Molecular Plant Advance Access originally published online on September 4, 2009
Molecular Plant 2009 2(5):990-999; doi:10.1093/mp/ssp065
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Pectin May Hinder the Unfolding of Xyloglucan Chains during Cell Deformation: Implications of the Mechanical Performance of Arabidopsis Hypocotyls with Pectin Alterations
a Max-Planck-Institute of Colloids and Interfaces, Department of Biomaterials, Potsdam, Germany
b College of Forestry and Natural Resources, University of the Philippines Los Baños, Philippines
c Department of Socio-Environmental Energy Science, Kyoto University, Japan
d Max-Planck-Institute for Molecular Plant Physiology, Potsdam, Germany
e Laboratoire de Biologie Cellulaire, UR501, Institute Jean-Pierre Bourgin, INRA, Versailles, France
f Michigan State University, Plant Research Laboratory, East Lansing, Michigan, USA
1 To whom correspondence should be addressed. E-mail ingo.burgert{at}mpikg.mpg.de, fax +49 331 567 9402.
 |
Abstract |
|---|
 TOP Abstract INTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
Plant cell walls, like a multitude of other biological materials, are natural fiber-reinforced composite materials. Their mechanical properties are highly dependent on the interplay of the stiff fibrous phase and the soft matrix phase and on the matrix deformation itself. Using specific Arabidopsis thaliana mutants, we studied the mechanical role of the matrix assembly in primary cell walls of hypocotyls with altered xyloglucan and pectin composition. Standard microtensile tests and cyclic loading protocols were performed on mur1 hypocotyls with affected RGII borate diester cross-links and a hindered xyloglucan fucosylation as well as qua2 exhibiting 50% less homogalacturonan in comparison to wild-type. As a control, wild-type plants (Col-0) and mur2 exhibiting a specific xyloglucan fucosylation and no differences in the pectin network were utilized. In the standard tensile tests, the ultimate stress levels (
tensile strength) of the hypocotyls of the mutants with pectin alterations (mur1, qua2) were rather unaffected, whereas their tensile stiffness was noticeably reduced in comparison to Col-0. The cyclic loading tests indicated a stiffening of all hypocotyls after the first cycle and a plastic deformation during the first straining, the degree of which, however, was much higher for mur1 and qua2 hypocotyls. Based on the mechanical data and current cell wall models, it is assumed that folded xyloglucan chains between cellulose fibrils may tend to unfold during straining of the hypocotyls. This response is probably hindered by geometrical constraints due to pectin rigidity.
Key Words: Arabidopsis thaliana • mutants • cellulose • xyloglucan • pectin • cyclic loading tests
Received for publication May 5, 2009. Accepted for publication July 14, 2009.
|
INTRODUCTION |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
The primary cell wall of plants is a unique engineering structure that combines conflicting characteristics such as rigidity as well as plasticity and compliance (Rose and Bennett, 1999; Whitney et al., 1999; Cosgrove, 2000). Rigidity is needed to withstand the osmotic pressure of the living cell (Taiz, 1984) and to cope with external loads, whereas sufficient plasticity and compliance are needed for cell wall expansion during growth (Baskin, 2005). Furthermore, the primary wall is specifically designed to provide a rigid barrier against pathogenic intrusions (Creelman and Mullet, 1997) and, at the same time, performs dynamic tasks in absorption, transport, and secretion of substances throughout plant growth and development (Eckardt, 2003).
In dicotyledonous plants, the primary wall consists of approximately 30% cellulose, 30% hemicelluloses, 35% pectin, and 1–5% structural proteins on a dry weight basis (Vorwerk et al., 2004). The structures of the individual polymers are well known; however, their specific arrangement and bonding patterns in the entire cell wall are not fully understood yet.
The complex assembly of stiff cellulose fibrils and pliable matrix components can be characterized in the same way as fiber-reinforced composites (Kerstens et al., 2001; Fratzl et al., 2004). Based on deep-etching methods and NMR spectroscopy, several cell wall models have been proposed (Keegstra et al., 1973; Hayashi, 1989; Fry, 1989a; Talbott and Ray, 1992; Ha et al., 1997; Cosgrove, 2000). Generally, the cellulose microfibrils are thought to be tethered by xyloglucan mainly through hydrogen bonding (Hayashi, 1989). Xyloglucan attaches itself on the fibril surface as well as in between fibrils (Pauly et al., 1999). Thereby, it coats the fibrils serving as a spacer that prevents direct hydrogen bonding between cellulose chains (Carpita and Gibeaut, 1993). Pectic polysaccharides form a co-extensive network that interpenetrate this network (McCann et al., 1990; Talbott and Ray, 1992) and interact to a certain degree with hemicelluloses via both non-covalent and covalent bonding (Fry, 1989b; Thompson and Fry, 2000).
From the 1960s onwards, mechanical measurements have been performed on plant cell walls using extensometers to elucidate factors that influence the extensibility of the cell walls as well as the mechanical role of the cell wall components and their interaction in the entire cell wall (Cleland, 1967, 1984; Cleland and Rayle, 1977; Cosgrove 1988, 1989). Uniaxial tensile tests have been also performed on various Arabidopsis mutants (Köhler and Spatz, 2002; Ryden et al., 2003; Peña et al., 2004; Cavalier et al., 2008). Ryden et al. (2003) compared stiffness and strength of Arabidopsis hypocotyls of GDP–fucose biosynthesis mutant mur1, the xyloglucan fucosyltransferase mutant mur2, and the xyloglucan galactosyltransferase mutant mur3. The results were interpreted in a way that the mechanical performance of primary walls depends on both galactosylated xyloglucan side chains and borate-complexed rhamnogalacturonan II. However, a mechanistic model that proposes how xyloglucan and pectin influence the stiffness and strength of Arabidopsis hypocotyls has not been proposed yet.
The approach reported here aims to gather further insight into the principle deformation mechanisms of the primary cell wall and the mechanical interactions of the polymer networks, particularly the interactions of xyloglucan and pectin. To elucidate the mechanical role and the interplay of the structural networks in the primary cell wall, standard tensile tests and cyclic loading experiments were carried out on mur-mutants (mur1 and mur2) and qua2. The latter mutant contains 50% less homogalacturonan in comparison to the wild-type (Mouille et al., 2007; Ralet et al., 2008). Based on the mechanical responses of the various hypocotyls, a simple structural model is proposed that extends existing models on the deformation of the cellulose fibril–xyloglucan network (Passioura, 1994; Passioura and Fry, 1992; Veytsman and Cosgrove, 1998) by a possible interplay of xyloglucan chains with the pectin network. This model shows some analogies to the so called ‘hidden length mechanism’, which, for instance, explains the high toughness of bone by an additional deformability of matrix polymers due to their specific structural alignment in the assembly and polymer interactions by means of ionic sacrificial bonds (Fantner et al., 2005; Gupta et al., 2007).
|
RESULTS |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
Figure 1A shows a representative stress–strain curve of a 4-day-old Arabidopsis wild-type (Col-0) hypocotyl illustrating its mechanical behavior and how the mechanical parameters used in this study were determined.
|
plastic) were determined.The stress–strain curve of the standard tensile test shows an initial phase, which is followed by an almost linear phase and a regime of non-linear deformation after yield. The curve ends at the point of rupture. Stiffness was calculated from the slope of the curve in segment 2 and the ultimate stress value can be taken as an approximate measure of the strength of the hypocotyl.
In Figure 1B, an exemplary stress–strain curve of a 4-day-old Arabidopsis mur1 hypocotyl in a cyclic loading experiment is shown. Cyclic loading tests can further provide important information on the deformation behavior of a sample. Stiffness was calculated for the upward loading phases. Stiffness 1 equates to the stiffness in the standard loading experiment presented in Figure 1A. Stiffness 2, stiffness 3, and so forth reflect the material response when the sample is re-loaded after unloading in the cycles. Cyclic loading experiments also allow the distinction between the elastic and the plastic fraction of a material response (Cleland, 1984). In a pure elastic deformation, all energy is returned after unloading, which means that the unloading curve should hit the abscissa in the initial point of the experiment. The ‘plastic strain’, as indicated in Figure 1B, was calculated as a qualitative measure of irreversible deformation. The given example also shows that the initial slope (stiffness 1) of a hypocotyl does not necessarily reflect a pure elastic material response (Young's Modulus).
Standard tensile tests according to Figure 1A were carried out on the hypocotyls to determine their ultimate stress levels (strength) and the stiffness (stiffness 1). Figure 2 shows the mechanical behavior of the 4-day-old mur hypocotyls (Fig. 2A) and the 6-day-old qua2 hypocotyls (Fig. 2B). Ultimate stress levels and the stiffness of Col-0 hypocotyls of both respective ages are shown for reference.
|
In the ultimate stress-versus-stiffness plots (Figure 2), only mur2 shows a significant difference in the ultimate stress level from Col-0, whereas no significant differences between Col-0 and the mutants with pectin alterations, mur1 and qua2, were observed. However, in terms of stiffness, all mutants were significantly different from Col-0. While mur2 showed only a moderate reduction in stiffness (
13%), the stiffness of the two mutants with pectin alterations was noticeably decreased. The stiffness of mur1 was reduced by 40% and the stiffness of qua2 by 34% compared to the Col-0 hypocotyls. Above the other hypocotyls, several hypocotyls of the mur1 mutant showed a pronounced non-linear deformation phase in the beginning of the tensile tests, which made it necessary to determine the stiffness at rather high strain levels (probably after a certain re-stiffening). Therefore, the stiffness shown for mur1 is likely to be the upper limit. Although mutants were tested at different ages, one can observe a cluster of Col-0 and mur2 and a cluster of mur1 and qua2, which differ noticeably in stiffness but not in ultimate stress levels.
To better understand the deformation mechanisms of the primary cell walls of the hypocotyls, cyclic loading tests were carried out in axial tension according to Figure 1B. After several cycles, the samples were stressed until failure. Figure 3A–3E show exemplary stress–strain curves of the 4 and 6-day-old hypocotyls, whereas, in Figure 3F–3J, the changes in stiffness are quantified for the first, second, and third cycle.
|
The exemplary stress–strain curves clearly point to a different mechanical response of the hypocotyls with pectin alterations, but only within the first loading phase of the experiment. The differences in the slopes between the first and second cycles are much more pronounced for mur1 and qua2 compared to Col-0 and mur2. In all further cycles, the deformation pattern seems to be rather consistent for all types of hypocotyls. The apparent differences in slopes between the first and the second cycles were quantified in the vertical bar graphs in Figure 3F–3J. All samples show a pronounced increase in stiffness from the first to the second cycle, whereas only little change in stiffness can be detected from the second to the third cycle.
Normalizing the stiffness of the first cycle (stiffness 1) as 100%, Col-0 (4-day-old), Col-0 (6-day-old), and mur2 showed an increase in stiffness from the first to the second cycle to 136, 123, and 122%, respectively, whereas the relative stiffness to mur1 and qua2 was considerably higher, reaching 186 and 168%, respectively (Figure 4A, the normalized stiffness of Col-0 is shown only for the 4-day-old hypocotyls). Another important parameter that reflects the different behavior of the hypocotyls is the plastic deformation (
plastic, in Figure 1B) during the first loading (Figure 4B).
|
Both hypocotyls with pectin alterations, mur1 and qua2, show notably higher plastic strain levels compared to Col-0 and mur2. However, this finding has to be qualified by saying that the determination of the plastic strain from the stress–strain curves is limited by experimental uncertainties. In fact, it is not possible to distinguish between plastic deformation and additional elongation due to sample reorientation in the initial phase of the experiment. Moreover, the loading and unloading cycles were performed continuously and therefore not all viscoelastic deformation that is characteristic for plant cell walls (Cleland, 1984) might have been relaxed before reloading. Hence, these data should be taken qualitatively.
In order to consider the mechanical properties of the hypocotyls as being indicative of specific deformation patterns of the cell walls, it is necessary to include the impact of the cell wall modification on hypocotyl turgor pressure (Figure 5). Differences in hydrostatic pressure would be of crucial relevance on the mechanical response of the hypocotyls, because turgor pressure can influence the longitudinal stiffness of the hypocotyls mainly by its impact on the Poisson's ratio of the hypocotyl by means of increasing the stiffness of the hypocotyl in its transverse direction.
|
The data indicate that the cell wall modifications did not result in pronounced differences in turgor pressure between the hypocotyls of the mutants and the wild-type. However, since turgor pressure was determined indirectly by the difference of water potential and osmotic pressure, two aspects that qualify the results need to be addressed. The indirect calculation of turgor pressure leads to rather large standard deviations because of error propagation and stress relaxation of the cell walls that is likely to occur in the psychrometer also affects turgor pressure (Cosgrove et al., 1984). Therefore, in terms of the latter point, it is important additional information whether the osmotic pressures of the different hypocotyls are in the same range (Col-0: 0.94 ± 0.13 MPa, mur1: 0.92 ± 0.16 MPa, mur2: 0.92 ± 0.15 MPa, qua2: 0.83 ± 0.04 MPa). This is the case, even though qua2 is slightly lower and significantly different from Col-0 (non-parametric U-test 0.05 level) but not different from mur1 and mur2. However, values for mur1 strongly coincide with Col-0 and mur2.
|
DISCUSSION |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
The primary cell wall consists of three networks that are made up of cellulose fibrils and xyloglucan, pectin, and structural proteins. These networks are described as being independent but interrelated (Schindler, 1998). For the mechanical performance of primary cell walls, it is believed that mainly the cellulose and xyloglucan network plays a crucial role (Carpita and Gibeaut, 1993; Peña et al., 2004; Cosgrove, 2005); however, also the pectin network seems to be important. Mechanical tests by Ryden and co-workers imply that borate-complexed rhamnogalacturonan II formation influences the mechanical properties of the cell wall as a stiffening and strengthening agent (Ryden et al., 2003). Although these investigations provided an indication of the mechanical relevance of the individual networks, it is not yet understood how the networks mechanically interact in the complex cell wall assembly and what the specific mechanical role of a certain network component is.
The analysis of the mechanical behavior of plants with cell walls of different structural/chemical composition may help to get better insight into the mechanical interaction of the cell wall macromolecules. In our study, we compared the mechanical behavior of well characterized Arabidopsis hypocotyls with alterations in the xyloglucan side chain structure and the pectin composition, respectively. In comparison to the wild-type, the xyloglucan fucosyltransferase mutant mur2 lacks the terminal fucose sugar unit. It has already been shown that this alteration rather marginally affects the mechanical performance of the hypocotyl whereas more severe xyloglucan side chain alterations, such as that found in the xyloglucan galactosyltransferase mutant mur3, result in a pronounced decrease in strength and stiffness (Ryden et al., 2003; Peña et al., 2004). In the hypocotyls of the GDP–fucose biosynthesis mutant mur1, the amount of fucose units is 60% of that in wild-type hypocotyls (Ryden et al., 2003). This not only leads to the absence of terminal fucose units in the xyloglucan side chains, but has an additional effect on pectic polysaccharides. The deficiency of fucose in rhamnogalacturonan II reduces its ability to form borate diesters through the apiosyl residue (Kobayashi et al., 1996). Qua2 shows an exclusive alteration in the pectin network by means of 50% less homogalacturonan in comparison to the wild-type (Mouille et al., 2007; Ralet et al., 2008).
From the standard tensile tests, two clusters of material properties were observed, one consisting of Col-0 and mur2 and one of mur1 and qua2. In terms of tensile stiffness of the mur-mutants, a similar trend was reported by Ryden et al. (2003), although their values were somewhat higher (50%). This is likely due to differences in the nutrient concentration of the medium, the growth temperature (25°C (Ryden et al., 2003) versus 22°C here), and the test-setup (in particular testing velocity). Indeed, when Col-0 was grown at 25°C, stiffness was 42% higher than the ones grown at 22°C (data not shown). For ultimate stress, however, the results were partly inconsistent with the Ryden data. They found Col-0 to have the highest strength and mur2 to be stronger than mur1, whereas, in our study, Col-0 and the two mur-mutants as well as the qua2 showed rather similar ultimate stress levels. These inconsistencies should be taken into account here, although they cannot be explained by different hypocotyl processing at this stage.
One limitation of the mechanical characterization of hypocotyls is that the system is fairly complex and cell wall properties are not measured directly. However, our main interest was not in stiffness and ultimate stress values of the cell walls, but in the change of deformation patterns of the hypocotyls and relative changes in stiffness and ultimate stress in the course of cell wall modifications. In order to calculate cell wall properties, several structural parameters (e.g. tissue density, cell length) and water interactions would need to be measured. One important parameter that can be expected to influence also the mechanical deformation pattern of the hypocotyls is the turgor pressure. However, as we could not detect considerable differences in turgor pressure between the hypocotyls of the mutants and the wild-type, the different mechanical behaviors of the mutants with pectin alterations (mur1, qua2) are not likely to be due to changes in turgor pressure. Another aspect to be considered is that water may be pressed out of the cells during mechanical testing and that pectin alterations may facilitate this process by increasing the permeability of the cell wall (Fleischer et al., 1999). However, such a process could not explain the more pronounced re-stiffening in the loading cycles of hypocotyls with pectin alterations compared to the wild-type and the mur2 mutant. Therefore, we conclude that differences in the mechanical behaviors are mainly indicative for changes in structure–property relationships due to the cell wall modifications.
Although cell wall properties could have not been directly measured, our finding that stiffness rather than ultimate stress of the hypocotyls is more affected by the pectin alterations (see Figure 2) has several implications for the possible spatial organization of pectin in the cell wall. The observations rule out that pectin is the matrix component that prominently contributes to a direct tethering of cellulose fibrils, as both parameters (stiffness and ultimate stress) would be affected in this configuration. On the contrary, in terms of mur2, stiffness and ultimate stress are reduced by almost the same relative amount compared to the wild-type. However, the data clearly point to the mechanical relevance of pectin as a structural element, and, although mur1 and qua2 possess alterations of different pectin components, they showed a comparable mechanical response.
To specify the possible mechanical role of pectin, the specific deformation patterns of the hypocotyls under cyclic regimes have to be considered. All mutants and the wild-type displayed a stiffening effect from the first to the second cycle during cyclic loading but the increase in stiffness was more pronounced for mur1 and qua2. Moreover, while Col-0 and mur2 showed only little plastic deformation, the irreversible deformation of mur1 and qua2 in the first cycle was noticeably higher (see Figure 4).
The increase in stiffness from the first to the second cycle should be generally associated with polymer reorientation in the cell wall towards the stress axis (Richmond et al., 1980). According to several authors (Preston, 1974; Reiterer et al., 1999), the stiffness of a plant cell wall is a function of its cellulose microfibril orientation. However, assuming almost transverse-oriented fibrils, an increase in stiffness due to microfibril reorientation is unlikely, since much higher strains than applied are needed to induce a noticeable passive reorientation of cellulose fibrils (Burgert and Fratzl, 2009). Although different deformation mechanisms should be dominant at various strain rates, it is interesting to note that also Marga et al. (2005) could not detect a change in cellulose fibril orientation even after up to 30% straining of cucumber hypocotyls in a creep experiment.
In a network of transversely oriented cellulose fibrils interconnected by xyloglucan chains, the latter are likely to be the main load-bearing part. The length of the xyloglucan chains is much longer than the spacing between cellulose fibrils (McCann and Roberts, 1991; McCann et al., 1992), which makes it likely that there is some folding of the xyloglucan chains. During straining of the hypocotyls, the xyloglucan chains may unfold. Hence, a possible explanation for the general stiffening effect during the first cycle is that the xyloglucan chains are straightening and thereby stiffening the entire wall. Thus, the relative increase in stiffness would depend on the absolute axial extension, since it dictates the degree of straightening of the xyloglucan backbone.
The hypocotyls with pectin alterations show an entirely different deformation pattern compared to the wild-type by means of stiffening behavior and plastic deformation. These differences seem to disappear after the second cycle. If pectins hinder the unfolding of the xyloglucan chain, then they would have the capability to stiffen the network and reduce the amount of chain flexibility in the first loading cycle. Although Thompson and Fry (2000) could show that xyloglucan and pectin are able to bind together covalently, an extensive cross-linking and, in particular, an inter- and intra-chain connection between xyloglucan chains and pectin seems unlikely. Therefore, besides some cross-linking, mainly geometrical constraints in pectin–xyloglucan interactions may influence the flexibility of the xyloglucan chains. If pectin, surrounding xyloglucan chains, has a high rigidity, the unfolding process is hindered or limited. The rigidity of the pectin should depend on the amount of pectin (qua2) and the number of ion-mediated cross-links (mur1), as cross-linking in general stiffens polymer networks (Boyd and Phillips, 1993). By modifying the pectin in the cell wall (mur1 and qua2), the amount of pectin and ion-mediated cross-linking decreases, which softens the system and allows the xyloglucan chains to unfold during deformation.
In Figure 6, the influence of pectin rigidity on xyloglucan chain unfolding and thereby cell wall deformability is illustrated schematically. Figure 6A shows the initial state as assumed for the two clusters, with and without pectin alteration, respectively. The xyloglucan chains are laterally bonded to the cellulose fibrils and folded in the space between the fibrils. The xyloglucan chains are embedded in a dense mesh of pectin. The pectin alterations are simply illustrated by a wider pectin mesh, since the model does not distinguish between the different pectin components, the number of ion-mediated bonds in pectin, and cross-links to the xyloglucan. As the alterations of two different pectin polymers lead to similar mechanical responses and the nature of the interconnections between the xyloglucan and pectin networks are not too clear, the mechanical tests alone are not sufficient to draw a more specific model. Figure 6B illustrates the deformation status of the cell wall after the first loading cycle. The cell walls with pectin alteration—lower amount of pectin and ion-mediated cross-links—show larger plastic deformation.
|
The larger unfolding and further alignment of the xyloglucan chains would result in a stiffening of the cell wall, which is consistent with theories on polymer network processing (Ward, 1997). Hence, the higher stiffening observed in the hypocotyls with pectin alterations may be explained by a lower pectin rigidity, which allows more unfolding and xyloglucan chain straightening.
An alternative hypothesis or a parallel mechanism could be that the pectin may contribute directly to the stiffness of the cellulose–matrix composite. As shown by Zykwinska et al. (2005), neutral sugar side chains of pectin are able to bind in vitro to the cellulose fibril surface. According to Proseus and Boyer (2007), the number of calcium-mediated cross-links determines the stiffness and strength of pectin, which influences the cell wall deformability during growth. Therefore, pectin alterations may also affect the capacity of pectin in tethering cellulose fibrils. However, Cleland and Rayle (1977) could not find a direct influence of calcium ion concentrations on the stiffness of cell walls.
Our study showed that by changes in the pectin network, the hypocotyl stiffness is far more affected than its ultimate stress. Thus, it seems unlikely that the cell wall properties are strongly dependent on a cellulose tethering function of pectin, although failure of the cell wall is rather controlled by the bonding pattern of the cellulose–xyloglucan network.
The model of the xyloglucan and pectin interactions in some ways resembles the mechanical effect of unfolding of macromolecules seen for many biological systems (e.g. DNA, titin, etc.), which has been investigated by many authors both experimentally and theoretically (e.g. Rief et al., 1997; Lu et al., 1998). However, in contrast to ‘sticky chain models’ (Jäger, 2001) exhibiting sacrificial bonds that inter-connect a folded chain, here, it is probably the rigidity of the embedding medium that may control the unfolding process. The proposed model, but also a model for a direct tethering of cellulose fibrils by pectin, point to an interesting analogy to the mechanics of bone of which the high toughness has been explained by a hidden length mechanism and sacrificial bonds (Fantner et al., 2005; Gupta et al., 2007). As in bone, ion-mediated bonds are likely to control the pectin rigidity either by Ca2+ ions in homogalacturonan or by borate in RG II and thereby influence the cell wall deformability.
In consequence, by ion-mediated links in pectin, plants would not only be able to modulate cell wall structure, permeability, swelling ability, and porosity (Zehirov and Georgiev, 2003; Jarvis, 1992; Zwieniecki et al., 2001; Fleischer et al., 1999), but also influence the plastic deformability of the cell wall at small strains. This finding may have implications on the current understanding of the underlying mechanisms that facilitate cell wall elongation during cell growth (reviewed by Cosgrove, 2005). Proseus and Boyer (2007) showed that the growth rate of Chara could be controlled by the deformability and strength of pectin adjusted by the number of calcium-mediated cross-links. In addition, our model is consistent with a theory that cells might be able to initiate cell wall loosening by regulating the ion-mediated cross-linking in the pectin. The advantage in the process of cell wall elongation driven by turgor pressure would be that it allows increasing deformability of the cell wall without severely affecting its strength.
|
METHODS |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
Arabidopsis thaliana (L.) Heynh. hypocotyls of wild-type (Col-0), mur1, mur2, and qua2 were utilized for this study. Mur1 is defective in the GDP-D-mannose-4,6-dehydratase (Bonin et al., 1997), leading to an altered structure of both xyloglucan and rhamnogalacturonan II (RG II) (Reiter et al., 1993), while mur2 shows a specifically altered xyloglucan structure particularly in xyloglucan fucosyltransferase (Vanzin et al., 2002). Qua2 possesses 50% less homogalacturonan in comparison to wild-type (Mouille et al., 2007). Sterilized seeds were plated in 8.8 g l–1 Murashige and Skoog basal medium in 0.8% agar, incubated in the dark at 22°C for 4 d in terms of the mur mutants and 6 d in terms of qua2. Hypocotyls of Col-0 were tested after both growth periods. Tests on older qua2 hypocotyls were necessary because the 4-day-old hypocotyls were very short and showed yielding at stress levels that were too low for running consistent cyclic loading tests.
Wild-type and the mutant hypocotyls were individually glued onto foliar frames, fixing them by a stepwise combination of rapid cyanoacrylate adhesive and Glass Ionomer Luting Cement (3M ESPE KetacTM Cem µ). The free length of the hypocotyl was located between the basal and the middle part. The foliar frames were specifically designed to be mounted onto a microtensile apparatus via pin-hole assembly. This is of crucial relevance, since the highly sensitive load cell with a maximum capacity of 500 mN forbids other mechanical clamping. The microtensile apparatus consists of a linear table driven by a step motor that allows feed rates of between 0.5 and 30 µm s–1. Black markers on the foliar frame allow following elongation more precisely compared to the machine path of the testing device (Burgert et al., 2003).
Hypocotyls were tested at room temperature at strain rates of either 10 or 15 µm s–1; the gauge lengths of the 4-day-old hypocotyls were, on average: Col-0 2.5 mm; mur1 2.2 mm; mur2 2.4 mm; the gauge lengths of the 6-day-old hypocotyls were, on average: Col-0 2.6 mm; qua2 2.7 mm. The duration of a simple loading test was 30 s; cyclic loading tests until completing the third cycles took 60 s. Vapor was constantly applied to the specimens to inhibit sample drying during the measurement. In the successive loading–unloading cycles, hypocotyls were subjected to several cycles before the sample was stressed until failure.
For transferring force–elongation curves into stress–strain curves, the cross-sections of the hypocotyls were calculated on the basis of diameter measurements under the microscope, assuming a circular outline of the hypocotyls. From the stress–strain curve generated in the standard tensile tests (see Figure 1A), sample stiffness (slope of the curve at a linear segment) and the ultimate stress level (curve's peak) of the hypocotyls were derived. In terms of cyclic loading (see Figure 1B), stiffness was determined in the upward loading phase for the first three cycles.
|
Turgor Pressure Determination |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
Turgor pressure was determined by the difference of water potential and osmotic pressure. Both parameters were measured with a C-52 sample chamber (Wescor, Logan, UT) in a sample holder with a diameter of 9.5 mm and a depth of 4.5 mm. The psychrometer was read by an automated datalogger (Wescor, Logan, UT). The system was calibrated with a sodium chloride solution (1000 mmol kg–1) used as a water potential standard of 2500 kPa (25°C). Psychrometers were allowed to equilibrate under each set of conditions. Cooling time to initiate condensation on the psychrometric junction was 30 s and the temperature depression from evaporative cooling was measured 30 s after active cooling ceased for a period of 30 s. The measured values were averaged. For the determination of the water potential, 15 seedlings were measured after at least 20 h of equilibration in one sample chamber.
The osmotic pressure was determined as described by Kutschera (1991). The hypocotyls were excised from the seedlings. Thereafter, they were frozen in liquid nitrogen, homogenized, and centrifuged for 2 min at 10 000 rpm. After centrifugation, the osmotic concentration of the supernatant was measured after an equilibration time of at least 2 h in the sample chamber.
Statistical Evaluation
Statistical evaluation of the mechanical behavior of the different types of hypocotyls was performed by t-tests at 
= 0.05, = 0.01, and = 0.001 confidence levels.
|
FUNDING |
|---|
|
TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSTurgor Pressure DeterminationFUNDING |
|---|
The financial support from the Max Planck Society and the EU grant 028974, project CASPIC, is gratefully acknowledged.
|
Acknowledgements |
|---|
We would like to thank Norma Funke, MPI-MP, as well as Annemarie Martins, Petra Leibner, and Susann Weichold, MPI-KG, for excellent technical support. No conflict of interest declared.
-
Baskin TI. Anisotropic expansion of the plant cell wall. Annu. Rev. Cell Dev. Biol. (2005) 21:203–222.[CrossRef][Web of Science][Medline]
Bonin CP, Potter I, Vanzin GF, Reiter W-D. The MUR1 gene of Arabidopsis thaliana encodes an isoform of GDP-D-mannose-4,6-dehydratase, catalyzing the first step in the de novo synthesis of GDP-L-fucose. Proc. Natl Acad. Sci. U S A (1997) 94:2085–2090.
Boyd RH, Phillips PJ. The Science of Polymer Molecules (1993) Cambridge: Cambridge University Press.
Burgert I, Fratzl P. Plants control the properties and actuation of their organs through the orientation of cellulose fibrils in their cell walls. Integrative and Comparative Biology (2009) 49:69–79.
Burgert I, Frühmann K, Keckes J, Fratzl P, Stanzl-Tschegg SE. Microtensile testing of wood fibers combined with video extensometry for efficient strain detection. Holzforschung (2003) 57:661–664.[CrossRef][Web of Science]
Carpita NC, Gibeaut DM. Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant J (1993) 3:1–30.[CrossRef][Web of Science][Medline]
Cavalier DM, et al. Disruption of two Arabidopsis thaliana xylosyltransferase genes results in plants deficient in xyloglucan, a major primary cell wall component. Plant Cell. (2008) 20:1519–1537.
Cleland RE. Extensibility of isolated cell walls: measurement and changes during cell elongation. Planta (1967) 74:197–209.[CrossRef][Web of Science]
Cleland RE. The Instron technique as a measure of immediate-past wall extensibility. Planta (1984) 160:514–520.[CrossRef][Web of Science]
Cleland RE, Rayle DL. Reevaluation of the effect of calcium ions on auxin-induced elongation. Plant Physiol (1977) 60:709–712.
Cosgrove DJ. Mechanism of rapid suppression of cell expansion in cucumber hypocotyls after blue-light irradiation. Planta (1988) 176:109–116.[CrossRef][Web of Science][Medline]
Cosgrove DJ. Characterization of long-term extension of isolated cell walls from growing cucumber hypocotyls. Planta (1989) 177:121–130.[CrossRef][Web of Science][Medline]
Cosgrove DJ. Expansive growth of plant cell walls. Plant Physiol. Biochem (2000) 38:109–124.
Cosgrove DJ. Growth of the plant cell wall. Nature Reviews—Molecular Cell Biology (2005) 6:850–861.[CrossRef]
Cosgrove DJ, Van Volkenburgh E, Cleland RE. Stress relaxation of cell walls and the yield threshold for growth. Planta (1984) 162:46–54.[CrossRef][Web of Science][Medline]
Creelman RA, Mullet JE. Oligosaccharins, brasshinolides and jasmonates: nontraditional regulators of plant growth, and development and gene expression. Plant Cell. (1997) 9:1211–1223.[CrossRef][Web of Science][Medline]
Eckardt NA. Cellulose synthesis takes the CesA train. Plant Cell. (2003) 15:1685–1688.
Fantner GE, et al. Sacrificial bonds and hidden length dissipate energy as mineralized fibrils separate during bone fracture. Nature Materials (2005) 4:612–616.[CrossRef][Web of Science][Medline]
Fleischer A, O'Neill MA, Ehwald R. The pore size of non-graminaceous plant cell walls is rapidly decreased by borate ester cross-linking of the pectic polysaccharide rhamnogalacturonan II. Plant Physiol (1999) 121:829–838.
Fratzl P, Burgert I, Gupta HS. On the role of interface polymers for the mechanics of natural polymeric composites. Phys. Chem. Chem. Phys. (2004) 6:5575–5579.[CrossRef][Web of Science]
Fry SC. Cellulases, hemicelluloses and auxin-stimulated growth: a possible relationship. Physiol. Plant (1989a) 75:532–536.
Fry SC. The structure and functions of xyloglucan. J. Exp. Bot (1989b) 40:1–11.
Gupta HS, Fratzl P, Kerschnitzki M, Benecke G, Wagermaier W, Kirchner HOK. Evidence for an elementary process in bone plasticity with an activation enthalpy of 1eV. J. Royal Society Interface (2007) 4:277–282.[CrossRef]
Ha M-A, Apperley DC, Jarvis MC. Molecular rigidity in dry and hydrated onion cell walls. Plant Physiol (1997) 115:593–598.[Abstract]
Hayashi T. Xyloglucans in the primary cell wall. Annu. Rev. Plant Physiol. Plant Mol. Biol. (1989) 40:139–168.[CrossRef]
Jäger I. The ‘sticky chain’: a kinetic model for the deformation of biological macromolecules. Biophysical J (2001) 81:1897–1906.[Web of Science][Medline]
Jarvis MC. Control of thickness of collenchyma cell walls by pectins. Planta (1992) 187:218–220.[Web of Science]
Keegstra K, Talmadge KW, Bauer WD, Albersheim P. The structure of the plant cell walls. III. A model of the walls of suspension-cultured sycamore cells based on the interconnections of the macromolecular components. Plant Physiol (1973) 51:188–196.
Kerstens S, Decraemer WF, Verbelen JP. Cell walls at the plant surface behave mechanically like fiber-reinforced composite materials. Plant Physiol (2001) 127:381–385.
Kobayashi M, Matoh T, Azuma J. Two chains of rhamnogalacturonan II are cross-linked by borate–diol ester bonds in higher plant cell walls. Plant Physiol (1996) 110:1017–1020.[Abstract]
Köhler L, Spatz H-CH. Micromechanics of plant tissues beyond the linear–elastic range. Planta (2002) 215:33–40.[CrossRef][Web of Science][Medline]
Kutschera U. Osmotic relations during elongation growth in hypocotyls of Helianthus annuus L. Planta (1991) 184:61–66.[Web of Science]
Lu H, Isralewitz B, Krammer A, Vogel V, Schulten K. Unfolding of titin immunoglobulin domains by steered molecular dynamics simulation. Biophysical J (1998) 75:662–671.[Web of Science][Medline]
Marga F, Grandbois M, Cosgrove DJ, Baskin TI. Cell wall extension results in the coordinate separation of parallel microfibrils: evidence from scanning electron microscopy and atomic force microscopy. Plant J (2005) 43:181–190.[CrossRef][Web of Science][Medline]
McCann MC, Roberts K. Architecture of the primary cell wall. In: The Cytoskeletal Basis of Plant Growth and Form—Lloyd CW, ed. (1991) New York: Academic Press. 109–129.
McCann MC, Wells B, Roberts K. Direct visualization of cross-links in the primary cell wall. J. Cell Sci. (1990) 96:323–334.
McCann MC, Wells B, Roberts K. Complexity in the spatial localization and length distribution of plant cell-wall matrix polysaccharides. J. Microscopy (1992) 166:123–136.[Web of Science]
Mouille G, et al. Homogalacturonan synthesis in Arabidopsis thaliana requires a Golgi-localized protein with a putative methyltransferase domain. Plant J (2007) 50:605–614.[CrossRef][Web of Science][Medline]
Passioura JB. The physical chemistry of the primary cell wall: implications for the control of expansion rate. J. Exp. Bot (1994) 45:1675–1682.[Web of Science]
Passioura JB, Fry SC. Turgor and cell expansion: beyond the Lockhart equation. Aust. J. Plant Physiol (1992) 19:565–576.
Pauly M, Albersheim P, Darvill A, York WS. Molecular domains of the cellulose/xyloglucan network in the cell walls of higher plants. Plant J (1999) 20:629–639.[CrossRef][Web of Science][Medline]
Peña MJ, Ryden P, Madson M, Smith AC, Carpita NC. The galactose residues of xyloglucan are essential to maintain mechanical strength of primary cell walls in Arabidopsis during growth. Plant Physiol (2004) 134:443–451.
Preston RD. The Physical Biology of Plant Cell Walls (1974) London: Chapman and Hall.
Proseus TE, Boyer JS. Tension required for pectate chemistry to control growth in Chara corallina. J. Exp. Bot (2007) 58:4283–4292.
Ralet MC, Crépeau MJ, Lefèbvre J, Mouille G, Höfte H, Thibault JF. Reduced number of homogalacturonan domains in pectins of an Arabidopsis mutant enhances the flexibility of the polymer. Biomacromolecules (2008) 9:1454–1460.[CrossRef][Web of Science][Medline]
Reiter WD, Chapple CCS, Somerville CR. Altered growth and cell-walls in a fucose-deficient mutant of Arabidopsis. Science (1993) 261:1032–1035.
Reiterer A, Lichtenegger H, Tschegg S, Fratzl P. Experimental evidence for a mechanical function of the cellulose microfibril angle in wood cell walls. Phil. Mag. A. (1999) 79:2173–2184.
Richmond PA, Metraux J-P, Taiz L. Cell expansion patterns and directionality of wall mechanical properties in Nitella. Plant Physiol (1980) 65:211–217.
Rief M, Gautel M, Oesterhelt F, Fernandez JM, Gaub HE. Reversible unfolding of individual titin immunoglobulin domains by AFM. Science (1997) 276:1109–1112.
Rose JKC, Bennett AB. Cooperative disassembly of the cellulose-xyloglucan network of plant cell walls: parallels between cell expansion and fruit ripening. Trends Plant Sci. (1999) 4:176–183.[CrossRef][Web of Science][Medline]
Ryden P, Sugimoto-Shirasu K, Smith AC, Findlay K, Reiter W-D, McCann M. Tensile properties of Arabidopsis cell walls depend on both a xylogucan cross-linked microfibrillar network and rhamnogalacturonan II-borate complexes. Plant Physiol (2003) 132:1033–1040.
Schindler TM. The new view of the primary cell wall. Z. Pflanzenernähr. Bodenk (1998) 161:499–508.
Taiz L. Plant cell wall expansion: regulation of cell wall mechanical properties. Annu. Rev. Plant Physiol (1984) 35:585–657.[CrossRef][Web of Science]
Talbott LD, Ray PM. Molecular size and separability features of pea cell wall polysaccharides: implications for models of primary wall structure. Plant Physiol (1992) 98:357–368.
Thompson JE, Fry SC. Evidence for covalent linkage between xyloglucan and acidic pectins in suspension-cultured rose cells. Planta (2000) 211:275–286.[CrossRef][Web of Science][Medline]
Vanzin GF, Madson M, Carpita NC, Raikhel NV, Keegstra K, Reiter W-D. The mur2 mutant of Arabidopsis thaliana lacks fucosylated xyloglucan because of a lesion in fucosyltransferase AtFUT1. Proc. Natl Acad. Sci. U S A (2002) 99:3340–3345.
Veytsman BA, Cosgrove DJ. A model of cell wall expansion based on thermodynamics of polymer networks. Biophys. J. (1998) 75:2240–2250.[Web of Science][Medline]
Vorwerk S, Somerville S, Somerville C. The role of plant cell wall polysaccharide composition in disease resistance. Trends Plant Sci. (2004) 9:203–209.[CrossRef][Web of Science][Medline]
Ward IM. Structure and properties of oriented polymers (1997) London: Chapman and Hall.
Whitney SEC, Gothard MGE, Mitchell JT, Gidley MJ. Roles of cellulose and xyloglucan in determining the mechanical properties of primary plant cell walls. Plant Physiol (1999) 121:657–663.
Zehirov GT, Georgiev GI. Effects of boron starvation on the apoplastic and total solute concentrations influencing nodule growth and acetylene reduction rate. Bulg. J. Plant Physiol (2003) 367–373.
Zwieniecki MA, Melcher PJ, Holbrook NM. Hydrogel control of xylem hydraulic resistance in plants. Science (2001) 291:1059–1062.
Zykwinska AW, Ralet MC, Garnier CD, Thobault JFJ. Evidence for in vitro binding of pectin side chains to cellulose. Plant Physiol (2005) 139:397–407.
CiteULike 
Connotea 
Del.icio.us What's this?
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||