About the textbook

Welcome to Plants in Action. This online text book, produced by the Australian and New Zealand societies of plant sciences, makes high-quality, cutting-edge, peer-reviewed research freely available to users across the world.

Edition 2, Plants in Action. You are now on the home page of Edition 2. Ten of the original twenty chapters have been fully revised, as shown on the navigation panel.

Edition 1, Plants in Action is also available free and on-line at http://plantsinaction.science.uq.edu.au/edition1. Twenty chapters provide over a thousand illustrations designed for teaching that are easily downloaded.

If you wish to contact the editorial team with queries or suggestions, please email the Australian Society of Plant Scientists: admin@asps.org.au. To use figures or text for non-commercial purposes, please acknowledge the source as: "Reproduced from Plants in Action, http://plantsinaction.science.uq.edu.au, published by the Australian Society of Plant Scientists." Reproduction of this material in another language or for commercial purposes requires permission.

Editorial committee


Professor Rana Munns is an honorary fellow at CSIRO Agriculture in Canberra, and professor in the School of Plant Biology, and the ARC Centre of Excellence in Plant Energy Biology, at the University of Western Australia. She uses physiological insights and molecular genetics to improve growth and yield of crop plants in dry or saline soils. Rana is a Fellow of the Australian Academy of Science, and an editor of PrometheusWiki, the plant methods wiki.

Susanne Schmidt2lt-115px.jpg

Professor Susanne Schmidt is a researcher and educator at The University of Queensland and Alexander von Humboldt fellow. Susanne has a passion for ecophysiology and plant nutrition and leads a vibrant group researching plant-microbe-soil interactions. Fundamental to applied research addresses environmental problems at the interface of ecology and agriculture.


Professor Christine Beveridge is a researcher and educator at The University of Queensland and current Future Fellow of the Australian Research Council. Christine’s research team uses advanced molecular, physiology and computer modelling technologies to understand how mobile plant signals and resources interact to control shoot and root architecture and development.

Contents Page

Chapter 1 - Light use and leaf gas exchange


Leaves come in a variety of shapes and sizes. Soybean (Glycine max) has a trifoliate leaf with broad laminae designed for capturing the maximum amount of light in a dense canopy.

John R Evans and Susanne von Caemmerer, Research School of Biology, Australian National University


Leaves come in a great variety of shapes and sizes. The photosynthetic processes that occur within leaves also show considerable variation. All of these variations represent different adaptive responses to different environmental conditions leading to altered gene expression.  

Despite such variation, leaves fulfil a common purpose: to capture energy from sunlight and convert that energy currency into chemically useful forms to drive CO2 assimilation and subsequent growth. CO2 assimilation broadly refers to the first steps in the production of sugars from CO2 and water, that is the initial incorporation of inorganic CO2 into biological molecules. Light absorption and energy utilisation is considered at progressively finer levels of organisation from leaves (Section 1.1) to chloroplasts (Section 1.2).

Section 1.1 encompasses anatomy, light interception and leaf gas exchange and includes a case study on development of a process-based model for photosynthetic CO2 assimilation using A:Ci curves.

1.1 - Leaf anatomy, light interception and gas exchange


Leaves experience a mix of demands under frequently adverse conditions. They must intercept sunlight and facilitate the uptake of CO2, which exists at levels around 390 ppm (µL L-1) in the atmosphere, while restricting water loss. The wide variety of shapes, sizes and internal structures of leaves imply that many solutions exist to meet these mixed demands.

In nature, photon irradiance (photon flux density) can fluctuate over three orders of magnitude and these changes can be rapid. However, plants have evolved with photosynthetic systems that operate most efficiently at low light. Such efficiency confers an obvious selective advantage under light limitation, but predisposes leaves to photodamage under strong light. How then can leaves cope? First, some tolerance is achieved by distributing light over a large population of chloroplasts held in architectural arrays within mesophyll tissues. Second, each chloroplast can operate as a seemingly independent entity with respect to photochemistry and biochemistry and can vary allocation of resources between photon capture and capacity for CO2 assimilation in response to light climate. Such features confer great flexibility across a wide range of light environments where plants occur and are discussed in Chapter 12.

Photon absorption is astonishingly fast (single events lasting 10–15 s). Subsequent energy transduction into NADPH and ATP is relatively ‘slow’ (10–4 s), and is followed by CO2 fixation via Rubisco at a sedate pace of 3.5 events per second per active site. Distributing light absorption between many chloroplasts equalises effort over a huge population of these organelles, but also reduces diffusion limitations by spreading chloroplasts over a large mesophyll cell surface area within a given leaf area.  The internal structure of leaves (shown in the follwing section) reflects this need to maximise CO2 exchange between intercellular airspace and chloroplasts and to distribute light more uniformly with depth than would occur in a homogeneous solution of chlorophyll.

1.1.1 - Leaf Structure


Figure 1.1 A scanning electron micrograph of an uncoated and rapidly frozen piece of tobacco leaf showing a hairy lower leaf surface and cross-sectional anatomy at low magnification. Notional values for resistances to CO2 diffusion are given in units of m2 s mol-1. Corresponding values for CO2 concentration are shown in µL L-1. Ci is routinely inferred from gas exchange measurements and used to construct A:Ci curves for leaf photosynthesis. Scale bar = 100 µm. (Image courtesy J-W. Yu and J.Evans)

In a typical herbaceous dicotyledon (Figure 1.1) lower leaf surfaces are covered with epidermal outgrowths, known to impede movement of small insects, but also contributing to formation of a boundary layer. This unstirred zone of air immediately adjacent to upper and lower epidermes varies in thickness according to surface relief, area and wind speed. Boundary layers are significant in leaf heat budgets and feature in the calculation of stomatal and mesophyll conductances from measurements of leaf gas exchange.  

The diffusion of CO2 into leaves can be modelled using an analogue with electrical resistance (R) and conductance (the reverse of resistance), as in Figure 1.1, right hand side. This shows a series of resistances (r) that would be experienced by CO2 molecules diffusing from outside (ambient) air, through the boundary layer (b), the stomata (s), the intercellular airspaces (i), the cell walls and liquid phase (w) to fixed sites inside chloroplasts. These values emphasise the prominence of stomatal resistance within the series.

Corresponding values for CO2 concentration in ambient air (a), the leaf surface (s), the substomatal cavity (i), the mesophyll cell wall surface (w) to the sites of carboxylation with the chloroplasts (c) reflect photosynthetic assimilation within leaves generating a gradient for inward diffusion.

In transverse fracture as shown below in Figure 1.2(A) the bifacial nature of leaf mesophyll is apparent with columnar cells in the palisade layer beneath the upper surface and irregular shaped cells forming the spongy mesophyll below. Large intercellular airspaces, particularly in the spongy mesophyll, facilitate gaseous diffusion. The lower surface of this leaf is shown in Figure 1.2(B). On the left-hand side, the epidermis is present with its irregular array of stomata. Diagonally through the centre is a vein with broken-off hair cells and on the right the epidermis has been fractured off revealing spongy mesophyll cells beneath. Light micrographs of sections cut parallel to the leaf surface (paradermal) through palisade (C) and spongy (D) tissue reveal chloroplasts lying in a single layer and covering most of the internal cell wall surface adjacent to airspaces. Significantly, chloroplasts are rarely present on walls that adjoin another cell. Despite the appearance of close packing, mesophyll cell surfaces within the palisade layer are generally exposed to intercellular airspace. Inward diffusion of CO2 to chloroplasts is thereby facilitated. 

Leaves that develop in sunny environments and have high photosynthetic capacities are generally thicker than leaves from shaded environments. This is achieved with more elongate cells within the palisade layer and/or several layers of cells forming the palisade tissue. Thicker leaves in a sunny environment enable more Rubisco to be deployed which confers a higher photosynthetic capacity. Fitting more Rubisco into a unit of leaf area with good access to intercellular airspace requires an increase in mesophyll cell surface which is possible by increasing the thickness of the mesophyll tissue and hence leaf thickness. A thicker leaf in sunny environments is energy effective because enough photons reach chloroplasts in lower cell layers to keep their Rubisco gainfully employed. By contrast, in a shaded habitat, less Rubisco is required for a leaf with lower photosynthetic capacity and this can be fitted into thinner leaves.


Figure 1.2 A scanning electron micrograph of an uncoated and rapidly frozen piece of tobacco leaf fractured in (A) to reveal columnar mesophyll cells of the palisade layer beneath the upper leaf surface and spongy mesophyll in the lower half. Chloroplasts can be clearly seen covering the inner faces of cell walls. Looking onto the lower surface (B), the epidermis and stomata are present on the left side of the vein, whereas the epidermis was fractured away on the right side, revealing spongy mesophyll tissue. Light micrographs (C, D) of sections cut parallel to the leaf surface are shown for palisade (C) and spongy mesophyll (D) with solid lines showing where the paradermal sections align with (A). Chloroplasts form a dense single layer covering the cell surfaces exposed to intercellular airspace, but are rarely present lining walls where two cells meet. Scale bar in (A) = 50 µm and (B) = 200 µm.  (C) and (D) have same magnification as (A). (Images courtesy J. Evans and S.von Caemmerer)


1.1.2 - Light absorption


Figure 1.3 Light absorption by pigments in solution and by leaves. Absorbance (A) refers to attenuation of light transmitted through a leaf or a solution of leaf pigments, as measured in a spectrophotometer, and is derived from the expression A = log I0/I where I0 is incident light, and I is transmitted light. The solid curve (scale on right ordinate) shows absorbance of a solution of pigment—protein complexes equivalent to that of a leaf with 0.5 mmol Chl m-2. The dotted curve shows absorptance (scale on left ordinate), and represents the fraction of light entering the solution that is absorbed. Virtually all light between 400 and 500 nm and around 675 nm is absorbed, compared with only 40% of light around 550 nm (green). The dashed curve with squares represents leaf absorptance, which does not reach 1 because the leaf surface reflects part of the incident light. Leaves absorb more light around 550 nm than a solution with the same amount of pigment (75 versus 38%, respectively) because leaves scatter light internally. This increases the pathlength and thereby increases the probability of absorption above that observed for the same pigment concentration in solution. (Based on K.J. McCree, Agric Meteorol 9: 191-216, 1972; J.R. Evans and J.M. Anderson, BBA 892: 75-82, 1987)

Pigments in thylakoid membranes of individual chloroplasts (Figure 1.7) are ultimately responsible for strong absorption of wavelengths corresponding to blue and red regions of the visible spectrum (Figure 1.3). Irradiated with red or blue light, leaves appear dark due to this strong absorption, but in white light leaves appear green due to weak absorption around 550 nm, which corresponds to green light. Ultraviolet (UV) light (wavelengths below 400 nm) can be damaging to macromolecules, and sensitive photosynthetic membranes also suffer. Consequently, plants adapt by developing an effective sunscreen in their cuticular and epidermal layers. 

Overall, absorption of visible light by mesophyll tissue is complex due to sieve-effects and scattering. Sieve-effect is an outcome from packaging pigments into discrete units, in this case chloroplasts, while remaining leaf tissue is transparent. This increases the probability that light can bypass some pigment and penetrate more deeply. A regular, parallel arrangement of columnar cells in the palisade tissue with chloroplasts all vertically aligned means that about 80% of light entering a leaf initially bypasses the chloroplasts, and measurements of absorption in a light integrating sphere confirm this. Scattering occurs by reflection and refraction of light at cell walls due to the different refractive indices of air and water. Irregular-shaped cells in spongy tissues enhance scattering, increasing the path length of light travelling through a leaf and thus increasing the probability of absorption. Path lengthening is particularly important for those wavelengths more weakly absorbed and results in nearly 80% absorption, even at 550 nm (Figure 1.3). Consequently, leaves typically absorb about 85% of incident light between 400 and 700 nm; only about 10% is reflected and the remaining 5% is transmitted. These percentages do of course vary according to genotype x environment factors, and especially adaptation to aridity and light climate.

Sunlight entering leaves is attenuated with depth in much the same way as light entering a canopy of leaves shows a logarithmic attenuation with depth that follows Beer’s Law (Section 12.4). Within individual leaves, the pattern of light absorption is a function of both cell anatomy and distribution of pigments. An example of several spatial profiles for a spinach leaf is shown in Figure 1.4. Chlorophyll density peaks in the lower palisade layer and decreases towards each surface. The amount of light declines roughly exponentially with increasing depth through the leaf. Light absorption is then given by the product of the chlorophyll and light profiles. Light absorption initially increases from the upper surface, peaking near the base of the first palisade layer, then declines steadily towards the lower surface. Because light is the pre-eminent driving variable for photosynthesis, CO2 fixation tends to follow the light absorption profile (see 14C fixation pattern in Figure 1.4). However, the profile is skewed towards the lower surface because of a non-uniform distribution of photosynthetic capacity. Chloroplasts near the upper surface have ‘sun’-type characteristics which include a higher ratio of Rubisco to chlorophyll and higher rate of electron transport per unit chlorophyll. Chloroplasts near the lower surface show the converse features of ‘shade’ chloroplasts. Similar differences between ‘sun’ and ‘shade’ leaves are also apparent. Chloroplast properties do not change as much as the rate of absorption of light. Consequently, the amount of CO2 fixed per quanta absorbed increases with increasing depth beneath the upper leaf surface. The lower half of a leaf absorbs about 25% of incoming light, but is responsible for about 31% of a leaf’s total CO2 assimilation.


Figure 1.4 Profiles of chlorophyll, light absorption and photosynthetic activity through a spinach leaf. Cell outlines are shown in transverse section (left side).Triangles represent the fraction of total leaf chlorophyll in each layer. The light profile (dotted curve) can then be calculated from the Beer—Lambert law. The profile of absorbed light is thus the product of the chlorophyll and light profiles (solid curve). CO2 fixation, revealed by 14C labelling, follows the absorbed light profile, being skewed towards slightly greater depths. (Based on J.N. Nishio et al., Plant Cell 5: 953-961, 1993; J.R. Evans, Aust J Plant Physiol 22: 865-873, 1995)

1.1.3 - CO<sub>2</sub> diffusion to chloroplasts

Leaves are covered with a barrier or ‘cuticle’ on the outer walls of epidermal cells that is impermeable to both water and CO2. To enable CO2 entry into the leaf for photosynthesis, the epidermis is perforated by pores called stomata (Figure 1.5). As CO2 molecules diffuse inwards they encounter an opposite flux of H2O molecules rushing outwards that is three to four orders of magnitude stronger. This problem of transpirational water loss is a particular problem for plants in hot, dry climates, such as in most of Australia. Leaves control this gas exchange by adjusting the aperture of stomata which can vary within minutes in response to changes in several environmental variables including light, humidity and CO2 concentration (see Chapter 15 for more details). Air-spaces inside leaves are effectively saturated with water vapour (equivalent to 100% relative humidity at that leaf temperature) and because air surrounding illuminated leaves is almost universally drier, water molecules diffuse outwards down this concentration gradient from leaf to air. 


Figure 1.5 Diagram of a transverse section through an isolateral Eucalyptus pauciflora leaf which is normally pendulant. Palisade tissue occurs beneath both surfaces with spongy tissue and oil glands (not shown) in the middle. Putative pathways for diffusion of H2O out of substomatal cavities are shown by the solid curved arrows. CO2 diffuses inwards and H2O diffuses outwards in response to concentration differences between the leaf and air. Such gas exchange is restricted by a boundary layer (the unstirred layer of air at the leaf surface) and by stomata. One stoma is shown on each surface. CO2 diffusion continues inside the leaf mesophyll through airsapces between cells (curved dashed arrows) to reach cell walls adjacent to each chloroplast where CO2 dissolves and then diffuses into the chloroplast top reach the carboxylating enzyme Rubisco. Bundle sheath extensions (bottom of diagram) reach both epidermis and create an internal barrier to lateral diffusion. (Based on J.R. Evans et al., Planta 189: 191-200, 1993)

The diffusion pathway for H2O out of a leaf is usually divided into two parts, namely the boundary layer of still air at the leaf surface and stomatal pores (Figure 1.5). Boundary layer thickness depends on windspeed, leaf dimensions and the presence of surface structures (e.g. hairs in Figure 1.1). Positioning of stomata also varies between species. Leaves of terrestrial plants always have stomata on their lower (abaxial) surface but many species have stomata on both surfaces, especially if they have high photosynthetic rates and are in sunny locations such as pendulant leaves of eucalypts. Adaptations for arid environments include having surface structures like hairs and waxes, which increase the thickness of the boundary layer, and leaf rolling and encryption of stomata by placing them in crevices in the leaf surface. While these features restrict water loss, they also impose an increased resistance (decreased conductance) to CO2 uptake. 

The flux of water escaping from a leaf, called transpiration rate, can be understood from Fick’s law. It depends on the product between conductance and the gradient in water vapour from the inside of the leaf to the surrounding air. The vapour pressure gradient depends on both the humidity of the surrounding air and leaf temperature. Dry air (low humidity), or hotter leaf temperatures will result in greater transpiration rates for a given conductance. Maximum leaf conductance depends on the number and size of stomata per unit leaf area which is a leaf property that becomes fixed during development. However, the aperture of stomata can be varied, so stomatal conductance can vary over the timescale of minutes. Stomatal conductance responds to light, CO2 and humidity. The sensitivity of a leaf to these variables is not fixed but can change over time in response to, for example, drought. Transpiration rate can be measured by a variety of means. With the availability of portable instruments, it is now most commonly obtained by measuring the increase in water vapour content of air from a leaf enclosed in a chamber. Stomatal conductance can then be calculated from Fick’s law by dividing the transpiration rate by the vapour pressure gradient between the leaf and the air.

CO2 molecules diffusing inwards from ambient air to chloroplasts encounter restrictions additional to boundary layer and stomata (Figure 1.5). CO2 must also diffuse from substomatal cavities throughout the mesophyll, dissolve in wet cell walls, cross the plasma membrane to enter the cytosol, diffuse into chloroplasts across a double membrane (outer envelope in Figure 1.7) and finally reach fixation sites within the stroma of those chloroplasts. The combination of these restrictions from intercellular airspace to the sites of fixation within chloroplasts has been termed mesophyll conductance.

There is considerable variation in leaf anatomy and hence potential restriction to CO2 diffusion, but in general leaves with high rates of photosynthesis tend to have more permeable leaves (e.g. tobacco in Figure 1.2) and this complex anatomy ensures a greatly enlarged surface area for diffusion across interfaces. Indeed the total mesophyll cell wall area can be 20 times that of the projected leaf surface.

Chloroplasts tend to be appressed against cell walls adjacent to intercellular spaces (Figure 1.2 C, D) which improves access to CO2 and they contain carbonic anhydrase which speeds up diffusion of CO2 by catalysing interconversion of CO2 and bicarbonate within the stroma of chloroplasts. Although CO2 rather than HCO3 is the substrate species for Rubisco, the presence of carbonic anhydrase enables bicarbonate ions, which are more abundant under the alkaline conditions (pH 8.0) that prevail inside chloroplasts, to diffuse to Rubisco in concert with diffusion of CO2. By sustaining a very rapid equilibration between CO2 and HCO3 immediately adjacent to active sites on Rubisco, carbonic anhydrase enhances inward diffusion of inorganic carbon.

1.1.4 - Light and CO<sub>2</sub> effects on leaf photosynthesis


Figure 1.6 Photosynthetic response to photon irradiance for a Eucalyptus maculata leaf measured at three ambient CO2 concentrations, 140, 350 and 1000 µmol mol-1. Irradiance is expressed as µmol quanta of photosynthetically active radiation absorbed per unit leaf area per second, and net CO2 assimilation is inferred from a drop in CO2 concentration of gas passing over a leaf held in a temperature-controlled cuvette. CO2 evolution in darkness is shown on the ordinate as an extrapolation below zero. The irradiance at which net CO2 exchange is zero is termed the light compensation point (commonly 15-50 µmol quanta m-2 s-1, shade to sun species respectively). The initial slope of light-response curves for CO2 assimilation per absorbed quanta represents maximum quantum yield for a leaf. (Based on E. Ögren and J.R. Evans, Planta 189: 182-190, 1993)

Light impinging on plants arrives as discrete particles we term photons, so that a flux of photosynthetically active photons can be referred to as ‘photon irradiance’. Each photon carries a quantum of electromagnetic (light) energy. In biology the terms photon and quantum (plural quanta) tend to be used interchangeably.

CO2 assimilation varies according to both light and CO2 partial pressure. At low light (low photon irradiance in Figure 1.6) assimilation rate increases linearly with increasing irradiance, and the slope of this initial response represents maximum quantum yield (mol CO2 fixed per mol quanta absorbed). Reference to absorbed quanta in this expression is important. Leaves vary widely in surface characteristics (hence reflectance) as well as internal anatomy and chlorophyll content per unit leaf area. Therefore, since absorption of photosynthetically active quanta will vary, quantum yield expressed in terms of incident irradiance does not necessarily reflect the photosynthetic efficiency of the mesophyll. In the case of comparisons between sun and shade leaves, it has led to a widely held but mistaken belief that shade leaves (thinner and with higher chlorophyll content) are more efficient. Expressed in terms of absorbed quanta, sun and shade leaves have virtually identical quantum efficiencies for CO2 assimilation. 

Assimilation rate increases more slowly at higher irradiances until eventually a plateau is reached where further increases in irradiance do not increase the rate of CO2 assimilation (Figure 1.6). Chloroplasts are then light saturated. Absolute values for both quantum yield and light-saturated plateaux depend on CO2 concentration. Quantum yield increases as CO2 concentration increases as it competes more successfully with other species such as oxygen, at the binding site on Rubisco. Leaf absorptance has a hyperbolic dependence on chlorophyll content. For most leaves, 80–85% of 400–700 nm light is absorbed and it is only in leaves produced under severe nitrogen deficiency where there is less than 0.25 mmol Chl m–2 that absorptance falls below 75%. 

The plateau in Figure 1.6 at high irradiance is set by maximum Rubisco activity. With increasing CO2 partial pressure, the rate of carboxylation increases. The transition from light-limited to Rubisco-limited CO2 assimilation as irradiance increases becomes progressively more gradual at higher CO2 partial pressures. In part, this gentle transition reflects the fact that a leaf is a population of chloroplasts which have different photosynthetic properties depending on their position within that leaf. As discussed above, the profile of photosynthetic capacity per chloroplast changes less than the profile of light absorption per chloroplast (Figure 1.4). This results in an increase in CO2 fixed per quanta absorbed with increasing depth. A transition from a light to a Rubisco limitation therefore occurs at progressively higher incident irradiances for each subsequent layer and results in a more gradual transition in the irradiance response curve of a leaf compared to that of a chloroplast. 

Photosynthetic capacity of leaves varies widely according to light, water and nutrient availability and these differences in capacity usually reflect Rubisco content. Leaves in high light environments (‘sun’ leaves) have greater CO2 assimilation capacities than those in shaded environments and this is reflected in the larger allocation of nitrogen-based resources to photosynthetic carbon reduction (PCR cycle; Section 2.1). Sun leaves have a high stomatal density, are thicker and have a higher ratio of Rubisco to chlorophyll in order to utilise the larger availability of photons (and hence ATP and NADPH). Shade leaves are larger and thinner, but have more chlorophyll per unit leaf dry weight than sun leaves. They can have a greater quantum yield per unit of carbon invested in leaves, but with a relatively greater allocation of nitrogen-based resources to photon capture, shade leaves achieve a lower maximum rate of assimilation.

Despite such differences in leaf anatomy and chloroplast composition, leaves sustain energy transduction and CO2 fixation in an efficient and closely coordinated fashion. Processes responsible are discussed below (Section 1.2).

Case Study 1.1 - Development of <em>A:C<sub>i</sub></em> curves

Susanne von Caemmerer, Research School of Biology, Australian National University

CO2 assimilation rate at a whole-leaf level can be analysed in terms of the underlying biochemistry. Traditionally, photosynthesis has been divided into light and dark reactions. The light reactions describe photosynthetic electron flow which generates reducing power (NADPH) and the formation of ATP. The dark reactions consist of the photosynthetic carbon reduction and oxidation cycles which start with Rubisco as the primary catalyst.

In this essay A:Ci refers to CO2 assimilation rate (A) as a function of intercellular CO2 (Ci) which can either be expressed in terms of concentration (µL of CO2 per litre of gas, µL L–1, or ppm) or partial pressure (µbar, or Pa). Multiplying concentration by atmospheric pressure converts it to partial pressure (e.g. 400 µL L–1 x 0.95 bar = 380 µbar). Partial pressures are preferred as this is the form that relates best to Rubisco performance and takes into account the altitude where the measurement was made. At sea level where atmospheric pressure averages one bar, the values for concentration and partial pressure are the same. A:Ci curves are created by measuring A in various atmospheric CO2 concentrations.

Physical concepts of leaf gas exchange

Penman and Schofield (1951) put diffusion of CO2 and water vapour through stomata on a firm physical basis. Their ideas were taken up at Wageningen by Pieter Gaastra in the 1950s and modern analytical gas exchange is often attributed to this seminal work (Gaastra 1959) where he even constructed his own infrared gas analyser and other equipment necessary to make measurements of CO2 and water vapour exchange. His work was a landmark because it examined CO2 assimilation and water vapour exchange rates of individual leaves under different environmental conditions, and he distinguished between stomatal and internal resistances. Gaastra calculated resistances to water vapour and CO2 diffusion from two equations (here in our simplified notation) which are based on Fick’s Law for the diffusion of gases.

\[ E=\frac{w_i-w_a}{r_{sw}} \text{ and } A=\frac{C_a-C_i}{r_{sc}} \tag{1} \]

where E and A are the fluxes of water vapour and CO2 and \(w_i\) and \(c_i\) and \(w_a\) and \(c_a\) are the mole fractions of water vapour and CO2 in intercellular air spaces and ambient air respectively. The denominator terms, \(r_{sw}\) and \(r_{sc}\), represent stomatal resistances to H2O and CO2 diffusion respectively. Gaastra assumed that \(w_i\) was equivalent to the saturated vapour pressure at the measured leaf temperature. By rearranging equation 1, \(r_{sw}\) could be calculated:

\[ r_sw=\frac{w_i-w_a}{E} \tag{2} \]

Knowing that resistances to CO2 and water vapour are related by the ratio of their diffusivities, he calculated stomatal resistance to CO2 diffusion, \(r_{sc}\). Gaastra realised that the diffusion path for CO2 is longer than that of water vapour, as CO2 had to diffuse from the intercellular airspaces through the cell wall across membranes to the chloroplast stroma where CO2 fixation by Rubisco takes place. He therefore extended the equation for CO2 assimilation to:

\[ A=\frac{C_a-C_c}{r_{sc}+r_m} \tag{3} \]

where Cc represented CO2 concentration in the chloroplasts.

Gaastra analysed the dependence of CO2 assimilation rate on light, CO2 and temperature, and observed that at low CO2 concentrations the rate of CO2 assimilation was independent of temperature whereas it was strongly influenced by temperature at higher CO2 concentrations. This led him to conclude that the rate of CO2 uptake was completely limited by CO2 diffusion processes at low CO2 and that biochemical processes became limiting only at high CO2. The belief that CO2 diffusion was limiting gave rise to the assumption that chloroplastic CO2 concentration was close to zero. This led to the erroneous simplification of the above equation such that the total resistance to CO2 diffusion could be calculated from CO2 assimilation rate and the ambient CO2 concentration alone. Since stomatal resistances could be calculated from measurements of water vapour diffusion, it was also possible to calculate mesophyll resistance to CO2 diffusion. In Australia particularly, there was great interest in determining the relative importance of stomatal versus mesophyll resistance in limiting CO2 assimilation rates under adverse conditions of high temperature and water stresses. In global terms, much of the pioneering work was undertaken in this country (see, for example, Bierhuizen and Slatyer 1964).

Calculation of intercellular CO2, Ci and the first A versus Ci curves


Figure 1.  An early A:Ci curve showing the CO2 assimilation rate of cotton at a range of cell wall CO2 concentrations (redrawn from Troughton and Slayter (1969) and retaining original units for CO2 flux). For comparative purposes, 10 × 10-8 g cm-2 s-1 would be equivalent to 22.27 µmol CO2 m-2 s-1, and 1 µg L-1 would be equivalent to 0.54 µL L-1 (assuming a gram molecular weight of 44 for CO2, and measurements at normal temperature and pressure). (a) Leaf temperature influences the overall shape of CO2 response curves (measured in O2-free air) but has no effect on the initial slope where response to CO2 is limited by Rubisco activity. This family of curves comes from repeated measurements of gas exchange by the same leaf at five different temperatures (values shown) and indicated in the figure by five different symbols. (b) CO2 response curves for two leaves of cotton measured in O2-free air at 25°C and three levels of relative water content. Legend: ● leaf 1, 92% water content; O leaf 1, 56%; ▲ leaf 2, 92%; Δ leaf 2, 69%. Identical slopes regardless of treatment mean that variation in relative water content over this range is without effect on CO2 assimilation within mesophyll tissues. By implication, reduction in CO2 uptake as commonly observed on whole leaves under moisture stress would be attributable to stomatal factors.

Although CO2 concentration in intercellular airspaces, Ci, was explicit in Gaastra’s equations, this term was first specifically calculated by Moss and Rawlings in 1963, and the first extensive use of the parameter was made by Whiteman and Koller in 1967, who examined stomatal responses to CO2 and irradiance, concluding that stomata were more likely to respond to Ci rather than Ca. The first bona fide response curves of CO2 assimilation rate to Ci rather than Ca were those of Troughton and Slatyer (1969) (Figure 1). In Figure 1(a), Ci was derived from measurements of CO2 uptake in an assimilation chamber where air passed through a leaf, rather than over both surfaces concurrently (as became commonplace in subsequent designs), and such estimates would differ slightly. More importantly, those measurements were made at different temperatures and confirmed that CO2 assimilation was not greatly affected by temperature at low Ci. Later, this lack of temperature dependence was explained by the kinetics of Rubisco (von Caemmerer and Farquhar 1981). Figure 1(b) shows the initial slope of CO2 response curves measured at different stages of water stress. In this case, water stress has affected stomatal resistance (as the Ci obtained at air levels of CO2 occur at progressively lower Ci) but not the relationship between CO2 assimilation rate and Ci. A versus Ci response curves thus provided an unambiguous distinction between stomatal and non-stomatal effects on CO2 assimilation and, provided stomata respond uniformly across both leaf surfaces, that distinction can be made quantitative.

Before we head further into a discussion of our understanding and interpretation of more comprehensive CO2 response curves, we must take an important digression into development of mathematical models of C3 photosynthesis.

Biochemistry of photosynthesis and leaf models

Gas exchange studies focused initially on physical limitations to diffusion, but it was not long before persuasive arguments were being brought forward to show that leaf biochemistry must influence the rate of CO2 fixation even at low CO2 concentrations. Björkman and Holmgren (1963) made careful gas exchange measurements of sun and shade ecotypes of Solidago growing in Sweden, and noted strong correlations between photosynthetic rate measured at high irradiance and ambient CO2 and the nitrogen content of leaves, and later also related it to different concentrations of Rubisco (then called carboxydismutase). Anatomical studies implied that thin shade leaves would have less internal diffusion resistance to CO2 than thicker sun leaves where cells were more densely packed, but the opposite was observed. Furthermore, following earlier discoveries that CO2 assimilation rate was enhanced under low-O2, Gauhl and Björkman (1969), then at Stanford, showed very elegantly that O2 concentration affected CO2 assimilation rate but not water vapour exchange (i.e. stomata did not respond to a change in O2). Clearly, the increase in CO2 assimilation rates seen with a decrease in O2 concentration could not be explained via a limitation on CO2 diffusion.


Figure 2. Comparison of measured and modelled CO2 response curves. (a) CO2 assimilation rate (A) v. intercellular CO2 partial pressure (Ci) in Phaseolus vulgaris measured at two irradiances and a leaf temperature of 28°C. Arrows indicate points obtained at an external CO2 partial pressure of 330 µbar, which was the ambient CO2 partial pressure in Canberra around 1980. (b) Modelled CO2 response curves. The solid curve extending from the x axis represents the Rubisco-limited rate of CO2 assimilation.

\[ A=\frac{\left(C_i - \Gamma_{*} \right) V_{cmax}}{C_i + K_c \left( 1 + O / K_o \right)} -R \]

The dashed lines and their extensions represent the electron-transport-limited rates of CO2 assimilation at the two irradiances.

\[ A=\frac{J \left( C_i - \Gamma_{*} \right)}{4.5C_i + 10.5 \Gamma} - R \]

For further details, see von Caemmerer and Farquhar (1981). (c) CO2 assimilation rate v. intercellular CO2 partial pressure in Phaseolus vulgaris measured at two O2 partial pressures at a leaf temperature of 28°C. Arrows indicate points obtained at an external CO2 partial pressure of 330 µbar. (d) Modelled CO2 response curves for conditions applied in (c) using the equations given in (b).

Central importance of Rubisco

Early mathematical models of leaf photosynthesis were extensions of Gaastra’s resistance equation, and could not accommodate the O2 sensitivity of CO2 assimilation. They were quickly followed by development of more biochemical models in the early 1970s and the discoveries by Bowes et al. (1971) that Rubisco was responsible for both carboxylation and oxygenation of RuBP (a five-carbon phosphorylated sugar, regenerated by the photosynthetic carbon reduction (PCR) cycle of chloroplasts). This crucial observation of dual function put Rubisco at centre stage. Laing et al. (1974) were first to compare the gas exchange of soybean leaves with the in vitro kinetics of Rubisco and suggested the following equation for the net CO2 assimilation rate:

\[ A=V_c \left(1-0.5\frac{V_o}{V_c} \right) \tag{4} \]

where \(V_c\) and \(V_o\) are the rates of Rubisco carboxylation and oxygenation (later on a term for mitochondrial respiration was added to most models). Laing et al. related a ratio of the rates of carboxylation to oxygenation of RuBP to the concentration of its substrates, CO2, \(C\), and O2, \(O\), and showed that:

\[ \frac{V_o}{V_c} = \frac{V_{omax} K_c}{V_{cmax} K_o} \frac{O}{C} = \frac{2 \Gamma_{*}}{C} \tag{5} \]

where \(K_c\), \(K_o\), \(V_{cmax}\), \(V_{omax}\) are the corresponding Michaelis Menten constants and maximal activities of carboxylase and oxygenase functions respectively and \(\Gamma_{*}\)is the CO2 compensation point in the absence of mitochondrial respiration.

A note on \(\Gamma\): illuminated leaves held in a closed circuit of recirculating air will reduce CO2 to a ‘compensation point’ where uptake and generation of CO2 are balanced; this is commonly 50–100 ppm for C3 plants and referred to as \(\Gamma\). A CO2 response curve for leaf photosynthesis will show a similar value as an intercept on the abscissa. \(\Gamma\) can thus be measured empirically, and will be an outcome of interactions between photosynthesis, photorespiration and dark (mitochondrial) respiration (R). If allowance is made for R, the CO2 compensation point would then be slightly lower, and is termed \(\Gamma_{*}\). As with measured \(\Gamma\), this inferred CO2 compensation point, \(\Gamma_{*}\) , is linearly related to O2, an observation that intrigued earlier observers but was easily reconciled with the dual function of Rubisco. Laing et al. (1974) used Equations 4 and 5 to predict this linear dependence of \(\Gamma_{*}\) on O2, and with subsequent confirmation Rubisco became a key player in photosynthetic models. (Equation 4 assumes that for each oxygenation, 0.5 CO2 are evolved in the subsequent photorespiratory cycle, although there has been some debate over this stoichiometry.) If the enzyme reaction is ordered with RuBP binding first, the rate of carboxylation in the presence of the competitive inhibition by O2 at saturating RuBP concentration can be given by

\[ V_c=\frac{CV_{cmax}}{C+K_c \left( 1+O⁄K_o \right)} \tag{6} \]

When combined with Equation 4 this gave a simple expression of net CO2 fixation rate:

\[ A = \frac{\left(C_i - \Gamma_{*} \right) V_{cmax}}{C_i + K_c \left( 1 + O / K_o \right)} \tag{7} \]

which depends on the maximal Rubisco activity and provided the quantitative framework for comparing rates of CO2 assimilations with the amount of Rubisco present in leaves (von Caemmerer and Farquhar 1981). Difference in CO2 assimilation rates observed under different growth conditions could then be explained according to variations in the amount of Rubisco present in leaves. In Figure 2 the dotted line shows a CO2 response curve modelled by Equation 7. Chloroplast CO2 partial pressure was then assumed to be similar to that in the intercellular airspaces. Using on-line discrimination between 13CO2 and 12CO2, and deriving an estimate of CO2 partial pressure at fixation sites within chloroplasts, we subsequently learned that a further draw down can occur, but the general applicability of Equation 7 was not compromised. As an aside, these equations became basic to most photosynthetic models long before the order of the reaction mechanism of Rubisco had been unequivocally established. Had CO2 and O2 bound to Rubisco before RuBP, or the reaction not been ordered, our equations would have been much more complex with both Km(CO2) and Km(O2) dependent upon RuBP concentration.


Figure 3. Transgenic tobacco with reduced amount of Rubisco shows no limitation by the rate of RuBP regeneration. CO2 assimilation response curves in wild-type tobacco ■ and in transgenic tobacco with reduced amount of Rubisco □, were measured at a photon irradiance of 1000 µmol quanta m-2 s-1 and a leaf temperature of 25°C. Lines show Rubisco-limited rates of CO2 assimilation (see legend to Figure 2). The reduction in Rubisco in transgenic tobacco was achieved with an antisense gene directed against the mRNA of the Rubisco small subunit (Hudson et al. 1992). Arrows indicate the points obtained at an external CO2 partial pressure of 350 µbar.

Regeneration of RuBP and electron transport rate

Equation 7 could mimic CO2 assimilation rate at low Ci , as well as O2 effects on CO2 uptake, but measured rates of CO2 assimilation saturated much more abruptly at high CO2 concentrations than could be predicted from Rubisco kinetics (Figure 2). Using a novel approach in Estonia, Laisk and Oja (1974) proposed that CO2 assimilation was limited by RuBP regeneration rate at high Ci. They had fed brief pulses of CO2 to leaves that had been previously exposed to low CO2 (conditions under which RuBP concentrations were presumably high), and obtained rates up to 10 times higher than the steady-state rates of CO2 assimilation! Lilley and Walker (1975) at Sheffield reached a similar conclusion after comparing the CO2 responses of illuminated isolated chloroplasts with those obtained upon lysing chloroplasts in a medium containing saturating RuBP.

In our model of C3 photosynthesis (Farquhar et al. 1980), the way we handled rate limitation by RuBP regeneration was probably the most important decision made in that context. Both ATP and NADPH were required for RuBP regeneration, and this fundamental need formed a connection with light in our model. From a mathematical perspective there were two options: (1) RuBP and CO2 could always colimit the rate of carboxylation, and this we would express in a double Michaelis Menten equation, or (2) carboxylation rate could be limited by either RuBP or else be saturated and thus independent of RuBP. The in vivo kinetics of Rubisco suggest the second option.

Peisker (1974) and Farquhar (1979) pointed out that Rubisco was unusual in that it was present in the chloroplast at very high concentrations. Given such a low Km(RuBP), this meant that the in vivo kinetics with respect to chloroplastic RuBP were those of a tight binding substrate. That is, the rate of Rubisco would depend linearly on RuBP concentration when chloroplastic RuBP concentration was below Rubisco catalytic site concentration, and once RuBP exceeded Rubisco site concentration carboxylase would be RuBP saturated. We also knew that irradiance affected CO2 assimilation rate mainly at high intercellular CO2. This supported option 2 (see Figure 2a, b). Given these insights, the more complex link between chloroplastic electron transport rate and RuBP pools used by Farquhar et al. (1980) was quickly simplified to a description of CO2 assimilation that was limited by RuBP regeneration, and utilisation of ATP and NADPH for photosynthetic carbon reduction or oxygenation. RuBP regeneration was in turn driven by the electron transport rate, J (dependent on irradiance and its own maximal capacity), and stoichiometry of ATP or NADPH use by the photosynthetic carbon reduction and oxygenation cycle. For example, when electron transport rate, \(J\), was limiting (in view of ATP use) carboxylation rate could proceed at:

\[ V_c = \frac{J}{4.5 + 10.5 \Gamma_{*} / C} \tag{8} \]

Dashed lines in Figure 2 give modelled electron-transport-limited rates of CO2 fixation according to:

\[ A = \frac{J \left( C_i - \Gamma_{*} \right)}{4.5C_i + 10.5 \Gamma} \tag{9} \]

This simplified formulation of C3 photosynthesis (Equations 7 and 9) now provides a meaningful framework for analysis of leaf photosynthesis, and has focused our interpretation of CO2 response curves on leaf biochemistry. For example, von Caemmerer and Farquhar (1981) related the initial slopes of CO2 response curves to in vitro Rubisco activity, and the CO2-saturated rates of A:Ci curves to in vitro measurements of electron transport rates. Such studies validate Equations 7 and 9, demonstrating that CO2 response curves could be used as a meaningful and non-invasive tool to quantify these biochemical components under a wide variety of conditions. Subsequent comparisons between wild-type tobacco and transgenic tobacco with a reduced amount of Rubisco have confirmed our concepts. When Rubisco alone is reduced in transgenic plants, RuBP regeneration capacity remains unchanged and no longer limits the rate of CO2 assimilation at high CO2. Rubisco then constitutes the sole limitation (Figure 3).


Both Rubisco and electron transport components are expensive in terms of leaf nitrogen. For example, Rubisco represents up to 25% of a leaf’s protein nitrogen, with energy transduction components a further 25%. At a Ci where the transition from a Rubisco limitation to RuBP regeneration limitation occurs, both capacities are used efficiently and colimit net CO2 assimilation. That is, assimilation can only be increased if both sets of component processes are increased. Where then should the balance lie if a plant is to use nitrogen-based resources to best effect? The transition obviously varies with irradiance and temperature so that an optimal balance will vary with habitat. However, surprisingly little variation has been observed and plants appear unable to shift this point of balance. As an example, important in the context of rising atmospheric CO2 concentrations, plants grown in a high CO2 environment should manage with less Rubisco and thus put more nitrogen into the capacity of RuBP regeneration. Surprisingly, such adjustments have not been observed experimentally, but given prospects of global change, our need for understanding gains urgency.


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Whiteman PC, Koller D (1967) Interactions of carbon dioxide concentration, light intensity and temperature on plant resistance to water vapour and carbon dioxide diffusion. New Phytol 66: 463–473

1.2 - Chloroplasts and energy capture


Chloroplasts dividing (dumbbell figures) within an enlarging cell of a young spinach leaf, resulting in about 200 chloroplasts per cell at leaf maturity. (Micrograph courtesy John Possingham: Nomarski optics)

In thermodynamic terms, O2-generating photosynthesis in vascular plants is an improbable process! Improbable, because a weak oxidant (CO2) must oxidise a weak reductant (H2O), thereby producing a strong oxidant (O2) and a strong reductant (carbohydrate). To achieve this ‘uphill’ reaction, a massive and continuous input of chemical energy is required. However, in nature, only radiant energy is available on that scale. How then can green plants achieve this conversion? Chloroplasts are responsible, and in the most significant process in our biosphere, photosynthetically active quanta are trapped and converted into chemically usable forms. This captured energy sustains plant growth and provides a renewable resource base for life on earth.

Thanks to the pioneering work of Calvin and Benson at Berkeley on 14CO2 fixation products by Chlorella which began in the 1950s, biochemical aspects of photosynthetic carbon reduction (Calvin cycle) are now comprehensively understood. The transduction of light energy into chemical potential energy is not so well understood, while events surrounding photosynthetic electron flow are defined in some detail and are described here, biophysical processes within the water-splitting apparatus of chloroplasts, and indeed the manner in which photons are captured and their quantum energy harnessed for photolysis, remain something of an enigma and fall outside the scope of our present account.

1.2.1 - Chloroplast structure and composition


Figure 1.7 A mature and functional chloroplast in an immature leaf of bean (Phaseolus vulgaris) with an extensive network of photosynthetic membranes (thylakoids), parts of which are appressed into moderate granal stacks, and suspended in a gel-like matrix (stroma).The chloroplast containing a pair of starch grains (S) is encapsulated in a double membrane (envelope) and suspended within a granular cytoplasmic matrix adjacent to a mitochondrion (M) and in close proximity to the cell wall (CW). Scale bar = 1 µm. (Micrograph courtesy S. Craig and C. Miller)

Chloroplasts are easily recognised under a light microscope in leaf sections as distinctive green organelles suspended in the cytoplasm and usually appressed against cell walls. Chloroplasts are abundant in mesophyll tissue (commonly 200–300 in each palisade cell) and functional organelles can be isolated from homogenates of leaf tissue. 

Chloroplasts are surrounded by a double membrane, or envelope, just visible in transmission electron micrographs (Figure 1.7). This envelope encapsulates a soluble (gel-like) stroma which contains all the enzymes necessary for carbon fixation, many enzymes of nitrogen and sulphur metabolism and the chloroplast’s own genetic machinery. 

The inner membrane of a chloroplast envelope is an effective barrier between stroma and cytoplasm, and houses transporters for phosphate and metabolites (Section 2.1.8) as well as some of the enzymes for lipid synthesis. By comparison, the outer membrane of the chloroplast envelope is less complex and more permeable to both ions and metabolites. 

Suspended within the stroma, and entirely separate from envelope membranes, is an elaborately folded system of photosynthetic membranes or ‘thylakoids’ (literally ‘little sacs’). Embedded within these membranes are the complexes that enable light harvesting and electron flow from H2O molecules to NADP+, thereby converting light energy into chemically usable forms. There are four basic complexes comprising two types of photosystem (with interlinked protein and pigment molecules), cytochrome b/f complexes (pivotal for photosynthetic electron transport) and ATP synthase complexes (responsible for proton egress from thylakoid lumen to stroma, and consequent ATP generation). These complexes are densely packed within the thylakoids. This remarkable transduction of energy, with such profound implications for life as we know it, starts with selective absorption of incoming light by chlorophylls and accessory pigments (certain carotenoids) that operate within both photosystems.

1.2.2 - Chlorophyll absorption and photosynthetic action spectra


Figure 1.8 Upper curves: Diethylether solutions of chlorophyll a (Chl a, solid line) and chlorophyll b (Chl b, dotted line) show distinct absorption peaks in
 blue and in red regions of the visible spectrum (redrawn from Zscheile and Comar’s (1941) original data). Fluorescence emission spectra (inset, redrawn from Lichtenthaler 1986) show peaks only in red, and at wavelengths characteristically longer than corresponding absorption peaks, namely 648 cf. 642 nm for Chl b, and 668 cf. 662 nm for Chl a. Lower curves: In situ absorption spectra (eluted from gel slices) for pigment-protein complexes corresponding to photosystem II reaction centre (PSII RC) and light-harvesting chlorophyll (a,b)-protein complexes (LHC). A secondary peak at 472 nm and a shoulder at 653 nm indicate contributions from Chl b to these broadened absorption spectra which have been normalised to 10 µM Chl solutions in a 1 cm path length cuvette. (Based on J.R. Evans and J.M. Anderson, BBA 892: 75-82, 1987)

Chlorophylls are readily extracted from (soft) leaves into organic solvent and separated chromatographically into constituent types, most notably chlorophyll a (Chl a) and chlorophyll b (Chl b). These two chemical variants of chlorophyll are universal constituents of wild vascular plants and express highly characteristic absorption spectra (Figure 1.8, upper curves). Both chlorophylls show absorption maxima at wavelengths corresponding to blue and red, but chlorophyll assay in crude extracts, which inevitably contain carotenoids as well, is routinely based on absorption maxima in red light to avoid overlap with these accessory pigments that show strong absorption below 500 nm. Absorption maxima at 659 and 642 for Chl a and Chl b respectively would thus serve for assay in diethylether, but these peaks will shift slightly according to solvent system, and such shifts must be taken into account for precise measurement (see Porra et al. 1989 for details). Additional chlorophylls have been discovered that exist in cyanobacteria which extends their absorption spectrum into the infrared (Figure 1.9).

Chl a and Chl b differ with respect to both role and relative abundance in higher plants. Chl a/b ratios commonly range from 3.3 to 4.2 in well-nourished sun-adapted species, but can be as low as 2.2 or thereabouts in shade-adapted species grown at low light. Such variation is easily reconciled with contrasting functional roles for both Chl a and Chl b. Both forms of chlorophyll are involved in light harvesting, whereas special forms of only Chl a are linked into energy-processing centres of photosystems. In weak light, optimisation of leaf function calls for greater investment of leaf resources in light harvesting rather than energy processing. As a result the relative abundance of Chl b will increase and the Chl a/b ratio will be lower compared with that in strong light. Conversely, in strong light, photons are abundant and require greater capacity for energy processing by leaves (hence the higher Chl a/b ratio).  As a further subtlety, the two photosystems of higher plant chloroplasts (discussed later) also differ in their Chl a/b ratio, and this provided Boardman and Anderson (1964) with the first clue that they had achieved a historic first in the physical separation of those two entities. 

Carotenoids also participate in photosynthetic energy transduction. Photosystems have an absolute requirement for catalytic amounts of these accessory pigments, but their more substantive involvement is via dissipation of potentially harmful energy that would otherwise impact on delicate reaction centres when leaves experience excess photon irradiance (further details in Chapter 12). Carotenoids are thus regarded as ‘accessory’ to primary pigments (chlorophylls) and in molar terms are present in mature leaves at about one-third the abundance of Chl (a + b). 

Chlorophyll in leaves is not free in solution but is held in pigment-protein complexes, each with a different absorption spectrum (see Evans and Anderson 1987). In particular, light-harvesting Chl a, b–protein complexes (LHC in Figure 1.8, lower curves) develop a secondary absorption peak at 472 nm with a shoulder at 653 nm, while the Chl a of photosystem II reaction centres shows absorption peaks at 437 and 672 nm (compared with 429 and 659 nm for purified Chl a in ether; Figure 1.8, upper curves).

Subtle alterations in the molecular architecture of chlorophyll molecules according to the particular protein to which they bind in either light-harvesting or energy-processing centres are responsible for these shifts in absorption peaks, and for a general broadening of absorption spectra (compare lower and upper curves in Figure 1.8). Such effects are further accentuated within intact leaves by accessory pigments and greatly lengthened absorption pathways resulting in about 85% of visible wavelengths being absorbed (Figure 1.10). Any absorbed quanta at wavelengths below 680 nm can drive one electron through either reaction centre. Maximum quantum yield (Figure 1.10) occurs when both reaction centres absorb equal numbers of such quanta. When one photosystem population (PSII) absorbs more quanta than the other (PSI), excess quanta cannot be used to drive whole-chain (linear) electron flow. Quantum yield is reduced as a consequence, and leads to a slight discrepancy between in vivo absorption maxima (Figure 1.8) and quantum yield (Figure 1.10).

Although UV wavelengths are absorbed by leaves and would be capable of driving photosynthesis, such short wavelengths are damaging to biological systems and plants have adapted by developing a chemical sunscreen. Consequently, the quantum yield from these wavelengths drops off markedly below about 425 nm. Beyond 700 nm (infrared band) absorption drops to near zero, and forestalls leaf heating from this source of energy. However, quantum yield falls away even faster, and this ‘red drop’, though puzzling at first, led subsequently to a comprehensive model for photosynthetic energy transduction, outlined below.


Figure 1.9 Absorption spectra for the four types of chlorophyll found in photosynthetic organisms with respect to the visible spectrum. Chlorophyll d and f are found in a cyanobacteria which allows it to utilise infrared light between 700-750 nm, beyond the range normally absorbed by photosynthetic organisms. The chlorophylls are dissolved in methanol which alters their spectra compared to in vivo. The extinction coefficients for the long wavelength peak of each chlorophyll are: Chl a 665.5 nm 71.4 L mmol-1 cm-1, Chl b 652 nm 38.6 L mmol-1 cm-1, Chl d 697 nm 63.7 L mmol-1 cm-1, Chl f 707 nm 71.1 L mmol-1 cm-1. (Based on Chen and Blankenship, Trends Plant Sci 16: 427-431, 2011; Li et al., BBA Bioenergetics, 2012; Porra et al., BBA Bioenergetics 975: 384-394, 1989).


Figure 1.10 Leaves absorb visible light very effectively (85% for the waveband between 400 and 700 nm; solid curve).Wavelengths corresponding to green light are  absorbed less effectively (absorptance drops to c. 0.75). Beyond 700 nm  (infrared band) absorptance drops to near zero, and forestalls leaf heating  from this source of energy. Quantum yield is referenced to values obtained  in red light (600-625 nm), which is most effective in driving photosynthesis, requiring about 10 quanta per CO2 assimilated (based on high-precision leaf  gas exchange) compared with about 12 quanta at the blue peak (450 nm). Quantum yield shows a bimodal response to wavelength. Absorptance drops  beyond 700 nm but quantum yield drops off even faster because PSII (responsible for O2 generation) absorbs around 680 nm and cannot use quanta at longer wavelengths in this measuring system. UV wavelengths (below 400 nm) are capable of driving photosynthesis, but as a protective adaptation  vascular plants accumulate a chemical ‘sunscreen’ in response to UV exposure. Field-grown plants are especially rich in these substances so that  absorbed UV is dissipated harmlessly, lowering quantum yield compared  with growth-chamber plants. (Based on K.J. McCree, Agric Meteorol 9: 191-216, 1972)

1.2.3 - Cooperative photosystems and a ‘Z’ scheme for electron flow

Plants and many algae contain two distinct protein complexes for trapping and processing photons of light; photosystems I and II (PSI and PSII). These two systems can be separated and identified using a combination of biochemical and chemical techniques. Within the chloroplast, however, these two systems must work cooperatively and sequentially to absorb photons and convert their quantum energy into a flow of electrons. Interestingly, although PSI was discovered first, in cyanobacteria, photosynthetic electron flow is initiated in PSII and then proceeds to PSI. In PSII electrons are provided through the splitting of water molecules. PSI is responsible for finally delivering these electrons to NADH+.

This section presents a historical account of the discovery of the two photosystems and how they work together to split water and produce NADH+.

Prior to the advent of high-precision leaf gas exchange methods (as employed for Figure 1.10), O2 evolution was taken as a measure of photosynthetic activity. Action spectra were measured on a number of plants and algae over the range of visible radiation. A crucial and consistent observation was that O2 evolution dropped off much faster in the long-wavelength red region (>690 nm) than did absorption. Put another way, more quanta were being absorbed at longer wavelengths than could be used for photosynthesis. It seemed at these longer wavelengths as though a light absorber was being robbed of energy-processing capacity.

Anticipating that bimodal absorption implied a two-step process, and knowing that chlorophyll also absorbed photons at shorter wavelengths, Robert Emerson (working at Urbana in the mid-1950s) supplemented far-red light with shorter wavelength red irradiance and demonstrated that the relatively low photosynthetic rate in far-red light could be significantly increased. In fact the photosynthetic rate achieved with the two light qualities combined could be 30–40% higher than the sum of the rates in far-red or shorter red when measured separately (Emerson et al. 1957). This phenomenon became known as the ‘Emerson Enhancement Effect’ and contributed to a working hypothesis for photosynthetic energy conversion based upon two photochemical acts (proposed by Duysens et al. 1961), but additional lines of evidence were impacting on this outcome.

At about the same time as Emerson was establishing his enhancement effect, Myers and French observed ‘sequential enhancement’; that is, a disproportionate increase in photosynthetic rate or efficiency when the two light qualities were separated in time. The upper limits of dark intervals between two flashes of different light quality were 6 s for far-red after green and 1 min for green after far-red. Clearly, the ‘product’ of photochemical act 1 was stable for 1 min, that of act 2 for only 6 s. This discovery implied that chemical intermediates, rather than an altered physical state, were involved in a two-step cooperation (see Clayton 1980).

According to physical laws of photochemical equivalence, there should be a 1:1 yield in converting light energy to chemical energy by a perfect system. Quantum requirement for such events would be 1. However in photosynthesis the absolute quantum requirement for O2 is much greater than I. In the 1950s, Robert Emerson (at Urbana) and co-workers determined that 8-10 quanta were required. Hill and Bendell (1960) suggested a 'Z' scheme that was consistent with a requirment of 8-10 quanta, the cooperation of 2 quanta in the separation of one strong reducing and one strong oxidising equivalent, and the operation of two sequential photochemical acts. Figure 1.11 is a greatly developed version of their original model.


Figure 1.11. A highly diagrammatic zig-zag or ‘Z’ scheme of photosynthetic electron transport from water to NADP+ showing the sequence of electron/proton carriers and their association with either PSII or PSI. Linear electron flow is shown as solid lines; cyclic electron flow is indicated by dashed lines. All of these electron transport chains operate within thylakoid membranes with electron flow following a sequence dictated by redox potential (shown in volts on the ordinate). Cyclic electron flow in PSII diverts electrons from pheophytin to cytochrome b559 (and possibly back to P680+). Cyclic electron transport around PSI moves electrons from ferredoxin through cytochrome b565 and plastoquinone (PQ), while pseudocyclic electron transport takes electrons from ferredoxin to O2. (Original drawing courtesy C. Critchley).

In linear flow, water molecules are split in PSII, liberating O2 and providing a source of electrons. M is the manganese—containing cluster which oxidises water, Z is tyrosine-161 of the D1 protein which in turn represents the primary electron donor to P680+ (a special pair of Chl a molecules with an absorption peak at 680 nm). Pheo is the primary electron acceptor pheophytin a, a chlorophyll molecule lacking magnesium; QA is the first stable and permanently bound plastoquinone electron acceptor; QB is the second, temporarily bound, plastoquinone electron acceptor which actually leaves PSII in a reduced form (PQH2). Further along, FeS = Rieske iron—sulphur centre; Cyt f = cytochrome f; PC = plastocyanin; P700 = reaction centre chlorophyll a of PSI; A0, A1, FX, FB and FA are electron acceptors of PSI; Fd = ferredoxin; Cyt b559 = cytochrome b559; Cyt b563 = cytochrome b563. Also shown as tapered arrows is H+ accumulation in the lumen associated with water and plastoquinol oxidations.

The original version of this ‘Z’ scheme was further validated by unequivocal evidence from Australia that the two (inferred) photosystems were indeed separate physical entities. Using sophisticated biochemical chloroplast purification and subfractionation methods, coupled with detergent solubilisation of membranes, Boardman and Anderson (1964) achieved the first physical separation of photosystem II (PSII) and photosystem I (PSI), thus confirming the separate identities of those complexes. 

A source of electrons had long been recognised as basic to the operation of this ‘Z’ scheme, with H2O molecules an obvious source, but were photosynthetic membranes capable of photolysis? Early experiments by Robin Hill and colleagues at Cambridge had established this capability. They used isolated thylakoid membrane preparations and showed that O2 could be evolved in the absence of CO2 as long as external electron acceptors were present (Hill reaction). Intact leaves or whole chloroplasts have no need for an artificial acceptor because electron flow is directed to NADP+ and subsequent reduction of CO2 (first demonstrated with intact chloroplasts; see Arnon 1984). The O2-evolving function of photosynthesis was found to be associated with PSII in experiments with isolated thylakoids using external (artificial) electron donors and acceptors and specific electron transport inhibitors. As one outcome of those early Cambridge experiments, O2 evolution is now measured routinely in vitro (and in vivo on leaves) with O2 electrodes (Walker 1987). 

Chloroplast structure and function is by now sufficiently well defined to consider photosynthetic electron flow in detail. Figure 1.11 applies equally well to vascular plants or to algae with oxygenic photosynthesis, where in either case two photosystems work cooperatively and sequentially in absorbing photons and converting their quantum energy into a flow of electrons. Paradoxically, convention has it that photosynthetic electron flow initiates in PSII and proceeds to PSI. PSII was so named because PSI had already been described in single-celled (prokaryotic) organisms and, owing to the rules of nomenclature, was accorded priority. 

Both photosystems are large multi-subunit complexes, quite different structurally and functionally, and operating in series. In PSII, electrons are provided from a water-splitting apparatus via a manganese complex which undergoes oxidation from a valency state of +2 to +4. These oxidation states are made possible by P680+ (a special form of Chl a with an absorption peak at 680 nm). P680+ is a powerful oxidant generated by absorption of energy from a photon. P680 is referred to as a ‘special pair’ because it is a pair of Chl a molecules. Electrons from P680 pass to pheophytin (Pheo in Figure 1.11) and on to a bound quinone molecule, QA. From there a second transiently bound quinone, QB, receives two electrons in succession and requires protonation. The entire, fully reduced, quinone molecule leaves PSII and enters a plastoquinone pool (2PQ).

In PSI, absorption of quantum energy from a photon causes oxidation of P700, the PSI reaction centre equivalent of P680. In contrast to PSII, where electrons are drawn from a water-splitting apparatus, P700 accepts electrons from PC (reduced form PC in Figure 1.12). Electrons then pass through three iron–sulphur (FeS) centres and out of PSI to ferredoxin (Fd). The reaction centre of PSI contains several proteins, but most of the electron transfer cofactors are bound to large heterodimeric proteins which in turn bind the inner Chl a antenna. The LHCI complex consists of possibly eight polypeptides of between 24 and 27 kDa which carry Chl a and Chl b plus carotenoids.


Figure 1.12. Light harvesting, photosynthetic electron transport from H2O to NADP+ and generation of ATP are achieved via four types of complexes which show a lateral heterogeneity within thylakoid membranes. A small part of a continuous network of interconnected thylakoids is shown here diagrammatically where PSI complexes and ATP synthase are restricted to non—appressed regions. Most PSII complexes and the light-harvesting assemblages associated with PSII (LHCII) are held within appressed regions of this network. Cytochrome b/f complexes (Cyt b/f) are more generally located. (Based on J.M. Anderson and B. Andersson, Trends Biochem Sci 13: 351-355, 1988)

A chemiosmotic coupling mechanism is responsible for ATP synthesis. Protons are ‘pumped’ across the thylakoid membrane from outside (stroma) to inside (lumen) by a complex arrangement of electron carriers embedded within the membrane. A prodigious concentration of protons builds up within the lumen, partly from photolysis of water molecules (water-splitting apparatus on PSII) and partly from oxidation of plastoquinone (PQ) on the inner face of the membrane. Hence, energy originally carried by incident photons is transduced into energy stored within an electrochemical gradient acrosss the thylakoid membrane. The protonmotive force from inside (lumen) to outside (stroma) is used to generate ATP within the stroma via an ATP synthase complex (CF0 and CF1) that straddles the thylakoid membrane. OEC = oxygen-evolving complex; Pheo = pheophytin a.These two photosystems are juxtaposed across thylakoid membranes in such a way that linear electron transport is harnessed for charge separation, leading to a massive accumulation of H+ ions within the lumen of illuminated thylakoids, which is then employed in ATP generation.

Combining concepts of photolysis and photosynthetic electron flow outlined earlier (Figure 1.11) and putting that conceptual framework into a thylakoid membrane system (Figure 1.12), a picture emerges where electrons generated from splitting H2O molecules on the inner surface of PSII are transferred from plastoquinol (PQH2) to the Rieske iron– sulphur centre (Rieske FeS) of the cytochrome b6/f complex (Cyt b6/f) and further to cytochrome f (Cyt f). The pivotal importance of Cyt f in facilitating electron transport from PSII to PSI was demonstrated by Duysens and colleagues (see Levine 1969), who showed that preferential energisation of PSII (light at <670 nm) caused reduction, whereas preferential energisation of PSI (light at >695 nm) caused oxidation. This elegant ‘push–pull’ experiment confirmed the cooperative and sequential nature of PSII and PSI, as well as indicating overall direction of photosynthetic electron flow. 

Proteins which bind the Rieske FeS centre and Cyt f together with cytochrome b563 (Cyt b6) form a large electron transfer complex. This complex (Figure 1.12) spans the membrane and is located between the two photosystems. Electrons are transferred to PC (forming PC), a copper-containing soluble protein extrinsic to the thylakoid membrane and located in the lumen. On the other side of the membrane, attached to the stromal side, is ferredoxin (Fd) which accepts electrons from PSI and passes them on to ferredoxin–NADP reductase, an enzyme, also extrinsic to thylakoids, and attached on the stromal side of the thylakoid membrane. This enzyme accomplishes the final electron transfer in an overall linear chain and reduced NADP is then protonated.

While linear electron transport from water to NADP+ is the main and most important path, electrons can also be transferred to O2 in a so-called pseudocyclic or Mehler reaction (Figure 1.11). This pathway probably operates in vivo as a sink for electrons when synthetic events call for more ATP than NADPH. Electrons can also be cycled around both PSII and PSI. Electrons cycling around PSI will produce ATP but with no accompanying NADPH. Cyclic electron flow around PSII may have a completely different role and may be related to the downregulation of this photosystem during photoinhibition (Chapter 12).

According to this multistage scheme, electrons are transferred from donor (reductant) to acceptor (oxidant). The direction of that transfer depends upon a difference in oxidation–reduction potential between a given donor and a given acceptor (as indicated on the ordinate in Figure 1.11). A more positive potential implies stronger oxidative power (i.e. capacity to accept electrons); a more negative potential implies stronger reducing power (i.e. capacity to donate electrons). P680* thus has a strong capacity to donate electrons (a strong reductant); P700* has an even stronger capacity to donate electrons (an even stronger reductant). 

Molecules which accept electrons are immediately protonated. In aqueous systems, such as chloroplasts in vivo, hydrogen ions (H+) are ubiquitous, and these ions combine with electron acceptors to generate hydrogen atoms (i.e. H+ ion + electron  H atom). In Figure 1.11, some events involve electron transfer, while others include transfer of hydrogen atoms. As a simplifying convention, all such events are referred to as electron transfers. Ironically, the end result of all these reactions is a net transfer of hydrogen atoms!

1.2.4 - ATP synthesis

During photosynthetic electron transfer from water to NADP+, energy captured in two photoacts is stored as an electrochemical potential gradient of protons. First, such reduction of QB requires protonation with protons drawn from the stromal side of the membrane. Reoxidation (and deprotonation) occurs towards the thylakoid lumen. In addition, protons are lost from the stromal side via protonation of reduced NADP and they are also generated in the lumen during photolysis. A massive ΔpH, of approximately 3–4 pH units, equivalent to an H+ ion concentration difference of three to four orders of magnitude, develops across the thylakoid membrane. This immense gradient drives ATP synthesis (catalysed by ATP synthase) within a large energy-transducing complex embedded in the thylakoid membrane (Figure 1.12). 

ATP synthesis in chloroplasts (photophosphorylation) proceeds according to a mechanism that is basically similar to that in mitochondria. Chemiosmotic coupling (Mitchell 1961) which links the movement of protons down an electro-chemical potential gradient to ATP synthesis via an ATP synthase applies in both organelles. However, the orientation of ATP synthase is opposite. In chloroplasts protons accumulate in thylakoid lumen and pass outwards through the ATP synthase into the stroma. In mitochondria, protons accumulate within the intermembrane space and move inwards, generating ATP and oxidising NADH within the matrix of these organelles (Figure 2.24).

In chloroplasts, ATP synthase is called the CF0CF1 complex. The CF0 unit is a hydrophobic transmembrane multiprotein complex which contains a water-filled proton conducting channel. The CF1 unit is a hydrophilic peripheral membrane protein complex that protrudes into the stroma. It contains a reversible ATPase and a gate which controls proton movement between CF0 and CF1. Entire CF0CF1 complexes are restricted to non-appressed portions of thylakoid membranes due to their bulky CF1 unit. 

Direct evidence for ATP synthesis due to a transthylakoid pH gradient can be adduced as follows. When chloroplasts are stored in darkness in a pH 4.0 succinic acid buffer (i.e. a proton-rich medium), thylakoid lumen equilibrate to this pH. If the chloroplasts, still in the dark, are rapidly transferred to a pH 8.0 buffer containing ADP and Pi, ATP synthesis then occurs. This outcome confirms a central role for the proton concentration difference between thylakoid lumen and stroma for ATP synthesis in vitro; but does such a process operate on that scale in vivo

Mordhay Avron, based in Israel, answered this question in part during the early 1970s via a most elegant approach (Rottenberg et al. 1972). Working with thylakoid preparations, Avron and colleagues established that neutral amines were free to exchange between bathing medium and thylakoid lumen, but once protonated in illuminated preparations they became trapped inside. By titrating the loss of such amines from the external medium when preparations where shifted from dark to light, they were able to infer the amount retained inside. Knowing that the accumulation of amine depended upon H+ ion concentration in that lumen space, the difference in H+ ion concentration and hence ΔpH across the membrane were established. 

At saturating light, chloroplasts generate a proton gradient of approximately 3.5 pH units across their thylakoid membranes. Protons for this gradient are derived from the oxidation of water molecules occurring towards the inner surface of PSII and from transport of four electrons through the Cyt b/f complex, combined with cotranslocation of eight protons from the stroma into the thylakoid space for each pair of water molecules oxidised. Electrical neutrality is maintained by the passage of Mg2+ and Cl across the membrane, and as a consequence there is only a very small electrical gradient across the thylakoid membrane. The electrochemical potential gradient that yields energy is thus due almost entirely to the concentration of intrathylakoid H+ ions. 

For every three protons translocated via ATP synthase, one ATP is synthesised. Linear electron transport therefore generates about four molecules of ATP per O2 evolved. Thus eight photons are absorbed for every four ATP molecules generated or for each O2 generated. Cyclic electron transport is slightly more efficient at producing ATP and generates about four ATP per six photons absorbed. However, linear electron transport also generates NADPH, which is equivalent, in energy terms, to six ATP per O2 released.

As implied in Figure 1.12, the four thylakoid complexes, PSII, PSI, Cyt b/f and ATP synthase, are not evenly distributed in plant thylakoid membranes but show a lateral heterogeneity. This distribution is responsible for the highly characteristic structural organisation of the continuous thylakoid membrane into two regions, one consisting of closely appressed membranes or granal stacks, the other of non-appressed stroma lamellae where outside surfaces of thylakoid membranes are in direct contact with the stroma. This structural organisation is shown on a modest scale in Figure 1.7, but extreme examples are evident in chloroplasts of shade-adapted species grown in low light (Chapter 12). Under such conditions, membrane regions with clusters of PSII complexes and Cyt b/f complexes become appressed into classical granal stacks. Cyt b/f complexes are present inside these granal stacks as well as in stroma lamellae, but PSI and ATP synthase are absent from granal stacks. Linear electron transport occurs in granal stacks from PSII in appressed domains to PSI in granal margins. Nevertheless, shade plants have only a low rate of linear electron transport because they have fewer Cyt b/f and to a lesser extent fewer PSII complexes compared to PSI, a consequence of investing more chlorophyll in each PSII to enhance light harvesting (see Anderson (1986) and Chapter 12 for more detail).

1.2.5 - Chlorophyll fluorescence


Figure 1.13 Catching the Light is a demonstration of photosynthesis in action. Photosynthesis begins when light is absorbed by chlorophyll. The flask contains chlorophyll extracted from spinach leaves. When a beam of light passes through the extract, the chlorophyll absorbs this energy. But because the chlorophyll in the flask has been isolated from the plant, energy cannot be converted and stored as sugar. Instead it is released as heat and red fluorescence. Note the green ring below the flask which is transmitted light, the colour we normally perceive for chlorophyll. The colour of a leaf is green because it reflects and transmits green light but absorbs the blue and red components of white light. (Image courtesy R. Hangarter)

A dilute solution of leaf chlorophyll in organic solvent appears green when viewed in white light. Wavelengths corresponding to bands of blue and red have been strongly absorbed (Figure 1.8), whereas mid-range wavelengths corresponding to green light are only weakly absorbed, hence the predominance of those wavelengths in transmitted and reflected light. However, when viewed at right angles to the light source, the solution will appear deep red due to energy re-emitted as fluorescence (Figure 1.13). The red colour is evident regardless of the colour of the source light.

Chlorophyll within the two photosystems can absorb energy from incident photons. This absorbed energy can be dissipated by driving the processes of photosynthesis, as heat, or re-emitted as fluorescence radiation. These are all complementary processes so that fluorescence provides an important tool in the study of photosynthesis. The normal processes of photochemistry and electron transport within intact leaves typically reduce the amount of fluorescence, a process referred to as quenching. In the demonstration shown in Figure 1.13 the chlorophyll has been isolated from the plant these processes are disrupted, minimizing the quenching effects.

Fluorescence spectra are invariate, and the same spectrum will be obtained (e.g. Figure 1.8 inset) regardless of which wavelengths are used for excitation. This characteristic emission is especially valuable in identifying source pigments responsible for given emission spectra, and for studying changes in their photochemical status during energy transduction. 

Fluorescence emission spectra (Figure 1.8 inset) are always displaced towards longer wavelengths compared with corresponding absorption spectra (Stoke’s shift). As quantum physics explains, photons intercepted by the chromophore of a chlorophyll molecule cause an instantaneous rearrangement of certain electrons, lifting that pigment molecule from a ground state to an excited state which has a lifetime of c. 10–9 s. Some of this excitation energy is subsequently converted to vibrational energy which is acquired much more ‘slowly’ by much heavier nuclei. A non-equilibrium state is induced, and molecules so affected begin to vibrate rather like a spring with characteristic periodicity, leading in turn to energy dissipation as heat plus remission of less energetic photons of longer wavelength.

Apart from their role in photon capture and transfer of excitation energy, photosystems function as energy converters because they are able to seize photon energy rather than lose as much as 30% of it through fluorescence as do chlorophylls in solution. Moreover, they can use the trapped energy to lift an electron to a higher energy level from where it can commence a ‘downhill’ flow via a series of electron carriers as summarised in Figure 1.11.

Protein structure confers very strict order on bound chlorophylls. X-ray crystallographic resolution of the bacterial reaction centre has given us a picture of the beautiful asymmetry of pigment and cofactor arrangements in these reaction centres, and electron diffraction has shown us how chlorophylls are arranged with proteins that form the main light-harvesting complexes of PSII. This structural constraint confers precise distance and orientation relationships between the various chlorophylls, as well as between chlorophylls and carotenoids, and between chlorophylls and cofactors enabling the photosystems to become such effective photochemical devices. It also means that only 2–5% of all the energy that is absorbed by a photosystem is lost as fluorescence. 

Fig 1.14.png

Figure 1.14 Fluorescence emission spectra from a leaf measured at room temperature or in liquid nitrogen. Spectra have been normalised to the peak at 748 nm.

If leaf tissue is held at liquid nitrogen temperature (77 K), photosynthetic electron flow ceases and chlorophyll fluorescence increases, including some emission from PSI (Figure 1.14). Induction kinetics of chlorophyll fluorescence at 77 K have been used to probe primary events in energy transduction, and especially the functional state of photosystems. Present discussion is restricted to room temperature fluorescence where even the small amount of fluorescence from PSII is diagnostic of changes in functional state. This is because chlorophyll fluorescence is not emitted simply as a burst of red light following excitation, but in an ordered fashion that varies widely in flux during continuous illumination. These transient events (Figure 1.15) are referred to collectively as fluorescence induction kinetics, fluorescence transients, or simply as a Kautsky curve in honour of its discoverer Hans Kautsky (Kautsky and Franck 1943). 

At room temperature and under steady-state conditions, in vivo Chl a fluorescence from leaves show a characteristic emission spectrum with two distinct peaks around 680–690 nm and 750 nm, both of which mainly originate from photosystem II (Figure 1.14). Because  other chlorophyll molecules can reabsorb fluorescence emitted at 680–690 nm within a leaf, the spatial origin of fluorescence can differ between the 680 and 750nm fluorescence that is detected. The fluorescence waveband measured by room temperature fluorometers differs between instruments. 

(a) Fluorescence induction kinetics


Figure 1.15 A representative chart recorder trace of induction kinetics for Chl a fluorescence at room temperature from a mature bean leaf (Phaseolus vulgaris). The leaf was held in darkness for 17 min prior to excitation (zig-zag arrow) at a photon irradiance of 85 µmol quanta m-2 s-1. The overall Kautsky curve is given in (b), and an expanded version of the first 400 ms is shown in (a). See text for explanation of symbols and interpretation of variation in strength for these ‘rich but ambiguous signals’! (Based on R. Norrish et al., Photosyn Res 4: 213-227, 1983)

Strength of emission under steady-state conditions varies according to the fate of photon energy captured by LHCII, and the degree to which energy derived from photosynthetic electron flow is gainfully employed. However, strength of emission fluctuates widely during induction (Figure 1.15) and these rather perplexing dynamics are an outcome of some initial seesawing between photon capture and subsequent electron flow. Taking Figure 1.11 for reference, complexities of a fluorescence transient (Figure 1.15) can be explained as follows. At the instant of excitation (zig-zag arrow), signal strength jumps to a point called \(F_0\) which represents energy derived largely from chlorophyll molecules in the distal antennae of the LHCII complex which fail to transfer their excitation energy to another chlorophyll molecule, but lose it immediately as fluorescence. \(F_0\) thus varies according to the effectiveness of coupling between antennae chlorophyll and reaction centre chlorophyll, and will increase due to high-temperature stress or photodamage. Manganese-deficient leaves show a dramatic increase in \(F_0\) due to loss of functional continuity between photon-harvesting and energy-processing centres of PSII (discussed further in Chapter 16). 

Returning to Figure 1.15, the slower rise subsequent to \(F_0\) is called \(I\), and is followed by a further rise to \(F_m\). These stages reflect a surge of electrons which fill successive pools of various electron acceptors of PSII. Significantly, Fm is best expressed in leaves that have been held in darkness for at least 10–15 min. During this dark pretreatment, electrons are drawn from QA, leaving this pool in an oxidised state and ready to accept electrons from PSII. An alternative strategy is to irradiate leaves with far-red light to energise PSI preferentially, and so draw electrons from PSII via the Rieske FeS centre. The sharp peak (\(F_m\)) is due to a temporary restriction on electron flow downstream from PSII. This constraint results in maximum fluorescence out of PSII at about 500 ms after excitation in Figure 1.15(a). That peak will occur earlier where leaves contain more PSII relative to electron carriers, or in DCMU-treated leaves. 

Photochemistry and electron transport activity always quench fluorescence to a major extent unless electron flow out of PSII is blocked. Such blockage can be achieved with the herbicide 3-(3,4-dichlorophenyl)-1,1-dimethyl urea (DCMU) which binds specifically to the D1 protein of PSII and blocks electron flow to QB. DCMU is a very effective herbicide because it inhibits photosynthesis completely. As a consequence, signal rise to \(F_m\) is virtually instantaneous, and fluorescence emission stays high. 

Variation in strength of a fluorescence signal from \(F_0\) to \(F_m\) is also called variable fluorescence (\(F_v\)) because scale and kinetics of this rise are significantly influenced by all manner of environmental conditions. \(F_0\) plus \(F_v\) constitute the maximal fluorescence (\(F_m\)) a leaf can express within a given measuring system. The \(F_v/F_m\) ratio, measured after dark treatment, therefore reflects the proportion of efficiently working PSII units among the total PSII population. Hence it is a measure of the photochemical efficiency of a leaf, and correlates well with other measures of photosynthetic effectiveness (discussed further in Chapter 12).

(b) Fluorescence relaxation kinetics

Both the patterns of initial induction of fluorescence, and its subsequent decay once the light has ceased, are important indicators of the underlying structure and function of photosynthetic systems. The latter is referred to as the relaxation kinetics of a fluorescence event. In a typical experiment the chlorophyll is exposed to repeated pulses of light and the relaxation kinetics measured (Figure 1.16).


Figure 1.16 Induction and relaxation kinetics of in vivo Chl a fluorescence from a well-nourished radish leaf (Raphanus sativus) supplied with a photon irradiance of actinic light at 500 µmol quanta m-2 s-1 and subjected to a saturating pulse of 9000 µmol quanta m-2 s-1 for 0.8 s every 10 s. Output signal was normalised to 1.0 around the value for \(F_m\) following 30 min dark pretreatment. Modulated light photon irradiance was <1 µmol quanta m-2 s-1. See text for definition of symbols and interpretation of kinetics. (Original data from J. Evans generated on a PAM fluorometer - Heinz Walz GmbH, Germany)

Excellent fluorometers for use in laboratory and field such as the Plant Efficiency Analyser (Hansatech, King’s Lynn, UK) make accurate measurements of all the indices of the Kautsky curve and yield rapid information about photochemical capacity and response to environmental stress. Conventional fluorometers (e.g. Figure 1.15) use a given source of weak light (commonly a red light-emitting diode producing only 50–100 µmol quanta m–2 s–1) for both chlorophyll excitation and as a source of light for photosynthetic reactions.

Even more sophisticated is the Pulse Amplitude Modulated (PAM) fluorometer (Walz, Effeltrich, Germany) which employs a number of fluorescence- and/or photosynthesis-activating light beams and probes fluorescence status and quenching properties. These fluorimeters measure fluorescence excited by a weak source of light that is modulated: that is a beam that applies short, square pulses of saturating light for chlorophyll excitation on top of a constant beam of light that sustains photosynthesis (actinic light). A combination of optical filters plus sophisticated electronics is used to tune the detector to detect only fluorescence excited by the modulated light beam.

In this way, most of the continuous background fluorescence and reflected long-wavelength light is disregarded. Most significantly, relative fluorescence can be measured in full sunlight in the field. The functional condition of PSII in actively photosynthesising leaf tissue is thus amenable to analysis. This instrument also reveals the relative contributions to total fluorescence quenching by photochemical and non-photochemical processes and will help assess any sustained loss of quantum efficiency in PSII. Photosynthetic electron transport rates can be calculated concurrently. These techniques have revolutionised the application of chlorophyll fluorescence to the study of photosynthesis.

Photochemical quenching (\(q_p\)) varies according to the oxidation state of electron acceptors on the donor side of PSII. When QA is oxidised (e.g. subsequent to dark pretreatment), quenching is maximised. Equally, \(q_p\) can be totally eliminated by a saturating pulse of excitation light that reduces QA, so that fluorescence yield will be maximised, as in a PAM fluorometer. Concurrently, a strong beam of actinic light drives photosynthesis (maintaining linear electron flow) and sustaining a pH gradient across thylakoid membranes for ATP synthesis. Those events are a prelude to energy utilisation and contribute to non-photochemical quenching (\(q_n\)). This \(q_n\) component can be inferred from a combination of induction plus relaxation kinetics.

In Figure 1.16, a previously darkened radish leaf (QA oxidised and ready to receive an electron from P680; 'traps open') initially receives weak modulated light (<1 µmol quanta m–2 s–1) that is insufficient to close traps but sufficient to establish a base line for constant yield fluorescence (\(F_0\)). This value will be used in subsequent calculations of fluorescence indices. The leaf is then pulsed with a brief (0.8 s) saturating flash (9000 µmol quanta m–2 s–1) to measure \(F_m\). Pulses follow at 10 s intervals to measure \(F_m^\prime\). Actinic light (500 µmol quanta m–2 s–1) starts with the second pulse and pH starts to build up in response to photosynthetic electron flow. Photosynthetic energy transduction comes to equilibrium with these conditions after a minute or so, and fluorescence indices \(q_n\) and \(q_p\) can then be calculated as follows:

\[ q_n=\frac{F_m - F_m^\prime}{F_m - F_0} \text{, and } q_p=\frac{F_m^\prime-F}{F_m^\prime - F_0} \tag{1.1} \]
Under these steady-state conditions, saturating pulses of excitation energy are being used to probe the functional state of PSII, and by eliminating \(q_p\) the quantum efficiency of light-energy conversion by PSII (\(\Phi_{PSII}\)) can be inferred:

\[ \Phi_{PSII} = \frac{F_m^\prime - F}{F_m^\prime} \tag{1.2} \]
If overall quantum efficiency for O2 evolution is taken as 10 (discussed earlier), then the rate of O2 evolution by this radish leaf will be: 

\[ \Phi_{PSII} \times \text{photon irradiance}/10 \;(\mu\text{mol O}_2 m^{-2}s^{-1}) \tag{1.3} \]

In summary, chlorophyll fluorescence at ambient temperature comes mainly from PSII. This photosystem helps to control overall quantum efficiency of electron flow and its functionality changes according to environmental and internal controls. In response to establishment of a ΔpH across thylakoid membranes, and particularly when irradiance exceeds saturation levels, some PSII units become down-regulated, that is, they change from very efficient photochemical energy converters into very effective energy wasters or dissipators (Chapter 12). Large amounts of the carotenoid pigment zeaxanthin in LHCII ensure harmless dissipation of this energy as heat (other mechanisms may also contribute). PSII also responds to feedback from carbon metabolism and other energy-consuming reactions in chloroplasts, and while variation in pool size of phosphorylated intermediates has been implicated, these mechanisms are not yet understood.

Case Study 1.2 - Five chlorophylls and photosynthesis

Min Chen

School of Biological Sciences, University of Sydney, Australia


Figure 1. Absorption spectra of photosynthetic organisms containing different chlorophylls and the quantum yield of photosynthesis using chlorophylls a and b (grey line). Green line, in vivo absorption spectrum of Synechocystis PCC 6803 in BG11 medium; Black line, isolated Prochloron cell suspension in seawater; Red line, in vivo absorption spectrum of Acaryochloris marine MBIC11017 in seawater medium; and Blue line, in vivo absorption spectrum of Halomicronema hongdechloris in seawater medium.

Solar radiation is a black body at a temperature of ~5800oK, covering all spectral regions. However, all known eukaryotic photosynthetic organisms (including plants and algae) are only able to use the same region of the solar spectrum that our eyes are sensitive to, covering the wavelength of 400 – 700 nm region, which is approximately 43% of the total solar radiation. This region is called photosynthetic active radiation (PAR) with estimated photon flux of 1.05 x 1021 photons m-2 s-1. Longer wavelengths up to 1000 nm can drive anoxygenic photosynthesis but not oxygenic (oxygen evolving) photosynthesis. The reason for the high threshold energy for oxygenic photosynthesis is the higher energy requirement for catalysing water oxidation and oxygen evolution in photosynthesis. The PAR input limit depends on the absorption of the photopigments. Chlorophylls a and b, the main chlorophylls in eukaryotic photosynthetic organisms, show their maximal absorption bands in the blue region of 430 - 455 nm and the red region of 645 -670 nm, thus leaving a “green window” and photons outside of “visible region” (Figure 1). The photons collected by chlorophylls a and b provide a strong enough redox potential for the oxidisation of water, while at the same time they also provide an negative enough excited state redox potentials for the reduction of the primary electron acceptor. 

There are five different chlorophylls that have been identified, chlorophylls a, b, c, d and f. Here, we focus on chlorophylls containing five rings (macrocycle) and an esterified 17-propionic acid side chain, the chlorin type chlorophylls, including chlorophylls a, b, d and f. The chemical difference among the different chlorophylls is either formyl substitution at the side chain of the macrocycle (chlorophyll b, d, and f) or the degree of unsaturation of the macrocycle (8-vinyl chlorophyll a and 8-vinly chlorophyll b).  Those chemical structural differences are also the spectral determinants and responsible for the different absorption spectral features (Figure 2).


Figure 2. Chemical structure of chlorophylls and their absorption spectra in 100% methanol. (A) Chemical structure of chlorophyll a and the structural differences of other chlorophylls from chlorophyll a. The carbon atoms are numbered using IUPAC system. (B) Absorption spectra of red-shifted chlorophylls, chlorophylls d and f, compared with chlorophyll a. (C) Absorption spectra of chlorophyll b and 8-vinyl chlorophyll a compared with chlorophyll a. (modified from reference 3)

Chlorophyll b is distinguished from chlorophyll a by a formyl instead of a methyl group on ring B at C7 position, which results in a blue-shift of the longest red absorbance band (Qy) from 665 nm to 652 nm. Chlorophyll d and chlorophyll f are distinguished from chlorophyll a by replacement of a peripheral substituent on ring A by a formyl group at C3 position and C2 position, respectively (Figure 2). The consequences of those formyl group substitutions at ring A result into a red-shifted Qy absorption wavelength, from 665 nm to 696 nm (chlorophyll d) and even further to 706 nm for chlorophyll f. Both chlorophylls d and f are named as red-shifted chlorophylls. Those red-shifted chlorophylls allow the organisms to use the light beyond 700 nm efficiently compared with the organisms containing chlorophylls a and b only.

Plants using chlorophyll a and b demonstrate the decreased quantum yield of photosynthesis using wavelength >700 nm, which is known as the “red drop” (Figure 1). The reason for the red-drop at ~700 nm is that the chlorophylls that absorb longer wavelength light beyond 700 nm will not do photosynthesis as efficiently as the chlorophylls that absorb shorter wavelength light. The 700 nm photons were considered as the red-edge of oxygenic photosynthesis. However, the newly discovered chlorophyll d-containing Acaryochloris marina and chlorophyll f-containing Halomicronema hongdechloris have forced a re-evaluation of what is the minimum threshold energy for oxygenic photosynthesis. Both red-shifted chlorophyll-containing cyanobacteria are found in the environment where visible lights are depleted by above layers of oxygenic photosynthetic organisms. The red-shifted chlorophylls allow them to absorb the longer wavelength light beyond 700 nm and do oxygenic photosynthesis as efficiently as above layers of chlorophyll a-containing organisms.  Accordingly, the minimum threshold energy for oxygenic photosynthesis has been at extended to at least 750 nm in those red-shifted chlorophyll-containing organisms. The PAR increment in the region of the solar spectrum of 700 – 750 nm increases the number of available light energy by 19%.  The potential additional photon flux in the infrared region (700-750 nm) could improve the light-harvesting efficiency by extending the PAR coverage to 400-750 nm if those red-shifted chlorophylls could be introduced into plants and algae.

In addition for the potential enhancement for efficient light collection and transfer to the reaction centres under weak irradiation, the second functional demands for the light-harvesting process is their protecting function at exposure to strong light, which will be covered in a following case study.

Further Reading

Chen M, Blankenship RB (2011) Expanding the solar spectrum used by photosynthesis. Trends Plant Sci 16: 427-431

Chen M, Schliep M, Willows R, Cai Z-L, Neilan BA, Scheer H (2010) A red-shifted chlorophyll. Science 329: 1318-1319

Chen M, Scheer H (2013) Extending the limit of natural photosynthesis and implications of technical light harvesting, J Porphyrins Phthalocyanines 17: 1-15

1.3 - Concluding remarks


Chloroplasts are sites of solar energy absorption and subsequent transduction into chemically usable forms. Splitting water molecules and developing a proton motive force of sufficient magnitude to drive ATP synthesis are energy-intensive processes. Consequently, photosynthetic organisms evolved with dual photosystems that work cooperatively and sequentially to extract sufficient quantum energy from parcels of absorbed photons to generate a sufficiently strong electrochemical potential gradient to synthesise the relatively stable, high-energy compounds ATP and NADPH. Such metabolic energy sustains cycles of photosynthetic carbon reduction (PCR) where CO2 is initially assimilated by one of three photosynthetic pathways, namely C3, C4 or CAM, but eventually fixed via a PCR cycle within the stromal compartment of chloroplasts. These photosynthetic pathways are described in the following chapter.  Section 2.1 describes C3 photosynthesis, Section 2.2 presents C4 photosynthesis and other photosynthetic modes, and Section 2.3 covers photorespiration.

Thermodynamically, the net outcome of photosynthetic energy transduction must be viewed as long-term storage of energy in the form of a product pair, namely free oxygen and reduced carbon (organic matter), rather than as separate molecules. Plants themselves or indeed any heterotrophic organisms subsequently retrieve such energy via metabolic ‘combustion’ of the organic matter where enzyme-catalysed reactions bring this pair of products together again in the process known as mitochondrial respiration. This is described in Section 2.4.

1.4 Further reading

Evans JR, von Caemmerer S (1996) Carbon dioxide diffusion inside leaves. Plant Physiol 110: 339-346

Evans JR, Kaldenhoff R, Genty B, Terashima I (2009) Resistances along the CO2 diffusion pathway inside leaves. J Exp Bot 60: 2235-2248

Kramer DM, Evans JR (2011) The importance of energy balance in improving photosynthetic productivity. Plant Physiol 155: 70-78

Sharkey TD (1985) Photosynthesis in intact leaves of C3 plants: physics, physiology and rate limitations. Bot Rev 51: 53-105

Syvertsen JP, Lloyd J, McConchie C, Kriedemann PE, Farquhar GD (1995) On the relationship between leaf anatomy and CO2 diffusion through the mesophyll of hypostomatous leaves. Plant Cell Environ 18: 149-157

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Roger Hangarter and Dennis DeHart http://plantsinmotion.bio.indiana.edu/usbg/photosyn.htm

Chapter 2 - Carbon dioxide assimilation and respiration


Tobacco plants (a) transformed with an antisense construct against Rubisco (anti-Rubisco) grow more slowly than wild types due to a 60% reduction in photosynthetic rate. Immunodetection of the large subunit polypeptide of Rubisco with an anti-Rubisco antiserum (b) shows that the anti—Rubisco transgenic plants contain less than 50% of the Rubisco detected in wild-type tobacco plants. Scale bar in (a) = 10 cm (Photograph courtesy Susanne von Caemmerer; original immunoblot courtesy Martha Ludwig)

Oula Ghannoum1, Susanne von Caemmerer2, Nicolas Taylor3 and A. Harvey Millar3
1Hawkesbury Institute of the Environment, University of Western Sydney
2Research School of Biology, Australian National University
3ARC Centre of Excellence inPlant Energy Biology, University of Western Australia

Rubisco (ribulose-1,5-bisphosphate carboxylase/oxygenase) is the most abundant single protein on earth and is pivotal for CO2 assimilation by all plants. In higher plants, the holoenzyme consists of eight large subunits, each with a molecular mass of 50-55 kD and eight small subunits of molecular mass 12-18 kD. Large subunits are encoded by a single gene in the chloroplast genome while a family of nuclear genes encode the small subunits. Any loss of catalytic effectiveness or reduction in amount translates to slower photosynthesis and reduced growth.

Life on earth is sustained by photosynthetic use of sunlight energy to convert atmospheric CO2 into carbohydrates. Billions of years ago, photosynthetic cyanobacterium-like prokaryotes were engulfed by early heterotrophic eukaryotes to produce aquatic photosynthetic organisms harbouring chloroplasts with double membranes. These gave rise to vascular plants which in turn adapted to changing terrestrial environments via distinctive modes of photosynthesis.

Most terrestrial plants fix atmospheric CO2 into carbohydrates via the C3 photosynthetic pathway and its initial three-carbon fixation product (Section 2.1). Millions of years of evolution under conditions of water limitation, temperature variations and glacial CO2 concentrations have produced higher plants with significant biochemical variants for fixation of atmospheric CO2 into carbohydrate, namely C4 (initial four-carbon fixation product), CAM (crassulacean acid metabolism) and SAM (submerged aquatic macrophytes) (Section 2.2).

Photosynthesis in C3 plants is inhibited by oxygen, initiating a series of metabolic reactions termed photorespiration (Section 2.3). Mitochondrial respiration converts the carbon gained for generation of energy to sustain growth and nutrient upake, as well as providing carbon skeletons for a multitude of synthetic events (Section 2.4).

2.1 - C<sub>3</sub> photosynthesis

Oula Ghannoum, University of Western Sydney, Australia

Despite much diversity in life form and biochemical process, all of the photosynthetic pathways focus upon a single enzyme which is by far the most abundant protein on earth, namely ribulose-1,5-bisphosphate carboxylase/oxygenase, or Rubisco (Figure 2.1a). Localised in the stroma of chloroplasts, this enzyme enables the primary catalytic step in photosynthetic carbon reduction (or PCR cycle) in all green plants and algae. Although Rubisco has been highly conserved throughout evolutionary history, this enzyme is surprisingly inefficient with a slow catalytic turnover (Vcmax), a poor specificity for CO2 as opposed to O2 (Sc/o), and a propensity for catalytic misfiring resulting in the production of catalytic inhibitors. This combination severely restricts photosynthetic performance of C3 plants under current ambient conditions of 20% O2 and 0.039% CO2 (390 μL L-1). Furthermore, Rubisco has a requirement for its own activating enzyme, Rubisco activase, which removes inhibitors from the catalytic sites to allow further catalysis. Accordingly, and in response to CO2 limitation, C4, C3-C4 intermediate, CAM and SAM variants have evolved with metabolic concentrating devices which enhance Rubisco performance (Section 2.2).

2.1.1 - Photosynthetic carbon reduction


Figure 2.1 Photosynthetic carbon reduction (PCR cycle, also termed the Calvin-Benson cycle) utilises ATP and NADPH produced by thylakoid electron transport to drive CO2 fixation by Rubisco (a). CO2 is incorporated into a 5-carbon sugar phosphate to produce two 3-carbon sugar phosphates which can either be exported from the chloroplast for sucrose synthesis, be recycled to make more 5-carbon acceptors, or be used to make starch. The appearance of radioactive carbon in 3-carbon sugar phosphates and then in starch and sucrose following photosynthesis in 14CO2 was evidence for the pathway of photosynthesis. (b) (Original drawing courtesy Robert Furbank).

The biochemical pathway of CO2 fixation was discovered by feeding radioactively labelled CO2 in the light to algae and then extracting the cells and examining which compounds accumulated radioactivity. Figure 2.1(b) shows a typical labelling ‘pattern’ for a C3 plant. Here, a short burst of labelled CO2 was given to the plants, then the label was ‘chased’ through the photosynthetic pathway by flushing with unlabelled air. Atmospheric CO2 is initially incorporated into a five-carbon sugar phosphate (ribulose-1,5-bisphosphate or RuBP) to produce two molecules of the phosphorylated three-carbon compound 3-phosphoglycerate, often referred to as the acidic form 3-phosphoglyceric acid (3-PGA). Hence, plants which use Rubisco as their primary enzyme of CO2 fixation from the air are called C3 plants. Consequently, in C3 plants, 3-PGA is the first labelled sugar phosphate detected after a pulse of 14CO2 has been supplied (Figure 2.1b). In the PCR cycle, 3-PGA is phosphorylated by the ATP produced from thylakoid electron transport (see Chapter 1) and then reduced by NADPH to produce triose phosphate. Triose phosphates are the carbon backbones, produced by the PCR cycle, for the synthesis of critical carbohydrate for the maintenance of plant growth and the productive yield of stored carbohydrate in seed.

Newly synthesised triose phosphate faces three options. It can be (1) exported to the cytosol for sucrose synthesis and subsequent translocation to the rest of the plant, (2) recycled within the chloroplast to produce more RuBP or (3) diverted to produce starch (Figure 2.1a). This is shown by the time-course of the appearance of radioactivity in starch and sucrose after it has passed through 3-PGA (Figure 2.1b). The energy requirements of the PCR cycle are three ATP and two NADPH per CO2 fixed, in the absence of any other energy-consuming processes.

Sucrose and starch synthesis

Most of the triose phosphate synthesised in chloroplasts is converted to either sucrose or starch. Starch accumulates in chloroplasts, but sucrose is synthesised in the surrounding cytosol, starting with the export of dihydroxyacetone phosphate and glyceraldehyde phosphate from the chloroplast. A condensation reaction, catalysed by aldolase, generates fructose-1,6-bisphosphate, and this is converted to fructose-6-phosphate after an hydrolysis reaction catalysed by fructose-1,6-phosphatase. Sucrose-6-phosphate synthase then generates sucrose-6-phosphate from the reaction of fructose-6-phosphate and UDP-glucose. The phosphate group is removed by the action of sucrose-6-phosphatase. This Pi is transported back into the chloroplast where it is available for ATP synthesis. For each molecule of triose phosphate exported from a chloroplast, one Pi is translocated inwards.

Sucrose synthesised within the cytosol of photosynthesising cells is then available for general distribution and is commonly translocated to other carbon-demanding centres via the phloem (see Chapter 5).

By contrast, starch synthesis occurs within chloroplasts. The first step is a condensation of glucose-1-phosphate with ATP. Starch synthase then transfers glucose residues from this molecule to the non-reducing end of a pre-existing molecule of starch. Starch consists of two types of glucose polymer, namely amylose and amylopectin. Amylose is a long, unbranched chain of D-glucose units connected via (α1–4) linkages. Amylopectin is a branched form, with (α1–6) linkages forming branches approximately every 24–30 glucose residues.

2.1.2 - RuBP regeneration


Figure 2.2. A simplified (above) and detailed (below) description of the photosynthetic carbon reduction (PCR) cycle. The fixation of CO2 by Rubisco to the acceptor molecule RuBP initiates the cycle with the production of two molecules of PGA. The subsequent, enzyme catalysed, generation of cycle intermediates are cycled to either regenerate RuBP or produce triose phosphates which are precursors for carbohydrate synthesis. The cycle is powered by the co-factors NADPH and ATP that are synthesised from the chloroplast electron transport chain. Enzymes include: PGK, phosphoglycerate kinase; GAPDH, glycraldehyde-3- phosphate dehydrogenase; TIM, triose phoshphate isomerase; ALD, aldolase; FBPase, fructose-1,6-bisphosphatase; TKL, transketolase, SBPase, seduheptulose-1,7-bisphosphatase; RPI, ribose-5-phosphate isomerase; RPE, ribose-5-phosphate epimerase and PRK, phosphoribulose kinase. (Courtesy Robert Sharwood).


Figure 2.2b

Ribulose bisphosphate (RuBP) is consumed in the carboxylating step of carbon fixation. If such fixation is to continue, RuBP must be regenerated, and in this case via the PCR cycle. The PCR cycle operates within the stroma of chloroplasts, and consists of a sequence of 11 steps where a three-carbon compound (3-phosphoglycerate) is phosphorylated, reduced to glyceraldehyde 3-phosphate and isomerised to dihydroxyacetone phosphate. Condensation of this three-carbon compound with glyceraldehyde 3-phosphate yields a six- carbon compound (fructose bisphosphate). Following a series of carbon shunts, involving four-, five- and seven-carbon compounds, RuBP is regenerated.

Important features of the PCR cycle include: (1) for every step of the cycle to occur once, three carboxylations must occur via ribulose bisphosphate carboxylase thus generating six moles of phosphoglycerate (18 carbons); (2) for one turn of the cycle, three molecules of RuBP participate (15 carbons) and thus a net gain of three carbons has occurred for the plant; (3) in regenerating three molecules of RuBP, nine ATP and six NADPH are consumed.

2.1.3 - Properties of Rubisco

Photosynthetic carbon fixation in air is constrained by the kinetic properties of Rubisco. Form I Rubisco in higher plants is a large protein (approximately 550 kDa) comprised of eight large (approx. 50-55 kDa) and eight small subunits (approx. 13-18 kDa) to form an L8S8 hexadecamer. Rubisco synthesis and assembly in higher plants is a complex process whereby the large subunit gene (rbcL) is encoded in the chloroplast genome, while the small subunit genes (rbcS) are encoded as a multi-gene family in the nucleus. The Rubisco small subunits are translated as precursors in thc cytosol and are equipped with a transit-peptide to target them to the chloroplast. Upon import in the chloroplast the transit-peptide is cleaved by a stromal peptidase and the N-terminus modified by methylation of the n-terminal methionine. The large subunits  are synthesised within the chloroplast and also post-translationally modified through the removal of the the N-terminal methionine and serine amino acids and the subsequent acetylation of proline at the N-terminus and the methylation of lysine at position 14. The assembly of large and small subunits into functional hexadecameric Rubisco is reliant on the coordination of chloroplast-localised chaperones.

Despite selection pressure over evolutionary history, Rubisco remains an inefficient catalyst (Spreitzer and Salvucci 2002). Therefore, to achieve a productive maximum CO2 assimilation rate (Amax), plants must compensate for catalytic inefficiency by investing large amounts of nitrogen in Rubisco. Consequently, Rubisco comprises more than 50% of leaf soluble protein in C3 plants. On a global scale, this investment equates to around 10 kg of nitrogen per person!

More than 1000 million years of evolution has still not resulted in a ‘better’ Rubisco adapted for the current and future concentrations of CO2. Such a highly conserved catalytic protein is an outcome of thermodynamic and mechanistic difficulties inherent to this reaction. Rubisco requires carbamylation of the absolutely-conserved residue K201 that is then stabilised by the binding of Mg2+. Without this activation step Rubisco is unable to function. The fixation of CO2 to RuBP to form two molecules of 3-PGA is a five step catalytic process that produces highly reactive transition state intermediates that bind CO2. The highly reactive transition states make Rubisco prone to generating misfiring products, which generate inhibitors within the active site. Therefore, Rubisco requires its own catalytic protection enzyme Rubisco activase. Plants devoid of this enzyme fail to grow properly in air as the activation and subsequent activity of Rubisco is impeded (Portis and Salvucci 2002). Rubisco activase is an ATP-dependent process that removes inhibitors from the active site of Rubisco allowing for activation and catalysis to proceed. Recently, the crystal structure of Rubisco activase has been solved, which will provide key insight into the molecular interaction between Rubisco and Rubisco activase (reviewed by Portis et al. 2008).

Rubisco first evolved when the earth’s atmosphere was rich in CO2, but virtually devoid of O2. With the advent of oxygen-producing photosynthesis by land plants, and the resulting increases in atmospheric O2, one key deficiency of this enzyme became apparent. Rubisco would not only catalyse fixation of CO2 but would also permit incorporation of O2 into RuBP to produce, instead of two molecules of 3-PGA, just one molecule of 3-PGA with one molecule of a two-carbon compound, 2-phosphoglycolate (Section 2.3). Indeed, CO2 and O2 compete directly for access to the active sites of Rubisco. So feeble is Rubisco’s ability to distinguish between these two substrates that in air (20% O2) approximately one molecule of O2 is fixed for every three molecules of CO2.

Fixation of O2 and subsequent photorespiration (Section 2.3) is an energy-consuming process, due to competition between O2 and CO2 for RuBP, plus the energy cost of converting the phosphoglycolate product to a form which can be recycled in the PCR cycle. This energy cost is increased at higher temperatures because O2 competes more effectively with CO2 at the active site of Rubisco. Such sensitivity to temperature × O2 explains why CO2 enrichment, which reduces photorespiration, has a proportionally larger effect upon net carbon gain at higher temperatures than at lower temperatures (Section 13.3).


Figure 2.3. Mechanisms underlying CO2 fixation by Rubisco have changed very little during evolution but Rubisco efficiency has improved. The enzyme in more 'highly evolved' species such as C3 angiosperms is able to fix more CO2 and less O2 in air, reducing photorespiratory energy costs. A measure of this is the relative specificity of Rubisco for CO2, shown here for a range of photosynthetic organisms. (Based on Andrews and Lorimer 1987).

Notwithstanding a meagre catalytic effectiveness in present day Rubisco, more efficient variants would still have had a selective advantage, and especially during those times in the earth’s geological history when atmospheric CO2 concentration was decreasing. Indeed there has been some improvement (Figure 2.3) such that specificity towards CO2 as opposed to O2 has improved significantly. Recently evolved angiosperms show a relative specificity almost twice that of 'older' organisms such as photosynthetic bacteria.

Despite such improvement, Rubisco remains seemingly maladapted to its cardinal role in global carbon uptake, and in response to selection pressure for more efficient variants of CO2 assimilation, vascular plants have evolved with photosynthetic mechanisms that alleviate an inefficient Rubisco. One key feature of such devices is a mechanism to increase CO2 concentration at active sites within photosynthetic tissues. Some of these photosynthetic pathways are dealt with below.

Feature essay 2.1 - The discovery of C<sub>4</sub> photosynthesis

By M.D. (Hal) Hatch

Discovering C4 photosynthesis is an instructive story because it says a lot about progress in science, about serendipity, as well as mindsets and our natural resistance to accept results that conflict with the dogma of the day.


Figure 1. Dr M.D (Hal) Hatch, FAA, FRS, primary discoverer of C4 photosynthesis.

As a rule, the major chemical transformations that occur in plants proceed by exactly the same series of steps in all species. For instance, take the process of respiration where sugars and starch are broken down to CO2 and H2O, yielding energy for living cells. It is almost certain that this proceeds by exactly the same 20 or so steps in species right across the Plant Kingdom. In fact, the same process also operates in yeast, mice and man.

During the 1950s Melvin Calvin and his colleagues at Berkeley resolved the mechanism of photosynthetic CO2 assimilation in the alga Chlorella. Later, they showed that similar steps, with similar enzymes, occurred in a few higher plants. So, by the end of the 1950s it was reasonably assumed that this process, termed the Calvin cycle or photosynthetic carbon reduction (PCR) cycle, accounted for CO2 assimilation in all photosynthetic organisms.

In retrospect, a very observant reader of the plant biological literature of the early 1960s should have noticed that a small group of grass species, including plants like maize, had a set of very unusual but correlated properties, related in one way or another to the process of photosynthesis, that contrasted with the vast majority of other vascular plants. These included an unusual leaf anatomy, substantially higher rates of photosynthesis and growth, higher temperature and light optima for photosynthesis, a much higher water use efficiency, and a very low CO2 compensation point. From this, one might have reasonably concluded that these particular species could be using a different biochemical process for photosynthesis.

We now know that these unusual species fix CO2 by the C4 photosynthetic mechanism. However, the process was not discovered by following up these observations, and only later was the significance of these unusual, correlated features fully appreciated.

During the early 1960s, my colleague Roger Slack and I were working on aspects of carbohydrate biochemistry and sugar accumulation in sugar cane in the research laboratory of the Colonial Sugar Refining Company in Brisbane. Because of these particular interests, we were in regular contact with a laboratory in Hawaii that also worked on sugar cane. We learned from our Hawaiian colleagues, Hart, Kortschak and Burr, that they had seen some unusual results when they allowed sugar cane leaves to fix radioactive carbon dioxide (14CO2), that is, doing the same experiment that Calvin and his colleagues had done earlier with Chlorella. With this procedure radioactivity should be initially incorporated into the first products formed when CO2 is assimilated; in the case of the PCR cycle the radioactivity should appear in the three-carbon compound 3-phosphoglyceric acid (3-PGA) and then in sugar phosphates. However, when these Hawaiian workers first did this experiment as early as 1957 they saw only minor radiolabelling in 3-PGA after brief exposure to 14CO2 and later they showed that most of the radioactivity was located in the four-carbon dicarboxylic acids, malate and aspartate.

We were really intrigued by this result and had often discussed possible interpretations and significance. So when the Hawaiian group published their results a few years later in 1965 we set about repeating and extending these observations to see if we could find out what it all meant.

Before coming to that work it is worth recounting one other interesting twist to the story. In the late 1960s, and several years after we had begun studying C4 photosynthesis, we became aware of a report published some 10 years earlier in a somewhat obscure annual report of a Russian agricultural research institute. This report from a young Russian scientist, Yuri Karpilov, clearly showed that when maize leaves are exposed to radioactive CO2 most of the radioactivity incorporated after 15 s was not in 3-PGA but was in the same dicarboxylic acids, malate and aspartate, that the Hawaiians had found in labelled sugar cane leaves. In a publication about three years later, Karpilov and a more senior Russian scientist speculated that these results may have been due to faulty killing and extraction procedures. It seems doubtful that they appreciated the full significance of this earlier study.

Our initial experiments were designed to trace the exact fate of carbon assimilated by photosynthesis using 14CO2. Sugar cane leaves were exposed to 14CO2 for various periods under steady-state conditions for photosynthesis, then killed and extracted, and the radioactive products were separated by chromatography, identified and degraded to find out which carbons contained radioactivity. We confirmed the results of the Hawaiian group that most of the radioactivity incorporated after short periods in radioactive CO2 was located in the four-carbon acids malate and aspartate. Substantial radioactive labelling of the PCR cycle intermediates occurred only after longer periods (minutes, rather than seconds).

Critical information was subsequently provided by our so-called ‘pulse–chase’ experiments where a leaf was dosed briefly with 14CO2, and then returned to unlabelled air. The biochemical fate of previously fixed 14C can be followed in sequential samples of tissue. These experiments clearly showed a rapid movement of radioactivity from the four-carbon acid malate into 3-PGA and then later to sugar phosphates and finally into sucrose and starch. There were additional critical results from these initial studies: (1) a chemically unstable dicarboxylic acid, oxaloacetic acid, was rapidly labelled as well as malate and aspartate and was almost certainly the true first product formed; (2) fixed CO2 gave rise to the 4-C carboxyl of these four-carbon acids; and (3) this 4-C carboxyl carbon gave rise to the 1-C carboxyl of 3-PGA. Identification of oxaloacetic acid as an early labelled fixation product was an especially demanding task, and involved generation of a stable derivative that would remain intact during extraction and analysis of 14C fixation products.

Spurred on by this success, we then surveyed a large number of species and found radioactive labelling patterns similar to sugar cane in a number of other grass species, including maize, as well as species from two other plant families. This was an exciting result for us at the time since it clearly showed that this mode of photosynthesis was reasonably widespread taxonomically. The next step in determining the exact nature of this process was to discover the enzymes involved. In species such as sugar cane and maize, there proved to be seven enzyme-catalysed reactions involved in the steps unique to C4 photosynthesis, and these included two steps catalysed by enzymes that had never been described before!

Soon after, we named this process the C4 dicarboxylic acid pathway of photosynthesis — after the first product formed. This was later abbreviated to C4 pathway or C4 photosynthesis and the plants employing this process were termed C4 plants.

By 1970 we had a reasonably good understanding of how C4 photosynthesis worked in species like maize and sugar cane (see Section 2.2 for details), and suggested that the reactions unique to C4 photosynthesis might function to concentrate CO2 in the bundle sheath cells of C4 leaves, acting essentially as a CO2 pump. Later, we obtained direct experimental evidence that CO2 was indeed concentrated about 10- to 20-fold in these cells in the light.

As I mentioned earlier, a major departure from Calvin cycle photosynthesis was never expected. Imagine our surprise, therefore, when it was revealed during the early 1970s that there existed not one, but three different biochemical variants for C4 photosynthesis. On this basis C4 species were divided into three groups, and some connections between process and taxonomic background then emerged.

What advantages did all this offer plants over plants that fix CO2 directly by the PCR cycle — that is, using CO2 diffusing directly from air (and distinguished as C3 plants by virtue of their initial three-carbon fixation product phosphoglycerate). As Section 2.2 explains, concentrating CO2 in bundle sheath cells eliminates photorespiration. This, in turn, gives C4 plants distinct advantages in terms of growth and survival, especially at higher temperatures and under strong light. This can be seen most graphically in the distribution of grass species in Australia. In Tasmania, as well as the cooler and wetter southern-most tips of the continent, C4 species are in the minority. However, going north there is a rapid transition and for most of the continent most or all of the grass species are C4.

C4 photosynthesis also offers a potential for growth rates almost twice those seen in C3 plants, but this potential will only be seen at higher temperatures and higher light and this will not be evident in all C4 species. With this kind of growth potential, it is not surprising that C4 species also number among the world’s worst weeds!

As a parting note I should add that about 100 million years ago C3 plants were in their ‘prime’ with atmospheric CO2 concentrations between five and ten times present day levels. However, a new selection pressure then developed. Atmospheric CO2 declined over the next 50–60 million years to something close to our twentieth century levels of about 350 µL–1. This decline almost certainly provided the driving force for evolution of C4 photosynthesis. In other words, C4 photosynthesis was originally ‘discovered’ by nature in the course of overcoming the adverse effects of lower atmospheric CO2 concentration on C3 plants. In effect, C4 processes increase the CO2 concentration in bundle sheath cells to somewhere near the atmospheric CO2 concentration of 100 million years ago.

Further reading

Hatch MD (1987) C4 photosynthesis: a unique blend of modified biochemistry, anatomy and ultrastructure. Biochim Biophys Acta 895: 81–106

Hatch MD (1992) C4 photosynthesis: an unlikely process full of surprises Plant Cell Physiol 33: 333–342

2.2 - C<sub>4</sub> and CAM photosynthesis

Oula Ghannoum, University of Western Sydney, Australia

Approximately 85% of all terrestrial plant species perform C3 photosynthesis, while about 3% fix atmospheric CO2 via the C4 photosynthetic pathway. About 10% of plants carry out crassulation acid metabolism (CAM) and are usually found in highly xeric sites (deserts, epiphytic habitats). C4 plants predominate in open and arid habitats, and also include several important food crops such as maize and sugarcane. This section also covers other, less common photosynthetic modes, such as single-cell C4, C3-C4 intermediate and SAM photosynthesis.

A decline in atmospheric CO2 concentration during past millennia has likely provided the initial impetus for the evolution of C4 photosynthesis. High temperature and low water availability may have constituted additional evolutionary pressures. The key feature of C4 photosynthesis is the operation of a CO2 concentrating mechanism which elevates CO2 concentration around Rubisco sites. Hence, C4 plants have a competitive advantage over C3 plants at high temperature and under strong light because of a reduction in photorespiration and an increase in absolute rates of CO2 fixation at current ambient CO2. Such increase in photosynthetic efficiency results in faster carbon gain and commonly higher growth rates, particularly in subtropical and tropical environments. Consequently, and in response to the looming food security crisis, a global research effort led by IRRI (International Rice Research Institute) is underway to bioengineer C4 photosynthetic traits into major C3 crops, such as rice, in order to boost their photosynthesis, and thus, improve yield and resource use efficiency.

In response to CO2 limitation, not only C3-C4 intermediate, but also CAM and SAM variants have evolved with metabolic concentrating devices which enhance Rubisco performance (Sections 2.2.8 and 2.2.9).

2.2.1 - Evolution of C<sub>4</sub> photosynthesis


Figure 2.3. C4 photosynthesis is an evolutionary development where specialised mesophyll cells initially fix CO2 from the air into 4-carbon acids which are transported to the site of the PCR cycle in the bundle sheath. The bundle sheath cells are relatively impermeable to CO2, so that when the CO2 is released here from the 4-carbon acids, it builds up to high levels. The C4 photosynthetic mechanism is a biochemical CO2 pump. The pathway shown here is overlayed on a micrograph of a C4 leaf, showing bundle sheath and mesophyll cells. Rubisco and the other PCR enzymes are in the bundle sheath cells while phosphoenolpyruvate (PEP) carboxylase is part of the CO2 pump in the mesophyll cells. In C4 plants, after radioactive labelling, 14C appears first in a 4-carbon acid, rather than in 3-PGA. Scale bar = 10 µm. (Original drawings courtesy M.D. Hatch).

One hundred million years ago (Mid-Cretaceous), atmospheric CO2 was between 1500 and 3000 µL L–1, or four to ten times post-industrial levels. Atmospheric CO2 declined during the Oligocene (20-30 million years ago) from the high Tertiary levels (>1000 µL L-1), and oscillated between 180 and 300 µL L-1 for the last 1-3 million years. The Oligocene was also a time when the Earth was dry and the tropics were relatively hot. The earliest origins of C4 photosynthesis date back to this period. Curiously, C4 plants remained in low abundance for a long period of time. According to stable carbon isotopic data, a worldwide expansion of C4 grasslands and savannas occurred during the Late Miocene and Pliocene (3 to 8 million years ago), most probably through the displacement of C3 vegetation (Edwards et al. 2010).

Under the early high concentration of CO2, photorespiration of C3 plants was inhibited (Section 2.3) so that photosynthetic efficiency was higher than it is now. In addition, maximum photosynthetic rates were double twentieth century values, and the energy cost of photosynthesis would have been around three ATP and two NADPH per molecule of CO2 fixed. As atmospheric CO2 concentrations declined to approximately 250–300 µL L–1, photosynthetic rates were halved, photorespiration increased substantially, photosynthetic efficiency declined and the energetic costs of photosynthesis increased to approximately five ATP and 3.2 NADPH per CO2 molecule fixed. Such events would have generated a strong selection pressure for genetic variants with increased carboxylation efficiency and increased photosynthetic rates.

Angiosperms have a higher relative specificity of Rubisco for CO2 than ferns and mosses (see examples of other less evolutionarily advanced species in Figure 2.3). Such differences imply minor evolution in this highly conserved molecule of Rubisco and there is little variation between species of vascular plants. Consequently, alteration of Rubisco in response to a changing atmospheric CO2 concentration has not been an option.

By contrast, evolution of a new photosynthetic pathway (C4) has occurred independently and on many occasions in diverse taxa over 25 to 30 million years as CO2 levels declined. Despite its complexity, C4 photosynthesis evolved more than 60 independent times in 19 distantly related flowering families. About 50% of C4 species are grasses (Poaceae) with ~18 distinct origins distributed over 370 genera and ~4600 species (Sage et al. 2011). The oldest identifiable fossils with pronounced bundle sheath layers are seven million years old, although necessary metabolic pathways could have evolved earlier, prior to this adaptation in anatomy. C4 plants are known to differ from C3 plants in their discrimination against atmospheric 13CO2, and shifts in the stable carbon isotope signature of soil carbonate layers that reflect emergence of C4 plants have been dated at 7.5 million years bp. Modern evidence from molecular phylogeny places the origin of the main C4 taxa at 25-30 million years ago (Christin et al. 2009). By inference, C4 photosynthesis evolved in response to a significant decline in atmospheric CO2 concentration, from 1500–3000 µL L–1 to about 300 µL L–1. By evolving a CO2-concentrating mechanism, C4 plants presented their Rubisco with an elevated partial pressure of CO2 despite lower atmospheric CO2. As a consequence, photorespiration was inhibited, maximum photosynthetic rates increased and energetic costs reduced.

2.2.2 - The CO<sub>2</sub> concentrating mechanism in C<sub>4</sub> photosynthesis

The C4 pathway (Figure 2.3) is ‘a unique blend of modified biochemistry, anatomy and ultra-structure’ (Hatch 1987). The classical C4 syndrome in most terrestrial plants consists of two photosynthetic cycles (C3 (or PCR) and C4) operating across two photosynthetic cell types (mesophyll and bundle sheath), which are arranged in concentric layers around the vascular bundle, also known as the kranz anatomy.


Figure 2.5. CO2 photosynthesis response curves show that C4plants have a higher affinity for CO2. At common ambient levels of CO2, photosynthesis in a C4 leaf is almost fully CO2-saturated, whereas a C3 plant is operating at only one-half to two-thirds maximum rate. This contrast is due to the CO2-concentrating function of C4 photosynthesis. More sophisticated measurement of leaf assimilation as a function of intercellular CO2 (Figures 1-3 in Case study 1.1) can be used to reveal component processes. (Original drawings courtesy M.D. Hatch)

Initial and rapid fixation of CO2 within mesophyll cells results in the formation of a four-carbon compound which is then pumped to bundle sheath cells for decarboxylation and subsequent incorporation into the PCR cycle in that tissue. This neat division of labour hinges on specialised anatomy and has even resulted in evolution of distinct classes of chloroplasts in mesophyll compared with bundle sheath cells. Three biochemical variants of C4 photosynthesis (termed subtypes) are known to have evolved from C3 progenitor and in all cases with a recurring theme where the C4 cycle of mesophyll cells is complemented by a PCR cycle in bundle sheath cells, where Rubisco is exclusively localised. In effect, a biochemical ‘pump’ concentrates CO2 at Rubisco sites in bundle sheath cells thereby sustaining faster net rates of CO2 incorporation and virtually eliminating photorespiration. For this overall mechanism to have evolved, a complex combination of cell specialisation and differential gene expression was necessary. Figure 2.3a shows a low-magnification electron micrograph of a C4 leaf related to a generalised scheme for the C4 pathway.


Figure 2.6. Rubisco can be localised in transverse sections of leaves by indirect immunofluorescent labelling where treated sections are viewed in conjunction with autofluorescence controls. Tissues such as bundle sheath extensions and epidermes fluoresce naturally, and such emission has to be ‘subtracted’ from present images. Considering (a), all chloroplasts in this C3 grass leaf (Microlaena stipoides) show a strong yellow fluorescence, indicating general distribution of Rubisco, and hence operation of the PCR cycle. By contrast, in (b), the C4 grass (Digitaria brownii) has restricted Rubisco to bundle sheath cells. In that case, mesophyll cells are devoid of Rubisco, fixing CO2 via the action of phosphoenolpyruvate carboxylase into a four-carbon acid which moves to bundle sheath cells, there providing CO2 for subsequent relaxation via Rubisco and the PCR cycle. Scale bar = 100 µm. (Original light micrographs courtesy Paul Hattersley).

By analogy with Calvin’s biochemical definition of the C3 pathway at Berkeley in the 1950s, the C4 pathway was also delineated with radioactively labelled CO2 (see Feature Essay 2.1). Significantly, and unlike C3 plants, 3-PGA is not the first compound to be labelled after a 14C pulse (Figure 2.3b). Specialised mesophyll cells carry out the initial steps of CO2 fixation utilising the enzyme phosphoenolpyruvate (PEP) carboxylase. The product of CO2 fixation, oxaloacetate, is a four-carbon organic acid, hence the designation ‘C4’ photosynthesis (or colloquially, C4 plant). A form of this four-carbon acid, either malate or aspartate depending on the C4 subtype, migrates to the bundle sheath cells which contain Rubisco and the PCR cycle. In the bundle sheath cells, CO2 is removed from the four-carbon acid by a specific decarboxylase and a three-carbon product returns to the mesophyll to be recycled to PEP for the carboxylation reaction. Thus, label first appears in the four-carbon acid after 14C feeding, followed by 3-PGA and, finally, in sucrose and starch (Figure 2.1b).

A physical barrier to CO2 diffusion exists in the thickened walls of the bundle sheath cells (lined with suberised in some C4 species), preventing CO2 diffusion back to the mesophyll and allowing CO2 build up to levels at least 10 times those of ambient air. Build up of CO2 in the bundle sheath is also facilitated by the higher activity ratio (2-4 times) of PEP carboxylase relative to Rubisco in C4 plants. Rubisco is thus exposed to a saturating concentration of CO2 which both enhances carboxylation due to increased substrate supply, and forestalls oxygenation of RuBP (hence photorespiration) by outcompeting O2 for CO2 binding sites on Rubisco (Figure 2.5).

In leaves of C3 plants, the PCR cycle operates in all mesophyll chloroplasts, but in C4 plants the PCR cycle is restricted to bundle sheath cells (Figure 2.3). Rubisco is pivotal in this cycle, and can be used as a marker for sites of photosynthetic carbon reduction. Rubisco was visualised by localising this photosynthetic enzyme with antibodies via indirect immunofluorescent labelling (Hattersley et al. 1977; Figure 2.6). In this pioneering method, ‘primary’ rabbit anti-Rubisco serum (from rabbits injected with purified Rubisco) is first applied to fixed transverse sections of leaves. Rabbit antibodies to Rubisco bind to the enzyme in situ. Then ‘secondary’ sheep anti-rabbit immunoglobulin tagged with a fluorochrome (fluorescein isothiocyanate) is applied to the preparation. This fluorochrome binds specifically to the rabbit antibodies and fluoresces bright yellow wherever Rubisco is located (blue light excitation using an epifluorescence light microscope).

In the C3 grass Microlaena stipoides (Figure 2.6a), all chloroplasts are fluorescing bright yellow and this indicates wide distribution of Rubisco throughout mesophyll tissue. By contrast, only bundle sheath cells are equipped with Rubisco in the C4 grass Digitaria brownii (Figure 2.6b). These two native Australian grasses co-occur in the ACT but contrast in relative abundance. M. stipoides (weeping grass) is common in dry sclerophyll woodlands throughout southeast temperate Australia, whereas D. brownii (cotton panic grass) in the ACT is at the southern end of its distribution, being far more abundant in subtropical Australia and, in keeping with its C4 physiology, especially prevalent in semi-arid regions.

2.2.3 - Energetics of C<sub>4</sub> photosynthesis


Figure 2.7. Generalised light response curves for leaf photosynthesis show that C4 plants assimilate comparatively faster at high temperature (35°C), but that C3 plants are advantaged at low temperature (10°C). Photorespiration increases with temperature, and is largely responsible for this contrast. C4 plants are equipped with a CO2-concentrating device in their bundle sheath tissue which both enhances Rubisco’s performance at that location, and forestalls photorespiratory loss. (Original drawings coustesy M.D. Hatch).

One disadvantage of the C4 pathway is that an energy cost is incurred by C4 plants to run the CO2 ‘pump’. This is due to the ATP required for recycling PEP from pyruvate by the chloroplastic enzyme pyruvate, Pi dikinase in the mesophyll cells (Figure 2.4 and Hatch 1987). Under ideal conditions, five ATP and two NADPH are required for every CO2 fixed in C4 photosynthesis (two ATP are required to run the CO2 pump, i.e., regenerate PEP). In addition, a proportion (20-30%) of CO2 fixed by PEP carboxylase in the mesophyll is not fixed by Rubisco in the bundle sheath, and subsequently leaks back to the mesophyll. This leaked (or overcycled) CO2 represents an additional, inherent energetic cost of the C4 pathway.

From the previous section, the C4 pathway is obviously energetically more expensive than the C3 pathway in the absence of photorespiration. However, at higher temperatures the ratio of RuBP oxygenation to carboxylation is increased and the energy requirements of C3 photosynthesis can rise to more than five ATP and three NADPH per CO2 fixed in air (for these calculations see Hatch 1987).

Representative light response curves for photosynthesis in C3 cf. C4 plants (Figure 2.7) can be used to demonstrate some of these inherent differences in photosynthetic attributes. At low temperature (10°C in Figure 2.7) a C3 leaf shows a steeper initial slope as well as a higher value for light-saturated photosynthesis. By implication, quantum yield is higher and photosynthetic capacity is greater under cool conditions. In terms of carbon gain and hence competitive ability, C3 plants will thus have an advantage over C4 plants at low temperature and especially under low light.

By contrast, under warm conditions (35°C, upper curves in Figure 2.7) C4 photosynthesis in full sun greatly exceeds that of C3, while quantum yield (inferred from initial slopes) remains unaffected by temperature. Significantly, C3 plants show a reduction in quantum yield under warm conditions (compare 10°C and 35°C curves; right side of Figure 2.7). At 35°C C3 plants also show lower rates of light-saturated assimilation compared with C4 plants. Increased photorespiratory losses from C3 leaves at high temperature are responsible (Section 2.3). C4 plants will thus have a competitive advantage over C3 plants under warm conditions at both high and low irradiance.

2.2.4 - The biochemical subtypes of C<sub>4</sub> photosynthesis

C4 photosynthesis calls for metabolic compartmentation which is in turn linked to specialised anatomy (Figure 2.4). Three biochemical subtypes of C4 photosynthesis have evolved which probably derive from subtle differences in the original physiology and leaf anatomy of their C3 progenitors.

CO2 assimilation by all three C4 subtypes (Figure 2.8) involves five stages:

  1. carboxylation of PEP in mesophyll cells, thereby generating four-carbon acids (malate and/or aspartate);
  2. transport of four-carbon acids to bundle sheath cells;
  3. decarboxylation of four-carbon acids to liberate CO2;
  4. re-fixation of this CO2 via Rubisco within the bundle sheath, using the C3 pathway;
  5. transport of three-carbon acid products following decarboxylation back to mesophyll cells to enable synthesis of more PEP.


Figure 2.8. C4 plants belong to one of three subtypes represented here (left to right) as NADP-ME, NAD-ME and PCK. Each subtype has a distinctive complement and location of decarboxylating enzymes, and each differs with respect to metabolites transferred between mesophyll and bundle sheath. The path of carbon assimilation and intracellular location of key reactions are shown for each of these biochemically distinct subtypes. Heavy arrows indicate the main path of carbon flow and associated transport of metabolites. Enzymes involved (numbers shown in parentheses) are as follows: (1) PEP carboxylase, (2) NADP-malate dehydrogenase, (3) NADP malic enzyme, (4) pyruvate Pi dikinase, (5) 3-PGA kinase and GAP dehydrogenase, (6) aspartate aminotransferase, (7) NAD-malate dehydrogenase, (8) NAD-malic enzyme, (9) alanine aminotransferase, (10) PEP carboxykinase, (11) mitochondrial NADH oxidation systems. In PCK-type C4 plants, the PGA/DHAP shuttle would also operate between cells as indicated for NADP-ME and NAD-ME. Cycling of anaino groups between mesophyll and bundle sheath cells involves alanine and alanine aminotransferase. (Original diagram courtesy M.D. Hatch).

Recognising some systematic distinctions in whether malate or aspartate was transported to bundle sheath cells, C4 plants were further subdivided into three subtypes according to their four-carbon acid decarboxylating systems and ultrastructural features (Hatch et al. 1975). Members of each subtype contain high levels of either NADP-malic enzyme (NADP-ME), phosphoenolpyruvate carboxykinase (PCK) or NAD-malic enzyme (NAD-ME) (so designated in Figure 2.8). High NADP-malic enzyme activity is always associated with higher NADP-malate dehydrogenase activity, while those species featuring high activities of either of the other two decarboxylases always contain high levels of aminotransferase and alanine aminotransferase activities. As a further distinction, each of the decarboxylating enzymes is located in bundle sheath cells; NAD-malic enzyme is located in mitochondria but PEP carboxykinase is not.

In all three subtypes, the primary carboxylation event occurs in mesophyll cytoplasm with PEP carboxylase acting on HCO3 to form oxaloacetate. However, the fate of this oxaloacetate varies according to subtype (Table 2.1; Figure 2.8). In NADP-ME species, oxaloacetate is quickly reduced to malate in mesophyll chloroplasts using NADPH. By contrast, in NAD-ME and PCK species, oxaloacetate is transaminated in the cytoplasm, with glutamate donating the amino group, to generate aspartate. Thus, malate is transferred to bundle sheath cells in NADP-ME species and aspartate is transferred in NAD-ME and PCK species. The chemical identity of three-carbon acids returned to mesophyll cells varies accordingly.

In NADP-ME species, only chloroplasts are involved in decarboxylation and subsequent carboxylation via the PCR cycle (Figure 2.8). By contrast, in NAD-ME and PCK species, chloroplasts, cytoplasm and mitochondria are all involved in moving carbon to the PCR cycle of bundle sheath chloroplasts. In NAD-ME and PCK species, aspartate arriving in bundle sheath cells is reconverted to oxaloacetate in either mitochondria (NAD-ME) or cytoplasm (PCK) (Table 2.2). Reduction and decarboxylation of oxaloacetate occurs in mitochondria of NAD-ME species and CO2 is thereby released for fixation by chloroplasts of bundle sheath cells. In PCK species, oxaloacetate in the cytoplasm is decarboxylated by PCK, thereby releasing CO2 for fixation in bundle sheath chloroplasts (Figure 2.8).

Transport of metabolites to bundle sheath cells

A rapid transfer of malate and aspartate to bundle sheath cells from mesophyll cells is required if the CO2 concentration in bundle sheath cells is to stay high. A very high density of plasmodesmata linking bundle sheath cells to mesophyll cells facilitates this traffic. Consequently, the permeability coefficient of C4 bundle sheath cells to small metabolites such as four-carbon acids is about 10 times larger than that of C3 mesophyll cells (Table 2.2). However, coupled with this need for a high permeability to metabolites moving into bundle sheath cells is a low permeability to CO2 molecules so that CO2 released through decarboxylation in the bundle sheath does not diffuse rapidly into mesophyll air spaces. For some species, a layer of suberin in the cell wall of bundle sheath–mesophyll junctions (suberin lamella) significantly reduces CO2 efflux (Table 2.2).

Centrifugal versus centripetal chloroplasts

Not all species contain a suberin layer, but all C4 plants have a need to prevent CO2 from diffusing quickly out of bundle sheath cells, so that the location of chloroplasts of bundle sheath cells becomes critical in those species lacking a suberin layer (Figure 2.9). Where species have a suberin layer, chloroplasts are located in a centrifugal position, that is, on the wall furtherest away from the centre of the vascular bundle lying in the middle of the bundle sheath (Figure 2.9E, F). In those C4 species lacking a suberin layer, chloroplasts are located centripetally, that is, on the wall closest to the centre of the vascular bundle lying within the bundle sheath (Figure 2.9A, B). Such a location would help restrict CO2 diffusion from bundle sheath to mesophyll cells.

Fig 2.14.jpg

Figure 2.9. C4 plants belong to one of three subtypes shown here in cross-section as light micrographs (left side) and electron micrographs (right side). Top to bottom, these subtypes are designated NAD-ME (A, B) PCK (C, D) and NADP-ME (E, F). Features common to all subtypes include a vascular bundle (V), bundle sheath (B), mesophyll tissue (M) and chloroplasts (C).

Subtype NAD-ME (A, B) is represented by Amaranthus edulis and shows bundle sheath cells with centripetally located chloroplasts containing small starch grains and surrounded by mesophyll cells. The accompanying electron micrograph of a cytoplasmic region of a bundle sheath cell shows chloroplasts and numerous large mitochondria. Scale bar in A = 50 µm; in B = 2 µm.

Subtype PCK (C, D) is represented by Chloris gayana with chloroplasts arranged around the periphery of bundle sheath cells and adopting a centrifugal position. Mitochondria show well-developed internal membrane structures. Scale bar in C = 25 µm; in D = 3 µm.

Subtype NADP-ME is represented by Zea mays where the bundle sheath contains centrifugally located chloroplasts with numerous starch grains, but lacking grana. Chloroplasts in adjacent mesophyll cells are strongly granal. Bundle sheath cells contain few mitochondria and these show only moderate development of internal membrane structures. Scale bar in E = 25 µm; in F scale bar = 2 µm (Micrographs courtesy Stuart Craig and Celia Miller).

Regulation of C4 photosynthesis

Fixation of CO2 by C4 plants involves the coordinated activity of two cycles in separate anatomical compartments (Figure 2.8). The first cycle is C4 (carboxylation by PEP carboxylase), the second is C3 (carboxylation by Rubisco). Given this biochemical and anatomical complexity, close regulation of enzyme activities is a prerequisite for efficient coordination.

PEP carboxylase, NADP-malate dehydrogenase and pyruvate orthophosphate dikinase are all light-regulated and their activities vary according to irradiance. NADP-malate dehydrogenase is regulated indirectly by light via the thioredoxin system.

PEP carboxylase in C4 plants exists in the same homo-tetramer in light- and dark-acclimated leaves. This is in marked contrast to CAM species where different forms exist in light- and dark-acclimated leaves. In C4 plants, PEP carboxylase has extremely low activity at night, thus preventing uncontrolled consumption of PEP. Such complete loss of activity in darkness is mediated via divalent metal ions, pH plus allosteric activators and inhibitors. As a consequence, and over a period of days, C4 plants can increase or decrease PEP carboxylase in response to light regime.

2.2.5 - Environmental physiology of C<sub>3</sub> versus C<sub>4</sub> photosynthesis

Rubisco is characterised by its low affinity for its productive substrate, CO2 and slow catalytic turnover rate (i.e., 1-3 cycles per sec). Importantly, Rubisco reacts with O2 (photorespiration), and this culminates in loss of CO2 and energy. In C3 plants, photorespiration can drain more than 25% of fixed CO2 under non-stressful conditions. The ratio of photorespiration to photosynthesis increases with increasing temperature and decreasing intercellular CO2 such as occurs when stomatal conductance is reduced under water stress. C3 plants compensate for Rubisco’s inefficiencies by (i) opening their stomata to increase CO2 diffusion into chloroplasts, which increases water loss and lowers leaf-level water use efficiency, WUE; and (ii) investing up to 50% of leaf nitrogen in Rubisco, which lowers their leaf-level nitrogen use efficiency, NUE.

The C4 pathway supercharges photosynthesis and suppresses photorespiration by operating a CO2 concentrating mechanism which elevates CO2 around Rubisco. Although C4 photosynthesis incurs additional energy, the energy cost of photorespiration exceeds that of the CO2 concentrating mechanism above 25oC. Hence, higher radiation use efficiencies (i.e., efficiency of converting absorbed radiation into biomass) have been recorded for C4 than C3 crops. High bundle sheath CO2 concentration saturates C4 photosynthesis at relatively low intercellular CO2, allowing C4 plants to operate with lower stomatal conductance. Thus, leaf-level WUE is usually higher in C4 than C3 plants. Relative to C3 plants, Rubisco of C4 plants is faster (higher turnover rate) and operates under saturating CO2. Thus, C4 plants typically achieve higher photosynthetic rates with about 50% less Rubisco and less leaf nitrogen. Hence, photosynthetic NUE is higher in C4 than C3 plants. Accordingly, C4 plants are advantaged relative to C3 plants in hot and nitrogen-poor environments with short growing seasons, hence their great abundance in wet/dry tropics such as Northern Territory savannas.

As mentioned earlier, more than 50% of C4 plants are grasses. C4 grasses are confined to low latitudes and altitudes, whereas C3 species dominate at higher latitudes and altitudes. Generally, C4 species frequently occur in regions of strong irradiance. Ehleringer and colleagues (Ehleringer et al. 1997) proposed that these distribution patterns are best explained by the different responses of photosynthetic quantum yields to temperature between C3 and C4 plants.

C4 photosynthesis suppresses photorespiration by operating a CO2 concentrating mechanism that comes at additional energetic cost. This cost is independent of ambient CO2 and temperatures. In contrast, photorespiration (and its associated energy cost) increases steeply with temperature in C3 plants and is highly dependent on CO2 concentrations. Under saturating irradiance and current ambient atmospheric CO2 concentration, the threshold temperature where the cost of photorespiration in C3 plants exceeds that of the CO2 concentrating mechanism in C4 plants is estimated around 25oC. This model provides a physiological basis for understanding today’s contrasting geographic distribution between C3 and C4 grasses.

As an example, the C4 grasses of the northern Australian savannas are relatively un-shaded because of the low tree density and sparse canopy. Light is abundant and since the CO2 concentration inside C4 leaves is high, a potentially high rate of light-saturated assimilation can be exploited. Most C3 species reach light saturation in the range of one-eight to one-half full sunlight (Figure 2.7). In C4 species, canopy assimilation might not become light saturated even in full sunlight. Cplants thus maintain a competitive advantage over C3 plants in tropical locations, where average daily light receipt is much larger than in temperate zones, and associated with warmer conditions that also favour C4 photosynthesis (Figure 2.6). Given strong sunlight, warmth and seasonally abundant water, biomass production by C4 plants is commonly double the rate for C3 plants. Typically, C3 plants produce 15–25 t ha–1 but C4 plants easily produce 35–45 t ha–1.

Physiological characteristics of the C4 subtypes

As outlined in previous sections, characteristic biochemical, anatomical and physiological traits are associated with each of the three “classical” C4 subtypes (Table 2.3). However, it should be noted that many C4 plants have leaf structures that fall outside the “classical” subtype division (eg, NADP-ME tribes, Arundinelleae and Neurachneae). As many as 11 anatomical-biochemical suites have been identified in C4 grasses. A curious aspect about the subtypes of C4 grasses is their biogeography. In Australia and elsewhere, NADP-ME grasses are more frequent at higher rainfall, NAD-ME grasses predominate at lower rainfall, while the distribution of PCK grasses is even across rainfall gradients (Hattersley 1992).

2.2.6 - Single-cell C<sub>4</sub> photosynthesis

The fundamental paradigm underpinning the efficiency of C4 photosynthesis in terrestrial plants is the ‘division of labour’ between the initial fixation of CO2 into C4 acids, and their subsequent utilisation to generate high concentrations of CO2 for ultimate fixation by Rubisco. The basic model for C4 plants with classical kranz anatomy consists of two photosynthetic cycles (C3 and C4) operating across two photosynthetic cell types (mesophyll and bundle sheath), with strict cell- and organelle-specific localisation of key enzymes and with sufficient resistance to CO2 back-diffusion. Indeed, the discovery of the kranz anatomy by Haberlandt preceded that of C4 biochemistry by a century. The prevailing consensus has been that efficient C4 photosynthesis necessitates the collaboration of two cell types.

Recently, this notion has been challenged by the discovery of non-kranz or single-cell C4 photosynthesis in shrubs (Borszczowia aralocaspica and Bienertia cycloptera; Chenopodiaceae family) found in the salt deserts of Central Asia (Voznesenskaya et al. 2002, 2003). These plants show CO2 and O2 responses typical of C4 photosynthesis but lack the kranz anatomy. They perform C4 photosynthesis through the spatial localisation of dimorphic chloroplasts (as well as other organelles and photosynthetic enzymes) in distinct positions within a single chlorenchyma cell. Yet, the details of the partitioning differ between the two species (Edwards et al. 2004).

In Bienertia, the central cytoplasmic compartment of the chlorenchyma cell plays the role of bundle sheath cells in kranz-type C4 (NAD-ME) plants; it is filled with mitochondria surrounded by chloroplasts. The peripheral cytoplasm lacks mitochondria and plays the role of the mesophyll cell in kranz-type C4 plants. Accordingly, chloroplastic Rubisco and mitochondrial NAD-ME and glycine decarboxylase are restricted to the central compartment; chloroplastic pyruvate, Pi dikinase is restricted to the peripheral compartment, which is highly enriched with cytosolic PEP carboxylase. In Borszczowia, the compartmentation occurs at the distal (mesophyll equivalent) and proximal (bundle sheath equivalent) ends of the elongated, cylindrical chlorenchyma cell. The inter-connecting cytoplasm between the two intra-cellular compartments provides a liquid diffusion path, thus replacing the role of the bundle sheath cell wall in kranz-type C4 plants (Edwards et al. 2004).

A low conductance for CO2 diffusion out of the bundle sheath cells (or its equivalent cellular compartment) is critical for the efficient operation of C4 photosynthesis. The total diffusive resistance to CO2 has multiple components with different levels of contribution. These components include bundle sheath walls, membranes, bundle sheath chloroplast position, the site of C4 acid decarboxylation, and the liquid-phase diffusion path. For kranz-type C4 plants, calculated total bundle sheath resistance on a leaf area basis can range from 50 to 150 m2 s-1 mol-1 (von Caemmerer and Furbank 2003). Evidently, single-cell C4 plants have sufficient resistance to CO2 back-diffusion which is essentially made of the cytoplasmic liquid phase and the special localisation of the (Rubisco-containing) chloroplasts surrounding the mitochondria (site of C4 acid decarboxylation).

Thus, single-cell C4 plants have efficient photosynthesis which is not inhibited by O2, and their carbon isotope values are similar to kranz-type C4 plants. Although, single-cell C4 photosynthesis breaks away from the classical kranz anatomy, it remains within the general ‘division of labour’ paradigm.

2.2.7 - C<sub>3</sub>-C<sub>4</sub> photosynthesis

More than 40 eudicot and monocot species distributed over 21 lineages have been reported to possess intermediate C3 and C4 photosynthetic characteristics and CO2 compensation points. These intermediate species are likely remnants of the complex processes that led to the evolution of C4 plants from C3 ancestors, although reversions from the C4 condition have also been suggested. Moreover, a number of identified C3-C4 species occur in taxa that are not closely related to any C4 lineage, raising the possibility that the C3-C4 photosynthetic pathway may be a distinct adaptation. This, in addition to the small number of intermediate species found so far cast doubts over their physiological and ecological fitness, and whether they represent living fossils of evolutionary paths or evolutionary dead-ends (Rawsthorne 1992; Sage et al. 2011).

Leaves of all C3-C4 intermediates have partial or full kranz anatomy, with prominent bundle sheath cells containing chloroplasts and other organelles, and intermediate interveinal distances. Bundle sheath chloroplasts contain Rubisco and functional PCR cycle in both mesophyll and bundle sheath cells. Intermediate leaves also have CO2 compensation points that are lower than what is observed for C3 leaves and can be indistinguishable from C4 leaves, due to reduced photorespiration. Biochemically, C3-C4 intermediates differ in the level of activity of the C4 cycle and the extent to which CO2 is concentrated in bundle sheath cells.


Figure 2.10. Schematic representation of the ‘photorepiratory pump’ operating in C3-C4 photosynthesis. The intermediate photosynthetic pathway reduces photorespiration by refixing photorespired CO2 released locally in the bundle sheath cell. Mesophyll mitochondria lack glycine decarboxylase activity. Mesophyll and bundle sheath cells contain chloroplasts with functional Calvin cycle. Abbreviations: cp: chloroplast; GDC: glycine decarboxylase; PCO: photosynthetic oxidative cycle; PCR: photosynthetic reductive cycle; mt: mitochondrion.

C3-C4 intermediate plants reduce photorespiration (and hence, CO2 compensation point) using a ‘photorespiratory pump’ based on modified localisation of the mitochondrial photorespiratory enzyme, glycine decarboxylase (Figure 2.10). In these plants, glycine decarboxylase activity is restricted to bundle sheath cells and excluded from mesophyll cells. Consequently, photorespired CO2 is released in the bundle sheath where it is largely refixed by Rubisco and the bundle sheath PCR before it diffuses back to the mesophyll. Such a system may weakly elevate CO2 in bundle sheath cells. Intermediate species that rely on the ‘photorespiratory pump’ are termed C3-like or Type I intermediates (e.g., Panicum milioides, Flaveria pubescens) and have intermediate CO2 compensation points and negligible C4 cycle activity.

In Type II and C4-like intermediates (e.g., Flaveria brownii), up to 70% of atmospheric CO2 may be first fixed into C4 acids. These plants have C4-like CO2 compensation points but are not classified as C4 plants because they lack the strict localisation of photosynthetic enzymes (e.g., Rubisco is present in mesophyll cells) and their bundle sheath cell walls have high CO2 permeability, resulting in only a partial CO2 concentrating mechanism (Brown 1980; Ku et al. 1991; Rawthorne 1992; Vogan and Sage 2011).

The physiological advantages of the intermediate photosynthetic pathway in all its naturally occurring forms remain unclear. It may be hypothesised that lowered photorespiration may lead to reduced CO2 limitation of photosynthesis, and thus allow C3-C4 plants to operate with lower stomatal conductance, thus conferring higher water use efficiency relative to C3 counterparts. Moreover, increased nitrogen cost associated with ‘building’ another set of photosynthetic cells (bundle sheath) may reduce nitrogen use efficiency if the gains in CO2 uptake are not substantial.

Work conducted with C3-C4 species yielded inconclusive evidence on the likely advantages of C3-C4 photosynthesis relative to the ancestral C3 mode. Generally, these studies demonstrated that, short of substantial C4 cycle activity and advanced cell-specific localisation of C3 and C4 cycle enzymes between the mesophyll and bundle sheath cells, C3-C4 photosynthesis does not improve photosynthetic efficiency (Bolton and Brown 1980; Pinto et al. 2011, Vogan and Sage 2011). Therefore, partial recycling of photorespired CO2 or a partial CO2 concentrating mechanism reduce photorespiratory loss normally associated with C3 photosynthesis, without leading to significant gains in plant fitness or productivity.

2.2.8 - Crassulacean acid metabolism (CAM)

Joseph Holtum1, Klaus Winter2 and Barry Osmond3

1Centre for Tropical Biodiversity and Climate Change, James Cook University, Australia; 2Smithsonian Tropical Research Institute, Balboa, Ancón, Republic of Panama; 3School of Biological Sciences, University of Wollongong, and Research School of Biology, Australian National University, Australia

Crassulacean acid metabolism (CAM) is a water-conserving mode of photosynthesis that, like C4 photosynthesis, is a modification of the C3 photosynthetic pathway fitted with a CO2 concentrating mechanism (CCM) that can increase the [CO2] around ribulose bisphosphate carboxylase/oxygenase (Rubisco) by more than 10-fold and suppress photorespiration. The overall energy demand of the CAM pathway is only about 10% more than that of C3 photosynthesis, as costs of the CCM machinery are partially offset by reducing photorespiration.

In C4 plants, as explained earlier in Section 2.2.2, this CCM is most commonly achieved by an “in-line turbocharger” based on initial CO2 fixation by phosphoenolpyruvate carboxylase (PEPC) into C4 acids in the cytoplasm of outer mesophyll cells. These acids diffuse rapidly to adjacent relatively CO2-tight bundle-sheath cells (Figure 2.31 right) where CO2 is released again. High [CO2] builds up in this spatially separated compartment where it is refixed by Rubisco.


Figure 2.31 Leaf transverse sections of CAM versus C3 and C4 plants. Left: succulent CAM plant Kalanchoë daigremontiana. Centre: C3Atriplex hastata. Right: C4Atriplex spongiosa. Scanning electron  micrographs at similar magnification. (K. daigremontiana  image courtesy R.A. Balsamo and E.G. Uribe; Atriplex spp. images courtesy J. H. Troughton)

In CAM plants enzyme systems analogous to those in C4 plants achieve the same result through a “battery-like” dark accumulation of CO2 into the 2nd carboxyl group of malic acid (acidification phase) in the vacuole of large mesophyll cells (Figure 2.31 left). Malic acid can accumulate to very high concentrations, attaining concentrations of greater than 1 mole acid per litre in mesophyll cells of tropical tree-CAM plants (Clusia spp.). Indeed one can sometimes taste the acid, with acid taste-testing for the presence of CAM being possibly first recorded in Aloe sp. by Nehemiah Grew in 1682 and in field reports from India by Benjamin Heyne in 1815.

In the light, malic acid returns to the cytoplasm where it is rapidly decarboxylated (deacidification phase). The CO2 released, which accumulates to high internal [CO2] as stomata close, is refixed by Rubisco in chloroplasts of the same mesophyll cell where it is further assimilated by the photosynthetic carbon reduction (PCR) cycle (Figure 2.32).  


Figure 2.32  Schematic of the principal components of CAM, highlighting storage of malic acid in the vacuole (V) and the carbohydrate conundrum discussed later. (Diagram courtesy B. Osmond)

Ultimately three of the four carbons recovered from the malic acid must be stored as starch and/or sugars in order to provide to PEPC the C3 substrate required for CO2 uptake during the following night. The fourth carbon, effectively that obtained from the atmosphere, is available for growth. Deacidification may generate high [CO2] behind closed stomata but photorespiration is not completely abolished (Lüttge 2002) since photosynthesis also generates high internal [O2]. While exploring Lake Valencia in Venezuela in 1800, Alexander von Humboldt measured elevated [O2] in bubbles streaming from the cut base of presumably CAM Clusia leaves standing in water in the light.

Of course by closing stomata in the light CAM plants minimize water loss when evaporative demand is highest (von Caemmerer and Griffiths 2009). The biomass production per unit water utilized in CAM was 6 times higher than for C3 plants and 2 times higher than for C4 plants when plants exhibiting all three photosynthetic pathways were grown together in a garden outdoors (Winter et al. 2005). The attribute of water-use efficiency undoubtedly contributes significantly to the success of CAM photosynthesis in nature, with CAM species outnumbering C4 species by about two to one. The paradoxes of CAM, a mode of photosynthesis that involves stomatal opening and CO2 uptake during the dark, continue to inform many aspects of plant biochemistry, physiology, ecology and evolution. This article draws heavily on two recent reviews (Borland et al. 2011; Winter et al. 2015). - Biochemical attributes distinctive to CAM

Although CAM and C4 photosynthesis share common enzyme machineries, the physiological bases of spatially-separated and time-separated CCMs are very different and involve complex suites of distinctive regulatory processes ranging from allosteric modulation of enzyme activities, through cell and organelle membrane metabolite transport systems, to long-term responses to stress. The resulting metabolism is rarely at steady state. It is thus helpful to reference the principal biochemical interacting components of CAM to the CO2 exchange patterns and the pool sizes of acidity and carbohydrates in the archetypical Kalanchoë daigremontiana as outlined in Figure 2.33.


Figure 2.33 Schematic outline of the phases of CAM, showing net CO2 exchange and malic acid and carbohydrate (glucan) metabolism in Kalanchoë daigremontiana leaves. (Diagram courtesy B. Osmond)

The four phases of CAM metabolism are:

  • Phase I - acidification in the dark (PEPC active and stomata open)
  • Phase II - a transitional phase with  stomata open and both carboxylases active
  • Phase III - deacidification (PEPC inhibited, Rubisco active and stomata closed)
  • Phase IV - C3 photosynthesis (stomata open, Rubisco active and PEPC inhibited)

Within these four phases, the distinctive underlying biochemistry of CAM involves the up-regulation of cytoplasmic PEPC activity during phase I in the dark. Up-regulation is catalysed by PEPC kinase which phosphorylates PEPC making it less sensitive to inhibition by malic acid as it accumulates in the vacuole. Towards night’s end, CO2 fixation by PEPC declines as its carbohydrate substrates are exhausted (Figure 2.33). PEPC kinase is degraded during phase II and PEPC becomes increasingly sensitive to malic acid (declining Ki malate; Figure 2.34). It remains inhibited throughout phases III and IV.

CAM also involves the up-regulation of Rubisco in the light by ATP-dependent Rubisco activase as photosynthetic electron transport (ETR) increases in phase II and is maintained throughout phases III and IV (Figure 2.34).


Figure 2.34 Regulation of enzyme activities in deacidification phases of CAM as photosynthetic ETR increases in the light. (Diagram based on K. Maxwell et al. Plant Physiol 121: 849-856, 1999)

Partitioning of carbohydrate metabolism occurs in the light to retain chloroplast starch or vacuolar sugars as substrates for the next nocturnal acidification phase (phase I) while diverting sugars for phloem transport and growth. In pineapple, for example, degradation of starch in the chloroplast may provide the substrate for PEPC despite the large diel turnover of soluble sugars. The complexity of this “conflict of interest” (Borland and Dodd 2002) in carbohydrate metabolism varies between CAM plants with different deacidification pathways.

Sophisticated interactions occur between metabolite transporters in membrane systems of the vacuole, mitochondria and chloroplasts. Many of these are unique to CAM but of 48 such transporters required to support known variations of CAM (including Clusia spp. that also accumulate citric acid) up until 2005, only 8 had been demonstrated in at least one species (Holtum et al. 2005).

When studied under constant conditions, many of the above distinctive biochemical processes in CAM exhibit circadian rhythms. The extent to which endogenous oscillators orchestrate the clearly interacting biochemical, physiological and environmental controls seems likely to remain a challenging area of research. - Physiological attributes distinctive to CAM


Figure 2.35 Diel expansion growth of Opuntia oricola cladodes during drought in the desert biome of the Biosphere 2 Laboratory in Oracle Arizona USA measured by time-lapse photography. Heterogeneity of growth rate throughout the day is colour coded as red = 2.0% to blue = 0.5% per hour (Diagram by B. Osmond based on Gouws et al. Funct Plant Biol 32: 421-428, 2005)

Compared to the photosynthetic biochemistry and physiology in leaves of C3 and C4 plants, the 6% of taxa estimated to exhibit CAM (in at least 35 families and >400 genera) express it with staggering variety (Winter et al. 2015). That is, the distinctive biochemical attributes of CAM outlined above, derived from a handful of research-compliant leafy model species, are but the tip of an iceberg of what really qualifies as a CAM plant (Borland et al. 2011).

The following summary of some distinctive physiological attributes of CAM underscores this conundrum:

Biochemical and physiological determinants of stable isotopic composition of plants with CAM. Fixation of CO2 by PEPC and Rubisco in vitro show clearly different discriminations against the heavier, naturally occurring, non-radioactive (stable) 13C isotope of carbon when expressed as a \(\delta\)13C value. Thus total carbon in C4 plants reflects a small discrimination against 13C resulting in \(\delta\)13C values of about –12.5 ‰, with more negative values in C3 plants (about –27 ‰). It is therefore not surprising that CAM plants tend to fall between these values depending on the balance between total carbon assimilated by PEPC in phase I and that added by Rubisco in phase IV. Partial closure of stomata adds a diffusional discrimination to the biochemical discrimination associated with Rubisco, so \(\delta\)13C values in C3 plants (and CAM plants) become less negative under water stress (Griffiths et al. 2007). Recently it has been suggested that unequivocal identification of CAM can be assigned on the basis of net nocturnal CO2 assimilation, acidification and \(\delta\)13C values less negative than -20 ‰. If some dark CO2 uptake and net acidification is detectable, but \(\delta\)13C is more negative than -20 ‰, these plants would be designated as C3-CAM species, indicating that CAM is present but the contribution of the CAM pathway to net 24h carbon gain is small in comparison to the contribution of daytime CO2 uptake (Winter et al. 2015).

  1. Stomatal opening in the dark is a fundamental physiological feature of CAM. Dark CO2 fixation lowers internal [CO2] and promotes stomatal opening. Stomata retain their responsiveness to external CO2 in phases I and IV, opening further when external CO2 is reduced. They do not respond to low external CO2 during phase III, when closure occurs in response to high internal [CO2] from deacidification of malic acid but do seem to sense the completion of deacidification itself.
  2. CAM is essentially a single cell phenomenon and succulence (low surface to volume ratio) is a feature of many but not all CAM plants. Succulence makes at least two important contributions to the physiology of CAM: large vacuoles for malic acid storage in mesophyll cells and tight packing of cells with small intercellular spaces. The latter means that CO2 diffusion internally is largely confined to wet cell walls and is thus 3 to 4 orders of magnitude slower than in the gas phase, potentially mitigating CO2 fixation by Rubisco in phase IV.  It remains to be seen whether Clusia rosea may have resolved this trade-off by anatomical/physiological differentiation. Enlarged, tightly-packed PEPC-enriched upper palisade cells have a potential for nocturnal CO2 fixation and acidification whereas lower spongy mesophyll cells exhibit predominantly C3 metabolism (Zambrano et al. 2014).  
  3. Whereas leaf expansion growth of C3 plants in the hot dry desert usually occurs at night, leaves and cladodes of some CAM plants grow in the light during phase III (Figure 2.35). This is not surprising because this phase coincides with reliable availability of CHO, maximum temperature and highest cytoplasmic acidity required for growth (Gouws et al. 2005). On the other hand Mesembryanthemum crystallinum (a facultative CAM plant) shows maximum growth in the dark. - Ecological attributes distinctive to CAM

Appreciation of the remarkable plasticity in expression of CAM in response to development and environment has greatly advanced the understanding of the ecological attributes of this photosynthetic pathway. One attempt to bring order to the complexity of CAM expression was the designation of constitutive and facultative categories of CAM. Assignation of these terms requires close monitoring of CAM attributes throughout the life-cycle in response to stochastic environmental events such as water availability.

Constitutive CAM seems securely associated with many massive succulents such as the emblematic columnar cacti in the desert South Western USA but current research also shows it to be prevalent in tropical orchids and bromeliads. Young photosynthetic tissues of constitutive CAM plants are often C3 but CAM is always present at maturity, when the magnitude of the phases of CAM nevertheless remains responsive to stress, light and temperature.

Facultative CAM describes the reversible up-regulation of CAM in response to drought or salinity stress in plants that are otherwise C3 or display low-level CAM. In these, the up-regulated CAM activity is reversible, being reduced (or lost) on removal of stress (Winter et al. 2008; Winter and Holtum 2014). Facultative CAM has been demonstrated in annual plants of seasonally arid environments (e.g. Australia’s desert Calandrinia; Winter and Holtum 2011) as well as in tropical trees of the genus Clusia. The diel patterns of growth in facultative CAM Clusia minor shift from night when in C3 mode to phase III when in CAM mode (Walter et al. 2008).

Slow incremental increase in biomass through vegetative reproduction is a feature of CAM-dominated ecosystems. In CAM plants such as Agave and Opuntia, essentially all of the aboveground tissues are photosynthetic, and this partially compensates for lower rates of CO2 fixation on an area basis. With the noted exception of pineapple and Agave, few CAM species are domesticated, but others have been proposed as potential low-input biofuel crops on land not arable for C3 and C4 crops (Borland et al. 2011; Yang et al. 2015). There is no doubt that communities dominated by CAM plants can attain high biomass (Figure 2.36) and nowhere was this more obvious than during the invasion of 25 million hectares of central eastern Australia during 1846-1926 by prickly pear (Opuntia stricta).


Figure 2.36 High biomass of invasive Opuntia stricta (left) at the Chinchilla, Queensland site C27 before and after release of Cactoblastis cactorum larvae (photos courtesy Queensland Department of Lands 1980) and (right) heritage listed Cactoblastis memorial hall at nearby Boonarga, perhaps the only memorial building to commemorate the achievements of an insect. (Photograph courtesy B. Osmond)

After 2 decades of heroic chemical warfare (hand to hand stabbing or spraying with 10-15% arsenic pentoxide in sulphuric acid at close quarters) failed to restrain the “incubus”, an estimated 1.5 billion tonnes of prickly pear succumbed to trillions of larvae of the diminutive moth Cactoblastis cactorum in about 3 years. Eighty years later, this biological control system remains functionally intact thanks to the remarkably sensitive CO2 detectors in the mouth parts of the female moth that identifies the CAM plant as a target for oviposition by its distinctive nocturnal, inwardly directed CO2 flux in Australian ecosystems (Osmond et al. 2008). The hunger of emerging larvae does the rest. Nevertheless, around 27 species of opuntioid cacti remain naturalised across a range of soil types and climatic zones in the mainland states of Australia. It is not known why Cactoblastis cactorum does not attack a broad range of other feral opuntioid cacti.  In South Australia, with an estimated 1,000,000 ha affected (Chinnock 2015), a control management plan has been enacted (Harvey 2009).

Until the 1980s it was thought that the Australian native flora possessed few CAM plants and that prickly pear had occupied an “empty niche”. Field and laboratory studies by Klaus Winter using acid titration and \(\delta\)13C values demonstrated CAM in the desert succulent Sarcostemma australe as well as drought and salinity induced facultative CAM in Dysphyma clavellatum and Carpobrotus aequilaterus. He also found CAM in rainforest epiphytes and in a diminutive succulent Calandrinia polyandra from sandy and rocky desert habitats. The latter was recently shown to display one of the most overt transitions from C3 photosynthesis when well watered to classic CAM when drought stressed (Figure 2.37).  The impression persists that the warm, dry continent of Australia is either CAM-depauperate or ripe for CAM exploration. On the basis of the size of the Australian flora one might predict around 1,300 Australian CAM species, only about 80 have been documented.


Figure 2.37 Small is beautiful. Diminutive Calandrinia polyandra seems set to become the Arabidopsis analog for CAM research. The diel CO2 exchange patterns on the right were obtained from the same plant well watered for 33 days (top) and 46 days after water had been withheld (bottom). (Photograph courtesy K. Winter; data from K. Winter and J.A.M. Holtum, Funct Plant Biol 38: 576-582, 2011) - Speculations on the origins of CAM


Figure 2.38 A clump of Isoetes andicola symbolizes the extraordinary functional biodiversity of CAM. (Photograph courtesy J. E. Keeley)

From the above it will be clear that tracking the origins of CAM autotrophy in plants will involve no mean feat (“a laudable triumph of great difficulty”). From a holistic perspective, CAM tests the extremities of most aspects of the physiology and ecology of terrestrial plants, as testified in a comprehensive recent collection of reviews and research papers over-viewed by Sage (2014).  With all the emphasis on water-use efficiency in arid environments as a dominant selective pressure for CAM it is often overlooked (and perhaps ironic) that this pathway today is found in aquatic plants, including the fern-ally Isoetes. The origins of Isoetes, though not the present-day taxa themselves, are Triassic, some 100 x106 years before the commonly imagined emergence of CAM in terrestrial plants (Keeley 2014).

The selective pressure for nocturnal storage of CO2 in malic acid by CAM in terrestrial plants may well be closure of stomata to conserve water loss in a dry atmosphere in daylight. In aquatic plants the selective pressure may be the slow diffusion of CO2 in water and its depletion from solution by photosynthesis. In between we have Isoetes andicola from the high Andes of Peru, in which non-functional stoma-like epidermal structures seem literally stitched up (Figure 2.38).  

Clumps of I. andicola are embedded in mounds of peat, with the tips of leaf-like structures forming small rosettes (~5 cm diam.) on the surface. These contain chloroplast-containing cells surrounding large air spaces that evidently maintain gas-phase connections through their large “drinking straw-like” roots to high [CO2] in the peat (~ 4%). The green tips can’t fix CO2 from the air, but when 14CO2 is supplied to the peat it is fixed within leaves into malic acid in the dark and metabolized to photosynthetic products in the light. Understanding how the habit of I. andicola manages to “CAMpeat” in these high elevation ecosytems remains a challenge.          

Any comment on the ecological and evolutionary attributes of CAM must acknowledge the often remarkable features of sexual reproduction, especially in orchids so highly prized in horticultural and gardening contexts. It is also fair to observe that this popular zoocentric fascination pays little or no heed to the distinctive autotrophic metabolism that supports such ecological exotica. One must concede that nocturnal pollination of saguaro by bats is not very amenable to experiment, so plant ecophysiologists might be excused their preference to focus on the resilience of these organisms in the face of environmental stress. 

However, few would deny that the cameo performances of night-blooming cacti are an astonishingly beautiful reward for the nightshift efforts that have unraveled our current understanding of CAM (Figure 2.39).


Figure 2.39 A standing ovation for several centuries of CAM research? The spectacular night blooming cactus Epiphyllum oxypetalum. (Photograph courtesy B. Osmond)

The chapter is dedicated to the memory of Thomas Neales (1929-2010) who pioneered Australian research on CAM with Opuntia stricta in the Botany Department, University of Melbourne. - References for CAM

Balsamo RA, Uribe EG (1988) Plasmalemma- and tonoplast-ATPase activity in mesophyll protoplasts, vacuoles and microsomes of the Crassulacean-acid-metabolism plant Kalanchoe daigremontiana. Planta 173: 190-196

Borland AM, Dodd AN (2002) Carbohydrate partitioning in crassulacean acid metabolism plants. Funct Plant Biol 29: 707-716

Borland AM, Zambrano VAB, Ceusters J et al. (2011) The photosynthetic plasticity of crassulacean acid metabolism: an evolutionary innovation for sustainable productivity in a changing world. New Phytol 191: 619-633

Chinnock RJ (2015) Feral opuntioid cacti in Australia. The State Herbarium of South Australia, Adelaide.

Gouws LM, Osmond CB, Schurr U et al. (2005) Distinctive diel growth cycles in leaves and cladodes of CAM plants: differences from C3 plants and putative interactions with substrate availability, turgor and cytoplasmic pH. Funct Plant Biol 32: 421-428

Griffiths H, Cousins AB, Badger MR et al. (2007) Discrimination in the dark: resolving the interplay between metabolic and physical constraints to phosphoenolpyruvate carboxylase during the crassulacean acid metabolism cycle. Plant Physiol 143: 1055-1067

Harvey A (2009) Draft state opuntioid cacti management plan. Government of South Australia, Adelaide.

Holtum JAM, Smith JAC, Neuhaus HE (2005) Intracellular transport and pathways of carbon flow in plants with crassulacean acid metabolism. Funct Plant Biol 32: 429-439

Keeley JE (2014) Aquatic CAM photosynthesis: a brief history of its discovery. Aquatic Bot 118: 38-44

Lüttge U (2002) CO2 concentrating: consequences in crassulacean acid metabolism. J Exp Bot 53: 2131-2142

Osmond CB, Neales T, Stange G (2008) Curiosity and context revisited: crassulacean acid metabolism in the Anthropocene. J Exp Bot 59: 1489-1502

Sage R (2014) Photosynthetic efficiency and carbon concentration in terrestrial plants: the C4 and CAM solutions. J Exp Bot 65: 3323-3325

von Caemmerer S, Griffiths H (2009) Stomatal responses to CO2 during a diel crassulacean acid metabolism cycle in Kalanchoe daigremontiana and Kalanchoe pinnata. Plant Cell Environ 32: 567-576

Walter A, Christ MM, Rascher U et al. (2008) Diel leaf growth cycles in Clusia spp. are related to changes between C3 photosynthesis and crassulacean acid metabolism during development and water stress. Plant Cell Environ 31: 484-491

Winter K, Holtum JAM (2011) Induction and reversal of crassulacean acid metabolism in Calandrinia polyandra: effects of soil moisture and nutrients. Funct Plant Biol 38: 576-582

Winter K, Holtum JAM (2014) Facultative crassulacean metabolism (CAM) plants: powerful tools for unravelling the functional elements of CAM photosynthesis. J Exp Bot 65: 3425-3441

Winter K, Aranda J, Holtum JAM (2005) Carbon isotope composition and water-use efficiency in plants with crassulacean acid metabolism. Funct Plant Biol 32: 381-388

Winter K, Garcia M, Holtum JAM (2008) On the nature of facultative and constitutive CAM: environmental and developmental control of CAM expression during early growth of Clusia, Kalanchoë and Opuntia. J Exp Bot 59: 1829-1840

Winter K, Holtum JAM, Smith JAC (2015) Crassulacean acid metabolism: a continuous or discrete trait? New Phytol 208:  73-78

Yang X, Cushman JC, Borland AM et al. (2015) A roadmap for research on crassulacean acid metabolism (CAM) to enhance sustainable food and bioenergy production in a hotter, drier world. New Phytol 207: 491-504

 Zambrano VAB, Lawson T, Olomos E et al. (2014) Leaf anatomical traits which accommodate the facultative engagement of crassulacean metabolism in tropical trees of the genus Clusia. J Exp Bot 65: 3513-3523

2.2.9 - Submerged aquatic macrophytes (SAM)

Vascular plants often inhabit regions subject to tidal submergence while others carry out their entire life cycle under water. Examples of common submerged aquatic macrophytes are pond weeds and seagrasses. Once again, an evolutionary selective pressure for these plants has been the availability of CO2. Low levels of dissolved CO2 are common in both inland and marine waters, particularly at more alkaline pH. In more productive inland lakes, CO2 content can vary enormously, requiring considerable flexibility in the actual mode of carbon acquisition. At high pH, HCO3 becomes the more abundant form of inorganic carbon, whereas dissolved CO2 will predominate at low pH (Section 18.2). Consequently, when SAM plants evolved from their C3 progenitors on land, there was some adaptive advantage in devices for CO2 accumulation because CO2 rather than HCO3 is substrate for Rubisco. The nature of this ‘CO2 pump’ and the energetics of carbon assimilation are not fully characterised in SAM plants but considerable CO2 concentrations do build up within leaves, enhancing assimilation and suppressing photorespiration.

In summary:

Regardless of photosynthetic mode, and despite catalytic limitations, Rubisco is ubiquitous and remains pivotal to carbon gain in our biosphere. As a corollary, carbon loss via photorespiration is an equally universal feature of C3 leaves, and the evolution of devices that overcome such losses have conferred significant adaptive advantages to C4, CAM and SAM plants.


2.10 References for photosynthesis

Bolton JK, Brown RH (1980) Photosynthesis of grass species differing in carbon dioxide fixation pathways. V. Response of Panicum maximum, Panicum milioides, and tall fescue (Festuca arundinacea) to nitrogen nutrition. Plant Physiol 66: 97-100

Brown DA (1980). Photosynthesis of grass species differing in carbon dioxide fixation pathways. IV. Analysis of reduced oxygen response in Panicum milioides and Panicum schenckii. Plant Physiol 65: 346-349

Christin PA, Samaritani E, Petitpierre B et al. (2009) Evolutionary insights on C4 photosynthetic subtypes in grasses from genomics and phylogenetics. Genom Biol Evol 1: 221–230

Edwards GE, Franceschi VR, Voznesenskaya EV (2004) Single-cell C4 photosynthesis versus the dual-cell (Kranz) paradigm. Annu Rev Plant Biol 55: 173–196

Ehleringer JR, Cerling TE, Helliker BR (1997) C4 photosynthesis, atmospheric CO2, and climate. Oecologia 112: 285-299

Ghannoum O, von Caemmerer S, Conroy JP (2002) The effect of drought on plant water use efficiency of nine NAD–ME and nine NADP–ME Australian C4 grasses. Funct Plant Biol 29: 1337-1348

Ghannoum O, Evans JR, Chow WS et al. (2005) Faster rubisco is the key to superior nitrogen-use efficiency in NADP-malic enzyme relative to NAD-malic enzyme C4 grasses. Plant Physiology 137: 638-650

Hatch MD (1987) C4 photosynthesis: a unique blend of modified biochemistry, anatomy and ultrastructure. Biochim Biophys Acta 895: 81–106

Hatch MD, Kagawa T, Craig S (1975) Subdivision of C4-pathway species based on differing C4 acid decarboxylating systems and ultrastructural features. Aust J Plant Physiol 2: 111-128.

Hattersley PW, Watson L, Osmond CB (1977) In situ immunofluorescent labelling of ribulose-1,5-bisphosphate carboxylase in leaves of C3 and C4 plants. Aust J Plant Physiol 4: 523-539

Hattersley PW (1992) In ‘Desertified Grasslands: their Biology and Management’ (ed. Chapman GP) pp 181-212. Academic Press: London

Ku MSB, Wu JR, Dai ZY et al. (1991) Photosynthetic and photorespiratory characteristics of Flaveria species. Plant Physiol 96: 518-528

Pinto H, Tissue DT, Ghannoum O (2011) Panicum milioides (C3-C4) does not have improved water or nitrogen economies relative to C3 and C4 congeners exposed to industrial-age climate change. J Exp Bot 62: 3223-3234

Portis AR Jr, Salvucci ME (2002). The discovery of Rubisco activase – yet another story of serendipity. Photosyn Res 73: 257–264

Portis AR Jr, Li C, Wang D, Salvucci ME (2008) Regulation of Rubisco activase and its interaction with Rubisco. J Exp Bot 59: 1597-1604

Rawsthorne S (1992) C3-C4 intermediate photosynthesis - Linking physiology to gene expression. Plant J 2: 267-274

Sage RF (2004). The evolution of C4 photosynthesis. New Phytol 161: 341-370

Sage RF, Christin PA, Edwards EJ (2011) The C4 plant lineages of planet Earth. J Exp Bot 62, 3155-3169

Spreitzer RJ, Salvucci ME (2002) Rubisco: structure, regulatory interactions, and possibilities for a better enzyme. Annu Rev Plant Biol 53: 449-475

Vogan PJ, Sage RF (2011) Water-use efficiency and nitrogen-use efficiency of C3-C4 intermediate species of Flaveria. Plant Cell Environ 34: 1415–1430.

von Caemmerer S, Furbank RT (2003) The C4 pathway: an efficient CO2 pump. Photosyn Res 77: 191–207

Voznesenskaya EV, Franceschi VR, Kiirats O et al. (2002) Proof of C4 photosynthesis without Kranz anatomy in Bienertia cycloptera. Plant J 31: 649–662

Voznesenskaya EV, Edwards GE, Kiirats O et al. (2003) Development of biochemical specialization and organelle partitioning in the single celled C4 system in leaves of Borszczowia aralocaspica. Amer J Bot 90: 1669-1680

2.3 - Photorespiration


Figure 2.1a (also shown in the first section of this chapter). Diagram of the chloroplast showing Rubisco’s carboxylation reaction of RuBP with CO2 to produce two 3-PGA molecules, and the oxygenation reaction to produce one 3-PGA and one P-glycolate molecule.

Rubisco is a bifunctional enzyme capable of reacting either CO2 or O2 to RuBP in the active sites. Although Rubisco’s affinity for CO2 is an order of magnitude higher than that for O2, the high O2 concentration (20%) relative to CO2 (0.004%) in the Earth’s atmosphere leads to a ratio of 3:1 of carboxylation:oxygenation in C3 plants exposed to air. The carboxylation reaction yields two molecules of 3-PGA while the oxygenation of RuBP yields one molecule of 3-PGA and one molecule of phosphoglycolate (P-glycolate), as shown in this figure.

The 3-carbon compound 3-PGA enters the Calvin cycle, but the 2-carbon compound 2-phosphoglycolate is a dead-end metabolite. Consequently, plants have evolved a series of metabolic reactions, termed photorespiration, aimed at salvaging some of the carbon stored in 2-phosphogylcolate and evolving the rest as CO2. This process of CO2 evolution is different from mitochondrial respiration which is described in Section 2.4. The historical evidence for photorespiration is presented below.

2.3.1 - History of photorespiration research

(a) Historical evidence for photorespiration

The first line of evidence for photorespiration came from the different compensation points of C3 and C4 plants. When air is recirculated over an illuminated leaf in a closed system, photosynthesis will reduce CO2 concentration to a low level where fixation of CO2 by photosynthesis is just offset by release from respiration. For many C3 plants this compensation point is around 50 µL L–1 but is markedly affected by oxygen, photon irradiance and leaf temperature (Tregunna et al. 1966; Zelitch 1966). In low concentrations (1–2% O2) the CO2 compensation point of C3 plants is near zero. Significantly, early researchers in this area had already noted that some tropical grass species appeared to have a compensation point at or close to zero CO2, even in normal air (20% O2). This was first reported for corn (Zea mays) (Meidner 1962) and raised a very perplexing question as to whether these species even respired in light. However, we now know that C4 photosynthesis is responsible for the low evolution of CO2 (Section 2.2) and that C4 plants have a CO2 concentrating mechanism that forestalls photorespiration, resulting in a CO2 compensation point close to zero.


Figure 2.12. Photosynthesising leaves show a post-illumination burst of CO2 which varies in strength according to surrounding O2 concentration. This positive response to O2 was found at 105 µmol quanta m-2 s-1 and is functionally linked to O2 effects on the CO2 compensation point as measured under steady-state conditions. (Based on Krotkov 1963)

A second line of evidence for leaf respiration in light was provided by a transient increase in release of CO2 when leaves are transferred from light to dark. This ‘post-illumination CO2 burst’ was studied extensively during the early 1960s by Gleb Krotkov and colleagues at Queens University (Kingston, Ontario). The intensity of this burst increased with the photon irradiance during the preceding period of photosynthesis. Understandably, the Queens group regarded this post-illumination burst as a ‘remnant’ of respiratory processes in light, and coined the term ‘photorespiration’. A functional link with the CO2 compensation point was inferred, because the burst was abolished in low O2 (Figure 2.12). A competitive inhibition by O2 on CO2 assimilation was suspected and was subsequently proved to be particularly relevant in defining Rubisco’s properties. Nevertheless, for many years a biochemical explanation for interaction between these two gases remained elusive.

(b) Biochemistry

Significant progress came when Ludwig and Krotkov designed an open gas exchange system in which 14CO2 was used to separate the fixing (photosynthetic) and evolving (respiratory) fluxes of CO2 for an illuminated leaf (Ludwig and Canvin 1971). Results using this steady-rate labelling technique were particularly revealing and provided the first direct evidence that respiratory processes in light were qualitatively different from those in darkness. They were able to show that CO2 evolved during normal high rates of photosynthesis by an attached sunflower leaf was derived from currently fixed carbon. The specific activity of evolved CO2 (ratio of 14C to 12C) was essentially the same as that of the CO2 being fixed, indicating that photorespiratory substrates were closely related to the initial products of fixation. Ludwig and Krotkov concluded that 14CO2 supplied to a photosynthesising leaf was being re-evolved within 28–45 s! Furthermore the rate of CO2 evolution in light was as much as three times the rate in darkness, and while early fixed products of photosynthesis (intermediates of the PCR cycle) were respired in light, this was not the case in darkness.

The radiolabelling method of Ludwig and Krotkov had, for the first time, provided measurements of what could be regarded as a true estimate of light-driven respiration which was not complicated by transient effects (as the post-illumination burst had been), or by changes in CO2 concentration (as was the case for measurements in closed gas exchange systems or in CO2-free air) or by difficulties associated with detached organs. Ludwig and Canvin (1971) subsequently concluded that processes underlying photorespiration re-evolved 25% of the CO2 which was being fixed concurrently by photosynthesis. Such a rate of CO2 loss was not a trivial process so a biochemical basis for its operation had to be established, and particularly when photorespiration seemed to be quite different from known mechanisms of dark (mitochondrial) respiration.

The search for the substrates of ‘photorespiration’ occupied many laboratories worldwide for many years. Much work centred on synthesis and oxidation of the two-carbon acid glycolate because as early as 1920 Warburg had reported that CO2 fixation by illuminated Chlorella was inhibited by O2, and under these conditions the alga excreted massive amounts of glycolic acid (Warburg and Krippahl 1960). Numerous reports on the nature of the 14C-labelled products of photosynthesis showed that glycolate was a prominent early-labelled product. A very wide variety of research with algae and leaves of many higher plants established two significant features of glycolate synthesis: formation was enhanced in either low CO2 or high O2. Both of these features had been predicted from Ludwig’s physiological gas exchange work and eventually proved a key to understanding the biochemistry of photorespiration.

(c) Source of glycolate production

The photosynthetic carbon reduction (PCR) cycle for CO2 fixation (Section 2.1) involves an initial carboxylation of ribulose-1,5-bisphosphate (RuBP) to form 3-PGA, but makes no provision for glycolate synthesis. However, Wang and Waygood (1962) had described the ‘glycolate pathway’, namely a series of reactions in which glycolate is oxidised to glyoxylate and aminated, first to form glycine and subsequently the three-carbon amino acid serine. The intracellular location of this pathway in leaves was established in a series of elegant studies by Tolbert and his colleagues who also established that leaf microbodies (peroxisomes) were responsible for glycolate oxidation and the synthesis of glycine. Kisaki and Tolbert (1969) suggested that the yield of CO2 from the condensation of two molecules of glycine to form serine could account for the CO2 evolved in photorespiration. This idea was incorporated in later formulations of the pathway.

What remained elusive was the source of photosynthetically produced glycolate. Many studies had suggested that the sugar bisphosphates of the PCR cycle could yield a two-carbon fragment which, on the basis of short-term 14CO2 fixation, would have its two carbon atoms uniformly labelled (if the two carbons were to be derived directly from 3-PGA this would not be the case as PGA was asymmetrically labelled in the carboxyl group). The mechanism was likened to the release of the active ‘glycolaldehyde’ transferred in the thiamine pyrophosphate (TPP)-linked transketolase-catalysed reactions of the PCR cycle. In some cases significant glycolate synthesis from the sugar bisphosphate intermediates of the cycle were demonstrated in vitro; however, the rates were typically too low to constitute a viable mechanism for glycolate synthesis in vivo.

A more dynamic approach to carbon fixation was needed to resolve this impasse. In particular, the biochemical fate of early products would have to be traced, and using a development of the open gas exchange system at Queens, Atkins et al. (1971) supplied 14CO2 in pulse–chase experiments to sunflower leaf tissue under conditions in which photorespiration was operating at high rates (21% O2) or in which it was absent (1% O2). A series of kinetic experiments showed that synthesis of 14C-glycine and 14C-serine was inhibited in low O2 and that the 14C precursor for their synthesis was derived from sugar bisphosphates of the PCR cycle, especially RuBP. Indeed RuBP was the obvious source of glycine carbon atoms and the kinetics of glycine turnover closely matched those of RuBP. As these authors concluded, ‘we can no longer view this (glycolate) pathway as an adjunct to the Calvin cycle but must incorporate it completely into the carbon fixation scheme for photosynthesis’ (Atkins et al. 1971).

The question was finally and most elegantly resolved by Ogren and Bowes (1971) who demonstrated that the carboxylating enzyme of the PCR cycle, RuBP carboxylase, was both an oxygenase and a carboxylase! During normal photosynthesis in air, this enzyme thus catalysed formation of both P-glycolate (the precursor of glycolate) and 3-PGA from the oxygenation of RuBP as well as two molecules of PGA from carboxylation of RuBP. In effect, CO2 and O2 compete with each other for the same active sites for this oxygenation/carboxylation of RuBP, at last providing a biochemical mechanism which had confused and perplexed photosynthesis researchers since the 1920s. This primary carboxylating enzyme of the PCR cycle, which had hitherto rejoiced in a variety of names (carboxydismutase, fraction 1 protein, RuDP carboxylase and RuBP carboxylase), was renamed Rubisco (ribulose-1,5-bisphosphate carboxylase/oxygenase) to reflect its dual activity.

A scheme for the PCR cycle and photosynthetic carbon oxidation (PCO) pathways then represents the synthesis of almost 70 years of research effort, and integrates the metabolism of P-glycolate with the PCR cycle. This is shown in the following section. 


2.3.2 - Photorespiration needs three organelles


Figure 2.13. The photorespiratory carbon oxidation (PCO) cycle involves movement of metabolites between chloroplasts, peroxisomes and mitochondria. (Original drawing courtesy lan Woodrow)

A scheme for photorespiration involving the photosynthetic carbon reduction (PCR) cycle and the photosynthetic carbon oxidation (PCO) pathways represents the synthesis of almost 70 years of research. It integrates the metabolism of P-glycolate with the PCR cycle. This scheme is shown in Figure 2.13.

Specialised reactions within three classes of organelles in leaf cells are required, namely chloroplasts, peroxisomes (originally called microbodies) and mitochondria. Their close proximity in leaf cells (Figure 2.14 below) plus specific membrane transporters facilitate the exchange of metabolites.

Within choroplasts, oxygenase activity by Rubisco results in formation of phosphoglycolate which then enters a PCO cycle, and is responsible for loss of some of the CO2 just fixed in photosynthesis.

Within peroxisomes, O2 is consumed in converting glycolate to glyoxylate, and aminated to form glycine.

Within mitochondria, CO2 is released during conversion of glycine to serine. Subsequently, serine is recovered by peroxisomes where it is further metabolised, re-entering the PCR cycle of chloroplasts as glycerate. About 75% of carbon skeletons channelled into photorespiration are eventually recovered as carbohydrate.

Transport of glycerate and glycolate across the inner membrane of chloroplasts may involve separate translocators as shown in Figure 2.13, or it may involve a single translocator that exchanges two glycolate molecules for one molecule of glycerate. Transport of metabolites across the peroxisomal membrane most likely occurs through unspecific channel proteins, similar to those in the outer membranes of mitochondria and chloroplasts. These outer membranes are not included in this diagram. Mitochondria take up two molecules of glycine and release one molecule of serine. A specific translocator most probably mediates the exchange of these amino acids.

Not only does the photosynthetic oxidation pathway consume O2 and release CO2 but ammonia is also produced by mitochondria during synthesis of serine from glycine (Figure 2.13). This ammonia would be extremely toxic if it were not metabolised by either cytosolic or chloroplastic glutamine synthetase. A very effective herbicide that blocks glutamine synthetase has been developed (phosphinothricin, also known as glufosinate or Basta), and when it is applied to (or expressed in) actively growing plants they are killed by their photorespiratory ammonia release.

Fig 2.11.jpg

Figure 2.14. A transmission electron micrograph showing close juxtaposition of chloroplast (C), mitochondrion (M) and peroxisome (P) in a mesophyll cell of an immature leaf of bean (Phaseolus vulgaris). This group of organelles is held within a granular cytoplasmic matrix adjacent to a cell wall (CW) and includes a partial view of a small vacuole (V). Scale bar = 1 µm (Electron micrograph courtesy Stuart Craig and Celia Miller)

In summary, participation of photorespiration in leaf gas exchange, and thus dry matter accumulation by plants, reflects kinetic properties of Rubisco, and in particular a relatively high affinity for CO2 (Km = 12 µM) compared with a much lower affinity for O2 (Km = 250 µM). That contrast in affinity is, however, somewhat offset by the relative abundance of the two gases at catalytic sites of the enzyme where the ratio of O2:CO2 partial pressures approaches 1000:1! Thus, CO2 assimilation always prevails over CO2 loss in photorespiration. 

2.3.3 - C<sub>4</sub> plants and unicellular algae avoid photorespiration

At around the same time as the nature of photorespiration was becoming clearer Hatch and Slack (1966) demonstrated that in tropical grasses (initially sugar cane) the first-formed products of photosynthetic CO2 fixation were the four-carbon acids oxalacetate, malate and aspartate, rather than the 3-PGA formed in the PCR cycle. Furthermore, the carboxylation reaction involved PEP carboxylase and carbon was subsequently transferred to PCR cycle intermediates. As noted earlier (Section 2.2) C4 plants show no apparent CO2 release in light. The explanation lies in their anatomy and multiple carboxylation reactions rather than in the absence of the pathway of photorespiration. Bundle sheath cells are equipped with a CO2-concentrating mechanism that favours carboxylation over oxygenation reactions due to increased partial pressure of CO2, while photorespiratory release of CO2 is further prevented through the activity of PEP carboxylase which refixes any respired CO2 formed from the oxygenase function of Rubisco.

Unicellular green algae also posed a problem for the simple extrapolation of early models for photorespiratory metabolism in C3 leaves. Although organisms such as Chlorella had been used to establish the PCR cycle, and indeed provided much early evidence for effects of O2 on photosynthesis and formation of glycolate in light, they also appeared to lack CO2 evolution in light (Lloyd et al. 1977). In this case the explanation lies in a CO2-concentrating mechanism which effectively increases the internal pool of inorganic carbon (CO2 and HCO3) thereby favouring the carboxylase function of Rubisco over its oxygenase function.

2.3.4 - Does photorespiration represent lost productivity?

Such a substantial loss of carbon concurrent with CO2 fixation raises the question of whether eliminating or minimising photorespiration in C3 plants could enhance their yield, and specifically that of major crop plants such as rice, wheat, grain legumes, oil seeds and trees, none of which are C4 species. Faced with an expanding world human population and an increasing demand for food and animal feed, enhanced agricultural productivity is a global necessity. In its most obvious form a scenario which alters or removes the oxygenase function of Rubisco could achieve such a goal. In an early review of the process of photorespiration in plants, Ogren (1984) noted that ‘the sequence of reactions constituting the photorespiratory pathway in C3 plants appears to be firmly established’ and he went on to suggest that, although reducing the loss of fixed carbon as CO2 in the process may be a valid goal to improve the yield of crop plants, it is not clear whether or not this can be achieved by specific changes to the kinetic and catalytic properties of Rubisco alone.

Photorespiration may be loosely considered as a wasteful process because previously fixed carbon is lost and energy is dissipated. Ideal destinations for photoassimilates include synthetic pathways leading to fixed biomass and respiratory pathways for re-release of fixed energy in a controlled sequence of reactions leading to ATP and NAD(P)H for use in other synthetic events.

However, situations do exist where energy dissipation via photorespiration can be beneficial. For example, photo-oxidative damage can be alleviated in shade-adapted plants that experience strong irradiance if photorespiratory processes are allowed to proceed. Depriving such plants of an external oxygen supply, and hence preventing photosynthetic carbon oxidation, will exacerbate chloroplast lesions due to strong irradiance. Photosynthetic variants which obviate photo-respiratory loss, and most notably C4 plants, integrate structure and function in a way that forestalls photo-oxidative damage and leads to their outstanding performance under warm conditions. Environmental factors that constitute selection pressure for such photosynthetic adaptation are reviewed by Sage et al (2012). 

2.3.5 - References for photorespiration

Atkins CA, Canvin DT, Foch H (1971) Intermediary metabolism of photosynthesis in relation to carbon dioxide evolution in sunflower. In MD Hatch, CB Osmond, RO Slatyer, eds, Photosynthesis and Photorespiration. John Wiley, New York, pp 497-505

Hatch MD, Slack CR (1966) Photosynthesis by sugarcane leaves. A new carboxylation reaction and the pathway of sugar formation. Biochem J 101: 103-111

Kisaki T, Tolbert NE (1969) Glycolate and glyoxylate metabolism by isolated peroxisomes and chloroplasts. Plant Physiol 44: 242-250

Lloyd ND, Canvin DT, Culver DA (1977) Photosynthesis and photorespiration in algae. Plant Physiol 70: 1637-1640

Ludwig LJ, Canvin DT (1971) The rate of photorespiration during photosynthesis and the relationship of the substrate of light respiration to the products of photosynthesis in sunflower leaves. Plant Physiol 48: 712-719

Meidner HA (1962) The minimum intercellular-space CO2 concentration of maize leaves and its influence on stomatal movements. J Exp Bot 13: 284-293

Ogren WL (1984) Photorespiration: pathways, regulation and modification. Annu Rev Plant Physiol 35: 415-442

Ogren WL, Bowes G (1971) Ribulose diphosphate carboxylase regulates soybean photosynthesis. Nature New Biol 230: 159-160

Sage RF, Sage TL, Kocacinar F (2012) Photorespiration and the evolution of C4 photosynthesis. Annu Rev Plant Biol 63: 19-47.

Tregunna EB, Krotkov G, Nelson CD (1966) Effect of oxygen on the rate of photorespiration in detached tobacco leaves. Physiol Plant 19: 723-733

Warburg O, Krippahl G (1960) Glykolsaurebildung in Chlorella. Z Naturforsch 15b: 197-199

Wang D, Waygood ER (1962) Carbon metabolism of 14CO2 labelled amino acids in wheat leaves. I. A pathway of glyoxylate-serine metabolism. Plant Physiol 37: 826-832

Zelitch I (1966) Increased rate of net photosynthetic carbon dioxide uptake caused by the inhibition of glycolate oxidase. Plant Physiol 41: 1623-1631

2.4 - Respiration and energy generation

Nicolas L. Taylor1,2 and A. Harvey Millar1
1ARC Centre of Excellence in Plant Energy Biology and 2School of Chemistry and Biochemisty, The University of Western Australia.

During photosynthesis the carbon assimilated is either retained in the chloroplast as starch or converted to sucrose and directed for export to sites of growth. Starch is degraded by a series of enzymes in the chloroplast, with sucrose degradation mainly occurring in the cytosol and both leading to glycolysis and the oxidative pentose pathway which produce respiratory substrates. These carbon rich compounds are prime sources of respiratory substrates in plants, although other carbohydrates such as fructans and sugar alcohols are also used.

During respiration, metabolites are oxidised and the electrons released are transferred through a series of electron carriers to O2. Water and CO2 are formed and energy is captured as ATP which is harnessed to drive a vast array of cellular reactions.

In comparison to sucrose and starch, the contribution of proteins and lipids as sources of respiratory substrates in most plant tissues is minor; exceptions to this generalisation are the storage tissues of seeds such as castor bean and soybean, in which amino acids and lipids may provide respiratory substrates, and during the processes of senescence in plant tissues where protein and lipid degradation increases.

2.4.1 - Starch and sucrose degradation

Starch is the principal storage carbohydrate in plants and this carbon reserve plays a number of important roles in plants. It is composed of two polymers of glucose, amylose and amylopectin and is stored in the plastid (chloroplast in leaves, amyloplasts in non-photosynthetic tissues) as insoluble, semi-crystalline granules. Starch is accumulated during rapid growth in the day and is almost completely degraded at night to mostly glucose and maltose, which is exported from the chloroplast and metabolised in the cytosol (Figure 2.19). Starch degradation is initiated by the addition of phosphate groups at the C6-position and C3-position of individual glucosyl residues that act to disrupt the packing of the glucans at the granule surface. These phosphate additions are catalysed by two enzymes, glucan water dikinase (GWD) and phosphoglucan water dikinase (PWD) respectively. The hydrolysis of the resulting glucan and phosphoglucan chains is carried out by a suite of enzymes including the phosphoglucan phosphatases (SEX4/LSF2), β-amylases (BAM1/BAM3), debranching enzymes (DBE; ISA3/LDA), α-amylase (AMY3), α-glucan phosphorylase and the disproportionating enzyme 1 (D-enzyme 1; an α-1,4-glucanotransferase).). The resulting maltose and glucose are exported to the cytosol by the glucose transporter (pGlcT) and maltose transporter (MEX1) and glucose-1-phosphate is thought to be exported by a similar but yet unknown mechanism. Once in the cytosol the maltose and glucose are converted to substrates for either sucrose synthesis, glycolysis or the oxidative pentose phosphate pathway by a number of enzymes including the disproportionating enzyme 2 (D-enzyme 2; an α-1,4-glucanotransferase), α-glucan phosphorylase, hexokinase and phosphoglucomutase.


Figure 2.19. Pathways of starch degradation. ATP, adenosine triphosphate; AMP, adenosine monophosphate; Pi, inorganic phosphate; GWN, glucan water dikinase; PWD, phosphoglucan water dikinase; SEX4/LSF2, phosphoglucan phosphatases; ISA3/LDA debranching enzymes (DBEs); BAM1/BAM3, β-amylases; D-Enyzme 1/2, disproportionating enzyme (α-1,4-glucanotransferase); pGlcT, glucose transporter; MEX1, maltose transporter, HK, hexokinase; PGM, phosphoglucomutase; Glu-1-P, Glucose-1-phosphate; Glu-6-P, Glucose-6-phosphate; OPP, Oxidative pentose phosphate pathway. (Original drawing courtesy Nicolas Taylor & Harvey Millar)

Sucrose is the world’s most abundant disaccharide, it is only produced by photosynthetic organisms and serves a role as a transportable carbohydrate and sometimes as a storage compound. The reactions in plant tissues leading to degradation of sucrose to hexose monophosphates are outlined in Figure 2.20.


Figure 2.20. The pathways of sucrose breakdown. SPP, sucrose phosphate phosphatase; S-6-P, sucrose-6-phosphate; SPS, sucrose phosphate synthase; UDP, uridine diphosphate; HK, hexokinase, ATP, adenosine triphosphate; ADP, adenosine diphosphate; Pi, inorganic phosphate; HPI, hexose phosphate isomerase; OPP, Oxidative pentose phosphate pathway. (Original drawing courtesy Nicolas Taylor & Harvey Millar)

The first step is cleavage of the glycosidic bond by either invertase (Equation 2.1) or sucrose synthase (Equation 2.2).

\[\text{Sucrose + H}_2\text{O} \rightarrow \text{D-Glucose + D-Fructose}\tag{2.1}\]

\[\text{Sucrose + UDP} \rightarrow \text{UDP-Glucose + D-Fructose}\tag{2.2}\]

Plant tissues contain distinct invertases located in the vacuole, cell wall (acid invertases) cytosol, mitochondria, nucleus, and cholorplast (neutral/alkaline invertases) which hydrolyse sucrose to glucose and fructose in an irreversible reaction. The invertases are differentially regulated by a number of mechanisms including pH to allow them to function in cell expansion, supply of carbon skeletons and energy metabolism. Multiple isoforms of sucrose synthase are located in the cytosol or cytosolic membranes that catalyse a thermodynamically reversible reaction, but this reaction probably acts only to breakdown sucrose in vivo. Their activity is developmentally regulated and they have functions in the supply of activated glucose for starch and cellulose biosynthesis. While both invertase and sucrose synthase can both breakdown sucrose, research using knockouts of multiple isoforms of both enzymes has shown that sucrose synthase is not required for normal growth in Arabidopsis, whereas invertase is indispensable. However this does not rule out the requirement of sucrose synthase in certain tissues of crop plant tubers, seeds and fruits where it has been shown to be crucial. Glucose and fructose are metabolised further by phosphorylation to the corresponding hexose-6-P by hexokinase. Hexokinase in plant tissues is associated with the outer surface of mitochondria.

2.4.2 - The glycolytic pathway

The glycolytic pathway involves the oxidation of the hexoses and hexose phosphates molecules produced from the breakdown of starch or sucrose to generate ATP, reductants and pyruvate (Figure 2.21).


Figure 2.21 The glycolytic pathway. OPP, Oxidative pentose phosphate pathway; PPi, pyrophosphate, Pi, inorganic phosphate; TPI, triose phosphate isomerase; NAD+, nicotinamide adenine dinucleotide (oxidised); NADH, nicotinamide adenine dinucleotide (reduced); NADP+, nicotinamide adenine dinucleotide phosphate (oxidised); NADPH; nicotinamide adenine dinucleotide phosphate (reduced); ATP, adenosine triphosphate; ADP, adenosine diphosphate;  MDH, malate dehydrogenase; PEPC, phosphoenolpyruvate carboxylase; PK, pyruvate kinase. (Original drawing courtesy Nicolas Taylor & Harvey Millar)

The process of glycolysis occurs within both the cytosol and plastid, with reactions in the different compartments catalysed by separate enzyme isoforms. The first step in the pathway is the phosphorylation of glucose by hexokinase to form glucose-6-phosphate in an ATP consuming reaction. This glucose-6-phosphate is converted to fructose-6-phosphate by glucose phosphate isomerase to form fructose-6-phosphate, which is also the entry point for fructose that can be phosphorylated by hexokinase also forming fructose-6-phosphate. The fructose-6-phosphate is then further phosphorylated to fructose-1,6-bisphosphate by one of two enzymes capable of catalysing this step: an ATP-dependent phosphofructokinase (PFK), which catalyses an irreversible reaction and occurs in the cytosol and plastids, and a pyrophosphate-dependent phosphofructokinase, (PPi-PFK), which occurs only in the cytosol and utilises pyrophosphate (PPi) as the phosphate donor in a reaction that is readily reversible.

Regulation of PFK and PPi-PFK is achieved by a combination of mechanisms, including pH, the concentration of substrates and effector metabolites and changes in subunit association. Phosphoenolpyruvate (PEP) is a potent inhibitor of both of PFK and PPi-PFK, inhibiting at µM concentrations and Pi can activate the cytoplasmic PFK, whereas the plastidic form is slightly inhibited by Pi. A number of other effectors of PFK have been identified including ADP, 3-phosphoglycerate and phosphoglycolate as well as it ability to accept ribonucleoside triphosphates other than ATP as the phosphate donor. PPi-PFK, has a catalytic potential higher than that of PFK and is strongly activated by Fructose-2,6-bisphosphate, but has no effect on PFK.

Fructose-1,6-bisphosphate is cleaved by fructose bisphosphate aldolase to form glyceraldehyde-3-phosphate and dihydroxyacetone phosphate, and these triose phosphates can be interconverted in a reaction catalysed by triose phosphate isomerase. Glyceraldehyde-3-phosphate is oxidised to 1,3-bisphosphoglycerate by a nicotinamide adenine dinucleotide (NAD+)-dependent glyceraldehyde 3-P dehydrogenase. Glyceraldehyde 3-P dehydrogenase is sensitive to inhibition by the reduced pyridine nucleotide cofactor (NADH), which must be reoxidised to maintain the flux through the glycolytic pathway. A phosphate group is then transferred from 1,3-bisphosphoglycerate to ADP forming ATP and 3-phosphogylcerate by phosphoglycerate kinase. In the cytosol a bypass is present that can convert glyceraldehyde-3-phosphate directly to 3-phosphoglycerate without phosphorylation by a non-phosphorylating NADP dependent glyceraldehyde 3-phosphate dehydrogenase. The resulting 3-phosphoglycerate is then converted to phosphenolpyruvate (PEP) by the action of phosphoglycerate mutase and then enolase.  

The end-products of glycolytic reactions in the cytosol are determined by the relative activities of the two enzymes that can utilise PEP as a substrate: pyruvate kinase, which forms pyruvate and a molecule of ATP, and PEP carboxylase, which forms oxaloacetate (Figure 2.21). Both of these reactions are essentially irreversible and there are fine controls that regulate the partitioning of PEP between these reactions. Pyruvate kinase is controlled post translationally by a partial C-terminal truncation which may yield altered regulatory properties and a phosphorylation and ubiquitin conjugation that targets the protein to the 26S proteasome for complete degradation, it is also inhibited by ATP. Whereas PEP carboxylase is inhibited by malate and thus its regulation is independent of cell energy status. The sensitivity of PEP carboxylase to malate is regulated by phosphorylation of a N-terminal serine of the enzyme, with the phosphorylated form less sensitive to malate inhibition. Oxaloacetate is then reduced by malate dehydrogenase to malate which, along with pyruvate, can be taken up into mitochondria and metabolised further in the TCA cycle (see below). The reduction of oxaloacetate in the cytosol could provide a cytosolic mechanism for oxidising NADH formed by glyceraldehyde 3-P dehydrogenase (Figure 2.21).

Another level of regulation of components of glycolysis is their physical location within the plant cell. Under conditions of high respiratory activity, a greater proportion of the cytosolic enzymes of glycolysis are present on the surface of mitochondria. In contrast when respiration is experimentally inhibited, a decrease in the association of glycolytic enzymes with the mitochondria is observed. It is likely that the glycolytic enzymes associate dynamically with mitochondria to support respiration and that this association restricts the use of glycolytic intermediates by competing metabolic pathways.

2.4.3 - The oxidative pentose phosphate pathway

An alternative route for the breakdown of glucose-6-phosphate is provided by the oxidative pentose phosphate pathway (OPP) (Figure 2.22). This pathway functions mainly to generate reductant (i.e. NADPH) for biosynthetic processes including the assimilation of inorganic nitrogen and fatty acid biosynthesis and to maintain redox potential to protect against oxidative stress. In addition, the reversible oxidative section of the pathway is the source of carbon skeletons for the synthesis of a number of compounds. For example ribose-5-phosphate provides the ribosyl moiety of nucleotides and is a precursor for the biosynthesis of purine skeletons and erythrose-4-phosphate, which is the precursor for the biosynthesis of aromatic amino acids by the shikimic acid pathway.



Figure 2.22. The oxidative pentose phosphate pathway. NADPH, nicotinamide adenine dinucleotide phosphate (reduced); R-5-p I, ribose-5-phosphate isomerase; R-5-p Epimerase, ribulose-5-phosphate epimerase. (Original drawing courtesy Nicolas Taylor & Harvey Millar)

The pathway begins with the dehydrogenation of glucose-6-phosphate catalyzed by glucose-6-phosphate dehydrogenase to produce 6-phosphoglucolactone and is the first step of the oxidative phase of the pathway. The 6-phosphoglucolactone is then hydrolysed to 6-phosphogluconate by 6-phosphogluconolactonase and then undergoes oxidative decarboxylation by 6-phosphogluconate dehydrogenase to produce ribulose-5-phosphate in the final step of the oxidative phase. Overall this phase of the pathway produces two molecules of NADPH from the conversion of glucose-6-phosphate to ribulose-5-phosphate. The non-oxidative phase begins with the reaction of ribulose-5-phosphate with either ribulose-5-phosphate isomerase or ribulose-5-phosphate epimerase followed by a series of reactions catalyzed by transaldolase and transketolase. These reactions result in the production of two molecules of fructose-6-phosphate and one glyceraldehyde 3-phosphate. The glyceraldehyde 3-phosphate and fructose-6-phosphate in the oxidative pentose phosphate pathway may be exchanged with enzymes of glycolysis.

As with glycolysis, reactions of the pentose phosphate pathway are catalysed by different isoforms of the enzymes that occur either in the cytosol or in plastids. Although transketolase and transaldolase may be absent from the cytosol of some species, the activity is maintained by phosphate translocator proteins on the plastid inner-envelope membrane that have the capacity to translocate sugar phosphates.

2.4.4 - Mitochondria and organic acid oxidation (the TCA cycle)

Organic acids such as pyruvate and malate produced in the cytosol by processes described above are further oxidised in mitochondria by the tricarboxylic acid (TCA) cycle and subsequent respiratory chain. Energy released by this oxidation is used to synthesise ATP which is then exported to the cytosol for use in biosynthesis and growth.

(a)  Mitochondrial structure


Figure 2.23. Transmission electron micrograph of a parenchyma cell in a floral nectary of broad bean (Vicia faba) showing an abundance of mitochondria, generally circular in profile and varying between about 0.75 and 1.5 µm in diameter. Each mitochondrion is encapsulated by an outer and inner membrane which is in turn infolded to form cristae. Scale bar = 0.5 µm. (Original electron micrograph courtesy Brian Gunning)

Plant mitochondria (Figure 2.23) are typically double-membrane organelles where the inner membrane is invaginated to form folds known as cristae to increase the surface area of the membrane. The outer membrane contains relatively few proteins (<100) and is permeable to most small compounds (< Mr=5 kDa) due to the presence of the pore-forming protein VDAC (voltage dependent anion channel) which is a member of the porin family of ion channels. The inner membrane is the main permeability barrier of the organelle and controls the movement of molecules by means of a series of carrier proteins many of which are members of mitochondrial substrate carrier family (MSCF). The inner membrane also houses the large complexes that carry out oxidative phosphorylation and encloses the soluble matrix which contains the enzymes of the TCA cycle and many other soluble proteins involved in a myriad of mitochondrial functions.

Mitochondria are semi-autonomous organelles with their own DNA and protein synthesis machinery. However, the mitochondrial genome encodes only a small portion of the proteins which make up the mitochondrion; the rest are encoded on nuclear genes and synthesised in the cytosol. These proteins are the transported into the mitochondrion by the protein import machinery and assembled with the mitochondrially synthesised subunits to form the large respiratory complexes. The number of mitochondria per cell varies with tissue type (from a few hundred in mature differentiated tissue to some thousands in specialised cells). Understandably, more active cells, with high energy demands, such as those in growing meristems are generally equipped with larger numbers of mitochondria per unit cell volume, and consequently show faster respiration rates.

(b)  Mitochondrial substrates

Two substrates are produced from glycolytic PEP for oxidation in mitochondria: malate and pyruvate (Figure 2.21). These compounds are thought to be the most abundant mitochondrial substrates in vivo. However, amino acids may also serve as substrates for mitochondrial respiration in some tissues, particularly in seeds rich in stored protein or under conditions of sugar depletion such as extended darkness, shading and senescence. β-oxidation of fatty acids typically does not occur in plant mitochondria, this oxidation is principally carried out in peroxisomes in plants.

(c) Carbon metabolism in mitochondria

Malate and pyruvate enter the mitochondrial matrix across the inner membrane via separate carriers. Malate is then oxidised by either malate dehydrogenase (a separate enzyme isoform from that in the cytosol), which yields oxaloacetate (OAA) and reduced nicotinamide adenine dinucleotide (NADH), or NAD+-linked malic enzyme, which yields pyruvate and NADH and releases CO2 (Figure 2.24). Cytosolic pyruvate carboxylase is an alternative means of providing substrate to mitochondria by combining pyruvate with HCO3 to yield OAA that can then be imported into mitochondria.


Figure 2.24. The tricarboxylic acid cycle. NAD+, nicotinamide adenine dinucleotide (oxidised); NADH, nicotinamide adenine dinucleotide (reduced); ATP, adenosine triphosphate; ADP, adenosine diphosphate; Pi, inorganic phosphate; GTP, guanosine triphosphate; GDP, guanosine diphosphate; Q, qunione; QH2, dihydroquinone. (Original drawing courtesy Nicolas Taylor & Harvey Millar)


Pyruvate formed either from malate and malic enzyme or transported directly from the cytosol is oxidised inside mitochondria by the pyruvate dehydrogenase complex (PDC) to form CO2, acetyl-CoA and NADH. This enzyme, which requires coenzyme A, thiamine pyrophosphate and lipoic acid as cofactors, effectively links the TCA cycle to glycolysis. PDC comprises three enzymes E1 (2-oxo acid dehydrogenase), E2 (acyltransferase) and E3 (lipoamide dehydrogenase). This complex is regulated by phosphorylation of the E1 subunit, lowering PDC activity in the day and increasing PDC activity at night. Pyruvate dehydrogenase is also subject to feedback inhibition from acetyl-CoA and NADH.

(d) Tricarboxylic acid (TCA) cycle

The TCA cycle begins with the condensation of acetyl-CoA and OAA, to form the six-carbon molecule citrate and the release coenzyme A (CoA) (Figure 2.24) in a reaction catalysed by citrate synthase. Aconitase catalyses the next step, converting citrate to isocitrate in a two-step reaction (dehydration/hydration) with cis-aconitate as an intermediate.

NAD-linked isocitrate dehydrogenase then oxidatively decarboxylates isocitrate to form CO2 and 2-oxoglutarate, and reduce NAD+ to NADH. The 2-oxoglutarate formed is also oxidatively decarboxylated to succinyl-CoA in a reaction catalysed by the enzyme 2-oxoglutarate dehydrogenase. This enzyme complex has similarities to pyruvate dehydrogenase and its reaction is analogous to the formation of acetyl-CoA from pyruvate by pyruvate dehydrogenase. The reaction mechanisms are also very similar, with 3 subunit enzymes, but 2-oxoglutarate dehydrogenase is not subject to the phosphorylation control that regulates pyruvate dehydrogenase. Succinyl-CoA synthase then catalyses the conversion of succinyl-CoA to succinate, with the concomitant phosphorylation of ADP to ATP, the only substrate-level phosphorylation step in the mitochondrion. This enzyme in plants differs from its mammalian counterpart in that it is specific for ADP rather than GDP.

Succinate dehydrogenase (SDH, Complex II), which catalyses the oxidation of succinate to fumarate, is the only membrane-bound enzyme of the TCA cycle and is part of the respiratory electron transfer chain (Figure 2.25). SDH is a large complex consisting of four core subunits, as well as number other associated subunits.

Fumarase catalyses the hydration of fumarate to malate followed by malate dehydrogenase that catalyses the final step of the TCA cycle, oxidising malate to OAA and producing NADH. The reaction is freely reversible, although the equilibrium constant strongly favours the reduction of OAA, necessitating rapid turnover of OAA and NADH to maintain this reaction in a forward direction.

Overall, during one turn of the cycle, three carbons of pyruvate are released as CO2, one molecule of ATP is formed directly, and four NADH and one FADH2 are produced. The strong reductants are oxidised in the respiratory chain to reduce O2 and produce ATP. Although most of the TCA cycle enzymes in plant mitochondria are NAD linked, NADP-dependent isoforms of isocitrate and malate dehydrogenases also exist, and these may play a role in a protective reductive cycle in the matrix.

Regulation of carbon flux through the TCA cycle probably occurs via phosphorylation/dephosphorylation of pyruvate dehydrogenase, which will depend in turn on mitochondrial energy status and feedback inhibition of various enzymes by NADH and acetyl-CoA. The rate of cycle turnover thus depends on the rate of electron flow through the respiratory chain (to reoxidise NADH) and the utilisation of ATP in the cell to provide ADP for substrate level and oxidative phoshorylation. TCA cycle turnover will also depend on the rate of substrate provision by reactions in chloroplasts and cytosol. A number of studies of TCA cycle mutants have demonstrated the wide impact these enzymes have not simply on TCA cycle function but as steps for the delivery of organic acids for other processes in plant cells such a photosynthetic performance, plant biomass, root growth, photorespiration, nitrogen assimilation, amino acid metabolism, and stomatal function.

2.4.5 - Electron transport chain

The respiratory electron transfer chain (ETC) of mitochondria consists of a series of large membrane-bound protein complexes (Complexes I, II, III, IV) which together with a small lipid ubiquinone (UQ) and the small protein cytochrome c catalyse the transfer of electrons from NADH and succinate to O2, forming H2O (Figure 2.25). Electron flow from NADH and succinate to oxygen is coupled to proton translocation out of the matrix to the intermembrane space which establishes a proton electrochemical gradient (DµH+) across the inner membrane that is used to drive phosphorylation of ADP to form ATP by the F1FO ATP synthase (Complex V, Figure 2.25).


Figure 2.25. The electron transport chain of plant mitochondria. Numbers (I-V) identify the large respiratory complexes located on the inner mitochondrial membrane. Complex I, NADH:ubiquinone oxidoreductase; complex II, succinate dehydrogenase; complex III, ubiquinone-cytochrome c oxidoreductase; complex IV, cytochrome c oxidase; complex V, ATP synthase. Letters identify alternative pathway enzymes, eN, external NAD(P)H dehydrogenase; iN, internal NAD(P)H dehydrogenase; A, alternative oxidase and UQ, ubiquinone/ubiquinole pool; c, cytochrome c; U, uncoupling protein. NADH, nicotinamide adenine dinucleotide (reduced); NAD+ nicotinamide adenine dinucleotide (oxidised); ATP, adenosine triphosphate; ADP, adenosine diphosphate; Pi, inorganic phosphate.  Unbroken arrows indicate pathways of electron flow; broken arrows indicate proton translocation sites. (Original drawing courtesy Nicolas Taylor & Harvey Millar)

(a) Complex I (CI)


NADH-UQ oxidoreductase, is responsible for the oxidation of matrix NADH and reduction of ubiquinone (UQ) in the inner mitochondrial membrane (Figure 2.25). In plants it is a large multi-subunit complex composed of 49 subunits, up to ten of which are synthesised in the mitochondrion whilst the others are imported from the cytosol. One of the subunits, a 50 kDa protein, contains flavin mononucleotide as a cofactor and is the dehydrogenase which oxidises NADH and passes electrons to iron-sulphur containing subunits of the complex, and eventually to ubiquinone. The passage of electrons through the complex is accompanied by H+ translocation across the membrane. Complex I is inhibited specifically by the flavonoid rotenone and its analogues. The NADH-binding site is exposed to the matrix and the complex oxidises NADH produced by the TCA cycle and other NAD-linked enzymes (Figure 2.25). Studies of mutations of CI subunits have shown that plants can survive without CI due to the activity of alternative NAD(P)H dehydrogenases (see below). Such mutants have a variety of interesting phenotypes including viral infection tolerance, prolonged hydration under water-deficient conditions and altered organic and amino acid concentrations. Using a series of Arabidopsis CI subunit knockout mutants, a number membrane arm subcomplexes (of 200, 400, 450 and 650 kDa) have been identified using BN-PAGE and antibodies. It is proposed that at least some of these subcomplexes may be assembly intermediates during CI formation, and these are seen to accumulate when specific subunits are absent.

(b) Complex II (CII)

Succinate dehydrogenase, is an enzyme of both the respiratory ETC and the TCA cycle (see above) (Figure 2.25). It is composed of four core subunits: a flavoprotein (SDHI), an iron-sulphur subunit (SDH2) and two membrane anchor subunits (SDH3 and SDH4) in most organisms. In plants, the purification of the complex has revealed the common core subunits, but also additional proteins of unknown function that co-migrate with the complex. All SDH subunits are encoded in the nuclear genome in Arabidopsis. SDH contains FeS and flavin centres which participate in electron transfer from succinate to ubiquinone. Unlike complex I, complex II does not pump H+ and succinate oxidation is therefore linked to the synthesis of less ATP per O2 reduced (see below). Malonate, an analogue of succinate, is a strong competitive inhibitor of succinate dehydrogenase. Knockout mutants of the SDH1 gene have been shown to be embryo lethal, but knockdown of SDH1 and SDH2 leads a array of phenotypes including altered stomatal aperture, mitochondrial ROS production and nitrogen use efficiency. SDHAF1 and SDHAF2 assist in CII assembly in plants and knockdown of the SDHAF2 homolog lowers SDH assembly and reduces root growth.

(c) Complex III (CIII)

Ubiquinone-cytochrome c oxidoreductase or the cytochrome b/c1 complex as it is sometimes known, contains 10 subunits including a number of bifunctional core proteins. These proteins act both in CIII function and as a matrix processing peptidase, removing targeting presequences from imported matrix synthesized proteins (Figure 2.25). A single subunit of this complex, cytochrome b, is encoded by the plant mitochondrial genome, whilst the others are encoded by the nuclear genome. It contains two b-type cytochromes, b566 and b562, cytochrome c1, an iron-sulpher protein named the Rieske iron–sulphur protein and several other polypeptides. Electron flow from ubiquinol to cytochrome c is accompanied by the translocation H+ across the membrane, via the so-called Q cycle. Various inhibitors of complex III have been discovered, with antimycin A and myxothiazol most widely used in research. The assembly of CIII is modular and includes an early core subcomplex, a late core subcomplex and the final dimeric CIII. Approximately 13 assembly factors implicated in aiding one or more of the different stages of CIII assembly in yeast, however little is known about CIII assembly or functional assembly factors in plants.

(d) Complex IV (CIV)

Cytochrome c oxidase is the final step of electron transfer of the classical ETC. As the name implies, cytochrome c oxidase accepts electrons from cytochrome c and transfers them to O2 which is reduced to form H2O. Purification of CIV in plants has identified a complex containing 14 protein subunits (Figure 2.25). Eight of these proteins are homologous to known CIV subunits from other organisms, together with a further six proteins that are probably plant specific. Two cytochrome haem centres, a and a3, and two copper atoms make up its redox active components and like complex I, cytochrome oxidase is a proton pump. Cytochrome c oxidase is sensitive to a number of inhibitors, the best known of which are carbon monoxide and cyanide. Plants, however, show resistance to both carbon monoxide and cyanide because they are equipped with an alternative oxidase (see below). Studies of human and yeast CIV has shown an assembly pathway that involves the sequential incorporation of CIV subunits, initiated by subunit 1 and assisted by over 40 assembly factors. Research investigating the plant homolog of the yeast assembly factor COX19 has found it is capable of complementing the yeast cox19 null mutant. This suggests it might play a role in the biogenesis of plant cytochrome c oxidase or in the replacement of damaged forms of the enzyme. However, our knowledge of the assembly of CIV in plants is still incomplete.

(e) Ubiquinone and Cytochrome c

These large multi-subunit complexes (I, II, III, IV) of the respiratory ETC chain are embedded in the inner mitochondrial membrane by virtue of their hydrophobic subunits, and interact with one another via two smaller molecules: ubiquinone and cytochrome c. The lipid-soluble ubiquinone also known as coenzyme Q10 is a small mobile electron carrier which moves rapidly along and across the membrane, and participates in H+ transport across the membrane via the Q-cycle as well as shuttling electrons from complexes I and II to complex III. Cytochrome c is a small haem-containing protein located on the outer surface of the inner membrane, which shuttles electrons between complexes III and IV. In this respect, the respiratory chain is similar in layout to the photosynthetic electron transport chain: three large complexes which communicate by a quinone and a small mobile protein (Cyt c or plastocyanin). However, orientation of components in the membrane is inverted and the net reaction catalysed is opposite to that in chloroplasts (Figure 1.12).

2.4.6 - ATP synthesis (oxidative phosphorylation)

When electrons are transferred from NADH to O2, a large release of redox energy enables ATP formation in complex V of the respiratory chain (Figure 2.24). Energy release associated with electron transport is conserved by H+ translocation across the membrane to form a proton electrochemical gradient (ΔµH+) that has both an electrical membrane potential (Δψ) and a pH component (ΔµH+ = Δψ + ΔpH). This is known as the chemiosmotic theory and was originally proposed by Peter Mitchell in 1960s. In plant mitochondria, ΔµH+ exists mainly as a Δψ of ~150–200 mV, with a pH gradient (ΔpH) of ~0.2–0.5 units. ATP synthesis occurs as H+ move from a compartment of high potential (the intermembrane space) to one of low potential (the mitochondrial matrix) through the ATP synthase complex. Oxidation of NADH via the cytochrome pathway has three associated H+ translocation sites and is linked to synthesis of up to three ATP molecules for each molecule of NADH oxidized. By contrast, both succinate and alternative NADH oxidation by the rotenone-insensitive NADH dehydrogenases (see below) are linked to the synthesis of only two ATP molecules per NADH or succinate, as these events are associated with only two H+ pumping sites.


Figure 2.26. Stylised O2 electrode recording of respiring plant mitochondria illustrating respiratory control. Oxygen consumption is measured as a function of time. The isolated mitochondria are depleted of substrates and are therefore dependent on added substrates and effectors. Addition of ADP (Pi is in the reaction medium) allows oxidative phosphorylation to proceed, dissipating some of the electrochemical gradient (ΔµH+) and thereby stimulating electron transport; the enhanced rate of O2 uptake is called State 3. When all the ADP is phosphorylated, electron transport slows to what is known as State 4. Addition of more ADP stimulates O2 uptake further, but addition of oligomycin, which blocks the ATP synthase, lowers O2 uptake to the State 4 rate. The addition of an uncoupler (protonophore) fully dissipates the electrochemical gradient (ΔµH+) and stimulates O2 uptake; no ATP synthesis occurs in the presence of the uncoupler. When the O2 concentration falls to zero, respiration ceases (Original drawing courtesy David Day)


(a) Complex V (CV)

ATP synthase is a membrane-bound F1F0 type H+-ATP synthase that harnesses the ΔµH+ generated by the ETC to produce ATP. It is composed of a hydrophobic F0 component which channels protons through the inner mitochondrial membrane and also anchors the complex to the membrane and a hydrophilic F1 component which catalyses ATP formation and protrudes into the matrix. The core subunits of the enzyme are highly conserved in both prokaryotic and eukaryotic organisms. In plants, the majority of mitochondrial F1 subunits are encoded in the nucleus and translated in the cytosol before being imported into the mitochondria (including  α, β, γ and ε subunits), while most of the F0 subunits are encoded in the plant mitochondrial genome and translated in the mitochondrial matrix (including a, b, c and A6L subunits).  The reaction mechanism of the ATP synthase is known as the three-site alternating binding site mechanism. According to this model, F1 has three nucleotide-binding sites which can exist in three configurations: one with loosely bound nucleotides, one with tightly bound nucleotides and the third in a nucleotide-free state. H+ movement through F0 results in rotation of F1, causing a conformational change during which the site with loosely bound ADP and Pi is converted to one which binds them tightly in a hydrophobic pocket in which ATP synthesis occurs. Further H+ movement then causes another rotation of F1 and the ATP binding site is exposed and releases the nucleotide. In the meantime, the other nucleotide-binding sites are undergoing similar changes, with ADP and Pi being bound and converted to ATP. Thus H+ translocation drives the three sites through three different configurations and the main expenditure of energy is in the induction of a conformational change that releases tightly bound ATP, rather than in ATP synthesis itself. The F0 complex also contains a protein known as the oligomycin-sensitivity-conferring protein (OSCP) because it binds the antibiotic oligomycin that prevents H+ translocation through F0 and inhibits ATP synthesis. Therefore, adding oligomycin to mitochondria oxidising a substrate in the presence of ADP restricts O2 uptake. Generally knockouts of ATP synthase core subunits are lethal in plants, however inducible knockdowns have enabled investigations into the tissue-specific phenotypes incurred by slowing the rates of mitochondrial ADP:ATP cycling at a number of different developmental stages. It has been proposed that the assembly of plant CV comprises of three steps, the first being the formation of a rapidly turned over F1 subcomplex in the matrix, followed by an intermediate stage where F1 associates with the inner membrane and still turns over at a fast rate, and then finally a union of F1 with FO to form functional CV. A number of assembly factors (Atp10, Atp11, Atp12, Atp22, Atp23 and Fmc1) have been discovered for yeast ATP synthase, however, a detailed study of the presence and conservation of CV assembly factors in plants has not been undertaken.

(b) Respiratory control

Electron transport through the respiratory chain, and therefore rate of O2 uptake, is controlled by availability of ADP and Pi, a phenomenon described as ‘respiratory control’. In the absence of ADP or Pi, the proton pore of ATP synthase is blocked and a ΔµH+ builds up to a point where it restricts further H+ translocation across the inner membrane. Since electron transport is functionally linked to H+ translocation, this elevated ΔµH+ will also restrict O2 consumption. That outcome is easily seen with isolated mitochondria (Figure 2.26) where O2 uptake is stimulated by adding ADP (‘State 3’ respiration). When all of the added ADP has been consumed, O2 uptake decreases again (‘State 4’). In steady state, the rate of electron flow is determined by the rate of flow of H+ back across the membrane: when ADP and Pi are available the backflow is rapid and occurs via ATP synthase; in the absence of these compounds, backflow is by slow diffusion through the membrane.

The ratio of State 3 to State 4 (the respiratory control ratio) is thus an indication of coupling between ADP phosphorylation and electron transport. Larger values represent tighter coupling. The proton leak can be dramatically stimulated by some compounds which act as protonophores or proton channels; these compounds collapse the ΔµH+ and increase O2 uptake up to the State 3 rate (Figure 2.26). However, no ATP is formed and these compounds are called uncouplers because they uncouple the linked processes of electron transport and phosphorylation.

2.4.7 - Alternative electron transport pathways

Plant mitochondria have a respiratory chain which is more complicated than that of animals and contains alternative NADH dehydrogenases, alternative oxidases which catalyse cyanide-insensitive O2 consumption and uncoupling proteins that acts to dissipate the ΔµH+. The alternative NADH dehydrogenases and alternative oxidase do not translocate protons and therefore are not linked to ATP synthesis; they are often referred to as the non-phosphorylating bypasses of the plant respiratory chain. These pathways were initially identified in plant mitochondria as they are able to continue to respire in the presence of the CIV inhibitor, cyanide and the CI inhibitor, rotenone and by their ability to exhibit natively uncoupled respiration in the absence of an ADP source.

(a) Alternative oxidase

Cyanide-insensitive respiration is catalysed by the alternative oxidase (AOX). This alternative terminal oxidase is a diiron quinol oxidase that branches from the classical respiratory chain at UQ and reduces oxygen to water without an associated proton translocation. The oxidase exists in mitochondria as a dimer which can be inactivated by covalent linkage via disulphide bonds. The reduced enzyme is stimulated allosterically by pyruvate and some other 2-oxo acids (such as glyoxylate), which interact directly with the oxidase. The exact role of AOX continues to be debated but it appears to play an antioxidant role in plant mitochondria. Research has shown it is actively induced by oxidative stress and the different genes for the oxidase have been shown to be both development- and tissue-specific. Knockout of AOX leads to reactive oxygen species and anthocyanin accumulation in the leaves exposed to a combination of high light and drought stress. AOX can be inhibited by hydroxamic acids such as n-propylgallate (nPG) and salicyl hydroxamic acid (SHAM).

(b) Alternative NAD(P)H dehydrogenases

Alternative NAD(P)H dehydrogenases have been shown to be present on both sides of the inner mitochondrial membrane. These type II NAD(P)H dehydrogenases oxidise external or cytosolic and matrix NADH and NADPH and are insensitive to the classical CI inhibitor rotenone. As with AOX, these enzymes do not translocate protons and therefore are not linked to ATP synthesis. The Arabidopsis genome contains seven genes encoding NAD(P)H dehydrogenases, although it appears that some of these isoforms are present in multiple subcellular compartments in addition to mitochondria.

(c) Uncoupling proteins

Uncoupling proteins (UCPs) are members of the mitochondrial carrier family of proteins. They act to dissipate the ΔµH+ built up the ETC by transporting H+ back across the inner membrane uncoupling proton and electron transport. The reactive oxygen species superoxide activates UCPs and this suggests a possible mechanism for the engagement of this enzyme in vivo. Analysis of knockouts of UCP (AtUCP1) showed that its absence led to localized oxidative stress but did not impair the ability of the plant to withstand a wide range of abiotic stresses. However, knockout of UCP1 did limit the photorespiration rate of plants and led to a reduction in photosynthetic carbon assimilation. This suggests that the main role of UCP1 in leaves is to maintain the redox poise of the mitochondrial ETC to facilitate photosynthesis.

2.4.8 - Energetics of respiration

(a)  Efficiency

Respiration represents a substantial loss of carbon from a plant, and under adverse conditions can be as high as two-thirds of the carbon fixed daily in photosynthesis. Both the rate and the efficiency of respiration will therefore affect plant growth significantly. The overall process of respiration results in the release of a substantial amount of energy which may be harnessed for metabolic work. In theory, the energy released from the complete oxidation of one molecule of glucose to CO2 and H2O in respiratory reactions leads to the synthesis of 36 molecules of ATP. However, in plants, because there are alternative routes for respiration, this yield can be greatly reduced. Mechanisms for regulating respiration rates in whole plants remain unclear. Convention has it that the rate of respiration is matched to the energy demands of the cell through feed-back regulation of glycolysis and electron transport by cytosolic ATP/ADP. However, since plants have non-phosphorylating bypasses in their respiratory chain that are insensitive to ATP levels, and since PEP carboxylase and PFP might be involved in sucrose degradation, the situation in vivo is not so simple. For example, the rotenone-insensitive alternative NADH dehydrogenases requires high concentrations of NADH in the matrix before it can operate and seems to be active only when substrate is plentiful and electron flow through complex I is restricted by lack of ADP. Alternative oxidase activity also depends on carbon and ADP availability and its flux is very dependent on the degree of environmental stress of the plant. In other words, non-phosphorylating pathways act as carbon or reductant ‘overflows’ of the main respiratory pathway and will only be active in vivo when sugar levels are high and the glycolytic flux rapid, when the cytochrome chain is inhibited, or when the bypasses have been induced significantly during stress. In glycolysis, the interaction between environmental signals and key regulatory enzymes, as well as the role of PFP and its activator fructose-2,6-P2, will be important.

(b)  Allocation of respiratory energy to process physiology

One way of viewing respiratory cost for plant growth and survival is by subdividing measured respiration into two components associated with (1) growth and (2) maintenance. This distinction is somewhat arbitrary, and these categories of process physiology must not be regarded as discrete sets of biochemical events. Such energy-dependent processes are all interconnected because ATP represents a universal energy currency for both, while a common pool of substrates is drawn upon in sustaining production of that ATP. Nevertheless, cells do vary in their respiratory efficiency, while genotype × environment interactions are also evident in both generation and utilisation of products from oxidative metabolism. The benefit of a high respiration rate is that more ATP is produced, which provides vital energy for growth of new tissue and defence processes, such as antioxidant activation, metabolite transport or production of resistant protein isoforms. However, the cost of high respiration rates is that carbon is expended on respiration instead of being allocated to synthesis of new tissue, therefore limiting growth capacity. Variation in respiration rate has implications for growth and resource use efficiency in plants during drought, temperature and salinity responses of plants.

2.4.9 - Further Reading

Araujo WL, Nunes-Nesi A, Nikoloski Z, Sweetlove LJ, Fernie AR (2012) Metabolic control and regulation of the tricarboxylic acid cycle in photosynthetic and heterotrophic plant tissues. Plant Cell Environ 35: 1-21

Atkin OK, Macherel D (2009) The crucial role of plant mitochondria in orchestrating drought tolerance. Ann Bot 103: 581-597

Atkin OK, Tjoelker MG (2003) Thermal acclimation and the dynamic response of plant respiration to temperature. Trends Plant Sci 8: 343-351

Jacoby RP, Li L, Huang S, Pong Lee C, Millar AH, Taylor NL (2012) Mitochondrial composition, function and stress response in plants. Journal of Integrative Plant Biology 54: 887-906

Jacoby RP, Taylor NL, Millar AH (2011) The role of mitochondrial respiration in salinity tolerance. Trends Plant Sci 16: 614-623

Kruger NJ, von Schaewen A (2003) The oxidative pentose phosphate pathway: structure and organisation. Curr Opin Plant Biol 6: 236-246

Mannella CA (2008) Structural diversity of mitochondria functional implications. Mitochondria and Oxidative Stress in Neurodegenerative Disorders 1147: 171-179

Millar AH, Whelan J, Soole KL, Day DA (2011) Organization and regulation of mitochondrial respiration in plants. Annual Review of Plant Biology 62: 79-104

Moller IM (2001) Plant mitochondria and oxidative stress: electron transport, NADPH turnover, and metabolism of reactive oxygen species. Annu Rev Plant Physiol Plant Mol Biol 52: 561-591

Patrick JW, Botha FC, Birch RG (2013) Metabolic engineering of sugars and simple sugar derivatives in plants. Plant Biotechnol J 11: 142-156

Plaxton WC (1996) The organization and regulation of plant glycolysis. Annu Rev Plant Physiol Plant Mol Biol 47: 185-214

Stitt M (2013) Progress in understanding and engineering primary plant metabolism. Curr Opin Biotechnol 24: 229-238

Stitt M, Zeeman SC (2012) Starch turnover: pathways, regulation and role in growth. Curr Opin Plant Biol 15: 282-292

Streb S, Zeeman SC (2012) Starch metabolism in Arabidopsis. Arabidopsis Book 10: e0160

Sweetlove LJ, Beard KF, Nunes-Nesi A, Fernie AR, Ratcliffe RG (2010) Not just a circle: flux modes in the plant TCA cycle. Trends Plant Sci 15: 462-470

Sweetlove LJ, Fernie AR (2013) The spatial organization of metabolism within the plant cell. Annu Rev Plant Biol 64: 723-746

Taylor NL, Day DA, Millar AH (2004) Targets of stress-induced oxidative damage in plant mitochondria and their impact on cell carbon/nitrogen metabolism. Journal of Experimental Botany 55: 1-10

Tcherkez G, Mahe A, Hodges M (2011) (12)C/(13)C fractionations in plant primary metabolism. Trends Plant Sci 16: 499-506

Zeeman SC, Kossmann J, Smith AM (2010) Starch: its metabolism, evolution, and biotechnological modification in plants. Annu Rev Plant Biol 61: 209-234

Chapter 3 - Water movement in plants


Karri (Eucalyptus diversicolor) forest in Pemberton, Western Australia. Karri may reach the height of 80 m, and is the second highest hardwood tree in the world (Photograph courtesy A. Munns)

Chapter editors: Brendan Choat and Rana Munns

Contributing Authors: B Choat1, R Munns2,3,4, M McCully2, JB Passioura2, SD Tyerman4,5, H Bramley6 and M Canny*

1Hawkesbury Institute of the Environment, University of Western Sydney; 2CSIRO Agriculture, Canberra; 3School of Plant Biology, University of Western Australia; 4ARC Centre of Excellence in Plant Energy Biology;  5School of Agriculture, Food and Wine, University of Adelaide; 6Facutly of Agriculture and Environment, University of Sydney; *Martin Canny passed away in 2013

Evolutionary changes were necessary for plants to inhabit land. Aquatic plants obtain all their resources from the surrounding water, whereas terrestrial plants are nourished from the soil and the atmosphere. Roots growing into soil absorb water and nutrients, while leaves, supported by a stem superstructure in the aerial environment, intercept sunlight and CO2 for photosynthesis. This division of labour results in assimilatory organs of land plants being nutritionally inter-dependent; roots depend on a supply of photoassimilates from leaves, while shoots (leaves, stems, flowers and fruits) depend on roots to supply water and mineral nutrients. Long-distance transport is therefore a special property of land plants. In extreme cases, sap must move up to 100 m vertically and overcome gravity to rise to tree tops.

This chapter explains the mechanism by which water can rise to the top of a tall tree, and the cellular processes essential for plant cells to maintain turgor.


3.1 - Plant water relations


Figure 3.1 Surface view of cleared whole mount of a wheat leaf showing large and small parallel veins (mauve) with transverse veins connecting them. Lines of stomates (shown by the orange colour of the guard cells) lie along the flanks of these veins. Water evaporates from the wet walls of mesophyll cells below the stomates, drawing water from the veins. Distance between veins is 0.15 mm; scale bar is 100 µm. (Photograph courtesy M. McCully)

Water is often the most limiting resource determining the growth and survival of plants. This can be seen in both the yield of crop species and the productivity of natural ecosystems with respect to water availability.

The natural distribution of plants over the earth’s land surface is determined chiefly by water: by rainfall (\( R \)) and by evaporative demand (potential evapotranspiration, \( PE \)) which depends on temperature and humidity. This leads to such diverse vegetation groups as the lush vegetation of tropical rainforests, the shrubby vegetation of Mediterranean climates, or stands of tall trees in temperate forests. Climates can be classified according to the Thornthwaite Index: \( (R-PE)/PE \).

Agriculture also depends on rainfall. Crop yield is water-limited in most regions in the world, and agriculture must be supplemented with irrigation if the rainfall is too low. Horticultural crops are usually irrigated.

Plants require large amounts of water just to satisfy the requirements of transpiration: a large tree may transpire hundreds of litres of water in a day. Water evaporates from leaves through stomates, which are pores whose aperture is controlled by two guard cells. Plants must keep their stomates open in order to take up CO2 as the substrate for photosynthesis (Chapter 2). In the process, water is lost from the moist internal surfaces of the leaf through the stomatal pores (Figure 3.1). Water loss also has a benefit in maintaining the leaf temperature through evaporative cooling.

The ratio of water lost to CO2 taken up is around 300:1 in most land plants, meaning that plants must transpire large quantities of water on a daily basis in order to take up sufficient CO2 for normal development.

In this section we will examine plant water relations and the variables that plant physiologists use to describe the status and movement of water in plants, soil and the atmosphere.

One of the challenging aspects of understanding plant water relations is the range of pressures from positive to negative that occur within different tissues and cells. Positive pressures (turgor) occur in all living cells and depend on the semipermeable nature of the plasma membrane and the elastic nature of the cell walls. Negative pressures (tensions) occur in dead cells and depend on the cohesive strength of water coupled with the strength of heavily lignified cell walls to resist deformation. These play an important role in water transport through the xylem.

3.1.1 - The power of turgor pressure


Figure 3.3 Wilted squash plants demonstrating loss of cell turgor. (Photograph courtesy of Home and Garden Information Center, University of Maryland Extension).

Well-watered plants are turgid, and their leaves and stems are upright and firm, even without woody tissue to support them. If water is lost from leaves through the stomates at a faster rate than it is resupplied by roots, then plants wilt (Figure 3.3)

Well-watered plants are turgid because their cells are distended by large internal hydrostatic pressures (Figure 3.4a). This internal hydrostatic pressure (also called turgor pressure) is typically 0.5 MPa or more. Lack of water causes cells to shrink until the pressure inside equals that of the atmosphere (zero), and the cells thus have zero turgor (Figure 3.4b). The initial shrinkage while turgor drops from 0.5 to zero MPa is determined by the properties of the cell wall: cell walls are slightly elastic, and the relation between volume change and turgor pressure depends on the “elastic modulus” of the wall. This involves little change in whole cell volume for a drop in turgor pressure to zero. However, further water loss causes the wall to shrink and deform inwards, and the whole cell volume decreases markedly.


Figure 3.4 Turgid leaf cell and flaccid cell (zero turgor). (a) In the turgid cell in a well-watered plant, the cell is distended by a large internal hydrostatic pressure, usually 0.5 MPa - 1 MPa. (b) In the flaccid cell of a wilted plant, the cell wall is rather dry, and water has been lost to the atmosphere until the pressure inside is that of the atmosphere, zero.

The turgor pressure of a fully turgid cell may even exceed 1 MPa, about five times the pressure in a car tyre, and ten times the pressure in the atmosphere. In a physically unconstrained cell, the turgor pressure is borne by the cell wall, which develops a large tension within it. But in cells that are physically constrained, such as those of a tree root whose growth becomes hampered by the presence of a slab of concrete, the tension in the cell walls is relieved and the pressure is applied directly to the constraint (Figure 3.5).


Figure 3.5 Roots lifting slab of concrete. (Photograph courtesy L. Atmore, Daily Bruin, UC Davis)

It is easy to see how a constrained tree root could eventually lift a slab of concrete: 1 MPa applied over 100 cm2 is equivalent to a weight of one tonne. Pressure is force/area, and 1 MPa is approximately equal to 10 kg weight per cm2.

A definition of all these terms is summarised at the end of this section (Section 3.1.7).

3.1.2 - Osmotic pressure and water potential

How is it that plant cells can have such large turgor pressures? The essential reason is that the cells contain large concentrations of solutes. These solutes attract water into the cells through a process known as osmosis, which involves water flowing in through semipermeable membranes that prevent the passage of solutes but not of water. The inflow of water swells the cells until a hydrostatic pressure is reached at which no more water will flow in. In cells bathed in fresh water, such as algal cells in a pond, this equilibrium hydrostatic pressure is known as the osmotic pressure (\( \pi \)) of the cell contents, and is commonly about 500 kPa or 0.5 MPa.

This osmotic pressure can be measured directly with an osmometer, or it can be calculated from the solute concentration in the cell (\( C \)) from the van‘t Hoff relation:

\[ \pi = RTC \tag{1} \]

where \( R \) is the gas constant, \( T \) is the absolute temperature (in degrees Kelvin) and \( C \) is the solute concentration in Osmoles L-1. At 25 ºC, \( RT \) equals 2.5 litre-MPa per mole, and \( \pi \) is in units of MPa. Hence a concentration of 200 mOsmoles L-1 has an osmotic pressure of 0.5 MPa.

However, land plants are different from algae in a pond. Their leaves are in air, and the water in their cell walls, unlike the water in a pond, is not free. It has a negative hydrostatic pressure (discussed further in the next section). Thus, for a given osmotic pressure (\( \pi \)) within a cell, the hydrostatic pressure, \( P \), will be lower than if the cell were bathed in free water. This difference is known as the water potential (\( \psi \)) of the cell. It is zero in an algal cell in fresh water, but it is always negative in land plants. Its value is the difference between \( P \) and \( \pi \), that is:

\[ \psi=P-\pi \tag{2} \]

An alternative notation for equation (2) used commonly by plant physiologists is:

\[ \psi_w = \psi_p + \psi_s \tag{3} \]

In this case, \( \psi_w \) is the total water potential, \( \psi_s \) is the solute potential and \( \psi_p \) is the pressure potential. Thus \( \psi_s \) is equal, but opposite in sign, to \( \pi \).

The notion of water potential can be applied to any sample of water, whether inside a cell, in the cell wall, in xylem vessels, or in the soil. Water will flow from a sample with a high water potential to one with a low water potential provided the samples are at the same temperature and provided that no solutes move with the water. Water potential thus defined is always zero or negative, for by convention it is zero in pure water at atmospheric pressure.

3.1.3 - Positive and negative hydrostatic pressures

Positive values of hydrostatic pressure occur in the living cells of plants, in the symplast, and as explained above are induced by high solute concentrations and the resultant osmotic pressure. However, large negative values are common in the apoplast of plants and the soil they are growing in. These negative values arise because of capillary effects - the attraction between water and hydrophilic surfaces at an air/water interface, the effects of which can be seen in the way that water wicks into a dry dishcloth. This attraction reduces the pressure in the water, and does so more intensely the narrower are the water-filled pores. It accounts for how cell walls, which have very narrow pores, can remain hydrated despite very low water potentials in the tissue they are part of. For a geometrically simple cylindrical pore, the relation between the induced pressure and the radius of the pore can be derived as follows:

Take a glass capillary tube with a radius \( r \) (m) and place it vertically with one end immersed in water. Water will rise in the tube against the gravitational force until an equilibrium is reached at which the weight of the water in the tube is balanced by the force of attraction between the water and the glass. A full, hemispherical, meniscus will have now developed, i.e. one with a radius of curvature equal to that of the tube (Figure 3.6).


Figure 3.6 A fully-developed meniscus in a cylindrical tube showing the equality between the upward pull of surface tension and the downward pull of the suction in the water from which the relation \( \Delta P = 2\gamma/r \) can be derived.

The meniscus is curved because it is supporting the weight of the water - much as a trampoline sags when several people are sitting on it. There is a difference in pressure, \( \Delta P \) (Pa), across the meniscus, with the pressure in the water being less than that of the air. The downward acting force (N) on the meniscus is the difference in pressure multiplied by the cross-sectional area of the tube, i.e \(\pi r^2 \Delta P\). The upward acting force is equal to the perimeter of contact between water and glass (\( 2 \pi r \)) multiplied by the surface tension, \( \gamma \) (N m-1), of water, namely \( 2 \pi r \gamma \) (provided the glass is perfectly hydrophilic, when the contact angle between the glass and the water is zero, otherwise this expression has to be multiplied by the cosine of the angle of contact). Thus, because these forces are equal at equilibrium, we have \(\pi r^2 \Delta P = 2 \pi r \gamma \), whence

\[ \Delta P = 2\gamma/r \tag{4}\]

The surface tension of water is 0.075 N m-1 at about 20°C, so \( \Delta P \) (Pa) equals 0.15 divided by the radius \( r \) (m):

\[ \Delta P = 0.15/r \tag{5}\]

Thus a fully-developed meniscus in a cylindrical pore of radius 0.15 mm would have a pressure drop across it of 1.0 MPa. The pressure, \( P \), in the water would therefore be -1.0 MPa if referenced to normal atmospheric pressure, or -0.9 MPa absolute pressure (given that standard atmospheric is approximately 100 kPa).  

This argument applies not only to cylindrical pores. It is the curvature of the meniscus that determines the pressure drop, and this curvature is uniform over a meniscus occupying a pore of any arbitrary shape. It is such capillary action that generates the low pressures (large suctions) in the cell walls of leaves that induce the long-distance transport of water from the soil through a plant to the sites of evaporation. The pores in cell walls are especially small (diameters of the order of 15 nm), and are therefore able to develop very large suctions, as they do in severely water-stressed plants. Such pores can hold water against a suction of 10 MPa. (Table 3.1)

In plants, other water-filled pores vary in size from large xylem vessels with diameters of 100 mm or more down to a few mm, so for them to remain water-filled requires that they have no air/water interfaces.

3.1.4 - Turgor loss, cytorrhysis, and plasmolysis

When a cell in an intact plant growing in soil loses water, turgor declines and solute concentrations increase. As explained before (3.1.1), at turgor loss point, when turgor becomes zero, the hydrostatic pressure in the cell sap is equal to the atmospheric pressure, meaning that no net force is exerted on the cell wall, and the plant is wilting. If water continues to be lost from the cell, the pressure within the cytoplasm drops below atmospheric pressure, resulting in a force imbalance that collapses the cell wall. The deformation of living cells upon desiccation is called cytorrhysis. Note that the plasma membrane remains in close contact with the cell wall throughout desiccation ie plasmolysis does not occur, because the hydrostatic pressure in the cytoplasm remains greater than the hydrostatic pressure in the apoplast.

Plasmolysis only occurs in cells that are completely immersed in solution and have no air spaces around them, as in epidermal strips floating on water. Plasmolysis starts when the osmotic pressure of the solution is increased above that of the cells, causing the protoplast to shrink, and the plasma membrane separates from the wall (Fig. 3.7). Large gaps created between the plasma membrane and the wall fill with the bathing solution. This cannot occur in normal tissues as the cells have air spaces between them that are not filled with water. This includes root cells of intact plants growing in hydroponic solution or in waterlogged soil, as they still have air spaces.

Air cannot enter the cell through the cell walls as the small pore size, about 15 nm, would need a suction of 20 MPa to drain the pores (Table 3.1, in previous section) which is impossible.        


Figure 3.7. Turgid leaf cell (turgor about 0.5 MPa) and flaccid cell (zero turgor) that has lost some water. With further water loss, the cell collapses. The collapse of the wall is called cytorrhysis. Plasmolysis occurs when a cell is placed in a solution of osmotic strength greater than that of the cell. Water is withdrawn from the cell until its concentration of solutes equals that of the bathing solution. If the bathing solution is sucrose or NaCl or any small molecule (smaller than the pores in the cell wall), solution enters the cell through pores in the cell wall which prevents cytorrhysis.

During plasmolysis, the plasma membrane is stretched into strands that remain tethered to the wall at particular sites (Figure 3.8). Plasmolysis has been used by microscopists to demonstrate the tethering of the plasma membrane to specific sites on cell walls, by floating tissue such as epidermal peels of onion bulbs on high concentration of solution of sucrose (Figure 3.8). When the protoplast shrinks away from the cell wall, and solution with small molecular weight solutes pentrates the cell wall and floods into the space between the wall and the proplasts, some parts of the plasma membrane stay tethered and the rest become pulled into very fine strands.


Figure 3.8. Cells in onion bulb scale leaf epidermis before and after plasmolysis, viewed by confocal microscopy and stained with fluorescent dye DIOC(6). (A) Cytoplasm is seen as pale strands at the cell surface, traversing the large vacuole. Cell boundaries are bright because the surface cytoplasm is intensely fluorescent. (B) Precisely the same field of view after plasmolysis in 0.6 M sucrose. The cell walls are now visible as dark lines between the shrunken protoplasts, which still show brightly fluorescent surfaces. (C) A reconstruction of many planes of focus at higher magnification to show some of the hundreds of stretched strands of plasma membrane that remain tethered to the wall during plasmolysis. (Micrographs courtesy B.E.S. Gunning)

The difference between cytorrhysis versus plasmolysis is most easily seen in leaves with single cell layers like mosses. Figure 3.9 shows that in the hydrated leaflet of Physcomytrella, when the central vacuole is distended, the chloroplasts line the cell wall. Rapid water loss causes a general shrinkage and eventually a collapse at the central parts of the cells. In cytorrhysis, the plasma membrane always remains in close contact with the cell wall. In contrast, when cells are bathed in a solution of small molecules like sucrose, glycerol, or low molecular weight polyethylene glycol, PEG, the solutes pass through the cell wall but not the plasma membrane, causing shrinkage of the protoplast and detachment of the plasma membrane from the cell wall. In plasmolysis, the gaps between the cell wall and the plasma membrane are filled with plasmolytic solution (Figure 3.9).


Figure 3.9. Cytorrhysis versus plasmolysis in Physcomitrella patens. Left: The single cell layer of a hydrated moss leaflet. Centre: Cytorrhysis, where water loss causes a general shrinkage and eventually a collapse at the central parts of the cells. White areas appear where the upper and lower cell walls meet; the chloroplasts are pushed towards the side walls. Right: Plasmolysis in 10% glycerol, which passes through the cell wall but not the plasma membrane. Water loss causes shrinkage of the living protoplast and the detachment of the plasma membrane from the cell wall. (Photographs courtesy I. Lang)

Cytorrhysis also occurs during freezing, when water is withdrawn from cells (Buchner and Neuner 2010).

Plasmolysis is a laboratory phenomenon and does not occur in nature. It is an experimental artifact.

3.1.5 - What drives water flow?

Water flows throughout the plant in three different ways: (a) in bulk, (b) by diffusion in a liquid, and (c) by diffusion as a vapour. Different mechanisms are involved in these three types of flow.

Bulk flow is driven by gradients in hydrostatic pressure. It is much faster than diffusive flow because the molecules are all travelling in the same direction and hence their movement is cooperative. This is the flow that occurs in xylem vessels, in the interstices of cell walls, and in water-filled pores in soil. The resistance to such flow depends very strongly on the size of the flow channels.

Tall trees and fast-growing cereal crops like maize have large xylem vessels, of 100 µm in diameter or more. Flow rates are fast because the rate of volume flow increases in proportion to the fourth power of the radius for a given pressure gradient. Volume flow rate (m3s-1) in a cylindrical tube of radius \( r \) is proportional to \( r^4 \) and to the gradient in pressure along the tube, and inversely proportional to the viscosity \( η \) (Pa s) (Poiseuille’s Law)

\[ \text{Volume flow rate} = \left(\frac{\pi r^4}{8\eta}\right) * \left(\frac{\Delta P}{\Delta x}\right) \tag{6}\]

Where \( η \) is the solution viscosity and \( ΔP/Δx \) (Pa m-1)is the gradient in hydrostatic pressure. From equation (5) we understand that wide tubes are enormously more effective than narrow tubes. The importance of the relationship between tube radius and conductive efficiency becomes apparent when we examine long distance transport of water through the xylem (Section 3.2).

Diffusive flow in the liquid phase is driven by gradients in osmotic pressure. It is much slower than bulk flow because the net flows of solute and water molecules are in opposite directions and therefore impede each other. Where two liquid phases are separated by a semi-permeable membrane the flow of water across the membrane to the phase with the higher osmotic pressure is essentially diffusive, and the flow is driven by the difference in water potential across the membrane.

Vapour flow, for example through the stomata, is driven by gradients in vapour concentration, which are usually expressed in terms of partial pressure, but are nevertheless mechanistically concentrations.

As water in the transpiration stream moves from the soil to the roots, through the plant, and out through the stomata, all three types of flow are involved at various stages.

3.1.6 - The influence of gravity

The potential energy of water is affected by gravity: unconstrained water runs down hill. In most plants the effect of gravity is small relative to common values of the water potential, but in tall trees it can dominate. Where the effect is important it is convenient to introduce the notion of a total water potential, \( Φ \), which is the sum of the water potential, \( ψ \), and a gravitational term, thus

\[\Phi = \psi + \rho gh = P - \pi + \rho gh  \tag{7}\]

where \( ρ \) (kg m-3) is the density of water, \( g \) (m s-2) is the acceleration due to gravity, and \( h \) (m) is the height (relative to some reference) in the gravitational field. \( Φ \) is constant in a system at equilibrium with respect to water even when height varies. The value of \( g \) is approximately 10 m s-2, so the gravitational term, \( ρgh \), increases by 10 kPa for each metre increase in height. Hence, at equilibrium, when \( Φ \) and \( π \) are uniform (at least, in a system without semipermeable membranes) the hydrostatic pressure falls by 10 kPa for each metre increase in height.

In the tallest trees, for example a Eucalyptus regnans 100 m tall, equation (6) predicts that, even when the tree is not transpiring, the water potential at the top is about 1.0 MPa lower than at the base.

3.1.7 - Definitions and further reading

Definition of Terms

Pressure is force per unit area, Newtons per square meter, or N m-2. Its unit is the Pascal (Pa). 1 MPa is approximately equal to 10 kg weight per cm2.

Hydrostatic pressure is the pressure in a stationary fluid. (Note that hydrostatic pressure is usually quoted as the difference from atmospheric pressure, and is therefore taken to be zero when it equals atmospheric pressure).

Turgor pressure is the term used for the hydrostatic pressure in the cells’ contents.

Osmotic pressure (\( π \)) is the hydrostatic pressure in a compartment containing an aqueous solution that will just prevent pure water at atmospheric pressure flowing into that compartment through its membrane that is permeable to the water but not to the solutes within.

Water potential is the difference between \( P \) and \( π \).

Further Reading

Kramer PF, Boyer JS (1995) Water relations of plants and soils.   http://udspace.udel.edu/handle/19716/2830

Nobel PS (2005) Physicochemical and environmental plant physiology (3rd edition). Elsevier Academic Press, Burlington, MA

Passioura JB (1980) The meaning of matric potential. J Exp Bot 31:1161-1169

Passioura JB (2010) Plant–Water Relations. In: Encyclopedia of Life Sciences. Wiley, Chichester. DOI: 10.1002/9780470015902.a0001288.pub2

3.2 - Long distance xylem transport


Figure 3.10 Cross section from a seminal wheat root stained with toluidine blue, showing the cortex, endodermis (black arrow), late metaxylem (LMX) central vessel, and peripheral xylem (red arrow). Many root hairs can be seen. Scale bar, 100 µm. (Image courtesy H. Bramley)

In vascular plants, water absorbed by roots is transported up the plant in the mature (dead) tracheary elements (xylem vessels and tracheids) of roots and stems (Figure 3.10).

Plants are capable of rapidly transporting water to heights in excess of 100 m, even from extremely dry soils and highly saline substrates. They can transport water from soils to leaves at velocities of up to 16 m per hour (4 mm per second) if they have wide xylem vessels in the range of 100 µm. With the more common xylem vessel size of 25-75 µm, peak velocities are 1-6 m per hour. What biophysical mechanism allows plants to achieve this? We know that plants do not possess a pump to move water to the canopy under positive pressure. Instead, plants suck!

Plants have evolved a transport system that relies on water sustaining a tensile force while under suction. The xylem sap in transpiring plants is under negative pressure. This elegant, but counter intuitive mechanism, described by the Cohesion-Tension theory, allows plants to move large quantities of water from the soil to the transpiring leaf surface with little input of metabolic energy. The following section describes the experimental history of how the Cohesion Theory came to be accepted.

3.2.1 - Cohesion Theory for the Ascent of Sap


Figure 3.11 (a) Mercury (black) sucked into tracheids of pine (Pinus radiata) by transpirational pull generated in the shoots. The water—mercury interface is powerful enough to hold this vertical column of mercury in stems. The height to which the dark column of mercury rises is used to calculate suctions created in xylem vessels. Note the generally small heights, reflecting the high specific gravity of mercury. About 2 MPa suction is produced in these xylem vessels. (b) Mercury enters bordered pits but remains connected to the vertical column of mercury in xylem vessels. While mercury can pass through the pit apertures, it cannot pass the finely porous ‘pit membranes’ because it is much more cohesive than water. Seen laterally, the bordered pits appear as discs.

Around 1905, great plans were made to resolve the mystery of the ascent of sap in trees by Professor E.J. Ewart in Melbourne, using eucalypts as a model plant. At that time, Australian mountain ashes (Eucalyptus regnans) vied with American coast redwoods (Sequoia sempervirens) as the tallest trees in the world, being well over 100 m tall. Using special scaffolding, Ewart climbed eucalypt trees, removed lengths of branch and measured the pressures required to push water through these stems. These investigations led him to conclude ‘The ascent of water is, therefore, a vital problem in so far as it depends upon conditions which hitherto can only be maintained in living wood’. If water transport required living cells, it could not be supported by discovery of a pump akin to that in animals. Even roots, which sometimes could pump water by root pressure, lacked the necessary positive pressures to push water so far aloft, especially around midday when water was most needed.

Suction from the shoots was an alternative explanation, but manmade suction pumps cannot do this without inducing formation of air bubbles (embolisms) in the xylem and blocking flow. One clue to the solution came from Dixon and Joly (1894) who claimed that very pure water molecules would be held together by powerful cohesive forces provided the water was especially clean (much cleaner than in manmade pumps).

Ewart did not agree with the unorthodox proposal that the suction of pure water through xylem vessels underpinned transpiration. However, Dixon (1914) ultimately postulated the Cohesion Theory, based on those properties of water which distinguish it as an ideal biological solvent. Cohesion (due to hydrogen bonding between molecules of water), adhesion to walls of the vessels, and surface tension, are central features. In short, in the absence of microscopic gas bubbles water could withstand quite enormous tensions.

Evaporation from wet cell walls of substomatal cavities in leaves creates a large tension (also called negative pressure or suction), which is transmitted via xylem conduits, pulling more sap from roots to leaves. Fine pores in cell walls provide sufficient suction to draw water to the crown of even a lofty tree: a curved interface in a 10 nm pore can store a pressure of -30 MPa. This value can be derived from equation (5) in the previous section: DP = 0.15/r where P is in Pa and r in this case is 5 x 10-9 m.

Through the evaporative power of the atmosphere, a continuous ‘chain’ or ‘catena’ of water, well below atmospheric pressure, could be drawn up to a leaf canopy. The tensions created in this way could even suck water from the surrounding soil. We now recognise that the evaporative energy is supplied as the latent heat of vaporisation ultimately derived from solar radiation. This cohesive property of water gave rise to the ‘Cohesion Theory for the Ascent of Sap’.

Two other properties of water are also essential for long-distance water transport: surface tension, and the adhesion of water to solid surfaces such as the xylem vessels within trees. Dixon claimed that if water could ‘hang together’, the enormous evaporative energy of the air (the same power which dries the washing hanging on a line) could be harnessed to lift sap, which is mainly water, vertically. This would entail no metabolic energy on the part of the plant. This theory of sap flow accorded with earlier experiments by Professor E. Strasburger in 1893 showing that a tall oak tree trunk, severed at the base, could draw poisons and dyes up to the leaves by some wick-like action. If metabolism energised sap flow, poison should have inhibited it. This was well illustrated in later experiments (Figure 3.11a) in which mercury was drawn through fine tracheids of pine stems purely through the suction created by transpirational water loss from the shoot above. The tension required to achieve this is about 2 MPa.

However, the physical properties of plants had to be more complex than those of simple pipes conducting water. As mentioned, manmade pumps failed through embolisms if used to suck water higher than 10 m, whereas hundreds of litres of water reaches the canopies of tall trees daily. Even overlapping sawcuts in tree trunks, which should allow a massive quantity of air to flow into xylem vessels when under suction and cause trees to die from embolisms, did not stop all water flow to leaves. If water was under such suction, how could trees keep air bubbles out of the sap when the trunk was cut? This additional problem was not resolved until the very complex anatomical structures of trunks were much better understood. The highly compartmentalized, extensively redundant structure of the xylem network performs the critical role of isolating gas voids while water transport continues in adjacent conduits. In reality, the complex structure of the xylem network is what makes reliable water transport under tension possible. 

Xylem is not composed merely of pipes: it is made up of partially sealed units (technically vessels, tracheids and fibres, called collectively conduits), which most effectively limit the spread of introduced gases and thus, maintain water flow in some conduits despite very severe disruption from embolisms in others.

3.2.2 - Xylem as an effective conduit for sap


Figure 3.12 Cross section from barley root grown in soil; coleoptile node axile root, bar 100 µm. C is cortex, arrow points to peripheral xylem and arrowhead points to inner xylem. Extensions from the epidermis (red) are root hairs. Section was stained with rhodamine B and viewed with UV fluorescence optics. Micrograph, M. Watt. (Reproduced from New Phytol 178: 135-146, 2008)

The diameter of xylem vessels can be as small as 10 µm as in Arabidopsis, 60-100 µm in the roots of wheat and rapid growing annuals like maize, to over 100 µm in trees. Remembering that trees can be over 100 m in height, the conductive efficiency of xylem conduits is essentials for plants to move water to the canopy at rates that satisfy the transpirational water loss at the leaf surface. These dimensions are for the vessels with maximum diameter, the late metaxylem in the central part of the stele (e.g. Figure 3.12).

The diameter of xylem vessels in a given species varies greatly with root type (Watt et al. 2008). For example, in wheat and barley, the diameter varies from 10 to 60 µm depending on position within the stele (central or peripheral), and the type of root (seminal or nodal). Figure 3.12 shows a section of a nodal root from barley.

A wider xylem diameter translates to an increase in conductive efficiency that can be appreciated by revisiting equation (6). From the Hagen-Poiseuille Law, which shows that flow increases with the fourth power of the radius, we can see that a four-fold increase in the radius of a tube leads to a 256 fold increase in the volumetric flow rate.

In addition to allowing for high rates of water flow, the xylem must also protect the plant against formation and spread of gas bubbles. For xylem sap to sustain tensions required in tall trees, there must be no gas bubbles in the system. Cohesion breaks down if there is a single ‘nucleation site’ on which bubbles can form and enlarge. On the other hand, sap normally contains dissolved gases which, surprisingly, do not disrupt the system provided there are no nucleation sites available. Even the rigid walls of xylem vessels are compatible with high xylem tensions, attracting water by adhesion, which is essential for transport.

Surface tension acts as an interfacial water–air stopper, preventing air from being sucked into the many millions of tiny pores present in all plant cell walls. For example, water delivered to leaf cells by xylem vessels passes through these tiny menisci, which act effectively as non-return valves, so preventing air from being sucked into the xylem (Section 3.3). Surface tension also explains how water in leaves remains under strain within an essentially porous system through which water flows.


Figure 3.13 (a) Scanning electron micrograph (SEM) shows a transverse section of xylem tissue in Brachychiton australis. Large xylem vessels are surrounded by fibres and parenchyma; scale bar, 500 μm. Xylem vessels are dead at maturity and form long hollow tubes that minimise the resistance to water flow through the plant. Connecting intervessel walls contain bordered pits, cavities in the lignified secondary cell walls that allow for transfer of water between vessels. (b) Longitudinal section showing vessels in the xylem tissue of Fraxinus americana. Vessels are made up of repeated individual units (vessel elements) that are joined end to end by perforation plates; scale bar, 400 μm  (c) SEM of the finely sculptured scalariform perforation plates in Betula ermanii xylem. Water passes easily from one xylem vessel to another by this route; scale bar, 20 μm (d) SEM showing surface view of the pit membrane with secondary wall removed by sectioning; scale bar, 2 μm. Tiny pores allow the movement of water between vessels but limit the movement of gas and pathogens. Bordered pits act as the safety valves of the plant hydraulic system. (Images courtesy B. Choat and S. Jansen).

Vascular transport systems have evolved to become amazingly reliable despite the metastable condition of the sap (existing as a liquid below its vapour pressure). From primitive, thickened, hollow cells, increasing specialisation has produced greater elongation and thickening of the tubes (Figure 3.13a,b). Xylem walls contain pits, in which zones of the primary wall known as ‘pit membranes’ allow water to be transmitted between vessels efficiently, while preventing a gas phase spreading through the interconnected system of vessels and blocking transport through embolisation (the blockage of a fluid channel with a bubble of gas) (Figure 3.13d). No living membrane is present in these wall structures. The efficiency with which pit membranes isolate adjacent vessels is shown in Figure 3.11b (in the previous section) where mercury, a highly cohesive liquid, is drawn into specialised bordered pits of pine tracheids without being able to exit into neighbouring tracheids.

Vascular systems have evolved from plant species possessing only fibres and tracheids, for example the more primitive Tasmannia, to more advanced plants possessing vessels which resemble the unicellular tracheids in structure but are much wider and longer and originate from a number of cell initials fused together. Lignin thickening patterns have also evolved. Some thickening designs, such as annular and spiral, allow the tubes to extend longitudinally while supplying growing organs.

When elongation growth has ceased, an organ can be provided with more efficient pipes of larger bore and with stronger thickenings, in reticulate and scalariform patterns (Figure 3.13c). Pit fields which allow water transport across vessel walls can also be simple, unreinforced structures (simple pits) or more elaborate bordered pits in which secondary cell walls mechanically support the pit membrane. All these forms of pits can prevent air in an air-filled conduit from spreading to adjacent conduits which are conducting water under strong suction. Reinforcement of the walls around pits allows pit membranes to be as large as possible and thereby maximise water exchange between vessels.

3.2.3 - Axial flow in the xylem - where does it start?

The late metaxylem carries the bulk of the water to the shoot because of its greater diameter - four-fold increase in the radius of a tube lead to a 256-fold increase in the volumetric flow rate (3.1.5). However, it does not mature until well back from the root tip, and so the younger part of the root is not functional in providing water to the rest of the plant.

Root tips take up enough water for their own cell expansion but they cannot pass this onto the rest of the plant as their xylem vessels are still alive and not able to function as a conduit. Only when they die, mature, and lose the integrity of their plasma membranes, and the cell walls in the transverse plane disintegrate, can they function as conduits.

Xylem conduits (vessels and tracheids) are dead at maturity. They do not mature until sometime after they are fully elongated, and so they remain alive long after they leave the growing zone of the root. The last-formed xylem vessels in angiosperms, the late metaxylem, may be found alive for some distance from the root tip. In Arabidopsis they remain alive until the root hair zone, but in most species they can only be seen further than 5 or even 10 cm from the root tip.


Figure 3.15 A longitudinal face 15 cm from the tip of a main root of a 21 day-old soybean. Portions of 4 developing elements of what will become a late metaxylem vessel (LMX) are shown. The face grazes a mature LMX element (upper left) with very thick wall. The base of the root is toward the left. Diameter of immature xylem is about 50 µm. SEM image, M. McCully. (Reproduced from Protoplasma 183: 116-125, 1994)


Figure 3.16 Cytoplasmic strands in differentiating LMX elements 50 mm from the root tip of barley. Scale bar, 10 µm. Hand cut section of fresh material, Nomarski optics, C.X. Huang and R.F.M. van Steveninck. (Reproduced from Physiol Plant 73: 525-533, 1988)

In soybeans, immature vessels were found as far as 150 mm from the root tip (McCully 1994). In barley the late metaxylem vessel (LMX) was still differentiating 100-150 mm from the tip (Huang and van Steveninck, 1988). Light microscopy of hand-cut sections showed the presence of cytoplasmic strands and intact cross walls in LMX up to 100 mm from the tip (Figure 3.16).

Living/immature xylem vessels can be recognised by their high K+ concentrations, 100 mM or more.  In contrast, xylem sap that flows through the roots has  K+ concentrations of only 5-10 mM, as shown in Table 3.2 in the following section.

When the vessels mature, and their end walls disintegrate, their cellular contents are carried away by the transpiration stream. This leaves hollow tubes that greatly increase the conductance of water flow through the xylem.

Figure 3.17 shows the effect of xylem differentiation and maturity on axial conductance in roots of two crop species, wheat and lupin. Xylem are continually differentiating in lupin within a short region of a young root, which results in dramatic increase in axial conductance. In contrast, once the central metaxylem of wheat has matured there is little change in the root’s axial conductance.


Figure 3.17 Cross sections of wheat (A) and lupin (B) roots stained with berberine-aniline blue and viewed under UV optics. Scale bars, 50 µm. Mature xylem vessels fluoresce due to lignification of their cell walls, which increases with root development (compare upper panels taken 2 cm from the root tip with lower panels taken 18 cm from root tip). (C) shows change in axial conductance as vessels mature in young roots. Images and graph, H. Bramley. (Modified from Plant Physiol 150: 348-364, 2009)

3.2.4 - Solutes in xylem sap

Xylem sap contains all the inorganic nutrients needed for plant growth, in the proportions in which they are needed. The concentrations of some nutrients are dilute when compared to phloem sap, in particular potassium and nitrogen-containing solutes (Table 3.2). The concentrations also vary at different times of day, being lowest in the middle of the day when transpiration is highest, and quite high at night when stomates are closed so there is very little flow of sap to the shoots. The osmotic pressure of xylem sap therefore ranges from less than 0.05 MPa during the day to about 0.15 MPa (60 mOsmol L–1) during the night. The flux of solutes (the concentration multiplied by the flow rate of the sap) is maintained at a steady rate over the whole 24 hours.

Most solutes in xylem sap are inorganic ions (e.g. nitrate, potassium, magnesium and calcium), but organic solutes are also present (Table 3.2). Organic acids and amino acids in xylem sap can be present in substantial concentrations, and sugars (but not sucrose) reaching 5 mM in some perennial species. Many trees including eucalypts are host to boring insects at particular times of the year, when sugar and nitrogen content of the sap is nutritionally valuable. Even though sugar concentrations in xylem sap are much lower than in phloem sap, the high nitrogen to sugar ratios and low osmotic pressures make it a good substrate for many herbivores. More extreme examples of the carbohydrate content of xylem sap are temperate deciduous trees such as maple, which have traditionally been tapped to yield a sugary solution in the period prior to budburst. This indicates that xylem can be a conduit for carbon remobilisation in addition to its central role as a pathway for water and nutrient transport. Xylem parenchyma cells load ions into xylem vessels in roots (Section 3.6) and also contribute to modification of ion levels along the xylem pathway. In secondary tissues, rapid transfer of solutes into and out of the xylem is partly achieved through close association of living ray cells and xylem vessels.

Other organic molecules act to transport inorganic nutrients to the shoots. Nitrate and ammonium are assimilated into organic forms, such as amino acids, in the roots of many plants. In legumes, nodules deliver an even wider selection of nitrogenous compounds to the xylem, including ureides and amides. These often constitute the dominant form of nitrogen reaching shoots and are therefore, a major component of the sap. Other examples of complexed forms of inorganic nutrients in xylem sap are metal ions such as zinc, copper and iron, which are almost exclusively chelated to organic acids.

Phytohormones such as abscisic acid and cytokinins are present in xylem sap and serve as signals from roots to shoots affecting growth and development, and other physiological responses.

3.3 - Leaf vein architecture and anatomy

A rapidly transpiring leaf can evaporate its own fresh weight of water in 10 to 20 min, though many plants such as cacti, mangroves and plants in deep shade have much smaller rates of water turnover. Leaf veins must carry this water to all parts of a leaf to replace evaporated water, and maintain cell hydration and turgor. When water supply fails to meet this demand, shoots wilt.


Figure 3.20 Typical leaf vein pattern of broad-leaf species versus grasses. Left: Leaf of Eucalyptus crenulata showing the arrangement of supply and distribution veins. Large veins with large vessels, in which water is moved rapidly across the lamina, surround islets of small veins with small vessels in which water is slowly distributed locally. Scale bar, 1 mm. Right: Leaf of wheat (Triticum aestivum) showing one large and three small longitudinal veins, with transverse veins connecting them. Scale bar, 0.1 mm. Distance between small veins is about 0.15 mm in both species. (Photographs courtesy of M. McCully and M. Canny)

Vein distribution patterns differ markedly between broad-leaf species and grasses. Broad-leaf species generally have a highly branched network while grass species have parallel veins (Figure 3.20).

Veins consist typically of tightly packed xylem and phloem tissues surrounded by a parenchymatous or fibrous sheath. Both xylem and the phloem contain living parenchyma cells as well as their characteristic transporting conduits: vessels and/or tracheids in the xylem tissue, plus sieve tubes in the phloem tissue. There are no intercellular air-spaces, or only very small ones.


Figure 3.21 Transverse section of small intermediate vein of wheat leaf, after being fed the fluorochrome sulphorodamine, which acts as a tracer for water movement. Water passes out of the xylem through paths in the walls of the mestome sheath cells (M) and enters the parenchymatous sheath cells (P) leaving red crystals in the intercellular spaces. Scale bar, 15 µm. Image, M. Canny. (Reproduced from New Phytologist, Tansley Review No 22, 1990)

The ring of cells forming the sheath around the xylem and phloem tissue acts both as a mechanical barrier that may confine pressure within the vein, and a permeability barrier that can control rates and places of entry and exit of materials (Figure 3.21).

3.3.1 - Vein architecture of conifers and angiosperms

The simplest vein architecture is found in conifer needles where a single unbranched strand of xylem and phloem is surrounded by mesophyll (Figure 3.22).


Figure 3.22 Transverse section of a pine needle. A single central vein has two strands of phloem (p) and xylem (x), embedded in transfusion tissue (t). These vascular tissues are separated from the chlorophyll-containing mesophyll (red) by an endodermis (e) with Casparian strips and suberised lamellae. Stomata in the epidermis appear bluish. Rhodamine B stain, fluorescence optics. Scale bar, 0.1 mm.  (Photograph courtesy M. McCully)


Figure 3.23 Wheat leaf showing one large supply vein and three small distribution veins, connected by transverse veins. Small veins could carry two thirds of the evaporated water. Dark-field optics, blue filter. Scale bar, 0.1 mm. (Photograph courtesy M. McCully)

Vascular strands are enclosed by an endodermis that separates them from the mesophyll, and are embedded in a mixture of parenchyma cells and tracheids called transfusion tissue. Water from the xylem permeates radially outward through transfusion tissue, endodermis and mesophyll to evaporate below lines of stomata in the epidermis.

Leaves of angiosperms have much more complicated venation than conifer needles. If you look at a grass leaf with your hand lens, parallel veins run the length of the leaf, but they are not all the same size. A few large veins have several small veins lying between them. On closer inspection with a light microscope, all these parallel veins are connected at intervals by very small transverse veins (Figure 3.23).

There are in fact two vein systems with different functions: large veins supply water rapidly to the whole length of a leaf blade while small veins and their transverse connections distribute water locally, drawing it from the large veins. Water in large veins flows only towards the tip, but in small veins it can flow either forwards or backwards along the leaf blade or transversely between adjacent parallel veins. The distinction of flow patterns in large and small veins arises as a result of different vessel sizes. Large veins have wide vessels (about 30 µm diameter), while small veins have narrow vessels (about 10 µm diameter). As the volume of water flowing along pipes is proportional to the fourth power of the radius (Equation 6; Poiseuille’s Law), volume flow in the larger vessels will be 3 to the fourth power (= 81) times greater than the flow in the smaller vessels. Put another way, pressure gradients along the leaf in large vessels will be very slight, but steep pressure gradients can develop locally in narrow vessels that will direct local flows into the mesophyll. Large veins supply water rapidly over the whole lamina while small veins distribute it locally and slowly. The slower flux along minor veins is compensated by their far greater number, which results in them having a greater length per unit leaf area than major veins.

With a hand lens, you do not see the vessels, instead you see the sheaths that surround xylem and phloem, much as an endodermis surrounds vascular strands of a conifer needle. Grass leaf veins have two sheaths of cells surrounding the parallel veins and containing the xylem and phloem tissues. In the leaf anatomical development typical of C3 species (e.g. wheat), both sheaths are parenchymatous and lack chloroplasts (Figure 3.24).


Figure 3.24 Wheat leaf showing a single large (supply) vein comprising three large vessels (V). The vein is surrounded by two sheaths of living cells, the inner mestome sheath (arrowheads), and outer parenchyma sheath (stars). The mestome sheath of these veins is impermeable to water. Water enters the symplasm at the inner boundary of the parenchyma sheath (Canny 1990). Phloem (P). Transverse hand-section, Toluidine blue stain, bright-field optics. Scale bar, 0.1 mm. (Photograph courtesy M. McCully and M. Canny)


Figure 3.25 Minor vein of maize, consisting of one xylem vessel (v), five vascular parenchyma cells (vp), bundle sheath (bs), companion cell (cc), and sieve tube (st) with intercellular space (is). Scale bar, 4.2 µm. Transmission electron micrograph, R.F. Evert. (Reproduced from Planta 138: 279-294, 1978)

The large veins of a C3 species is surrounded by two sheaths of living cells, the inner mestome sheath and outer parenchyma sheath (Figure 3.24). The mestome sheath of these large veins is impermeable to water. There is no apoplastic path for water through the mestome sheath of large veins, except through a connecting transverse vein. In small veins, by contrast, two or three mestome sheath cells next to the xylem permit flow of water and solutes through the cell wall apoplasm.

In C4 species (e.g. maize), only the inner (mestome) sheath is without chloroplasts. The outer ring of sheath cells contains large chloroplasts and is known as the bundle sheath (Figure 3.25). This is the cell layer in which CO2 fixation in the Calvin cycle takes place in C4 plants. This is an essential part of the carbon concentrating mechanism, and the special anatomy of C4 photosynthesis, as detailed in Chapter 2, Section 2.2.2

A dicotyledonous leaf contains the same two vein systems as a grass leaf, but these are differently arranged. Large supply veins are prominent, comprising a midrib and two orders of branches off it, often standing out from the surface of the lamina. These contain wide vessels and carry water rapidly to the leaf margins. Between them lie distribution veins, another two branch orders of small veins dividing the mesophyll up into islets about 1–2 mm across (Figure 3.26), and within these islets a fifth and final order of branches of the finest veins. The fourth- and fifth-order branches have only narrow vessels (Figure 3.26).

In dicotyledenous leaves, as in grass leaves, these vascular tissues are enclosed by bundle sheath cells through which water and solutes must pass when leaving the xylem.


Figure 3.26 Whole mount of a cleared leaf of Eucalyptus crenulata showing the complicated arrangement of supply and distribution veins characteristic of a dicotyledonous leaf. Islands marked out by large veins with large vessels, in which water is moved rapidly all over the lamina, surround islets of small veins with small vessels in which water is slowly distributed locally to the mesophyll. Partial phase-contrast optics. Scale bar, 1 mm. (Photograph courtesy M. McCully and M. Canny)

Any distribution network such as the branching vessels of decreasing size in leaves is found to obey Murray’s Law. This states that the cube of the radius of a parent vessel is equal to the sum of cubes of the radii of the daughter vessels (e.g. a 50 µm vessel would branch into five 30 µm vessels). Such a pattern of branching produces optimal flow in several senses: minimum energy cost of driving that flow, minimum energy cost of maintaining the pipeline, constant shear stress at the walls of pipes, and rapid flow in supply pipes with slow flow in distributing pipes to permit exchange through the pipe walls (LaBarbera 1990).


Figure 3.27 Soybean leaf showing two of the smallest veins surrounded by bundle sheath cells. Veins of this size are the end of the branching network shown in Figure 3.26, and supply most water that is evaporated. This leaf was transpiring in a solution of fluorescent dye for 40 min. The small vessels in each vein contain dye solution which has become concentrated by water loss to the symplasm and out through the bundle sheath. Dye has started to diffuse away from small vessels in the cell wall apoplasm of bundle sheath cells. Anhydrous freeze-substitution and sectioning, fluorescence optics. Scale bar, 50 µm. (Photograph courtesy M. McCully and M. Canny)

The ring of cells forming the sheath around the xylem and phloem tissue (Figure 3.27) acts both as a mechanical barrier that may confine pressure within the vein, and a permeability barrier that can control rates and places of entry and exit of materials. Exceptions are to be found at the ultimate ends of some dicotyledonous fine veins where tracheids or sieve elements may be unaccompanied by other cells, in the transverse veins of grasses which have no sheath, and in some special veins at leaf margins where the sheath is absent on the xylem side. There is evidence of a suberised layer in walls of these sheaths cells in some species, but not in others. It is uncertain whether xylem sap must traverse the cell membranes, as in the suberised endodermis of roots, or if it can travel along the cell walls.

3.3.2 - Damage control

Transpiration operates by suction, but leaves are especially liable to damage by grazing, mechanical forces, and extreme negative pressures in the xylem water column due to high rates of transpiration. Leaf vessels therefore need special protection against air embolisms spreading in the vessel network that would block liquid flow. This is achieved at all points in the leaf distal to the node (i.e. petiole, large veins, small veins) by the vessels being very short. That is, files of vessel elements joined to make a single pipe with a terminal end-wall are much shorter in the leaf than in the rest of the plant. Water flows through vessel end-walls with little extra resistance, but an air–water interface cannot be pulled through an end-wall or pit membrane because the pores are so small. The force needed to curve the interface into a meniscus small enough to pass through the end-wall is similar to that generated by evaporation from wet cell walls (Section 3.1.3). From equation (4), (DP = 0.15/r) we see that a pressure of roughly 0.3/d (MPa) is required to pull the interface through a hole of diameter d (µm). While 0.1 MPa can pull an interface through a 3 µm hole, 6 MPa is required to pull air through a 0.05 µm hole. Suctions of 6 MPa are not known to occur in transpiring plants, and cell walls have pores much smaller than 0.05 µm - in the order of 0.003 µm. So an embolism formed from cavitating water fills one vessel but does not spread beyond it.

The length of xylem vessels is demonstrated by allowing a leaf to transpire in a fine colloidal suspension that cannot pass end-walls. Latex paint, diluted 100 times with water and allowed to settle for a week or two, provides such a suspension. Leaves that have drawn up this suspension for an hour or so during transpiration can be cleared by dissolving out the chlorophyll and soaking in lactic acid. Progress of the paint is then readily seen (Figure 3.28). Very few vessels exceed 1 cm in length.


Figure 3.28 Cleared whole mount of a wheat leaf demonstrating the frequent occurrence of end-walls in leaf vessels. The leaf was fed an emulsion of green latex paint in the transpiration fluid from a cut surface 6.5 mm to the right of the picture. At the right side of the picture, two vessels in the central large vein are carrying paint. Halfway across the picture (at arrowhead) the upper vessel is blocked by an end-wall through which the paint particles could not pass, although water continued to flow. The paint in the lower vessel continued for another 3 mm beyond the left of the picture, where an end-wall in that vessel limited its further progress. Note that paint has not passed out of the large vein into transverse veins where water flowed because pit membranes (Section 3.2) filtered out paint particles. Bright-field optics. Scale bar, 100 µm. (Photograph courtesy M. McCully and M. Canny)

3.3.3 - Water extraction from the xylem


Figure 3.11d SEM showing surface view of the pit membrane with secondary wall removed from most pits by sectioning. Tiny pores allow the movement of water between vessels but limit the movement of gas and pathogens. Bordered pits act as the safety valves of the plant hydraulic system. (Photograph courtesy B Choat and S Jansen)

Xylem vessels (and tracheids) are not just pipes to carry water, they are pipes with holes in them (pits) through which water can leak out, fulfilling a principal leaf function of water distribution through the transpiration stream to places where it will evaporate. The branching network of vessels is beautifully adapted to achieve this.

Think about flow in a leaky vessel. As explained earlier, the rate of volume flow varies as the fourth power of the radius (Poiseuille’s Law). The frequency of leaks through vessel walls varies with surface area of the walls, that is, as the first power of the radius (2πr). So if the vessel is wide, forward flow is much larger than leakage. The wide vessels in large veins supply water all over the leaf without losing much on the way. As the width of a vessel becomes smaller, the forward flow (a function of r4) is reduced much more strongly than the leaks (a function of r). The proportion of water lost to leakage therefore increases as vessels become smaller. Indeed, for a fixed pressure gradient there is a critical radius at which all water entering the vessels supplies leaks, and there is no forward flow at all. The finest veins of dicotyledenous leaves have vessels of a radius that is close to this critical value. As sap disperses into the fine ramifications of the network, it moves more and more slowly forward, and leaks increasingly outwards through the sheath to the mesophyll. This is the rationale of the distributing networks of the small branched veins of both dicotyledons and grasses.


Figure 3.29 Fresh paradermal hand-section of a leaf of Eucalyptus crenulata which had been transpiring for 80 min in a solution of sulphorhodamine G. The dye solution is present at low concentration in the vessels of the larger veins, but is not visible at that concentration. In the smallest veins the dye has become so concentrated by loss of water to the symplasm that dye crystals have formed inside vessels. Sectioned in oil, bright-field optics. Scale bar, 100 µm. (Photograph courtesy M. McCully and M. Canny)

Extraction of water from fine veins can be readily demonstrated. Stand a cut leaf in an aqueous solution of dye, such as 0.1% sulphorhodamine G, and allow it to transpire for an hour. The solution moves rapidly through large veins all over the leaf in a few minutes. Then it moves increasingly slowly into the network of distributing small veins. By the end of an hour it has reached the ends of the finest veins and, as water is extracted from them, the dye becomes more and more concentrated (Figure 3.29). Movement of dyes from the finest veins to leaf surfaces 100 µm away takes about 30 min, suggesting that diffusion rather than mass flow is responsible for solute distribution to cells.

3.3.4 - Solute extraction from the xylem


Figure 3.33 Leaf of horse chestnut, Aesculus hippocastanum, showing as with all strongly-toothed leaves, conspicuous veins running to each tooth. (Photograph courtesy M. Waters)

(a) Phloem export

Potassium is known to be extracted from the xylem and re-exported via the phloem (Section 5.1). In leaves, the concentrated sap flowing through the narrow xylem vessels of fine veins is separated from sieve tubes of the phloem by only two or three parenchyma cells, frequently enhanced by transfer cells. Exchange from xylem to phloem is probably made by this direct path. Solutes travel in the phloem back to stems, either distally to younger developing leaves and the stem apex or proximally towards roots.

(b)  Scavenging cells  

Recycling of solutes out of leaves can also be fed by a variety of special structures adapted for processing rather larger volumes of sap, and so suited for collecting solutes present in the stream at quite low concentrations. The transfusion tissue of conifer needles is one of these structures. As sap leaves the xylem of a vascular strand it moves through a bed of tracheids mixed intimately with transfusion parenchyma cells (Figure 3.22 shown earlier). The endodermis acts as the ultrafiltration barrier, allowing water to pass through while leaving dilute solutes to accumulate in transfusion tracheids. Transfusion parenchyma cells have very active H+-ATPases in their cell membranes which accumulate selected solutes (certainly some amino acids) back into the symplasm for return to the phloem and re-export. Such actively accumulating cells are called scavenging cells.

A tissue that acts in the same way is a special layer of cells in the central plane of many legume leaves (extended bundle sheath system or paraveinal mesophyll). It consists of scavenging cells with active H+-ATPases and accumulates amino acids, stores them and forwards them to developing seeds via the phloem.

Jagged ‘teeth’ on the margins of many leaves also contain scavenging cells (Figure 3.33). Veins carry large volumes of the xylem sap to these points, where evaporation is especially rapid. Within a ‘tooth’, xylem strands end in a spray of small vessels among a bed of scavenging cells. Scavenging cells can thereby collect amino acids and load them into the phloem.

(c) Solute excretion

Not all the solutes of the transpiration stream are welcome back in the plant body. Some, such as calcium, are immobilised in insoluble compounds (calcium oxalate crystals) and shed when leaves fall. Others are excreted through the surface of living leaves. A striking excretion system is found along the margin of maize leaves. Here the outermost vein has a single very wide vessel. Rapid evaporation from the exposed leaf edge cooperates with this low-resistance vessel to draw to the leaf margin all residual solutes that have not been taken out of the stream by other veins. Thus foreign material (like dyes) accumulates in this outermost vessel. The vein sheath is missing from the outer part of this vein so that vessels abut directly the airspace at leaf margins (Canny 1990). Solutes are excreted from this marginal vein, dissolved out by rain and dew, and, more actively, at night time by guttation fluid when there is positive pressure in the xylem. A similar accumulation of dye from the transpiration stream is shown along the margin of a eucalyptus leaf in Figure 3.34.


Figure 3.34 Whole mount of the living margin of the eucalypt leaf sectioned in Figure 3.11 prepared after dye had been fed to the cut petiole for 90 min. Dye has spread to the leaf margin in large veins where it accumulates at high concentrations. By analogy with maize leaves (Canny 1990) this is likely to be a system for excreting unwanted solutes. Bright-field optics. Scale bar, 1 mm. (Photograph courtesy M. McCully and M. Canny)


3.4 - Water movement from soil to roots

Plants lose water by evaporation through their leaves to the atmosphere. The loss is made good by water flowing from the soil into the roots, and thence within the xylem to the leaves. The factors that provide resistances to the flow of water from soil to root are set by the nature of the soil, the proliferation of roots within it, and the contact between the soil and the root. Root system architecture and the proliferation of lateral roots are covered in the following chapter, section 4.1.

Water removed by transpiration results in drier soil around roots compared with bulk soil. This provides the gradient in suction necessary for water flow towards roots to continue.  As soil dries near the root surface, water flows radially from bulk soil to replenish it. When the bulk soil becomes drier, greater suctions are necessary to sustain the flow. The next section describes the capacity of different types of soils to hold water when wet, and release it to plants as they dry. It explains the terms field capacity and permanent wilting point. Following sections quantify the uptake of water by roots, and describe a major barrier to water uptake – the soil:root interface.

3.4.1 - Water in soils

Water drains through soil due to gravity, leaving medium-sized pores still filled with water, and with thin films around soil particles. Soil is porous and holds water in its pores by capillary forces. As a soil dries, the larger pores drain and the remaining pores hold water ever more tenaciously. Water in these pores is under suction (negative hydrostatic pressure, P) and this suction typically ranges from about 10 kPa to about 1.5 MPa in soils supplying water to plants. At suctions of less than 10 kPa, water is held in such large pores that it is likely to drain quickly away; at suctions greater than 1.5 MPa, most plants are at their limit for exerting sufficient suction to extract the water.

Field capacity

When substantial rain falls on a freely draining soil, it may fill all the pores, even to the point of run-off. Thereafter, water quickly drains out of the largest pores under the influence of gravity into the drier soil below. This process of redistribution continues for days, even weeks, but it does so ever more slowly as the remaining largest pores continue to drain. During this time the hydraulic conductivity of the soil falls by several orders of magnitude because the resistance to viscous flow through water-filled pores depends inversely on the 4th power of their diameter.

After about 2 days of redistribution, the rate of drainage typically drops to only a few mm per day, which in the field is similar to other forms of water loss from the soil: uptake by roots, or evaporation from the soil surface. At this point there will be a suction (negative pressure) in the soil water of somewhere between 10 and 30 kPa, and the water content then is known as “field capacity” or “drained upper limit”.

Soil types

Soil scientists classify soils according to particle size as:

Gravel: > 2 mm
Sand: 0.05 – 2 mm
Silt: 0.002 – 0.05 mm
Clay: < 0.002 mm (2 µm)

The structure and texture of the soil determines how much water can be held in it. Sand particles are large (50 mm - 2 mm) and hold onto little water, but a sandy soil provides good aeration. Sand holds little water because the pores between the grains are large. Therefore, its field capacity, the amount of water in the soil remaining a day or two after irrigation, can be as low as 10% (weight per volume; or 10 mL water per 100 mL soil volume).

Clay particles are much smaller (less than 2 mm) and easily compacted. That makes clay a good material for building bricks, but not so good for allowing water, air, and plant roots through. Clay particles have stacked platelets which provide large amounts of surface area. Clay soils store large amounts of water and have a field capacity above 40%, though much of that may be in pores too small for plants to extract water, as shown in the table below.

Silt is the medium size particle (2-50 mm), with better water retention than sand, but less nutrients than clay. Loam is a soil comprised of roughly equal amounts of sand and silt and a little less clay.

Different types of soil are comprised of various proportions of these particle sizes, as illustrated in Figure 3.37.


Figure 3.37 Textural classification of soils. Loam is a soil comprised of roughly equal amounts of sand and silt and a little less clay. http://imgkid.com/soil-texture-triangle.shtml

The diameter of pores in soil is roughly equal to one fifth of the diameter of the soil particles. Thus, at the boundaries between these classes, the approximate diameters of the pores would be as shown in Table 3.3 with the suction needed to drain them.

This table shows that many pores in a gravelly soil would drain at a suction of only about 1 kPa, but that a clay soil would need ten times this suction, close to 1 MPa, to drain many pores. Transpirational pull by plants can provide suctions above this, but there is a limit at which plants can maintain suction without the risk of embolism.

Wilting point

This term is used by crop physiologists, and often called the “lower limit of water extraction” by agronomists and crop modellers. When the water content of a soil is below the wilting point, plants have difficulty accessing it, as they cannot generate a sufficiently low water potential. In practice, this critical water potential has been found to be about -1.5 MPa, which effectively marks the lower limit of the available soil water. 

Soil at wilting point is not dry. Different soil types hold very different amounts of moisture at their wilting points. As plant remove water and the soil starts to dry, the small pores make it difficult for roots to extract all the water, so clay soils hang onto their water more strongly than sandy or loamy soils, and results in the plant available water being less for clay soils than for loamy soils which have a lower clay content (Table 3.4).  Silt contains pore sizes intermediate to clay and sand, and makes up about 50% of the composition of loamy soils (Figure 3.37) and give it the favourable property of retaining more water than sand as the soil but releasing it more easily than clay.


Figure 3.38 Clay soils are fine textured and have a greater water-holding capacity than do sandy soils that are composed of larger particles. Clay soils hold a greater amount of plant-available water between field capacity (Ψsoil about -0.02 MPa) and wilting point (Ψsoil about -1.5 MPa) than do sandy soils. In both situations, water is held with progressively stronger forces as the soil mass dries out. (R O. Slatyer and I.C McIlroy, 1961).

The relationship between soil water content and soil water potential (Ψsoil) is shown in Figure 3.38 for the three different soil types. Water is held with progressively stronger suctions as the soil dries, and at the permanent wilting point of -1.5 MPa clay soils retain much more water than sands or sandy loams that is not available to plants.

Another feature of clay soils that makes them less desirable for agriculture or horticulture is that root length density (length of roots per unit volume of soil) tends to be lower than in the lighter textured loamy and sandy soils. Despite their substantial moisture reserves, fine-textured clay soils are generally less hospitable to plant roots than loamy or sandy soils. As a result, plant-extractable moisture in clay soils is somewhat less than the soil’s physical properties alone would imply.

3.4.2 - Water in pots

Field capacity is not a term relevant to pot experiments, unless they are very tall. After irrigation, a drained pot will contain much more water than soil in the field.

The soil at the bottom of a freshly watered and drained pot is inevitably saturated with water; if the pot is short the whole of the medium may have such little air-filled porosity that it becomes hypoxic. This is a special problem with field soils used in pots, for these typically do not contain many pores large enough to be drained at the small water suctions that prevail. Such suctions are zero at the bottom of a freshly drained pot and rise by 1 kPa for every 100 mm increment in height, leading to suctions at the top of a typical pot that are much less than the 10–30 kPa that occurs in soils in the field that have drained to ‘field capacity’ (Passioura 2006).


Figure 3.39 The relationships, at equilibrium, between soil water suction (a), effective diameter of largest water-filled pores (b), and height above a free water surface, such as a water table within the soil or the bottom of a recently drained pot. The dotted lines exemplify conditions at a height of 100 mm. (J.B. Passioura, Funct Plant Biol 33: 1075–1079, 2006)

When a pot has finished draining, the soil at the bottom is saturated: it has zero water suction. The suction varies linearly with height, as illustrated in Fig. 3.39, from zero at the bottom of the pot, to 10 H Pa, where H is the height of the pot in mm. Thus, a freshly watered and drained pot that is 100 mm high has a water suction at its top of 1 kPa, very much less than the 10–30 kPa that occurs in a freshly watered and drained soil in the field. Similarly, a pot that is 200 mm high has a suction of 2 kPa at the top. Thus the average suction and thence average soil water content of a freshly watered and drained pot depends on its height.

It is worth noting that the size and number of drainage holes in the bottom of the pot have no bearing on the distribution of suction with height at equilibrium. The rate at which equilibrium is attained will also depend little on the number and size of the drainage holes — just one hole of a few mm diameter is adequate.

The implication of these small suctions is that much larger pores contain water than in a drained soil in the field. For example, at the top of a pot that is 100mmhigh, and where the suction is 1 kPa, all pores less than 0.3 mm wide will contain water. This contrasts greatly with the field soil, in which the suction will be at least 10 kPa and the diameter of the largest water-filled pore will be 0.03 mm or even less.

This has considerable implications for doing pot experiments on the availability of soil water to plants using field soils. Standard watering techniques, which result in adding excess water that then drains from pots, lead to initial soil water contents that are much more than they would be in the freshly drained soil in the field. To match what happens in the field, it would be necessary to water the pots by weight to reproduce a water content consistent with a suction somewhere between 10 and 30 kPa. Further, the large initial water content can produce problems with aeration.

The plant nursery industry is well aware of the difficulties in adequately aerating pots. That is why their potting mixes contain peat or vermiculite or other bulky materials that create large pores of 1mm or more in diameter, which contain air at heights greater than 30mm above the bottom of a freshly drained pot. These large pores protect the roots from hypoxia in a material that might otherwise have a dangerously small air-filled porosity. Nevertheless, despite these large air-filled pores giving the roots growing in them access to oxygen, the interior of aggregates in the soil or potting mix will still be essentially saturated and possibly hypoxic.

Data for three different types of potting mix are shown in Figure 3.29: horticultural topsoil, commercial potting mix is designed for growing plants for sale in garden nurseries, and fine potting mix designed for growing Arabidopsis thaliana in small pots. The topsoil is the worst aerated, with the bottom 150mm of a freshly drained pot in danger of becoming hypoxic. It is notable that in the small pots (70 mm tall) often used for growing Arabidopsis, all of the medium is in danger of becoming hypoxic, for the porosity ranges from 0 to only 7%within the pot (Figure 3.40). With a frequent watering regime, say twice daily, the medium could be permanently hypoxic. Commercial potting mixes overcome this problem by using coarse materials in the mix, which create many large pores (>1 mm diameter) that drain at small suctions.


Figure 3.40 Air-filled porosity of three examples of potting media as functions of height above a free water surface such as the bottom of a freshly drained pot. There is danger of hypoxia at air-filled porosities less than 10%. (J.B. Passioura, Funct Plant Biol 33: 1075–1079, 2006)

Field capacity should therefore not be confused with the water content of a drained pot, which should be called “pot capacity”.

3.4.3 - Uptake of water by roots

Flow of water through soil is induced by gradients in hydrostatic pressure, P. The rate of flow, F (m s–1), depends on both the gradient in P and on hydraulic conductivity, K  (m2 MPa–1 s–1), of the soil:

\[F=K \frac{dP}{dx} \tag{8}\]

where x (m) is distance. This equation is Darcy’s Law. Conductivity, K, varies enormously, by about a thousand-fold, over the range of available water content. The reason for this large range is that water flows much more easily in a large pore than in a small one. As explained earlier (Section 3.1), the flow rate varies according to Poiseuille’s Law — the flow in capillary tubes depends on the fourth power of the diameter. As the soil dries, the large pores are empty and water is drawn from smaller and smaller pores.

Water removed by transpiration results in drier soil around roots compared with bulk soil. As soil dries near the root surface, water flows radially from bulk soil to replenish it. Calculated distributions of water content and pore water pressure with radial distance from an absorbing root predict a pronounced increase in suction adjacent to the root

(a) Theory of water uptake

Roots are cylindrical, so the flow of water to them is radial. This has considerable consequences: the flow lines converge as they approach the root, which requires increasing gradients of pressure in the soil water to keep the water flowing (Figure 3.41).


Figure 3.41 Calculated distributions of volumetric soil water content (left) and pressure in water-filled pores (right) as functions of distance from a model root. Pressures become more negative over time, indicating increasing suction. Horizontal lines denote water status on each of six successive days (day 1 is uppermost). The steepening of curves at later times reflects how transport of water from bulk soil to the root surface becomes increasingly difficult as the soil dries. r = distance from central axis of root.

A radial flow of water from the bulk soil towards roots of transpiring plants is generated by suction at the root surface. However, because K decreases with falling water content, there is a limit to how fast roots can extract water from soil. Once this limit has been reached, increasing suction by roots simply steepens the gradient in P to match the fall in K close to root surfaces so that the product of the two (Equation 3.1) remains the same.

Although soil water is driven by gradients of pressure, it is necessary when water content is changing to describe this flow in terms of gradients in volumetric water content, θ (m3 m–3). The coefficient relating flow rate to the gradient in water content is known as diffusivity, D (m2 s–1), and the appropriate equation is formally analogous to Fick’s First Law of diffusion:

\[F=D \frac{d\theta}{dx} \tag{9}\]

This equation can be elaborated to allow for the cylindrical flow, and then solved to derive an approximate relationship for the uptake of water by a population of roots:

\[Q \approx 2D*\Delta \theta * L \tag{10}\]

where Q is the flow rate of water through the soil, (m3 m–3 s–1), now expressed as the overall rate of change of θ in the sample soil, and L is the rooting density, average length of absorbing root per unit volume of soil (m m–3).

Like K, D varies with the soil water content, although not so widely.  Laboratory measurements of  D, which are so far the only ones that have been made with some accuracy, show that D is about 10-7 m2s-1 when the soil is fairly wet, and falls to about 10-9 m2s-1 once the soil has dried far enough for the soil water suction to exceed about 300 kPa.  Where D is of the order of 10-9 m2 s-1 one can calculate from eqn 3.3 that wherever the density of roots exceeds about 1 cm per cm3 of soil (104 m m-3) the flow of water through the soil is not likely to limit uptake until there is almost no available water left in the soil. But where there is less than about 1 mm of root per cm3 (103 m m-3) of soil, uptake is likely to be sufficiently limited by flow through the soil that it would take several days for the roots to extract most of the available water.

 (b) The uptake of water by roots - in practice


Figure 3.42 Sample of dense subsoil containing a biopore occupied by a root. Such biopores are passages for roots through otherwise almost impenetrably dense soil.

Many experiments have been done to explore this theory, with mixed results.  Most have involved growing plants in repacked soil in controlled laboratory conditions. Some have confirmed the theory, many have not. The most likely reasons for the discrepancies is that interfacial effects - at the junction between the surface of the root and the soil - come into play.  These are discussed in the next section.

In field soil, the concentration of roots in the topsoil is usually so high that the local rate of uptake of water is unlikely to be ever limited by the properties of the soil.  It is in the subsoil that the rooting density drops to a level at which flow through the soil may limit the rate of uptake. Further, the rate of uptake is often much lower than what the simple theory would predict, even when the often sparse rooting density is taking into account.

In subsoil, there are other possible reason for the discrepancy between theory and observation.  One is that, in contrast to disturbed soil, either in ploughed topsoil or repacked in lab experiments, roots in the subsoil do not ramify more or less randomly through the soil. Subsoils are usually dense and difficult to penetrate.  Roots grow predominantly in pre-existing fissures or in continuous large pores, biopores, made by previous roots or soil fauna (Figure 3.42). 

A second reason is that it may be wrong to extrapolate laboratory measurements of D in repacked soil to undisturbed soil in the field. The undisturbed structure of the soil may inhibit the flow of water.  For example, the formation of aggregates of particles in the soil often results in particles of clay (which are usually in the form of small plates) becoming oriented parallel to the surface. Such orientation is likely to increase greatly the flow path for the water, but so far no reliable measurements of D have been yet made on undisturbed subsoil.

In summary, the dense root systems common in topsoils extract water effectively from surface soil layers. Extracting water from subsoil layers is more difficult. Australian subsoils are typically inhospitable to roots. They are dense, have a large resistance to penetration. They are often sodic, that is, sodium dominates the exchange complexes on soil particles, altering the soil structure so that it sets like concrete when dry, and becomes impermeable to water when wet. Moreover, subsoils can be acutely deficient in some nutrients that are required locally by roots. Native vegetation overcomes these difficulties by forming deep biopores in the subsoil. For example, roots of jarrah trees can create and maintain a path to water deep in the subsoil, possibly even as far as a water table 20 m below the surface.

3.4.4 - Soil:root interface

The junction between root and soil is more than just two surfaces touching.  It is often marked by a mucilaginous layer, rich in bacteria and fungi, that may be a few hundred micrometres thick, and in which adjacent soil particles are half or even wholly embedded. This is called the rhizosphere (Chapter 4). The hydraulic conductivity of this layer is not known, but there are two properties of this interfacial zone that may influence the flow of water across it.  The first is that extensive gaps may occur within it, which strongly impede the flow.  The second is that any exclusion of ions from the soil solution may result in large increases in the concentrations of these ions at the surface of the roots and this, because of the osmotic effects, would also impede the flow of

Water usually flows from soil to root as liquid, carrying with it dissolved nutrients.  But there may be times when the hydraulic continuity between root and soil is so poor that a substantial proportion of the flow of water is as vapour.  So far, nobody has managed to measure such flows, or even the relative importance of liquid and vapour flow, but their possible importance can be appreciated with the help of a few

The flow of vapour through soil is by diffusion, in contrast to that of liquid water, which moves in bulk.  The consequence of this, and the fact that the vapour pressure of water is close to saturated in moist soil so that gradients in it are slight, is that water flows as vapour many times more slowly, in response to a given difference in water potential, than it does as

There are two main reasons why a root may be in poor hydraulic contact with the soil.  The first is that it may be growing in a pre-existing pore larger than itself, so that it makes only glancing contact with the wall of the pore (see Figure 3.42 above).  The second is that it has shrunk within a pore that it had itself made and which it had previously fitted snugly.  What would induce a root to shrink?  A fall in the water potential of the cortex would do so, for the cells there are thin-walled and separated in part by intercellular gas spaces, so they are likely to buckle.  Observations made in rhizotrons (glass-walled tunnels for observing the behaviour of roots in the field) clearly show a diurnal shrinkage in cotton roots of up to 40% where the roots are growing in large pores, but we still do not know, because they are so difficult to observe, whether roots growing in intimate contact with the soil are similarly prone to shrink.  A few observations made using neutron autoradiography have shown no shrinkage in roots growing in the

Because of the small size of the interfacial zone, it is very difficult to measure concentrations of ions within it.  Rough estimates have been made by carefully separating the roots from the bulk of the soil, leaving only tenacious soil attached, then shaking off this remaining soil and analysing it.  Such analyses do show somewhat higher concentrations (compared with those in the bulk soil) of ions such as sodium that we would expect to be largely excluded by the roots.  But because the gradients in concentration are likely to be very large close to the root, these analyses are hard to assess for their consequences on the water relations of the plant.  The gradients are likely to be especially large when the soil is becoming fairly dry, for the diffusion coefficient for the solutes falls by a few orders of magnitude as the soil dries, so that any excluded solutes will diffuse away from the root surface only very slowly.


3.5 - Water and nutrient transport through roots


Figure 3.46. Transverse section of a mature maize root, showing many layers of cortical cells outside a distinctive suberised endodermis bounding the stele. Late metaxylem vessels with large diameters are the dominant feature of the stele. Note a hypodermal layer underlying the epidermis. Pink Toluidine blue staining characterises suberised cells. (Photograph courtesy A.W.R. Robards)

Water and nutrients travel radially across roots from the soil to the xylem vessels and from there, they travel axially to the shoot. Anatomical and morphological features determine how effective roots are at absorbing and transporting water and nutrients from the soil to the shoot. Root tissue needs to form a low resistance pathway for transport, but also minimise the loss of water to the soil under adverse conditions.

The complexity of the root tissue that water and nutrients must cross is shown below in Figure 3.46. Notable features are the many layers of cortical cells, and the stele containing large and small xylem vessels. The stele is enclosed by the endodermis, a cell layer that contains deposits of suberin and lignin in the anticlinal walls between cells, called a Casparian strip, which blocks diffusion of water and solutes from the cortex into the stele. All roots have an endodermis, except at the tip where they are still growing.

Figure 3.46 also shows a layer of cells beneath the epidermis, which is often suberised and called exodermis. The suberin forms in older roots of many species or under drought conditions. Water and solutes cannot flow across the suberised cells of the exodermis and are restricted to passage cells that are not suberised.

Across the cortex, water could flow along cells walls and through interstitial spaces, the so-called apoplastic route. Alternatively it could enter the epidermal or hypodermal layer and flow symplastically across the cortex. Another alternative is the transmembrane (or transcellular) route. These three pathways are described in the following section. The solutes in the soil solution may also take the same route, or move independently of water.

Radial root conductance to water varies with stage of root development. The radial pathway is considered as more limiting than the axial pathway, and in the younger part of the root where the endodermis and hypodermis are not suberised, the radial conductance to water is higher than when these cell layers are suberised.

Within the stele, water and solutes move symplastically from the endodermis to the adjacent layer of pericycle cells, which are not suberised, so water can then move apoplastically to the xylem vessels. Solutes would most likely stay within the symplasm and be transported via plasmodesmata to the xylem parenchyma cells, from where they are loaded into the xylem vessels. If the xylem vessels are mature and the plant is transpiring, the solutes are carried swiftly to the shoots. If the xylem vessels are immature, the solutes are later released after rupture of the end walls of the immature xylem elements.

3.5.1 - Radial pathways across roots


Figure 3.47. Sketch of a transverse section of a young root showing three proposed pathways of ion and water flow across roots. 1. Apoplastic, via cell walls. 2.  Transmembrane (transcellular), via cytoplasm within cells and crossing membranes via efflux and influx membrane transporters. 3. Symplastic, via cytoplasm all the way, crossing cell walls via plasmodesmata. (Drawing courtesy B. Berger, based on Plett and Moller 2010). Note that at this stage the endodermis is not suberised and has just a Casparian strip (red). Deposition of suberin in the endodermal wall, and the development of an exodermis, will restrict the apoplastic and transcellular pathways.

Water and ions may move through root tissues along one of three routes: symplastic, apoplastic, or transmembrane, the latter also known as the “transcellular pathway” (Figure 3.47). An update on the anatomical and molecular bases of these pathways in Arabidopsis is given by Barberon and Geldner (2014).

The proportion of water flowing through the different pathways undoubtedly varies with the plant’s rate of transpiration.

Apoplastic Flow

Water and soil solutes might move through the cell walls of the cortical cells until the endodermis, where they would cross the plasma membrane of the endodermal cells. Water would cross via aquaporins, and solutes via ion channels or transporters (see Section 3.6). This is possible in roots without a suberised hypodermis and when the endodermis is in the primary stage of development, as described in the following page. Entry of anions to deep layers of the cortex is likely to be restricted by charge repulsion from dissociated, negative carboxyl groups in cell walls (Donnan Free Space). In general, cations also pass through cell walls more readily than anions, particularly if many of the carboxyl groups in cell walls are not occupied by Ca2+ ions. Nonetheless, apoplastic flow of water through roots can sustain large ion fluxes during periods of high transpiration.

It is likely that the bulk of the soil water taken up by the plant moves in the apoplast across the cortex, but that solutes are taken up by epidermal or outer cortical cells and then move in the symplasm across the cortex.

When the endodermis is suberised, water and solutes cannot enter from the apoplast and instead are taken up into the adjacent cortical cells and move via plasmodesmata into the endodermis.

Symplastic Flow

Symplastic flow occurs when water or solutes are taken up by the epidermis or an outer cortical cell, and then subsequently cross the rest of the cortex via plasmodesmata. Plasmodesmata are extensions of the plasma membrane that cross cell walls and provide cytoplasmic continuity between cells, allowing neighbouring cells to communicate and exchange materials more freely. Water and ions can move through the cortex via a series of plasmodesmal connections, thereby remaining in the cytoplasm until reaching the stele. Rate of water and nutrient transport is largely regulated by plasmodesmal resistance, which depends on size and frequency of plasmodesmata in the cell membranes. Alternatively, water and some ions enter vacuoles and are therefore, subject to transport properties of the tonoplast.

Transmembrane (transcellular) Flow

A third possibility is the transmembrane or transcellular route, in which water and nutrients cross membranes, passing repeatedly between symplasm and apoplasm as they are transported from one cell to the next. For nutrients, this form of transport is facilitated by carriers that are distributed in a polar fashion, with influx carriers located on the outer walls and efflux carries on the inner walls. Water transport across cell membranes is via aquaporins for which there is no “rectification”, or intrinsic directionality of transport. They are therefore most likely to be distributed evenly along the plasma membrane.

This pathway may increase resistance to flow of water, but also provides a more effective mechanism for controlling the flow of solutes and water across the root than the purely symplastic route via plasmodesmata when only the first membrane crossing is available to control ion and water transport.

3.5.2 - Variable barriers: endodermis and exodermis

The epidermis and root hairs probably mediate most of the selective uptake of solutes from the soil solution. Nutrients are selectively taken up, and potential toxins excluded.  The cortical cell layers continue this selective uptake if solutes travel apoplastically towards the stele.

The endodermis is the innermost cortical layer that surrounds the central vasculature, and forms a barrier preventing water flow and free diffusion of solutes in the apoplast because of its Casparian strip and suberin lamellae. It thus, prevents soil solution moving into the stele without crossing a membrane, and also prevents water from within the stele being lost back to the soil at night.  Although, in some regions of the endodermis that are opposite xylem poles, water and solutes may bypass the endodermis through unsuberised “passage cells”. The endodermis also functions in structural support for the stele, particularly in drying soil, and minimises shrinkage or swelling of the cells of the stele. Its role in ion selectivity is minor compared to the cells of the cortex and epidermis.

In many plants, the cortical layer under the epidermis (the  hypodermis) develops Casparian bands and suberin lamellae, and develops into the exodermis. This forms another barrier for the apoplastic movement of water and solutes. Just like in the endodermis, some passage cells in the exodermis may be free of suberin and can take up water and solutes.

(a) Endodermis

An endodermis with a Casparian strip is a feature of roots of all land plants from ferns upwards. As the innermost cortical layer that surrounds the central vasculature of roots, the endodermis acts as a barrier to the free diffusion of solutes from the soil into the stele, and the stele into the soil. The protective functions range from efficient water and nutrient transport to defence against soil-borne pathogens. The genes and regulation mechanisms that drive the differentiation of this intricately structured barrier have been reviewed by Geldner (2013).

The development of the endodermis has three stages: the primary stage in which the Casparian strip forms, the second stage when suberin lamellae encases the entire endodermal cell, and the tertiary stage when the inner tangential walls thicken and a layer of cellulose is deposited over the suberin lamellae.

The Casparian strip is made of lignin and suberin and deposited as a ring in the radial walls of the endodermal cells, like a hoop around a barrel of beer. It reaches from the plasma membrane to the outermost part of the wall and adjoins adjacent cells so there are no air spaces between the cells at this point. It therefore blocks the flow of water through the cell walls. It differentiates as root cells mature about 5-10 mm from the tip, and so an entirely apoplastic pathway from soil to central stele can occur only in very young root parts or at sites where the Casparian strip is disrupted such as site of lateral root development, which starts at the pericycle, the cell layer beneath the endodermis.

In the first stage of endodermis development, the Casparian strip in the primary wall prevents flow of water and solutes through the wall from inner cortex to stele. Any solutes remaining in the apoplastic water must enter the endodermis via membrane transporters. Water in the apoplast must enter the endodermis via aquaporins as illustrated in the lower pathway shown in Figure 3. 50. 


Figure 3.50. The alternative pathways for water and nutrient solute flow across roots with the endodermis at Stage 1 of development, with a primary wall with Casparian strip (c). The apoplastic pathway is blocked at this point, and water, along with any nutrients still in the walls, enters the endodermal cells via membrane transporters (lower panel). If the hypodermis also has a Casparian strip, water and nutrients must also enter via membrane transporters. Alternatively, water and nutrients can move via the symplastic (dotted red line, lower panel) or transmembrane pathways (upper panel). (Diagram courtesy H. Bramley)

In the second stage of development, the endodermis becomes suberised as the secondary walls develop, and suberin lamellae are formed all over the cell, underneath the primary wall that contains the Casparian strip. This occurs at varying distances from the root tip depending on species and is often induced under drought to form in younger parts of the root.

Suberin is a hydrophobic polymer, deposited in the secondary cell wall in lamellae. It therefore seals off the plasma membrane from solutes, as water and ion channels are sealed. So the transcellular and apoplastic pathways are curtailed, and water and solutes enter the endodermis only through plasmodesmata from neighbouring cells (Figure 3.51). Any function in selective control of particular nutrients into or out of the endodermal cell through the suberin lamellae is unclear.


Figure 3.51. Alternative pathways for water and nutrient flow across roots with a suberised (Stage 2) endodermis. A complete layer of suberin around the endodermis means that water and solutes can enter only via plasmodesmata from the inner cortical cells. (Diagram courtesy H. Bramley)

Some endodermal cells remain at the primary stage even late in root development, and are called passage cells.

Stage 3 of the endodermis involves further deposition of cellulosic wall material, further impeding flow of solution through walls.

(b) Exodermis

Exodermis is the name given to a hypodermis with Casparian strips and suberised lamellae.


Roots of most species form an exodermis with time. First, a Casparian band is laid down in the primary wall of the hypodermis, then all the wall is suberised especially the inner wall, as for the endodermis. Some cells in this hypodermal layer (‘passage cells’) remain unsuberised.

The development of the exodermis is very common in the plant kingdom (Perumalla et al. 1990). In a study of 180 angiosperm species, the great majority (89 %) showed a clearly suberised exodermis with Casparian strip. It was found in roots of primitive and advanced plant families, from hydrophytic, mesophytic and xerophytic habitats, but was lacking in some Poaceae. It was notably absent in oat, barley and wheat (Perumalla et al. 1990), but generalizations within cereals cannot be made as subsequently maize was found to have an exodermis (Hose et al. 2001),

The Casparian band can develop close to the root tip. For example, in aeroponcially grown maize a complete exodermal layer formed 30 mm above the root tip. In roots elongating more slowly due to abiotic stress or low temperature, it can be found closer to the tip. The extent at which apoplastic barriers form depends on the stage of development of the root system and also the habitat: drought, waterlogging, salinity, nutrient deficiency or toxicity may strongly influence the degree of suberisation (Hose et al. 2001).

Permeability to water and solutes


Figure 3.54. Schematic diagram of a longitudinal section through a root indicating the stage at which the critical structures in radial and axial transport of water develop. Not to scale. (Diagram courtesy H. Bramley)

When an exodermis forms, it also imposes a restriction to radial transport. Complete layers of suberin constrain water and solute flow only via plasmodesmata from the epidermis and inner cortical cells.  Whether or not it can be considered as a barrier depends on the degree of suberisation and the number of passage cells within it. Its properties as a barrier are variable (Hose et al. 2001).

The exodermis represents a resistance to the radial flow of both water and solutes, much like the endodermis in Stage 1 development. It restricts radial apoplasmic movement and may also restrict transmembrane transport of nutrients. Exodermal layers become functionally mature 20–120 mm from the apex, where lateral roots are initiated, and therefore constitute a barrier to apoplastic ion flow only in root zones where an endodermis is already present. In a similar way to the endodermis, maturation of an exodermis involves further deposition of suberin and cellulosic wall material, further impeding flow of solution through walls.

The exodermis is not totally impermeable to water or nutrients and may have a differential selectivity that varies with the environment. How much the root is sealed off depends on the type of exodermis: (1) uniform exodermis where the cells are uniform in shape (suberin deposition is patchy and develops late) or (2) dimorphic exodermis, which consists of long and short passage cells (the former of which are suberised). Suberin lamellae enclosing the long cells disrupt the cytoplasmic continuity through plasmodesmata and the cell dies, but suberin lamellae in the uniform-type exodermis do not affect plasmodesmata. Individual passage cells allow passage of solutes and water via uptake carriers, not via the apoplast as flow there is blocked by the Casparian strip. They have an active role in ion uptake and often become the only plasmalemma facing the soil solution especially when the epidermis dies. Some families (e.g. irises) have large numbers of passage cells while others have very few.

The diagram of a root to the right (Figure 3.54) indicates the stage of development at which the various structures are functional.

The anatomy of roots and the alternative pathways of water and solute flow across roots and into the xylem indicates that the relationship between water and nutrient flow to the shoots could be quite complex. Water may take different pathways than nutrients. The distribution of water in the different pathways depends not only on the age of the root, but may also vary with the flux of water moving through, that is, the rate of transpiration.

3.5.3 - Relation between water and nutrient flux


Figure 3.55. Effect of volume flux (sap flow rate) on K+ concentration and K+ flux in xylem sap from barley. Arrow indicates the external K+ concentration, and dotted line indicates the range of sap flow rate for which  K+ in the sap was at a higher concentration than the external solution. (Munns, 1985)


The various pathways taken by water and nutrients, and the environmental factors that influence them, are not well understood. The fundamental processes governing the relationship between water and nutrient flow through roots are complex. One thing is clear: nutrients do not move passively with the transpiration stream. But neither is their movement entirely independent of it.

This section looks at the relation between water and solute fluxes to the shoot: how the rate of transpiration affects solute concentrations in the xylem, and how active solute transport affects water flow rates at night (root pressure).

(a) Effect of transpiration on solute flux in the xylem

Solute fluxes in the xylem usually respond to changes in transpiration rate, although the relationship is not proportional. The figure below (Figure 3.55) indicates an apparently strong relation between water flux and  K+ flux, but it must be noted that the flux of  K+ is already high at very low flow rates, increases linearly with moderate increases in flow rate, but then tapers off at high flow rates.

A similar relation has been found for other nutrients like nitrate.

The importance of transpiration in carrying nutrients to the shoot has long been debated. However, experimental evidence showed that transpiration was not necessary to get nutrients to the shoot, as growth rates and net nutrient transport rates were unaffected by humidity and other environmental conditions that would reduce transpiration (Tanner and Beevers 2001). Transpiration was not a prerequisite for long-distance transport of nutrients, as root pressure (see below) in the absence of transpiration can supply the shoot with the required nutrients.

(b) Effect of transpiration on solute concentration in the xylem


Figure 3.56. Effect of transpiration rate on osmotic pressure (left hand axis) and the corresponding solute concentration (right hand axis) of xylem sap of young barley plants. Transpiration rates were imposed by varying vapour pressure deficit around the shoots and xylem sap was sampled by applying sufficient pressure to roots to cause a cut leaf tip to bleed. Arrow indicates the external osmotic pressure. (Munns and  Passioura, 1984)

Transpiration rate (the volume flux in the xylem) has a marked effect on the concentration of solutes in the xylem sap. Plants which are transpiring rapidly have a low concentration of nutrients in their xylem sap compared with slowly transpiring plants.  This relationship between concentration and volume flux holds for most nutrients, see example with K+ (Figure 3.55 above) in which increasing flow rates decreased the concentration of  K+ in xylem sap.

The data also shows that an apoplastic or diffusive uptake of K+ is not important because the concentration of K+ in the xylem was greater than the external K+ over a range of volume fluxes which produced an increasing flux (Figure 3.54). These data strongly suggest that ion concentrations in xylem sap are a result of either a single variable pump, or more likely of several sequential processes. The composition of the transpiration stream could be modified after the first passage across a membrane as it flows towards the stele or upwards through the stele by either active or passive ion fluxes from cortical or xylem parenchyma cells.  K+ could be pumped in (or diffuse in) at rates which increase with increasing volume flux.

Similar relationships occur between most solutes in the sap and transpiration rate, as shown in Figure 3.56.

This shows how much influence the transpiration rate has on the concentration of solutes in the xylem, and that concentrations in sap collected from exuding cut stumps are not typical of concentrations in transpiring plants.

(c) Root pressure

Plants that have had shoots removed have very concentrated xylem fluid, which exudes from the cut stumps under positive hydrostatic pressure from the roots (‘root pressure’). Root pressure is also responsible for the droplets of water seen on the margins of guttating leaves early in the morning (Figure 3.57).


Figure 3.57. Guttation droplets from a eucalypt leaf, E. tetragona. (Photograph courtesy C. Hooper)

This occurs because nutrient uptake is an active process, independent of water uptake from the soil, and continues during the night – the rate of nutrient uptake varies little over 24 h. When nutrients are pumped into the stele, water flows in by osmosis, and the pressure builds up. Positive pressures of 30-300 kPa can be achieved in this way.

The Casparian strip and the suberisation of the endodermis is important as it provides a barrier to prevent back-flow of water and also structural support so that the root can contain a positive pressure at night. The pathways of nutrient and water flow across the root cortex may change at night, for instance the apoplastic pathway may be of lesser importance at low rates of transpiration. At zero transpiration induced by a root pressure probe and when the shoot is removed, pathways may be different again.

Root pressure is not just a phenomenon, it is an essential process responsible for moving nutrients to the shoot during the night when transpiration is low. It may also have a function in dissolving gas bubbles that might have caused cavitation in the xylem during the day.

3.6 - Membrane transport of water and ions


Figure 3.61. Guard cells are structured so that high turgor pressure opens the stomatal pore, and low turgor pressure closes the pore. Left (Lower image), Partially open Tradescantia virginiana stoma with 0.2 MPa guard cell turgor, as measured directly with a guard cell pressure probe (shown). Right (upper image) same stoma showing the aperture almost fully open, with 3.6 MPa guard cell turgor. (Images and data courtesy P.  Franks).

Water uptake by cells is driven by solute uptake. Osmotically-driven water uptake generates the turgor pressure needed for maintenance of cell volume, and for specific cell functions that depend on controlled changes in turgor.  The rapid influxes or effluxes of water that cause rapid changes in cell volume in key cells are brought about by ion influxes or effluxes.

Rapid changes in turgor cause the swelling or shrinking of guard cells in leaves that controls the opening and closing of stomata. The cell walls of guard cells are structured so that high turgor pressure pushes them apart and opens the stomatal pore. Low turgor pressure allows them to collapse and close the pore (Figure 3.61). Influxes or effluxes of K+ along with accompanying anions causes the osmotically-driven water uptake or loss

Whole leaves or parts of leaves can move quickly. Changes in the turgor of a group of cells at the base of leaves, the pulvinus, cause leaves to fold quickly in response to touch, as in the ‘sensitive plant’ Mimosa pudica, (Figure 3.62)


Figure 3.62. Seismonastic movement of pinnae and pinnules in leaves of the sensitive plant (Mimosa sensitiva) (a) before and after touch stimulation. (Photographs courtesy J.H. Palmer)

Turgor changes also change the curvature of hairs of insectivorous plants. In the case of the Venus fly trap, sensory hairs coupled to an electrical signalling system require stimulation at least twice within a 30 s period (Simons 1992). This appears to allow the plant to discriminate single pieces of debris from an insect crawling within the trap. Most seismonastic movements result from the explosive loss of water from turgid ‘motor’ cells, causing the cells temporarily to collapse and inducing very quick curvature in the organ where they are located.

Similar mechanism causes the slower folding of leaves at night into special positions to reduce heat loss, as in the prayer plant. This also occurs in many legumes. Charles Darwin measured the folding of leaves at nightfall in white clover, and wrote: “The two lateral leaflets will be seen in the evening to twist and approach each other, until their surfaces come into contact, and they bend downwards. This requires a considerable amount of torsion in the pulvinus. The terminal leaflet merely rises up without any twisting and bends over until it forms a roof” (Figure 3.63).


Figure 3.63 The nictitropic movements of leaves of white clover (Trifolium repens) from daytime (A) to ‘sleeping’ position (B). (Charles Darwin, The Power of Movement in Plants, 1881). (Diagram courtesy R. Purdie)

Movement of some plant organs can be staggeringly fast. The firing of the reproductive structure (“column”) of trigger plants (Stylidium genus) in response to the landing of a pollinating insect may take only 15 msec (Figure 3.64).


Figure 3.64 Sequence of superimposed images captures the flower column of a trigger plant (Stylidium crassifolium) as it ‘fires’ in response to a physical stimulus (insect landing, seen on upper right). The column rotates through 200° from a ‘cocked position’ to a relaxed position in 20 ms (photographs taken at 2 ms intervals). (Findlay and Findlay 1975)

The kinetic energy manifested in this rapid firing is derived from events controlled at a membrane level. Ions transported into specialised cells cause hydrostatic (turgor) pressure to develop which is suddenly dissipated following mechanical stimulation

Water transport into a cell anywhere in the plant is governed by solute uptake which in turn is governed by the permeability of membranes to water as well as solutes.

3.6.1 - Diffusion and permeability

This section covers diffusion of molecules, and permeability of cell membranes, essential to the process of osmosis. Cell membranes are bilayers of phospholipids in which transport proteins are embedded (Figure 3.65).


Figure 3.65 Diagram of a cell membrane with transport proteins embedded in the phospholiped bilayers. (Image courtesy M. Hrmova)

A plant membrane is often described as semi-permeable, meaning permeable to water but not biological solutes. However the membrane is not 100% permeable to water, as water can enter cells only by being transported through aquaporins, neither is it 100% impermeable to solutes, as solutes can slowly permeate the membrane particularly through specific transport proteins.

Unrestricted movement of water relative to solutes is the basis of osmosis, and in plants the generation of turgor pressure, \(P\). The principles of diffusion and selectivity, which are used to describe differential rates of molecular movement, provide a physical rationale for osmosis.

(a) Diffusion

In a homogeneous medium, net movement of molecules down their concentration gradient is described by Fick’s First Law of diffusion. The molecule and medium may be a solute in water, a gas in air or a molecule within the lipid bilayer. Fick’s Law holds when the medium is homogeneous in all respects except for the concentration of the molecule. If there was an electric field or a pressure gradient then Fick’s Law may not apply. Considering the case of a solute in water, say, sugar, Fick’s Law states that net movement of this solute, also called the net flux \(J_s\), is proportional to the concentration gradient of the solute \( \Delta C_s / \Delta x \):

\[ J_s = -D_s \frac{\Delta C_s}{\Delta x} \tag{12}\]

The diffusion coefficient (\(D_s\)) is a constant of proportionality between flux, \(J_s\), and concentration gradient (mol m-4), where solute concentration (\(C_s\), mol m-3) varies over a distance (\( \Delta x\), m). Flux is measured as moles of solute crossing a unit area per unit time (mol m–2 s–1), so \(D_s\) has the units m2 s–1. \(D_s\) has a unique value for a particular solute in water which would be quite different from \(D_s\) for the same solute in another medium, for example the oily interior of a lipid membrane.

Across a membrane, the movement of a molecule from one solution to another can be described by Fick’s Law applied to each phase (solution 1–membrane–solution 2). However, flux across the membrane also depends on the ability of the molecule to cross boundaries (i.e. to partition) from solution into the hydrophobic membrane and then from the membrane back into solution. Another difficulty is that the thickness of membranes is relatively undefined and we need to know this for Fick’s equation above (\( \Delta x \)). The two solutions might differ in pressure and voltage and these can change steeply across a membrane; however, if for simplicity we consider a neutral solute at low concentration, these factors are not relevant (see below for charged molecules). A practical quantitative description of the flux of neutral molecules across membranes uses an expression intuitively related to Fick’s Law stating that flux across a membrane (\( J_s \)) of a neutral molecule is proportional to the difference in concentration \(\Delta C_s\):

\[ J_s = P_s \Delta C_s \tag{13}\]


Figure 3.66. The range of permeability coefficients for various ions, solutes and water in plant membranes (vertical bars) and artificial phospholipids (arrows). Note that the permeability of ions as they cross plant membranes is higher than through the artificial lipid bilayer.

The constant of proportionality in this case is the permeability coefficient (\( P_s \)), expressed in m s–1. When \( P_s \) is large, solutes will diffuse rapidly across a membrane under a given concentration gradient. \( P_s \) embodies several factors: partitioning between solution and membrane, membrane thickness and diffusion coefficient of the solute in the membrane which may be largely depend on specific transporters. It can be used to compare different membranes and to compare treatments that might alter the ability of a solute to move across the membrane.

Note that Equation 13 assumes that the concentration gradient is only across the membrane, and that when the permeability coefficient is measured, concentration gradients leading to diffusion in solutions adjacent to the membrane will not be significant. If the two solutions are stirred rapidly then this will help to justify this assumption. However, there is always an unstirred layer adjacent to the membrane through which diffusion occurs, and for molecules that can permeate the membrane very rapidly the unstirred layer can be a problem for the correct measurement of permeability.

(b) Permeabilities

Solute movement across membranes can be across the lipid phase of the membrance, depending on size, charge and polarity, and it can be assisted by transport proteins embedded in the membrane.

The range of permeability coefficients for various ions, solutes and water in plant membranes is shown in Figure 3.66, along with the permeability of artificial phospholipid bilayers. The permeability of ions as they cross membranes is higher than that through the artificial lipid bilayer, especially for potassium, indicating the presence of specialised permeation mechanisms, ion transporters, in plant membranes. Water permeabilities are high in both plant and artificial membranes but can range over an order of magnitude in plant membranes. This variability may be partially accounted for by the activity of aquaporins.

A comparison of artificial lipid membranes with biological membranes supports this notion because it shows that many molecules and ions permeate biological membranes much faster than would be predicted on the basis of oil solubility and size (Figure 3.66). For these solutes there are transport proteins in biological membranes that increase solute permeability.

(c) Reflection coefficient - water versus solute permeability

Plant membranes are ideally semipermeable, that is, water permeability is much larger than solute permeability.

The degree of semi-permeability that a membrane shows for a particular solute is measured as the reflection coefficient, \( \sigma \):

\[ \sigma = 1 - \frac{\text{Solute Permeability}}{\text{Water Permeability}} \tag{15}\]


Figure 3.67. Turgor pressure (\( P \)) in a Tradescantia virginiana epidermal cell as a function of time after the external osmotic pressure was changed with different test solutes. Measurements were made with a pressure probe. (Tyerman and Steudle 1982)

If a plant cell or an epidermal strip is bathed in solution, the reflection coefficient for a particular solute can also be considered as the ratio of the effective osmotic pressure versus the actual osmotic pressure in the bathing solution.

The reflection coefficient usually ranges between zero and one, being zero for molecules with properties similar to water, like methanol, to one for large non-polar molecules like sucrose.

Figure 3.67 shows turgor pressure (\( P \)) in a Tradescantia virginiana epidermal cell as a function of time after the external osmotic pressure was changed with different test solutes. The initial decrease in \( P \) is due to water flow out of the cell and is larger for solutes with a reflection coefficient near one (sucrose and urea). Propanol induces no drop in \( P \), indicating that its reflection coefficient is zero. Subsequent increase in \( P \) is due to penetration of particular solutes such as alcohol across the cell membrane. Water flows osmotically with the solute thereby increasing \( P \) to its original value. Removing solutes reverses osmotic effects. That is, a decrease in \( P \) follows the initial inflow of water as solutes (e.g. alcohols) diffuse out of cells.

The pressure probe apparatus is illustrated in Figure 3.68(a).


Figure 3.68 (a) A miniaturised pressure probe. An oil-filled capillary of about 1 µm diameter is inserted into a cell whose turgor pressure (\( P \)) is transmitted through the oil to a miniature pressure transducer. The voltage output of the transducer is proportional to \( P \). A metal plunger acting as a piston can be used via remote control to vary cell \( P \).

Using the pressure probe to measure turgor pressure, \( P \), the membrane is found to be ideally semipermeable for sucrose (\( \sigma = 1 \)); that is, the membrane almost totally ‘reflects’ sucrose. Over long periods, sucrose is taken up slowly but permeability relative to water is negligible. In this case, the change in \( P \) would be equivalent to the change in \( \pi \). If \( \sigma \) is near zero, then water and the solute (say, propanol) are equivalent in terms of permeability. No change in \( P \) can be generated across a cell wall if \( \sigma \) is zero.

3.6.2 - Chemical potential

Diffusion of neutral molecules at low concentrations is driven by differences in concentrations across membranes, as explained above. There are other forces that may influence solute diffusion, including the voltage gradient when considering movement of charged molecules (ions) and the hydrostatic pressure when considering movement of highly concentrated molecules (such as water in solutions). These forces can be added to give the total potential energy of a particular molecule (\( \mu_s \)) relative to a reference value (\( \mu_s^* \)):

\[ \mu_s = \mu_s^* + RT\ln C_s + z_s FE + V_sP \tag{16}\]

Gravitational potential energy could also be added to this equation if we were to examine the total potential over a substantial height difference, but for movement of molecules across membranes this is not relevant.

The concentration term (\( RT\ln C_s \)) is a measure of the effect on chemical potential of the concentration of solutes (actually the activity of the solutes which is usually somewhat less than total concentration). The gas constant, \(R\) (8.314 joules mol–1K–1), and absolute temperature, \(T\) (in degrees Kelvin, which is equals 273 plus temperature in degrees Celsius), account for the effects of temperature on chemical potential.

Incidentally, from this term the well-known van‘t Hoff relation is derived for osmotic pressure \(\pi\), as given at the beginning of this chapter:

\[ \pi = RTC \tag{1} \]


Figure 3.69. Illustration of how electrical and concentration terms contribute to electrochemical potential of ions. Calcium (top) commonly tends to leak into cells and must be pumped out whereas chloride tends to leak out and must be pumped in to be accumulated.

where \(R\) is the gas constant (8.31 joules mol–1K–1), \(T\) is the absolute temperature in degrees Kelvin (273 plus degrees Celsius), and \( C \) is the solute concentration (Osmoles L-1). At 25 ºC, \(RT\) equals 2.48 liter-MPa per mole, and (\( \pi \)) is in units of MPa. Hence a concentration of 200 mOsmoles L-1 has an osmotic pressure of 0.5 MPa.

The electrical term (\( z_s FE \) ) is a measure of the effect of voltage (\( E \) ) on chemical potential. The charge on a solute (\( z \)) determines whether an ion is repelled or attracted by a particular voltage. Electrical charge and concentration are related by the Faraday constant (\( F\) ) which is 96,490 coulombs mol–1. The electrical and concentration terms form the basis of the Nernst equation (see below).

The pressure term measures the effect of hydrostatic pressure on chemical potential, where \(P = \text{Pressure}\) and \( \overline{V}_s \) is the partial molar volume of the solute.

Molecules diffuse across a membrane down a chemical potential gradient, that is, from higher to lower chemical potential. Diffusion continues until the difference in chemical potential equals zero, when equilibrium is reached. The direction of a chemical potential gradient relative to transport of a molecule across that membrane is important because it indicates whether energy is or is not added to make transport proceed (Figure 3.69). Osmotic ‘engines’ must actively pump solutes against a chemical potential gradient across membranes to generate \(P\) in a cell. Sometimes ions move against a concentration gradient even when the flux is entirely passive (no energy input) because the voltage term dominates the concentration term in Equation 16. In this case, ions flow according to gradients in electrical and total chemical potential. For this reason, the chemical potential of ions is best referred to as the electrochemical potential.

3.6.3 - Ions, charge and membrane voltages

Ions such as potassium and chloride (K+ and Cl) are major osmotic solutes in plant cells. Deficiency of either of these two nutrients can increase a plant’s susceptibility to wilting. Most other inorganic nutrients are acquired as ions and some major organic metabolites involved in photosynthesis and nitrogen fixation bear a charge at physiological pH. For example, malic acid is a four-carbon organic acid that dissociates to the divalent malate anion at neutral pH. Calcium (Ca2+) fluxes across cell membranes are involved in cell signalling and although not osmotically significant they play a crucial role in the way cells communicate and self-regulate. Finally, some ions are used to store energy but need not occur at osmotically significant concentrations. Cell membranes from all kingdoms use hydrogen (H+) ions (protons) in one way or another to store energy that can be used to move other ions or to manufacture ATP (Chapter 2). The highest concentration of H+ that occurs is only a few millimoles per litre yet H+ plays a central role in energy metabolism.

To understand ion movement across membranes, two crucial points must be understood: (1) ionic fluxes alter and at the same time are determined by voltage across the membrane; (2) in all solutions bounded by cell membranes, the number of negative charges is balanced by the number of positive charges. Membrane potential is attributable to a minute amount of charge imbalance that occurs on membrane surfaces. So at constant membrane potential the flux of positive ions across a membrane must balance the flux of negative ions. Most biological membranes have a capacitance of about 1 microFarad cm–2 which means that to alter membrane voltage by 0.1 V, the membrane need only acquire or lose about 1 pmol of univalent ion cm–2 of membrane. A univalent ion is one with a single positive (e.g. K+) or negative (e.g. Cl) atomic charge. In a plant cell of about 650 pL, this represents a change in charge averaged over the entire cell volume of 12 nmol L–1!

The membrane voltage or membrane potential difference, as it is sometimes called, can be measured by inserting a fine capillary electrode into a plant cell (Figure 3.68b). Membrane voltage is measured with respect to solution bathing the cell and in most plant cells the voltage is negative across the plasma membrane. That is, the cytoplasm has a charge of –0.1 to –0.3 V (–100 to –300 mV) at steady state with occasional transients that may give the membrane a positive voltage. The tonoplast membrane that surrounds the central vacuole is generally 20 to 40 mV more positive than the cytoplasm (still negative with respect to the outside medium).


Figure 3.68. (a) Techniques employing fine glass capillaries to probe plant cells. (b) A probe for measuring membrane voltage. The capillary is filled with 1 mol L-1 KCl and connected to a silver/silver chloride electrode that acts as an interface between solution voltage and input to the amplifier. A voltage is always measured with respect to a reference (in this case, a bath electrode). The headstage amplifier is close to the electrodes to minimise noise.

Cell membrane voltages can be affected by ion pumps, diffusion potential and fixed charges on either side of the membrane.

Special mention needs to be made of one such fixed charge which arises from galacturonic acid residues in cell walls. Although cations move to neutralise this fixed negative change, there is still a net negative potential associated with cell walls (Donnan potential). In spite of being external to the plasma membrane, the Donnan potential is in series with it and probably adds to what we measure as the membrane potential with electrodes.

Most charge on macromolecules in the cytoplasm is also negative (e.g. nucleic acids, proteins) and because of their size it can be regarded as a fixed negative charge. This has consequences on the water relations of the cytoplasm in that they exert a significant osmotic potential, even though not in solution, as do the clay particles in soil (Passioura 1980).

Different ions have different permeabilities in membranes. Potassium, for example, is usually the most permeable ion, entering under most conditions about 10 to 100 times faster than Cl Since ions diffuse at different rates across membranes, a slight charge imbalance occurs and gives rise to a membrane voltage (Figure 3.69). This voltage in turn slows down movement of the rapidly moving ion so that the counter-ion catches up. The result is that when net charge balance is achieved, a diffusion potential has developed that is a function of the permeabilities (\( P_\text{ion} \)) of all diffusible ions present and concentrations of each ion in each compartment.


Figure 3.70. How a diffusion potential develops through differential movement of an ion across a membrane that is permeable to K+ but not Cl-.  The left hand compartment (representing the cytoplasm) has the higher concentration of K+ and Cl-, as indicated by the size of the letters. The right hand compartment represents the apoplast (cell wall). Initially (a), a minute amount of K+ crosses the membrane along its concentration gradient, and creates a positive charge in the right-hand compartment as K+ concentration there rises above Cl- concentration. At equilibrium (b), a diffusion potential is established, and further movement of K+ is prevented. Concentrations never equalise on both sides because K+ is the only species able to move through the membrane.

The Goldman equation describes this phenomenon and gives the membrane voltage (\( \Delta E \) ) that would develop due to diffusion of ions. The Goldman equation for the ions that mostly determine this diffusion potential (K+, Na+ and Cl) is given by:

\[ \Delta E = \frac{RT}{F} \ln \frac{P^{}_K C_K^o + P^{}_{Na}C_{Na}^o + P^{}_{Cl}C_{Cl}^i}{P^{}_K C_K^i + P^{}_{Na}C_{Na}^i + P^{}_{Cl}C_{Cl}^o} \tag{17}\]

The superscripts refer to the inside (i) or outside (o) of the membrane and \( R \), \( T \), \( F \) and \( C \) are defined elsewhere (Equation 4.5). Note that the concentration terms for Cl are reversed in the numerator and denominator compared to the cations. This is because Cl is the only anion represented. Many texts do not include H+ in the Goldman equation because, in spite of high permeability of H+, diffusion of H+ is unlikely to have a strong effect on ΔE at such low (micromolar) concentrations. However, membrane potential is occasionally dominated by the diffusion of H+, indicating that H+ permeability must be exceedingly high. For example, local variations in pH cause alkaline bands to form on Chara corallina cells and in the leaves of aquatic plants at high pH.

The Nernst equation

When one ion has a very high permeability compared to all other ions in the system the membrane will behave as an ion-sensitive electrode for that ion (e.g. Figure 4.7). A pH electrode which is sensitive to H+ flux across a glass membrane serves as an analogy. In the case of a single ion, the Goldman equation can be reduced to the simpler Nernst equation that yields the equilibrium membrane potential which would develop for a particular concentration gradient across a membrane.

\[ \Delta E = \frac{RT}{zF} \ln \frac{C_o}{C_i} \tag{18}\]

where \( R \) and \( T \) are the gas constant and temperature (degrees Kelvin) and \( F \) is the Faraday constant. Typical charges on ions (\( z \)) would be –1 for Cl-, +1 for K+) and so on. This term in the Nernst equation gives the correct sign for the calculated membrane potential.

The Nernst equation is routinely used by electro-physiologists to calculate the equilibrium potential for each ion. Theoretical equilibrium potentials can then be compared with the actual membrane potential in order to decide whether the membrane is highly permeable to one particular ion. For example, in many plant cells there are K+ channels that open under particular circumstances. When this occurs, the membrane becomes highly permeable to K+ and the measured membrane potential very nearly equals the Nernst potential for K+. The Nernst equation can also be used as a guide in deciding whether there is active transport through a membrane. For example, when the measured membrane potential is less negative than the most negative Nernst potential, an electrogenic pump must be engaged for K+ to enter the cell (Table 4.1). If the membrane potential is less negative than the Nernst potential and if a K+ channel were open then K+ would leak out of the cell. For K+ uptake to occur with such a gradient for passive efflux then energy would need to be generated.

Equation 4.7 can be rewritten with constants solved and log10 substituted for the natural logarithm. This yields a useful form as follows,

\[ \Delta E = \frac{58}{z} \log_{10} \frac{C_o}{C_i} \tag{19} \]

showing that 10-fold differences in concentration across a membrane are maintained by a 58 mV charge separation for monovalent ions. For example, -58 mV will keep K+ concentrations 10 times higher inside a cell than in the external medium and Cl concentrations 10 times lower. Plasma membranes are normally about -116 mV, which would keep K+ concentrations inside a cell 100 times higher and Cl- concentrations 100 times lower than in the external solution.

Internal membranes have a different electrical potential, the mitochondria being more negative than the plasma membrane (around -180 mV) and the chloroplast and tonoplast being slightly positive (around +50 mV).

The concentration of an ion inside a cell membrane (\( C_i \)) that would occur at equilibrium for any \( C_o \) and \( \Delta E \) can be calculated by rearranging the above equation as:

\[ \log_{10} C_i = \log_{10} C_o - \frac{z\Delta E}{58} \tag{20} \]

remembering that \( z \) and \( \Delta E \) can be positive or negative, depending on the ion and the particular cell membrane being considered.

3.6.4 - Aquaporins (water channels)

Plant and animal membranes have much higher permeability to water than can be explained by diffusion rates through a lipid bilayer. Furthermore, the activation energy for diffusion of water across a plant membrane is lower than would be expected across a lipid bilayer, where water has to overcome the high-energy barrier of partitioning into a very hydrophobic oily layer. Some reports put the activation energy for water flow across membranes as low as the value for free diffusion of water. In other words, water enters the membrane about as readily as it diffuses through a solution. This suggests that water is moving across the membrane through a pathway other than the lipid, perhaps some kind of water pore or water channel. Since the discovery of water channel proteins in animal cell membranes, molecular biologists discovered that similar proteins exist in plants.

Water channels, like ion channels, are proteins embedded in membranes that facilitate the passive transport (non-energised flow) of water or ions down their respective energy gradients. Movement of a solute or water through these transport proteins is not coupled to movement of any other solute, and does not require ATP. The proteins that facilitate passive transport are diverse; some are specific for particular ions and allow high transport rates per protein molecule (ion channels), some are specific for water (water channels or aquaporins) and some are specific for neutral solutes and may have slower transport rates per protein molecule.

Why are there water channels in membranes when the lipid itself is already somewhat permeable to water? There are several rationales for the presence of water channels in plant membranes. One is that specialised transport proteins can control water flow. That is, a water channel protein may be turned on and off, for example by phosphorylation, while water permeability of the lipid is essentially constant. In animal cells, such as in the kidney, water channels are controlled by antidiuretic hormones. Plant hormones could also influence the function of water channels. A second rationale for the presence of water channnels is to balance water flow and prevent bottlenecks. In the root, water channels are most abundent in the endodermis and inner stele where water flow across membranes is rapid. 

The approach to studying water channels has been to inject genetic material from plants into Xenopus oocytes (a particular type of frog’s egg). The Xenopus oocyte is particularly useful because it is large, enabling observations of cell response to foreign proteins. It is one of several expression systems along with Chara (giant algal cells) and yeast cells. cDNA arising from screens of cDNA libraries can be injected into the Xenopus nucleus, or poly (A)+-RNA can be injected into the oocyte cytoplasm where it is translated. Plant water transport proteins expressed in the oocyte plasma membrane result in physiological changes; for example, the oocyte swells rapidly when the external osmotic pressure in the bathing medium is lowered (Figure 3.71a). The first plant aquaporin gTIP (Tonoplast Integral Protein) that was discovered occurs in the tonoplast and probably accounts for its high water permeability. Provided that the increase in water permeability is not a consequence of some other change or a side effect of other types of transport, it can be concluded that the protein catalyses transport of water across membranes.

Figure 3.71 Evidence for presence of aquaporins (water channels) in plant membranes. (a) Change in volume of Xenopus oocytes injected with two TIP proteins after lowering osmotic pressure of the external medium (Maurel et al. 1995). (b) Sensitivity of two TIP proteins expressed in Xenopus oocytes to mercuric chloride (HgCl2), a general inhibitor of aquaporins (Daniels et al. 1996). (c) Inhibitory effect of HgCl2 on hydraulic conductivity of the freshwater alga Chara corallina measured with a pressure probe (Schütz and Tyerman 1997).

Water channels can be inhibied by mercuric chloride when expressed in Xenopus occytes (Figure 3.71b), and also in plant cells (Figure 3.71c). Inhibition is reversed by applying mercaptoethanol to block the action of HgCl2. Under mercury inhibition, the activation energy of water flow increases markedly indicating that water flow is now restricted to diffusional flow across the lipid bilayer, that is, aquaporins are blocked.

Plants have many aquaporin genes. For example, Arabidopsis thaliana has 35 and rice (Oryza sativa) has 33. Proteins encoded by aquaporin genes are localized in the plasma membrane, tonoplast and endo membranes and classed as Plasma membrane Intrinsic Proteins (PIPs), Tonoplast Intrinsic Proteins (TIPs), Nodulin26-like Intrinsic Proteins (NIPS), Small basic Intrinsic Proteins (SIPs) or X Intrinsic Proteins (XIPs) (Luu and Maurel 2013). Within membranes, clustering of PIPs in membrane rafts has been observed, and there is variation in the lateral mobility for different aquaporins. Of the many aquaporin proteins in plants, some transport water only (up to 109 molecules per second) and others are permeable to a range of neutral solutes such as gases (carbon dioxide, ammonia), metalloids (boron, silicon, arsenic), or reactive oxygen species (hydrogen peroxide). The transcription, translation, trafficking, and gating of PIPs are regulated by environmental and developmental factors involving signalling molecules, phytohormones and the circadian clock (Chaumont and Tyerman 2014). The transcription, translation, trafficking, and gating of PIPs are regulated by environmental and developmental factors involving signalling molecules, phytohormones and the circadian clock (Chaumont and Tyerman 2014).

When PIP genes are transcribed, their mRNA is translated in the rough endoplasmic reticulum (ER), and the proteins targeted to the plasma membrane. PIPs belonging to the PIP2 group form homo-oligomers, or hetero-oligomers by associating with PIP1 isoforms. Some PIP2s contain a diacidic motif in their N terminus that acts as an ER export signal. It may be recognized by Sec24 which is the main “cargo” selection protein of the coat protein complex COPII that mediates vesicle formation at the ER export sites. PIP oligomers then transit through the Golgi apparatus and trans-Golgi network and are then loaded into secretory vesicles and routed to the plasma membrane (Figure 3.72). Insertion of PIPs into the plasma membrane is mediated by a protein that regulates vesicle fusion (the “syntaxin” SYP121). The plasma-membrane localized PIPs can be recycled internally: once internalized in vesicles, PIPs are delivered to the trans-Golgi network before being routed back to the plasma membrane or directed into lytic vacuoles for degradation (Figure 3.72). Salt stress causes dephosphorylation and internalization of PIPs, and drought stress induces ubiquitylation of PIPs, which are then degraded in the proteasome.

Aquaporins assemble as homo- or hetero-tetramers, each monomer acting as an independent water channel. The structure of an aquaporin monomer (Figure 3.72) consists of six membrane-spanning α-helices connected by five loops, with both N and C termini facing the cytosol. Two loops form two short hydrophobic α-helices dipping halfway into the membranes, which, together with the membrane-spanning helices, create a pore with high specificity (Murata et al. 2000).

Figure 3.72 Formation and trafficking of PIPs within a plant cell. PIPs belonging to the PIP2 group (in yellow) form homo- or heterooligomers by associating with PIP1 isoforms (in green). PIP oligomers transit through the Golgi apparatus and trans-Golgi network (TGN) and are then loaded into secretory vesicles and routed to the plasma membrane. In the circle is shown the topological structure of the aquaporin monomer AQP1 (Murata et al. 2000), with six membrane-spanning α-helices (1-6) connected by five loops (A-E). The loops B and E form two short hydrophobic α-helices (in red) dipping halfway into the membranes, which, together with the membrane-spanning helices, create a pore with high specificity. From Chaumont and Tyerman (2014). Reproduced from F. Chaumant and S.D. Tyerman, Plant Phys 164: 1600-1618, plantphysiol.org. Copyright American Society of Plant Biologists.

Aquaporins play a central role in regulating plant water relations. Water diffusion across cell membranes is facilitated by aquaporins that provide plants with the means to rapidly and reversibly modify water permeability. This is done by changing aquaporin density and activity in the membrane, including post-translational modifications and protein interaction that act on their trafficking and gating. At the whole organ level aquaporins modify water conductance and gradients at key “gatekeeper” cell layers that impact on whole plant water flow and plant water potential. In this way they may act in concert with stomatal regulation.

PIP and TIP expression is higher during the day than the night, and correlates with diurnal changes in transpiration. It is likely under circadian regulation. Expression can correlate with changes in hydraulic conductivity, Lp, so that aquaporins are more abundant or have higher activity at times when stomatal conductance is higher. Over a diurnal cycle, Lp can change 2-5 fold and so can PIP transcript activity or protein abundance (Chaumant and Tyerman 2014). This occurs in both roots and shoots.

This section has shown how the flow of water and ions across membranes are linked. The function of membrane transport in regulating nutrient supply is covered in Chapter 4.

3.7 - References

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Chapter 4 - Nutrient uptake from soils

Chapter editors: Rana Munns and Susanne Schmidt

Contributing Authors: MC Brundrett1,2, BJ Ferguson3, PM Gressshoff3, S Filleur4, U Mathesius5, R Munns1,6, A Rasmussen7, MH Ryan1, P Ryan6, S Schmidt8, M Watt5

1School of Plant Biology, University of Western Australia; 2Department of Parks and Wildlife, Western Australia; 3Centre for Integrative Legume Research, University of Queensland, 4CNRS, Gif sur Yvette, France; 5Research School of Biology, Australian National University; 6CSIRO Agriculture, Canberra; 7Centre for Plant Integrative Biology, University of Nottingham, UK; 8School of Agriculture and Food Science, University of Queensland

With acknowledgements to authors of the original edition Chapter 3, sections 3, 4 and 5 respectively: BJ Atwell, JWG Cairney and KB Walsh

Plants require at least 14 essential minerals for growth, along with water and carbohydrates. The processes by which plants convert CO2 to carbohydrate are described in Chapters 1 and 2 of this text book, and Chapter 3 explains how plants take up water from the soil and transport it to leaves. Chapter 4 describes the fundamental processes by which plants acquire minerals from the soil, with N and P as main examples. In most situations, roots do not take up minerals directly from the soil, but work in association with soil microbes that make the minerals more available to plants.

This chapter first explains the concept of plant nutrition, then describes the various root structures and symbioses with microorganisms that allow plants to take up essential nutrients. These adaptations include specialised root architecture, cluster roots, rhizosphere organisms, mycorrhizas, and symbiotic nitrogen fixation. Finally the principles of membrane transport which require ATP hydrolysis or specialised membrane transporters are described with a focus on uptake of nitrate, different forms of organic N, and phosphate.

Nutrient application to soils via fertilizers, and the ecophysiology of nutrient relations are covered in Chapter 16.

4.0-Ch-Fig-4.0-NoNum v2 resized.jpg

Different nutrient acquisition strategies. (figure by S. Buckley)

4.1 - Nutrient requirements and root architecture

Amanda Rasmussen1 and Susanne Schmidt,2
1Centre for Plant Integrative Biology, University of Nottingham, UK; 2School of Agriculture and Food Science, University of Queensland

Plants require at least 14 essential elements called ‘mineral nutrients’ to sustain life function and complete their life cycle, in addition to carbon (in the form of CO2), oxygen, and hydrogen (in the form of water). Some plants have specific requirements for additional elements. The acquisition via the roots and use of these elements are the topic of plant nutrition.


Figure 4.1 Young wheat seedlings with a mass of elongated seminal roots that were excavated and washed (left) or imaged in situ using microCT (right). Note that in situ imaging preserves the lateral growth of lateral roots. Images courtesy of E. Delhaize (left) and S. Mairhofer (right).

Nutrients are taken up by roots via active or passive transport across membranes, and travel from the bulk soil to the roots via diffusion or mass flow. However, in order to access all the available nutrients, plants have evolved dynamic and plastic root systems that explore the soil for maximum nutrient uptake. In monocots, lateral roots grow into the volume of soil between seminal roots, as shown by in situ CT imaging (Figure 4.1).

By responding to signals and gradients in the soil, the root system can maximise growth in local nutrient patches while minimising growth in areas of deficiency. This is extremely important for plant survival particularly in deficient or marginal soils. Efficient root growth is also an important factor in maximising yield with lower fertiliser applications because ‘wasted’ root growth costs energy that could otherwise be invested in the crop of interest (whether seeds, leaves, stems or tubers). For this reason understanding the root environmental responses and breeding crops with efficient root systems for the conditions of interest are currently highly active areas of agronomic research.

This section covers the different nutrients required for plant growth, and the different root architectures and structures which help the plants maintain sufficient nutrient uptake to support the above ground biomass.

4.1.1 - Plant nutrition

Although the absolute quantities of nutrients required vary between plant species, genotypes and growth environments, essential nutrients are categorised into so-called macronutrients (N, K, Ca, P, Mg, S) that plants require in larger quantities, and micronutrients (Fe, Cl, Mn, B, Zn, Cu, Mo, Ni) that are needed in small amounts (Figure 4.2). Additional beneficial elements include Si (e.g. for grasses) and Na (for many sea-shore plants).


Figure 4.2 Mineral nutrient concentrations in plant leaves. Concentrations can be larger or smaller than shown here. Note, lower concentrations occur in roots and wood. (Adapted from Marschner 1995)

Macronutrients form the structural components of proteins, cell walls, membranes, nucleotides and chlorophyll, and have roles in energy and water maintenance. The macronutrient potassium has a special function in regulating the osmotic potential of plant cells. Under saline or dry conditions, Cl (and for some plants Na) is important in plant water relations.

Micronutrients mainly provide functional groups in enzymes (BOX 1 shows how Ni forms the active site in urease, as an example).

BOX 1 – Nickel (Ni) at the centre of Urease

In 1926 James B. Sumner from Cornell University studied the structure of Urease from Jack Bean plants and demonstrated that the enzyme is also a protein. This work led to the recognition that most enzymes are in fact proteins and in 1946 Sumner was awarded the Nobel Prize in Chemistry.

Urease is an enzyme that breaks down urea to ammonia and carbon dioxide in plants, bacteria and fungi and contains a nickel active site.

(NH2)2CO + H2O -> CO2 + 2NH3

For 3D structure see http://www.proteopedia.org/wiki/index.php/Urease

Further reading: Follmer 2008; Carter et al. 2009.

In terrestrial ecosystems and in agriculture, the availability of nitrogen (N) and phosphorus (P) are often limiting and so affect plant growth and productivity most strongly. However, other elements can also be limiting. Plants showing nutrient deficiencies will exhibit symptoms such as stunted growth, leaf or shoot tip chlorosis, and defoliation, and will die if supplements are not provided. Fertilisers are applied to supply essential elements in agriculture to maximise plant growth and enhance yields. Along with the discovery of ‘dwarfing genes’ and development of short stature crop varieties, it was the use of synthetic nitrogen fertilisers that played a significant role in the Green Revolution of the 1930s-1960s.

The acquisition of mineral nutrients starts with their movement from the surrounding soil to root surfaces. The movement of nutrients from bulk soil towards plant roots occurs via diffusion or mass flow (Figure 4.3). Root interception occurs as the root comes in contact with, and displaces the soil through which it is growing. Nutrient availability in soils and the physical and chemical factors influencing their movement from soils to the root surface is comprehensively described in a review by Marschner and Rengel (2012). Nutrients are taken up into roots by active or passive transport across cell membranes, which is described later in this chapter (Section 4.5).


Figure 4.3 Element movement to root surface. Root interception occurs as the root displaces soil; diffusion occurs along a concentration gradient; mass flow is driven by plant transpiration and is the movement of soil solution along a water potential gradient. (Based on Marschner and Rengel 2012)

Diffusion occurs along a concentration gradient, over relatively short distances (in the order of 1 cm). As roots take up nutrients and ions from the soil a depletion zone can be established allowing diffusion to occur into the depletion zone. The rate of diffusion depends on how fast the roots are taking up the nutrient, how much of the nutrient is present in the soil (this determines the steepness of the concentration gradient that forms) and also on the mobility of the ions by diffusion. Soluble ions would take about a day to diffuse 1 cm; ions bound to the soil matrix would take longer. For examples, Marscher and Rengel (2012) show that nitrate by diffusion in a ‘typical’ soil travels 3 mm in a day, potassium about 1 mm in a day, and phosphate moves only about 0.1 mm in a day. This illustrates the importance of root hairs in intercepting and accessing phosphate.

Mass flow is driven by the uptake of water caused by the transpiration rate of the plants and can occur over long distances. Many soluble nutrients such as nitrate are dissolved in the soil water and as the plant pulls the water from the soil, the nutrients move too. Some nutrients move by mass flow faster than their uptake rate so they build up on the surface of the root during daylight hours (Marschner and Rengel, 2012). The rate of movement by mass flow of solution depends basically on the rate of transpiration of the plant, so there is little movement at night. It is also influenced by soil water content and soil texture (see Chapter 3, Section 3.4).

Nutrients are unevenly distributed in the soil, generally being concentrated in the topsoil due to decomposition of leaf litter, but also dispersed elsewhere in pockets. Uneven surface enrichment arises from diverse sources such as dead fauna, urine patches from grazing animals, and localised application of fertiliser. Phosphorus and all cations are relatively immobile as they bind to the soil while nitrate and other anions (except phosphorus) are soluble and can readily be leached to deeper soil layers.

Because the soil is so heterogeneous, plants have developed adaptable (plastic) root systems so that the roots proliferate close to the nutrients for uptake.

4.1.2 - Root system architecture


Figure 4.4 Root systems of young (left) wheat and (right) lupin plants. Wheat, a monocot, has a dual root system. Seminal roots emerge from the seed and nodal roots (thicker roots on the outside of the picture) emerge from the crown, a group of closely packed nodes from which tillers emerge. Lupin, a dicot, has a tap root from which lateral roots emerge and which thickens with time as continued cambial activity leads to secondary growth.

The root system architecture is the arrangement of different roots in solid space. Just like a building has walls, roof, and floors, plant root systems also contain different structures including root types (primary, lateral, adventitious), root hairs, and specialized features such as nodules and cluster roots (see case study). In contrast to a fixed structure like a building, the root system is dynamic with new structures forming as needed to explore the soil and old structures breaking down when their use has expired. This four dimensional architecture within soil can now be visualized using technology such as X-ray microscale computed tomography (microCT) and magnetic resonance imaging (MRI).

In order to understand root architecture it is important to understand the different structures that make up the root system. This section will focus on the different root types while cluster roots are explained in the case study that follows. Root hairs are described in the section on the rhizosphere (Section 4.2) and the formation of N2-fixing nodules in Section 4.4.


Figure 4.5 MicroCT images of crop root systems. A, Single wheat plant with primary root (P), lateral roots (L) and seminal roots (S). Roots are false-coloured in white, and soil is false-coloured in brown. Scale bar = 1 cm. (Reproduced from Atkinson et al. (2014) Plant Physiol 166: 538-550. ​doi:​10.​1104/​pp.​114.​245423. Copyright American Society of Plant Biologists). B) Two wheat plants grown in the same pot. Roots are false-coloured in blue or orange; each colour shows a single root system. (Image courtesy S. Mairhofer)



Figure 4.6 Root types. White = primary root, cream = seminal roots, blue = first order lateral roots, pink = second order lateral roots, yellow = crown roots, orange = brace roots. Crown and brace roots are both adventitious root types. (Original drawing courtesy A Rasmussen)

Primary root: the first root to emerge from a germinating seed (starts as the radicle). Since the primary root is present within the embryo, this class of root is embryonic.

Lateral roots: roots that form from other roots. The lateral roots that form from the primary root are first order lateral roots; the lateral roots that form from the first order laterals are second order laterals and so on. This class of root is post-embryonic.

Seminal roots: form adjacent to the radicle and dominate the early root growth in monocots. This root type is embryonic.

Adventitious roots: any root that forms from anything other than another root. This includes roots that form on the base of stem cuttings, from leaf explants, from stems in flooded plants and also from nodes of cereal crops (often called crown roots). These root types are very diverse (Steffens and Rasmussen, 2015) so can include both embryonic and post-embryonic roots.


Figure 4.7 Examples of adventitious root types. This figure highlights a few examples of the diversity of adventitious roots; A and B show types of adventitious roots that form during normal development while C and D are examples of stress-induced adventitious roots. A, Those potentially established in the embryo. B, The dominant root system of monocots including maize (top image) crown roots (yellow) and brace roots (orange) and nodal roots on other grasses (lower image) and on eudicots such as strawberry. C, Low or no light (e.g. Arabidopsis used as a model for adventitious root regulation) or flooding (lower image) can induce adventitious roots from either nodal or non-nodal stem positions. D, Wounding such as taking a cutting induces de novo adventitious root development. Primary and seminal roots are depicted in white, first order lateral roots in blue and second order laterals in pink. (Based on Steffens and Rasmussen (2016) Plant Physiol. 170: 603-617. doi:10.1104/pp.15.01360. Copyright American Society of Plant Biologists)

Root hairs: single-cell, hair-like extrusions from the epidermis which increase root surface area for nutrient uptake (Jones and Dolan, 2012) and are important for nodulation (Section 4.4).

The combination of different root types present in the root system differs across species. In particular the root systems of cereal crops (monocots) differ dramatically to the root systems of tree crops (eudicots) (Figure 4.8).


Figure 4.8 Eudicot and monocot root systems. A ,C and E represent eudicot roots while B, D and F represent monocot roots. Schematic showing the dominant root types of tomato (A) and maize (B). SR = stem roots, P = primary root, L = lateral root, Br = brace root, Cr = crown root, S = seminal root (original diagram courtesy A. Rasmussen). MicroCT images of tomato (C) and maize (D) (images courtesy J. Johnson and S. Mairhofer). Cross section schematics of the eudicot Arabidopsis (E) and rice (F) showing where new roots initiate. (Reproduced from Atkinson et al. (2014) Plant Physiol 166: 538-550. ​doi:​10.​1104/​pp.​114.​245423. Copyright American Society of Plant Biologists)

Eudicots typically develop a primary (tap) root from a single radicle that emerges from a seed. This primary root, plus the first order lateral roots which emerge from it, provide a framework on which higher-order lateral roots are formed. Such a framework strengthens due to secondary thickening when division of the cambium gives rise to more cell layers, leading to massive roots that are often seen radiating from the base of a tree trunk (Figure 4.9).

Monocots such as grasses and cereal crops do not have a cambium for secondary thickening and develop a fibrous root system. This root system begins with the radicle which grows into the primary root. Adjacent to the radicle, several seminal roots also emerge and combined with the primary root these roots dominate the young root systems of monocots. Next nodal adventitious roots (often called crown roots) emerge from lower stem nodes and these thicker roots gradually dominate the root system. Finally in some monocots, such as maize (corn), nodal adventitious roots emerge above the soil level (brace roots) to provide additional structural support. Stems of monocots are typically anchored by the nodal roots, which are more numerous than seminal roots (Hochholdinger et al. 2004; Hochholdinger 2009).

Despite these structural differences between monocot and eudicot root systems, they can all vary the soil volume which they explore depending on water and nutrient availability. In this way the root distribution in the soil can vary both vertically and locally depending on available resources.

Root distribution


Figure 4.9 Dimorphic root system of a six-year-old Banksia prionotes tree growing in Western Australia in a deep sand. The trunk (T) is connected through a swollen junction (J) to the root system which comprises a dominant sinker root (S) with smaller sinkers (S2). A system of lateral roots (L) emerge horizontally from the junction, some bearing smaller sinker roots (arrows). Other laterals give rise to cluster roots (CR). (W.D. Jeschke and J.S. Pate, J Exp Bot 46: 907-915, 1995)

The amount of roots present in a volume of soil varies both vertically and locally depending on resource availability and physical restrictions. This is often measured as the total length of all roots present per unit volume of soil (root length density, L, expressed in km m–3).


Figure 4.10 Root length density in relation to depth in the soil for a wheat crop and a jarrah forest (Eucalyptus marginata). (Jarrah data from B.A. Carbon et al., Forest Sci 26: 656-664, 1980; wheat data courtesy F.X. Dunin)

Vertically, the root length density is often large in surface layers of the soil and typically decreases with increasing depth. Commonly, hundreds of kilometres of root per cubic metre of soil are observed near the soil surface.

Figure 4.10 shows root length density, L, as a function of depth in a wheat crop in early spring, and under a jarrah forest, also in spring. Both have a dense population of roots near the surface but wheat roots barely penetrate below 1 m, whereas jarrah roots penetrate to well below the 2.5 m shown here, often to 20 m. Dense root proliferation near the soil surface probably reflects an adaptation of plants to acquire phosphorus, potassium and other cations such as the micronutrients zinc and copper. These nutrients do not move readily in soil as they are bound to the soil surfaces, hence roots branch prolifically to ensure close proximity (a few millimetres) between adsorbing surfaces and these soil-immobile ions. Roots of jarrah are also concentrated near the soil surface (Figure 4.10) to access phosphate and nutrients released by litter decomposition, but some roots penetrate very deeply to tap subsoil moisture.

Nutrients are distributed unevenly in the soil. Root systems respond to enriched zones of nutrients by high levels of branching. Figure 4.11 shows an example of such a proliferation; the dense roots in the centre of the figure are a response by the row of wheat plants to application of a large pellet of nitrogen fertiliser (see arrow).


Figure 4.11 Excavated root system of wheat plants whose roots were provided with a concentrated band of ammonium sulphate fertiliser at the head of the arrow. (Photograph courtesy J.B. Passioura)

Such proliferations around bands of fertilizer ensure plants maximise nutrient uptake with the minimum cost to plant development. This efficiency fits within optimal partitioning theory which states that plants respond to environmental variation by partitioning biomass among the plant organs to optimize the acquisition of nutrients, light, water and carbon to maximize plant growth (Reich, 2002). This means that in low nutrient conditions the plants will put more energy into growing roots and less into shoot growth (Reich, 2002). Likewise when light is limiting, plants will invest more energy in leaf area and less in root development (Weaver and Himmel, 1929; Reich, 2002). Maintaining the balance between root and shoot is important as the roots must be extensive enough to supply nutrients and water in proportion to the demand and hydraulic pull from the leaves and vice versa the leaves must produce enough sugars to continue the growth of the root system (Weaver and Himmel, 1929). Consistent with this, Butler et al. (2010) found in Sitka spruce forest, the root absorbing area was correlated with the tree stem diameter and to the transpiring leaf area index. This highlights the link in hydraulics between leaf and root areas.

Young roots absorb nutrients more rapidly than old roots. New roots supply annual plants with abundant sites for nutrient uptake, especially during establishment. A feature of the roots of perennials is that they have a large turnover of the fine, high-order lateral roots that emerge from the secondarily thickened framework each year. This turnover draws heavily on photoassimilate, equivalent to half the CO2 fixed in annuals and up to 90% of the standing biomass of temperate forests. Production of fine (and often ephemeral) roots ensures uptake of nutrients over many years.

Because many soils are deficient in key nutrients, plants have developed a special relationship with certain fungi called mycorrhizae (Section 4.4). In this symbiosis the fungi obtain fixed carbon from the host plant, and in turn supply the host with poorly mobile nutrients, especially phosphorus. This is achieved by proliferating their hyphae to provide a much greater surface area for nutrient uptake than could be provided by roots alone. Another adaptation, common in the Proteaceae, and also occurring in some species of lupin, is proteoid roots, clusters of tiny rootlets that greatly enlarge the available surface area for ion uptake and which are inducible by low levels of phosphorus (see Case study 4.1).

Case Study - Cluster (proteoid) roots

M. Watt


Figure 1 Cluster roots in Banksia serrata growing on Hawkesbury Sandstone hillslopes in the Sydney region. a, Roots that have grown across a dead eucalypt leaf extract nutrients remaining in the decaying leaf. b, Clusters of fine rootlets at the tips of roots increase the surface area for nutrient extraction from surrounding soil. Scale bar = 100 µm. (Scanning electron micrograph courtesy S. Gould)

Cluster or proteoid roots (Figure 1) are found in many species originating from nutrient-deficient soils (Dinkelaker et al. 1995). They enhance uptake of nutrients, especially phosphate. Species which develop these “dense clusters of rootlets of limited growth” include members of the Australian family Proteaceae, where they were first described by Purnell (1960). Other families such as the Casuarinaceae, Cyperaceae, Mimosaceae and Restionaceae also contain species with heavily branched root systems (Lamont 1993). Significantly, few species with cluster roots are mycorrhizal, implying that root clusters fulfil a similar role to mycorrhizal fungi.

Australian soils generally contain low concentrations of plant-available phosphate, much of it bound with iron–aluminium silicates into insoluble forms or concentrated in the remains of decaying plant matter. Because very little of this phosphate is soluble, most roots extract it only slowly. Plants with cluster roots gain access to fixed and organic phosphate through an increase in surface area and release of phosphate-solubilising exudates. Hence plants with cluster roots grow faster on phosphate-fixing soils than species without clusters.

Cluster roots have a distinct morphology. Intense proliferation of closely spaced, lateral ‘rootlets’ occurs along part of a root axis to form the visually striking structures. Root hairs develop along each rootlet and result in a further increase in surface area compared to regions where cluster roots have not developed.

In the Proteaceae, clusters generally form on basal laterals so that they are abundant near the soil surface where most nutrients are found. For example, Banksia serrata produces a persistent, dense root mat capable of intercepting nutrients from leaf litter and binding the protecting underlying soil from erosion (Figure 1a). New clusters differentiate on the surface of this mat after fires and are well placed to capture nutrients. In contrast, Banksia prionotes forms ephemeral clusters which export large amounts of nutrients during winter. Lupinus albus has more random clusters which appear on up to 50% of roots (Figure 2).


Figure 2 Basal roots of a two-week-old Lupinus albus plant grown in nutrient culture with 1 µM phosphate. Proteoid roots have emerged along the primary lateral roots (arrowhead) and the oldest proteoid rootlets have reached a determinate length of 5 mm. As rootlets approach their final length, they exude citrate for 2-3 d. (mm scale on left side) (Photograph courtesy M. Watt)

Rootlets not only represent an increase in surface area but also exude protons and organic acids, solubilising phosphate and making it available for uptake (Watt and Evans 1999a). Exudates from cluster roots represent up to 10–23% of the total weight of an L. albus plant, suggesting that they constitute a major sink for photoassimilates. However, not all this additional carbon comes from photosynthesis because approximately 30% of the carbon demand of clusters is met by dark CO2 fixation via phosphoenolpyruvate carboxylase. Because cluster roots form on roots of L. albus even when phosphate supply is adequate, growth of L. albus in soils with low phosphate availability is not restricted by an additional carbon ‘drain’ to roots. On the other hand, the great many species which produce cluster roots in response to environmental cues like phosphate deficiency might experience a carbon penalty to support these roots.

Cluster roots on L. albus are efficient with respect to carbon consumption by generating citrate on cue. Most of the citrate exuded by clusters is released during a two to three day period when the cluster is young (Watt et al. 1999b). A large root surface area in clusters works in concert with this burst of exudation to solubilise phosphate before it is re-fixed to clay surfaces or diffuses away.

Cluster roots can mine a pocket of phosphate-rich soil which would otherwise not yield its nutrients. They are an elegant adaptation of root structure and biochemistry to nutrient-poor soils.


Dinkelaker B, Hengeler C, Marschner H (1995) Distribution and function of proteoid roots and other root clusters. Bot Acta 108: 183–200

Lamont BB (1993) Why are hairy root clusters so abundant in the most nutrient impoverished soils of Australia? Plant Soil 156: 269–272

Purnell HM (1960) Studies of the family Proteaceae 1. Anatomy and morphology of the roots of some Victorian species. Aust J Bot 8: 38–50

Watt M, Evans JR (1999a) Proteoid Roots. Physiology and Development. Plant Physiol 121: 317–323

Watt M, Evans JR (1999b) Linking development and determinacy with organic acid efflux from proteoid roots of Lupinus albus L. grown with low phosphorus and ambient or elevated atmospheric CO2 concentration. Plant Physiol 120: 705–716

4.2 - Soil-root interface

Ulrike Mathesius, Research School of Biology, Australian National University

As a general rule, the surface area of a root system exceeds the leaf canopy it supports. Even disregarding root hairs, the interface between roots of a three-week-old lupin plant and soil is about 100 cm2 while a four-month-old rye plant under good conditions has more than 200 m2 of root surface (Dittmer 1937). Trees’ root systems are difficult to quantify but kilometres of new roots each year generate hundreds of square metres of root surface. Such a root–soil interface arises through the simultaneous activity of up to half a million root meristems in a mature tree.


Figure 4.12 Transverse view of a young, soil-grown wheat root, sectioned by hand and stained with Toluidine Blue. Most soil in the rhizosheath was washed away during preparation, revealing many long root hairs extending from the main axis (diameter 0.6 mm). Root hairs allow this root to explore 21 times more soil volume than would be possible without hairs. A lateral root can be seen extending from the pericycle which surrounds the stele. (Photograph courtesy M. Watt)

Many roots form fine extensions to epidermal cells called root hairs, amplifying the effective surface area of the soil–root interface many times. Dittmer (1937) estimated that the surface area of root hairs in rye plants was more than that of the root axes on which they grew; similar observations have been made for trees. The aggregate length of root hairs in the rye plants studied by Dittmer increased 18 times faster than that of the main axes. Thus, up to 21 times more soil is explored when root hairs are present (Figure 4.12).

Root hairs are particularly important in taking up mineral nutrients that are not readily soluble and therefore not mobile in the soil solution, like phosphate. Measurements of the phosphate concentration in soil at different distances from roots show that soil phosphate is depleted only in the zone close to roots, the 1 mm zone, the typical length of root hairs (Figure 4.13).

Anchorage and extraction of inorganic soil resources both call for a large area of contact between roots and soil. However, this vast interface is much more than a neutral interface; events within it allow resources to be extracted from the most unyielding soils. Intense chemical and biological activity in a narrow sleeve surrounding roots, particularly young axes, give rise to a rhizosphere, the volume of soil influenced by the root, a concept first introduced in 1904 by Lorenz Hiltner (Hartmann et al. 2008). The rhizosphere has been estimated to contain up to 1011 microbial cells per gram of soil, and harbour up to or above 30,000 different microbial species, undoubtedly the moxt complex ecosystem on earth (Berendsen et al. 2012). The rhizosphere concept has been extended to include symbiotic mycorrhizal fungi associate with the root (See Section 4.3 on mycorrhiza), and this has been named the ‘mycorrhizosphere’, as most land plants are colonised by mycorrhizal fungi most of the time.


Figure 4.13 Young root tip with elongating root hairs. Root tip of Medicago truncatula showing the approximate zones of root elongation and differentiation relative to the tip. Root hairs are protuberances of epidermal cells that first emerge approximately 4 mm behind the root tip and elongate over the span of about two days until they are fully elongated in the differentiation zone of the root. (Photograph courtesy U. Mathesius)

4.2.1 - The rhizosphere

The rhizosphere is the narrow zone of soil surrounding plant roots that is characterised by root exudation and an abundance of micro-oganisms which can be beneficial or harmful to plants, or have no effect on root growth and function. These microbes are saprophytic, pathogenic or symbiotic bacteria and fungi, including rhizobia forming nodules and arbuscular mycorrhizal fungi (Figure 4.14).


Figure 4.14 The rhizosphere is the narrow zone of soil surrounding plant roots that is characterised by root exudation and an abundance of saprophytic, pathogenic and symbiotic bacteria and fungi. These include rhizobia that form nodules, and arbuscular mycorrhizal fungi (AMF). The rhizoplane describes the root surface in contact with the soil. Root cap and root border cells near the root tip provide lubrication as the root expands into the soil. (Reproduced by permission from Macmillan Publishers Ltd from L. Philippot et al. Nature Rev Microbiol 11: 789-799, 2013)

Many root phenomena suggest specific roles for the rhizosphere. For example, roots have long been thought to find a relatively frictionless path through soils because of exudation of organic substances and cell sloughing, but the chemical and physical processes that underpin this phenomenon are still quite unclear (McKenzie et al. 2012). Production of new roots around local zones of enrichment (Section 4.1) is made far more effective through rhizosphere activity associated with these young roots. Phosphate availability is particularly likely to be improved by the presence of a rhizosphere. Potential mechanisms will be discussed below.

Enhancement of root growth under conditions which favour high root:shoot ratios and the attendant rhizosphere surrounding those roots (rhizosheath) require a substantial input of organic carbon from shoots. Some is used in structural roles, while roots and microbes also require large amounts of carbon to sustain respiration. Even in plants growing in nutrient-adequate, moist soils, 30–60% of net photosynthate finds its way to roots (Marschner 1995). Carbon allocation to roots can be even greater in poor soils or during drought. The rhizosphere accounts for a large amount of the root carbon consumption (Jones et al. 2009). Barber and Martin (1976) showed that 7–13% of net photosynthate was released by wheat roots over three weeks under sterile conditions while 18–25% was released when roots were not sterile. This difference might be considered carbon released because of microbially-induced demand in the rhizosphere, and therefore made unavailable for plant growth.


Figure 4.15 Concentration of Enterobacter cloacae (RP8) around wheat roots when the bacterium was introduced by inoculating seeds (circles) or soil (triangles). Uninoculated controls are shown as diamonds. Approximately 3 mm of the soil around roots supports an elevated bacterial population (A.F. Dijkstra et al. Soil Biol Biochem 19: 351-352, 1987)

Rhizosphere chemistry and physics differ from the adjacent soil matrix and root tissues. Gradients in solutes, water and gases combine with microbial activity to produce a unique compartment through which roots perceive bulk soil. This zone of influence typically extends not more than 3 mm from the root axis (Figure 4.15), partly due to the low diffusion coefficients of most solutes that move through the rhizosphere (10–12 to 10–15 m2 s–1 for ions such as orthophosphates). Even a relatively mobile ion such as nitrate, with a diffusion coefficient (D) of around 10–9 m2 s–1 in soil solution, diffuses through about 1 cm of soil in a day. Because the time required (t) for diffusion of ions is a function of the square of distance traversed (l), where t = l2/D a nitrate ion would take four days to travel 2 cm, nine days to travel 3 cm and so on. Similarly, organic carbon diffuses away from roots only slowly, sustaining a microbial population as it is consumed in the rhizosphere.

Roots advancing through soil perceive a wide range of chemical and biological environments: a rhizosphere simultaneously fulfils buffering, extraction and defence roles allowing roots to exploit soils. A rhizosphere is thus a dynamic space, responding to biological and environmental conditions and often improving acquisition of soil resources. New roots develop an active rhizosphere which matures rapidly as the root axis differentiates.

4.2.2 - Rhizosphere chemistry

Photoassimilate diffuses from roots into the rhizosphere where it is either respired by microorganisms, volatilized, or deposited as organic carbon (‘rhizodeposition’). Some of this photoassimilate loss is in the form of soluble metabolites, but polymers and cells sloughed off the root cap also provide carbon substrates. Grasses undergo cortical cell death as a normal developmental process, providing further carbon substrates to support a rhizosphere microflora. Nitrogen and some other inorganic nutrients which are co-released with plant carbon are often reabsorbed by roots. Extraction of minerals from bulk soil also relies strongly on rhizosphere processes, especially near the root apices. Compounds exuded from roots interact with soil components in direct chemical reactions (e.g. adsorption reactions), through microbially mediated events (e.g. immobilisation reactions) and volatilisation. In addition, complex polysaccharides and glycoproteins of microbial and root origin give rise to a gelatinous mucilage which associates with soil particles to form a rhizosheath.


Figure 4.16 Roots of a young wheat plant showing soil attached to roots, the rhizosheath. Only the root tips, without hairs, have no rhizosheath. (Photograph courtesy E. Delhaize)

The rhizosheath is known as the soil that adheres to roots when they are removed from the pot or field Figure 4.16). The amount of soil can vary depending on how gently or roughly the roots are removed. For wheat, at least, the size of the rhizosheath correlates with root hair length. Mutants without root hairs have no rhizosheath. The distinction between the terms rhizosheath and rhizosphere are that the first term refers to the soil that physically adheres, and the second term the volume of soil influenced by the root. Mutants without root hairs would still have a rhizosphere of sorts since the root would still chemically influence its surrounding soil.

Rhizosheaths have physical and chemical implications for root function. Hydraulic continuity between soil and roots is, for example, thought to be enhanced by the hydrated mucigel, which facilitates water uptake by roots in dry soils. Negatively charged groups on side-chains of mucilagenous polysaccharides attract cations like Ca2+, providing exchange sites from which roots might absorb nutrients. The mucigel between the sloughed root cap and root border cells also acts as a lubricant for reducing penetration resistance of the expanding root tip in soil (McKenzie et al. 2012). For example, root elongation through hard soil is greatly reduced if the root cap cells are removed. Once the soil and the mucigel dry up, this lubrication effect is significantly reduced.

Such a diversity of chemical reactions in the rhizosphere is largely an outcome of the array of root-derived exudates. For example, phenolic compounds can be released by root cells in large amounts (Marschner 1995), both as a result of degradation of cell walls and from intracellular compartments. Flavonoids are a group of phenolics that can be specifically exuded into the rhizosphere as signal molecules to attract rhizobia (See section 4.4 on nitrogen fixation). Release of organic acids (principally citric, fumaric and malic acids) solubilises phosphate from surfaces to which they are adsorbed in many species, including those of the family Proteaceae. A modest release of organic acids accounting for about 0.1% of the root mass each week is sufficient to enhance phosphate acquisition in a selection of annual legumes (Ohwaki and Hirata 1992). In more extreme cases, up to a quarter of the dry weight of Lupinus albus plants is released from cluster roots, mostly as citrate (see Case study 4.1). Even the fungal hyphae of mycorrhizal eucalypt and pine roots can secrete photoassimilates, in the form of oxalic acid, causing phosphorus to be solubilised from insoluble calcium apatite (Malajczuk and Cromack 1982).

The main families of low molecular weight compounds which react with inorganic ions are phenolics, amino acids and organic acids. Heavy metals such as aluminium, cadmium and lead are complexed by phenolics, affecting the mobility and fate of these ions in contaminated soils. Flavonoids can chelate iron and make iron oxides available to plants. Manganese is complexed by organic acids, as are ferric ions, which also interact chemically with phenolic compounds and amino acids. For example, highly specialised amino acids (phytosiderophores) can complex ferric ions and enhance uptake from soils by rendering iron soluble. Low iron status actually stimulates release of phytosiderophores into the rhizosphere (Marschner 1995). Other metals such as zinc and copper might also be made more available to the plant through the chelating action of phytosiderophores. Chemical processing by chelating agents is dependent on plant perception of nutrient deficiencies, leading to an ordered change in rhizosphere chemistry. A significant demand on photoassimilates is required to sustain chelation of nutrient ions.

Enzymes are also released from roots, particularly phosphatases, which cleave inorganic phosphate from organic sources. The low mobility of orthophosphates means that phosphatases can be an important agent in phosphorus acquisition, especially in heathland soils where the native phosphorus levels are low relative to the phosphorus-rich remnants of decaying plant material.

pH is another important rhizosphere property. Roots can acidify the rhizosphere by up to two pH units compared to the surrounding bulk soil through release of protons, bicarbonate, organic acids and CO2 (Figure 4.17). In contrast, the rhizosphere of roots fed predominantly with nitrate was more alkaline than bulk soil. A distinct rhizospheric pH arises because of the thin layer of intense biological activity close to roots, especially young roots. In addition to proton fluxes, release of CO2 by respiring roots and microbes is likely to cause stronger acidification of the rhizosphere near root apices where respiration is most rapid.


Figure 4.17 Root-induced changes in the rhizosphere. a, Oxygen profiles across a growing root of Juncus effusus (in white). b, pH profiles across growing roots of intercropped durum wheat (dashed white) and chickpea (solid white). (Adapted from L. Philippot et al. Nature Rev Microbiol 11: 789-799, 2013, S. Blossfeld et al. Soil Biol Biochem 43: 1186-1197, 2011, and S. Blossfield et al. Ann Bot 112: 267-276, 2013, with permissions respectively from Macmillan Publishers Ltd, Elsevier, and Oxford University Press)

Rhizosphere acidification affects nutrient acquisition by liberating cations from negative adsorption sites on clay surfaces and solubilising phosphate from phosphate-fixing soils. Furthermore, micronutrients present as hydroxides can be released at low pH, conferring alkalinity tolerance on those species with more acidic rhizospheres. So, the rhizosphere is a space which ensheathes particularly the youngest, most active parts of a root in a chemical milieu of the root’s making. In this way, acquisition of soil resources is strongly controlled by processes within roots. Local variations within soil are buffered by rhizosphere chemistry, enabling roots to exploit heterogeneous soils effectively.

4.2.3 - Rhizosphere biology


Figure 4.18 Mature rhizosphere from roots of clover (Trifolium subterraneum L.). The outer cortex has been crushed and epidermal cells (EP) have become distorted, leading to leakage of substrates into the rhizosphere. The rhizosphere is rich in microorganisms with bacteria (B) clearly visible. Soil (Q) and clay (CL) particles are held together in the inner rhizosphere by a mucilage of polysaccharides. Sustained losses of carbon required to maintain this microflora are thought to come from exudation and senescence of root cells. (× 10,000) (Courtesy R.C. Foster, A.D. Rovira and T.W. Cock)

Microbial activity, sustained by photoassimilates secreted from roots, contributes substantially to rhizosphere properties. The level of microbial activity is also influenced by availability of nitrogen as a substrate for microbial growth. Soils with high fertility and biological activity have microbial densities 5–50 times greater in the rhizosphere than in bulk soil. The diversity of rhizosphere microflora is spectacular (Figure 4.18) and still incompletely described. An initial hurdle in the identification of rhizosphere microbes was the fact that most of them are unculturable on most known growth media. Metagenomics allows species to be sequenced from soil samples without culturing, largely overcoming this bottleneck. Next Generation sequencing studies of microbes present on root surfaces and in the rhizosphere soil have discovered thousands of different bacterial and fungal species living in close association with plant roots (e.g. Bakker et al. 2013; Bulgarelli et al. 2012; Lundberg et al. 2012). Some of these are very abundant and found in association with many plant species, others are less abundant and highly variable (Figure 4.19).


Figure 4.19 The composition of the bacterial community in the rhizosphere. The figure shows examples of the composition of the bacterial community in the rhizosphere of three maize genotypes (Mo17, B73 and III14h) and of sugarbeet. The distribution of the different bacterial phyla is based on data obtained by 454 sequencing (maize) and G3 PhyloChip analyses (sugarbeet). The bacterial community composition was characterized in the rhizosphere of 27 maize genotypes cultivated in five fields located in three states in the USA. Here, three genotypes displaying contrasted rhizosphere microbiota in a given field are depicted for illustration and the sugar beet rhizosphere microbiota presented is from seedlings grown in a disease-conducive soil in The Netherlands. (Reproduced by permission from Macmillan Publishers Ltd from L. Philippot et al. Nature Rev Microbiol 11: 789-799, 2013)

These microbes can almost be viewed as an extension of the plant into the soil. Like the human gut microbiome, the plant rhizosphere microbiome appears to be an essential part of the plant with multiple functions in nutrition and pathogen defense; it is inseparable from the plant and has been dubbed the plants second genome. The rhizosphere community is highly structured and not a random collection of species – it is strongly influenced by plant species and even ecotypes, by the type of the soil, availability of nutrients and the exudation of chemicals from the root (Bakker et al 2013). Plant mutants with altered chemical composition of root exudates have been found to attract significantly altered microbial communities. It will be fascinating to discover to what extent this is an active strategy of the plant to attract the most appropriate rhizosphere microbiome to help the plant survive in a given environment.

Rhizosphere microorganisms are also not uniformly distributed along roots. Apices are almost free of microbes but densities can increase dramatically in subapical zones. Very mature root axes with lateral branches are sparsely populated with microbes. Even within these zones, there are large variations in distribution, with radial epidermal walls of roots secreting exudates which can support huge microbial populations, up to 2 × 1011 microbes cm–3. Composition of microbial communities varies with their distribution along the root as well, likely reflecting different nutrient sources along the root. Fluorescence in situ hybridization (FISH) can be used to visualise different taxa of bacteria on the root surface (rhizoplane; Figure 4.20).


Figure 4.20 Arabidopsis root-inhabiting bacteria are detectable on the rhizoplane. a to e, Scanning electron micrographs of bacteria-like structures. Bars, 1 mm. f to j, Detection of bacteria by fluorescence in situ hydridisation (FISH) using probes against specific bacterial groups (bacteria in green due to AlexaFluor488) on the root surface (red, root autofluorescence) by confocal laser scanning microscopy. f, Most Eubacteria detected with probe EUB338. g, Negative control with reverse complementary probe of EUB338 (NONEUB). h, Betaproteobacteria detected with probe BET42a. i, Bacteroidetes detected with probe CF319a. j, Actinobacteria detected with probe HGC69a. Bars, 20 mm. (Reproduced by permission from Macmillan Publishers Ltd from D. Bulgarelli D et al., Nature 488: 91-95, 2012)

Roots do of course influence adjacent soil throughout their length by setting up gradients of water, gases and ions. For example, in waterlogged soils leakage of O2 from aerenchymatous roots leads to oxidation of metal ions and local build up of aerobic microflora around roots of agricultural plants (Chapter 18). In general, however, the most active microbial populations and rates of chemical transformation in the rhizosphere occur in the subapical zones of the root. In supporting these processes, root-associated microbes metabolise inorganic nitrogen, depositing protein nitrogen in the process of immobilisation. Microbial activity also produces plant growth regulators such as auxin, cytokinins and gibberellins, sometimes in amounts sufficient to influence root morphogenesis. Ethylene can also be produced by rhizospheric fungi, potentially influencing root morphological changes such as lateral root initiation. Some bacteria have been found to promote plant growth by reducing ethylene levels around roots through production of an enzyme degrading an ethylene precursor, 1-aminocyclopropane-1-carboxylate (ACC) deaminase.

4.2.4 - Costs and benefits of a rhizosphere

Root function and overall plant performance can benefit conspicuously from processes in the rhizosphere. Infection by rhizobia (Section 4.4) and mycorrhizal fungi (Section 4.3) improve the nutritional status of many species. Rhizobial strains have even been used to manipulate rhizosphere biology. A significant proportion of photoassimilate is used to support a rhizosphere, reflecting the high cost of microbial activity and polymer exudation. This pattern is repeated in many species with up to 20% of plant carbon consistently lost by roots, however, this value can vary substantially with the biotic and abiotic conditions. Relative rates of microbial and root respiration are almost impossible to estimate in roots growing in undisturbed soils because of the intimacy of roots and microbes. In addition to consuming large amounts of plant carbon, some microbes can produce phytotoxins, which can impose further restrictions on root function. Some microbes also contribute to nutrient depletion in the rhizosphere, for example by converting usable forms of nitrogen, i.e. nitrate or ammonium, into unusable forms like N2.

Mechanisms describing how a rhizosphere benefits its host are even more elusive because of the diversity of reactions in such a small space. Chelation is identified as a major influence on nutrient acquisition and might also help ameliorate ion toxicities. Physical properties of the rhizosphere are even less well understood, with questions such as root lubrication, root–mucilage shrinkage and interfacial water transport not yet resolved. Physical properties of mucilage do not suggest it is an ideal lubricant. Whether the dynamic properties of a rhizosphere bring constant benefits to a plant or simply passively coexist with growing roots remains a critical question.

One demonstrated benefit of the rhizosphere microbiome is the protection of the plant from diseases. Several mechanisms have been suggested for this effect (Bakker et al. 2013; Berendsen et al. 2012): Disease suppressiveness, the ability of the microbial inhabits of the rhizosphere to suppress the infection of plants by soil-swelling pathogens, has been ascribed to the production of antimicrobial substances by bacteria, to competition between beneficial and pathogenic microbes and to the induction of systemic resistance by beneficial bacteria. An intriguing example is the colonization of plants by pathogens, which can lead to changes in the exudation of organic acids that then attract beneficial bacteria that induce systemic resistance in the plant, reducing pathogen infection. The induction of systemic resistance to pathogens can also be triggered by specific signaling molecules of bacteria, quorum sensing signals, which bacteria used to ‘talk’ to each other to coordinate multicellular-like behaviours of bacterial colonies. Perception of quorum sensing signals from rhizosphere bacteria by plants can increase systemic resistance to pathogens in the shoot, and can also enhance symbiosis with nitrogen-fixing bacteria. Quorum sensing signal perception also triggers the production of so called quorum sensing mimic compounds – signals that interfere with bacterial communication in the rhizosphere (Teplitski et al. 2011). While we still need to identify most of the signals, signal mimics and exudate components in the interaction of roots with their microbiome, it is clear that plants actively create the rhizosphere, and that this is likely to benefit the plant in its environment.

4.3 - Mycorrhizal associations

Megan H Ryan1 and Mark C Brundrett1,2
1School of Plant Biology, University of Western Australia; 2Department of Parks and Wildlife, Western Australia.

The roots of around 90% of higher plants form a symbiotic association with mycorrhizal fungi (Figure 4.21). These fungi colonise roots, with the colonised root being termed a “mycorrhiza”. The fungi benefit from the provision of plant carbon. The host plant may benefit in many ways, but the primary benefit is most often the ability to access inorganic nutrients from soil beyond the rhizosphere due to their transport into the root by hyphae of the fungi. Mycorrhizal associations are present in plants in both natural ecosystems and modern agricultural systems; although their occurrence in the latter may be reduced by common management practices, especially the addition of fertiliser.


Figure 4.21 Relative importance of mycorrhizal associations for all flowering plants. About 94% of plants can form mycorrhizas of various types. Arbuscular mycorrhizas (AM) are the most common type. Shown in light green are the 8% of species with inconsistent associations that vary with habitat or soil conditions and can be nonmycorrhizal or AM. (Based on Brundrett 2009)

Mycorrhizal fungi are thought to have aided the first plants to colonise land, but most or all species in some plant genera have subsequently lost the ability to form mycorrhizas (e.g. Lupinus, Brassica and Banksia). Nonmycorrhizal plants may have roots that are consistently free of mycorrhizal fungi or have inconsistent associations. The former tend to have alternative nutrition strategies, the latter occur in soils where fungal activity is inhibited, at least part of the time. On a global scale, nonmycorrhizal plants tend to be more common in colder arctic and alpine habitats, and wetland and aquatic habitats, as well as in saline soils and arid habitats (Figure 4.22). These habitats also include many plants with facultative mycorrhizal associations that are present in some cases and not others (called “nonmycorrhizal or AM” in Figure 4.21). In other cases, plants loose the capacity to form mycorrhizas because they are redundant. These include parasites and carnivores, which do not need to acquire nutrients directly from soil (Figure 4.22).


Figure 4.22 Categories and numbers of nonmycorrhizal plants. Most nonmycorrhizal plants occur in specialized habitats where fungal activity is likely to be restricted or have specialized nutritional uptake mechanisms such as carnivory, parasitism or cluster roots. (Data from M. Brundrett, Plant Soil 320: 37-77, 2009)

4.3.1 - Main types of mycorrhizas

Mycorrhizal associations are classified according to the way in which the fungi interact with the host plant root, in particular, the structure of the interface that forms between host cells and fungal hyphae. This classification leads to a number of distinct types of mycorrhizal association, as defined in Table 4.1. However, only two of these are widely distributed in the plant kingdom: arbuscular mycorrhizas (AM) and ectomycorrhizas. Orchid and ericoid mycorrhizas are confined to genera within the Orchidaceae and Ericaceae families, respectively.

Mycorrhizal types generally form with a characteristic group of plant species, but there are occasional examples of overlap, such as many Australian plants in the families Fabaceae and Myrtaceae, which have both arbuscular mycorrhizas and ectomycorrhizas. Arbuscular mycorrhizas occur in a vast array of herbaceous genera. In fact, as shown in Figure 4.21 above, some 75% of all plant species form arbuscular mycorrhizas, including most major crop species, that is, all cereals and most grain legumes and pasture legumes.

Table 4.1 shows that the main types of mycorrhizas differ in host preference and in the structures they form during association with the host root, but they are similar in the ways by which they enhance host plant nutrition. Each type of mycorrhiza can be formed by many species of fungi and a single root may often be colonised by more than one species.

Plants with arbuscular mycorrhizas are common in most ecosystems, but are more likely to be dominant in regions of relatively high mean annual temperatures and rates of evapotranspiration, where phosphorus availability is often the major limiting factor for plant growth. However, in some soils with very low phosphorus availability, nonmycorrhizal plant species with cluster roots may be locally dominant and these plants seem to be more efficient at obtaining phosphorus from these soils than mycorrhizal species (e.g. south west Western Australia).

Ectomycorrhizas are most common in tree species, but also occur in some shrubs. In the Northern hemisphere, ectomycorrhizal associations are typically dominant in boreal forests where temperatures and evapotranspiration are relatively low, leading to slow rates of decomposition and accumulation of plant litter in soil and low nitrogen availability. However, ectomycorrhizal plants are also dominant or co-dominant in many other temperate forests, as well as some tropical and subtropical areas, where soil properties are not substantially different from habitats where only arbuscular mycorrhizal plants occur.

Each type of mycorrhizal association has evolved separately to enhance growth and survival of both the host plants and the mycorrhizal fungi. While the primary role of these associations is to increase nutrient supply to the host plant, mycorrhizas have also been shown under some circumstances to enhance plant water status, confer protection against root pathogens, contribute to soil structure through hyphal binding of soil particles and other processes, and render plants less susceptible to toxic elements. The relative importance of these secondary roles is very difficult to determine since they are difficult to separate from nutritional benefits to plants in experiments and they will not be considered in detail here.

4.3.2 - Development and structure of mycorrhizas


Figure 4.23 Schematic diagram of an arbuscular mycorrhiza. (Courtesy M. Brundrett)

Arbuscular mycorrhizal associations are formed by fungi from division Glomeromycota, class Glomeromycetes. DNA sequence evidence shows they are closely related to the Zygomycota. These fungi simultaneously exist in both the soil and roots, with different forms of hyphae in each environment, as shown in the diagram below (Figure 4.23).

During the infection process, fungal hyphae penetrate the epidermal cell layer, often forming distinctive large hyphae within the root at the point of entry (Figure 4.24). From the entry point, hyphae then spread through the root cortex by growing either through the intercellular spaces or from cell to cell by penetrating the cell walls. Hyphae do not, however, penetrate the endodermis or enter the stele.


Figure 4.24 Left, Root of clover (Trifolium subterraneum) colonised by an arbuscular mycorrhizal fungus. The fungus has formed thick entry hyphae in the epidermis before spreading through the cortex cells, forming an arbuscule (A) in many cortex cells and some vesicles (V). Roots were cleared (to make them transparent) and then stained with Trypan blue. Right, Root of leek (Allium porrum) colonised by indigenous mycorrhizal fungi showing hyphae, arbuscules and many large vesicles. (Photographs courtesy M. Brundrett)

Within individual cortex cells, hyphae may form a distinctive structure called an arbuscule. From the base of each arbuscule, hyphae repeatedly branch, becoming thinner and thinner as they do so (Figure 4.25). The host cell plasma membrane is never penetrated by the fungus.


Figure 4.25 Longitudinal view of a mature arbuscule of an arbuscular mycorrhizal fungus which has formed in a root cortical cell of leek (Allium porrum). Roots were cleared, stained with Chlorazol black E and viewed with interference contrast microscopy. (Photograph courtesy M. Brundrett)

Thus in a cell with an arbuscule, the host cell plasma membrane remains intact and functional, but proliferates to surround the arbuscular branches (Figure 4.26). The highly-branched nature of arbuscules is thought to increase the surface area to volume ratio of the host plant plasma membrane by up to 20-fold, relative to unoccupied root cells, thus providing an extensive interface across which nutrient exchange can take place (Figure 4.26).


Figure 4.26 A transverse view of cortex cells in frozen roots of white clover (Trifolium repens) colonised by indigenous arbuscular mycorrhizal fungi. In uncolonised cells (UC), the plasma membrane is so closely aligned to the cell wall that it cannot be distinguished and what can be seen inside the cell are dots and lines formed by solutes frozen in the cell vacuole. However, some cells contain the many small hyphae of mature arbuscules (Arb) which have the plasma membrane encasing them and large hyphae are evident occupying most of the intercellular spaces surrounding these cells (arrows). Roots were frozen in liquid nitrogen and viewed using cryo-scanning electron microscopy. (Photograph courtesy M. McCully)

When we consider that all fungal biomass was built using plant carbon, it becomes evident that considerable carbon is needed to maintain the symbiosis.

Arbuscular mycorrhizal fungi store the carbon they obtain from the host plant root primarily in the form of lipids. Lipids are particularly dense in vesicles and spores, which also act as inoculum. Vesicles and spores may form within or outside of roots and often develop most prolifically when roots begin to senesce. The morphology of the fungal infection, particularly the vesicles and spores, differs with the species of fungi.

Ectomycorrhizal symbioses are formed primarily by higher fungi in the Basidiomycotina and Ascomycotina, which form mycorrhizas with the short lateral roots of trees (Table 4.1). Unlike arbuscular mycorrhizas and ericoid mycorrhizas, hyphae of ectomycorrhizal fungi do not normally penetrate host cell walls. Rather, they form an entirely extracellular interface, with highly branched hyphae growing between epidermal or cortical cells, forming a network known as the Hartig net (Figure 4.27).


Figure 4.27 Schematic diagram of an ectomycorrhiza showing structures visible in the soil at three levels of magnification (Diagram courtesy M. Brundrett)

In Gymnosperms such as Pinus, the Hartig net may extend through most of the root cortex, but in most Angiosperms it is confided to the epidermis (Figure 4.28). In both cases, the highly branched hyphae of the Hartig net provide a substantial surface area for nutrient exchange between the fungus and the plant. Ectomycorrhizas are further differentiated from the other mycorrhizal types by the fact that the fungus usually forms a dense hyphal mantle around each short lateral root, greatly reducing its contact with the soil (e.g. Figure 4.28).


Figure 4.28 Transverse section of ectomycorrhizas showing labyrinthine Hartig net hyphae (arrows) in roots of Pinus sp. (left) and Populus sp. (right). Fungal hyphae are structurally modified, making intimate contact with root cortex (C) (left) or epidermal cells (E) (right) and enabling exchange of resources through the interface between fungus and host. A mantle of fungal hyphae (M) surrounds both roots. (Photographs courtesy M. Brundrett)

Orchid mycorrhizal associations consist of coiling hyphae within cells of a root or stem (Figure 4.29). The most common fungi involved are members of the Rhizoctonia alliance, but ectomycorrhizal fungi are also found in some orchids, especially in achlorophyllous species lacking photosynthesis. A key feature of orchid mycorrhizas is the capacity of fungi to germinate the tiny seeds of orchids to form tiny protocorms which lack roots or leaves. It is thought that orchids start out by exploiting fungi, but then may develop more mutualistic associations as they grow larger and develop leaves.


Figure 4.29 Orchid mycorrhizal association in the underground stem of an Australian greenhood orchid (Pterostylis sanguinea). This cross section is stained by Trypan blue and shows individual cells filled with densely packed hyphal coils (arrow). (Photograph courtesy M. Brundrett)

Ericoid mycorrhizal fungi (largely ascomycetes) form an interface within cells, consisting of dense hyphal coils which are surrounded by host plasma membrane which is similar to orchid mycorrhizas (Figure 4.30). Many members of the Ericaceae host these associations in very fine lateral roots called hair roots, which are only a few cells wide (Figure 4.30).


Figure 4.30 Several hair roots of Leucopogon verticillata, an Australian member of the Ericaceae, with nearly every cell containing intracellular hyphal coils of an ericoid mycorrhizal fungi (arrows). Many soil hyphae can also be seen leaving the hair roots. Hair roots stained with Chlorazol black E and viewed with interference contrast. (Photograph courtesy M. Brundrett)

4.3.3 - Functional aspects of mycorrhizas

The association between fungus and plant delivers nutrients to the host plant via: (a) mobilisation and absorption by fungal mycelia in the soil; (b) translocation to the fungus–root interface within the root and (c) transfer across the fungus–root interface into the cytoplasm of root cells. As shown in Figure 4.31, both roots and mycorrhizas can absorb nutrients such as phosphorus from the soil, so plants with highly branched fine roots and long root hairs are less likely to benefit substantially from mycorrhizal associations.


Figure 4.31 Diagrammatic summary showing the impact of roots hairs or arbuscular mycorrhizal fungal hyphae on phosphorus uptake from the soil. Compare the upper and lower pairs of drawings to see how soil hyphae increase the size of phosphorus depletion zones in soil much more if plants lack highly branched roots with long root hairs. (Based on Brundrett et al. 1996)


Figure 4.32 Root hairs of an Australian sundew (Drosera erythrorhiza), a carnivorous plant with nonmycorrhizal roots that have extremely long root hairs (1 mm) in relation to the diameter of the root. (Photograph courtesy M. Brundrett)

The majority of plants in natural ecosystems have relatively thick and unbranched roots without long root hairs so are likely to be highly dependent on mycorrhizal associations in the soils where they normally grow. Some crop and garden plants, such as many grasses and members of the Brassicaceae have long root hairs so tend to benefit less from mycorrhizas or have nonmycorrhizal roots (e.g. Figure 4.32).

(a) Nutrient uptake from the soil by the fungi

In addition to hyphae in direct contact with the root surface, all mycorrhizal fungi produce soil hyphae (extramatrical mycelium) which extend into the surrounding soil. Both arbuscular mycorrhizal and ectomycorrhizal fungi can produce copious soil hyphae, that extends well beyond the nutrient depletion zone for immobile nutrients around individual roots and display a complex architecture that renders them an efficient nutrient-collecting and transport network (Figure 4.33). The soil hyphae of many ectomycorrhizal fungi form hyphal aggregations, known as mycelial strands and rhizomorphs, that play a major role in transport of inorganic nutrients or photoassimilates.


Figure 4.33 Abundant mycelium (M) of Scleroderma forms a sheath (S) around roots of an eucalypt and explores surrounding soil (E). These ectomycorrhizas benefit the host through enhanced nutrient uptake (especially phosphorus) from surrounding soil. (Photograph courtesy I. Tommerup)

Extramatrical mycelium is also primarily responsible for spread of the association to new roots and translocation of energy from the plants for fungal reproduction. Fungi forming mycorrhizal associations can also spread by germination of wind or animal dispersed spores and, in some cases, from old root pieces.

In arbuscular mycorrhizas, fine highly branched soil hyphae (diameter 1–5 µm) provide surface area for nutrient absorption, while larger diameter hyphae (up to 10 µm) form a transport network in the soil for moving solutes from bulk soil to the root (Figure 4.34). Absorption of phosphate by the fungus is maximised by the action of a high-affinity transporter which is expressed only in the soil hyphae of arbuscular mycorrhizal fungi during symbiosis with the plant. The fungi take up inorganic phosphate and quickly convert it to polyphosphate, a macromolecule where the charge of the phosphate ions is balanced by cations including those of potassium and magnesium. Polyphosphate allows phosphorus to be transported to the plant without affecting hyphal osmotic balance. For instance, within the root, concentrations of phosphorus in hyphae may be up to 350 mM, but plant cell vacuoles generally have < 10 mM. Once the polyphosphate reaches an arbuscule in the plant root it is converted back to phosphate and released into the peri-arbuscular space where it is absorbed by the host plant. Active transporters in the host cell plasma membrane maintain a concentration gradient across the plant-fungus interface.


Figure 4.34 Soil hyphae of an arbuscular mycorrhizal fungus growing through the surrounding rhizosphere soil and forming spores. Such hyphae ramify through the soil and likely influence soil chemistry and microbial functioning. Root stained with Chlorazol black E. (Photograph courtesy M. Brundrett)

Many experiments have demonstrated a relationship between arbuscular mycorrhizal infection and improved plant phosphorus status, particularly under glasshouse and laboratory conditions (Figure 4.35). Arbuscular mycorrhizal fungi do not appear to have access to sources of soil phosphorus that are otherwise unavailable to nonmycorrhizal roots. Thus, increased plant absorption in the presence of arbuscular mycorrhizal fungi of phosphorus, nitrogen and other macronutrients such as calcium and sulphur, and micronutrients including zinc and copper, seems to primarily reflect the increased absorptive surface of the soil hyphae. However, soil hyphae also provide a conduit for rapid transport of carbon from plants into soil and there is evidence that hyphal exudation may promote breakdown of organic nutrient sources by other microorganisms (see below). Note that the effects of individual species or strains of fungi on plant nutrition will vary, in part due to different morphology of soil hyphae. Under some circumstances, the presence of arbuscular mycorrhizal fungi may decrease plant growth especially in heavily fertilised crop plants.


Figure 4.35 The typical growth response for an Australian Cassia species to inoculation with arbuscular mycorrhizal fungi. The greater shoot dry weight of the inoculated plants is due to the fungi enhancing plant uptake of phosphorus. There is no benefit for the plant from the fungi at the lowest level of phosphorus as the fungi are also likely limited by phosphorus, but benefit is substantial at low-intermediate phosphorus levels. (Based on Brundrett et al. 1996, data courtesy of David Jasper and Karen Clarke)

The soil hyphae of ectomycorrhizal fungi increase the absorptive area of a root system substantially, extending the volume of soil explored by the host plant and consequently the quantity of minerals available. Ectomycorrhizal fungi, however, use additional strategies to enhance nutrient acquisition. Some secrete extracellular proteinases, peptidases, phosphomonoesterases and phosphodiesterases that effectively hydrolyse organic nitrogen and phosphorus sources to liberate some nitrogen and phosphorus compounds which can be absorbed by the fungi. Some ectomycorrhizal fungi produce hydrolytic enzymes within the cellulase, hemicellulase and lignase families that may facilitate hyphal entry to moribund plant material in soil and access to mineral nutrients sequestered therein. In these ways, ectomycorrhizal fungi short-circuit conventional nutrient cycles, releasing nutrients from soil organic matter with little or no involvement of saprotrophic organisms. Ectomycorrhizal fungi also release siderophores capable of complexing iron and oxalate to improve potassium uptake and have also been implicated in promoting weathering of rocks to release mineral nutrients for plants.

(b) Carbon uptake from the plant by the fungi

Arbuscular mycorrhizal fungi depend completely on the host plant for carbon and are unable to grow without being associated with a host plant. This has made the culture of these fungi difficult and proved a significant barrier to development of cheap technologies for inoculation (as might be desirable in land rehabilitation or agriculture) on a large scale. Transfer of carbon from the host to an arbuscular mycorrhizal fungus likely takes place in the arbuscule where the plant releases simple sugars (hexoses) which are absorbed by the fungi. These sugars are rapidly converted into trehalose, glycogen and lipids. The lipids and, to a lesser extent, glycogen are transported to the soil hyphae. Once in the soil hyphae, lipids are progressively broken down into hexoses and trehalose and used to fuel the growth of the fungus. As the lifecycle of the fungus progresses, large amounts of lipid are stored, particularly in vesicles and spores which may be inside or outside of the roots.

In contrast to arbuscular mycorrhizal fungi, ectomycorrhizal fungi can utilise carbon substrates other than those provided by the host plant. It seems that most ectomycorrhizal fungi have some ability to use lignin and cellulose, along with various other substrates including starch, glycogen and sugars such as glucose. Ability to utilise various substrates differs among fungal species. As a result of these abilities, ectomycorrhizal fungi are able to be isolated and grown in culture. For ectomycorrhizal fungi associated with host roots, sucrose is thought to be hydrolysed in root cell walls and glucose to be then absorbed by hyphae from the interface apoplasm.

It has been estimated that 20-50% of plant photosynthate is allocated to mycorrhizal fungi, much of which is allocated to soil hyphae. The soil hyphae of the fungi exude carbon compounds which will influence soil processes including the growth, composition and function of the soil microbial community. Recent research suggests that roots and mycorrhizas may differentially affect soil carbon pools. Thus, overall, the fungi provide a significant pipeline for the movement of carbon from the plant shoot into the soil and may greatly influence soil processes and microbial activity both within and away from the rhizosphere. Indeed, it is now thought that the fungi may significantly influence the global carbon cycle (e.g. their cell walls include some components that are very slow to decompose in soils). In addition, some compounds exuded by the soil hyphae of mycorrhizas, such as glomalin, play an important role in maintenance of soil structure through gluing together soil particles, especially in sandy soils. In addition, colonisation can change the amount and composition of compounds exuded by roots. For instance, the presence of arbuscular mycorrhizal fungi can result in the amount of carboxylates in the rhizosphere being reduced by 50% or more (Figure 4.36). Carboxylates are low molecular weight organic anions which are thought to play a role in release of highly sorbed phosphorus into forms that plant roots or the hyphae of arbuscular mycorrhizal fungi can absorb. Hence, the presence of the fungi enhances the ability of the host to access orthophosphate, but perhaps at the expense of its ability to release phosphorus from sorbed sources.


Figure 4.36 Amount of carboxylates in the rhizosphere of 12 species of Kennedia grown with and without mycorrhizal fungi. (Adapted from Ryan et al. 2012)

Overall, there are many fascinating complexities to the relationships among host plants, fungi, nutrients such as phosphorus and nitrogen, carbon and the soil microbial community. For instance, plants make “trade-offs” among nutrient acquisition strategies, probably due to the carbon costs of each strategy. For example, colonisation by arbuscular mycorrhizal fungi often results in a reduction in root / shoot ratio, and root hair production and fine root production tends to be greater in nonmycorrhizal plants. While there are substantial costs to plants in supporting mycorrhizal associations, the cost of producing roots that function well without them seems to be even greater. Overall, the best evidence that mycorrhizal roots are more efficient than nonmycorrhizal roots is provided by the data in Figures 4.21 and 4.22. This global dataset shows that mycorrhizal plants are normally dominant in ecosystems, with the exception of habitats where conditions are likely to suppress fungal activity (e.g. waterlogged, saline, or very cold soils and epiphytic habitats).

Case Study 4.3 - Regulation of legume nodule numbers

Brett J Ferguson and Peter M Gresshoff, Centre for Integrative Legume Research, University of Queensland, Australia

Forming nodules and supplying rhizobia with food for nitrogen fixation are energy demanding processes for the host plant. As a result, legumes have developed innate mechanisms to balance their need to acquire nitrogen with their ability to expend energy (reviewed in Ferguson et al. 2010; Reid et al. 2011).

Nitrogen availability: why form nodules when there is plenty of nitrogen available?

When ample nitrogen is available in the soil, legume plants require fewer nodules to meet their nitrogen demands. Accordingly, they have an inbuilt mechanism in the root to detect the nitrogen content of the surrounding soil. Nitrogen-based compounds, such as nitrate, trigger the production of a small, hormone-like peptide signal, called Nitrate-Induced CLE 1 (NIC1) in soybean. NIC1 is predicted to be perceived by the receptor, Nodule Autoregulation Receptor Kinase (NARK). This perception results in the production of a novel inhibitor signal that acts locally in the root to prevent further nodulation events.

Stress: nodulation is a luxury for a stressed plant.

Nodulation is reduced in plants experiencing stress. This likely helps the plant to conserve its resources for combating the stress and for all-important seed development. To date, a number of stress-related factors have been found to inhibit nodule formation locally in the root, including ethylene, salicylic acid and various reactive oxygen species (reviewed in Ferguson and Mathesius 2014). Acidic soil conditions also reduce nodulation, with low pH also causing elevated soil Al3+ levels that negatively affect root growth.

Autoregulation of nodulation: too much of a good thing is not good.

Less than 10% of rhizobia infection events result in the formation of a fully-developed nodule. This is largely due to the Autoregulation Of Nodulation (AON) mechanism that the host plant uses to control its nodule numbers (reviewed by Ferguson et al. 2010; Reid et al. 2011). Mutant legume plants lacking a functional AON pathway are unable to regulate their nodule numbers and as a result exhibit a supernodulation phenotype (i.e. they develop an excessive number of nodules, with up to 25,000 per plant scored, Figure 1). When these supernodulating mutants are induced to form nodules, they are typically reduced in stature and often yield about 20 – 30% less, likely a direct result of their resources being used to form excess nodule structures (Figure 1).


Figure 1 Legume nodulation and autoregulation. A, Soybean plants grown in nitrogen-poor conditions. Wild-Type (WT) plants form functional root nodules when inoculated with compatible, nitrogen-fixing, Bradyrhizobium japonicum. Non-nodulating mutant plants are unable to form nodules (nod-) and supernodulating mutant plants (nod++) are unable to regulate the number of nodules they form. B, Roots of wild-type (WT) and supernodulating mutant (nod++) common bean plants exhibiting mature nodule structures.

The AON process is triggered within hours of rhizobia inoculation. It involves long-distance signalling between the root and shoot, commencing with the production of a root-derived signal (Figure 2). Recent work has indicated that this signal is a CLE peptide hormone, highly similar to NIC1. In soybean, two candidate CLE peptide signals have been identified and are called Rhizobia Induced CLE1 (RIC1) and RIC2. The use of grafting and over-expression experiments demonstrated that RIC1 and RIC2 travel to the shoot, likely via the xylem. They are thought to be perceived in the shoot by the same receptor kinase that detects NIC1 in the root, namely NARK, possibly in combination with other receptors, such as CLAVATA2 and KLAVIER. Indeed, this has now been confirmed for orthologous peptides of Lotus japonicus, called LjCLE-RS1 and 2, which travel in the xylem and are perceived by the orthologue of NARK in L. japonicus, HAR1 (Okamoto et al., 2013). The peptides are post-translationally modified with three arabinose sugars attached to a central proline residue, likely by an arabinotransferase encoded by the NOD3/RDN1 gene in Pisum sativum and Medicago truncatula (Ogawa-Ohnishi et al. 2013). Key domains and amino acid residues of soybean RIC1 that are required for effective suppression of nodulation have also been identified. Perception of either RIC1 or RIC2 triggers the production of a new signal, called the Shoot Derived Inhibitor (SDI). SDI is subsequently transported down to the root where it acts to inhibit further nodulation events (Figure 2).


Figure 2 Regulation of legume nodule development. Legume roots exude flavonoid molecules, which attracts compatible rhizobia and triggers them to produce a Nod factor signal. Stress and Nitrogen locally inhibit nodule development. Nitrogen triggers the production of a CLE peptide, called GmNIC1 in soybean, that acts through the GmNARK receptor to suppress nodulation. The Autoregulation Of Nodulation (AON) acts systemically through the shoot. CLE peptides, called GmRIC1 and GmRIC2 in soybean, are produced in the root in response to the first nodulation events. These signals are transported to the shoot where they are perceived by GmNARK, which triggers the production of a shoot-derived inhibitor (SDI) signal that travels to the root to prevent further nodulation.

Although it has not yet been identified, SDI is reported to be a small, heat-stable, Nod factor- and NARK-dependent molecule that is not likely an RNA or protein (Lin et al. 2010) and has been recently been proposed to be the classical phytohormone, cytokinin (Sasaki et al. 2014). Additional factors acting downstream of SDI include Too Much Love, a Kelch-repeat transcription factor whose role in the AON process is yet to be fully defined.

An interesting finding is that of Wang et al. (2014), who suggest that the AON pathway may involve a microRNA after the tentative cytokinin signal. Specifically miR172c was shown to negatively regulate the transcript abundance of a gene (GmNNC1) encoding a transcription factor, being part of the AP2 family. GmNNC1 negatively targets the early nodulin gene ENOD40, needed for nodulation progress in soybean, Medicago truncatula, and Lotus japonicus. It now appears critical to connect the function of a peptide-activated receptor (GmNARK) with the reported cytokinin signal and the subsequent negative regulation cascade described by Wang et al. (2014).


Ferguson BJ, Indrasumunar A, Hayashi S et al. (2010) Molecular analysis of legume nodule development and autoregulation. J Integr Plant Biol 52: 61-76

Ferguson BJ, Mathesius U (2014) Phytohormone regulation of legume-rhizobia interactions. J Chem Ecol 40: 770-790

Lin Y-H, Ferguson BJ, Kereszt A, Gresshoff PM (2010) Suppression of hypernodulation in soybean by a leaf-extracted, NARK- and Nod factor-dependent small molecular fraction. New Phytol 185: 1074-1086

Okamoto S, Shinohara H, Mori T et al. (2013) Root-derived CLE glycopeptides control nodulation by direct binding to HAR1 receptor kinase. Nature Comms 4, doi:10.1038/ncomms3191

Ogawa-Ohnishi M, Matsushita W, Matsubayashi Y (2013). Identification of three hydroxyproline O-arabinosyltransferases in Arabidopsis thaliana. Nature Chem Biol 9: 726-730

Reid DE, Ferguson BJ, Hayashi S et al. (2011) Molecular mechanisms controlling legume autoregulation of nodulation. Ann Botany 108: 789-795

Sasaki T, Suzaki T, Soyano T et al. (2014). Shoot-derived cytokinins systemically regulate root nodulation. Nature Comms 5, doi:10.1038/ncomms5983.

Wang Y, Wang L, Zou Y et al. (2014) Soybean miR172c targets the repressive AP2 transcription factor GmNNC1 to activate GmENOD40 expression and regulate nodule initiation. Plant Cell 26: 4782–4801

4.4 - Symbiotic nitrogen fixation

Ulrike Mathesius, Research School of Biology, Australian National University

Nitrogen is an important nutrient for all plants. While there is an abundance of nitrogen in the atmosphere, plants are unable to convert N2 into a usable form. Fixation of nitrogen gas into ammonia is an ability restricted to nitrogen-fixing bacteria, which contribute most of the inorganic nitrogen to the Earth’s nitrogen cycle.

This chapter explores the importance, evolution and regulation of biological nitrogen fixation, especially of bacteria that have evolved symbiotic associations with higher plant plants. The symbiosis of legumes with nitrogen-fixing soil bacteria called rhizobia has become a model for our understanding of plant-microbe interactions.


Figure 4.38 An infected nodule of Medicago truncatula. Rhizobia are shown as green inside the large nodule, which is outlined in blue. The root is outlined in red. (Photograph courtesy U. Mathesius)

Research over the last decade and beyond has revealed major principles and molecular mechanisms of how plants have evolved to recognise their symbiotic partners, how they allow them entry into their root systems, how nutrients are exchanged between the partners and how the symbiosis is controlled systemically to balance demand and supply.

4.4.1 - Acquiring atmospheric nitrogen

Plant growth is frequently limited by nitrogen. Plants generally obtain nitrogen from soil reserves of nitrate or ammonium (so-called mineral nitrogen) but these reserves are often scarce.

Natural ecosystems can ‘run down’ with respect to nitrogen through soil leaching and fire. Relative abundance of nitrogen-fixing species will then increase. For example, a walk from east to west across Fraser Island, Queensland, will take you across progressively older and more nitrogen deficient sand dunes, and from rainforest to heathland.

Agriculture and horticulture, with its harvest of nitrogen-rich grains or leaves, removes site nitrogen that must be replaced by further mineralisation of soil nitrogen, import of mineral nitrogen (fertiliser) or fixation of atmospheric nitrogen (N2).

The earth’s atmosphere is rich in N2 (about 78% N2) which is very unreactive, due to its stable triple bond. No known eukaryotes have the ability to fix nitrogen, i.e. to reduce atmospheric nitrogen into a usable form like ammonia (NH3). Hydrogen (H2) will react with N2 at very high temperatures and pressures on a catalyst by the Haber-Bosch process, where the pressures needed are 10–100 MPa, the temperatures are 400–550 ºC, and the catalyst is Fe. Then:

\[ N_{2} + 3H_{2} \rightarrow 2NH_{3} \tag{1} \]

Large quantities of ammonia are produced by this method for industrial and agricultural use. Some nitrogen is also fixed in the atmosphere during lightning strikes (Fowler et al. 2013).

Amazingly, some bacteria have the ability to catalyse this reaction with an enzyme complex called nitrogenase donating at least four pairs of electrons to every N2 molecule to effect reduction to two NH4+ and at least one H2. The reaction takes place at ambient conditions, catalysed by the Fe–Mo-containing enzyme, nitrogenase.

\[N_{2} + 16ATP + 8e^– + 10H^+ \rightarrow 2NH_{4}^+ + H_{2} + 16ADP + 6P_{i} \tag{2}\]

Biological N2 fixation is energetically expensive even though it occurs at ambient conditions — estimates fall between 3 and 7 g carbon respired gram of nitrogen fixed (Layzell 1992). Photoassimilate consumed to support N2 fixation is unavailable for other processes such as growth. Consider a crop fertilised with 140 kg N ha–1. An N2 fixer could replace this fertiliser, but only at a cost of at least 420 kg C ha–1. As most plant dry matter contains 40% carbon, this is equivalent to a loss of one tonne of dry matter per hectare! However, in natural ecosystems where no nitrogen fertiliser is applied, and with rising costs of synthetic nitrogen fertiliser, which consumes about 2% of global fossil fuels annually, biological nitrogen fixation confers a distinct advantage to plants that associate with N2-fixing bacteria (Figure 4.39). In fact, our use of nitrogen fertilisers in agriculture has expanded so enormously that pollution of aquatic habitats by nitrogen run-off has moved beyond safe limits for our planet (Steffen et al. 2015). During the last 30 years, research into biological nitrogen fixation, especially in the symbiosis of legumes with rhizobia, has advanced to a point where transfer of this symbiosis to non-legume crops is now becoming a serious goal (Rogers and Oldroyd 2014).


Figure 4.39 Benefit of nitrogen fixation in legumes depending on N fertilisation. Total nitrogen content (A) and plant biomass (B) of subclover (Trifolium subterraneum) plants grown in a glasshouse for four weeks in the presence and absence of nitrogen fertiliser (10 mM potassium nitrate, ‘N’) and/or the rhizobium symbiont (Rhizobium leguminosarum bv. trifolii, ‘R’). Plants benefit from rhizobial inoculation only in the absence of N fertilizer, as shown by the increases in N content and biomass. Rhizobia confer a similar benefit to the unfertilised subclover plants as a 10 mM nitrate addition does, although this varies with different rhizobial strains and with legume species. (Data modified from C.-H. Goh et al., Plant Cell Environ, 2015)

4.4.2 - A range of N<sub>2</sub>-fixing associations

Many bacterial species from diverse phyla have the ability to fix nitrogen, including many (but not all!) cyanobacteria, actinobacteria and proteobacteria. The nitrogen fixation genes are thought to have spread between distantly related bacteria by horizontal gene transfer of clusters of nitrogen-fixation (nif) genes. Nitrogen-fixing bacteria, also called ‘diazotrophs’, can be free-living in water or on solid substrates like soil or rocks. More than half of the biological nitrogen fixation on earth stems from nitrogen-fixing marine bacteria, the rest from terrestrial sources (Fowler et al. 2013). Terrestrial N2-fixing bacteria are found in the soil, in aquatic, and often extreme, habitats, such as hot springs, and nutrient-poor areas. Much of the action of N2-fixing bacteria happens in soils. Plants can then use nitrogen released by decay of such organisms.

However, some plants have evolved a tighter relationship with N2-fixing bacteria, involving an exchange of carbon and nitrogen between plant host and bacterial partner (Santi et al. 2013). Several different symbioses of this type have evolved independently (Table 4.2), from primitive algae to higher vascular plants. These associations are characterised by active attraction of the symbiotic partner through metabolites or signals from the host, as well as a physical housing of the symbiont inside the host (Delaux et al. 2015). For example, algae can form a symbiosis with N2-fixing cyanobacteria to form lichens, and Bryophytes like Anthoceros harbor N2-fixing cyanobacteria in cavities of their thalli. Roots or leaves of some plants form a loose association with N2-fixing bacteria, with plant exudates used as a carbon source by the bacteria. In the water fern Azolla, the cyanobacterium Anabaena is located in cavities on the underside of modified leaves, with a secretory trichome delivering sugars and absorbing fixed nitrogen. This symbiosis is especially important in rice paddy fields, where Azolla densely covers the water surface and can then be incorporated into the field soil. In other plants, the N2 fixers are located in intercellular spaces of the host plant, as reported for sugarcane. These less intimate associations supply host plants with substantial amounts of nitrogen. While such associations have been explored for other grasses, the likely limitation for substantial nitrogen fixation in these loose associations is the limited amount of carbon that the plant provides for its symbionts.

In more highly developed associations, plants localise the symbiotic association within a modified root or ‘nodule’. In cycads, the microsymbiont Anabaena is located in intercellular spaces of the mid-cortex of short, highly branched, modified roots (Figure 4.40a and b). In another class of symbioses, the actinorhizal plants, the micro-symbiont Frankia (an actinomycete, or filamentous bacterium) is located within the cortical cells of a modified root. This group includes the genera Casuarina, Allocasuarina, Alnus, Datisca and Myrica from eight plant families, all belonging to the Rosid I class, as do legumes (Figure 4.40c and d). These plants often grow in nutrient-poor habitats where nitrogen fixation provides a nutritional advantage. Parasponia, a tropical tree native in New Guinea, is the only non-legume known to form a symbiosis with the rod-shaped bacterium Rhizobium. Unlike legumes, the Parasponia nodule has a central vascular bundle and the microsymbiont is always encapsulated within cellulosic material (termed a ‘persistent infection thread’). In legumes, nodules typically have a central infected zone, and rhizobia are enveloped by plasma membrane-derived vesicles called symbiosomes (Figure 4.40e and f).


Figure 4.40 Nodule anatomy showing: a, A cycad (Macrozamia miquellii) nodule consisting of a central vascular strand (VB) and an infected cortical region (stars); b, A cyanobacterium, a microsymbiont, is located in the intercellular spaces of this infected cortex; c, A nodule of the river-oak (Casuarina cunninghamii) consisting of a central vascular bundle with infected cells in the cortex (arrows) identified by subersation and lignification of their walls (section stained with berberine sulphate and viewed under epi-fluorescence optics); d, Scanning electron micrograph of an actinomycete microsymbiont (a filamentous bacterium) encapsulated within threads (arrow) throughout the plant cytoplasm; e, A legume (Macroptilium lathyroides) nodule consisting of a central infected region with scattered infected cells (arrows) enclosed in a cortex. Vascular strands (VB) are present in the cortex; f, Transmission electron micrograph of a soybean (Glycine max L.) nodule containing a microsymbiont enveloped by plasma membrane to form ‘symbiosomes’ — packets of bacteria within the cell cytoplasm (arrows). Scale bar = 100 µm in a, b, c and e; 5 µm in d and f. (Images courtesy K. Walsh)

4.4.3 - Rhizobium associations

The symbiosis of legumes with rhizobia is the most effective and agriculturally the most important nitrogen-fixing symbiosis. Rhizobia are soil bacteria from the α- and β-proteobacteria (including Burkholderia sp.) that can occur free-living in the soil, but benefit greatly from symbiosis with legume partners, which provide a carbon source, shelter inside a nodule as well as a niche outside the species-rich and competitive zone of the rhizosphere (See previous section on the Soil-root interface). Many species of rhizobia only effectively fix nitrogen inside a nodule (Figure 4.41).


Figure 4.41 Nodules on the root system of a legume. A, Nodules on a root system of a broad bean. B, An infected nodule of Medicago truncatula. Rhizobia express the green fluorescent protein (GFP) and appear in green inside the nodule. New rhizobia infect the tip of the nodule through infection threads (arrow). The blue colour originates from flavonoid autofluorescence in the nodule cortex. Red fluorescence in the main root stems from chlorophyll autofluorescence, as roots were exposed to some light. Magnification bars = 1 cm in A, and 1 mm in B. (Photographs courtesy U. Mathesius)

The ‘precursor’ of symbiosis evolved approximately 100 million years ago, and led to the similar symbioses of actinorhizal plants with Frankia, and the legume symbiosis with rhizobia. Nitrogen fixing symbioses in legumes evolved at a time of relatively high atmospheric CO2. Nitrogen fixation is likely to have conferred an advantage in using this increased CO2 for photosynthesis (Sprent 2007). Phylogenetic reconstruction of symbiotic nitrogen fixation suggests that it evolved once, and was subsequently gained and lost several times in legumes (Figure 4.42; Werner et al. 2014).


Figure 4.42 Origin of root-nodule symbiosis with rhizobia in angiosperms. Angiosperm phylogeny of 3,467 species showing reconstruction of node states. Branches are coloured according to the most probable state of their ancestral nodes. A star indicates precursor origin. Turquoise and yellow band indicate the legumes and the so-called nitrogen-fixing clade, which contains all known nodulating angiosperms. Grey and white concentric circles indicate periods of 50 million years from the present. The positions of some important angiosperms are indicated with drawings (illustrations by Floortje Bouwkamp). (Reproduced by permission from Macmillan Publishers Ltd from G.D.R. Werner et al. Nature Comms 5: 5087, 2014)

Nodules formed by members of the family Leguminoseae have a central zone of infected cells, surrounded by a cortex of uninfected cells. A root vascular strand branches within the cortex of the nodule. This structure is quite distinct from nodules of the cycads or actinorhizal plants, which have a central vascular bundle and an infected cortex (a typical root vascular anatomy). Different legume species display various nodule growth patterns, but they can be roughly classified as either of indeterminate growth (i.e. with an apical meristem and consequent elongated shape) or determinate growth (i.e. a spherical meristem which ceases activity at nodule maturity).

The association between rhizobia and legumes is a controlled infection. Typically, the bacterial partner infects the plant through root hairs, and is then encapsulated by polysaccharide material produced by the host plant, forming infection threads. Infection threads then grow into the root cortex, while bacteria multiply within each thread. Finally, bacteria are released from the infection threads and engulfed by plant cells in a form of phagocytosis. This process results in a bacterium (sometimes several) encapsulated by a plant cell membrane. Encapsulating membranes control the delivery of photoassimilate to bacteria, thus ensuring a symbiotic rather than a parasitic relationship. These units are termed ‘symbiosomes’ (see also later Section 4.4.5)

Evolution of this partnership might be similar to that of other endosymbiotic organelles such as mitochondria and chloroplasts. Perhaps a future step in the evolution of a legume–rhizobium symbiosis will be retention of bacteria within plant cells to create a new organelle! If this were to happen, the legume would no longer be dependent on the presence of a microsymbiont for infection. Cells could maintain a low resident population of the new organelle, like plastids in non-photosynthetic tissue, and allow proliferation under set conditions within nodule structures.

In some legume symbioses, bacteria are not released from infection threads. This character is one of several that distinguish each of the three legume subfamilies Caesalpinoideae, Mimosoideae and Papilionoideae (e.g. cassia, acacia and soybean, respectively). The Caesalpinoideae are largely trees or shrubs, and the few species which nodulate have little nodule mass proportional to plant biomass (Sprent and Raven 1985). In most of the caesalpinoid species that do nodulate, the microsymbiont remains encapsulated in an infection thread throughout the life of a nodule. In some species the infection threads are thin walled, while in others bacteria are released into the cytoplasm. The Papilionoideae is considered the most advanced of the legume subfamilies.

Biological interactions between host plant and bacterium are subtle. Just as legumes vary genetically, so do the rod-shaped bacteria (rhizobia) that infect various legumes. Not all rhizobia are equally infective (able to infect and form nodules) or effective (able to fix N2) on all legumes. An appropriate bacterial partner must therefore be matched genetically with each legume for optimal N2 fixation. Pure cultures of rhizobia are produced commercially, generally in a peat-moss-based medium or as a seed coating, for inoculating legume seed prior to planting.

4.4.4 - The Rhizobium-legume symbiosis is fine-tuned by a molecular dialogue

Over the last 30 years, the molecular, cellular and genetic analysis of the association between legumes and rhizobia has given us a detailed insight into the specificity and regulation of this symbiosis. As a prerequisite for the symbiosis, the symbionts have to first recognise each other in the soil; the plant host then attracts specific rhizobial symbionts that only nodulate one or a few specific hosts; in response, the rhizobia initiate infection and nodule development. On a whole-plant level, the plant strictly regulates the number of nodules on the root system depending on carbon availability relative to nitrogen demand (See ‘Case Study 4.3 - Regulation of nodule numbers’).

The attraction of rhizobia to the root system of its appropriate host is initiated by the exudation of flavonoids by the plant roots. Flavonoids are a diverse class of phenolic compounds that share a common 15-carbon skeleton consisting of two phenyl rings and a heterocyclic ring, but differ in their exact final structure, for example through modification of the flavonoid ‘backbone’ through hydroxylation, methylation or glycosylation. Over ten thousand different flavonoid molecules have been identified from different plant species. Each legume species exudes a specific mixture of flavonoids into the soil that chemotactically attract their rhizobial symbionts (Figure 4.44). Flavonoids have evolved into specific signalling molecules in the symbiosis by binding specifically to a transcription factor, NodD, inside the rhizobia. This binding of the correct flavonoid activates NodD, and allows it to bind to and activate the promoter regions of a large number of nodulation genes. These nodulation genes of rhizobia are responsible for synthesising a specific signalling molecule, the Nod factor, as well as other genes necessary for symbiosis.


Figure 4.44 Examples of flavonoids produced by different legume hosts to activate Nod gene expression in their symbiont. Luteolin is a Nod gene inducer produced by Medicago sp., while daidzein is made by bean and soybean.

Nod factors are lipochitin-oligosaccharides that are composed of a short ‘backbone’ of N-acetylglucosamine units (the same building blocks that are polymerised to chitin in fungal cell walls, insect exoskeletons and crustacean shells). This backbone typically varies in length between 3-6 units, and is additionally ‘decorated’ with chemical substitutions like acetylation, sulfation or glycosylation (Figure 4.45). Importantly, an acyl side chain of specific length and saturation pattern gives each Nod factor molecule its specificity. Different strains of rhizobia produce specific (mixtures of) Nod factor molecules, which are only recognised by their specific hosts. The Nod factor molecule is crucial for nodulation; mutants defective in Nod factor synthesis cannot nodulate their hosts. Nod factor molecules are also partially sufficient for many of the nodulation steps: application of purified Nod factors to the correct hosts initiates root hair curling and nodule development, but will not lead to infection threads or fully formed nodules. For the complete symbiosis to succeed, other signalling molecules from rhizobia are necessary. For example, surface molecules like exopolysaccharides are necessary to evade host defence responses (Figure 4.45), and communication signals like quorum-sensing signals are necessary to coordinate bacteria behaviours during nitrogen fixation.


Figure 4.45 Nod factor and Myc factor signalling. The structure of the predominant Nod factor from Sinorhizobium meliloti, the symbiont of the legume Medicago truncatula, is shown (top), in comparison with the structure of the non-sulphated lipochitooligosaccharide Myc factor produced by the arbuscular mycorrhizal fungus Glomus intraradices (bottom). These molecules induce the symbiosis signalling pathway, that shares a calcium spiking response. There are two different interpretations for the genetic overview of this signalling pathway. These interpretations are an amalgamation of genetic analyses in Lotus japonicus and Medicago truncatula; gene names are indicated for the model legume L. japonicus. In the first interpretation (top), the symbiosis signalling pathway transmits the signal through the transcription factors encoded by nodulation signalling pathway 2 (NSP2), NSP1and RAM1 (required for arbuscular mycorrhization 1), whereas in the second interpretation (bottom), symbiosis signalling occurs in parallel to the signalling through these GRAS domain transcription factors. Names for these genes in M. truncatula are as follows: Nod factor receptor 1 (NFR1) is LysM domain receptor kinase 3 (LYK3); NFR5 is Nod factor perception (NFP); symbiosis receptor-like kinase (SYMRK) is DMI2; POLLUX is DMI1; calcium- and calmodulin-dependent serine/threonine protein kinase (CCAMK) is also known as DMI3; CYCLOPS is interacting protein of DMI3 (IPD3). (Reproduced by permission from Macmillan Publishers Ltd from G.E.D. Oldroyd, Nature Rev Microbiol 11: 252-632, 2013)

Nod factors are recognised by their host by Nod factor receptors. Binding of the correct Nod factor structure activates a signalling cascade in the infected root hairs. A critical part of the signalling cascade is a periodic spiking of calcium concentrations in the root hair (‘calcium spiking’), which starts within minutes of Nod factor perception (Figure 4.45). Calcium spiking activates downstream transcription factors that lead to the activation of cytokinin signaling. The plant hormone cytokinin is necessary, and in some cases sufficient to induce cell divisions in the root cortex, the first step of nodule initiation.


Figure 4.46 Rhizobial and mycorrhizal colonization. a, Flavonoids released by the legume root signal to rhizobia in the rhizosphere, which in turn produce Nod factors that are recognized by the plant. Nod factor perception activates the symbiosis signalling pathway, leading to calcium oscillations, initially in epidermal cells but later also in cortical cells preceding their colonisation (See Figure 3.21). Rhizobia gain entry by infection root hair cells that grow around the bacteria attached at the root surface, trapping the bacteria inside a root hair curl. Infection threads are invaginations of the plant cell that are initiated at the site of root hair curls and allow invasion of the rhizobia into the root tissue. The nucleus relocates to the site of infection, and an alignment of ER and cytoskeleton, known as the pre-infection thread, predicts the path of the infection thread. Nodules initiate below the site of bacterial infection and form by de novo initiation of a nodule meristem in the root cortex. The infection threads grow towards the emergent nodules and ramify within the nodule tissue. In some cases, the rhizobia remain inside the infection threads, but more often, the bacteria are released into membrane-bound compartments inside the cells of the nodule, where the bacteria can differentiate into a nitrogen-fixing state. b, Strigolactone release by the plant root signals to arbuscular mycorrhizal fungi (AMF) in the rhizosphere. Perception of strigolactones promotes spore germination and hyphal branching. AMF produce mycorrhizal factors (Myc factors), including lipochitooligosaccharide (LCOs) and, possibly, signals that activate the symbiosis signalling pathway in the root, leading to calcium oscillations. AMF invasion involves an infection peg from the hyphopodium that allows fungal hyphal growth into the root epidermal cell. The route of hyphal invasion in the plant cell is predicted by a pre-penetration apparatus, which is a clustering of ER and cytoskeleton in a zone of the cell below the first point of fungal contact. The fungus colonizes the plant root cortex through intercellular hyphal growth. Arbuscules are formed in inner root cortical cells from the intercellular hyphae. (Reproduced by permission from Macmillan Publishers Ltd from G.E.D. Oldroyd, Nature Rev Microbiol 11: 252-632, 2013. Part b image is modified by permission from M. Parniske Nature Rev Microbiol 6: 736-775, 2008)

One of the most significant findings of recent years has been the discovery that the same early machinery for Nod factor signal transduction is also required to signal a successful interaction between mycorrhizal fungi and plants (Figure 4.45), and both symbioses share similar mechanisms of bacterial or fungal invasion (Figure 4.46). Mycorrhizal fungi have evolved symbioses with the majority of plant plants much earlier (~400 MYA) than the emergence of the Rhizobium-legume symbiosis (~100 MYA) (see previous section 4.3 on mycorrhiza). Mycorrhizal fungi produce signals that are structurally closely related to Nod factors, and these have been termed ‘Myc factors’. Myc factors are thought to be perceived by different receptors to the Nod factor receptors. Despite some of the shared signals, including calcium spiking, activated by both Myc factors and Nod factors, the later responses in the root differ, leading either to nodulation or to mycorrhization (Figure 4.45). It is still not known what enables legumes to form nodules in response to Nod factors, but other plants not; however, the knowledge that mycorrhizal signalling is part of the response machinery present in all plants that form mycorrhizal symbioses might make it easier to engineer non-legume crops with the ability to form rhizobial symbioses (Rogers and Oldroyd 2014).

Having established the initial contact with the legume host, rhizobia invade the young, just emerging root hairs of their legume partners. In some species, infection can occur intercellularly, and this may have been also the initial mode of infection at the early evolutionary stage of the symbiosis (Held et al. 2014). The infected root hairs curl around small colonies or single cells of rhizobia to form a typical ‘Sheppard’s crook’. The rhizobia locally digest the cell wall of the root hair and induce the formation of ‘infection threads’, tubular invaginations of the root hair in which the rhizobia multiply and travel towards the cortex of the root. This infection step requires correct Nod factor and exopolysaccharide structures from rhizobia, otherwise infection threads are aborted by the plant through defence responses (Figure 4.47). From the infection threads, rhizobia are released into cortical cells as small vesicles, in which rhizobia remain separated from the plant cytoplasm by the plasma membrane. In these vesicles, called symbiosomes, rhizobia differentiate into bacteroids, the nitrogen-fixing forms of rhizobia (Figure 3.49).


Figure 4.47 Successful and aborted infection threads. a, A successful infection thread of compatible rhizobia on their host. Rhizobia are expressing the Green Fluorescent Protein. b, An aborted infection thread as a result of infection with a rhizobium strain unable to synthesise the correct exopolysaccharides, leading to a defence response. (Reproduced by permission from Macmillan Publishers Ltd from K.M. Jones et al. Nature Rev Microbiol 5: 619-633, 2007)

Concurrent with the growth of the infection thread towards the root cortex, Nod factors produced by the invading rhizobia trigger the re-initiation of cell division in the cortex. These divisions are activated through cytokinin and auxin gradients that the plant forms in response to Nod factors (Desbrosses and Stougaard 2011). In some legumes and actinorhizal plants, the invading rhizobia do not initiate nodules de novo from cortical cells, but target an emerging lateral root primordium that is then ‘converted’ into a nodule.

One surprising recent discovery has been the identification of rhizobia that nodulate legumes in the absence of Nod factors. Sequencing of a large number of Rhizobium species uncovered rhizobial genomes that do not contain any of the canonical nodulation genes that would be required to make Nod factors. It is still unclear what the signal is that these rhizobia use to regulate infection and nodule formation. Similarly, Frankia symbionts do not produce Nod factors, yet use some of the same symbiotic (SYM) signal transduction cascade as legumes. Unravelling the identity and mode of action of these unknown nodulation signals will be a rewarding future challenge.

4.4.5 - Linking functions with structures

(a) Protecting nitrogenase from O2

A basic conflict arises in biological N2 fixation: nitrogenase is destroyed by O2, yet aerobic respiration is essential to sustain the high energy demand of N2 fixation. Nitrogen-fixing bacteria must be protected from O2, while a level of aerobic respiration occurs in the host cell cytoplasm. In cycads, cyanobacteria provide their own O2 protection. Nitrogenase is located in specialised cells (heterocysts) which have an O2-impermeable lining of glycolipid. An analogous structure (a vesicle) affords protection to nitrogenase in the microsymbiont Frankia within most actinorhizal nodules. In Parasponia and most caesalpinoid nodules the persistent infection threads provide O2 protection to nitrogenase (Sprent and Raven 1985).

There is one major problem with structures of ‘fixed’ resistance. As respiration rate varies (i.e. O2 flux), O2 concentration inside the structure must also vary: following Fick’s Law of diffusion, O2 flux into the nodule will change in proportion to the O2 concentration gradient at constant resistance. Free water bathing the nodule is equilibrated with the atmosphere (20.8% O2), therefore containing approximately 360 µM O2 in solution, while nitrogenase is destroyed by submicromolar concentrations of dissolved O2. For practical purposes then, an O2 gradient of 360 (outside) to 0 (inside) µM must be maintained. If respiration rate was halved without a change in resistance, the O2 concentration gradient would also halve from 360 to 180 µM. An O2 concentration of 180 µM O2 inside would destroy nitrogenase. So, resistance must vary too.

A legume-nodule cortex copes with variations in respiration rate by providing a variable level of O2 protection (Layzell and Hunt 1990). According to their model, a layer of cells adjacent to the infected zone either lacks radial intercellular spaces (preventing inflow of O2) or has intercellular spaces filled with water. The thickness of this layer could vary under osmotic control to set nodule permeability. Diffusivity of O2 through water is about 10,000 times slower than through air, so flooding of radial air spaces in the nodule cortex would be an effective way of decreasing O2 diffusion into infected tissue.

Any O2 leaking through this cortex can diffuse freely in the intercellular airspaces of infected tissue and dissolve in the cytoplasm of infected cells. O2 gradients which might be expected within infected cells because of rapid bacterial respiration are largely avoided by the presence of leghemo-globin (Lb) (a molecule similar to the hemoglobin in mammalian blood). O2 diffuses to Lb molecules where it is bound to form high concentrations of oxygenated Lb (estimated at 0.7 mM by Bergersen 1982). Effective nodules are pink because of oxygenated Lb (Figure 4.48); indeed this colour change can be used to estimate free O2 concentrations. Soybean nodules seem to regulate the free O2 in infected cells at between 5–60 nM (e.g. Layzell and Hunt 1990). Finally, residual O2 diffuses through the symbiosome to the bacteroids, supporting a level of aerobic respiration.


Figure 4.48 Leghemoglobin in nitrogen fixing nodules. The pink colour in the centre of the nodule (arrows) originates from the presence of leghemoglobin in the nitrogen-fixing zone of mature nodules. Leghemoglobin binds oxygen inside the nodules to protect the activity of nitrogenase. It also transports oxygen. Magnification bar = 1 mm. (Photograph courtesy U. Mathesius)

‘Conventional’ chemistry may not be appropriate when describing O2 movement in cells because O2 molecules in cellular compartments are so scarce. A sphere of 1 µm radius — roughly the size of a mitochondrion or bacterium — containing a solution with 10 nM O2 will contain only 24 molecules of O2.

(b) Carbon supply and nitrogen export

Nodules are metabolically highly active. A typical maximum rate of nitrogenase activity in soybean nodules, as measured by gas exchange (discussed below) is 300 µmol electron pairs g–1 (nodule) h–1. This value is useful to bear in mind when reading the literature about N2 fixation, with low values possibly indicating unhealthy or disturbed plants. As nitrogenase is at best 75% efficient, with respect to N2 fixation (Equation 2), this rate is equivalent to the fixation of some 150 µmol N g–1 (dry weight) h–1. Reduced nitrogen is exported from nodules to the host plant while carbon is imported into the nodule, supporting energy needs of fixation (through respiration) and providing carbon skeletons for packaging nitrogen as an organic molecule.

Photoassimilate (host to nodule) and nitrogen-based resources (nodule to host) must pass through the endodermis of nodule vascular bundles. Radial walls of this endodermis have Casparian bands and tangential walls have relatively few plasmodesmata, so this cell layer restricts apoplasmic and symplasmic flow of carbon into nodules and nitrogen out of nodules.

The transport of solutes, including C and N metabolites but also metal ions, e.g. Fe and Mo required for nitrogenase, are exchanged via both plant and bacterial transporters on the symbiosome and bacterial membrane, and this transport is carefully controlled to balance supply and demand (Figure 4.49; Udvardi and Poole 2013). Not all rhizobia that invade and inhabit legume nodules fix nitrogen efficiently, with the legume host able to restrict C flow to those nodules that do not fix sufficient N.


Figure 4.49 Transport and metabolism in an infected nodule cell. Sucrose from the shoot is converted to malate in the plant and imported across the symbiosome membrane and into bacteroids, where it fuels nitrogen fixation. The product of the nitrogen fixation is then exported back to the plant, where it is assimilated into asparagine (Asn) for export to the shoot (blue arrows). In many legumes, such as soybean, the export products are ureides instead of Asn. The plant must provide metals and ions to the bacteroid, although only some of the transport systems on the symbiosome and bacteroid membranes are defined. Many rhizobia lack the ability to make homocitrate or become symbiotic auxotrophs for supply of branched-chain amino acids and become dependent on the plant. (Reproduced with permission by Annual Reviews from M. Udvardi and P.S. Poole, Annu Rev Plant Biol 64: 781-805, 2013)

The concentration of nitrogenous solutes in the xylem apoplasm causes a hydrostatic pressure to develop, and this results in a mass flow of nodule xylem sap to adjacent roots. The water that accompanies sucrose entering the nodule as phloem sap is re-exported with assimilated nitrogen in the xylem. Nodules are thus analogous to ‘glands’ that secrete nitrogenous compounds.

4.4.6 - Measuring N<sub>2</sub> fixation

Rates of N2 fixation can be measured by a number of techniques to address questions of nodule efficiency and nitrogen cycling in agricultural and natural plant systems. Nitrogenase is pivotal for initial reduction of N2 but this same enzyme will also reduce acetylene (C2H2) to ethylene (C2H4). Acetylene is an effective competitor with N2 for nitrogenase so the rate of C2H4 synthesis is proportional to nitrogenase activity. Acetylene reduction gives an instantaneous estimate of the N2 fixation rate. Another instantaneous technique requires flushing nodulated roots with an argon : oxygen gas mixture (79:21) to displace all N2. All electron flux through nitrogenase is then diverted to the reduction of protons to H2 rather than N2 to NH4+ (Equation 2). The rate of H2 evolution by roots can thus be used to estimate nitrogenase activity.

Alternative approaches to ‘instantaneous’ estimates of N2 fixation provide an integrated rate of fixation over periods of hours or days. The proportions of inorganic and organic nitrogen compounds in xylem sap are affected by the ratio of inorganic nitrogen taken up to symbiotic N2 fixation; this can be exploited in genera of legumes in which amides and ureides are major products of N2 fixation. Soybean, for example, exports less than 10% of nitrogen to shoots in the form of ureides when supplied nitrate but more than 80% when all nitrogen is biologically fixed. Thus, relative ureide levels in sap give an estimate of N2 fixation.

Many experiments now rely on 15N-based techniques to obtain an integral of fixation over the life of a plant. These techniques rely on a difference in ratio of the stable isotopes of nitrogen (15N and 14N) in soil and atmosphere (Figure 4.50). The soil must be enriched in 15N relative to the atmosphere — either naturally (the process of denitrification causes a fractionation of the two isotopes, leaving the soil enriched in 15N) or by artificial 15N addition. The N2-fixing plant of interest is sampled, together with an adjacent non-N2-fixing plant (e.g. grass) whose 15N enrichment represents that of soil nitrogen. 15N enrichment in digested plant material and soil is analysed isotopically in a mass spectrometer and contribution of biological N2 fixation calculated.


Figure 4.50  Basis of the natural abundance method for assessing the contribution of N2 fixation to legume nutrition. This method entails measuring plant 15N/14N ratio by mass spectrometry. Natural differences in 15N/14N ratio between soil and atmospheric nitrogen are exploited. Legumes to the left and right of the figure each have a unique source of nitrogen, while a test plant in the middle relies on both fixed nitrogen and soil inorganic nitrogen. Plants (left) denied a source of inorganic nitrogen (e.g. nitrate) fix atmospheric nitrogen and therefore have low 15N/14N ratios. Plants without nodules (right) take up only soil-derived nitrogen and are enriched with 15N (high 15N/14N ratios). 15N 'signatures' of these two sets of plants can be used to estimate the relative contributions of soil and atmospheric nitrogen as nitrogen sources in the test plant, and therefore to assess the significance of N2 fixation. (Based on Peoples et al. 1989; reproduced with permission of ACIAR)

A typical ‘good’ rate of fixation for a (non-irrigated) field of subtropical legumes in northern Australia is c. 60–100 kg N ha–1 year–1. About the same amount of nitrogen is harvested as seed from a crop of cowpea, soybean or chickpea, so growing these legumes does not add net nitrogen to the soil; it does, however, spare nitrogen which would otherwise be removed at harvest. Irrigated legume-based pastures in temperate Australia or New Zealand fix 250–300 kg N ha–1 year–1 and make a substantial contribution to the low energy costs of agriculture in these regions. Selection of appropriate biological N2 fixers could greatly improve N2 fixation in tropical legume crops.

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Chapter 5 - Phloem transport


Early development of a pineapple. Phloem conduits from the leaves distribute sugars to the growing inflorescence, with flower buds arranged in spirals, which will later develop into the large juicy fruit.

Chapter editor: Yong-Ling Ruan1

Contributing Authors: Craig Atkins2; Yong-Ling Ruan1. 1University of Newcastle, Australia. 2School of Plant Biology, University of Western Australia

This Chapter is updated from a previous version written by John W Patrick, Ian Wardlaw and Tina Offler for Plants in Action 1st Edition.

A plant is a coordinated network of assimilatory regions (sources) linked to regions of resource utilisation (sinks). The phloem vascular system provides a path for assimilate transport from source to sink.

The phloem conduits distribute the sugars made in the leaves to growing tissues and organs that cannot carry out photosynthesis. These ‘sinks’ include shoot and root apices, flower buds, and developing fruit and seed.

Xylem conduits are responsible for delivery of water, inorganic nutrients and organic forms of nitrogen to transpiring leaves (Chapters 3 and 4).

Section 5.1 describes the pathway of the distribution of sugars made in chloroplasts, as well as nitrogen assimilates made in the leaves, to growing organs and other non-photosynthetic tissues. Section 5.2 describes the composition of phloem sap and how to collect it. Quantitative information is presented on the speed of phloem transport from sources to sinks, and the controls of long-distance transport.

Cellular and regulatory mechanisms of phloem loading in leaves are shown in Section 5.3, and mechanisms of phloem unloading at sinks in Section 5.4 with particular reference to developing seeds.

The focus of this chapter is on the transport of sugars. The transport of amino acids and other nitrogen-containing compounds is equally important, and the same general principles apply to nitrogen-containing or phosphorus-containing compounds that are synthesised in the leaf.

5.1 - Distribution of photoassimilates within plants

CO2 fixed by photosynthesis in chloroplasts has several possible fates, but most ends up as sucrose or starch. Starch is stored in chloroplasts, and sucrose is stored in vacuoles of mesophyll cells. Both starch and vacuolar sucrose serve as temporary storage pools from which the cytoplasmic sucrose pool is replenished. Sucrose, along with amino acids and mineral nutrients, is loaded into the phloem tissue which consists of sieve element—companion cell (se—cc) complexes for long-distance transport to growing tissues and other non-photosynthetic sinks. These solutes are exchanged reversibly between se-cc complexes and short- and long-term storage pools along the axial pathway. Short-term storage pools include phloem apoplasm, and the protoplasm of non-transport cells provides a long-term storage pool. At the end of the pathway, sucrose and other transported solutes are consumed in respiration and growth, or are stored as solutes in vacuoles or polymers in amyloplasts (starch) or protein bodies.

The overall flow of photoassimilates throughout the plant can therefore be called a source–path–sink system (Figure 5.1).


Figure 5.1. Schematic diagram of transfer and transport processes contributing to the flow of assimilates acquired from aerial or soil environments, through the source-path-sink system. CO2 fixed by photosynthesis in chloroplasts gives rise to sucrose and starch. Sucrose, amino acids and mineral nutrients are loaded into sieve element—companion cell (se—cc) complexes of leaf phloem for long-distance transport to non-photosynthetic sinks. These solutes are exchanged reversibly between se-cc complexes and short- and long-term storage pools along the axial pathway. Short-term storage pools include phloem apoplasm, whereas the protoplasm of non-transport cells provides a long-term storage pool. In sink tissues, solutes are used for respiration, growth or storage.

5.1.1 - Source–path–sink transport processes

(a) Source processes


Figure 5.2 Time course of sucrose and phosphorus (P) net import and export from a leaf during its development. As a cucumber leaf expands, net sucrose export coincides with the rise in net leaf photosynthetic rate (O) to meet photoassimilate demands of young leaves. Once a leaf has reached some 30% of its final area, net photosynthesis by the whole leaf exceeds photoassimilate demand by growth and so excess sucrose can be exported. Thereafter, the rate of sucrose export closely follows photosynthetic rate, reaching a maximum when the leaf reaches its final size and gradually declining thereafter. Import of P (and other mineral nutrients) continues throughout leaf expansion and P export only starts once the leaf is fully expanded. Sucrose import and export were calculated from the difference between rates of whole-leaf photosynthesis and dry matter gain (Based on Hopkinson 1964; reproduced with permission of Journal of Experimental Botany)


Net export of photoassimilates occurs from fully expanded leaves (Figure 5.2) and long-term storage pools located along the axial transport pathway. Chloroplasts of C3 plants (Chapters 1 and 2) partition photoassimilates between the photosynthetic oxidative cycle and starch biosynthesis or release them immediately to the cytosol as triose phosphate for sucrose synthesis. In non-starch-forming leaves, high concentrations of sugars can be accumulated in the vacuoles of mesophyll cells or made available for immediate loading into the phloem and export. Leaves also serve as secondary sources for nutrients and amino acids previously delivered in the transpiration stream. Nutrients and amino acids can be exported in the phloem immediately, or after accumulation in short-term storage pools.

An additional source of photoassimilates is located along the axial phloem path (petioles, stems, peduncles, pedicels and roots) as a result of leakage from the vascular tissues. Leaked photoassimilates accumulate in short- or long-term storage pools which serve as secondary sources to buffer photo-assimilate supplies to the sinks against shifts in export rates from the primary photoassimilate sources.

(b) Path processes

Assimilates including sucrose, amino acids are transferred into sieve elements of fully expanded leaves against significant concentration and electrochemical gradients. This process is referred to as phloem loading. The cellular pathways of phloem loading, and hence transport mechanisms and controls, vary between plant species. Longitudinal transport of assimilates through sieve elements is achieved by mass flow and is termed phloem translocation. Mass flow is driven by a pressure gradient generated osmotically at either end of the phloem pathway, with a high concentration of solutes at the source end and a lower concentration at the sink end. At the sink, assimilates exit the sieve elements and move into recipient sink cells where they are used for growth or storage. Movement from sieve elements to recipient sink cells is called phloem unloading. The cellular pathway of phloem unloading, and hence transport mechanisms and controls, vary depending upon sink function.

(c) Sink processes

Many sink organs are characterised by low rates of transpiration (an exception is a developing leaf) so that most assimilates are delivered by the phloem. Having reached the sink cell cytoplasm through the post-sieve-element transport pathway, assimilates are either metabolised to satisfy the energy, maintenance and growth requirements of sink cells or are compartmented into polymer or vacuolar storage. Collectively, metabolism and compartmentation create a demand for assimilates which is ultimately responsible for driving phloem import.

5.1.2 - Photoassimilate transport and biomass production

(a) Whole-plant growth

Sink and source strength must be in balance at a whole-plant level. Thus, an increase in whole-plant sink strength must be matched by an equal increase in source strength, either through increases in source activity or source size. Prior to canopy closure in a crop, much of the increase in source strength comes from increased source size, source activity remaining relatively constant. Significantly, until a leaf has reached some 30% of its final size, photoassimilates for leaf production are exclusively imported through the phloem from fully expanded leaves (Figure 5.2).

(b) Photoassimilate transport and crop yield

During domestication of crop plants, plant breeders selected for crop yield via maximum investment into harvested organs (mostly seeds). Total plant biomass production of advanced wheat is the same as its wild progenitors yet grain yield has increased some 30-fold through breeding. That is, whole-plant source and sink strength have not changed. Increases in wheat yield are associated with a diversion of photoassimilates from vegetative organs to the developing grain, as illustrated by the relative accumulation of 14C photoassimilates exported from the flag leaf.

Final grain yield is not only determined by partitioning of current photoassimilates, but also depends upon remobilisation of non-structural carbohydrates stored in stems, particularly under conditions where environmental stress impairs leaf photosynthesis (Wardlaw 1990). In fact, remobilisation of reserves affects yield in many food plants. For example, deciduous fruit trees depend entirely on remobilised photo-assimilates to support flowering and fruit set as do early stages of pasture regrowth following grazing.

5.1.3 - Whole-plant distribution of photoassimilate

Fig 5.14.jpg

Figure 5.3 Photoassimilate distribution in a rooted cutting of Washington Navel orange (mounted specimen shown on left; matching autoradiograph on right). 14CO2 was supplied to source leaves (boxed area top left) for a day, and movement of 14C-labelled assimilate followed by autoradiography of harvested plant material. 14C photosynthates were distributed widely via vascular conduits to sinks including some roots and a fruit on an adjacent shoot (note stem labelling between sources and sinks). Nearby mature leaves failed to import; they were additional sources of photosynthate. Scale bar = 2 cm (Unpublished material courtesy P.E. Kriedemann)

Photoassimilate transport to harvestable organs plays a central role in crop yield brought about by greater harvest indices. This raises questions about transport and transfer processes that collectively influence photoassimilate partitioning between competing sinks.

Historically, these questions were elucidated by observing partitioning patterns of photoassimilates exported from specified source leaves labelled with 14C supplied as a pulse of 14CO2. Following a chase period, in which 14C photoassimilates are transported to and accumulated by recipient sink organs, the plant is harvested. The pattern of photoassimilate partitioning operating during the pulse is deduced from 14C activity accumulated by sinks (Figure 5.3).

Photoassimilates are partitioned from source leaves to sinks in characteristic and reproducible patterns. For instance, in a vegetative plant, lower leaves are the principal suppliers of photoassimilate to roots, whereas upper leaves are the principal suppliers to the shoot apex. Leaves in an intermediate position export equal quantities of photoassimilates in either direction. However, the pattern of photoassimilate partitioning is not static, it changes with plant development. In vegetative plants, the direction of flow from a leaf changes as more leaves above it become net exporters. Furthermore, at the onset of reproductive development, growing fruits or seeds become dominant shoot sinks for photoassimilates at the expense of vegetative apices.

Photoassimilate partitioning patterns can be altered experimentally by removal of selected sources (e.g. leaves) or sinks (e.g. fruits). These manipulative experiments demonstrate that photoassimilate partitioning reflects the relative strengths of individual sources and sinks. Properties of the phloem pathway connecting sources with sinks are shown in the following Section 5.2.

5.2 - Phloem transport


Figure 5.4 The role of bark (phloem) in sugar movement in plants. Mason and Maskell (1928) demonstrated that removing a complete ring of bark (a) while leaving the wood (xylem) intact prevented downward movement of sugars. When a strip of bark was retained between upper and lower stem parts (b), sugars flowed downwards in direct proportion to the width of the remaining bark

Photoassimilate, mainly in the form of sucrose, is loaded into phloem of photosynthetically active leaves for long distance transport to nonphotosynthetic sink tissues.  Figure 5.4  shows that assimilate transport occurs in phloem but not xylem. Key characteristics of phloem transport along with its chemical composition and regulation are described below.

5.2.1 - Phloem structure and function

(a) Phloem structure

In most plant species, phloem is made up of phloem fibres, phloem parenchyma, sieve cells (sieve elements) and their accompanying companion cells (Figure 5.5a). Sieve elements are ideally suited for rapid transport of substances at high rates over long distances. They are elongated and are arranged end to end in files referred to as sieve tubes (Figure 5.16b). Abutting sieve elements are interconnected through membrane-lined pores (sieve pores) with large diameters (1 to 15µm). These pores collectively form sieve plates (Figure 5.16c). The transport capacity of sieve tubes is dependent on a developmentally programmed degeneration of the sieve element protoplasm (cell contents) leaving an open, membrane-bound tube. In mature conducting sieve elements, the protoplast is limited to a functional plasma membrane enclosing a sparse cytoplasm containing low densities of plastids, mitochondria and smooth endo-plasmic reticulum distributed along the lateral walls (Figure 5.16d). These relatively empty sieve tubes provide a longitudinal network which conducts phloem sap (Figure 5.5b).

Fig 5.16ann.jpg

Figure 5.5 (a) spatial arrangement of cell types in a vascular strand from the primary stem of Phaseolus vulgaris (French bean); electron micrographs of stem phloem of Curcurbita maxima (b,c) and P. vulgaris (d) illustrating significant structural characteristics of sieve elements and companion cells. (a) Conducting cells of the phloem (sieve elements) and accompanying companion cells form groups of cells that are separated by phloem parenchyma cells. This mosaic of cells is located between the cortex and xylem and capped by phloem fibres. Scale bar = 7.3 μm. (b) A longitudinal section through two sieve elements arranged end to end to form part of a sieve tube. Companion cells can also be seen. The abutting wall (sieve plates) displays characteristic membrane-lined sieve pores (arrowheads). Cytoplasm of the sieve elements has largely degenerated leaving only endoplasmic reticulum (arrows) and a few plastids around the mature sieve element. Scale bar = 5 μm. (c) A face view of part of a sieve plate showing sieve pores (arrowheads). Scale bar = 0.5 μm. (d) Transverse section through a sieve element and its accompanying companion cell illustrating the sparse cytoplasm and low density of organelles in the sieve element contrasting with the dense ribosome-rich cytoplasm of the nucleated companion cell. Note the mitochondria and rough endoplasmic reticulum. Scale bar = 1.0 μm. c, cortex; cc, companion cell; e, epidermis; er, endoplasmic reticulum; m, mitochondrion; n, nucleus; p, pith; pf, phloem libres; pp, phloem parenchyma; se, sieve element; sp, sieve plates; vc, vascular cambium; x, xylem

Sieve elements are closely associated with one or more companion cells, forming a sieve element–companion cell (se–cc) complex (Figure 5.5d) that plays an important role in transport. These distinct cell types result from division of a common procambial mother cell. In mature se–cc complexes, relatively open sieve elements contrast with adjacent companion cells containing dense, ribosome-rich cytoplasm with a prominent nucleus and abundant mitochondria and rough endoplasmic reticulum (Figure 5.5d). High densities of extensively branched plasmodesmata in contiguous walls of sieve elements and companion cells (Figure 5.6) account for intense intercellular coupling in se–cc complexes (van Bel 1993). Thus, companion cells are considered to perform the metabolic functions surrendered by, but required for, maintenance of viable sieve elements. This functional coupling has led to the concept of se–cc complexes being responsible for phloem transport.


Figure 5.6 (a) Electron micrograph and (b) diagrammatic interpretation of a secondary plasmodesma interconnecting a mature sieve element and its companion cell in a tobacco leaf. Note the characteristic branching of the plasmodesma within the wall of a companion cell. Scale bar = 0.2 μm. c, callose; other symbols as for Figure 5.16 (Based on Ding et al. 1993; reproduced with permission of Blackwell Science)

(b) Visualising the translocation stream

Fig 5.18-ann.jpg

Figure 5.7 Microautoradiographs of (a) transverse and (b) longitudinal sections of Phaseolus vulgaris stem tissue illustrating localisation of 14C-labelled photosynthate in sieve tubes. These sections are obtained by snap freezing plant tissue and removing frozen water by sublimation (e.g. freeze-drying or freeze substitution). 14C-labelled compounds do not move during preparation. Tissues are embedded in absolute dryness and thin sections are cut, mounted dry on microscope slides and overlain with a thin film of photographic emulsion. Silver grains are visible in the emulsion where 14C, an ideal radioisotope for these experiments, irradiates the film. Abbreviations: se, sieve element; pp, phloem parenchyma; vb, vascular bundle; other symbols as for Figure 5.6. Scale bar in (a) = 20μm; in (b) = 10 μm

Transport of radioactively labelled substances through phloem has been demonstated using microautoradiography (Figure 5.7), providing irrefutable evidence that sieve elements are conduits for transport of phloem sap. Experimentally, a pulse of 14CO2 is fixed photosynthetically and 14C-labelled sugars are given time to reach the stem, which is then excised and processed for microautoradiography. As 14C first moves through the stem, most of the isotope is confined to the transport pathway and very little has had time to move laterally into storage pools. High densities of 14C-labelled sugars are found in sieve elements (Figure 5.7), demonstrating that these cells constitute a transport pathway.

(c) Phloem sealing mechanisms

Herbivory or environmental factors causing physical damage could pose a threat to transport through sieve tubes and has undoubtedly imposed strong selection pressure for the evolution of an efficient and rapid sealing mechanism for damaged sieve tubes. Since sieve tube contents are under a high turgor pressure (P), severing would cause phloem contents to surge from the cut site, incurring excessive assimilate loss in the absence of a sealing mechanism. For dicotyledonous species, an abundant phloem-specific protein (P-protein) provides an almost instantaneous seal. P-protein is swept into sieve pores where it becomes entrapped, thus sealing off the damaged sieve tubes. Production of callose (β-1,3 glucan) in response to wounding or high-temperature stress is another strategy to seal off damaged sieve tubes. Callose also seals off sieve pores during overwintering in deciduous plants. Callose is deposited between the plasma membrane and cell wall, eventually blocking sieve pores. Whether deposited in response to damage or overwintering, callose can be degraded by β-1,3 glucanase, allowing sieve tubes to regain transport capacity.

5.2.2 - Techniques to collect phloem sap

Since phloem translocation is confined to sieve elements embedded within a tissue matrix, it is difficult to obtain uncontaminated samples of translocated sap. The least equivocal approach has been to take advantage of the high P of sieve tube contents. Puncturing or severing sieve tubes should cause exudation of phloem sap provided a sealing mechanism is not activated.


Figure 5.8 Aphids can be used to collect phloem sap. Top photograph: a feeding aphid with its stylet embedded in a sieve tube (see insert); scl, sclerenchyma; st, stylet; x, xylem; p, phloem. Note the drop of ‘honeydew’ being excreted from the aphid’s body. Plates (a) to (e) show a sequence of stylet cutting with an RF microcautery unit at about 3-5 s intervals (a to d) followed by a two-minute interval (d to e) which allowed exudate to accumulate. The stylet has just been cut in (b); droplets of hemolymph (aphid origin) are visible in (b) and (c); once the aphid moves to one side the first exudate appears (d), and within minutes a droplet (e) is available for microanalysis. Scale bars: top = 1 mm, bottom = 1.5 mm (Courtesy D. Fischer)


Figure 5.9. Exudation of phloem contents from lupins. A, following incision of the vasculature at the stylar tip and of the ventral suture of fruits of Lupinus angustifolius. B, following incision of the vasculature of a stem of L. angustifolius. C, following incisions to the vasculature of pre and post anthesis stage flowers on the inflorescence of L. angustifolius. D. exudation at the abscission zone following abscission of two flowers 5 minutes earlier on the raceme of L. mutabilis. Photographs courtesy Craig. Atkins.

For some plant species, sieve-pore sealing develops slowly, or can be experimentally down-regulated by massage or repeated excisions (Milburn and Kallarackal 1989) or slowed by puncturing the vasculature while it is snap frozen in liquid N2 (Pate et al 1984). Carefully placed incisions that do not disturb the underlying xylem, which in any case is more likely to be under tension, permit collection of relatively pure phloem exudate through the severed sieve tubes. Nevertheless, contamination with the contents of cells other than sieve tubes damaged at the site of incision is inevitable.  For the major solutes of phloem such as sugars or amino acids that are present in high concentrations this problem is minimal but for less abundant molecules like hormones or other signals, particularly proteins or nucleic acids, conclusions about the origin and functions of these must be made with caution. The ‘natural hemophiliacs’ of the plant world are few and include a number of cucurbits, some brassicas, castor bean, species of the genus Yucca and some species of lupin (Lupinus albus, L. angustifolius, L. mutabilis and L. cosentinii). The excision technique has been expanded to plant species that do not readily exude, by chemically inhibiting the sealing mechanism. Callose production is blocked when wounded surfaces are exposed to the chelating agent ethylenediaminetetraacetic acid (EDTA) by complexing with calcium, a cofactor for callose synthase. Immersing whole, excised organs in EDTA solution, which is essential to inhibit blockage, risks contaminating sap with solutes lost from the apoplast as well as non-conducting cells. This is not an ideal technique.

Enlisting sap-sucking aphids or leaf hoppers to sample sap has been more successful. Aphids can guide a long syringe-like mouthpart (a stylet) into conducting sieve elements (Figure 5.8). Pressure normally forces sieve-tube sap through the stylet into the aphid’s gut where it becomes food or is excreted as ‘honeydew’. By detaching the aphid from its mouthpart pure phloem contents can be collected from the cut end of the implanted stylet. Detaching the aphid body can be achieved by surgery following rapid anesthesia in high CO2 or by severing the stylet using a laser.  While stylectomy has been successful with a number of monocotyledons (rice, wheat and barley) the technique has proved more difficult to use with dicotyledons, yielding at best a few microlitres of phloem contents. On the other hand collection of milliliter volumes of exudate from one of the natural hemophiliacs is possible permitting extensive analysis of solutes and macromolecules. In the case of lupins, exudation occurs readily at many sites on the plant so that solutes translocated from source tissues as well as entering sinks can be collected and analysed (Figure 5.9).

5.2.3 - Chemical nature of translocated material

(a) Chemical analysis of phloem exudate

Chemical analyses of phloem exudate collected from a wide range of plant species have led to a number of generalisations (e.g. Milburn and Baker 1989) about the contents of sieve tubes. Phloem exudate is a concentrated solution (10–12% dry matter), generating an osmotic pressure (Π) of 1.2 to 1.8MPa. pH is characteristically alkaline (pH 8.0 to 8.5). The principal organic solutes are non-reducing sugars (sucrose), amides (glutamine and asparagine), amino acids (glutamate and aspartate) and organic acids (malate). Of these solutes, non-reducing sugars generally occur in the highest concentrations (300–900 mM). Nitrogen is transported through the phloem as amides and amino acids; nitrate is absent and ammonium only occurs in trace amounts. Calcium, sulphur and iron are scarce in phloem exudate while other inorganic nutrients are present, particularly potassium which is commonly in the range of 60–120 mM. Physiological concentrations of auxins, gibberellins, cytokinins and abscisic acid have been detected in phloem exudate along with nucleotide phosphates. The principal macromolecule group is protein but low levels of peptides and nucleic acids are also present.  While in cucurbits the protein in exudate is comprised largely of P-protein, a diverse array of proteins, many of them enzymes, have also been detected.

(b) Significance of the chemical forms translocated 

Phloem sap provides most inorganic and all organic substrates necessary to support plant growth. Non-transpiring tissues are particularly dependent on resources delivered in the phloem (Section 5.1). That translocated sugars represent the major chemical fraction of the phloem sap is consistent with the bulk of plant dry matter (90%) being composed of carbon, hydrogen and oxygen. Carbon transport is further augmented by transport of nitrogen in organic forms.

Carbohydrate is translocated as non-reducing sugars in which the metabolically reactive aldehyde or ketone group is reduced to an alcohol (mannitol, sorbitol) or combined with a similar group from another sugar to form an oligosaccharide. Apart from sucrose, transported oligosaccharides belong to the raffinose series. In this series, sucrose is bound with increasing numbers of galactose residues to form raffinose, stachyose and verbascose respectively. However, sucrose is the most common sugar species transported. In a small number of plant families, other sugar species predominate. For example, the sugar alcohol sorbitol is the principal transport sugar in the Rosaceae (e.g. apple) and stachyose predominates in the Cucurbitaceae (e.g. pumpkin and squash). Exclusive transport of non-reducing sugars probably reflects packaging of carbohydrate in a chemical form which protects it from being metabolised. Metabolism of these transported sugars requires their conversion to an aldehyde or ketone by enzymes which are thought to be absent from sieve-tube contents.

Plant physiologists have long regarded the two long distance translocation streams of xylem and phloem as having functions additional to the distribution of nutrients and assimilates. Specifically, each serves as a means of communication between the source and sink organs such that systemic signals are thought to transmit molecular responses to endogenous and environmental cues. Furthermore, evidence is accumulating that some of these signals regulate gene expression as a consequence of their translocation (see below). 

(c) Macromolecule composition of phloem

Proteomic and transcriptomic analyses have demonstrated a widely diverse composition of proteins, peptides and nucleic acids, including mRNA and small RNAs, in phloem exudates. While the origin of each individual protein or nucleic acid remains to be verified the limited compositional data available from stylectomy confirms that indeed each group of macromolecules is present in phloem. In cucurbit phloem exudate some 1110 different proteins have been detected along with a large number of mRNAs and similar data have been obtained for exudates from other species (Brassica napus, Ricinus communis and Lupinus albus). Compositional data for phloem proteins of these species show a common complement that includes phloem-specific P proteins together with proteins involved in sugar metabolism and transport, protein turnover and transport, detoxification of reactive oxygen species, as well as proteins that provide defence against insect herbivores and pathogens (Figure 10).  Some undoubtedly play a role in maintenance of the SE system while others, such as the Flowering Locus T (FT) protein associated with the flowering response (‘florigen’), appear to be systemic ‘signals’ (Rodriguez-Medina et al 2011) and there may be many more. Because sieve tubes are enucleate and lack ribosomes (5.2.2 a), proteins in the translocation stream are not formed in situ but are transported from sites of synthesis in the companion cells.


Figure 5.10. Two dimensional polyacrylamide gel electrophoresis separation of proteins in phloem exudate from Lupinus albus.  The gel was developed in the first dimension by isoelectric focusing with a linear pH gradient of 3-10 followed by separation due to differences in molecular mass.  The positions of mass standards are shown on the right hand side of the gel.  After staining with Coomassie Blue to locate the spots they were excised for digestion with trypsin. The peptides were then analysed by partial sequence determination using MS/MS and identified using database searches. (Courtesy of Craig Atkins)

Functional analysis of the cDNA identified in transcriptome studies of phloem exudates revealed transcripts involved in a wide range of processes that include metabolism, plant responses to stresses, transport, DNA/RNA binding and protein turnover.  The presence of transcripts in phloem exudate supports the idea of an RNA-based signalling network that is thought to function in control of processes associated with plant growth and development (Lough and Lucas, 2006).  However, the functional role of transcripts in the contents of sieve tubes as well as their actual translocation is yet to be determined.

Small RNA molecules (18-25 nt) have been identified in phloem exudate collected from rape, white lupin, pumpkin, castor bean and Yucca filamentosa as well as in aphid stylet exudate collected from apple stems. The population includes both microRNAs (miRNA) and small interfering RNAs (siRNA) a large number of which target mRNA of transcription factors that themselves regulate genes expressions. miRNAs are also involved in mediating environmental responses, including responses to salinity, drought, nutrient limitations, as well as hormone interactions. Their small size and powerful functions in targeting mRNAs to regulate expression suggest that those in phloem exudate are likely to be systemic signals.

An important question that relates to the significance of macromolecules in the contents of sieve tubes is proof that they are translocated and that translocation is essential for their function at a sink. A diversity of studies that have exploited cucurbit root stocks and grafted scions has provided clear evidence that P proteins among others are graft transmissible. In a series of elegant experiments Aoki et al. (2005) labelled and injected two isolated pumpkin phloem proteins (CmPP16‑1 and CmPP16-2) into the vasculature of intact rice plants through severed leaf hopper stylets and showed their translocation as well as some evidence for specificity in protein translocation.  The Flowering Locus T (FT) protein formed in leaves mediates the flowering transition of shoot apical meristems and the evidence that it is translocated is compelling. The long distance movement of RNA molecules was first demonstrated for plant viruses and there is now good evidence for phloem translocation of a number of transcripts (Lough and Lucas 2006). A recent compilation identified 13 miRNAs involved in plant responses to drought/salt stress (Covarrubias and Reyes 2010).  Eight of these were identified in lupin phloem exudate (Rodriguez-Medina et al. 2011) and, importantly, six were also recovered from PCR amplification of apple stylet exudate (Varkonyi-Gasic et al 2010).  There is thus a possibility that the responses to drought and salinity are mediated through miRNAs translocated from sites where the stress is sensed to sites where a response is initiated. 

The most convincing case for a translocated miRNA in phloem regulating gene expression relates to Pi homeostasis. While both local and systemic signals are involved, miR399 is phloem mobile and acts directly in roots to down regulate the expression of PHO2 (a ubiquitin conjugating enzyme) that results in greater expression of Pi transporters to increase Pi uptake under conditions of deficiency. Systemic signaling has also been implicated in homeostasis of other nutrients, including N, S and Cu with, in each case, miRNAs involved. 

5.2.4 - Phloem flux

Phloem flux can be estimated in a number of ways. The simplest is to determine dry weight gain of a discrete organ connected to the remainder of a plant by a clearly definable axis of known phloem cross-sectional area. Developing fruits or tubers meet these criteria. Sequential harvests from a population of growing fruit or tubers provide measures of the organ’s net gain of dry matter imported through the phloem. Net gains or losses of dry matter resulting from respiration or photosynthesis are incorporated into calculations to give gross gain in dry matter by the organ. Flux of dry matter through the phloem (specific mass transfer — SMT; Canny 1973) can then be computed on a phloem or preferably on a sieve-tube lumen cross-sectional area basis. Area estimates can be obtained from histological sections of the pedicel or stolon that connects a test organ to its parent plant. Expressed on a phloem cross-sectional area basis, SMT estimates are normally in the range of 2.8–11.1 g m–2 phloem s–1 (Canny 1973). Flux on the basis of sieve-tube lumen cross-sectional area is preferable but relies on identification of sieve tubes and the assumption that they are equally functional as transport conduits. Sieve tubes account for some 20% of phloem cross-sectional area, suggesting fluxes are about five-fold higher through a sieve-tube lumen.

Speed of phloem translocation can be determined from simultaneous measurements of SMT and phloem sap concentrations as shown in Equation 5.1 below:

\[\mathrm{Speed} (m \cdot s^{-1}) = \mathrm{SMT}(g \cdot m^{-2} \cdot s^{-1})/ \mathrm{concentration} (g \cdot m^{-3}) \tag{5.1} \]

For a sucrose concentration of 600 mM (or 2.16 x 105 g m-3) and the highest SMT values shown above, Equation 5.1 estimates that phloem sap can move at speeds of up to 56 × 10–5 m s–1 or 200 cm h–1. These estimates have been verified by following the movement of radioisotopes introduced into the phloem translocation stream.

These estimates of transport rates and speeds tacitly assume that phloem sap moves through sieve tubes by mass flow (water and dissolved substances travel at the same speed). Independent estimates of transport rate, concentration of phloem sap and translocation speed lend support to, but do not verify, the assumption that movement occurs as a mass flow.

A simple and direct test for mass flow is to determine experimentally whether water and dissolved substances move at the same speed. This test should be relatively easy to apply using radioactively labelled molecules. Unfortunately, in practice it turns out that different molecular species are not loaded into the sieve tubes at the same rates and the plasma membranes lining the sieve tubes are not equally permeable to each substance. Thus, the analysis is complicated by the necessity to use model-based corrections for rates of loading into and losses from the sieve tubes. Nevertheless, the speed estimates obtained from such experiments are found to be similar for dissimilar molecules, supporting the proposition that mass flow accounts for most transport through sieve tubes.

Phloem translocation is generally believed to be driven by pressure. Münch (1930) proposed that a passive mass flow of phloem sap through sieve tubes was driven by the osmotically generated pressure gradient between source and sink regions (Figure 5.11). At source regions, the principal osmotica of phloem sap are actively or passively loaded into sieve tubes from companion cells or mesophyll cells (see 5.3.2), thereby driving water towards the lower water potentials within sieve tubes. As water enters, P rises. Unloading of solutes from sieve tubes at sink regions reverses water potentials; water flows out of sieve tubes and P falls relative to that of sieve tubes in source regions.

The pressure-flow hypothesis can be modelled using the relationship that rate of mass flow (Ff) of a substance is given by the product of speed (S) of solution flow, path cross-sectional area (A) and its concentration (C). That is:

 \[F_f = S \cdot A \cdot C \tag{5.2}\]

 Speed (m s–1) has the same units as volume flux (Jv — m3 m–2 s–1) of solution passing through a transport conduit. Poisseuille’s Law describes the volume flux (Jv) of a solution of a known viscosity (h) driven by a pressure difference (DP) applied over the length (l) of pathway of radius (r) as:

\[J_v = \pi r^4 \Delta P/8 \eta l \tag{5.3} \]


Figure 5.11 Scheme describing the pressure flow hypothesis of phloem transport (Based on Münch 1930)

The term πr4/8ηl in Equation 5.3 provides an estimate of hydraulic conductivity (Lp) of the sieve-tube conduit which is set by the radius of the sieve pores. Raised to the fourth power, small changes in the sieve-pore radius will exert profound effects on the hydraulic conductivity of the sieve tubes (Section 5.2). The viscosity of sieve-tube sap is determined by the chemical species (particularly sugars) and their concentrations in the phloem sap.

Key features of the pressure-flow hypothesis are encapsulated in Equation 5.3. The central question is whether a pressure gradient exists in sieve tubes with the expected direction and of sufficient magnitude to support observed rates of sap flow. Indirect estimates of P in sieve tubes made through determination of intra- and extracellular P support the pressure-flow hypothesis. Direct measurements of sieve-tube P are technically challenging because of the inaccessibility of these small, highly turgid cells. They are, for instance, too small for pressure-probe measurements. However, manometric pressure measurements obtained using severed aphid stylets agree with indirect estimates (Wright and Fisher 1980). Experimental manipulation of the pressure gradient between the source and sink also results in alterations in phloem translocation rates consistent with the pressure-flow model.

Whether the pressure gradient is sufficiently steep is a more vexing question. The pressure gradient required to drive phloem translocation at observed rates is determined by the transport resistance of the phloem path, according to Ohm’s Law. Dimensions of the sieve pores set a limiting radius for volume flux of transported sap (Equation 5.3) and hence transport resistance. If the sieve pores were open and unoccluded by P-protein, a number of studies have demon-strated that the measured pressure gradients are sufficient to support the observed rates of flow. However, the in situ radii of sieve pores remain unknown.

Overall, the pressure-flow hypothesis accounts for many observed features of phloem translocation, including distribution of resources. While conclusive evidence supporting this hypothesis is still sought, less attention is now focused on this issue with a growing appreciation that the phloem pathway has spare transport capacity. Evidence from Kallarackal and Milburn (1984), for example, showed that the specific mass transfer (SMT – see preceding section) to an intact fruit of castor bean could be doubled on removal of competing fruits. Moreover, if P of sieve elements at the sink end of the phloem path was reduced to zero, by severing the pedicel and allowing exudation, SMT rose to an incredible 305 g m–2 sieve-tube area s–1! In another experiment, when half the conducting tissue was removed from the peduncle of sorghum or wheat plants, grain growth rate was not impaired (Wardlaw 1990). Together, these observations imply that phloem has excess carrying capacity in both dicotyledons and monocotyledons. Particularly in monocotyledonous plants, a strong selection pressure for spare transport capacity must exist because there is no vascular cambial activity to replace damaged sieve elements.

5.2.5 - Control of assimilate transport from source to sink

Loading of sugars, potassium and accompanying anions into sieve tubes at sources determines solute concentrations in phloem sap (Table 5.1). The osmotic pressure (Π) of these solutes influences P generated in sieve tubes. Thus, source output determines the total amount of assimilate available for phloem transport as well as the pressure head driving transport along the phloem path to recipient sinks. Withdrawal of assimilates from sieve tubes at the sink end of the phloem path, by the combined activities of phloem unloading and metabolism/compartmentation (Table 5.1), determines Π of phloem sap. Other sink-located membrane transport processes influence Π around sieve tubes. The difference between intra- and extracellular Π of sieve tubes is a characteristic property of each sink and determines P in sink sieve tubes.

The pressure difference between source and sink ends of the phloem pathway drives sap flow (Equation 5.3) and hence phloem translocation rate (Equation 5.2) from source to sink. The source and sink processes governing the pressure dif-ference (Table 5.1) are metabolically dependent, thus rendering phloem translocation rates susceptible to cellular and environmental influences. The pressure-flow hypothesis predicts that the phloem path contribution to longitudinal transport is determined by the structural properties of sieve tubes (Table 5.1). Variables of particular importance are cross-sectional area (A) of the path (determined by numbers of sieve pores in a sieve plate and sieve-tube numbers) and radius of these pores (sets r in Equation 5.3). These quantities appear in Equations 5.2 and 5.3. Thus, the individual properties of each sink and those of the phloem path connecting that sink to its source will determine the potential rate of assimilate import to the sink (Figure 5.12).


Figure 5.12. Scheme describing photoassimilate flow from a source leaf linked to two competing sinks, Sink 1 and Sink 2. Assimilate flows through alternative phloem paths (Path 1 and Path 2) each with its own conductance (Kpath) and pressure difference (P) between source and sink. Hence Path 1 is distinguished by Kpath1 and Psink1 and Path 2 by Kpath2and Psink2

The transport rate (R) of assimilate along each phloem path, linking a source with each respective sink, can be predicted from the pressure-flow hypothesis (see Equations 5.2 and 5.3) as:

\[ R = K_{path} (P_{source} - P_{sink}) C \tag{5.4}\]

where path conductance (Kpath) is the product of path hydraulic conductivity (Lp) and cross-sectional area (A). Hence, the relative flows of assimilates between hypothetical sinks (sink 1 and sink 2) shown in Figure 5.12 may be expressed by the following ratio:

\[ \frac{K_{path1} (P_{source} - P_{sink1}) C} {K_{path2} (P_{source} - P_{sink2}) C} \tag {5.5} \]

Partitioning of assimilates between two competing sinks is thus a function of path conductance and P at the sink end of the phloem path (Equation 5.5). Since phloem has spare capacity, any differences in the conductance of the inter-connecting paths (Figure 5.12) would exert little influence on the rate of phloem transport to the competing sinks. Assimilate partitioning between competing sinks would then be determined by the relative capacity of each sink to depress sieve-tube P at the sink end of the respective phloem path. Even when differences in path conductance are experi-mentally imposed, phloem transport rates are sustained by adjustments to the pressure differences between the source and sink ends of the phloem path (Wardlaw 1990).

These conclusions have led to a shift in focus from phloem transport to phloem loading and unloading, which are instrumental in determining the amount of assimilate translocated and its partitioning between competing sinks, respectively.

5.3 - Phloem loading

Photoassimilates are loaded along the entire phloem transport pathway, from photosynthetic leaves to importing sinks. While most photoassimilate loading occurs in photosynthetically active leaves, root-produced metabolites, such as amino acids, move readily from xylem to phloem particularly at the stem nodes. Phloem loading also occurs in storage organs during periods when reserves are remobilised and exported. Indeed, the membrane transport events contributing to phloem loading were first examined using export of sucrose remobilised from the endosperm of germinating castor bean seed as an experimental model (Kriedemann and Beevers 1967).

This section focuses on phloem loading in the leaves. It analyses the cellular pathways for assimilate loading, and the regulatory controls.

5.3.1 - Pathway of phloem loading in source leaves

(a) Delineating the transport path 

Phloem loading is used variously to describe transport events outside, and inside, phloem tissues of leaves. The broader general application is adopted here — that is, phloem loading describes photoassimilate transport from the cytoplasm of photosynthetic mesophyll cells to se–cc complexes of leaf phloem.

Phloem loading commences in mesophyll cells and ends in the leaf vascular system. The se–cc complexes occur in a wide array of vascular bundle sizes. In dicotyledonous leaves, veins undergo repeated branching, forming the extensive minor vein network described in Section 5.2. For example, sugar beet leaves contain 70cm of minor veins cm–2 of leaf blade, while the major veins contribute only 5.5cm cm–2 of leaf blade (Geiger 1975). These observations and physiological studies (van Bel 1993) show that the principal site of phloem loading is in the minor vein network of dicotyledonous leaves. In contrast, the major veins transport loaded photoassimilates out of leaves.

Fig 5.22-ann.jpg

Figure 5.13. Transmission electron micrograph through a minor vein of a source leaf of maize (Zea mays L.) This vascular bundle consists of two sieve elements (st), one xylem vessel (v) and five vascular parenchuma cells (vp). These sieve elements are of two types, one thin walled and accompanied by a companion cell, the other thick walled and adjacent to the xylem vessel. Other symbols are: bs, bundle sheath; cc, companion cell; is, intracellular space; st, sieve tube. Scale bar - 4.2 µm (Based on Evert et al. 1978; reproduced with permission of Planta)

Minor veins usually comprise a single xylem element, vascular parenchyma cells and one to two sieve elements surrounded by one to four companion cells (Figure 5.13). The se–cc complex in minor veins bears similarities to that of stems (Figure 5.6). Companion cells have dense cytoplasm containing many mitochondria and are often considerably larger than the sieve elements they accompany. Companion cells are symplasmically connected to the sieve elements by branched plasmodesmata.

Cross-sectional areas of veins in monocotyledonous leaves reveal large and small parallel veins. Photoassimilates are loaded into the small veins and conducted through large veins. Fine transverse veins carry photoassimilates loaded into small veins across to large veins for export.

(b) Cellular pathways — symplasmic versus apoplasmic


Figure 5.14. Scheme describing symplasmic and apoplasmic pathways of phloem loading. Lines without arrows joining boxes represent symplasmic continuity (i.e. plasmodesmata). Black arrows indicate symplasmic transport (i.e. through plasmdesmata); green arrows indicate apoplasmic transport requiring solutes to cross membranes. vpc, vascular parenchyma cell; se, sieve element; cc, companion cell (Based on van Bel 1993)

Photoassimilates could move intercellularly through interconnecting plasmodesmata from chloroplasts in mesophyll cells to the lumena of sieve elements (symplasmic phloem loading) or across plasma membranes, travelling part of the route through the cell wall continuum (apoplasmic phloem loading). These fundamentally different pathways are shown schematically in Figure 5.14. Debate persists over which cellular pathway of phloem loading prevails because experiments on transport from mesophyll cells to sieve elements are difficult.

Extraordinarily, the cellular pathway of phloem loading reflects evolutionary relationships. Species from ancient plant groups display symplasmic loading, while species of more modern plant groups appears to exhibit apoplasmic phloem loading (van Bel 1993). Evidence for respective routes of loading follows.

A symplasmic pathway depends upon development of extensive plasmodesmal interconnections between adjoining cells, forming a cytoplasmic continuum from mesophyll to se–cc complexes (Figure 5.14). Such symplastic continuity is found in leaves of plant families containing trees and shrubs as well as cucurbits such as squash (van Bel 1993). An abundance of plasmodesmal interconnections demonstrates potential for symplasmic transport but does not establish whether such transport actually occurs. Membrane-impermeant fluorescent dyes microinjected into mesophyll cells are transported to se–cc complexes, demonstrating that plasmodesmata can provide a route for photoassimilate transport. Furthermore, when leaves were fed14CO2 and treated with inhibitors that block sugar transport across plasma membranes, transport of 14C-labelled photo-assimilates continued unaffected along the enforced symplasmic unloading route (Figure 5.15; van Bel 1993). In this case, sugar levels are higher in the mesophyll than in the phloem and ions and molecules diffuse through plasmodesmata at each interface, without a concentrating step (Turgeon 2010). Therefore, this is a passive symplasmic phloem loading.

Symplasmic phloem loading may also be an active process occurring in some herbaceous eudicots. This model of phloem loading, called polymer trap mechanism, depends on sucrose being biochemically converted to raffinose oligosaccharides (RFOs) in specialized CCs (intermediary cells - ICs) (Turgeon 2010). The biochemical synthesis of RFOs from sucrose requires metabolic energy. The synthesized RFOs exceed size exclusion limits of plasmodesmata linking mesophyll cells with ICs and therefore are trapped and accumulate to high concentrations in SE/IC complexes of minor veins for long distance transport (Turgeon 2010).

Plant species that load phloem from the leaf apoplasm are characterised by a low abundance of plasmodesmata between se–cc complexes and abutting vascular cells. However, as for symplasmic loaders, mesophyll cells of these species are interconnected by abundant plasmodesmata (Figure 5.23). Herbaceous and many crop species belong to this group of phloem loaders, including grasses (van Bel 1993). Conventional physiological observations are consistent with phloem loading in leaves of these species including a membrane transport event located somewhere between mesophyll cells and the se–cc complexes of minor veins (Figure 5.15).


Figure 5.15. Testing whether photoassimilates move from mesophyll cells to se-cc complexes through (a) an entirely symplasmic route or (b) a route with an apoplasmic step. The approach is to use PCMBS as an inhibitor of membrane transport. PCMBS does not cross membranes but binds to the apoplasmic face of plasma membranes. Therefore, it blocks apoplasmic transport while symplasmic phloem loading is unaffected. PCMBS was introduced into the leaf apoplasm through the transpiration stream of excised leaves. Leaves were then exposed in a closed illuminated chamber to 14CO2. The 14C photoassimilate exported from labelled leaf blades was used to monitor phloem loading. PCMBS only reduced photoassimilate export (i.e. phloem loading) from those leaves with few plasmodesmata interconnecting se-cc complexes with surrounding cells. Thus, photoassimilate flow included a membrane transport step from the leaf apoplasm in certain plant species while others loaded via a symplasmic route. cc, companion cell; mc, mesophyll cell; msc, mesophyll sheath cell; PCMBS (para-chloromercuriben-zenesulphonic acid, also abbreviated to P); se, sieve element; vp, vascular parenchyma (Based on van Bel 1993)

Molecular biology has brought new insights to phloem loading. For instance, existence of an apoplasmic step demonstrated with PCMBS (Figure 5.15) has been elegantly confirmed using molecular biology to control activity of the sucrose/proton symporter responsible for sucrose uptake from phloem apoplasm into se–cc complexes. Specifically, potato plants were transformed with an antisense copy of the gene encoding the sucrose/proton symporter, producing a phenotype with low levels of the symporter in plasma membranes of se–cc complexes (Frommer et al. 1996). Excised leaves of transformed plants exported significantly less photoassimilates than wild-type plants, corroborating the inhibitory effect of PCMBS on apoplasmic phloem loading (Figure 5.15). This provides compelling evidence that passage of photoassimilates from mesophyll cells to se–cc complexes in potato leaves includes an apoplasmic step.

Vascular parenchyma cells are the most probable site for photoassimilate exchange to phloem apoplasm (van Bel 1993), ensuring direct delivery for loading into se–cc complexes. Furthermore, plasma membranes of se–cc complexes in minor veins have increased surface areas to support photoassimilate transfer from phloem apoplasm. Notably, the surface area of se–cc complexes in sugar beet leaves is surprisingly large—0.88 cm–2of leaf blade surface. By implication, these large membrane surfaces are involved in phloem loading. Further support comes from cytochemical studies, demonstrating a great abundance of proteins associated with energy-coupled sucrose transport (Section 5.3.3(b)).

Leaf anatomies in some plant species suggest a potential for simultaneous phloem loading through apoplasmic and symplasmic pathways (van Bel 1993). Whether these pathways connect the same sieve element, different sieve elements in the same minor vein order or sieve elements in different vein orders is still unknown.

5.3.2 - Mechanisms of phloem loading

(a) General characteristics

Any hypothesis of phloem loading must account for the following characteristics:

  1. Elevated solute concentration in se–cc complexes. Estimated solute concentrations in sap of se–cc complexes is much higher than concentrations in sap of surrounding cell types, irrespective of whether phloem loading is by an apoplasmic or a symplasmic route.
  2. Selective loading of solutes into se–cc complexes. Chemical analysis of phloem sap by techniques shown above in Section 5.2 reveals relative solute concentrations different from those in surrounding cells. Phloem loading is therefore a selective process.

(b) Symplasmic loading 

The above-described characteristics have been used to argue against loading of se–cc complexes through a symplasmic route on the grounds that plasmodesmata lack mechanisms for concentrating and selecting solutes. However, a contribution of plasmodesmata to concentrating and selecting solutes cannot be precluded from our current knowledge of plasmodesmal structure and function.

Plants that load se–cc complexes through a symplasmic route translocate 20–80% of sugars in the form of raffinose-related compounds such as raffinose, stachyose and verbascose (Section 5.2.3(c)). Grusak et al. (1996) proposed a model for symplasmic phloem loading that accounts for the general characteristics stated above. According to this model (Figure 5.16), sucrose diffuses from mesophyll and bundle sheath cells into intermediary (companion) cells through plasmodesmata. Within companion cells, sucrose is thought to be enzymatically converted to oligosaccharides (raffinose or stachyose) maintaining a diffusion gradient for sucrose from mesophyll cells into se–cc complexes. The molecular-size-exclusion limit of plasmodesmata interconnecting mesophyll and companion cells is such that it prevents back diffusion of stachyose and raffinose molecules, which are larger than sucrose. These oligosaccharides are able to diffuse through plasmodesmata with larger diameters linking companion cells with sieve elements (van Bel 1993). This model accounts for selective loading of sugars to achieve high photoassimilate concentrations in phloem elements.


Figure 5.16. Model of the ‘polymerisation trap mechanism’ to explain symplasinic phloem loading against a solute concentration gradient. Sucrose moves through a symplasmic path from photosynthetic cells into intermediary (companion) cells of the minor veins. Sucrose movement is by diffusion down a concentration gradient maintained by the polymerisation of sucrose into oligosaccharides (raffinose and stachyose) in intermediary cells. Diffusion of these oligosaccharides into mesophyll cells is prevented, as their size exceeds the molecular exclusion limit of plasmodesmata joining mesophyll and intermediary cells. However, the larger-diametered plasmodesmata linking intermediary cells with sieve elements permit oligosaccharides to be loaded into sieve elements for export from the leaf. [], glucose; Δ, fructose; • galactinol (Based on Grusak et al. 1996)

(c) Apoplasmic loading 

Phloem loading with an apoplasmic step is an attractive model, explaining both how solutes become concentrated in se–cc complexes (energy-coupled membrane transport) and how they could be selected by specific membrane transporters (see van Bel 1993). Identifying transport mechanisms responsible for photoassimilate transport to and from the leaf apoplasm has proved challenging.

Based on estimates of sucrose fluxes and high sucrose concentrations in phloem sap, there is little doubt that sucrose loading into phloem is energy dependent. The demonstration that PCMBS blocks loading of photoassimilates in whole leaves of certain species (Section 5.3.2(b)) points to carrier-mediated transport across plasma membranes. Genes encoding sucrose porters have been cloned from leaf tissue (Frommer et al. 1996) and shown to be specifically expressed in leaf phloem. Complementation studies in yeast defective in sucrose transport suggest that the phloem-located sucrose porter catalyses sucrose/proton symport in a similar way to that illustrated in Figure 5.32. Antisense transformants of potato with low abundance of this symporter have impaired sucrose transport (Section 5.32(b)).

In contrast to photoassimilate uptake from phloem apoplasm, very little is known about the mechanism of sugar efflux into the apoplasm until very recently. Estimates of photoassimilate flux to phloem apoplasm, based on rates of sucrose export from leaves, suggest that this transport event must be facilitated by other transport processes (van Bel 1993). This is now confirmed by the recent cloning of sucrose efflux protein that sheds a light on the molecular mechanisms of phloem loading (Chen et al. 2012).  

5.3.3 - Sink regulation of phloem loading


Figure 5.17. Time-course of photoassimilate export from source leaves of tomato plants. Control plants, in which fruits were a major sink for photoassimilates, were maintained at 20°C. Treatments involved (1) removing fruit or (2) exposing plants with fruits to 30°C. The proportion of 14C label remaining in source leaves after a radioactive pulse was monitored through time to show that (1) presence of major sinks or (2) more rapid metabolism accelerated 14C export from source leaves (Based on Moorby and Jarman 1975).

(a) Sink effects on export

The response of photoassimilate export to changes in sink demand depends upon whether photoassimilate flow is source or sink limited (Wardlaw 1990). A source-limited system does not respond rapidly to an increase in sink demand, depending more on the capacity of leaves to increase the size of the transport pool. In contrast, alterations in sink demand in a sink-limited system elicit immediate effects on photoassimilate export. Figure 5.17 shows how the presence of fruits accelerates 14C export, especially at high temperatures. For leaves that load the se–cc complexes from apoplasmic pools, changes in sink demand probably influence photoassimilate export by altering membrane transport properties. These changes in membrane transport entrain a flow of adjustments in biochemical partitioning within the leaf through substrate feedback (see below).

(b) Sink effects on membrane transport

Changes in the turgor pressure of phloem sap or altered phytohormone levels could serve as signals for sink demand.

Changes in the pressure of sink phloem sap are rapidly transmitted through sieve tubes to sources. Phloem loading in source tissues responds to this pressure signal by changes in solute transport rates mediated by membrane-associated porters (van Bel 1993). This is a proposed mechanism for phloem loading which would respond rapidly (within minutes) to changes in sink demand.

Phytohormone levels in leaves respond to changes in the source/sink ratio. For instance, gibberellin levels in leaves proximal to developing inflorescences increase at fruit set. In contrast, abscisic acid levels in soybean and grape leaves are inversely related to alterations in sink demand (Brenner 1987). Therefore, changes in leaf phytohormone levels could serve to signal shifts in sink demand for photoassimilates. In this context, direct application of auxin and gibberellic acid to source leaves results in a rapid enhancement of photoassimilate export (Table 5.2). Gibberellic acid did not stimulate leaf photosynthesis or alter photoassimilate partitioning, appearing instead to upregulate phloem loading. This was confirmed by faster 14C loading into isolated phloem strands (Table 5.2).

(c) Sink influences on biochemical partitioning within source leaves  

A substrate feedback response is elicited if the rate of photo-assimilate export from chloroplasts is limited by sink demand. If sucrose export from source pools is accelerated by phloem loading, substrate feedback inhibition of photoassimilate delivery is alleviated. A cascade of adjustments in the activity of key regulatory enzymes follows (see Section 2.3) with the final outcome of an increased flow of sucrose into transport pools. Conversely, if photoassimilate flow is limited by photosynthetic rate, the activity of enzymes responsible for sucrose biosynthesis is not subject to feedback inhibition by substrates. As a consequence, responses to increased sink demand can only be mediated by increases in photosynthetic enzyme activity.

5.4 - Phloem unloading and sink utilisation

Photoassimilate removal from phloem and delivery to recipient sink cells (phloem unloading) is the final step in photoassimilate transport from source to sink. Within sink cells, cellular metabolism and compartmentation are the end-users of phloem-imported photoassimilates. Combined activities of these sink-located transport and transfer events determine the pattern of photoassimilate partitioning between competing sinks and hence contribute to crop yield.

Phloem unloading describes transport events responsible for assimilate movement from se–cc complexes to recipient sink cells. A distinction must be made between transport across the se–cc complex boundary and subsequent movement to recipient sink cells. The former transport event is termed sieve element unloadingand the latter post-sieve element transport. On reaching the cytoplasm of recipient sink cells, imported photoassimilates can enter metabolic pathways or be compartmented into organelles (e.g. amyloplasts, protein bodies and vacuoles). Metabolic fates for photoassimilates include catabolism in respiratory pathways, biosynthesis (maintenance and growth) and storage as macromolecules (starch and fructans).

Compared with phloem loading, phloem unloading and subsequent sink utilisation of imported photoassimilates operate within a much broader range of configurations:

  1. morphological (e.g. apices, stems, roots, vegetative storage organs, reproductive organs);
  2. anatomical (e.g. provascular differentiating sieve elements, protophloem sieve elements lacking companion cells, metaphloem se–cc complexes);
  3. developmental (e.g. cell division, cell expansion);
  4. metabolic (e.g. storage of soluble compounds/polymers, growth sinks).

A correspondingly large range of strategies for phloem unloading and sink utilisation must be anticipated.

5.4.1 - Cellular pathways of phloem unloading

Most photoassimilates travel along one of three cellular pathways: apoplasmic, symplasmic or a combination of both with symplasmic transport interrupted by an apoplasmic step (Figure 5.19).


Figure 5.18. Scheme describing the cellular pathways of phloem unloading and their relationship with sink types. (a) Apoplasmic unloading showing direct transport of photoassimilates from se—cc complexes to the phloem apoplasm. (b) Symplasmic unloading pathway which may or may not have an apoplasmic barrier between sieve elements and recipient sink cells. (c) Symplasmic unloading with the intervention of an apoplasmic step at (i) the maternal-filial interface of developing seeds and (ii) the vascular parenchyma-sink cell interface. Circled numbers denote different sink types assigned to each pathway. 1, vegetative apex; 2, elongating axis of a dicotyledonous stem; 3, mature axis of a primary dicotyledonous stem i permanent storage; 4, mature axis of a primary monocotyledonous stem i permanent storage; 5, mature primary root; 6, fleshy fruit; 7, developing seed. ab, apoplasmic barrier; apo, apoplasm; gp, ground parenchyma; sc, sink cell; se—cc, sieve element—companion cell complex; vp, vascular parenchyma.

(a) Apoplasmic pathways

Fig 5.31.jpg

Figure 5.19. Fluorescent micrograph of the distribution of a membrane-impermeant fluorescent dye, carboxyfluorscein (CF), imported through the phloem into roots of French bean (Phaseolus vulgaris L.). The green/yellow fluorescence of CF is confined to the se-cc complexes of mature portions of roots as seen by the thin central band of fluorescence away from the root apex. In contrast, dye spreads through the apex itself apparently via the symplasm of young cells. Scale bar = 2 mm

Photoassimilates can move directly across plasma membranes of se–cc complexes to the surrounding apoplasm (Figure 5.19a). Apoplasmic unloading is important along the axial transport pathway of roots and stems where vascular parenchyma and ground tissues serve as reversible storage sinks.

(b) Symplasmic pathways

An entirely symplasmic path of photoassimilate transport from sieve elements to recipient sink cells (Figure 5.18b) operates in a wide range of morphological and metabolic sink types. Terminal growth sinks such as root (Figure 5.19) and shoot apices, as well as vegetative storage sinks such as stems, roots and potato tubers, demonstrate symplasmic unloading.

In most sinks that exhibit symplasmic unloading, photo-assimilates are metabolised into polymeric forms within the recipient sink cells. Sugar cane is a notable exception because it stores sucrose unloaded symplasmically from sieve elements in parenchyma cells of stems. Stem sucrose reaches molar concentrations by this unloading route.

(c) Symplasmic pathway interrupted by an apoplasmic step 

Symplasmic discontinuities exist at interfaces between tissues of differing genomes including biotrophic associations (e.g. mycorrhizas and mistletoes) and developing seeds (Figure 5.18c). In addition, within tissues of the same genome, plasmodesmata can close permanently or reversibly at points along the post-sieve-element pathway. This necessitates photoassimilate exchange between symplasmic and apoplasmic compartments (Figure 5.18c). For instance, photoassimilate exchange between apoplasm and symplasm has been detected in sinks that store high solute concentrations and have unrestricted apoplasmic transport between vascular and storage tissues. Developing seeds, particularly of cereals and large-seeded grain legumes (Patrick and Offler 1995), are another model for symplasmic/apoplasmic pathways.

The apoplasmic space between maternal (seed coat) and filial (embryo plus endosperm) tissues in seeds prevents symplasmic continuity in the unloading pathway (Patrick and Offler 1995). In these organs, photoassimilates are effluxed across membranes of maternal tissues and subsequently taken up across the membranes of filial tissues (Figure 5.18c). Photoassimilates are unloaded from sieve elements and transported symplasmically to effluxing cells where they are released to the seed apoplasm. Influx from the seed apoplasm by the filial generation is restricted to specialised cells located at the maternal–filial interface. The final transport of photoassimilates to the filial storage cells largely follows a symplasmic route.

(d) Pathway linkage with sink function and pathway switching

The symplasm is the most frequently engaged cellular pathway of phloem unloading. Even where an apoplasmic step intervenes (e.g. developing seeds), photoassimilates travel predominantly through the sink symplasm (Figure 5.18c). Symplastic routes do not involve membrane transport and therefore offer lower resistances than apoplasmic routes.

Apoplasmic pathways are restricted to circumstances where (1) symplasmic transport compromises phloem translocation and (2) photoassimilate transport is between genetically distinct (e.g. maternal–filial) tissues. Phloem translocation would be compromised when solutes accumulate to high concentrations in sink cells were it not avoided by symplasmic isolation of phloem from sinks. This is exemplified by the switch to an apoplasmic step during development of tomato fruit. In young fruit, imported sugars are converted into glucose or fructose to support cell division and excess photoassimilate is accumulated as starch. At this stage, phloem unloading of photoassimilates follows a symplasmic route (Figure 5.18b). However, once sugars commence accumulating during cell expansion, apoplasmic transport is engaged (Figure 5.18c). The apoplasmic path isolates pressure-driven phloem import from rising osmotic pressures (P) occurring in fruit storage parenchyma cells (Patrick and Offler 1996).

Radial photoassimilate unloading in mature roots and stems may switch between apoplasmic or symplasmic routes depending upon the prevailing source/sink ratio of the plant. At low source/sink ratios, photoassimilates remobilised from axial stores are loaded into the phloem for transport to growth sinks (Wardlaw 1990). Under these conditions, symplasmic unloading into axial stores might be blocked by plasmodesmal closure while photoassimilates are absorbed by se–cc complexes from the surrounding apoplasm. This would prevent futile unloading while stores are drawn upon. In contrast, net flow of photoassimilates into axial storage pools at high source/sink ratios would be facilitated by plasmodesmal opening.

5.4.2 - Mechanisms of phloem unloading

(a) Apoplasmic transport


Figure 5.20. Mechanistic model for plasma membrane transport of sucrose from the coat and into the cotyledons of a developing legume seed. Plasma membrane ATPases vectorially pump protons to the seed apoplasm from both the opposing seed coat and cotyledon cells. The proton gradient is coupled to drive sucrose efflux from the seed coats through a sucrose/proton antiporter and sucrose influx into the cotyledons by a sucrose/proton symporter.

Se–cc complexes contain high sugar concentrations (Section 5.2.3(b)). Thus, a considerable transmembrane concentration gradient exists to drive a passive leakage of sugars to phloem apoplasm. Sugars leaked to phloem apoplasm are often retrieved by an active sucrose/proton symport mechanism (Figure 5.20). Thus, net efflux of sugars from se–cc complexes is determined by the balance between a passive leakage and sucrose/proton retrieval.

Passive unloading (Ep) of sucrose from se-cc complexes to the phloem apoplasm (Equation 5.6) is determined by the permeability coefficient (P) of se–cc complex plasma membranes and the transmembrane sucrose concentration (C) gradient between sieve element lumena (se) and surrounding phloem apoplasm (apo).

 \[E_p=P(C_{se}-C_{apo}) \tag{5.6} \]

 Sinks containing extracellular invertase (e.g. developing tomato fruit, sugar beet tap roots, maize seeds) can hydrolyse sucrose, lowering Capo thereby enhancing sucrose unloading from se–cc complexes. Furthermore, hydrolysis of sucrose renders it unavailable for se–cc complex retrieval by sucrose/proton symport. The resulting hexoses can act as signals to promote cell division in many sinks such as developing seed of Vicia faba.

(b) Symplasmic transport


Figure 5.21. Externally supplied solutes have a marked effect on sucrose import into root tips of hydroponically grown pea seedlings. This was tested by immersing root tips in (a) sucrose or (b) mannitol solutions ranging up to 350 mM. Cotyledons, which supply these young roots with carbon, were fed 14C sucrose and 14C arriving in different root parts was measured (Bq per root segment). (a) Import of 14C into root tips was diminished when they were exposed to external sucrose concentrations of less than 100 mM but promoted by sucrose concentrations of 150 to 350 mM; Two effects operate. At low concentrations, sucrose might enter root tip cells and suppress phloem import by a feedback mechanism. At higher concentrations, sucrose might act mainly as an osmoticum (see (b)). Mannitol is not taken up or metabolised quickly and can therefore help answer these questions. (b) Import through the phloem was stimulated by exposing root tips to the slowly permeable sugar mannitol, at concentrations of from 13 to 350 mM. This demonstrates an osmotic dependence of import through the phloem pathway, presumably through progressively decreasing P of root tip cells as external solute concentrations rise (Based on Schultz 1994)

Symplasmic transport is mediated by cytoplasmic streaming in series with intercellular transport via plasmodesmata. Plasmodesmal transport is usually the overriding resistance determining transport rates between cells.

Root tips offer a useful experimental model to explore post-sieve-element symplasmic transport because of morphological simplicity and accessibility. Exposing pea root tips to low sucrose concentrations (<100mM) slowed photoassimilate accumulation (Figure 5.21a) by raising intracellular sucrose concentrations. This response to concentration gradients is consistent with a diffusion component to phloem unloading (Equation 5.7). When roots were bathed in much higher concentrations of either sucrose (Figure 5.22a) or a slowly permeating solute, mannitol (Figure 5.33b), turgor pressure (P) of sieve elements and surrounding tissues decreased and 14C import rose. This is consistent with a hydraulically driven (bulk) flow of photoassimilates into the root apex. Thus, photoassimilate movement from phloem through a symplasmic path can be mediated by diffusion and/or bulk flow. The relative contribution of each transport mechanism depends on the magnitude of concentration and pressure gradients (Equations 5.6 and 5.8).

Physical laws can be used to model diffusion and bulk flow of sucrose through a symplasmic route. Sucrose diffuses through symplasm at a rate (Rd) defined by the product of plasmodesmal number in the path (n), plasmodesmal conductivity to diffusion (Kd) and sucrose concentration difference (ΔC) between sieve elements and sink cell cytoplasm. That is:

 \[R_d=n \cdot K_d \cdot \Delta C \tag{5.7} \]

 Transport by bulk flow (Rf) is determined by the product of flow speed (S), cross-sectional area of the plasmodesmal flow path (A) and concentration (C) of sucrose transported (Equation 5.2). Flow speed (S), in turn, is a product of hydraulic conductivity (Lp) of a plasmodesma and turgor pressure difference (ΔP) between se–cc complexes and recipient sink cells (Equation 5.8). Flow over the entire pathway considers the number of interconnecting plasmodesmata (n). Thus, bulk flow rate (Rf) is given by:

 \[R_f=n \cdot L_p \cdot \Delta P \cdot A \cdot C \tag{5.8} \]

 Equations 5.7 and 5.8 predict that sink control of symplasmic photoassimilate transport resides in plasmodesmal conductivity and/or sucrose metabolism/compartmentation.

Sucrose metabolism within sink cells influences cytoplasmic sucrose concentration and Πsink. The difference between Πsink and Πapo determines P (Section 4.3). Sucrose metabolism and compartmentation can affect sucrose concentration gradients and ΔP, both driving forces for symplasmic transport from se–cc complexes to sink cells (Equations 5.6 and 5.8).

Fig 5.34.jpg

Figure 5.22. Cellular distribution of the apoplasmic tracer fluorescent dye 3-hydroxy—5,8,10—pyren—etrisulphonate (PTS) imported through the xylem in stem explants of sugar cane. (Left) Fluorescent micrograph of a longitudinal section of a stem with PTS (green fluorescence) localised to the vascular bundle. (Right) Fluorescent micrograph of a transverse section showing PTS confined to the vascular bundle. Retention of PTS in vascular bundles demonstrates that a barrier to lateral dye movement must be located in the walls of bundle sheath cells (bs). (Based on Jacobsen et al. 1992)

Transgenic plants which under- or over-express key sugar metabolising enzymes have allowed definitive experiments to be carried out on the role of sucrose metabolism in symplasmic phloem unloading. For example, reduction of sucrose synthase activity (Section 5.4.4) in tubers of transformed potato to 5–30% of wild-type levels depressed dry weight of tubers and starch biosynthesis (Table 5.3). Tubers of transformed plants had very high hexose levels (hence high P) which might contribute to downregulation of photoassimilate import. As a corollary, plants with enhanced starch biosynthesis through overexpression of the key starch synthesising enzyme, ADP-glucose pyrophosphorylase (Section 5.4.5), also had higher rates of photoassimilate import.

For sinks that store sugars to high concentrations (e.g. sugar cane stems), gradients in Π, and hence P, between se–cc complexes and sink storage cells could become too small to sustain transport. Instead, P in the apoplasm of storage tissues increases as sucrose (hence Π) in the storage cell sap rises. This maintains a lower P in storage cells than in sieve elements and sustains transport. High sucrose concentrations in the apoplasm of storage cells is achieved through an apoplasmic barrier which isolates storage parenchyma cells from sieve elements (Figure 5.22).

(c) Symplasmic transport interrupted by an apoplasmic step

Fig 5.35_0.jpg

Figure 5.23. Experimental systems used to determine sucrose fluxes in (a) attached caryopses of wheat and (b) coats of developing legume seed. In (a), sucrose effluxed from the maternal tissues was collected by infusing the endosperm cavity of an attached wheat grain with solutions delivered and retrieved through micro-capillaries (Wang and Fisher 1994). In (b), embryos are surgically removed from the coats which may be (i) attached to or (ii) detached from the pod wall. The space vacated by the embryo is filled with a wash solution that is changed at frequent intervals. The wash solution is used to deliver treatments to the seed coat and as a trap to collect the effluxed sucrose.

Phloem unloading in legume seed pods is one case of symplasmic and apoplasmic transport operating in series; the pathway is described in Section 5.4.2(c). Whether sucrose efflux requires energy remains unknown since concentration gradients between seed coats and apoplasm might be steep enough to drive facilitated diffusion. Indeed, using an elegant infusion technique (Figure 5.23a), Wang and Fisher (1994) concluded that efflux from the nucellar projection cells of wheat grain was unlikely to be energy dependent. In contrast, sucrose efflux from coats (maternal tissue) of surgically modified legume seeds (Figure 5.23b) is inhibited by about 50% in the presence of PCMBS, a membrane transport inhibitor. Efflux from legume seed coat cells exhibits charac-teristics of a sucrose/proton antiport. Sucrose uptake by filial tissues is mediated by sucrose/proton symport (Figure 5.23).

A fascinating aspect of phloem unloading in legume seed pods is how photoassimilate demand by filial tissues is integrated with supply from maternal tissues, itself an integration of photoassimilate efflux and import from phloem. One variable that could regulate rates of photoassimilate transport through seed coat symplasm and efflux into apoplasm of the maternal–filial interface is P of seed coat cells (Psc): this would sense depletion of apoplasmic sucrose through uptake by cotyledons, producing a signal in the form of a ΔPsc (Figure 5.24). Specifically, Psc is determined by ΔΠ between the seed coat (Πsc) and seed apoplasm (Πapo), which fluctuates according to photoassimilate withdrawal by cotyledons.

A pressure difference (ΔP) between the points of photo-assimilate arrival (sieve tubes) and efflux (seed coats) drives bulk flow of photoassimilates through the seed coat symplasm. Turgor pressure of seed coat efflux cells is maintained homeostatically at a set point (Pset) by P-dependent efflux into the seed apoplasm. Changes in apoplasmic assimilate concentrations and hence Π are sensed immediately as deviations of Psc from Pset. A rise in Psc produced by photoassimilate depletion around filial tissues elicits an error signal, activating P-dependent solute efflux (Figure 5.25b) and thereby raising photoassimilate concentrations in the apoplasm to meet demand by cotyledons (Figure 5.24c). Long-term increases of sucrose influx by cotyledons, for example over hours, are accompanied by adjustments in Pset (light to dark arrows in Figure 5.24b) which elicit commensurate increases in phloem import rates (light to dark arrows in Figure 5.24a).


Figure 5.24. A turgor-homeostat model describing the integration of photoassimilate transport to developing legume seeds. Photoassimilate import through phloem (a) and efflux from seed coat to seed apoplasm (b) is mediated by a turgor (P)-dependent efflux mechanism and uptake of sugars by cotyledons (c). Metabolic activity in growing seeds influences sucrose concentrations within cotyledon cells, possibly feeding back on activities of symporters located in plasma membranes of the cotyledon dermal cell complex. Graph (c) denotes increased influx (R) from light to dark curve as sugar demand increases. The apoplasmic solute pool size is small and turns over in less than one hour (Patrick and Offler 1995). Thus, faster photoassimilate withdrawal from the seed apoplasm by cotyledons will rapidly lower apoplasmic osmotic concentration. Since the osmotic difference between seed apoplasm (Πapo) and seed coat (Πsc) is only 0.1-0.2 MPa, a small decrease in osmolality of the apoplasm will elicit a significant increase in seed coat P (Psc).A shift in Psc above the turgor set point (Pset) results in an error signal (see model) which in turn induces an immediate compensatory increase in photoassimilate efflux to the apoplasm (light curve in graph b). Increased photoassimilate efflux acts to maintain a constant Πapo in spite of enhanced flux through the apoplasm (graph c). Consequently, the increased potential for photoassimilate uptake by cotyledons can be fully realised (dark curve in graph c). In the short term (minutes), the turgor-homeostat ensures that Psc is maintained and hence phloem import, which is driven by the turgor difference between sieve tubes and unloading cells (PstPsc) is also maintained. Under conditions where cotyledon demand is sustained, Pset in the seed coat adjusts downwards. This results from decreases in Πsc, while Πapo is homeostatically maintained. The decrease in Pset of efflux cells serves to enhance the pressure difference between these cells and the importing sieve elements. As a result, the rate of phloem import into seed coats (Rimport) is increased (graph a, light to dark curve). This new rate of import is commensurate with accelerated sucrose efllux from seed coats to the apoplasm (graph b, light to dark curve) and, ultimately, cotyledons.

5.4.3 - Sugar metabolism and compartmentation in sinks

The fate of imported photoassimilates depends on sink cell function. In broad terms, imported photoassimilates are primarily used to provide carbon skeletons or signals for growth or storage. Some photoassimilates provide energy for maintenance. Relative flows of photoassimilates to these fates change during cell development and sometimes over shorter time scales depending upon a plant’s physiological state.

(a) Cell maintenance

Irrespective of sink function, a portion of imported sugars is respired to provide energy (ATP) for maintenance of cell function and structure. Most of this energy is required for continual turnover of cellular constituents such as enzymes and mem-branes. Rates of synthesis and degradation of individual macromolecules vary widely, as does the energy invested in different molecular configurations, so sugar demand for maintenance respiration could differ substantially between tissues.

(b) Cell growth

In growing organs, photoassimilates become substrates for synthesis of new cell material either directly or after biochemical conversions. Other fates for sugars include catabolism in energy-generating pathways which support growth (growth respiration) and storage in vacuolar pools. Stored sugars make an osmotic contribution to growing cells and can act as energy stores in species such as sugar cane. In roots of young barley plants, 40% and 55% of imported sugars are respired and used in structural growth, respectively. Stored sugars turn over each 30 min but account for only 1% of root weight.

(c) Reserve storage in cells

In mature cells, imported sugars enter physical (e.g. vacuoles) and chemical (e.g. starch) storage pools with lesser amounts diverted to respiration (15–20%) and structural components. In contrast to growth sinks, stored carbohydrates are ultimately retrieved from storage pools and used by other storage sinks (e.g. germinating seeds) or translocated to support growth and storage processes elsewhere in the plant. Carbohydrate storage can be brief (hours, days) or extend over considerable periods (months to years). Short-term storage of carbohydrates in stems and roots buffers phloem sap sugar concentrations against changes in photoassimilate export from photosynthetic leaves.

Sugars can also be stored in soluble forms by compartmentation into vacuoles. In this case, the tonoplast provides a physical barrier to protect stored sugars from molecular interconversion by cytoplasmic sugar-metabolising enzymes. Vacuolar sugars are accumulated as sucrose, hexoses or fructans (short-chain polymers of fructose). Sucrose and hexoses can accumulate to molar concentrations (0.1–1.5M) in storage parenchyma cells of roots, stems and fruits. For instance, tap roots of sugar beet and stems of sugar cane accumulate 1M sucrose thereby providing 90% of the world’s sucrose. Hexoses are a common form of sugar storage in fruit, contributing to sweetness of edible fruits such as tomato, grape, orange and cucumber. The wine industry depends upon hexoses accumulating to high concentrations (1.5M) in grape berries to fuel fermentation of ‘must’ in wine making. Fructans are stored in significant quantities in leaf sheaths and stems of temperate grasses and cereals. In pasture species, they contribute to forage quality, and in cereals constitute an assimilate pool that is mobilised to support grain filling.

Alternatively, imported sugars may be stored as starch along the axial transport pathway (available for remobilisation to buffer phloem sap sugar concentrations) or in more long term storage pools of terminal sink organs such as tubers, fruits and seeds. The proportion of photoassimilates diverted into starch differs widely between species and organs. Starch accounts for some 90% of dry weight of potato tubers and cereal grains.

The chemistry of storage products can change during organ development. For instance, starch is the principal storage carbohydrate in young tomato fruit. Later in fruit development, stored starch is hydrolysed and contributes to hexose accumulation in vacuoles of fruit storage parenchyma cells. In other fruits, significant switches between hexose and sucrose accumulation occur during development. All these changes are brought about by ontogenetic shifts in activities of sugar-metabolising enzymes.

5.4.4 - Key transfer events in sugar metabolism and compartmentation

Phloem-imported sucrose can reach the cytoplasm of recipient sink cells chemically unaltered or be hydrolysed en route by extracellular invertase into its hexose moieties. These sugars may then enter a number of metabolic pathways or be compartmented to vacuolar storage (Figure 5.25).


Figure 5.25. Pathways of sugar metabolism and compartmentation within sink cells. Sugars can be delivered to sink cells through either apoplasmic or symplasmic pathways. Within the sink apoplasm, sucrose can be hydrolysed to hexoses by an extracellular invertase. Apoplasmic sugars are transported across plasma membranes of sink cells by proton/ sugar symporters. Alternatively, sucrose enters the sink cytoplasm through a symplasmic path. Within the sink cytoplasm, sucrose can be hydrolysed or compartmented into vacuolar storage. Sucrose hydrolysis provides substrates for energy metabolism or for synthesis of macro-molecules. Invertase activity is important to sustain hexose supply for glycolysis. Sucrose synthesis from the hexose pool is catalysed by sucrose phosphate synthase (SPS). Degradation of sucrose by sucrose synthase generates fructose and uridine diphosphate glucose (UDP-glucose) which enters various biosynthetic pathways including cellulose and starch synthesis. In the case of starch biosynthesis, UDP-glucose and fructose generate glucose-1-phosphate (G-1-P) which is transported across the amyloplast membrane. Accumulated G-1-P is interconverted into adenine diphosphate glucose (ADP-glucose) by the enzyme adenine diphosphate glucose pyrophosphorylase (ADP-glucose PPase). ADP-glucose is the substrate for starch polymer formation. Sucrose compartmentation into the vacuole is mediated by a sucrose/ proton antiporter located on the tonoplast. Within the vacuole, sucrose can be exposed to invertase hydrolysis with the hexose products accumulating or leaking back to the cytoplasm

(a) Sucrose metabolism

Sucrose is metabolically inert and, in order to be metabolised, must be hydrolysed to glucose and fructose. Only two enzymes are capable of metabolising sucrose in green plants. These are invertase and sucrose synthase (Figure 5.25) and they are paramount in sugar metabolism after phloem unloading.

Invertase catalyses irreversible hydrolysis of sucrose to its hexose moieties, glucose and fructose. Both acid and neutral invertases occur in plants, with pH optima of about 5 and 7.5, respectively. The activity of invertases varies with plant species, organ type and stage of development. Acid invertases, located in cell wall or in vacuole, are usually active in rapidly growing leaves, stems and fruits and seeds (Ruan et al. 2010), making hexoses available for regulating gene expression and for respiration and biosynthesis. Reduced acid invertase activity in vacuoles during development of sugar cane stems, and its absence from sucrose-accumulating tomato fruit, is a major factor in sucrose accumulation in vacuoles of these tissues. Suppression of cell wall invertase activity led to shrunken seed in maize and small fruit in tomato and loss of pollen fertility in tomato, wheat and rice, demonstrating its critical roles in these reproductive organs. Less is known about the physiological role of neutral invertases..

Sucrose synthase is mainly located in the cytoplasm but recent research also shows that the enzyme may also be associated with plasma membrane and even present in cell wall matrix.  It catalyses sucrose cleavage to fructose and UDP-glucose, a high-energy ester of glucose. UDP-glucose is a substrate for biosynthesis of cellulose and may be converted further for starch synthesis.High activities of sucrose synthase are found in both growing and starch storage tissues. In the cytoplasm of starchy tissues, UDP-glucose is converted by UDP-glucose pyro-phosphorylase to glucose-1-phosphate, which is transported across amyloplast membranes. In amyloplasts, glucose-1-phosphate provides glucose moieties for starch synthesis in a pathway comparable to starch formation in chloroplasts of photosynthetic leaves. The critical role of sucrose synthase in starch synthesis is demonstrated with potatoes transformed with an antisense construct of the gene encoding tuber-specific sucrose synthase. Tuber sucrose synthase activity in transformed plants was depressed significantly while the activities of key starch biosynthetic enzymes were unaltered. Low sucrose synthase activity was directly responsible for a proportional decrease in starch accumulation (Zrenner et al. 1995).

Sinks that accumulate soluble sugars have predictably low sucrose synthase activities. Contrastingly high sucrose synthase activities in phloem vessels may be responsible for energy production for phloem loading or unloading and maintaining cellular function of companion cells.

(b) Hexose metabolism

Hexoses transported to the sink cytoplasm are rapidly phosphorylated to hexose-6-phosphates by glucose- and fructose kinases. In these forms, hexoses can be used as substrates for respiration or for synthesis of new cell constituents. Alternatively, sucrose phosphate synthetase can convert them to sucrose, as in leaves (Chapters 1 and 2). Sucrose synthesized by this reaction can be accumulated in vacuoles (e.g. sugar beet tap roots, sugar cane stems) or be rehydrolysed into hexoses by a vacuolar acid invertase (e.g. grape berries).

5.4.5 - Sink control of photoassimilate partitioning

The pressure-flow hypothesis provides a compelling model to explain sink strength in plants. Evidence such as accelerated import of photoassimilate into roots with artificially lowered P lends empirical support to the model.

Knowledge of cellular and molecular events in phloem unloading and photoassimilate use begins to reveal the array of control steps which underlie photoassimilate unloading and relative sink strength. Photoassimilate import into sinks by apoplasmic pathways or by diffusion through symplasmic pathways (Figure 5.19) is controlled by P in se–cc complexes. In contrast, when phloem unloading is by bulk flow through a symplasmic route (e.g. legume seed coats), P in cells responsible for photoassimilate efflux to the apoplasm controls unloading. Unloading into storage tissues is controlled by P in sink (storage) cells.

These processes at the cell and tissue level must now be related to a whole-plant perspective of sink control of photoassimilate partitioning, taking into account influences of plant development and environmental factors. How plants use photoassimilates (e.g. switch from growth to storage) is accompanied by alterations in the cellular pathway of import. Phytohormones also play a role in photoassimilate partitioning through their influence on development and intercellular signalling.

Plant development generates new sinks, for example in meristems where cells undergo division or growth zones where enlarging cells import photoassimilates.

(a) Meristematic sinks

Potential sink size is set largely during the meristematic phase of development through determination of total cell number per organ. Photoassimilate supply has been implicated as a limiting factor in initiation of leaf primordia at the apical dome and subsequent early development by cell division. Substrate supply for developing seeds (endosperm, embryo) and root and floral apices might also be restricted.

Agricultural yields might therefore increase if plants could be modified to enhance the supply of photoassimilates to meristems. Which factors regulate photoassimilate supply to meristematic sinks? Rate equations describing mass flow of phloem sap (Section 5.2.5) predict that photoassimilate supply reaching a sink will be determined by source output (setting photoassimilate concentration in sap and P in sieve elements at the source) and modulated by Lp of the transport pathway. Increased photoassimilate output from source leaves increases growth activity of primary meristems. Even during reduced source output, photoassimilate import and meristematic sink strength can be maintained by remobilisation of storage reserves. Manipulating competition for photo-assimilates by more established sinks also suggests that source output influences sink behaviour.

Cultivated plants demonstrate these principles. For example, flushing CO2 into glasshouses increases flower set and hence yield of floral and fruit crops. Similarly, applying growth regulators to induce abscission of some floral apices lessens the number of sinks competing for photoassimilates at fruit set and leads to larger and more uniform fruit at harvest. Alternatively, breeding programs have reduced sink strength of non-harvestable portions of crops and hence the severity of competition. For instance, breeding dwarf varieties of cereals has reduced photoassimilate demand by stems with a consequent increase in floret numbers set and grain size.

These observations imply that increases in net leaf photosynthesis and phloem loading should set higher yield potentials. Yet meristems import only a small proportion of total plant photoassimilate. It may be that phloem conductance limits photoassimilate delivery to meristems; increases in source output would amplify the driving force for transport and hence bulk flow through a low-conductance pathway.

Given that mature phloem pathways have spare transport capacity (Section 5.2.5), any transport limitation imposed by low path conductance might be expected within immature sinks. Photoassimilate import into meristematic sinks involves transport through partially differentiated provascular strands that might extend up to 400µm. Movement through this partially differentiated path is symplasmic (Section 5.2.2(b)). Hence, plasmodesmal numbers and transport properties of plasmodesmata could play a critical role in photoassimilate supplies to sinks and determination of sink size (Equation 5.7).

(b) Expansion and storage sinks 

As cells expand and approach cell maturity, photoassimilates are increasingly diverted into storage products. Towards maturity, fully differentiated phloem pathways with spare transport capacity link expansion/storage sinks with photosynthetic leaves. Photoassimilate import by these sinks depends on duration of the storage phase. This can be short term for sinks located along the axial transport pathway and long term for sinks sited at the ends of transport pathways (e.g. tubers, fruits and seeds).

Storage along the axial pathway occurs mainly when photoassimilate production exceeds photoassimilate demands by terminal sinks. However, storage is not necessarily a passive response to excess photoassimilate supply. Stems of sugar cane store large quantities of photoassimilates (50% of dry weight is sucrose) even during rapid growth of terminal sinks. Photoassimilates might be stored as simple sugars (e.g. sugar cane stems) or as polymers (fructans in stems of temperate grasses; starch in stems and roots of subtropical cereals, herbaceous annuals and woody perennials). Photoassimilates stored along axial pathways buffer against diurnal and more long term fluctuations in photoassimilate supply to terminal sinks. In woody deciduous species, axially stored photoassimilates also provide a long-term seasonal storage pool that is drawn on to support bud growth following budburst. Remobilised photoassimilates can contribute substantially to biomass gain of terminal sinks. For instance, in some mature trees, over half the photoassimilates for new growth come from remobilised reserves; similar proportions of stem-stored fructans contribute to grain growth in cereals when photoassimilate production is reduced (e.g. by drought). Physiological switching between net storage and remobilisation is an intriguing regulatory question.

Growth and development of meristems is determined by phloem unloading events and metabolic interconversion of photoassimilates within recipient sink cells. These transport and transfer processes vary between sinks and can alter during sink development. Techniques now exist to alter expression of membrane porter proteins and possibly enhance photoassimilate import by sinks such as seeds which have an apoplasmic step in the phloem unloading pathway. Prospects of altering plasmodesmal conductivity will improve once plasmodesmal proteins are identified and their encoding genes known.

5.4.6 - Follow the flow: unloading of water and its destination

Phloem unloading of nutrients follows water release from vascular system.

As discussed before, because of their low transpiration rates, developing sinks typically import water through phloem, not xylem.  Water unloaded from phloem is used for cell growth or recycled back to the parental bodies.  Water is unloaded from se-cc complexes symplasmically in majority of sinks by bulk flow. For growth sinks such as shoot or root apices, continued symplasmic flow of phloem-imported water can drive cells expansion. In post-phloem unloading pathways interrupted with an apoplasmic step, water must exit cells of the unloading path across cell membranes, facilitated by aquaporins (AQPs). AQPs responsible for water flow across cell membranes are plasma membrane intrinsic proteins (PIPs) and tonoplast intrinsic proteins (TIPs). Strong PIP expression in expanding post-veraison grape berries has been shown to correlate with water flows into, and from, the berry apoplasm. For sinks that stop expansion but continuously accumulate biomass, water transported to storage sink apoplasms is recycled back to the parent plant body through a xylem route. Important roles played by AQPs in water recycling are indicated by their high expression at this stage in developing seed, particularly in the vascular parenchyma cells.

In conclusion, this chapter has shown how growth and development of meristems and other sinks is determined by phloem unloading events, and metabolism of the assimilates within the recipient sink cells. These transport and transfer processes vary between specific sinks, and can alter during development. Molecular techniques that alter expression of membrane transporters  can be used to study the pathways and limitations of photoassimilate transport into sinks such as seeds that have an apoplasmic step in the phloem unloading pathway, with the possibility of enhancing the rate of grain growth and crop yield in the future.

5.5 - References

Aoki K, Suzui N, Fujimaki S et al. (2005) Destination-selective long-distance movement of phloem proteins. Plant Cell 17: 1801-1814.

Brenner M (1987) The role of hormones in photosynthate partitioning and seed filling. In PJ Davies, ed, Plant Hormones and their Role in Plant Growth and Development, Kluwer, Dordrecht, pp 474-493

Canny MJ (1973) Phloem Translocation. Cambridge University Press

Chen LQ, Qu XQ, Hou BH, Sosso D, Osorio S, Fernie AR, Frommer WB (2012) Sucrose efflux mediated by SWEET proteins as a key step for phloem transport. Science 335: 207-211

Covarrubias AA, Reyes JL (2010) Post-transcriptional gene regulation of salinity and drought responses by plant microRNAs. Plant Cell Environ 33:481-489.

Daie J, Watts M, Aloni B, Wyse RE (1986) In vitro and in vivo modification of sugar transport and translocation in celery by phytohormones. Plant Sci 46: 35-41

Fisher DB, Frame JM (1984) A guide to the use of the exuding stylet technique in phloem physiology. Planta 161: 385–393

Frommer WB, Hirner B, Kuhn C, Harma K, Martin T, Reismeier JW, Schulz B (1996) Sugar transport in higher plants. In M Smallwood, JP Knox, DJ Bowles, eds, Membranes: Specialised Functions in Plants. Bioscientific, Oxford, pp 319-335

Geiger DR (1975) Phloem loading. In Zimmerman MH, Milburn JA, eds, Transport in Plants 1. Phloem Transport. Springer-Verlag, Heidelberg, pp 395-431

Grusak MA, Beebe DU, Turgeon R (1996) Phloem loading. In Zamski E, Schaffer AA eds, Photoassimilate Distribution in Plants and Crops. Source-Sink Relationships. Marcel Dekker, New York, pp 209-228 

Jacobsen KR, Fisher DG, Maretzki A, Moore PH (1992) Developmental changes in the anatomy of the sugarcane stem in relation to phloem unloading and sucrose storage. Bot. Acta 105: 70-80

Kallarackal J, Milburn JA (1984) Specific mass transfer and sink-controlled phloem translocation in castor bean. Aust J Plant Physiol 11: 483-490 

Kriedemann P, Beevers H (1967) Sugar uptake and translocation in the castor bean seedling. II. Sugar transformations during uptake. Plant Physiol 42: 174-180

Lough TJ, Lucas W. (2006) Integrative plant biology: role of phloem long-distance macromolecular trafficking. Annu Rev Plant Biol 57: 203–232

Milburn JA, Baker DA (1989) Physico-chemical aspects of phloem sap. In DA Baker, JA Milburn, eds, Transport of Assimilates. Longman, Harlow, pp 345-359

Milburn JA, Kallarackal J (1989) Physiological aspects of phloem translocation. In DA Baker, JA Milburn, eds, Transport of Assimilates. Longman, Harlow, pp 264-305

Moorby J, Jarman PD (1975) The use of compartmental analysis in the study of movement of carbon through leaves. Planta 122: 155-168

Münch E (1930) Die Stoffbewegungen in der Pflanze. Fischer, Jena 

Pate JS, Peoples MB, Atkins CA (1984) Spontaneous phloem bleeding from cryopunctured fruits of a ureide-producing legume. Plant Physiol 74:  499-505

Patrick JW (1997) Phloem unloading: sieve element unloading and post-sieve element transport. Annu Rev Plant Physiol Plant Mol Biol 48: 191–222

Patrick JW, Offler CE (1995) Post-sieve element transport of sucrose in developing seed. Aust J Plant Physiol 22: 761-769

Patrick JW, Offler CE (1996) Post-sieve element transport of photoassimilates in sink regions. J Exp Bot 28: 736-743

Rodriguez-Medina C, Atkins CA, Mann A, Jordan M, Smith PMC (2011) Macromolecular composition of phloem exudate from white lupin (Lupinus albus L.). BMC Plant Biol 11: 36-54

Ruan YL, Jin Y, Yang YJ, Li GJ, Boyer JS (2010)  Sugar input, metabolism, and signaling mediated by invertase: roles in development, yield potential, and response to drought and heat. Mol Plant 3: 942-955

Schulz A (1994) Phloem transport and differential unloading in pea seedlings after source and sink manipulations. Planta 192: 239-249 

Turgeon R (2010) The role of phloem loading reconsidered. Plant Physiol 152: 1817-1823

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Wang N, Fisher DB (1994) Monitoring phloem unloading and post-phloem transport by microperfusion of attached wheat grains. Plant Physiol 104: 7-16

Wright JP, Fisher DB (1980) Direct measurement of sieve tube turgor pressure using severed aphid stylets. Plant Physiol 65: 1133-1135

Varkonyi-Gasic E, Gould N, Sandanayaka M, Sutherland P, MacDiarmid RM (2010) Characterisation of microRNAs from apple (Malus domestica ’Royal Gala’) vascular tissue and phloem sap. BMC Plant Biol 10: 159-173

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Chapter 6 - Growth analysis: a quantitative approach


Highly productive multiple cropping in a CO2-enriched greenhouse at CSIRO Merbein (Original photograph courtesy E.A. Lawton)

Chapter editors: Charles Price1 and Rana Munns1,2

1School of Plant Biology, University of Western Australia, 2 CSIRO Plant Industry, Canberra

This chapter is updated from the original by PE Kriedemann, JM Virgona and OK Atkin (1st Edition)

Growth is an irreversible increase in plant size accompanied by a quantitative change in biomass (weight). Development is more subtle and implies an additional qualitative change in plant form or function such as a phase change from vegetative to reproductive growth.

Growth analysis is a conceptual framework for resolving the nature of genotype x environment interactions on plant growth and development.

In natural environments, growth and development cycles have to be completed within a time frame dictated by environmental conditions where light, moisture and nutrients often limit expression of genetic potential. Adaptive features that counter such constraints and help sustain relative growth rate can be revealed via growth analysis under contrasting conditions.

In managed environments, crop plants commonly experience similar restrictions, but in addition their economic yield is often only a small portion of total biomass at harvest and subject to genetic control. Crop scientists need to explore plant growth and reproductive development in quantitative terms. Sources of variation in productivity can then be resolved into those processes responsible for converting external resources into biomass and those responsible for partitioning biomass into usable sinks such as cereal ears or pumpkins. Both aspects are addressed here.

6.1 - Concepts and components of RGR

The most useful and widely used analysis is the concept of relative growth rate (RGR) and the simple RGR equation, which derives from the growth of cell populations with unrestricted resources – that is where light, space and nutrient supply are not limiting.

Growth models developed from populations of single cells can be extended mathematically to cover complex multicellular organisms where whole-plant growth is expressed in terms of leaf area and nutrient resources. Such growth indices are not intrinsic properties of plants, but rather mathematical constructs with functional significance. These concepts can be traced to the early 1900s and have proved increasingly useful for studies of growth and developmental responses in natural and managed environments.

6.1.1 - Cell populations

A small population of unicellular organisms presented with abundant resources and ample space will increase exponentially (Figure 6.1a). Population doubling time Td (hours or days) is a function of an inherent capacity for cell division and enlargement which is expressed according to environmental conditions. In Figure 6.1(a) doubling times for these two populations are 1 and 2 d for fast and slow strains respectively.

Fig 6.1.png

Figure 6.1. A population of cells unrestricted by space or substrate supply will grow exponentially. In this hypothetical case, a fast-growing strain of a single-celled organism with a doubling time of 1 d starts on day 0 with a population of n cells which increases to 120·n by day 7. In this example, n=10. The slow-growing strain with a doubling time of 2 d takes twice as long to reach that same size. When data for cell numbers are ln transformed, exponential curves (a) become straight lines (b) where slope = r.

Exponential curves such as those in Figure 6.1(a) are described mathematically as

\[N(t)=N_0e^{rt} \tag{6.1}\]

 where N(t) is the number of cells present at time tN0 is the population at time 0, r determines the rate at which the population grows, and e is the base of the natural logarithm. By derivation from Equation 6.1

\[r=\frac{1}{N}\frac{\text{d}N(t)}{\text{d}t} \tag{6.2}\]

and is called relative growth rate with units of 1/time. The doubling time is Td = (ln 2)/r.

If a population or an organism has a constant relative growth rate then doubling time is also constant, and that population must be growing at an exponential rate given by Equation 6.1. The ‘fast’ strain in Figure 6.1(a) is doubling every day whereas the ‘slow’ strain doubles every 2 d, thus r is 0.69 d–1 and 0.35 d–1, respectively.

If cell growth data in Figure 6.1(a) are converted to natural logarithms (i.e. ln transformed), two straight lines with contrasting slopes will result (Figure 6.1b). For strict exponential growth where N(t) is given by Equation 6.1,

 \[\text{ln }N(t)=\text{ln }N_0+rt\tag{6.3}\]

 which is the familiar slope-intercept form of a linear equation, so that a plot of ln N(t) as a function of time t is a straight line whose slope is relative growth rate r, and intercept is ln N0

In practice, r is inferred by assessing cell numbers N1 and N2 on two occasions, t1 and t2 (separated by hours or days depending on doubling time — most commonly days in plant cell cultures), and substituting those values into the expression

\[r=\frac{\text{ln }N_2-\text{ln }N_1}{t_2-t_1}\tag{6.4}\]

which expresses r in terms of population numbers N1 and N2 at times t1 and t2, respectively.

If growth is exponential, Eq. 6.3 will be linear and any two time points and the natural logarithm of their corresponding population sizes will give an estimate of the growth rate, r. However, if relative growth rate r is not constant, then growth is not exponential but the concept of relative growth rate is still useful for analysis of growth dynamics in populations or organisms. Equation 6.3 is then used to compute average relative growth rate between times t1 and t2 even though population growth might not follow Equation 6.1 in strict terms. In that case plots analogous to Figure 6.1(b) will not be straight lines.

6.1.2 - Plant biomass

In whole plants, cell number is an impractical measure of growth. Instead, fresh or oven-dried biomass (W) is generally taken as a surrogate for cell growth and referenced to the number of days elapsed between successive observations. Relative growth rate is now known as RGR rather than r and measured in days or weeks rather than hours.

Relative growth rate, RGR (d–1), can be expressed in terms of differential calculus as \(RGR=\frac{1}{W}\frac{\text{d}W}{\text{d}t}\) (compare Equation 6.2.) so that RGR is increment in dry mass (dW) per increment in time (dt) divided by existing biomass (W). Averaged over a time interval t1 to t2 during which time biomass increases from W1 to W2, RGR (d–1) can be calculated from

\[\text{RGR}=\frac{\text{ln }W_2-\text{ln } W_1}{t_2-t_1} \tag{6.5} \]

which is analogous to Eq, 6.4. Net gain in biomass (W) is the outcome of CO2 assimilation by leaves minus respiratory loss by the entire plant. Leaf area can therefore be viewed as a driving variable, and biomass increment (dW) per unit time (dt) can then be divided by leaf area (A) to yield the net assimilation rate, NAR (g m–2 d–1), where

\[\text{NAR}=\frac{1}{A}\frac{\text{d}W}{\text{d}T} \tag{6.6}\]

Averaged over a short time interval (t1 to t2 days) and provided whole-plant biomass and leaf area are linearly related (see Radford 1967),

\[\text{NAR}=\left(\frac{W_2-W_1}{t_2-t_1}\right)\left(\frac{\text{ln }A_2-\text{ln }A_1}{A_2-A_1}\right) \tag{6.7}\]

NAR thus represents a plant’s net photosynthetic effectiveness in capturing light, assimilating CO2 and storing photoassimilate. Variation in NAR can derive from differences in canopy architecture and light interception, photosynthetic activity of leaves, respiration, transport of photoassimilate and storage capacity of sinks, or even the chemical nature of stored products.

The following treatment assumes for simplicity that photosynthesis and the assimilation of CO2 occurs only in leaves, even though for many herbaceous or succulent species it occurs to a lesser degree also in stems. Since leaf area is a driving variable for whole-plant growth, the proportion of plant biomass invested in leaf area will have an important bearing on RGR, and can be conveniently defined as leaf area ratio, LAR (m2 g–1), where


LAR can be factored into two components: specific leaf area (SLA) and leaf weight ratio (LWR). SLA is the ratio of leaf area (A) to leaf mass (WL) (m2 g–1) and LWR is the ratio of leaf mass (WL) to total plant mass (W) (dimensionless). Thus,

\[\begin{align} \text{LAR} &=  \frac{A}{W_L}\frac{W_L}{W} \\
 &=  \text{SLA} \times \text{LWR} \end{align} \tag{6.9} \]

As an aside, average LAR over the growth interval t1 to t2 is

 \[\text{LAR}=\frac{1}{2} \left( \frac{A_1}{W_1}+\frac{A_2}{W_2} \right) \tag{6.10}\]

Expressed this way, LAR becomes a more meaningful growth index than A/W (Equation 6.8) and can help resolve sources of variation in RGR.

If both A and W are increasing exponentially so that W is proportional to A, it follows that


0r (substituting Equations 6.5, 6.6 and 6.8)

\[\text{RGR}=\text{NAR}\times \text{LAR} \tag{6.12}\]

As LAR can be broken into SLA and LWR (Equation 6.9) then

\[\text{RGR}=\text{NAR}\times \text{LWR} \times \text{SLA} \tag{6.13}\]

Sources of variation in RGR partitioned this way provide useful insights on driving variables in process physiology and ecology. For an expanded discussion on methodological issues associated with the determination of RGR in experimental populations see Poorter and Lewis (1986).

Increases in leaf area over time can be a more useful basis for measuring plant growth rates than biomass increases, particularly as non-destructive techniques for measuring leaf area are now available. Plant growth rate can be measured as the relative increase in leaf area over time, by substituting total plant leaf area for total biomass in the conventional RGR equation.

\[\text{RGR}_\text{A} =\frac{\text{ln } LA_2 - \text{ln } LA_1}{t_2-t_1} \tag{6.14} \]

where RGRA is relative leaf area expansion rate, LA is total plant leaf area and t is time at two time intervals, t1 and t2, preferably 2-3 days apart.

Growth indices in summary

Five key indices are commonly derived as an aid to understanding growth responses. Mathematical and functional definitions of those terms are summarised below.

Growth index

Mathematical definition


Functional definition

Relative growth rate




Rate of mass increase per unit mass present (efficiency of growth with respect to biomass)

Net assimilation rate



g m-2 d-1

Rate of mass increase per unit leaf area (efficiency of leaves in generating biomass)

Leaf area ratio



m2 g-1

Ratio of leaf area to total plant mass (a measure of ‘leafiness’ or photosynthetic area relative to respiratory mass)

Specific leaf area



m2 g-1

Ratio of leaf area to leaf mass (a measure of thickness of leaves relative to area)

Leaf weight ratio




Ratio of leaf mass to total plant mass (a measure of biomass allocation to leaves)


6.2 - Environmental impacts on RGR

Light, CO2, temperature, water and nutrients are key driving variables or limiting factors for growth responses in a wide range of species. Growth indices serve as an indicator of plant response, and of interactions between environmental factors where they occur. Variation in whole-plant RGR can be resolved into contributions from NAR (net assimilation rate) and LAR (ratio of leaf area to total plant mass). LAR in turn can be separated into SLA (ratio of leaf area to leaf mass) and LWR (ratio of leaf mass to total plant mass).

Ecological and agronomic implications for managed and natural communities are considered in this section in the context of growth responses to light, temperature and CO2 concentration. The same principles apply to the analysis of effects of nutrients on growth, and to interactions between environmental variables.

6.2.1 - Light

Light impacts both photosynthetic activity and morphology of individual leaves and of plant canopies. Leaves are larger at higher light. Leaf area increases because of more cells per leaf rather than by cells having a larger surface area. Cells volume, however, increases and gives rise to substantial increases in leaf thickness. This is usually achieved by a greater depth of palisade cells, either greater in depth or an extra layer of cells.

In the example shown for cucumber in Table 6.1, high light caused a three-fold increase in area, but the cell cross-section was the same, indicating that the leaves had three times the number of cells. The cell volume more than doubled under high light (3.11 × 10–5 mm3 at 3.2 MJ m–2 s–1 cf. 1.46 × 10–5 mm3 at 0.5 MJ m–2 d–1), and because cross-sectional area was virtually unchanged, cell depth was responsible. This greater depth of palisade cells in strong light confers a greater photosynthetic capacity (per unit leaf area) and translates into larger values for NAR and a potentially higher RGR. At lower irradiance (Table 6.1) leaves are thinner and SLA will thus increase with shading. Because LAR = SLA × LWR (Equation 6.9) a smaller leaf area at lower irradiance is offset to some extent by a higher SLA for maximum light capture with the most efficient use of resources.

The significance of LAR × NAR interaction for whole-plant growth was appreciated early by G.E. Blackman (Agriculture Dept, Oxford University), who in a series of papers analysed shade-driven growth responses for many species. RGR response to growing conditions in low light, and the degree to which upward adjustment in LAR could offset reduced NAR, was a recurring theme. In a series of 20 pot experiments, Blackman and Wilson (1951a) established a close relationship between NAR and daily irradiance where shade-dependent reduction in NAR was similar for 10 species. NAR was linearly related to log irradiance, and extrapolation to zero NAR corresponded to a light-compensation point of 6–9% full sun for eight species, and 14–18% full sun for two others. Significantly, neither slope nor intercept differentiated sun-adapted plants such as barley, tomato, peas and sunflower from two shade-adapted species (Geum urbanum and Solanum dulcamara). LAR proved especially responsive to light and accounted for contrasts between sun plants and shade plants in their growth response to daily irradiance.

Concentrating on sunflower seedlings, Blackman and Wilson (1951b) confirmed that NAR increased with daily irradiance (Figure 6.2) and that LAR was greatly decreased (Figure 6.2). Response in RGR reflected LAR and especially in young seedlings which also showed highest RGR and were most sensitive to shading. LAR appeared sensitive to both daily maxima as well as daily total irradiance.


Figure 6.2  A sun-adapted plant such as Helianthus annuus adjusts LAR to some extent in response to lower daily irradiance but not enough to maintain RGR. By contrast, a shade-adapted plant such as Impatiens parviflora with somewhat higher LAR and RGR in full sun makes further adjustment in LAR so that RGR does not diminish to the same extent in moderate or deep shade as does that of H. annuus (Based on Blackman and Wilson 1951b; Evans and Hughes 1961)

A comparison between sunflower (Blackman and Wilson 1951b) and the woodland shade plant Impatiens parviflora (Evans and Hughes 1961) confirms this principle of LAR responsiveness to irradiance (Figure 6.2). Sunflower achieved noticeably higher NAR in full sun than did I. parviflora, but LAR was considerably lower, and translated into a somewhat slower RGR for sunflower. This species contrast was much stronger in deep shade (12% full sun) where RGR for I. parviflora had fallen to 0.090 d–1 whereas sunflower was only 0.033 d–1. Clearly, I. parviflora is more shade tolerant, and retention of a faster RGR in deep shade is due both to greater plasticity in LAR as well as a more sustained NAR. Adjustments in both photosynthesis and respiration of leaves contribute to maintenance of higher NAR in shade-adapted plants growing at low irradiance.

6.2.2 - Temperature

Within a moderate temperature range readily tolerated by vascular plants (10–35°C, see Chapter 14) processes sustaining carbon gain show broad temperature optima. By contrast, developmental changes are rather more sensitive to temperature, and provided a plant’s combined responses to environmental conditions do not exceed physiologically elastic limits, temperature effects on RGR are generally attributable to rate of canopy expansion rather than rate of carbon assimilation. In the early days of growth analysis, Blackman et al. (1955) inferred from a multi-factor analysis of growth response to environmental conditions that NAR was relatively insensitive to temperature, but whole plant growth was obviously affected, so that photosynthetic area (LAR) rather than performance per unit surface area (NAR) was responsible. Such inferences were subsequently validated.

Using day/night temperature as a driving variable, Potter and Jones (1977) provided a detailed analysis of response in key growth indices for a number of species (Table 6.2). Data for maize, cotton, soybean, cocklebur, Johnson grass and pigweed confirmed that 32/21°C was optimum for whole-plant relative growth rate (RGR) as well as relative rate of canopy area increase (RGRA). Both indices were lowest at 21/10°C. This was true for C4 as well as C3 species.

C4 species had a higher RGR and RGRA than C3 species, especially under warm conditions (Table 6.2)

All populations described in Potter and Jones (1977) maintained strict exponential growth. NAR could then be derived validly and temperature effects on NAR could then be compared with temperature effects on RGR and RGRA (Figure 6.3). With day/night temperature as a driving variable, most values for NAR fell between 10 and 20 g m–2 d–1. Correlation between NAR and RGR was poor (Figure 6.3). By contrast, variation in both RGR and RGRA was of a similar order and these two indices were closely correlated (Figure 6.3).


Figure 6.3 Variation in whole-plant RGR is linked to relative rate of canopy expansion (RGRA). Nine species (including C3 and C4 plants) grown under three temperature regimes (21/10 °C, 32/21 °C and 21/27 °C day/night) expressed wide variation in RGR that showed a strong correlation with RGRA but was poorly correlated with variation in NAR. Extent rather than activity of leaves appears to be more important for RGR response to temperature. (Based on Potter and Jones 1977)

A later chapter in this book, Chapter 14, explains the effect of temperature on growth of different plant species, with particular focus on adaptation to very low and very high temperatures.

6.2.3 - Carbon dioxide


Figure 6.4 Early growth of cucumber (Cucumis sativus, top panel) and wong bok (Brassica pekinensis, bottom panel) is greatly enhanced in elevated CO2 (1350 ppm) compared with ambient controls (330-350 ppm). As shown here, that initial effect is still apparent after 52 d of greenhouse culture in nutrient rich potting mix. Scale bar = 10 cm. (Further details in Kriedemann and Wong (1984) and Table 6.3) (Photographs courtesy M. Whittaker)

Growth responses to elevated CO2 can be spectacular, especially during early exponential growth (Figure 6.4) and derive largely from direct effects of increased CO2 partial pressure on photosynthesis. C3 plants will be most affected, and especially at high temperature where photorespiratory loss of carbon has the greatest impact on biomass accumulation.

Global atmospheric CO2 partial pressure is expected to reach 60–70 Pa (600–700 ppm) by about 2050 so that growth response to a CO2 doubling compared with 1990s levels has received wide attention. Instantaneous rates of CO2 assimilation by C3 leaves can increase two- to three-fold in response to such elevated levels of CO2, but the short-term response is rarely translated into biomass gain by whole plants where growth and reproductive development can be limited by low nutrients, low light, low temperature, physical restriction on root growth (especially pot experiments) or strength of sinks for photoassimilate. Given such constraints, photosynthetic acclimation commonly ensues. Rates of CO2 assimilation (leaf area basis) by CO2-enriched plants, grown and measured under high CO2, will match rates measured on control plants at normal ambient levels.


Figure 6.5. A survey of growth response to elevated CO2 (ratio of growth indices in 600-800 cf. 300-400 ppm CO2) in 63 different C3 species (a) and eight C4 species (b) reveals systematic differences in median values for growth indices that relate to photosynthetic mode. C3 plants show a positive response in NAR that results in slightly faster RGR despite some reduction in LAR. C4 plants reduce RGR under elevated CO2 due to diminished NAR. SLA of C3 plants is generally lower under elevated CO2, but increased somewhat in C4. LWR is essentially unchanged in either group (Based on Poorter et al. 1996)

Acclimation takes only days to set in, and because plant growth analysis commonly extends over a few weeks, CO2-driven responses in growth indices tend to be more conservative compared with instantaneous responses during leaf gas exchange. C4 plants will be less affected than C3 plants (see Chapter 2) so that broad surveys need to distinguish between photosynthetic mode. For example, in Figure 6.5, average NAR for 63 different cases of C3 plants increased by 25–30% under 600–800 ppm CO2 compared with corresponding values under 300–400 ppm CO2. However, NAR increase was not matched by a commensurate response in RGR, and decreased LAR appears responsible. CO2-enriched plants were less leafy than controls (i.e. lower LAR), but not because less dry matter was allocated to foliage (LWR was on average unaltered). Rather, specific leaf area (SLA in Figure 6.5) decreased under high CO2 so that a given mass of foliage was presenting a smaller assimilatory surface for light interception and gas exchange. Accumulation of non-structural carbohydrate (mainly starch; Wong 1990) is commonly responsible for lower SLA in these cases, and in addition generally correlates with down-regulation of leaf photosynthesis.

By contrast, in C4 plants LWR was little affected by elevated CO2, but in this case SLA did show slight increase with some positive response in LAR. However, photosynthetic acclimation may have been more telling because NAR eased and RGR even diminished somewhat under elevated CO2.

Global change, with attendant increase in atmospheric CO2 over coming decades, thus carries implications for growth and development in present-day genotypes and especially the comparative abundance of C3 and C4 plants. Elevated CO2 also has immediate relevance to greenhouse cropping. In production horticulture, both absolute yield and duration of cropping cycles are factors in profitability. Accordingly, CO2 effects on rate of growth as well as onset of subsequent development are of interest.

Young seedlings in their early exponential growth phase are typically most responsive to elevated CO2, so that production of leafy vegetables can be greatly enhanced. This response is widely exploited in northern hemisphere greenhouse culture. In commercial operations, ambient CO2is often raised three- to four-fold so that growth responses can be spectacular (Figure 6.4) but they tend to be short lived (Table 6.3), as accelerated early growth gives way to lower RGR. During each cycle of growth and development, annual plants show a sigmoidal increase in biomass where an initial exponential phase gives way to a linear phase, eventually approaching an asymptote as reproductive structures mature. If CO2 enrichment hastens this progression, a stage is soon reached where RGR is lower under elevated CO2 due to accelerated ontogeny.

The fall in RGR with time of exposure to high CO2 is illustrated for wong bok (Brassica pekinensis) in Table 6.3. Wong bok is a highly productive autumn and winter vegetable that serves as ‘spring greens’ and is especially responsive to CO2 during early growth. RGRA at c. 330 ppm CO2 was initially 0.230 d–1 compared with 0.960 d–1 at c. 1350 ppm CO2, but by 40–52 d, RGRA had fallen to 0.061 and 0.020 d–1 for control and CO2 enriched, respectively (Table 6.3). CO2-driven response in NAR and RGR also diminished with age, and especially where these larger individuals failed to sustain higher RGR past 18 d (Table 6.3). Nevertheless, a response in NAR was maintained for a further two intervals so that a CO2 effect on plant size was maintained (Figure 6.4).

Component analyses of RGR as exemplified above are also useful for analysing whole plant responses to the supply of various nutrients and to interactions between nutrient supply and light, temperature or CO2. There are also strong interactions between these ambient conditions and abiotic stresses such as drought and salinity that can be quantitatively analysed.

The following two sections consider the developmental stages of leaf, shoot and reproductive organ formation, and how this information contributes to different types of quantitative growth analyses.

6.3 - Vegetative growth and development

Growth is an irreversible increase in plant size accompanied by a quantitative change in biomass. Development is more subtle and implies an additional qualitative change in plant form or function. Development thus lends ‘direction’ to growth and can apply equally well to a progressive change in gross morphology as to a subtle change in organ function, or to a major phase change from vegetative to reproductive development.

Increases in leaf area over time can be a useful basis for measuring plant growth rates than biomass increases, particularly as non-destructive and automated techniques for measuring leaf area are now available. Plant growth rate can be assessed as the relative increase in leaf area over time, by substituting total plant leaf area for total biomass in the conventional RGR equation.

\[\text{RGR}_\text{A} =\frac{\text{ln } LA_2 - \text{ln } LA_1}{t_2-t_1} \tag{6.14}\]

where RGRA is relative leaf expansion rate, LA is total leaf area and t is time at two time intervals, t1 and t2, preferably 2-3 days apart. This can be done by image analysis. This information can be extrapolated to whole plant growth rates as leaves, stems and roots generally maintain a balance in biomass that can be described by an allometric relationship.

In the first part of this section, growth of individual leaves is described at the cellular level of organisation, how this is influenced by light, and how much the photosynthetic activity of leaves changes with development.

The second part shows how root:shoot ratios change with availability of resources and the third part how these change with ontogeny (allometry).

6.3.1 - Patterns of leaf growth


Figure 6.6. Leaf expansion in sunflower shows a sigmoidal increase in lamina area with time where rate of area increase and final size both vary with nodal position, reaching a maximum around node 20. The curves were drawn by hand through all data points (two measurements of leaf length (L) and leaf breadth (B) per week with area A estimated from the relationship A = 0.73 (L × B). Based on Rawson and Turner (1982) Aust J Plant Physiol 9, 449-460

Growth rate of individual leaves provides much useful information on plant growth, especially in response to changes in environment, as leaf growth can be measured over hours or even minutes. Rates of leaf elongation or individual leaf area expansion cannot be used to calculate whole plant relative growth rates, but they can be used to assess current rates of individual leaf growth and effects of a treatment on the rate of leaf emergence (“phyllocron”). Leaf elongation (increase in length of a given leaf per hour or per day) is a sensitive measure of leaf growth and can be accomplished electronically with a transducer, over minutes, manually with a ruler over 4-24 hours, or automatically with a digital photographic technique over intervals of days. Linear measurements with a transducer or ruler are particularly sensitive for monocots whose growth is largely one-dimensional.

(a) Measurement of leaf expansion

Differences in canopy development result from the frequency of new leaf initiation and the time-course of lamina expansion. These can be inferred from comprehensive measurement of lamina expansion on successive leaves. Lamina expansion in both monocotyledons and dicotyledons is approximately sigmoidal in time and asymmetric about a point of inflexion which coincides with maximum rate of area increase. However there is a period of several days over which expansion rates are constant.

A determinate plant with large leaves such as sunflower (Figure 6.6) provides a typical example. Leaf area is shown as a function of time for eight nodes selected between node 6 and node 40. Final leaf area was greatest at node 20, but daily rates of expansion were uniform for leaves between nodes 10 and 25. Thus at any time between days 35 to 65, the daily rate of expansion of any leaf was the same. Slowest growth and smallest final size was recorded for node 40, adjacent to the terminal inflorescence.

Growth curves for monocots leaves are very similar, in that there is a period of several days during which the leaf has a constant rate of area expansion. Increases in leaf areas of cereals and other monocots are easier to measure than dicots as they grow only in length and not width, for the reason explained below.  

Frequency of leaf initiation can be inferred from a more comprehensive family of such curves where early exponential growth in area for each successive leaf is recorded and plotted as log10 area versus time. This results in a near-parallel set of lines which intersect an arbitrary abscissa (Figure 6.7). Each time interval between successive points of intersection on this abscissa is a ‘phyllochron’ and denotes the time interval between comparable stages in the development of successive leaves. This index is easily inferred from the time elapsed between successive lines on a semi-log plot (Figure 6.7). Cumulative phyllochrons serve as an indicator of a plant’s physiological age in the same way as days after germination represent chronological age.


Figure 6.7. Leaves of subterranean clover achieve a 10-million-fold increase in size from primordium to final area (volume of primordia shown as dotted lines; leaf fresh mass shown as solid lines). Successive leaves are initiated and enlarge in a beautifully coordinated fashion revealed here as a family of straight lines on a semi-log plot. Intervals along an arbitrary abscissa (arrow at 100 × 10-3 mm3) that intersects theses lines represent time elapsed (about 1.8 d) between attainment of a given development status by successive leaves (phyllochron). Full-sized leaves exceed about 100 mm3 in volume. Based on Williams (1975) J Aust Inst Agric Sci 41, 18-26


Figure 6.8. Leaves of cucumber (node 2 on plants in growth cabinets) show an approximately sigmoidal increase in area with time (broken lines) where final size and cell number vary with daily irradiance (0.6, 1.9 or 4.4 MJ m-2 d-1). During an initial exponential phase in area growth, cell number per leaf (solid lines) also increases exponentially. The slope of a semi-log plot (hence relative rate of cell division) is higher under stronger irradiance. Cell number per leaf approaches asymptote as the rate of leaf area increase becomes linear.  Based on Milthorpe and Newton (1963) J Exp Bot 14, 483-495

(b) Developmental stages of leaf expansion

Leaves are first discernible as tiny primordia which are initiated by meristems in accord with a genetically programmed developmental morphology. They undergo extensive cycles of cell division (peak doubling time about 0.5 d). Leaf growth is anatomically different in grasses (monocotyledonous species) and broad-leafed (dicotyledonous) plants.

Primordia of broad-leafed plants undergo extensive cycles of cell division and enlargement to form recognisable leaves with petioles that elongate and lamina that unfold and expand. Lamina expansion follows a coordinated pattern of further cell division and cell enlargement that is under genetic control but modified by the environment, particularly light. Early growth of the leaf is driven primarily by cell division, and cell number per leaf increases exponentially prior to unfolding. Cell division can continue well into the expansion phase of leaf growth, so that up to 90% of cells in a mature dicot leaf can have originated after unfolding. Cell division finishes about the time the leaf enters its period of linear rate of area expansion, so this period of maximum leaf expansion rate is due to expansion of pre-formed cells.

Primordia of grasses and other monocotyledonous species are hidden from view. All phases of cell growth occur at the base of the leaves which are usually not exposed to the environment. Cell division is confined to basal meristems which give rise to files of cells and a linear zone of cell expansion and differentiation. The emerging blade therefore is composed of cells that are fully expanded, and the elongation of that leaf takes place by addition of fully expanded cells from below.


Figure 6.9. Area of individual leaves on cucumber (Cucumis sativus) responds to daily irradiance and reaches a maximum above about 2.5 MJ m-2 d-1. Area increase (node 2 in this example) is due to greater cell number under stronger irradiance. Mean size of mesophyll cells is little affected and has no influence on area of individual leaves. Based on Newton (1963) J Exp Bot 14, 458-482

(c) Effects of light on leaf development

Light is the main variable affecting leaf growth rate, both the rate of leaf area expansion, final size, as well as cell shape as mentioned in the previous section.

Figure 6.8 shows the effect of light level on the rate of leaf area expansion in a cucumber leaf. As in all dicot leaves, the rate of lamina expansion is determined largely by the number of cells produced, with final cell area being unaffected (Figure 6.9). Rate of cell division during this early phase is increased by irradiance, so that potential size of these cucumber leaves at maturity is also enhanced. The upper curves in Figure 6.8 (highest irradiance), cell number per lamina reaches a plateau around 20 d, but area continues to increase to at least 30 d. Expansion of existing cells is largely responsible for lamina expansion between 20 and 30 d after sowing.

Figure 6.9 shows the effect of a range of light levels on final leaf area, and shows that area is strongly dependent on light level up to 2 MJ m-2 d-1, and that the increased area has been achieved by more cells rather than larger cells.

A similar light response curve would be shown by monocot leaves, and with similar contributions from cell number versus cell size. The difference between monocots and dicots is that the cell number is determined in the basal meristematic zone, before the lamina emerges. This zone is not exposed to the light environment, so cell division activity in monocots is controlled by substrates or signals arising in the older expanded leaves.

 (d) Leaf development and photosynthesis

When dicotyledonous leaves are very young and first unfold they have low rates of net photosynthesis (expressed per unit area) so have to import carbon from other leaves to support their growth. But as they expand their rates increase rapidly such that within a few days they can assimilate all their own carbon requirements and export excess (Figure 6.10).


Figure 6.10. Change in net photosynthesis rate as a cotton leaf unfolds, expands, reaches maximum area and ages. An initial phase of carbon import helps sustain early expansion but by the time the leaf is 70% of its final area it is self sufficient for carbon and exporting excess. From Constable and Rawson (1980) Aust J Plant Physiol 7, 89-100 and 539-553

In the example for cotton in the figure, this self-sufficiency occurs when the leaf is about 70% of its final area. Typically, net photosynthesis rate will reach a maximum before the leaf has fully expanded though this can range from 25 to 100% of final area across species. Photosynthesis rate will then remain at that maximum or start to decline with further leaf expansion before leaf aging, lessening requirement for the carbon produced, and environmental factors accelerate the decline. Because the amount of carbon produced by a leaf is the product of two largely independent variables, its photosynthesis rate x its area, leaf carbon production can continue to increase while photosynthesis rate is stable or even declining.

Monocotyledonous leaves grow from their base where the very young expanding parts of the leaf are fully enclosed inside a sheath created by the surrounding leaf bases. The emerged parts of the leaf blade are already approaching full expansion as they emerge from the sheath and unroll. Photosynthesis rates of those exposed parts are already close to their maximum. Once the whole blade is exposed, photosynthesis rate and leaf carbon production follow plateau and declining patterns similar to those described for dicots though magnitude and duration differ amongst species and environments.

When doing experiments that investigate the effects of environmental treatments on photosynthesis, it is important bear in mind the continuous progression in photosynthesis rate between growing, recently fully expanded and aging leaves. Leaves should be compared that are of equivalent age and stage of development, particularly if single leaves are being measured to represent a whole plant or a breeding line. If the eventual aim of the experiment is to compare or select for carbon production, the area of the leaf must also be known since photosynthesis rate and fully expanded area of a leaf are not linked. Leaves of some dicotyledonous species take a few days to reach full expansion while others take weeks.

6.3.2 - Root:shoot ratios

Roots, stems and leaves are functionally interdependent and these three systems maintain a dynamic balance in biomass which reflects relative abundance of above-ground resources (light and CO2) compared with root-zone resources (water and nutrients) (Poorter et al. 2012). Whole-plant growth rate and summary measures such as root:shoot ratio are thus an outcome of developmental stage and of environmental influences.

Change in root:shoot ratio during a plant’s life cycle is part of an intrinsic ontogeny, but growth rates of roots and shoots continually adjust to resource availability with photoassimilate (hence biomass). In herbaceous plants, root:shoot ratios typically decrease with age (size) due to sustained investment of carbon in above-ground structures (root crops would be a notable exception). Developmental morphology is inherent, but expression of a given genotype will vary in response to growing conditions (hence phenotypic plasticity).

Irradiance is a case in point where shoot growth takes priority in low light, whereas root growth can be favoured under strong light. For example, Evans and Hughes (1961) grew Impatiens parviflora at five light levels and demonstrated a steady increase in root mass relative to whole-plant mass (root mass ratio) from 7% to 100% full sun. Stem mass ratio showed the opposite sequence. Leaf mass ratio increased somewhat at low light, but increased SLA was far more important for maintenance of whole-plant RGR in this shade-adapted species.

If light effects on root:shoot ratio are translated via photosynthesis, then CO2 should interact with irradiance on root:shoot ratio because carbon assimilation would be maintained by a more modest investment in shoots exposed to elevated CO2Chrysanthemum morifolium behaved this way for Hughes and Cockshull (1971), returning a higher NAR due to CO2 enrichment under growth cabinet conditions despite lower LAR which was in turn due to smaller leaf weight ratio. Adjustment in SLA exceeded that of leaf weight ratio, and so carried more significance for growth responses to irradiance × CO2.

Consistent with shoot response to above-ground conditions, root biomass is influenced by below-ground conditions where low availability of either water or nutrients commonly leads to greater root:shoot ratio. For example, white clover (Trifolium repens) growing on a phosphorus-rich medium increased root:shoot ratio from 0.39 to 0.47 in response to moisture stress; and from 0.31 to 0.52 when moisture stress was imposed in combination with lower phosphorus (see Table 1 in Davidson 1969b). A positive interaction between low phosphorus and low water on root:shoot ratio was also evident in perennial ryegrass (Lolium perenne) grown on high nitrogen. In that case, root:shoot ratio increased from 0.82 to 3.44 in response to moisture stress when plants were grown on low phosphorus in combination with high nitrogen.

Adding to this nutrient × drought interaction, a genotype × phosphorus effect on root:shoot ratio has been demonstrated by Chapin et al. (1989) for wild and cultivated species of Hordeum. Weedy barleygrass (H. leporinum and H. glaucum) was especially responsive, root : shoot ratio increasing from about 0.75 to 1.5 over 21 d on low phosphorus. By contrast, cultivated barley (H. vulgare) remained between 0.5 and 0.75 over this same period. Held on high phosphorus, all species expressed comparable root:shoot ratios which declined from around 0.55 to about 0.35 over 21 d. High root:shoot ratios on low phosphorus in weedy accessions would have conferred a selective advantage for whole-plant growth under those conditions, thus contributing to their success as weeds.

Even stronger responses to phosphorus nutrition have been reported for soybean (Fredeen et al. 1989) where plants on low phosphorus (10 µM KH2PO4) invested biomass almost equally between roots and shoots, whereas plants on high phosphorus (200 µM KH2PO4) invested almost five times more biomass in shoots than in roots (daily irradiance was about 30 mol quanta m–2 d–1 and would have been conducive to rapid growth).

Root:shoot ratios are thus indicative of plant response to growing conditions, but ratios are not a definitive measure because values change as plants grow. Trees in a plantation forest would show a progressive reduction in root:shoot ratio, and especially after canopy closure where a steady increase in stem biomass contrasts with biomass turnover of canopy and roots and thus predominates in determining root:shoot ratio.

Broad generalisations are that root:shoot ratio increases with nutrient deficiency and moisture stress or under elevated CO2, but decreases in strong light. Too often, however, reports of treatment effects on root:shoot ratio have can overlooked differences in developmental ontogeny or size, and real responses may be obscured. Allometry then becomes a preferred alternative where repeated measurements of size or mass provide an unambiguous picture of carbon allocation.

6.3.3 - Allometry

During whole-plant growth in a stable environment, roots and shoots maintain a dynamic balance such that 

\[y=bx^k \tag{6.17}\]

Where y is root biomass and x is shoot biomass. More generally, x and y can be any two parts of the same organism that are growing differentially with respect to each other, but root–shoot relations are the most common candidate in such analyses of plant growth.

The allometric equation \(y=bx^k\)  (Equation 6.17) was formalised by Huxley (1924) and can be ln transformed to become

\[\text{ln } y= \text{ln }b + k\text{ln }x \tag{6.18}\]