10.1.2 Cellular integration

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Plant cells possess diverse organelles and other structures, all of which need to cooperate to ensure efficient functioning. Specific subcellular structures, collectively known as the cytoskeleton, control the positioning and repositioning of cellular components during the cell cycle, and later on during cell differentiation then finally in functioning of mature cells. In addition to control within each single cell, multicellular organisms require coordination between cells. Cytoplasmic bridges known as plasmodesmata allow local traffic of many molecules including some kinds of regulatory signals. Here, we provide some insight into these intracellular and inter-cellular processes.

(a) The cytoskeleton — intracellular organisation

The cytoskeleton in plant cells is a remarkable multifunctional internal network of two types of fibrous protein polymers:actin filaments (often referred to as microfilaments) and microtubules. Both polymers form dynamic arrays which organise cellular components and provide the structural frameworks necessary to perform many key subcellular events. In animal cells, actin filaments and microtubules are essential for cell migration and changes in cell shape, two central components of cell and tissue morphogenesis. In contrast, a rigid cellulose wall surrounds plant cells and restricts their movements relative to neighbouring cells. Plants achieve the development of form, or morphogenesis, largely through two other cellular processes, both under control of the cytoskeleton: the placement and orientation of new cell walls resulting from cell division, and control of the subsequent directions in which cells expand.

Cytoskeleton components: molecular fibres and intracellular motors

Actin filaments are polymers of the 43 kDa monomeric protein actin (or G-actin, for globular). The polymer, called F-actin (for filamentous), is composed of two helically arranged parallel strands with an average diameter of 7 nm. One turn of the filament helix occurs every 37 nm, involving about 13 monomers on each strand (Figure 10.9). Microtubules are polymers of tubulin heterodimers composed of one a- and one b-tubulin polypeptide, each with a molecular weight of approximately 50 kDa. The heterodimers align head to tail into protofilaments, 13 of which assemble side by side to form a hollow microtubule of 24 nm diameter (Figure 10.9). Actin filaments and microtubules both assemble and disassemble via exchange with cytoplasmic pools of free monomer, processes which require energy. G-actin binds to ATP and its incorporation into a growing actin filament is followed by ATP hydrolysis. Tubulin monomers instead bind to GTP (guanosine triphosphate). After incorporation into the growing polymer, hydrolysis of the bound nucleotide can cause the polymer to be destabilised, often resulting in rapid uncontrolled depolymerisation known as dynamic instability. Actin and tubulin monomers are both polarised molecules, and by adding to the growing polymer in specific orien-tations, they impose a defined directionality to the actin filament or microtubule. Monomer association and disas-sociation rates differ between the two ends of each polymer, manifested as dynamic growth and disassembly. In vitro experiments have achieved polymerisation of both actin filaments and microtubules from their constituent monomers. Within eukaryotic cells, hundreds of proteins have been identified which interact with these cytoskeletal polymers to regulate their assembly and disassembly, and their interaction with other cellular components. Animal cells and protists have provided most of the identifications of these actin-binding proteins (ABPs) and microtubule-associated proteins (MAPs), but we do know that plants possess some equivalent proteins. One important class of ABPs and MAPs is the so-called motor proteins. These proteins ‘walk’ along cytoskeletal polymers via cycles of attachment followed by energy-dependent con-formational changes. The conformational changes define the direction of movement of each motor protein along the polymer, and different classes of motor proteins relate to each direction of movement along either actin filaments (myosins) or microtubules (kinesins and dyneins). Motor proteins attach to the surface of different organelles and possibly the plasma membrane, and through their specific interaction with actin filaments or microtubules, provide the molecular driving force to move and position these organelles within the cell.


Figure 10.9 Structure of an actin filament (F-actin) and a microtubule. An actin filament is composed of G-actin subunits in parallel strands wound helically to form a filament with a diameter of 7.5 nm, with one complete turn every 37 nm. ATP-bound G-actin monomer is rapidly added to or removed from the plus end, but this occurs only slowly from the minus end (not shown). A microtubule is composed of heterodimers of α- and β-tublin polypeptides assembled with defined orientation into linear protofilaments, 13 of which align side by side to form the hollow microtubule with a diameter of about 24 nm. Assembyl of GTP-bound heterodimers is most rapid at the plus end, and depolymerisation of GDP-bound heterodimers is most rapid from the minus end (not shown).

(Reproduced from Gunning and Steer (1996), Plant and Cell Biology and Function, with permission of Jones and Bartlett Publishers)

Structure of cytoskeletal arrays — microtubules

Microtubules are multifunctional units which support cell growth and the cell cycle. The dynamic property of micro-tubules enables the transient formation of several distinct arrays as a cell progresses through different stages of the cell cycle and differentiation. Amazingly, the same microtubule molecules are recruited into four functions, which occur sequentially during the mitotic cell cycle. These appear as the interphase cortical array, preprophase band, mitotic spindle and phragmoplast (Figure 10.10). With the exception of the mitotic spindle, each array is unique to plants and each performs discrete functions central to plant cell morphogenesis.


Figure 10.10 Confocal laser scanning micrographs of dividing cells from wheat root tips showing microtubules labelled with anti-tubulin antibody (bright regions), and DNA stained with flourescent dye (darker regions) (see Colour Plate 40). (A) Interphase array of roughly parallel cortical microtubules arranged transversely across three visible faces of this cell, but more randomly oriented on the end wall. (B) Preprophase band a narrow band of tightly packed microtubules replacing the interphase array. (C) Metaphase spindle – bundles of microtubules aligned along the spindle axis attach to the chromosome kinetocores resulting in a configuration known as the metaphase plate. This is seen as a dark-blue region separating the two halves of the spindle (D) Early phragmoplast – after chromosome separation during anaphase, a new population of microtubules appears between the chromosome masses and the mid-zone. (E) Late phragmoplast – a tightly packed double ring of short microtubules viewed side on, separated by the cell plate (seen as a dark line). (F) Cytokenesis and new cell wall formation nearly complete – vestiges of the phragmoplast ring remain at the extreme margins and new microtubules are emanating from the surface of the two new nuclear envelopes. (G) Reinstated cortical arrays in the two interphase daughter cells.

(Reproduced from Gunning and Steer (1996), Plant and Cell Biology and Function, with permission of Jones and Bartlett Publishers)

Interphase cortical array During interphase, microtubules lying parallel to each other form an array immediately adjacent to the plasma membrane and usually are oriented perpendicular to the long axis of the cell (Figure 10.10A). Individual micro-tubules are relatively short, but by overlapping in roughly parallel alignment they create a continuous array which passes over four faces of the cell. The end faces of these cells instead contain fewer and more randomly organised microtubules. Some microtubules in these arrays may be physically cross-linked to each other and/or to the plasma membrane via as yet unidentified MAPs. Typically, the most recently deposited (innermost) layers of cellulose microfibrils in the adjacent cell wall are aligned parallel to these microtubules as discussed later.

Preprophase band  As the cell enters mitosis, the interphase array of microtubules is replaced swiftly by a narrow band of microtubules that encircles the nucleus (Figure 10.10B). This structure is termed the preprophase band and has a remarkable feature in that, even before the cell enters mitosis, it accurately predicts the site where the new cell wall will be deposited after cytokinesis. The molecular mechanisms determining this prediction are not understood, but the band may somehow chemically mark the future division site, perhaps by phosphorylation. The band itself disappears before metaphase.

Spindle microtubules  Disintegration of the preprophase band coincides with formation of the mitotic spindle (Figure 10.10C). These microtubules first emerge from broad, ill-defined polar regions of the nucleus, then progressively consolidate along the pole-to-pole axis of the spindle, finally invading the nucleus after nuclear envelope breakdown and attaching via kinetochores to the condensing chromosomes. All eukaryotic cells depend on the spindle system to maintain accurate inheritance of the entire genome at each mitosis. The spindle microtubules have responsibility first for organising chromo-somes during prometaphase, then physically segregating the two daughter chromatid arms to each pole of the spindle during anaphase.

Phragmoplast microtubules  After chromosome separation, the spindle microtubules disappear and instead numerous short microtubules aggregate in the equatorial zone of the spindle (Figure 10.10D). These microtubules form the phragmoplast, a structure consisting of a double ring of short, antiparallel microtubules which form a narrow overlapping region at the spindle equator (Figure 10.10E). The two sets of microtubule arrays deliver Golgi-derived vesicles containing cell wall precursor materials to the developing cell plate, most likely by a mechanism involving microtubule motor proteins attached to the vesicles. The expanding cell plate eventually fuses with the parent cell wall at the site previously marked by the preprophase band, thus forming two daughter cells (Figure 10.10F). The daughter cells establish new cortical arrays of microtubules which persist through interphase until each cell commits to a new round of mitotic division (Figure 10.10G).

Structure of cytoskeletal arrays — actin filaments

Actin filaments also form diverse and dynamic arrays in plant cells (Figure 10.11). They are readily detected as large bundles usually aligned longitudinally to the growth axis of the cell. Smaller bundles ramify throughout all parts of the cytoplasm, but often are prominent as transverse arrays (McCurdy et al. 1989). Because actin filaments often appear closely aligned with cortical microtubules under the electron microscope, we deduce there may be functional associations. In some cell types, especially highly vacuolate cells maintained as suspension cultures, the actin cytoskeleton consists of an interconnected and complex network of cortical filaments, larger transvacuolar cables and meshworks surrounding the nucleus (Lloyd and Traas 1988). A major role of the actin cytoskeleton is to generate protoplasmic streaming, which operates through myosin motor proteins with attached organelles moving along the filaments and bundles. This provides efficient mixing of structural and chemical components of cells, a process necessary to maintain metabolism in large plant cells. Protoplasmic streaming is visible in many living cells under the light micro-scope. In growing pollen tubes, spectacular two-way traffic over millimetre distances can be followed as organelles and metabolites are ceaselessly supplied to and from the sole growing point, the pollen tube tip.


Figure 10.11 Confocal laser scanning micrograph of actin filament bundles labelled with flourescent anti-actin antibodies. This is a longitudinal section through the zone of cell elongation of a wheat root tip, and shows bundles of actin running in a net longitudinal orientation throughout the cell, often passing close to the nucleus (darker oval shapes devoid of labelling). The borders of two individual cells are shown with dashed lines.

(Reproduced from Gunning and Steer (1996), Plant and Cell Biology and Function, with permission of Jones and Bartlett Publishers)

The cytoskeleton and morphogenesis

The development of cell shape, and in turn that of organs and tissues, is the result of both division plane determination and control of the direction of cell expansion. Because microtubules are relatively stable structures, and thus more easily detected by electron microscopy and immunofluorescence localisation techniques, their role in plant morphogenesis has been more readily appreciated than that of actin. However, we now know that actin filaments also participate in these processes, probably by influencing microtubule rearrangements.

Determination of the plane of division  Plant cells divide by constructing a new internal wall after mitosis has been completed. The placement of this new wall coincides precisely with the position of the preprophase band of microtubules that forms transiently before mitosis begins. The mechanisms determining where the preprophase band will form are not understood, nor is it clear how this structure acts as a predictor of subsequent cell division (Williamson 1991). Presence of the band is clearly required, since disruption by drugs such as colchicine or physical wounding alters the subsequent positioning of the new wall (Gunning and Wick 1985). The presence of the preprophase band may somehow condition or ‘tag’ the division site by chemically modifying the adjacent cell wall or cytoplasmic region with marker molecules (Mineyuki and Gunning 1990). Protein kinases are found localised to the preprophase band, suggesting phosphorylation of specific cellular components (Mineyuki et al. 1991). Once a division site has been determined, the production of a new cell wall involves both actin filaments and microtubules. The phragmoplast constructs the new wall by delivering precursor vesicles to the expanding cell plate. These vesicles move along microtubules, most likely using motor proteins such as kinesin (Liu et al. 1996), to the mid-region of the dividing cell where they fuse with the cell plate, thus enabling its centrifugal growth (i.e. centre outwards). In addition, actin filaments and myosin within the phragmoplast may contribute to vesicle migration and organisation. As the flattened disc-shaped cell plate expands, its leading edge is guided to and eventually fuses with the parent cell wall precisely at the site formerly occupied by the preprophase band. In large vacuolate cells where the mitotic spindle and phragmoplast are in a central raft of transvacuolar cytoplasm known as the phragmosome, the expanding cell plate must traverse a substantial distance before reaching the side wall. In such cells, actin filaments are visible connecting cell plate to parent wall along the phragmosome (Figure 10.12; Lloyd and Traas 1988), and presumably act as a guidance mechanism for the growing cell plate, since treatment of dividing cells with anti-actin drugs causes misaligned phragmoplasts and cell plates (Palevitz 1980; Lloyd and Traas 1988).

Direction of cell expansion  The final shape of a plant cell is dictated by the direction(s) of turgor-driven cell expansion. This direction is controlled by physical constraints imposed by cellulose microfibrils laid down in the growing cell wall (Green 1980). The innermost layer of microfibrils in the cell wall usually lies perpendicular to the axis of elongation, and these microfibrils act as hoop reinforcement elements to restrict lateral expansion of the cell while allowing elongation. The cytoskeleton is intimately linked to this process because the ordered deposition of cellulose microfibrils is tightly controlled by the orientation of cortical microtubules (Section 4.3.2).


Figure 10.12 Flourescence micrographs of the actin cytokinesis in carrot suspension cells. These cells are highly vacuolated and the phragmoplast forms within a complex meshwork of actin filaments. Actin filaments were visualised by labelling with a flourscent derivative of phallodin, a fungal compound which binds specifically to F-actin. (A) Intense flourescence of phragmoplast-associated actin in the middle of the cell (the two chromosome masses are indicated by double dots), together with pole-associated actin strands (arrows). (B) Phragmoplast suspended by the actin cytoskeleton in the transvacuolar phragmosome. The darker regions are the vacuole. Scale bar = 10 μm.

(Reproduced from Lloyd and Traas (1988), with permission of Company of Biologists Ltd)

Electron microscopy first revealed the co-alignment in growing cells of cortical microtubules and the inner-most layer of cellulose microfibrils in the cell wall (Ledbetter and Porter 1963). Control of microfibril orientation was revealed by the use of microtubule-destabilising drugs such as colchicine, which also disrupted the oriented deposition of the neighbouring cell wall microfibrils, with subsequent changes in cell shape (Brower and Hepler 1976; Hogetsu and Shibaoka 1978). Orderly deposition of microfibrils only returned after the colchicine had been washed away to allowed re-establish-ment of microtubule arrays, thus confirming the dependence of microfibril orientation on that of the cortical microtubules. Cellulose microfibrils are deposited on the outside of the plasma membrane by a membrane-bound enzyme complex called cellulose synthase. In higher plants, this complex appears as a rosette assembly of protein aggregates embedded within and spanning the plasma membrane (Figure 10.13). The complex is believed to move within the plane of the membrane, assembling a cellulose microfibril on the outer surface of the cell by incorporating precursor subunits (UDP-glucose) into the complex from the cytoplasmic side of the membrane. Forces resulting from continued polymerisation and distal crystallisation of the microfibrils drive the complexes through the plane of the plasma membrane. The direction of this movement, and thus the orientation of the cellulose micro-fibrils, is guided by the underlying cortical microtubules (Giddings and Staehelin 1991). The mechanism may involve protein cross-bridges between microtubules and plasma membrane, or perhaps between microtubules and synthase complexes themselves. Restriction of synthase complexes to defined membrane channels results in microfibril orientations reflecting that of the underlying cortical microtubules (Figure 10.13C).


Figure 10.13 Electron micrographs of freeze-etched and rotary shadowed cellulose microfibrils and celluolose synthase rosettes in differentiating Zinnia elegans tracheary elements. (A) Cellulose microfibrils in the secondary cell wall overlie the plasma membrane, a small patch of which is exposed on the right. Scale bar = 100 nm. (B) Rosette structure of putative cellulose synthase complexes in the plasma membrane underlying the cellulose-rich secondary wall thickenings in these cells. The rosette structure is composed of six globular units. Scale bar = 30 nm. (C) Diagram suggesting the spatial relationship between cortical microtubules and the most recently deposited layer of cellulose microfibrils. Cellulose microfibrils emerge from rosettes of cellulose synthesising complexes embedded in the plasma membrane (part of the lipid bilayer is peeled back to reveal the different faces of the membrane visualised in freeze-fracture preparation). The microtubules underlying the plasma membrane are shown cross-linked to each other or to the plasma membrane itself. The latter connections may define membrane channels which in turn restrict movement of cellulose synthase rosettes through the plane of the membrane in a direction reflecting the orientation of the underlying microtubules.

((A),(B) Images produced by C.H. Haigler and M.J. Grimson, Texas Tech University, and reproduced from Proceedings of the National Academy of Science, USA, 93, 12082-12085. © 1996, National Academy of Science USA, (C) reproduced with modifications from Gunning and Steer (1996), Plant Cell Biology, Structure and Function, with permission of Jones and Bartlett Publishers)

The relationship between cortical microtubules, cellulose deposition and cell shape is evident in certain differentiating cells such as developing tracheary elements or leaf mesophyll cells. Differentiated mesophyll cells in wheat have a highly lobed outline with regularly spaced constrictions. The devel-op-ment of this lobed morphology results initially from the bunching of interphase microtubules into discrete bands, often as many as five or six per cell (Figure 10.14) (Wernicke and Jung 1990). These localised ‘hoops’ of wall reinforcement laid down during the early stages of cell expansion act as restrictions to the expanding cell, resulting in the cell wall between these restrictions bulging out and thus producing the lobed morphology (Figure 10.14). Again, disruption of micro-tubules by inhibitors results in loss of the banded secondary wall thickenings. Actin also seems to be involved: disrupting actin filaments with cytochalasin B prevents cortical micro-tubules from rearranging into bands, thus preventing ultimately the localised deposition of secondary wall material (Wernicke and Jung 1992).


Figure 10.14 Developing mesophyll cells isolated from wheat leaf and double stained to visualise microtubules (A, C, E) and cellulose microfibrils of the cell wall (B, D, F), (A), (B) Young expanding cells 2-5 mm from the leaf base, showing typical transverse arrays of microtubules (A), and cellulose microfibrils with the same orientation (B). (C), (D) Between 5 and 10 mm from the the leaf base, cell shape has not changed markedly but cortical microtubules (C) have aggregated into discrete transverse bands, matched again by the pattern of cellulose deposition (D). (E), (F) Between 20 and 50 mm from the leaf base; the first indications of the lobed shape of mesophyll cells are seen. The microtubule bands have dispersed somewhat and fan out (E), but hoops of cellulose wall reinforcement are still evident (F), and clearly mark the isthmi where wall expansion has been restricted to create a highly lobed cell shape. Scale bar = 20 μm.

(Reproduced in part from Wernicke and Jung (1990), Protoplasma, 153, 141-148, with permission of Springer-Verlag)

The plant cytoskeleton responds to environmental factors

The cytoskeleton can respond to numerous environmental stimuli in complex and subtle ways. Reorganisation of cytoskeletal elements has been observed in response to light, gravity, hormonal signals, fungal attack and wounding. These changes can occur rapidly — minutes to hours — and often result in the reorganisation or repositioning of organelles. One example involves actin-controlled movement of chloroplasts in Selaginella leaf cells in response to varying light intensities (Cox et al. 1987; Dong et al. 1996). In low light conditions, chloroplasts are dispersed as a monolayer across the face of the cell that is perpendicular to the direction of incident illumi-nation. This array maximises light capture. Under potentially damaging high light intensities, however, chloroplasts aggregate to the side walls which are parallel to the incident light, thus minimising light interception.


Figure 10.15 Immunoflourescence images of epidermal cells from epicotyls of dwarf pea seedlings. Sections were labelled with anti-tubulin to visualise arrays of cortical microtubules. (A) Cells from control seedlings showing net longitudinal orientation of the microtubule arrays. (B) Cells from seedlings treated with gibberellic acid for 12 h showing coritcal microtubules orientated in a net transverse direction. This shift in microtubule orientation is mirrored int the orientation of new cellulose microfibrils in the cell wall, and consequently cell expansion becomes predominately longitudinal, thus promoting elongation growth of the epicotyl and restoration of a tall phenotype. Scale bar = 10 μm.

(Reproduced from Sakiyama and Shibaoka (1990) with permission of Springer-Verlag

Microtubules also mediate growth responses to plant hormones via their regulation of oriented cellulose microfibril deposition in the cell wall. Gibberellins and ethylene can affect orientation of cortical microtubules, and by doing so have profound effects on plant morphology. In gibberellin-deficient dwarf pea seedlings, cortical microtubules are abnormal by being oriented mainly longitudinally in epidermal cells of the epicotyl, thus restricting elongation of this organ. Application of gibberellin promotes a net transverse orientation of cortical microtubules (Figure 10.15; Sakiyama and Shibaoka 1990), which in turn leads to predominantly transverse cellulose microfibrils and hence promotes elongation growth resulting in longer, thin shoots. Similarly, ethylene inhibits stem elongation and stimulates stem thickening through reor-ganisation of cortical microtubules from transverse to net longitudinal orientations (Lang et al. 1982). In summary, changes in microtubule organisation, and subsequent effects on cellulose microfibril orientation, mediate several responses to hormonal signals that modify a plant’s growth habit.

(b) Plasmodesmata: intercellular communication


Figure 10.16 Plasmodesmata: intecellular cytoplasmic bridges. (a) Pit fields in the thick lignified walls of sclerenchyma cells, in this case from persimmon (Diospyros) endosperm, contain plasmodesmata which are visible under the light microscope as numerous file lines connecting cell to cell. Transmission electron micrographs of non-lignified cell walls from corn (Zeamays) leaf show longitudinal (b) and transverse (c) plasmodesmata profiles. The central strand of endoplasmic reticulum, the desmotubule, and surrounding plasma membrane are visible. Scale bar in (a) = 20 μm; in (b) = 0.2 μm; in (c) =0.2 μm.

(Based on Raven et al. 1992)

Integration of many cell functions, for example the co-ordinated growth of groups of cells within tissues, requires constant communication between cells. Some regulatory signals, such as plant hormones, are small molecules that can travel through the apoplasmic space in cell walls and may enter target cells by crossing the plasma membrane. Other signals travel only through the cytoplasm and move between cells via plasmodesmata, which are minute cytoplasmic channels traversing the wall between adjacent cells.

Tangl in 1879 first described plasmodesmata when pit fields were seen in the light microscope as fine strands in the walls of endosperm cells (Gunning and Robards 1976). Aggregations of plasmodesmata in pit fields are most apparent in very thick lignified walls (Figure 10.16a). However, their role in intercellular transport was not appreciated until the 1960s when electron microscopy revealed their fine ultra-structural details. Some early ideas included the proposal that plasmodesmata represented pieces of endoplasmic reticulum accidentally trapped in new cell walls during cytokinesis. Wider searches soon indicated, however, that almost all walls between adjacent plant cells contained these structures (Figure 10.16b, c).

In 1983, Gunning and Overall (see Feature essay 10.1) proposed a model for plasmodesmata that resolved earlier alternatives. Revising the model of López-Sáez et al. (1966), they showed that a narrow tube of endoplasmic reticulum — the desmotubule — passes through the middle of the plasmodesmal channel thereby connecting the endomembrane networks of adjacent cells (Figure 10.17a, d). They also supported the view of Tucker (1982) and others who suggested that the cyto-plasmic sleeve between the endoplasmic reticulum and the plasma membrane lining the plasmodesma could be a pathway for transport of cytoplasmic molecules. Under high-magnification electron microscopy, the cytoplasmic sleeve often appears partly occluded by particulate material (Figure 10.17b). This could be interpreted either as negatively stained particles in an electron-dense matrix or as positively stained spokes (Figure 10.17c) linking desmotubule to plasma membrane. Although cross-sections sometimes support the first view, the latter is supported by Ding et al. (1992) whose model is shown in Figure 10.17e. In neither case do we know the composition of these structures, nor yet of any other plasmodesmata-specific proteins.

Regulated transport through plasmodesmata enables coordination of like activities within a tissue, yet provides sufficient isolation from different adjoining cell types that can carry out specialised functions or enter alternative differ-entiation pathways. We can deduce that the size of the cytoplasmic channel is regulated, but because of the dimensions involved, even electron microscope sections cannot resolve the possible open and closed states. Electron microscope images do, however, sometimes show a ring of particles in the wall around the neck of plasmodesmata. Some have termed this a ‘sphincter’ apparatus because the structure is well positioned to squeeze the cytoplasmic channel closed or to relax and allow it to open (Olesen 1979). In addition, callose, a b-1,3 glucan, is found at the neck, especially after tissue wounding. This may be part of normal wound responses that seal off neighbouring cells and reduce leakage of cytoplasm. It is also possible that callose participates in normal regulation of plasmodesmal aperture (Robards and Lucas 1990). Cytochemistry shows that several enzymes, including ATPases, are abundant in the vicinity of plasmodesmata. The cytoskeleton protein actin is also detectable in plasmodesmata, from which we infer that either transport through the channel is actino-myosin based, or a contractile mechanism is responsible for regulating permeability (White et al. 1994).

Two different methods help us monitor transport through plasmodesmata. One involves watching membrane-impermeant dyes moving from cell to cell, and suggests that only molecules below 800–900 Da, with a size of 2 nm or less, are able to diffuse freely through plasmodesmata (Figure 10.18). This corresponds well with the measured 2–3 nm diameter of the small channels between particles in the cytoplasmic sleeve. Later on, we will see that despite this size limit, selective movement of much larger molecules is possible. The second method measures transmission of applied electrical pulses from cell to cell. Much more current passes between cells than is predicted if the current travelled across the high-resistance plasma membranes. We deduce that plasmodesmata act as a low-resistance pathway allowing diffusion of ions and other small molecules. We also know that calcium ions cause the channels to close briefly, but an induced pH shift has no such effect (Reid and Overall 1992). Induced osmotic pressure gradients between cells can also result in temporary closure. However, natural osmotic pressure differences, such as those between companion cell and sieve tube, do not induce closure. Indeed, the pressure flow mechanism in phloem depends on bulk flow between these cells.

There are also longer term changes in plasmodesmal permeability. For example, in the shoot apex of Silene, the molecular exclusion limit reduces from 656 Da to 536 Da following floral induction (Goodwin and Lyndon 1983). This size range is larger than most plant hormones (150–400 Da) but may partially isolate the floral apex from other mobile signals (e.g. short peptides or oligosaccharides) from surrounding cells and perhaps facilitates determination of the floral state.

The importance of plasmodesmata in maintaining coordination between adjacent cells is illustrated in leaf epidermis. Here, stomatal guard cells initially have plasmodesmal connections with surrounding subsidiary cells, but as they mature these bridges are severed. Isolation is probably essential for normal guard cell function which depends on rapid turgor, solute and volume changes. Temporary plasmolysis is a tool which breaks plasmodesmal links. In germinating spores of the fern Pteris this artificial symplasmic isolation causes each cell of the protonemal filament to form a small thallus. The normal pattern is for the apical cells to generate a single thallus, so we deduce that plasmodesmal signals usually restrict some cells from certain morphogenic pathways (Gunning and Robards 1976).


Figure 10.17 Models of plasmosdesmata structure and operation have been derived from high-magnification electron microscope images in longitudinal (a) and transverse (b, c) from Azolla pinnata root and adjascent phloem parenchyma cells in sugar cane leaf. PM = plasma membrane; ER = endoplasmic reticulum. In (b), there is a mottled layer of intervening cytoplasmic annulus between the central desmotublule and surrounding plasma membrane. In (c), the desmotubule is connected to the plasma membrane by 'spokes' that extend across the very open cytoplasmic annulus. In the Gunning and Overall (1983) model (d), most of the cytoplasmic channel between desmotubule and plasma membrane sleeve is occupied by globular proteins, whereas in Ding's model (e) spokes connect desmotubule to the plasma membrane, best seen in plasmodesmata with expanded regions in the centre of the wall. Both interpretations rely on the dark-light-dark anternation of staining in the cytoplasmic channel. Scale bars: (a) 100 nm; (b) 10 nm; (c) 100 nm.

(Based on (a), (b) Overall and Gunning 1982, (c) Robinson-Beers and Evert 1991, (d) Gunning and Overall 1983 and (e) Ding et al. 1992; (c) and (e) reproduced with perssion of Springer-Verlag; (d) reproduced with permission of Americal Institute of Biological Sciences)


Figure 10.18 Intercellular molecular traffic can be traced by low molecular weight fluorescent dyes, such as uranin (approx 800 Da) which has a molecular diamater of <2 nm, here injected into stamen hair cells of Setcreasea purpurea. Bright-field image before injection (a), 2 min after injecting the arrowed cell (b), 5 min later (c ). The dye is visible in the thin layer of cytoplasm around the edges of these highly vaculate cells. Because the dye cannot cross the plasma membrane, cell to cell movement must be by plasmodesmata. Contrast this with Figure 2 in Feature essay 10.1, showing similar experiments tracing movement of much larger macromolecules.

(Based on Tucker 1982; reproduced with permission of Springer-Verlag)

Counts of plasmodesmata in walls of cells of different ages demonstrate that in most tissues numbers in each cell wall are fixed at cytokinesis when plasmodesmal bridges of cytoplasm are retained within the advancing cell plate (Gunning 1978). Consequently, electrical coupling declines as cells enlarge because the number of plasmodesmata per unit membrane area decreases. There is some evidence that numbers of plasmodesmata between cell layers in shoot apical meristems reduce soon after transition to flowering, such as in Iris (Bergmans et al. 1997), which may indicate a further degree of isolation that relates to change in developmental pattern. In addition, some species exhibit ‘secondary’ plasmodesmata which increase numbers of connections during later cell development (Lucas et al. 1993). These have a modified structure with connections to several nearby plasmodesmata through an enlarged central region in the wall. New plasmodesmata also form in graft unions, often between tissues of two different species (Figure 10.19; Kollmann and Glockmann 1985). Grafting experiments indicate that some flowering stimuli have a strictly cytoplasmic route, only moving from induced to non-induced plant through fully established, plasmodesmal grafts.

Since the early 1990s, evidence has been accumulating that plasmodesmata sometimes also allow molecules of a much larger scale to pass through. The pioneering experiments were on viral movement proteins of about 30 kDa which become localised to plasmodesmata (Lucas et al. 1993). These proteins function to aid movement of viral components (protein and RNA) from cell to cell during infection spread. In transgenic plants expressing this movement protein, dye studies show that larger than normal molecules can pass between cells. If cells are injected with the protein itself, fluorescent molecules up to 9.4 kDa rapidly move out of the injected cell (Fujiwara et al. 1993). Some viruses induce much more drastic structural modification such as loss of the desmotubule, leaving an aperture wide enough for passage of entire virus particles.


Figure 10.19 Secondary plasmodesmata form later in cell development, well after the wall between two cells has been laid down. Although common in many cell types, this is neatly demonstrated in graft unions where the adjacent cells clearly had different ancestries. Here Vicia faba scions (V, left) were grafted into genetically unrelated Helianthus annuus rootstocks (H, right). In many instances, half plasmodesmata form in each cell wall but only sometimes do they match up (arrow) to form complete cytoplasmic bridges. Scale bar = 100 nm.

(Based on Kollmann and Glockmann 1985; reproduced with permission of Springer-Verlag)

Even in intact untreated tissue, larger molecules (up to 3 kDa) can sometimes move between cells, and there are reports of even 40 kDa molecules moving, albeit very slowly. Taken together with the virus studies, we are probably just starting to see a major revolution in our comprehension of intercellular signalling (Kragler et al. 1998). There is now evidence for selective transmission of regulatory molecules such as mRNA or proteins (e.g. Knotted1 in maize; Lucas et al. 1995) which may orchestrate development and functioning of small clusters of cells. Such macromolecular traffic is perhaps governed by ‘gate keeper’ recognition/receptor proteins at or in the cytoplasmic sleeve (McLean et al. 1997). This may be an additional layer of cellular control and perhaps as important as hormonal and other small molecule signalling systems.