Microanalysis of Plant Cell Wall Polysaccharides

Molecular Plant

•

Volume 2

•

Number 5

•

Pages 922–932

•

September 2009

RESEARCH ARTICLE

Microanalysis of Plant Cell Wall Polysaccharides Nicolai Obela,b, Veronika Erbenc, Tatjana Schwarza, Stefan Ku¨hneld, Andrea Fodora and Markus Paulya,e,1 a Max-Planck-Institut for Molecular Plant Physiology, Am Mu¨hlenberg 1, 14476 Golm, Potsdam, Germany b Present address: European Patent Office, Erhardtstrasse 27, Munich, Germany c Helmhotz Center Munich, Institute of Biological and Medical Imaging, Ingolstaedter Landstrass. 1, 85764 Neuherberg, Germany d Wageningen University, Department of Agrotechnology and Food Sciences, Laboratory of Food Chemistry, Bomenweg 2, 6703 HD Wageningen, The Netherlands e DOE Plant Research Lab., Michigan State University, East Lansing, MI 48864, USA

ABSTRACT Oligosaccharide Mass Profiling (OLIMP) allows a fast and sensitive assessment of cell wall polymer structure when coupled with Matrix Assisted Laser Desorption Ionisation Time Of Flight Mass Spectrometry (MALDI–TOF MS). The short time required for sample preparation and analysis makes possible the study of a wide range of plant organs, revealing a high degree of heterogeneity in the substitution pattern of wall polymers such as the cross-linking glycan xyloglucan and the pectic polysaccharide homogalacturonan. The high sensitivity of MALDI–TOF allows the use of small amounts of samples, thus making it possible to investigate the wall structure of single cell types when material is collected by such methods as laser micro-dissection. As an example, the analysis of the xyloglucan structure in the leaf cell types outer epidermis layer, entire epidermis cell layer, palisade mesophyll cells, and vascular bundles were investigated. OLIMP is amenable to in situ wall analysis, where wall polymers are analyzed on unprepared plant tissue itself without first isolating cell walls. In addition, OLIMP enables analysis of wall polymers in Golgi-enriched fractions, the location of nascent matrix polysaccharide biosynthesis, enabling separation of the processes of wall biosynthesis versus post-deposition apoplastic metabolism. These new tools will make possible a semi-quantitative analysis of the cell wall at an unprecedented level. Key words: Carbohydrate metabolism; xyloglucan; mass spectrometry; cell expansion; cell walls; Arabidopsis; laser microdissection.

INTRODUCTION Plant cells are surrounded by a cell wall that consists of complex heteroglycans, structural and catalytic proteins as well as polyphenolics (Carpita and McCann, 2000). This complex structure not only is responsible for the structural integrity of the cell, but it also represents the plant’s outer barrier against the environment, thus protecting the plant against biotic and abiotic stresses. The wall is a highly dynamic entity that undergoes reorganization and metabolism during the essential processes of cell elongation and differentiation (Carpita and Gibeaut, 1993; Cosgrove, 2005). Even though an enormous wealth of information has been obtained about the polysaccharide structures present in plant walls in general (Carpita and Gibeaut, 1993), very little is known about how the various components contribute to the diverse functions of the wall (Somerville et al., 2004). Major contributions have come from the model system Arabidopsis thaliana, which allowed the identification and characterization of wall mutants (Reiter et al., 1997). Moreover, the unique genetic resources available for Arabidopsis allowed reverse genetics approaches encompassing the identification of genes that are involved in wall biosynthesis and metabolism

(Lerouxel et al., 2006), genetic manipulation of their expression levels (e.g. by obtaining insertional knockout mutants), and subsequent characterization of those mutants (Cavalier et al., 2008; Jensen et al., 2008; Madson et al., 2003). With advances in such approaches, the molecular mechanisms of wall polysaccharide biosynthesis and metabolism are emerging (Farrokhi et al., 2006). Although the structure of the walls of Arabidopsis resembles that of many other dicots (Zablackis et al., 1995), one disadvantage of using Arabidopsis as a wall model system is its size and thus the small amount of rather heterogenous plant material. Arabidopsis, like other higher plants, consists of up to 40 functionally different cell types, most of which are identifiable by their wall structure (Carpita and McCann, 2000). Even a monosaccharide compositional analysis of various plant organs demonstrates the overall 1 To whom correspondence should be addressed. E-mail paulymar@msu. edu, fax +1-517-353-9168, tel. +1-517-353-4333.

ª The Author 2009. Published by the Molecular Plant Shanghai Editorial Office in association with Oxford University Press on behalf of CSPP and IPPE, SIBS, CAS. doi: 10.1093/mp/ssp046, Advance Access publication 6 July 2009 Received 29 April 2009; accepted 10 June 2009

Obel et al.

RESULTS Polysaccharide Analysis of Plant Organs OLIMP takes advantage of the specificity of glycosylhydrolases to solubilize specific polysaccharides from wall materials in the form of oligosaccharides, which are then analyzed by mass spectrometry (MS) (Lerouxel et al., 2002). OLIMP has been successfully performed on cell wall material for the analysis of XyG and the pectic polysaccharide homogalacturonan (HG; Obel et al., 2006). For XyG analysis, cell wall material is digested with xyloglucan-specific endo-glucanase (XEG), resulting in a detailed XyG oligosaccharide (XyGO) composition of the tissue including detailed sidechain distribution patterns (Figure 1A). When the pectins present in wall materials are digested with polygalacturonase (endoPG) together with pectin methylesterase (PME), OLIMP profiles can be generated

Cell wall microanalysis

|

923

Relative intensity

A XXXG

XXFG XXFG XXLG/ XLXG

XLFG XLFG/ XLFG

XXLG/ XXG

700

XLXG

GXXG 900

1100

1300

1500

1700

800

1000

1200

(GalA)7+Me5+Ac

(GalA)6+Me5

(GalA)6+Me4

(GalA)5+Me (GalA)5+Me2 (GalA)5+Me4 (GalA)5+Me5

B

(GalA)4+Me (GalA)4+Me4

m/z

Relative intensity

variance in wall composition (Richmond and Somerville, 2001). Such crude analyses as these demonstrate that techniques are urgently needed to analyze wall structure in detail in a cellular rather than organ or whole-plant context. One technique that enables us to look at wall structure on a cellular level is the labeling of tissue sections with polysaccharide-specific antibodies (Willats et al., 2000). Unfortunately, few antibodies towards defined wall epitopes are available to date. Moreover, the failure to detect an epitope by an antibody does not indicate an absence of the epitope, as it could also be masked by further substituents, polymer aggregation, or conformational changes. Another technique that can be used for elucidating changes in cell wall architecture is FTIR microscopy, but assigning observed changes to a specific polymer or linkage type remains elusive (McCann et al., 2001). Here, we describe the optimization and use of a cell wall analytical technique, oligosaccharide mass profiling (OLIMP, Lerouxel et al., 2002; Obel et al., 2006), which allows wall analysis even at cell type and organelle levels, or on plant tissue itself. We focus on the analysis of xyloglucan (XyG), the dominant cross-linking glycan in Arabidopsis, whose metabolism is thought to play a major role in cell elongation (Pauly et al., 2001). XyG sidechain structures can be quite diverse and are typically described using a single-letter nomenclature to represent the substitution pattern of each backbone glucosyl residue (Fry et al., 1993). For XyG occurring in Arabidopsis, the letters G and X denote an unbranched Glcp residue and a-DXylp-(1/6)-b-D-Glcp, respectively. The xylosyl residue may be substituted with a b-D-Galp (L sidechain), which is often further substituted with an a-L-Fucp residue to form the F sidechain. The Galp residue can also be O-acetylated indicated by underlining the one-code-letter (Hoffman et al., 2005). However, the precise distribution and modification of the sidechains in XyG is not known, presenting a major impediment to understanding the XyG structure and function during plant cell development.

d

1400

m/z Figure 1. OLIMP Spectra of Wall Material Derived from 8-Week-Old Rosette Leaves. (A) Spectrum of solubilized XyG oligosaccharides (XyGOs) facilitating a xyloglucanase (XEG). The structures of the various ion signals are shown using the nomenclature described by Fry et al. (1993). Underlined structures represent O-acetylated sidechains. (B) Spectrum of solubilized oligogalacturonides using a combination of endopolygalacturonase (endoPG) and pectinmethylesterase (PME). Putative structures are shown. GalA, galacturonic acid; Me, methylester; Ac, O-acetyl substituent.

for homogalacturonan (HG), the major component of the pectic polysaccharides (Figure 1B). The analysis results in a profile that includes the occurrence and distribution of the remaining methylesters and/or O-acetyl-esters. The peak areas of individual ion signals of the spectrum can be used to calculate the relative abundance of each structure. OLIMP on XyG was generated from tissue material derived from 10 different plant organs harvested, whereas pectin spectra could be obtained only from rosette leaves, various stem segments, petioles, and cauline leaves. The spectra looked qualitatively as shown in Figure 1—namely the same ions were observed—but the spectra varied in the quantities of the various ions. To highlight differences, we performed a principal component analysis (PCA) using the mean abundance of the various XyGOs and HG oligosaccharides (HGOs) generated from individual growth experiments (Figure 2). For XyG, principal components 1 and 2 represented 93.7% of the total variance. The loading indicated that component 1 represented mainly an abundance of the structures XXXG and XXFG, whereas component 2 represented mainly O-acetylated XXFG. The greatest variation in XyGO composition was observed between trichomes and the upper part of the stem, but there was

924

|

Obel et al.

d

Cell wall microanalysis

A 0.60

Factor 2

0.40 0.20 0.00

-0.20 0.05

0.10

0.15

0.20

0.25

Factor 1

B

Polysaccharide Analysis of Specific Leaf Tissues 0.60 0.40

Factor 2

a tetra-oligogalacturonide with a single methylester group, whereas component 2 mainly accounted for variation in penta-oligogalacturonides with varying degrees of methylesterification. PCA showed that the stem segments clustered differently from rosette leaves and petioles. Pair-wise comparison of rosette leaves with segments from the bottom of the stem showed significant changes in the homogalacturonan structure. In rosette leaves, single methylesterified tetraoligogalacturonides were more abundant, whereas in stem segments, more penta-oligogalacturonides with high methylesterification were present. Taken together, the data provide clear evidence that various organs of a plant have different XyG and HG structures.

0.20 0.00 -0.20 -0.40

0.16

0.20

0.24

0.28

0.32

Factor 1 Figure 2. Principal Component Analysis of OLIMP Derived from Different Arabidopsis Plant Organs. Various plant organs were analyzed by OLIMP with XEG (A) and the combination of endoPG and PME (B). Each point represents the average of at least three replicates from a single harvest, and each plant tissue was harvested three independent times. Black circles, trichomes; open triangles, petals; open circles, rosette leaf; black triangles, cauline leaves; open diamond, roots; open square, top stem segment; black square, middle stem segment; gray square, bottom stem segment; closed square, siliques; gray circle, petiole.

also a distinct difference between trichomes and the rosette leaves (Figure 2A). A pair-wise comparison of rosette leaves with trichomes showed that the O-acetylation level of XyG in trichomes is less than half of that in leaves. It was furthermore possible to observe that this change in O-acetylation was due to a lower acetylation of XXFG and XLFG, whereas the acetylation of XXLG/XLXG was almost identical in the two samples. In the PCA, we also observed that the three stem segments were placed differently along principal component 1 (open square, black square, gray square). We performed a further pair-wise comparison of the three segments and found an increasing abundance of the XXXG and XXFG fragments in the top of the stem. A higher level of XyG O-acetylation at the base of the stem was also observed when compared to the segments from the upper part of the stem. We performed OLIMP on HG from the wall materials of various organs and analyzed the spectra using PCA (Figure 2B). PC 1 and 2 represented 98% of the variance. Component 1 mainly represented a 759-Da fragment, which corresponded to

Wall material can be collected from specific leaf cell types using laser catapulting dissection microscopy. To ensure that the extensive sample preparation prior to laser dissecting and the heat generated by laser dissecting itself did not affect polysaccharide structure, leaf material with and without embedding was subjected to OLIMP of XyG. No significant differences were observed (data not shown). Furthermore, when OLIMP of entire leaves, extensively dissected and catapulted using the laser dissector, was compared to that of leaves prepared with the regular wall preparation method, no significant differences were observed in the profiles (data not shown). Thus, the laser dissecting protocol used here did not affect OLIMP analyses. Cell material was dissected and collected from the outer walls of an epidermis layer, an entire epidermis cell layer (also containing walls from the adjacent palisade mesophyll cells), palisade cells, spongy mesophyl cells, and cells of the vascular bundles (for examples of dissection, see Figure 3). Wall material of approximately 4000 cells was needed to obtain a clear spectrum. Differences of the recorded spectra were evaluated using PCA (Figure 4). Components 1 and 2 represented 61 and 22% of the variance, respectively. Component 1 represented mainly XXFG, XLFG, and XLFG, whereas component 2 represented largely low substituted XXG and XXXG. As can be seen in the PCA, the vascular tissue was very different from the other leaf samples distinguished mainly by component 1. A direct comparison of the spongy mesophyl and the vascular tissue (Figure 4B) showed no significant differences in the overall level of galactosylation and fucosylation, but a significant increase in overall O-acetylation. More specifically, a decrease of more than 50% in the abundance of XXFG was observed. That difference was offset with a similar increase in the corresponding O-acetylated fragment (Figure 4C). A similar pattern was also observed for the XLFG fragment where a decrease in the non-acetylated fragment was compensated for by a similar increase in the O-acetylated fragment, indicating distinct O-acetylation patterns in these tissue types. The walls from the outer walls of epidermis cells, the entire epidermis cell layer, and the palisade cells did not separate in the PCA. Similarly, the mesophyl spectra did not cluster separately from

Obel et al.

d

Cell wall microanalysis

|

925

In Situ OLIMP Wall Analysis of Plant Tissue

Figure 3. Laser Dissection and Catapulting of Specific Leaf Cells. 20 lm thick cross-sections of paraffin-embedded leaves were used for laser dissecting. (A–E) show 103 magnification of a leaf crosssection. (A) Leaf cross-section. (B) Leaf cross-section after removal and collection of whole epidermis cells. (C) Same as (B) after removal of only the outer layer of the epidermis. (D) Removal of palisade cells. (E) Mesophyl cells sampling. (F) and (G) show a 403 magnification of the midrib before (F) and after (G) removal of the vascular tissue.

the above-mentioned tissues, probably because of the large variability along principal component 2. Although the outer walls of the epidermis and walls from the entire epidermis cell layer grouped in the same part of the PCA, a pair-wise comparison of the spectra obtained from both tissues revealed distinct differences in XyGO composition (Figure 4C): a significant reduction in the overall level of XyG galactosylation and XyG fucosylation (Figure 4B). However, O-acetylation levels were not altered. These data suggest not only that the epidermal tissue contains a distinct XyG structure, but also that the inner and outer wall of the epidermis contain different XyGO compositions, indicating an altered XyG metabolism on opposite sides of the same cell.

Another way to generate XyG profiles is by what we term ‘in situ’ OLIMP analysis. As shown in Figure 5A and 5B, fresh plant tissue can be placed on a metal MALDI-target plate, and 0.5 ll droplets of enzyme can be added to various plant tissue sections (see halos indicated by arrows in Figure 5). After incubation of the tissue, the whole sample is dried under vacuum and matrix chemical added on the dried enzyme spots. Then, the entire target plate (plate, plant tissue, dried enzyme spots plus matrix) is inserted into the MALDI–TOF machine. Excellent high signal-to-noise spectra can then be obtained by guiding the laser to those spots to generate spectra. This method allows rapid wall analysis with a good spatial resolution. As examples, in situ XyG OLIMP was performed along the roots of Arabidopsis wild-type (Col-0) and mur1 (Figure 5A). Mur1 is defective in a GDP-mannose dehydratase involved in GDP-fucose formation leading to underfucosylated, but L-galactosylated XyG (Zablackis et al., 1996). This change in XyG structure is clearly visible in the spectra obtained from a spot on the root (Figure 5C). The mutant could thus have been identified in minutes. XyG profiles obtained from various positions along the roots of WT and mur1 showed no significant differences (data not shown), indicating that the respective XyG profile is constant along the root. Another example is in situ XyG OLIMP of 4-day-old etiolated hypocotyls (Figure 5B). Three tissues were analyzed, two on the above-ground tissue and one spot on the root. The obtained spectra are shown in Figure 5D. Both above-ground tissues exhibited similar spectra (Figure 5D, top). However, although the root spectra were qualitatively similar, they did show significant changes in abundance of various XyGOs. In particular, fucosylated XyGOs represented by XXFG, XLFG, and their acetylated counterparts (XXFG, XLFG) were more dominant in the root (Figure 5D, bottom).

Polysaccharide Analysis of the Plant Golgi Apparatus Plant matrix polysaccharides are synthesized in the Golgi apparatus (Lerouxel et al., 2006; Scheible and Pauly, 2004) and undergo further modification once deposited in the apoplast. Usually, only the apoplastic wall polymers are analyzed. However, owing to the sensitivity of OLIMP, oligosaccharide compositional data can also be obtained from preparations of Golgi-enriched fractions excluding the apoplastic cell wall (Figure 6A). Such an analysis allows the characterization of nascent polysaccharides. Compared to the XyGO composition of the apoplast, significant structural differences can be found in the Golgi-enriched fraction (Figure 6B). Smaller, lowsubstituted XyGOs such as XXG and GXXG are more abundant in the Golgi, whereas higher-substituted XyGOs are less abundant (overall degree of fucosylation and galactosylation, specifically XLFG). Such data could give us unique insights into the mode of biosynthesis of these polysaccharides and into their subsequent apoplastic modifications.

926

|

Obel et al.

d

Cell wall microanalysis

A

0.15

Factor 2

0.1 0.05 0 -0.05 -0.1 -0.1

-0.05

0

0.05

0.1

0.15

0.2

0.25

Factor 1

C

0.5

* *

0.25

*

*

XX G G XX XX G X XX G L XX G LG XX F XX G F XL G F XL G A FG ll A c A ll F A ll L

0

0.75

* 0.5

*

0.25

* 0 XX G G XX XX G X XX G L XX G LG XX F XX G FG XL F XL G A FG ll A c A ll F A ll L

0.75

Relative abundance

Relative abundance

B

Figure 4. PCA and Pair-Wise Comparison of Specific Plant Tissues as Shown in Figure 3. (A) Each point in the PCA represents the OLIMP profile of material collected using laser dissecting. The following material was collected: vascular tissue (star); outer wall of epidermis (circle), entire epidermis layer (diamond), palisade (triangle), and mesophyl (square). (B) Pair-wise comparison of vascular (gray bars) and mesophyl (white bars) samples.***(C) Pair-wise comparison of outer epidermis (gray bars) and whole epidermis (white bars) samples. Significant different abundance levels (p , 0.05) are indicated with an asterisk, n = 3.

DISCUSSION Established techniques for the analysis of wall polysaccharides are sequential extraction of wall material using chemicals or enzymes (Selvendran and O’Neill, 1987) followed by monosaccharide composition (Albersheim et al., 1967) or glycosidic linkage analysis (York et al., 1985). These techniques are usually used on material derived from whole plant organs and thus any tissue-specific structural information is diluted and averaged. In addition, these techniques are time-consuming and labor-intensive. One technique that can be used to look at wall structure on a cellular level is the labeling of tissue sections with polysaccharide-specific antibodies (Willats et al., 2000). Unfortunately, the number of defined wall polysaccharide epitopes that can be recognized is, at the moment, very limited. Moreover, negative results (non-binding of the antibody) may be due not only to absence of epitope, but also to epitope masked by substituents, polymer aggregation, and other polymer conformational changes. A fast and sensitive technique that can be used for elucidating changes in cell wall architecture at even a cellular level is FTIR-microscopy, but assigning observed changes to a specific polymer or linkage types remains elusive (McCann et al., 1997). A more promising technique is oligosaccharide profiling, which takes advantage of the specificity of glycosylhydrolases to solubilize

polysaccharides from wall materials in the form of oligosaccharides; these can then be analyzed with various methods including liquid chromatography (Kakita et al., 2002), gel electrophoresis (Goubet et al., 2002, 2003), and mass-spectrometry (MS) (Lerouxel et al., 2002). Oligosaccharide mass profiling (OLIMP) is limited by available specific wall hydrolytic enzymes, but their diversity is increasing (Bauer et al., 2006). Of these methods, OLIMP is particularly attractive, as it does not require any derivatization procedure and a spectrum can be obtained in less than a minute rather than an hour-long chromatography run. OLIMP generates a semiquantitative fingerprint of a particular wall polysaccharide (Figure 1). However, OLIMP does not allow the determination of absolute quantities present in a sample, and often only a portion of the given polymer can be solubilized from the wall when using a hydrolytic enzyme (Obel et al., 2006; Pauly et al., 1999a). OLIMP depends on the enzymatic accessibility of the cell wall polymers for the solubilization of the oligosaccharides. The high sensitivity of MALDI–TOF MS allows the use of very small amounts of sample and can, as described here, be coupled with laser micro-dissection,allowing investigation of the cell wall at the cellular level. Structural isomers can also not be distinguished by MS, but fragmentation of ions by MALDI post-source decay analysis and/or electrospray allows a distinction of isomers, such as XLXG and

Obel et al.

d

Cell wall microanalysis

|

927

XXLG (Yamagaki et al., 1998). Despite these limitations, OLIMP presents a rapid and particularly sensitive method for investigating wall structures (Obel et al., 2006). We took advantage of these attributes to investigate structural diversity of various polymers in plant organs, cell types, and even organelles.

OLIMP Analysis of Plant Organs Reveals Organ-Specific Wall Structures Even though it is known that plant tissues contain a variety of heterogeneous polymers (Carpita and McCann, 2000), very little is known about the diversity of the polymers on a detailed structural level. For example, one study determined only the monosaccharide sugar composition of various Arabidopsis organs (Richmond and Somerville, 2001), but not from which polymers these sugars originated. Here, the detailed substitution patterns of XyG and pectic HG were studied in various plant organs. All tissues investigated contained qualitatively the same oligosaccharide compositions. However, their quantity varied significantly. The biggest variation in XyGO composition was observed in the walls of trichomes, which are clearly set apart from the other organs. The main difference is the abundance of XXFG; trichomes contain less than half the O-acetylated XyGOs than leaves, for example. It also became evident that the abundance of XXXG and XXFG oligos increases in abundance from bottom to top of the stem. The abundance of the different sidechains in XyG has been investigated in previous studies. In pea, a large variation in the abundance of acetylated XLFG was observed when comparing rosette leaves with stem segments (Pauly et al., 2001) and an investigation of XyG fucosylation in Arabidopsis led to the discovery that the highest fucosylation level occurred in flowers and roots (Perrin et al., 2003). The root data were confirmed here, but petals had the lowest level of XyGfucosylation from all the organs investigated, suggesting a large variation in XyG structure within the flower. Similar to the report by Perrin et al. (2003), high levels of O-acetylation were found in leaves, the lower part of the stem, and siliques. When digesting cell walls with a combination of endoPG and PME, we were able to obtain reproducible fingerprints from different organs, and to observe specific variations between plant organs. However, a HG structural grouping

Figure 5. In Situ OLIMP Analysis of Various Plant Tissue.

(A) Roots of Arabidopsis WT and mur1 mutant placed on the target plate with added 0.5 ll of enzyme drops at various positions (halos, white triangles). (B) Etiolated 5-day-old Arabidopsis hypocotyl placed on the target plate with added enzyme drops at various positions (halos, white triangles). (C) OLIMP spectra obtained from the two encircled spots (white circles, (A)) of WT and mur1. (D) OLIMP spectra obtained from the two encircled spots (white circles, (B)) of cotyledons and root. The structures of the various ion signals are shown using the nomenclature described by Fry et al. (1993) including the J sidechain, where an L sidechain is extended with an a-L-Galp residue (Zablackis, 1996).

928

|

Obel et al.

A

Cell wall microanalysis

d

Microsomes

Relative intensity

XXXG

XXLG/ XLXG

XXFG XXFG XLFG XLFG/ XLFG

XXLG/ XLXG

1100

1300

1500

1700

m/z

Relative abundance

B

0.75

* 0.5

0.25

*

*

*

XX G G XX G XX XG XX LG XX LG XX FG XX FG XL FG XL F A G ll A c A ll F A ll L

0

Figure 6. XyG OLIMP Analysis of a Golgi-Enriched Fraction. (A) XyG OLIMP spectrum of an isolated Golgi-enriched fraction obtained from etiolated Arabidopsis hypocotyls. (B) Pair-wise comparison of XyGO profiles from cell walls (gray) and Golgi-enriched fraction (white). Significant different abundance levels (p , 0.05) are indicated with an asterisk, n = 3.

of the plant organs was not as clear as in the case of XyG, indicating a more homogeneous structure of homogalacturonan throughout the plant. In Arabidopsis stems, a high abundance of blockwise methyl-esterification of HG has been demonstrated (Goubet et al., 2003) and this might be reflected in this study by the high abundance of methyl-esterified penta-oligogalacturonides in the stem when compared to leaves. Studies using antibodies have shown variations of the cell wall structure at cellular levels for both pectin and xyloglucan (Freshour et al., 2003; Willats et al., 2001); thus, it was not surprising to find structural changes among plant organs. In contrast to antibody studies, OLIMP does not depend on the presence of a few epitopes, but gives a quantitative relative abundance of well defined XyG sidechains.

OLIMP Analysis of Various Leaf Tissues Reveals TissueSpecific Wall Structures Laser-microdissection catapulting is a recent technique that has allowed the characterization of specific tissues or even individual cells (EmmertBuck et al., 1996). In particular, RT–PCRamplified transcript levels can be determined on a single-cell

level (Brandt et al., 1999; Nakazono et al., 2003). Unfortunately, wall polymers cannot be amplified, but we took advantage of the sensitivity of mass spectrometry to investigate XyG on a cell-type level. Reproducible spectra with low noise ratios can be obtained when combining harvested walls of approximately 4000 cells. The laser in the microdissection instrument does not separate two cells, namely it does not cut through the middle lamella of two adjacent walls. Therefore, when harvesting cells from an epidermis cell layer, for example, one should keep in mind that those walls also contain the outer wall of the adjacent palisade mesophyll tissue. One advantageous aspect of this limitation is that one can cut through cells and collect specific local wall sections. For example, when cutting through an epidermis, the outer layer of this tissue can be specifically harvested. Importantly, the cutting of tissue with the laser did not seem to alter the oligosaccharide profile when compared, for example, to ball milling. Despite some limitations, laser micro-dissectioning does allow the tissuespecific harvest of wall material for detailed characterization in an unprecedented way. An investigation of the XyGO profile derived from various tissues present in the leaf (outer walls of the epidermis, entire epidermis cell layer, palisade tissue, spongy mesophyll tissue, and vascular tissue) showed significant alterations in XyG structure. Particularly, XyG in the vascular tissue was very different from other leaf tissue. More specifically, an overall increase in XyG O-acetylation was observed. Unlike the primary walls of other tissues that are mainly XyG-rich, secondary walls of the vascular tissue are rich in another cross-linking glycan, xylan. Hence, a change in XyG structure in the vascular tissue is not surprising, and one hypothesis could be that an increased abundance of O-acetylation leads to a more hydrophobic wall. Alternatively, the vascular tissue contains cells with primary walls and those walls might contain the more hydrophobic XyG to accommodate the needs to the cell types. Also, several structural differences could be observed between the outer epidermis wall preparation compared to that of an entire epidermis cell layer preparation, demonstrating for the first time that there are local structural XyG differences within a single cell. The reason in the case of the epidermis could be that the outer wall is in direct contact with the environment and thus has a different functionality (protection against water loss through the cuticle and epicuticular waxes, protection against pathogens) compared to the inner wall. The data presented here demonstrate that the (epidermis) cell has the capacity to affect XyG structure locally; yet, to be investigated is how this structural difference is manifested in metabolism, but correlations with transcriptome analyses of these cell types could be insightful (Brandt et al., 1999). Also, how the altered XyG structure mechanistically contributes to specific epidermal functionalities remains to be investigated.

In Situ OLIMP Wall Analysis Offers Exciting Possibilities MALDI–TOF MS ionizes molecules with the energy of a laser, which allows the ionization on a spatially defined area in

Obel et al.

the lm range. We took advantage of this feature to develop an ‘in situ’ OLIMP analysis. As demonstrated by two examples, roots and etiolated hypocotyls, OLIMP spectra could be obtained directly on various tissues eliminating wall preparation procedures and enhancing the spatial resolution of wall polymer analysis down to the tissue level. The results are quite promising: a mutant (mur1) was rapidly identified by its extensively altered XyG profile, and differences in the XyGO composition of cotyledons and roots were demonstrated. In situ analysis of the root showed no variation of XyG structure, regardless of the position on the root. However, when comparing in situ profiles to those of entire roots, we observed significant changes, which led us to suggest that the enzyme (XEG) is mainly active at the outer regions of the root, as in the epidermis cell layer. Freshour et al. (2003) demonstrated by antibody-labeling that the epidermal cells of the Arabidopsis root have a higher abundance of a-(1-2)-linked fucose than cortex walls. In contrast, the in situ analysis did not show a change in XyG fucosylation compared to whole roots. We hypothesize that the observed increase of the antibody-labeling in the epidermis may be due to the presence of other polymers that can be recognized by the antibody, such as RGI (Puhlmann et al., 1994), or that there is not a higher level of XyG fucosylation, but simply a higher abundance of overall XyG. However, one question that needs to be addressed in the future to assess the in situ method more thoroughly is how far the enzyme penetrates during the experimental conditions used. In situ plant tissue analysis has so far been achieved with metabolites (Cha et al., 2008), but, with our results here, the exciting possibility emerges of performing tissue ‘imaging’ (Francese et al., 2009) of the wall.

OLIMP Analysis of Nascent Polysaccharides Present in the Plant Golgi Apparatus Matrix polysaccharides such as cross-linking glycans and the pectic polysaccharides are synthesized in the plant Golgi apparatus (Lerouxel et al., 2006). After secretion into the apoplast, they form polymer networks and can undergo further modification through enzymes or radicals (Fry, 2004). Hence, when a particular wall polymer structure is characterized in the wall, the observed structure can be considered the product of two spatially distinct processes: biosynthesis in the Golgi and apoplastic modification. Based on the structure, one cannot deduce the individual processes. Taking advantage of the sensitivity of OLIMP, we were able to demonstrate for the first time the sidechain substitution pattern of nascent XyG present in a Golgi-enriched fraction, potentially contaminated with endoplasmic reticulum (Munoz et al., 1996) but certainly not with apoplastic cell wall fragments. The characterization of wall polymer structures specifically in the Golgi will allow a separation and thus definition of the biosynthetic from the apoplastic process. OLIMP makes possible such an analysis with much more structural detail than previous immuno-gold labeling procedures (Moore et al., 1986). However, one can assume that the Golgi-enriched fraction contains all parts of

d

Cell wall microanalysis

|

929

the Golgi—cis- medial-, and trans-cisternae—and an OLIMP characterization of a polymer will not be able to distinguish the structure within these sub-compartments. In the case of XyG, the data clearly demonstrate a lower degree of substitution of XyG in the Golgi compared to the apoplast, probably representing XyG in its various ‘unfinished’ stages. Analyses such as these will have great potential to give us vital clues about the molecular mechanism of wall polysaccharide biosynthesis, especially when combined with XyG biosynthesis mutants (Vanzin et al., 2002) and proteomic studies of the Golgi (Dunkley et al., 2006). In summary, the various OLIMP techniques developed here allow an assessment of changes in structural elements of polysaccharides at an unprecedented organ, cell type, and even organelle resolution and are complementary to existing techniques utilizing antibodies and FTIR-microscopy. OLIMP combines high sensitivity with short time of analysis. We expect that OLIMP will enable an enhanced understanding of cell wall metabolism and ultimately of the function of specific cell wall polymer substitutions in plant growth and development.

METHODS Materials Endopolygalacturonase (endoPG) was purchased from Megazyme International, Ireland (Bray, Ireland). Xyloglucanase (XEG) and pectin methyl esterase (PME) were kindly provided by Novozyme (Bagsværd, Denmark). Xylene was purchased from Merck (Darmstadt, Germany) and 2,5-dihydroxybenzoic acid was from Aldrich (Sigma-Aldrich Chemie Gmbh, Munich, Germany). Paraffin wax was provided by Fluka (Sigma-Aldrich Chemie Gmbh, Munich, Germany) and, if not otherwise stated, the chemicals were supplied by Sigma (Sigma-Aldrich Chemie Gmbh, Munich, Germany).

Plant Material and Growth Conditions Arabidopsis thaliana, ecotype Columbia 0, was grown on soil in climate chambers. For the first 4 weeks, plants were grown in light periods of 8 h and 20
C and dark periods of 16 h and 16
C and a constant humidity of 70%. For the following 4 weeks, plants were grown in 16 h light, 20
C and 8 h darkness, 16
C at a constant humidity of 75%. After 8 weeks, the plant organs (see below) were harvested and flash frozen in liquid nitrogen. Roots were grown for 13 d on vertical plates containing 4.4 g l 1 Murashige and Skoog basal medium with 1% sucrose and 0.8% agar. The seeds were placed on the agar, and the plates were wrapped in aluminium foil and stored for 2 d at 4
C before being transferred to a growth chamber with 16 h light and 22
C. Here, the plates were unwrapped and placed vertically. After 11 d in the growth chamber, roots were harvested and stored in 100% ethanol at 4
C. Etiolated seedlings were grown for 5 d on plates containing 4.4 g l 1 Murashige and Skoog basal medium with 1% sucrose

930

|

Obel et al.

d

Cell wall microanalysis

and 0.8% agar. The seeds were placed on the agar, and the plates were wrapped in aluminium foil and stored for 2 d at 4
C before being transferred to 22
C. For preparation of the Golgi-enriched fraction, etiolated Arabidopsis seedlings were grown in liquid culture containing 4.4 g l 1 Murashige and Skoog basal medium with 1% sucrose for 14 d at 22
C on a rotating shaker (100 rpm) in the dark.

Laser Dissection Leaves from 5-week-old plants were embedded without fixation using a modification of existing embedding protocols (Brandt et al., 1999). The leaf was harvested and immediately transferred into 1 ml 20% aqueous ethanol. In steps of 10%, the ethanol concentration was increased until 100% was reached. The last incubation in 100% ethanol was repeated twice, before incubating in a solution of ethanol and xylene (1:1, v:v) followed by three incubations in pure xylene. All incubations were done for at least 6 h at room temperature. Melted paraffin wax was mixed with xylene and the leaf was transferred from pure xylene to a xylene solution with 30% (v/v) paraffin wax and, after 8 h incubation at 42
C, the leaf was transferred to 50% and subsequently 80% and paraffin solutions and, at each step, incubated for 8 h at 52 and 58
C, respectively. Finally, the leaf was incubated twice in pure paraffin at 64
C for 7 h. The embedded leaf was transferred into an aluminium tray with melted paraffin, which was then allowed to solidify. The leaf was excised from the paraffin and mounted on a cutting block and 20 or 40-lm slices in the transverse or the longitudinal direction were made and placed on microscope glass slides (SuperFrost Plus, Menzel Gmbh, Braunschwieg, Germany). The paraffin was melted and removed from the specimen by carefully adding xylene and removing the solubilized paraffin with a paper towel. For one sample, approximately 4000 cells from the outer wall of the epidermis, the entire epidermis cell layer, palisade cells, mesophyls and vascular bundle were excised and collected in the cap of a 0.5-ml eppendorf tube using a P.A.L.M. laser dissector according to the manufacture’s instructions (P.A.L.M. Microlaser Technologies AG, Bernried, Germany). In the case of the palisade cells, the epidermis was removed prior to collection. The collected material was spun down and washed in xylene and chloroform/methanol, 1:1 v/v, before drying and digestion with XEG (see below).

Preparation of a Golgi-Enriched Fraction Arabidopsis seedlings grown in liquid culture were harvested and a Golgi-enriched fraction was obtained essentially as described in Munoz et al. (1996). The entire procedure was carried out on ice. Briefly, seedlings (8 g fresh weight) were finely cut with a razor blade and ruptured carefully with a mortar and pestle. The suspension was centrifuged at 2000 g, resulting in a pellet containing cell wall material and a supernatant containing microsomes. Fractionation of the microsomes was achieved by differential sucrose-centrifugation as described (Munoz et al., 1996) with the Golgi-enriched fraction collect-

ing between the 36 and 38% sucrose layer. The fraction was flash-frozen in liquid nitrogen before the OLIMP washing procedure (see below).

Generation of Wall Oligosaccharides The following plant organs were harvested: rosette leaf, cauline leaf, petiole, root, petals, silique, and three different segments of the stem, each 2 cm long, one from the base of the stem, a segment two-thirds up the stem, and a tip segment, segment 3. Leaf trichomes were isolated by gluing a leaf to a glass slide by a drop of water followed by dipping into liquid nitrogen. A pair of tweezers also dipped in liquid nitrogen was then used to collect the trichomes by stroking over the surface of the leaf without touching the epidermis. The collected trichomes were transferred to a 2-ml eppendorf tube with a drop of water. The plant/tissue/Golgi material was prepared for OLIMP as described previously (Lerouxel et al., 2002). Briefly, the frozen material was homogenized in a retsch-mill (model MM200, Retsch, Haan, Germany). The residue was washed with 70% aqueous ethanol, and pelleted through centrifugation. The pellet was washed with a chloroform–methanol mixture (1:1 v:v). For the profiling of XyG, after centrifugation, decanting, and evaporation of the solvent, the material was digested with a solution of 100 mM ammoniumformate, pH 4.5, containing 0.02 units of a purified recombinant XyG-specific endoglucanase (XEG; EC 3.2.1.151, Pauly et al., 1999b) and incubated for 17 h at 30
C. After digestion, the suspension was centrifuged and the supernatant containing the solubilized oligosaccharides was transferred to another tube. In the case of laser-dissected material or petals, the entire supernatant was dried in a speed-vac concentrator (Concentrator 5301, Eppendorf, Hamburg, Germany) and re-suspended in 5 ll water. An almost identical treatment of the sample was performed for profiling of homogalacturonan. Here, PME (Novozyme, Bagsvaerd, Denmark) 0.08 units (1 unit releases 1 lmol of methanol per min) and endoPG (0.15 units, Megazyme, Bray, Ireland) were added as enzymes instead of XEG.

Mass Profiling Ion exchange resin (Biorex MSZ 501(D), Bio-Rad Laboratories, Hercules, USA), ;10 beads, was added to 5 ll of the oligosaccharide containing supernatants to remove buffer salts. After an 8-min incubation, the supernatant (1 ll) was spotted onto a MALDI target plate that already contained vacuum-dried crystallized 2,5-dihydroxybenzoic acid (10 lg). Mass spectra were obtained using a Voyager DE-Pro MALDI– TOF MS (Applied Biosystems, Langen, Germany) in positive mode with an accelerating voltage of 20 000 V and a delay time of 350 ns. The spectra were recorded automatically and output-data files containing a list of the ion signals and their area were generated. Spectra were only used if the two major xyloglucan peaks in the spectrum had a signalto-noise ratio of 8 or above.

Obel et al.

In Situ OLIMP Harvested roots or etiolated seedlings were placed on a blank target plate and dried before 0.5 ll of purified XEG, 0.2 U, was placed at various positions (Figure 5A and 5B). The target plate was placed in a closed humid box and the sample was incubated at 30
C overnight. The target plate was then dried in a vacuum before 0.5 ll 2,5-dihydroxybenzoic acid, 10 mg ml 1, was added on top of the dried enzyme spots and allowed to stand for 2 min before the plate was dried under vacuum and loaded into the mass spectrometer. The laser was pointed at the positions where digestion had taken place, which were visible through the video monitor connected to the MALDI– TOF and spectra were obtained using the same MS settings as described above.

Data Analysis The recorded spectra were analyzed by an in-house-developed PERL program (Lerouxel et al., 2002), which calculated the relative area of the individual ion-signals of each individual spectra and performed a pair-wise comparison and statistical assessment by a student’s t-test. To asses the variation among the various plant organs, Hierarchical Cluster Analysis and PCA was performed using the program Pirouette 2.6 (Infometrix, Woodinville, USA). The data were mean-centered and the Euclidian distance was used for calculating the distance matrix.

FUNDING This work was supported by the German GABI project 0312277D.

ACKNOWLEDGMENTS We thank Dr Kirk Schnorr (Novozymes A/S, Bagsvaerd, Denmark) for the generous gift of XEG and pectin methyl esterase. Michael Stiebritz and Kerstin Zander are thanked for valuable technical support. No conflict of interest declared.

REFERENCES Albersheim, P., Nevins, D.J., English, P.D., and Karr, A. (1967). A method for the analysis of sugars in plant cell wall polysaccharides by gas-liquid chromatography. Carbohydr. Res. 5, 340–345. Bauer, S., Vasu, P., Persson, S., Mort, A.J., and Somerville, C.R. (2006). Development and application of a suite of polysaccharidedegrading enzymes for analyzing plant cell walls. Proc. Natl Acad. Sci. U S A. 103, 11417–11422.

d

Cell wall microanalysis

|

931

Carpita, N.C., and Gibeaut, D.M. (1993). Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant J. 3, 1–30. Cavalier, D.M., et al. (2008). Disrupting two Arabidopsis thaliana xylosyltransferase genes results in plants deficient in xyloglucan, a major primary cell wall component. Plant Cell. 20, 1519–1537. Cha, S.W., Zhang, H., Ilarslan, H.I., Wurtele, E.S., Brachova, L., Nikolau, B.J., and Yeung, E.S. (2008). Direct profiling and imaging of plant metabolites in intact tissues by using colloidal graphiteassisted laser desorption ionization mass spectrometry. Plant J. 55, 348–360. Cosgrove, D.J. (2005). Growth of the plant cell wall. Nat. Rev. Mol. Cell Biol. 6, 850–861. Dunkley, T.P.J., et al. (2006). Mapping the Arabidopsis organelle proteome. Proc. Natl Acad. Sci. U S A. 103, 6518–6523. EmmertBuck, M.R., Bonner, R.F., Smith, P.D., Chuaqui, R.F., Zhuang, Z.P., Goldstein, S.R., Weiss, R.A., and Liotta, L.A. (1996). Laser capture microdissection. Science. 274, 998–1001. Farrokhi, N., Burton, R.A., Brownfield, L., Hrmova, M., Wilson, S.M., Bacic, A., and Fincher, G.B. (2006). Plant cell wall biosynthesis: genetic, biochemical and functional genomics approaches to the identification of key genes. Plant Biotechnol. J. 4, 145–167. Francese, S., Dani, F.R., Traldi, P., Mastrobuoni, G., Pieraccini, G., and Moneti, G. (2009). MALDI mass spectrometry imaging, from its origins up to today: the state of the art. Combinatorial Chemistry & High Throughput Screening. 12, 156–174. Freshour, G., Bonin, C.P., Reiter, W.D., Albersheim, P., Darvill, A.G., and Hahn, M.G. (2003). Distribution of fucose-containing xyloglucans in cell walls of the mur1 mutant of Arabidopsis. Plant Physiol. 131, 1602–1612. Fry, S.C. (2004). Primary cell wall metabolism: tracking the careers of wall polymers in living plant cells. New Phytol. 161, 641–675. Fry, S.C., et al. (1993). An unambiguous nomenclature for xyloglucan-derived oligosaccharides. Physiol. Plant. 89, 1–3. Goubet, F., Jackson, P., Deery, M.J., and Dupree, P. (2002). Polysaccharide analysis using carbohydrate gel electrophoresis: a method to study plant cell wall polysaccharides and polysaccharide hydrolases. Anal. Biochem. 300, 53–68. Goubet, F., Morriswood, B., and Dupree, P. (2003). Analysis of methylated and unmethylated polygalacturonic acid structure by polysaccharide analysis using carbohydrate gel electrophoresis. Anal. Biochem. 321, 174–182. Hoffman, M., Jia, Z.H., Pena, M.J., Cash, M., Harper, A., Blackburn, A.R., Darvill, A., and York, W.S. (2005). Structural analysis of xyloglucans in the primary cell walls of plants in the subclass Asteridae. Carbohydr. Res. 340, 1826–1840.

Brandt, S., Kehr, J., Walz, C., Imlau, A., Willmitzer, L., and Fisahn, J. (1999). A rapid method for detection of plant gene transcripts from single epidermal, mesophyll and companion cells of intact leaves. Plant J. 20, 245–250.

Jensen, J.K., Sorensen, S.O., Harholt, J., Geshi, N., Sakuragi, Y., Moller, I., Zandleven, J., Bernal, A.J., Jensen, N.B., Sorensen, C., Pauly, M., Beldman, G., Willats, W.G.T., and Scheller, H.V. (2008). Identification of a xylogalactuonan xylosyltransferase involved in pectin biosynthesis in Arabidopsis. The Plant Cell. 20, 1289–1302.

Carpita, N., and McCann, M. (2000). The cell wall. In Biochemistry and Molecular Biology of Plants, Buchanan, B., Gruissem, W., and Jones, R., eds (Rockwell, MD: American Society of Plant Physiologists), pp. 52–108.

Kakita, H., Kamishima, H., Komiya, K., and Kato, Y. (2002). Simultaneous analysis of monosaccharides and oligosaccharides by high-performance liquid chromatography with postcolumn fluorescence derivatization. J. Chromatography A. 961, 77–82.

932

|

Obel et al.

d

Cell wall microanalysis

Lerouxel, O., Cavalier, D.M., Liepman, A.H., and Keegstra, K. (2006). Biosynthesis of plant cell wall polysaccharides: a complex process. Curr. Opin. Plant Biol. 9, 621–630. Lerouxel, O., Choo, T.S., Seveno, M., Usadel, B., Faye, L., Lerouge, P., and Pauly, M. (2002). Rapid structural phenotyping of plant cell wall mutants by enzymatic oligosaccharide fingerprinting. Plant Physiol. 130, 1754–1763.

xyloglucan fucosylation in Arabidopsis. Plant Physiol. 132, 768–778. Puhlmann, J., Bucheli, E., Swain, M.J., Dunning, N., Albersheim, P., Darvill, A.G., and Hahn, M.G. (1994). Generation of monoclonalantibodies against plant cell-wall polysaccharides. 1. Characterization of a monoclonal-antibody to a terminal alpha-(1-.2)linked fucosyl-containing epitope. Plant Physiol. 104, 699–710.

Madson, M., Dunand, C., Li, X., Verma, R., Vanzin, G.F., Caplan, J., Shoue, D.A., Carpita, N.C., and Reiter, W.D. (2003). The MUR3 gene of Arabidopsis encodes a xyloglucan galactosyltransferase that is evolutionarily related to animal exostosins. Plant Cell. 15, 1662–1670.

Reiter, W.D., Chapple, C., and Somerville, C.R. (1997). Mutants of Arabidopsis thaliana with altered cell wall polysaccharide composition. Plant J. 12, 335–345.

McCann, M.C., Chen, L., Roberts, K., Kemsley, E.K., Sene, C., Carpita, N.C., Stacey, N.J., and Wilson, R.H. (1997). Infrared microspectroscopy: sampling heterogeneity in plant cell wall composition and architecture. Physiol. Plant. 100, 729–738.

Scheible, W.R., and Pauly, M. (2004). Glycosyltransferases and cell wall biosynthesis: novel players and insights. Curr. Opin. Plant Biol. 7, 285–295.

McCann, M.C., et al. (2001). Approaches to understanding the functional architecture of the plant cell wall. Phytochemistry. 57, 811–821. Moore, P.J., Darvill, A.G., Albersheim, P., and Staehelin, L.A. (1986). Immunogold localization of xyloglucan and rhamnogalacturonan I in the cell walls of suspension-cultured sycamore cells. Plant Physiol. 82, 787–794. Munoz, P., Norambuena, L., and Orellana, A. (1996). Evidence for a UDP-glucose transporter in Golgi apparatus-derived vesicles from pea and its possible role in polysaccharide biosynthesis. Plant Physiol. 112, 1585–1594. Nakazono, M., Qiu, F., Borsuk, L.A., and Schnable, P.S. (2003). Lasercapture microdissection, a tool for the global analysis of gene expression in specific plant cell types: identification of genes expressed differentially in epidermal cells or vascular tissues of maize. Plant Cell. 15, 583–596. Obel, N., Erben, V., and Pauly, M. (2006). Functional wall glycomics through oligosaccharide mass profiling. In The Science and Lore of the Plant Cell Wall: Biosynthesis,Structure and Function, Hayashi T., ed. (Boca Raton, FL: Brownwater), pp. 258–266. Pauly, M., Albersheim, P., Darvill, A., and York, W.S. (1999a). Molecular domains of the cellulose/xyloglucan network in the cell walls of higher plants. Plant J. 20, 629–639. Pauly, M., Andersen, L.N., Kauppinen, S., Kofod, L.V., York, W.S., Albersheim, P., and Darvill, A. (1999b). A xyloglucan-specific endo-beta-1,4-glucanase from Aspergillus aculeatus: expression cloning in yeast, purification and characterization of the recombinant enzyme. Glycobiol. 9, 93–100. Pauly, M., Qin, Q., Greene, H., Albersheim, P., Darvill, A., and York, W.S. (2001). Changes in the structure of xyloglucan during cell elongation. Planta. 212, 842–850. Perrin, R.M., Jia, Z.H., Wagner, T.A., O’Neill, M.A., Sarria, R., York, W.S., Raikhel, N.V., and Keegstra, K. (2003). Analysis of

Richmond, T.A., and Somerville, C.R. (2001). Integrative approaches to determining Csl function. Plant Mol. Biol. 47, 131–143.

Selvendran, R.R., and O’Neill, M.A. (1987). Isolation and analysis of cell walls from plant material. In Methods of Biochemical Analysis, David G., ed. (Hoboken, NJ: John Wiley & Sons), pp. 25–153. Somerville, C., et al. (2004). Toward a systems approach to understanding plant-cell walls. Science. 306, 2206–2211. Vanzin, G.F., Madson, M., Carpita, N.C., Raikhel, N.V., Keegstra, K., and Reiter, W.D. (2002). The mur2 mutant of Arabidopsis thaliana lacks fucosylated xyloglucan because of a lesion in fucosyltransferase AtFUT1. Proc. Natl Acad. Sci. U S A. 99, 3340–3345. Willats, W.G.T., et al. (2001). Modulation of the degree and pattern of methyl-esterification of pectic homogalacturonan in plant cell walls: implications for pectin methyl esterase action, matrix properties, and cell adhesion. J. Biol. Chem. 276, 19404–19413. Willats, W.T., Steele-King, C.G., McCartney, L., Orfila, C., Marcus, S.E., and Knox, J.P. (2000). Making and using antibody probes to study plant cell walls. Plant Physiol. Biochem. 38, 27–36. Yamagaki, T., Mitsuishi, Y., and Nakanishi, H. (1998). Determination of structural isomers of xyloglucan octasaccharides using postsource decay fragment analysis in MALDI–TOF mass spectrometry. Tetrahedron Letters. 39, 4051–4054. York, W.S., Darvill, A.G., McNeil, T., Stevenson, T.T., and Albersheim, P. (1985). Isolation and characterization of plant cell walls and cell wall components. Meth. Enzymol. 118, 3–40. Zablackis, E., Huang, J., Muller, B., Darvill, A.G., and Albersheim, P. (1995). Characterization of the cell-wall polysaccharides of Arabidopsis thaliana leaves. Plant Physiol. 107, 1129–1138. Zablackis, E., York, W.S., Pauly, M., Hantus, S., Reiter, W.D., Chapple, C.C.S., Albersheim, P., and Darvill, A. (1996). Substitution of L-fucose by L-galactose in cell walls of Arabidopsis mur1. Science. 272, 1808–1810.

Copyright of Molecular Plant is the property of Oxford University Press / UK and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.