Anne Mie C. Emons. Tel: +31 317 484329, +31 317 482155; fax: +31 317 485005; e-mail: firstname.lastname@example.org
Plant cell wall production is a membrane-bound process. Cell walls are composed of cellulose microfibrils, embedded inside a matrix of other polysaccharides and glycoproteins. The cell wall matrix is extruded into the existing cell wall by exocytosis. This same process also inserts the cellulose synthase complexes into the plasma membrane. These complexes, the nanomachines that produce the cellulose microfibrils, move inside the plasma membrane leaving the cellulose microfibrils in their wake. Cellulose microfibril angle is an important determinant of cell development and of tissue properties and as such relevant for the industrial use of plant material. Here, we provide an integrated view of the events taking place in the not more than 100 nm deep area in and around the plasma membrane, correlating recent results provided by the distinct field of plant cell biology. We discuss the coordinated activities of exocytosis, endocytosis, and movement of cellulose synthase complexes while producing cellulose microfibrils and the link of these processes to the cortical microtubules.
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Plant cell walls are sophisticated, complex and dynamic structures composed of networks of various polysaccharides, glycoproteins and phenolic compounds (Carpita & Gibeaut, 1993; McCann et al., 2001). In many plant cell types, cellulose, the most abundant biopolymer on earth, is the main cell wall component. Cellulose is present in cell walls as long cellulose microfibrils (CMFs) that wrap around the cells providing them with a load-bearing framework and necessary rigidity to restrain the cell's turgor pressure that results from hydrostatic pressure exerted on the cell walls (Diotallevi & Mulder, 2007). The orientation of CMFs, which have a tensile strength comparable to that of steel, is central to regulation of cell elongation, and CMF deposition orientation after cell elongation is essential to ensure proper wall strength.
Cell walls are biphasic structures; they are composed of CMFs, embedded inside a matrix of other polysaccharides and glycoproteins. This biphasic structure is produced by two separate, but connected deposition mechanisms. The matrix materials are extruded by exocytosis into the cell wall. This same process also inserts the cellulose synthase complexes (CSCs) into the plasma membrane. These complexes are the nanomachines that produce the CMFs from UDP-glucose recruited from the cytoplasm, depositing them into the wall in a membrane-bound process.
The CMFs are crystalline arrays of β-1,4-linked chains of glucose residues (Delmer, 1999). Measurements with wide angle (WAXS) and small angle (SAXS) X-ray scattering and NMR have revealed that the diameter of cellulose microfibrils of celery collenchyma is between 2.4 and 3.6 nm (Kennedy et al., 2007), which is consistent with previously published data, including measurements from TEM images (Emons, 1988). Kennedy et al. (2007) argue that this diameter range corresponds to between 15 and 25 cellulose polymers per microfibril, rather than the commonly quoted number of 36, which is strongly based on the implications of the observed 6-fold symmetry of the membrane-embedded part of the CSC, the so-called particle rosette.
The angle with respect to the long axis of the cell at which the CMFs are deposited is an important determinant of cell growth behaviour and cell wall properties. The totality of all individual CMFs is organized into a variety of patterns, cell wall textures, which are species- and cell developmental stage-specific and thus regulated by the cells they surround. Therefore, the regulation of CMF angle is an important issue. In elongating cells, CMFs at the outside and cortical microtubules at the inside of the plasma membrane are often in the same net orientation, transverse to the cell's elongation direction (Lloyd & Chan, 2004), and cortical microtubules border the area in tracheary elements where wall thickening of almost 100% cellulose occurs (Oda et al., 2005). Topical work, in which both cortical microtubules and CSCs were visualized in living cells, epidermis cells of elongating etiolated hypocotyls, shows that cortical microtubule orientation and the oriented movement of CSCs are correlated (Paredez et al., 2006). This seems to reduce the question of CMF ordering to the question of the ordering of cortical microtubules, at least in these cells. However, in the absence of the cortical microtubules the CSCs move in a definite pattern as well, pointing to the existence of an underlying non-cortical microtubule based ordering mechanism for CMFs.
All this CMF production activity and alignment occurs in and at the plasma membrane while exocytosis and endocytosis are going on. Figure 1(A) shows the cytoplasmic side of the plasma membrane of the sub-apex of a growing radish root with microtubules longitudinal to the long axis of the root hair and an abundance of coated pits. In this review, we discuss the current knowledge of these processes. First, we discuss membrane turn over: the exo- and endocytosis mechanisms. Then we present the CSCs, physical considerations of CMF production, the organization of the CMFs, and confer attention to the organization of the polymers at the other side of the plasma membrane, the cortical microtubules. Lastly, we discuss the coordination of all this activity, taking into account the physical aspects pertaining to the processes occurring in that limited area of approximately 100 nm bordering the plasma membrane.
Exocytosis is the process in which the membrane of cargo containing vesicles fuses with the plasma membrane, while delivering their cargo to the outside of the cell. Figure 1(B) shows exocytotic structures at the plasma membrane of embryogenic carrot suspension cells. Exocytosis is an important part of the generally coupled processes of plant cell elongation and cell wall deposition. Material contained inside (Golgi) vesicles is extruded out of the cytoplasm into the existing cell wall, and this is combined with insertion of integral membrane proteins, such as for instance the CSCs, into the plasma membrane. The vesicle membrane encloses a three-dimensional sphere, whereas the plasma membrane by comparison is effectively a flat plane. This purely geometrical mismatch in surface-to-volume ratio between these two structures results in an excess of membrane that is inserted into the plasma membrane when compared with the deposition of the vesicle contents into the extra-cellular lumen. To ensure that the membrane lies straight against the cell wall, and does not ruffle, excess membrane retrieval is required. Indeed, the cytoplasmic side of the plasma membrane is covered with many coated pits, demonstrating that clathrin-mediated endocytosis occurs abundantly (Emons & Traas, 1986). Besides regulating the amount of plasma membrane, endocytosis has been hypothesized to be involved in rapidly generating membrane supply during cell plate formation (Dhonukshe et al., 2006). However, recently it has been shown that this hypothesis is not true and that also vesicles that form the cell plate are Golgi-derived (Reichardt et al., 2007).
Figure 1(C) is a detail of a coated pit in the sub-apex of a growing Equisetum hyemale root hair. Calculations have been made to estimate the number of exocytosis and/or endocytosis vesicles required for cell elongation to occur. Miller et al. (1997) calculated that in E. hyemale root hairs, in which the growth velocity is approximately 0.67 μm min–1 (Emons & Wolters Arts, 1983), the extension of the plasma membrane requires 148 exocytotic vesicles with a measured diameter of 300 nm (Emons, 1987) per minute. In combination with an estimated membrane turnover time of 20–40 min, calculated from the amount of coated pits (Emons & Traas, 1986), this leads to a requirement of 445 vesicles per minute to sustain membrane recycling and cell expansion. In this calculation, 67% of the inserted membrane needs to be recycled. In Lilium pollen tubes, Morre & Van der Woude (1974) estimated the amount of exocytotic vesicles by assuming that the volume of the exocytotic vesicles contributed to the cell wall expansion stoichiometrically. They obtained a value of approximately 1000 vesicles per minute. Ketelaar et al. (2008, this issue) assumed that the plasma membrane surface area and cell wall volume increase stoichiometrically by the membrane surface area and cell wall matrix contents of exocytotic vesicles. They estimated that 86.7% of the inserted plasma membrane in Arabidopsis root hairs and 79% in Arabidopsis pollen tubes are recycled by endocytosis. These values are higher than the value found by Morre & Van der Woude (1974), but of the same order of magnitude.
Picton & Steer (1983) used a different approach to estimate the amount of Golgi-derived vesicles that are required for pollen tube growth. Using 0.3 μg mL–1 (approximately 0.6 μM) of the actin depolymerizing drug cytochalasin D, they inhibited growth of Tradescantia pollen tubes. As the actin cytoskeleton serves as a transport system to deliver exocytotic vesicles to the apex of the pollen tube, the delivery of newly synthesized exocytotic vesicles from the Golgi apparatus to the growing tip should be inhibited. Assuming that the production of exocytotic vesicles continues at the normal rate when actin is depolymerized, the accumulation of vesicles around the Golgi stacks in the cytoplasm was counted. After 5 min, they calculated the increase in the density of Golgi-derived vesicles and used this value to calculate the number of exocytic vesicles that would be consumed for pollen tube growth during that time. They estimate a production of 3000–5000 exocytic vesicles per minute. The reliability of this approach depends on the assumption that exocytic vesicle production is not inhibited by actin depolymerization, which these authors have shown not to occur in these cells (Picton & Steer, 1981).
Besides sustaining cell expansion, maintaining the ratio between cell wall volume and plasma membrane surface area, the large numbers of exocytotic vesicles that fuse with the plasma membrane and the large percentage of membrane recycling that occurs could serve as a mechanism to tightly control the amount and type of trans-membrane proteins at the plasma membrane. One trans-membrane protein complex of which the amount (and/or activity) in the plasma membrane requires tight regulation for cell wall deposition is the CSC. In the next paragraph, we will discuss the current knowledge about this complex.
Cellulose synthase complexes
The supramolecular complexity of CMFs is reflected in their synthetic machinery, the CSCs (Somerville et al., 2004). CSCs are large transmembrane protein complexes, which move through the plasma membrane while producing CMFs (Paredez et al., 2006). They have been visualized by transmission electron microscopy of freeze-fractured membranes. These electron micrographs of plasmatic fracture faces of higher plant plasma membranes show six particles in a hexagonal arrangement, often at the end of a CMF. These arrangements of six particles have been called ‘particle rosettes’, or briefly, ‘rosettes’. In higher plant cells, only single rosettes have been reported. The composing six particles are each approximately 8 nm wide, and the total rosette diameter is approximately 24 nm (Mueller & Brown, 1980; Emons, 1985), both sizes including the 2- to 5-nm-thick platinum/carbon shadow of the freeze fracture/replication technique.
The catalytic enzyme cellulose synthase was the first component of the CSC that has been cloned (Pear et al., 1996). Direct genetic evidence that the CESA gene was the cellulose synthase came from the work of Arioli et al., who complemented the root swelling (rsw1) Arabidopsis mutant (Arioli et al., 1998). The availability of the CESA sequence enabled the manufacture of antibodies that were used by Kimura et al. to show that the particle rosettes indeed contain cellulose synthase proteins (Kimura et al., 1999).
In Arabidopsis thaliana, the cellulose synthase family contains at least ten different isoforms, AtCESA1 to AtCESA10, reviewed by Sommerville (2006), of which the encoded proteins have eight membrane-spanning regions, thought to form a pore. The catalytic centre contains the D,D,D,QxxRW motif, in which the D,D,D part could bind UDP-glucose (Richmond & Somerville, 2000). The N-terminus contains a double zinc finger motive called LIM domain. LIM domains in human cells are involved in protein–protein interactions. The zinc fingers of cellulose synthase could well be involved in protein interactions, for instance between CESA sub-units within the complex, or between a CESA and another protein.
During xylogenesis in Arabidopsis, three functionally different genes CESA4, -7 and -8 are required to make functional CSCs (Taylor et al., 2003). These findings were supported by microarray co-expression which showed very high co-expression of CESA4, -7 and -8 during secondary cell wall production (Birnbaum et al., 2003; Persson et al., 2005). Persson et al. (2005) also showed that genes involved in primary cell wall synthesis CESA1, -3 and -6 were highly co-expressed. The cesa1 and cesa3 mutants are gametophytic lethal but cesa2, -5, -6 and -9 have relatively mild phenotypes (Persson et al., 2007). Recent studies by Desprez et al. (2007) and Persson et al. (2007) show that the CESA6-like genes, CESA2, -5, -6 and -9, are at least partially redundant and cesa2, -6 and -9 triple mutants are gametophytic lethal. Very little is known about the function of the CESA10 gene.
It appears that there are two different classes of CSCs. One class of complexes, containing the CESA4, -7 and -8 proteins, is likely to be involved in secondary wall formation and the other class of complexes, containing CESA1, -3 complemented by either CESA2, -5, -6 or -9, is responsible for cellulose deposition during cell growth. The transcriptional regulation of the sub-units and subsequent assembly into functional CSCs is still poorly understood.
Physical aspects of cellulose production
Knowing from which components the CSC is built is not enough to understand its mode of operation. It is by now widely assumed that the force for CSC propulsion derives from the cellulose synthesis itself (Herth, 1980; Roberts et al., 1982). If that is indeed the case, we need to answer the question of how the energy released upon the synthesis of individual cellulose polymers by the CESAs and their subsequent aggregation into the CMFs is converted into motion. To that end, we must take into account the full set of physical constraints under which the CMF deposition process takes place.
Although many, if not most, of the details of this process are as yet experimentally inaccessible, we are still able to form a minimal model of CSC propulsion based on these constraints. The first is that the CSC itself is embedded in the plasma membrane. Both its ‘top view’ as seen in the inner plasma membrane leaflets (PF-face) of freeze fracture EM images (Herth, 1980; Mueller & Brown, 1980; Emons, 1985) and its ‘bottom view’ provided by the recent work of the laboratory of R.M. Brown Jr. (Bowling & Brown, unpublished work) show a laterally extended and roughly circular aspect and to a first approximation one can assume that this disk-like structure lies parallel to the membrane surface. From symmetry considerations, one also expects that the cellulose polymer chains emerge perpendicularly to the plane of the complex.
As the CMF formed in the wake of the CSC is trapped in the space between the plasma membrane and the already extant cell wall, and the first is pressed against the second by the turgor pressure, the CMF itself must also lie in the plane of the membrane. This implies that, independently of the exact sequence of events in the crystallization process, the individual cellulose microfibrils must be bent in their path from the CSC to the CMF. As this bending of the polymers, be it individually or, as has been suggested, already partially crystallized into sheets (Brown & Saxena, 2000), costs energy, forces are exerted at the location at which the chains are connected to the CSC and the CMF. At the connection point to the CSC, these forces can be decomposed into a downward component pointing into the plane of the CSC and to a lateral component parallel to this plane. The perpendicular forces counteract the extrusion process and hence decrease the polymerization rate, and effect which is readily modelled by the classical Brownian ratchet mechanism (Peskin et al., 1993). As a by-product of these perpendicular forces, the CSC as a whole and part of the membrane connected to it will be pressed downward, an effect probably responsible for the membrane depressions seen in some of the freeze fracture images. As the bending of the chains can be alleviated by moving the attachment point to the CSC farther away from the other constraint point formed by the attachment to the CMF, the in-plane components are responsible for moving the CSC, again by symmetry, in the direction into which the nascent CMF points. At the attachment point to the CMF, any force loading will tend to hinder the crystallization process. These forces are decreased whenever the CSC moves farther away, allowing the CMF to grow, tracking the motion of the CSC. The interplay between the physical forces generated and the polymerization and crystallization process allows a stationary state to be reached in which a constant force is exerted on the CSC that in turn is balanced by the viscous stresses in the fluid media (membrane and cytoplasm) in which it is immersed, leading to a constant speed of propagation.
Based on the heuristic mechanism described earlier, Diotallevi & Mulder (2007) were able to formulate an analytical model of CSC propulsion that correctly estimates the order of magnitude of the speed (102–103 nm min–1) of motion as observed in the recent experiments (Paredez et al., 2006). Furthermore, they provided a proof of principle of this mechanism by implementing a stochastic simulation, including all of the relevant ingredients explicitly, that shows how a CSC producing six cellulose strands progresses in the membrane.
Knowing how a single CSC is propelled, however, is still a long way from understanding how the often strikingly ordered textures of plant cell walls come about. To that end we must come to the grips with the question of which elements of the environment of a CSC determine its direction of motion within the plasma membrane.
Cell wall texture
In cell walls, CMFs are arranged in lamellae that together form the cell wall texture, the architecture of the totality of all the CMFs. The orientation of CMFs with respect to the long axis of the cell within a lamella is approximately constant, but may vary from lamella to lamella, depending on the type of texture. The distance between CMFs within a lamella varies from a few nanometres in cell wall thickenings of Lepidium xylem cells, up to 160 nm in helicoidal cell walls of root hairs, but exact numbers have hardly been reported (Emons, 1991). The most striking texture is the helicoidal cell wall texture, which consists of subsequent lamellae in which the orientation of the CMFs is shifted by a constant angle. Figure 1(F) shows cellulose microfibrils with different orientations in three subsequent lamellae of the helicoidal cell wall of an E. hyemale root hair. Other wall textures are the axial, helical, crossed-polylamellate, transverse and the random wall textures and combinations of these. These names refer either to the relative orientation of the CMFs with respect to each other or to their absolute orientation with respect to the long axis of the cell.
How is the biogenesis of wall textures regulated from within the cell? As mentioned earlier, CMFs are produced by CSCs, and it is by now widely assumed that these CSCs are propelled through the membrane by the force of the cellulose production itself, combined with the force that is generated by attachment of the nascent CMF to other cell wall components as soon as they are produced. CMFs are stiff, but have been seen to bend around pit fields (Mueller & Brown, 1980), showing that their intrinsic orientation of deposition can be over-ruled by mechanical obstacles. The CSCs indeed move in straight lines inside the plasma membrane in etiolated Arabidopsis hypocotyl epidermis cells, and the direction of movement of the CSCs appears to be forced upon them by cortical microtubules (Paredez et al., 2006), which are potential obstacles they encounter in their plane of movement (Emons et al., 2007). Cortical microtubules could be over-ruling the self-orientation mechanism of CSCs in all elongating cells. They ensure that CMFs are aligned consistently across the growing Arabidopsis root (Baskin et al., 2004).
When the microtubules are completely depolymerized, the complexes move inside the plasma membrane in ordered patterns as well (Paredez et al., 2006). In addition, examples are known of non-parallel cortical microtubules and CMFs in secondary wall deposition, that is wall thickening after cell elongation (Emons et al., 1992). The ordering of cellulose microfibrils during their deposition in such cells has been explained with a geometrical model (Emons 1994; Emons & Mulder, 1998; Mulder & Emons, 2001). This model provides a conceptual framework for the alignment mechanism of CMFs, which departs from the situation where cortical microtubules are not parallel to nascent CMFs. The basic assumption is that by default, CMFs go straight unless obstructed and that their alignment depends mainly on the number of CSCs simultaneously active at any position in the plasma membrane. Of course, this model does not rule out that cortical microtubules bind to the plasma membrane so tightly that synthase movement is obstructed (Emons et al., 2007). In addition, other work in which cellulose synthase and drugs against microtubules, and relevant mutants of Arabidopsis, were used does not point to a simple one-on-one relationship between the orientations of the two sets of polymers at both sides of the plasma membrane (Himmelspach et al., 2003; Sugimoto et al., 2003).
Despite these exceptions, and the, as yet, unexplained occurrences of incongruent CMFs and cortical microtubules, the work of Paredez et al. (2006) with CSCs and microtubules fluorescently labelled in the same cells, has made it clear that cortical microtubule orientation is an important regulator of CMF orientation, at least in epidermis cells of elongating etiolated Arabidopsis hypocotyls, and maybe in all elongating plant cells. This begs the question of how the cortical microtubules themselves are being organized.
In inter-phase plant cells, microtubules are localized in the cortex, just underneath the plasma membrane (Ledbetter & Porter, 1963; Hepler & Newcomb, 1964). Figure 1(E) shows the typical close connection of microtubules to the plasma membrane in a root hair of Vicia sativa. Although they are indirectly linked to the plasma membrane, they are at the same time dynamic (Hardham & Gunning, 1978; Wasteneys, 2002; Shaw et al., 2003; Vos et al., 2004; Hirase et al., 2006; DeBolt et al., 2007). They behave according to a hybrid treadmilling mechanism; their minus ends mostly pausing or shrinking and their plus ends alternating between growing and shrinking phases (Shaw et al., 2003). For this behaviour, the properties of dynamic instability must be carefully regulated through the interaction with microtubule-associated proteins (MAPs). For example, during G2/prophase, the microtubules become more dynamic and possibly longer by means of a hypothetical search and capture mechanism that helps the formation of the pre-prophase band (Vos et al., 2004). Ove-rexpression or knock out of a MAP can have severe effects. Typically, in such mutants, for example mor1, katanin, wvd, inter-phase cortical microtubules no longer organize perpendicularly to the growth axis of the cell, and the cells expand isotropically (Whittington et al., 2001; Burk & Ye, 2002; Yuen et al., 2003). The details of the mechanisms by which these MAPs determine the microtubule organization are largely unknown, but it could be that they control the stability and length of the microtubule and that these parameters in turn are e ssential for a self-organization mechanism that is based on microtubule–microtubule interactions.
So far, two distinct behaviours of cortical microtubules have been observed: zippering and catastrophic collisions (Dixit & Cyr, 2004). A microtubule that runs into another microtubule at a small angle (<40°) will alter its direction of polymerization and become bundled with it in a parallel or antiparallel fashion. When the polymerizing microtubule collides with a pre-existing one at a larger angle (>40°), the microtubule has a heightened chance of having a catastrophe: a switch from growing to shrinking. From this result, we can deduce that the unusually few, short or stable microtubules in several MAP mutants have fewer interactions with each other and therefore cannot align with each other. Nevertheless, although these two mechanisms together can align microtubules, they cannot dictate the direction of alignment.
We have shown, based on theoretical considerations and in vitro work, that microtubules can also change their direction by being bent and by polymerizing along the inner walls of (artificial) cells (Lagomarsino et al., 2007). However, confinement within the cell alone cannot organize the microtubules perpendicularly to the growth axis of the cell; rather the bending rigidity dictates the microtubules to become longitudinally arranged. This indicates that other, so far unknown, microtubule–microtubule interacting mechanisms must be at work to organize the microtubule array, or that the alignment perpendicular to the elongation axis of the cell is dependent on other, for example microtubule-plasma membrane mechanisms.
Chan et al. (2007) showed that domains of aligned microtubules move through the cortex of growing hypocotyl cells. The domains move and pivot a full 360°, on their way running into each other, splitting themselves or absorbing neighbouring domains. Whether or not these movements also occur in other cell types has not been shown yet, nor is it known if there is a relation between the moving domains and the formation of the helicoidal cell wall texture of CMFs. It is clear though that cortical microtubules do not exert a direct orienting influence on CMFs, but rather that they influence the path of the CSCs (Paredez et al., 2006). These complexes stick out below the plasma membrane into the cytoplasm where the cortical microtubules are located and which may act as tracks or as guard rail (Emons et al., 2007). The hypothesis that microtubules are ordered through a regulated cortical microtubule self-organization mechanism is still valid and the focus of research in several groups.
Activity at the plasma membrane
From the above, it is clear that cellulose deposition into the plant cell wall is a space–time process that involves a number of concurrent activities, all localized to a small area bordering the plasma membrane. In a drawing (Figs 1 & 2), we show the relevant structures occurring at the plasma membrane involved in the process of cellulose production. Exocytosis delivers new CSCs to the plasma membrane, at the same time depositing other wall polymers, whereas concomitant endocytosis recycles the excess membrane area created by the exocytosis. The CSCs, once inserted into the plasma membrane, move in the plane of the membrane, driven forward by the energy released in the cellulose polymerization process, leaving a CMF in their wake. The cortical microtubule network, which itself is dynamically self-assembled from its component parts, seems to play a role in regulating the motion of the CSCs, possibly by guiding their direction of motion. Understanding how these local activities are coordinated to produce textured cell walls that show spatially coherent patterns over many micrometres, in our opinion, requires a systems biological approach, in which the known biochemical, biophysical and cell biological data are integrated into mathematical models. We believe that only the detailed, quantitative, predictions that follow from such models will enable us to effectively select and design experiments aimed at unravelling the complex, collective, effects that underpin cell wall formation.
A.M.C.E. thanks the FOM Institute for Atomic and Molecular Physics (AMOLF), Amsterdam, for financial support for this project. This work was co-funded through EU grant 028974 CASPIC (JL, BM, AMCE). T.K. and J.V. were supported by VENI fellowships 863.04.003 and 863.02.009, respectively, from the Dutch Science Foundation (NWO). The work of B.M. is part of the research program of the ‘Stichting voor Fundamenteel Onderzoek der Materie (FOM)’, which is financially supported by the NWO.