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

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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