JXB Advance Access originally published online on February 13, 2004
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Journal of Experimental Botany, Vol. 55, No. 397, pp. 571-583, March 1, 2004
© 2004 Oxford University Press
Cell and Molecular Biology, Biochemistry and Molecular Physiology |
Solid-state 13C-NMR spectroscopy shows that the xyloglucans in the primary cell walls of mung bean (Vigna radiata L.) occur in different domains: a new model for xyloglucan–cellulose interactions in the cell wall
Received 24 June 2003; Accepted 5 November 2003
1 School of Biological Sciences, The University of Auckland, Private Bag 92019, Auckland, New Zealand
2 Food Science Programmes, The University of Auckland, Private Bag 92019, Auckland, New Zealand
3 Industrial Research Limited, PO Box 31-310, Lower Hutt, New Zealand
* To whom correspondence should be addressed. Fax: +64 9 373 7416. E-mail: p.harris{at}auckland.ac.nz
Abbreviations: CP/MAS, cross-polarization/magic-angle spinning; GalUA, galacturonic acid; NMR, nuclear magnetic resonance; PSRE, proton spin relaxation editing; PDMS, polydimethylsiloxane; SE-PSRE, spin echo-proton spin relaxation editing; SPE, single pulse excitation; TEM, transmission electron microscopy; XG, xyloglucans; XET, xyloglucan endotransglucosylase.
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Abstract |
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Xyloglucans (XG) with different mobilities were identified in the primary cell walls of mung beans (Vigna radiata L.) by solid-state 13C-NMR spectroscopy. To improve the signal:noise ratios compared with unlabelled controls, Glc labelled at either C-1 or C-4 with 13C-isotope was incorporated into the cell-wall polysaccharides of mung bean hypocotyls. Using cell walls from seedlings labelled with D-[1-13C]glucose and, by exploiting the differences in rotating-frame and spin-spin proton relaxation, a small signal was detected which was assigned to Xyl of XGs with rigid glucan backbones. After labelling seedlings with D-[4-13C]glucose and using a novel combination of spin-echo spectroscopy with proton spin relaxation-editing, signals were detected that had 13C-spin relaxations and chemical shifts which were assigned to partly-rigid XGs surrounded by mobile non-cellulosic polysaccharides. Although quantification of these two mobility types of XG was difficult, the results indicated that the partly-rigid XGs were predominant in the cell walls. The results lend support to the postulated new cell-wall models in which only a small proportion of the total surface area of the cellulose microfibrils has XG adsorbed on to it. In these new models, the partly-rigid XGs form cross-links between adjacent cellulose microfibrils and/or between cellulose microfibrils and other non-cellulosic polysaccharides, such as pectic polysaccharides.
Key words: Cellulose microfibrils, cell-wall models, primary cell wall, solid-state 13C NMR, Vigna radiata, xyloglucan.
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Introduction |
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Primary cell walls of dicotyledons contain cellulose microfibrils embedded in a matrix of non-cellulosic polysaccharides (Bacic et al., 1988). Models of the architecture of dicotyledon primary cell walls commonly assume that xyloglucans (XGs) are adsorbed on to and coat the cellulose microfibrils. In these models, reviewed by Cosgrove (2001), the XGs cross-link adjacent microfibrils (Fig. 1A), or cross-link microfibrils and other matrix molecules, for example, pectic polysaccharides (Fig. 1B). Such cross-links will constrain the movements of the microfibrils and must be disrupted to allow cell-wall expansion. This expansion is thought to occur either by expansin detaching XG adsorbed on to cellulose (Cosgrove, 2001) and/or by xyloglucan endotransglycosylase (XET) breaking and rejoining cross-linking XG molecules (Thompson and Fry, 2001).
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Several lines of evidence indicate that XGs are associated with cellulose microfibrils in primary cell walls. First, cross-links, possibly XG, between adjacent cellulose microfibrils have been observed in primary cell walls by TEM after preparing them using fast-freeze, deep-etch, and rotary shadowing (McCann et al., 1990; Fujino et al., 2000). Second, immunogold labelling using antibodies against XG showed that XG molecules were closely associated with cellulose microfibrils in the primary walls of suspension-cultured cells of Rubus sp. (Joseleau et al., 1992) and in suspension-cultured cells of tobacco (Nicotiana tabacum) (Sabba et al., 1999). In the latter cells, no labelling was found in the middle lamella where no cellulose could be detected using cellulase conjugated to colloidal gold. Third, the cell walls of suspension-cultured tobacco grown in the presence of 2,6-dichlorobenzonitrile, an inhibitor of cellulose synthesis, contained no cellulose and their XGs could be extracted easily without the use of alkali (Sabba et al., 1999). In vitro systems have also been used to show that XG can associate with cellulose. For example, if a solution of XG is passed down a column of cellulose, some of the XG associates with the cellulose (Mishima et al., 1998). Moreover, if the cellulose-producing bacterium Acetobacter xylinus is grown in the presence of XG, the latter associates with the cellulose ribbons produced (Whitney et al., 1995).
However, none of the above studies provide evidence for the cellulose microfibrils in primary cell walls being coated with XG. Indeed, it is possible that only a relatively short length of each XG molecule is actually adsorbed on to a cellulose microfibril, and only a small proportion of the total surface area of the cellulose microfibrils may have XG adsorbed on to it.
Solid-state 13C nuclear magnetic resonance (NMR) spectroscopy is a technique that can be used to identify polymers with different mobilities. These polymers may be cell-wall polysaccharides, such as xyloglucans, which are structurally similar, but have different mobilities because of their locations and interactions with other molecules within the cell wall. For example, the glucan backbones of the XG molecules, or parts of molecules, adsorbed on to the surface of cellulose microfibrils in muro would be predicted to adopt a rigid, flattened conformation, rather than the twisted-backbone conformation of free XG (Levy et al., 1997). These two different conformations can potentially be detected by solid-state 13C NMR spectroscopy (Horii et al., 1984; Jarvis, 1994). This technique has the added advantage that cell walls that have never been dried can be examined. By studying proton or 13C spin relaxation properties, it should be possible to detect differences in spin relaxation and in chemical shift between xyloglucans of different mobilities. However, solid-state 13C NMR studies of primary cell walls of six dicotyledon species have provided no conclusive evidence for rigid XG molecules (Newman et al., 1994, 1996; Koh et al., 1997; Smith et al., 1998a; Thimm et al., 2002). This may be because there are no rigid XG molecules in these cell walls, or because the amount is insufficient to be detected by solid-state 13C NMR spectroscopy using the natural abundance of 13C-isotope (1.1% of all carbon atoms).
In the present study, the proportion of 13C-isotope in the cell-wall polysaccharides was increased to improve the signal-to-noise ratio and thereby improve the sensitivity of the solid-state 13C-NMR spectroscopy. Etiolated hypocotyls of mung beans (Vigna radiata L.) were used as a source of 13C-enriched primary cell walls for these experiments as these seedlings are easy to incubate with 13C-labelled Glc, the hypocotyls consist mostly of cells with primary walls, and the XG in these cell walls has been characterized (Kato and Matsuda, 1976, 1980a, b). Plant tissues take up Glc and convert it to nucleoside diphosphate sugars, the precursors of the cell-wall polysaccharides (Fry, 1985). By using Glc labelled with 13C either at C-1 or at C-4, the monosaccharides in the cell-wall polysaccharides of mung bean seedlings can be labelled specifically at C-1 or C-4.
The mobility of XGs in the cell wall was investigated using solid-state cross-polarization/magic-angle spinning (CP/MAS) 13C-NMR spectroscopy in combination with proton spin relaxation editing (PSRE). This combination of techniques is useful in studying the mobilities of components in multi-component systems, such as the plant cell wall (Newman, 1999a; Tang and Hills, 2003). Two series of experiments were conducted. In the first series, cell walls from seedlings incubated with D-[1-13C]glucose were used to detect XGs in the cell-wall domain containing rigid polysaccharides. In the second series of experiments, the seedlings were incubated with D-[4-13C]glucose, and then a novel NMR technique was used on the cell walls that combined 13C spin-echo experiments with proton spin relaxation editing (SE-PSRE) to measure 13C spin-spin relaxation time constants (T2(C)). This technique was used to detect XGs in the cell-wall domain containing semi-rigid and mobile polysaccharides.
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Materials and methods |
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Chemicals
D-[1-13C]glucose (99%) and D-[4-13C]glucose (99%) were from Novachem Pty Ltd, South Yarra, Victoria, Australia.
Plant material
Seeds of mung bean (Vigna radiata L. cv. Berkin) were surface-sterilized in 30 g l–1 NaOCl for 20 min, soaked in sterile water in the dark for 7 h at 25 °C, and germinated on sterile, damp vermiculite in the dark at 25 °C.
Microscopy
Bright-field and UV-fluorescence microscopy was carried out on fresh transverse sections cut from midway along the hypocotyls as described by Carnachan and Harris (2000). Lignin was detected by the red colour given with phloroglucinol–HCl, by the blue autofluorescence in UV radiation when mounted in 0.1 M NH4OH, and by the green or blue staining produced with the polychromatic stain toluidine blue O (Carnachan and Harris, 2000). Starch was detected using I2 in KI.
Lignin was detected only in the annular and spiral thickenings of the few protoxylem vessels present in the hypocotyls. Small starch granules, detected in the layer of cortical parenchyma cells nearest the endodermis, were removed by filtration during cell-wall isolation.
Incubation of seedlings with 13C-labelled Glc
Batches of five seedlings with hypocotyls 3.5–4.5 cm long were selected, the primary roots were cut off, and the cut-ends placed in a solution of unlabelled Glc, D-[1-13C]glucose or D-[4-13C]glucose (7.5 mg Glc ml–1, 2 cm3). The seedlings were incubated (18 h at 25 °C) in the dark in a sealed glass container with the air saturated with water vapour. For each treatment, 15 batches of seedlings were used, making a total of 75 seedlings.
Isolation of cell walls
After incubation with 13C-labelled Glc or unlabelled Glc, the cotyledons were removed from the seedlings by cutting 0.5 cm below the cotyledons. All the steps in the isolation procedure were carried out at 4 °C. The hypocotyls were cut into pieces (0.5 cm long) and homogenized in 100 cm3 of 20 mM HEPES–KOH buffer (pH 6.7) containing 10 mM dithiothreitol, using a Polytron blender (Model PT10-35, Kinematica GmbH, Luzern, Switzerland) on full power (three times for 20 s). Further homogenization was done using a Tenbroeck ground-glass homogenizer (15 cm3; Kontes Glass Company, Vineland, NJ, USA). Breakage of the cells was monitored using bright-field microscopy after staining with Ponceau 2R, which stains cytoplasmic protein red ((Harris, 1983). The homogenate was centrifuged (250 g for 10 min), and the pellet washed three times by centrifugation with buffer (with dithiothreitol omitted), followed by six times with water. The pellet was resuspended, washed on to nylon mesh (pore size 11 µm), and the residue on the mesh washed with water (500 cm3). The residue (cell-wall preparation) was washed three times by centrifugation with 80% (v/v) ethanol and the final suspension was kept at 4 °C until NMR spectroscopy could be carried out.
An additional cell-wall preparation was obtained from the hypocotyls of seedlings incubated with D-[1-13C]glucose, but was not washed with 80% (v/v) ethanol. Instead, this preparation was frozen and dried under vacuum to a water content of approximately 40% (w/w) and NMR spectroscopy was carried out immediately. The exact water content of each sample was determined after completing the NMR spectroscopy by heating at 105 °C until dry (Table 1).
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NMR spectroscopy
Cell-wall preparations in 80% (v/v) ethanol were prepared for NMR by filtering portions of the suspensions onto a glass-fibre filter (GF/C, Whatman Scientific Ltd, Kent, UK) and part-drying in air to a water content of approximately 40% by weight. A sample was then packed in a 7 mm diameter, cylindrical silicon nitride rotor, and retained with Vespel end caps. An internal standard of PDMS (polydimethylsiloxane) was added to all samples. The rotor was spun at 4 kHz in a Doty Scientific magic-angle spinning probe for 13C NMR spectroscopy at 50.3 MHz on a Varian Inova-200 spectrometer.
In the cross-polarization (CP) NMR experiments, each 6 µs 90° proton preparation pulse was followed by a 1 ms CP contact time, 51 ms of data acquisition and a recovery delay of 1.0 s before the sequence was repeated. Newman (1987) showed that a 6 µs proton preparation pulse is sufficient to excite protons in the rigid domain. Preliminary T1(H) experiments indicated that a 1.0 s delay was adequate for the recovery of proton magnetization, as was also shown in the experiments on other cell walls (Newman et al., 1994, 1996). The proton transmitter output was increased to its maximum during data acquisition, corresponding to a precession frequency of 59 kHz.
Proton relaxation experiments
Proton rotating-frame spin relaxation with time constant T1
(H), and proton spin-spin relaxation with time constant T2(H) were characterized by inserting relaxation intervals of duration t1 or t2, respectively, between the proton preparation pulse and the CP contact time. The values for t1 and t2 were chosen to be within the ranges of values for T1(H) and T2(H), respectively, in order to optimize signal:noise ratios in separate subspectra (Newman and Hemmingson, 1990). Protons were spin-locked during t1, but the proton transmitter was switched off during t2.
PSRE NMR subspectra were generated by combining spectra labelled S, S' and S", where S was obtained by the normal cross-polarization pulse sequence, S' with t1=4 ms and S" with t2=15 µs. The principles behind PSRE NMR have been described elsewhere (Newman, 1999a; Tang et al., 2000). In the simplest case, a spectrum S is the sum of subspectra A and B from two distinct types of domains. A partly-relaxed spectrum S' is then
S'=faA+fbB(1)
where fa and fb are signal suppression factors. The subspectra can then be separated by computing:
A=kS+k'S'(2a)
B=(1–k)S–k'S'(2b)
where
k=fb/(fb–fa)(3a)
k'= –1/(fb–fa)(3b)
In the present context, A and B contain signals from the cellulose microfibrils and the non-cellulosic polysaccharides, respectively.
Two NMR signals were selected for proton rotating-frame relaxation experiments. The signal at 89 ppm (assigned to C-4 of cellulose) was selected as representative of cellulose crystallites, appearing at a chemical shift for which there is little risk of overlap with signals from other polysaccharides. A signal at 69 ppm (assigned to C-2, C-3, and C-5 of GalUA residues in pectic homogalacturonan (Jarvis and Apperley, 1995; Sinitsya et al., 1998)) was selected as representative of mobile polysaccharides. Because the T1(H) relaxation for the 69 ppm signal was non-exponential, it was not possible to achieve total elimination of signals from non-cellulosic polysaccharides without also suppressing signals from cellulose, so linear combinations of S and S' were generated to enhance signal suppression. The suppression factors and the resulting linear combinations used to generate the PSRE subspectra from the experimental PSRE spectra are shown in Table 1. The slight differences in these values may reflect differences in the water content of the cell-wall preparations (Hediger et al., 1999; Tang et al., 2000). The separation was successful in that the signals assigned to cellulose and non-cellulosic polysaccharides appeared in subspectra A and B, respectively.
The editing process was repeated for T2(H) relaxation for the cell walls from seedlings labelled with D-[1-13C]glucose. This spin relaxation process suppressed cellulose signals more than the other signals, but did not eliminate them entirely. Linear combinations were again generated to enhance the amount of suppression (Table 1). The separation was successful in that signals assigned to cellulose and non-cellulosic polysaccharides appeared in subspectra A and B, respectively.
The combinations of spectra were generated off-line using software written to run under MS-DOS. Spectra were then processed using MS Excel.
SE-PSRE experiments
T2(C) relaxation was characterized by a spin-echo sequence in which a delay of duration t2 was inserted between the CP contact time and commencement of data acquisition (Newman et al., 1996). A 180° refocusing pulse was applied halfway through t2, and values of t2 were chosen so that 0.5t2 was always a multiple of the rotor rotation period, and protons were decoupled with an attenuated power output corresponding to a precession frequency of 43 kHz throughout t2. PSRE NMR subspectra were generated for the T2(C) spectrum using the S and S' values calculated as described above.
Single pulse excitation/magic-angle spinning (SPE/MAS) experiments
The single pulse excitation/magic-angle spinning (SPE/MAS) spectrum was obtained with a pulse sequence in which each 6 µs 13C excitation pulse was followed by 51 ms of data acquisition time and a 1.0 s recovery delay (Smith et al., 1998b). The 1.0 s delay was used to maximize the response from mobile components and minimize the response from rigid components of the cell walls (Smith et al., 1998b). The proton decoupler transmitter power was increased to provide radiofrequency field strengths of 59 kHz during data acquisition.
Cellulose content of the an unlabelled cell-wall preparation
The cellulose content of the cell-wall preparation was determined by the method of Brendel et al. (2000).
Neutral monosaccharide composition of an unlabelled cell-wall preparation
An unlabelled cell-wall preparation was hydrolysed with sulphuric acid, and the released neutral monosaccharides reduced and acetylated to produce alditol acetates, which were quantified by gas chromatography (Smith and Harris, 1995).
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CP/MAS 13C NMR spectroscopy of cell-wall preparations from hypocotyls of seedlings incubated with 13C-labelled Glc
CP/MAS solid-state 13C NMR spectra of cell-wall preparations from seedlings incubated with D-[1-13C]glucose or D-[4-13C]glucose showed increases in signal strengths at C-1 or C-4, respectively, compared with the spectrum of a cell-wall preparation from seedlings incubated with unlabelled Glc (Fig. 2). Measurements of peak areas over the chemical-shift range assigned to C-1 or C-4 (93–108 ppm and 80–93 ppm, respectively) showed that isotopic enrichment for cell walls obtained from seedlings incubated with D-[1-13C]glucose was approximately 2.5 and that for cell walls obtained from seedlings incubated with D-[4-13C]glucose was approximately 2.3. These enrichment factors are averages for all polysaccharides responding to the cross-polarization pulse sequences. Moreover, they are only approximations as they do not take into account the efficiency of the magnetization transfer in the CP process, where molecules will have different mobility and strength of H-C dipolar interactions (Hediger et al., 1999). The signals include the dominant signals assigned to the rigid cellulose crystallites and the broader underlying signals assigned to non-cellulosic polysaccharides.
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PSRE of CP/MAS NMR spectra obtained for a cell-wall preparation from hypocotyls of seedlings incubated with D-[1-13C]glucose
Proton spin relaxation experiments on a cell-wall preparation from seedlings incubated with D-[1-13C]glucose were used to search for NMR signals with proton relaxation properties appropriate for rigid XG. PSRE involved exploiting the differences between the normal CP/MAS spectrum and experimental spectra obtained with proton spin relaxation (Fig. 3). The CP/MAS spectrum obtained for this cell-wall preparation was then edited using the T1
(H) time constants into two subspectra (Fig. 4A): subspectrum A contained signals with long values of T1(H), assigned to the rigid cell-wall polysaccharides, mostly cellulose, and subspectrum B contained signals with short values of T1(H), assigned to polysaccharides more mobile than cellulose. When the CP/MAS spectrum was edited using the T2(H) time constants (Fig. 4B), subspectrum A contained signals with short values of T2(H), assigned to rigid polysaccharides, mostly cellulose, and subspectrum B contained signals with long values of T2(H), assigned to more mobile polysaccharides. Amplification of the signals in the C-1 portion of subspectrum A separated using T1(H) and using T2(H) both showed a small signal at 100.4 ppm (Fig. 5A). This signal can be assigned to Xyl in XG, based on chemical shifts reported for XG (Table 2). Full spin relaxation curves could not be analysed for this signal, since the initial portions of these curves were dominated by an overlapping signal at 101 ppm from GalA of pectic homogalacturonans. It is also possible that the signal at 100.4 ppm may contain a contribution from pectic homogalacturonans. Proton spin diffusion can account for the transfer of spin information over distances of nanometers during the 1 ms cross-polarization contact time of a typical solid-state 13C-NMR experiment (Zumbulyadis, 1983), so the values of T1(H) and T2(H) are averaged within a monosaccharide residue and between adjacent residues (Hediger et al., 1999). Therefore, the signal strength observed through cross-polarization will reflect the molecular ordering over dimensions several times larger than a Glc residue (Newman, 1992). The presence of the 100.4 ppm signal in subspectrum A indicates that the polysaccharide producing this signal, which was assigned to XG, is in the vicinity of another rigid polysaccharide, namely cellulose. This signal at 100.4 ppm was small and only just above the background and it could not be detected in the total CP/MAS spectrum or subspectrum A of cell walls from unlabelled seedlings (results not shown). However, it was reproducibly detected in two different cell-wall preparations (Fig. 5A, B). The small Xyl signal (at 100.4 ppm) detected in subspectrum A accounted for less than 1% of the total C-1 signals. Therefore, it must be stressed that it would be difficult to detect this signal without using isotopic labelling.
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A quantitative estimate of the proportion of the cell-wall XG that is rigid is difficult because of the low strength of the signal at 100.4 ppm, and because it is not known whether the 13C-label on either C-1 or C-4 divides equally among the cell-wall polysaccharides, including XGs and cellulose. In addition, the quantification does not take into account the efficiency of the magnetization transfer in the CP process, where molecules will have differing mobility and strength of H-C dipolar interactions (Hediger et al., 1999). However, it was considered important to place an upper limit on the signal strength and then use this upper limit to obtain an estimate of the proportion of rigid XG. First, the contribution of this signal at 100.4 ppm to the total signal in the C-1 region of the rigid domain of the cell wall was obtained by integrating over an arbitrary 2 ppm range centred on 100.4 ppm. This was then compared with the area of the cellulose C-1 signal at 105.5 ppm which showed that for every C-1 atom in Xyl there were 30–40 cellulose C-1 atoms; the lower value was obtained using the T1
(H) subspectrum (Fig. 4A) and the higher using the T2(H) subspectrum (Fig. 4B), with a mean ratio of 1:35. As the cellulose content of the cell-wall preparations was 41% (w/w) (cell-wall dry weight), the amount of Xyl in the rigid domain was 1.2% (w/w). Moreover, these cell-wall preparations contained 4.4% (w/w) Xyl, which originated principally from XG, although the cell-wall preparations probably also contained small proportions of heteroxylans (Bacic et al., 1988). If it is assumed that all the Xyl was from XG that had the monosaccharide composition reported by Kato and Matsuda (1980a) for mung bean hypocotyl XG (Glc:Xyl:Gal:Fuc=10:5.7:2.5:2.9 anhydro w/w), then the maximum XG content of the cell-wall preparations was 13.5% (w/w). Approximately 27% of the total wall XG will be in the rigid domain, accounting for 3.9% (w/w) of the cell-wall preparation.
Because the xylose residues of XGs are attached to the glucan backbone by (1
6)-linkages, it is possible that the xylose is able to rotate around this linkage and so be under-represented in the rigid domain. However, this was considered unlikely because models of rigid XGs show straight backbones with the xylose residues protruding on either side; such Xyl residues cannot rotate freely around the (16)-linkage (Levy et al., 1991). Moreover, the Xyl residue may appear more rigid than it actually is because of the averaging effects of proton spin diffusion from dominant, inherently rigid polysaccharides.
Rigid XG does not account for all the signals between 92 ppm and the main C-1 peak of cellulose at 105.5 ppm (Fig. 5). Small amounts of Man (2.8% w/w) were found in these cell-wall preparations and the small signal at 102 ppm may be from the C-1 of Man in a gluco- or a galactogluco-mannan. A signal at 102 ppm has previously been assigned to C-1 of Man in rigid galactomannans (Newman and Hemmingson, 1998) and in galactoglucomannans in the cell walls of Arabidopsis thaliana (Davies et al., 2002).
Proton spin diffusion averages the spin information over dimensions of nanometres (Zumbulyadis, 1983; Hediger et al., 1999). Therefore, if the Xyl in the side-chains of XG is present in the NMR subspectra of the rigid domain of the cell walls (Fig. 4A, B), the Glc signals from the glucan backbone of this XG should also be present in these subspectra. A shoulder at 107.5 ppm was found on the side of the main cellulose peak in both the total spectrum and in the subspectra of the rigid domain (Fig. 5), which may be from the Glc of XG with a rigid, flattened, glucan backbone. The flattened backbone of rigid XG is predicted to have two Glc residues per helical turn and is, therefore, similar in conformation to ß-glucan chains in crystalline cellulose (Levy et al., 1997). The flattening of a ß-glucan backbone is predicted to cause increased torsion in the linkage carbons, C-1 and C-4, compared with that in a ß-glucan backbone in a twisted conformation, resulting in different 13C NMR chemical shifts (Horii et al., 1984; Jarvis, 1994). Data for cellulose provide an estimate of the likely chemical-shift displacement for the Glc residues of rigid XG. For the twisted conformation of the backbone, the C-1 chemical shift is 102.7 ppm, obtained from a solution of cellulose oligosaccharides generated by partial acid hydrolysis (Gast et al., 1980). However, for the flat-backbone conformation of cellulose I and II, the C-1 chemical shifts are in the range 105 to 108 ppm (Hemmingson and Newman, 1995), so the chemical-shift displacements are in the range 2 to 5 ppm. Similar chemical-shift displacements for the Glc residues in XG would move the C-1 signal from 103 ppm (Table 2) to the range 105 to 108 ppm, which includes the shoulder found in these subspectra at 107.5 ppm. Because the shoulder at 107.5 ppm was poorly resolved, it was not possible to estimate its area, but it is clearly no more than a minor component of the cell wall. The assignment of the shoulder at 107.5 ppm to the Glc in the flattened backbone of rigid XG is reasonable as it is consistent with the small signal at 100.4 ppm, assigned to Xyl in the sidechain of XG.
However, the signal at 107.5 ppm could also be assigned to C-1 of cellulose II. Hemmingson and Newman (1995) assigned a signal at 106.7 ppm to C-1 of cellulose II. They also assigned signals at 88.2 ppm and 89.4 ppm to C-4 of cellulose II crystallite-interior molecules, coinciding with the area assigned to the interior chains of cellulose I, and a signal at 87 ppm to C-4 in cellulose II crystallite-surface chains. This signal at 87 ppm would fall on the boundary between signal areas assigned to interior and surface chains of cellulose I, but no signal was detected at 87 ppm, suggesting that cellulose II is absent.
To confirm that the polysaccharide associated with the signal at 107.5 ppm was indeed rigid, complete relaxation curves were constructed by varying the relaxation intervals t1 and t2 in the T1(H) and T2(H) NMR pulse sequences ((Tekely and Vignon, 1987a, b). These curves (Fig. 6) show the relaxation characteristics for the signal at 107.5 ppm compared with those for the signal at 89 ppm (assigned to C-4 of Glc in cellulose crystallite interior) and for the signal at 69 ppm (assigned to C-4 of GalUA in pectic homogalacturonans). The T1(H) relaxation curve for the signal at 107.5 ppm was similar to that obtained for crystalline cellulose (Fig. 6A). The T1(H)) time constants for the signal at 89 ppm and 107.5 ppm were also similar, with T1(H)=8.0 and 6.4 ms, respectively. A curve for the C-4 signal at 85 ppm was also exponential with T1(H)= 7.4 ms (data not shown). This signal at 85 ppm was assigned to cellulose on the microfibril surfaces. The relaxation curve for pectic homogalacturonan (Fig. 6A) was distinctly different, with rapid initial decay (T1(H)=1.3 ms) followed by a slower phase attributed to replenishment of magnetization through spin diffusion from cellulose to pectic homogalacturonan. Furthermore, the Gaussian shape of the T2(H) relaxation curve (Fig. 6B) for the signal at 107.5 ppm was typical of the response of rigid, solid matter (Cheung and Gerstein, 1981) and was, therefore, consistent with the assignment of this signal to a rigid component of the cell wall. The T2(H) time constants for the signal at 89 ppm and 107.5 ppm were similar, with T2(H)=10.2 and 9.5 µs, respectively. A curve for the C-4 signal at 85 ppm was also Gaussian with T2(H)=9.7 µs (data not shown). By contrast, the exponential shape of the T2(H) curve for the signal at 69 ppm (Fig. 6B) is typical of mobile material, with T2(H)=24 µs.
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To prevent microbial contamination, the cell-wall preparations were kept in 80% (v/v) ethanol until NMR experiments could be carried out. To confirm that the signal at 100.4 ppm was not a result of this exposure to 80% (v/v) ethanol, a cell-wall preparation that had not been exposed to 80% (v/v) ethanol was also examined. This cell-wall preparation was also obtained from seedlings incubated with D-[1-13C]glucose. No differences were found in the PSRE NMR spectra for these two cell-wall preparations. The same signals at 100.4, 107.5 and 102 ppm were found in the rigid domain of the PSRE subspectra (Fig. 5B).
SE-PSRE NMR of a cell-wall preparation from hypocotyls of seedlings incubated with D-[4-13C]glucose
A novel combination of spin-echo relaxation and rotating-frame experiments were used to detect partly-rigid XGs. Unlike T1(H) and T2(H), T2(C) is sensitive to the segmental motion of the polysaccharide at the site of the 13C nucleus and is insensitive to the motion of more distant polysaccharides (Rothwell and Waugh, 1981). Therefore, in these SE-PSRE NMR experiments, the T2(C) values for a polysaccharide will indicate the mobility of that particular polysaccharide, whereas the T1(H) and T2(H) values will reflect the mean molecular motion of the surrounding polysaccharides. Because the spin relaxation of the 13C is dominated by the motion of the relevant carbon atom, the T2(C) time constants of these XGs should be different from those of polysaccharides with different mobilities in the matrix of the cell wall. The same suppression factors, obtained from rotating-frame experiments (Fig. 7A) and used to separate the subspectra of rigid from partly rigid and mobile domains (Fig. 7B) in PSRE, were also used to edit the SE spectrum into subspectra A and B. Figure 8 shows the SE spectrum and PSRE subspectra for a pulse sequence with a relaxation interval of t2=5 ms. The T2(C) time constants shown in Table 3 were determined by repeating the SE-PSRE experiment for other values of t2. Signals at 101, 80, and 69 ppm, assigned to pectic homogalacturonan (Table 3), relaxed with T2(C) values of 3–4 ms. The 13C spins of these mobile polysaccharides had almost completely relaxed within 5 ms and, therefore, the pectic homogalacturonan signals were not detected in subspectrum B of Fig. 8.
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XGs with partly-rigid glucan backbones should have a long T2(C) time constant compared with mobile cell-wall polysaccharides. However, if the XGs are surrounded by mobile polysaccharides, they should have a relatively short T1
(H) time constant as the proton spin is influenced by the other polysaccharides in the surrounding matrix. Therefore, when the T2(C) spectrum is edited using the T1(H) time constants, it would be expected that signals from the backbone of such partly-rigid XGs surrounded by mobile polysaccharides would separate into subspectrum B, whereas signals assigned to cellulose would separate into subspectrum A. Using this SE-PSRE technique, two signals at 105 and 84 ppm were detected in subspectrum B and assigned to C-1 and C-4, respectively, of the Glc residues in partly-rigid XG (Fig. 8) (Braccini et al., 1995). Both of these signals relaxed with the same time constant (9 ms), which was shorter than those of the cellulose signals, but longer than the time constants of the pectic homogalacturonans signals (Table 3). Hence, the 105 and the 84 ppm signals in subspectrum B (Fig. 8) are assigned to partly-rigid XGs surrounded by more mobile polysaccharides. Compared with the C-4 Glc signal of XG in the solution state, the C-4 signal assigned to Glc in XG is displaced from 80 to 84 ppm. This displacement is consistent with the XG being partly rigid and can be attributed to a degree of order in the XG backbone compared with the twisted conformation of the backbone of XG in solution (Table 2).
Other NMR chemical shifts in subspectrum B (Fig. 8) can also be assigned to partly-rigid XG. The shifts at 62 and 74 ppm can be assigned primarily to C-6 and C-2, 3, 5 of the Glc residues of the XG backbone, respectively (Braccini et al., 1995). The weakness of the 62 ppm signal may be due to T2(C) relaxation promoted by rotation of the –CH2OH group (Rothwell and Waugh, 1981). Shifts at 72 and 75 ppm superimposed in this C-2, 3, 5 region of the spectrum may result from minor contributions from other polysaccharides.
Unlike the Xyl residues in rigid XGs which, as indicated above, are unable to rotate freely around the (16)-linkage (Levy et al., 1991), the Xyl residues in the partly-rigid XGs would be expected to rotate around the (16)-linkage as occurs in XG in solution. Thus, the absence of a signal from these Xyl residues at 100.4 ppm is consistent with the XG being in this partly-rigid state.
It is also possible that the signals at 105 and 84 ppm originate from disordered cellulose. However, previous studies have not detected such cellulose in subspectrum B of PSRE-NMR spectra obtained from angiosperm primary cell walls, unless these cell walls had first been treated with strong alkali (Davies et al., 2002). Nevertheless, other researchers have published evidence indicating the presence of disordered cellulose in such cell walls (Wickholm et al., 1998; Hediger et al., 2002).
As indicated above, quantification of XGs with different mobilities is difficult. However, to obtain an indication of the proportion of the total XG in the cell wall which is partly rigid, the areas of the C-4 signals for Glc residues of both cellulose and XG were extrapolated back to t2=0. The ratio of Glc in partly-rigid XG (Fig. 8) to Glc in cellulose was 1:5. Based on a maximum XG content of the mung bean cell-wall preparation of 13.5% and a cellulose content of 41% (w/w), all of the cell wall XG could be accounted for by the 84 ppm signal.
Single pulse excitation/magic-angle spinning (SPE/MAS) NMR of cell-wall preparations from hypocotyls incubated with D-[1-13C]glucose
The CP/MAS NMR spectra of the cell-wall preparations were dominated by signals assigned to cellulose and the broader, underlying signals were assigned to non- cellulosic polysaccharides, along with signals at 173, 54, and 22 ppm assigned to carboxylic acid, methoxyl, and acetyl carbons, respectively (Fig. 2). As cellulose makes up less than half the weight of the cell wall, a portion of the non-cellulosic material must have been too mobile to respond to the cross-polarization pulse sequence. Although SPE/MAS NMR can potentially detect all components in the cell-wall preparations (Tang et al., 1999), the highly mobile cell-wall components, which are less responsive to CP/MAS NMR (Foster et al., 1996; Smith et al., 1998b) are particularly responsive to SPE/MAS NMR. By using a short recovery delay in the SPE/MAS pulse sequence (1 s), signals which have long T1(C) relaxations, such as the rigid cellulose molecules, are not detected (Smith et al., 1998b; Tang et al., 1999). SPE/MAS is thus an excellent complementary technique to CP/MAS NMR (Tang and Hills, 2003) for investigating mobile polysaccharides in cell-wall preparations. The SPE/MAS NMR spectra of the cell-wall preparations were dominated by signals assigned to galactans of the pectic polysaccharides (results not shown), so it was difficult to distinguish signals that could be assigned to highly mobile XG. Hence, although the two CP/MAS techniques accounted independently for all the estimated XG in the cell-wall preparations, the possibility that some XG also exists in a highly mobile domain cannot be excluded.
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Using solid-state 13C-NMR, XGs of two different mobilities in the primary cell walls of mung beans have been identified: XGs with a rigid glucan backbone, which account for the smaller proportion of the total cell-wall XG, and XGs with a partly-rigid glucan backbone. The latter XGs were surrounded by mobile polysaccharides. It is also possible that there were small quantities of highly mobile XGs; however, interference from signals from mobile pectic polysaccharides precluded their detection. These results are consistent with the findings of Mackay et al. (1988) and Taylor et al. (1990), who, using solid-state 1H-NMR, found that the XGs in primary cell walls of common bean (Phaseolus vulgaris) had constrained molecular mobility. Pauly et al. (1999) also identified XGs in different cell-wall domains based on their solubility in XG-specific endoglucanase, concentrated alkali and cellulase, applied sequentially to the cell walls. From the results reported in the present paper, it is not possible to identify the locations of the two different mobility types of XGs relative to other components in the cell wall, it is possible to make predictions based on current models of the primary cell walls of dicotyledons.
The XG molecules with the rigid glucan backbone may have been adsorbed on to the surfaces of the cellulose microfibrils. XG molecules adsorbed in this way would be expected to have a rigid, flattened backbone and not the twisted backbone conformation of free XG (Levy et al., 1997). Despite the difficulties involved in quantifying the amounts of the different mobility types of XGs, approximations can be made about the proportion of total cellulose microfibril surface area that has rigid XGs adsorbed to it. This was done by first expanding the NMR signals assigned to C-4 of cellulose in PSRE subspectra separated using T1(H) and T2(H) to estimate the total surface area of the cellulose microfibrils available for adsorption. Evidence has recently been obtained that the cellulose microfibrils in angiosperm primary cell walls consist of only one cellulose crystallite (Davies and Harris, 2003). Thus, in the present study, it may be assumed that the cellulose crystallite accounts for the entire width of the cellulose microfibril. The integrated signal areas assigned to interior and surface cellulose in the cellulose crystallites (Newman, 1999b) for all three cell-wall preparations showed that 64±1.4% (using T1(H)) or 62.7±1.3% (using T2(H)) of the cellulose was on the surface of the crystallite. As the cell walls contained 41% (w/w) (cell-wall dry weight) cellulose, and 63% of this cellulose was on the surface of the microfibrils, then 26% (w/w) of the cell walls was surface cellulose. As it was estimated that the cell walls contained an upper limit of 3.9% (w/w) rigid XG with a Glc:Xyl ratio of 10:5.7 (Kato and Matsuda, 1980a), then there would be one XG Glc residue for every 13 surface cellulose Glc molecules. Thus, a maximum of only 8% of the surface of the cellulose microfibrils has adsorbed XG.
This conclusion is inconsistent with present models of cell walls that show XGs coating the entire surface of the cellulose microfibrils (Fig. 1A, B). It is, therefore, suggested that, at least for mung bean cell walls, these models need to be modified to indicate this small proportion of the cellulose microfibril surfaces having XG adsorbed on to it (Fig. 1C, D). It is possible that these adsorbed XGs are equivalent to the small proportion of XGs that is difficult to remove from the primary cell walls by treatment with aqueous solutions of alkali (Edelmann and Fry, 1992). In addition to XGs, the small amounts of rigid gluco- or galactoglucomannans, tentatively identified in the mung bean cell walls, may also be adsorbed on to the microfibril surfaces.
Another possible location for at least some of the rigid XG in the mung bean cell walls is within the cellulose microfibrils. Cell-wall models show a small proportion of XG in this location (Cosgrove, 2001). However, it was found that the cellulose molecules at the surface of the cellulose crystallite were similar in rigidity to those in the interior of the crystallite. This was particularly evident in the T2(C) values for the interior and surface cellulose, where proton spin diffusion is not a confounding factor. Thus, there was no disruption to the integrity of the cellulose and no evidence to support a model in which some of the XG is within the microfibrils.
The XGs with partly-rigid glucan backbones surrounded by mobile polysaccharides that have been identified may be constrained within the mung bean cell walls. This property is consistent with them forming cross-links between adjacent cellulose microfibrils or loops on the same cellulose microfibril (Fig. 1A, C). It is also consistent with them forming cross-links between cellulose microfibrils and other non-cellulosic polysaccharides, such as pectic polysaccharides (Fig. 1B, D). Evidence has recently been obtained for covalent linkages between XG and pectic polysaccharides in the primary cell walls of rose suspension-cultured cells, where up to 30% (w/w) of the XG was estimated to be involved in these interactions (Thompson and Fry, 2000). It is possible that these are not the only locations for the partly-rigid XG molecules within the cell wall. In addition, although there is evidence that XG in cell walls is associated with the cellulose microfibrils, it is also possible that some of the partly-rigid XG molecules are constrained by being cross-linked between two non-cellulosic polysaccharides.
In conclusion, the results lead to the proposal that, at least for the primary cell walls of mung bean, a modification of current models is required. In these modified models (Fig. 1C, D), no XG is located within the cellulose microfibrils and only a small proportion of the total surface area of the cellulose microfibrils has XGs adsorbed on to it. Adsorbed XG occupying only a small proportion of the total surface area of the cellulose microfibrils in primary cell walls is consistent with recent models of 
-expansin action (Cosgrove, 2001). Expansins are thought to mediate wall expansion by breaking XG-cellulose interactions. These models of expansin action propose that slippage between XG and cellulose is induced at only a restricted number of sites in the cell wall; they are therefore inconsistent with abundant XG-cellulose interactions. However, they fit well with the finding that these interactions are infrequent.
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Acknowledgement |
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This work was supported by the Marsden Fund of the Royal Society of New Zealand (grant no. UOA806).
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