SEARCH

SEARCH BY CITATION

Keywords:

  • Arabidopsis thaliana;
  • cell wall ultrastructure;
  • cinnamoyl-CoA reductase;
  • laser capture microdissection;
  • lignin;
  • quantitative immunogold labeling

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results and Discussion
  6. Acknowledgements
  7. References
  • • 
    A cinnamoyl-CoA reductase 1 knockout mutant in Arabidopsis thaliana was investigated for the consequences of lignin synthesis perturbation on the assembly of the cell walls.
  • • 
    The mutant displayed a dwarf phenotype and a strong collapse of its xylem vessels corresponding to lower lignin content and a loss of lignin units of the noncondensed type. Transmission electron microscopy revealed that the transformation considerably impaired the capacity of interfascicular fibers and vascular bundles to complete the assembly of cellulose microfibrils in the S2 layer, the S1 layer remaining unaltered. Such disorder in cellulose was correlated with X-ray diffraction showing altered organization.
  • • 
    Semi-quantitative immunolabeling of lignins showed that the patterns of distribution were differentially affected in interfascicular fibers and vascular bundles, pointing to the importance of noncondensed lignin structures for the assembly of a coherent secondary wall.
  • • 
    The use of laser capture microdissection combined with the microanalysis of lignins and polysaccharides allowed these polymers to be characterized into specific cell types. Wild-type A. thaliana displayed a two-fold higher syringyl to guaiacyl ratio in interfascicular fibers compared with vascular bundles, whereas this difference was less marked in the cinnamoyl-CoA reductase 1 knockout mutant.

Abbreviations: 
CCR

cinnamoyl-CoA reductase

CMFs

cellulose microfibrils

CP/MAS

cross-polarization/magic angle spinning

G

guaiacyl unit

GS

mixed guaiacyl–syringyl structures

H

p-hydroxyphenyl unit

ifs

interfascicular fibers

KL

Klason lignin

LCM

laser capture microdissection

S

syringyl unit

SEM

scanning electron microscopy

TEM

transmission electron microscopy

TFA

trifluoroacetic acid

vbs

vascular bundles

Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results and Discussion
  6. Acknowledgements
  7. References

The precise contribution of lignins in the assembly of plant cell walls has not been elucidated completely. However, it is widely accepted that, during differentiation, the composition and structure of lignins have a specific impact on the assembly and structural cohesion of the secondary walls according to cell type (Joseleau & Ruel, 1997). The study of mutant plants altered in the biosynthesis of any one of the wall polymers offers promising opportunities to obtain an insight into plant cell wall macromolecular assembly and organization. The monolignols originate from the phenylpropanoid pathway, which includes many steps that are potential targets for the modification of lignin biosynthesis (Baucher et al., 1998; Dixon et al., 2001). Many transgenic or mutant plants affected in the expression of the corresponding genes have been studied (reviewed in Anterola & Lewis, 2002). Cinnamoyl-CoA reductase (CCR) is the first enzyme specific to the monolignol pathway (Lacombe et al., 1997). This enzyme has been shown to catalyze the NADPH-dependent conversions of p-coumaroyl-CoA, feruloyl-CoA and sinapoyl-CoA into the corresponding aldehydes. By analyzing CCR-down-regulated tobacco plants (Piquemal et al., 1998; Ralph et al., 1998), a 50% reduction in the lignin content, an increase in the syringyl to guaiacyl (S : G) ratio and an increase in the amounts of cell wall-bound phenolics, such as ferulic and sinapic acids, were observed. A higher concentration of chlorogenic acid and of other soluble phenolics was also observed in tobacco (Chabannes et al., 2001a). The plants with the lowest CCR activity presented a dwarf size and a collapsed vessel phenotype, accompanied by alterations of lignin deposition and of wall cohesion (Chabannes et al., 2001b; Pinçon et al., 2001). Lignins from these transgenic tobacco plants released lower yields of thioacidolysis monomers, which indicates that they were enriched in resistant interunit bonds, referred to as condensed bonds in the following (Piquemal et al., 1998; O’Connell et al., 2002).

Arabidopsis thaliana is widely considered to be a model plant for understanding lignin synthesis, deposition and function (Boudet, 2000; Chaffey et al., 2002; Goujon et al., 2003b; Laskar et al., 2006). The first mutant for CCR1 in A. thaliana was named irx4 (Jones et al., 2001) because of its irregular xylem. CCR1-down-regulated lines were obtained by the antisense strategy (Goujon et al., 2003a), and showed an important reduction in lignin content and modifications of lignin structure.

Most of these previous investigations have studied the impact of genetic modifications in plants using biochemical approaches and light microscopy. As the genetic transformations modify the structural components of the cell wall, it seems necessary to investigate the resulting modifications with the resolution of electron microscopy appropriate to the scale of the elementary cellulose microfibrils (CMFs). Previous ultrastructural investigations have demonstrated that lignin modification could have characteristic consequences on cell wall assembly (Chabannes et al., 2001b; Ruel et al., 2001, 2002).

Two knockout mutants for CCR1 have been identified recently. Both have a dwarf phenotype, delayed senescence, reduced lignin content, some alterations in lignin structure and a dramatically altered pool of soluble phenolics (Mir Derikvand et al., 2008). In this work, we examined the effects of the mutation on the morphology of the plant, and investigated the ultrastructural phenotype of the interfascicular fibers (ifs) and xylem vascular bundles (vbs). As the mutation originates from a gene related to lignin biosynthesis, we analyzed the distribution of lignin structures in planta using immunological probes directed against various lignin units and structures. One interest of these probes against lignins is in their capacity to discriminate between the principal lignin units and to show specificity for the type of linkage (either condensed or noncondensed) in which they are involved in the polymer (Ruel et al., 1994). Preparative laser capture microdissection (LCM) (Matsunaga et al., 1999; Asano et al., 2002), coupled to microanalysis (Angeles et al., 2006; Nakashima et al., 2008), was implemented to study the specific composition of ifs and vbs of A. thaliana floral stems. This led to the characterization of lignification in individual cell types, and also to the evaluation of the tissue-specific impact of genetic modifications.

Materials and Methods

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results and Discussion
  6. Acknowledgements
  7. References

Plant material

The Arabidopsis thaliana (L.) Heynh. ecotype Columbia (Col-0) control line and the ccr1 mutant plant (ccr1g; Mir Derikvand et al., 2008) in the same ecotype were grown together in the same glasshouse (20°C, 60% relative humidity) to ensure the same environmental conditions. For polysaccharide analyses, ultrastructural and microdissection experiments, we used the floral stems at the complete flowering stage, corresponding to stage 6.90 (Boyes et al., 2001). Lignin analyses were performed on floral stems at complete maturity. Both lignin and polysaccharide analyses were carried out on extract-free samples after milling and solvent extraction (toluene–ethanol, 2 : 1 v/v, ethanol and H2O).

Lignin and polysaccharide analyses

The lignin content was estimated by the gravimetric Klason procedure (Dence, 1992). The lignin structure was investigated using thioacidolysis (Lapierre et al., 1995).

For total carbohydrate composition, the procedure with 72% H2SO4 followed by 2 m H2SO4, adapted from Saeman et al. (1954), was used. The released monosaccharides were converted to their alditol-acetate derivatives (Chambat et al., 1997) and analyzed by gas–liquid chromatography using an Agilent 6850 Series GC System equipped with an SP 2380 macrobore column (25 m × 0.53 mm) (Agilent Technologies, Palo Alto, CA, USA).

Selective hydrolysis of the noncellulosic wall polysaccharides was carried out with 2 m trifluoroacetic acid (TFA) (0.5 ml; 120°C; 4 h). After evaporation of the excess acid, the dry carbohydrates were converted to their alditol-acetate derivatives and analyzed as above.

Cross-polarization/magic angle spinning (CP/MAS) 13C-NMR spectra

In order to avoid grinding, the stem segments were hand-sectioned and introduced as such in the NMR cell. The solid-state NMR experiments were performed according to Heux et al. (1999) on a Bruker (Karlsruhe, Germany) MSL spectrometer operated at a 13C frequency of 25 MHz using combined proton decoupling, MAS and CP. The spinning speed was 3000 Hz, the contact time was 1 ms, the acquisition time was 70 ms and the recycled delay was 4 s. About 10 000 scans were accumulated for each sample.

Preparative LCM coupled to chemical microanalyses

Stem segments (2–3 cm in length) from A. thaliana wild-type (Col-0) and ccr1g lines were fixed in 0.2% glutaraldehyde–4% paraformaldehyde in 0.05 m phosphate buffer, pH 7.0, washed in phosphate buffer and then trimmed to a final length of 10–15 mm. Embedding was performed at −20°C in Tissue-Tek 4583 OCT compound. The material was cut into sections, 10 µm thick, with a cryostat (MICROM, Waldorf, Germany), and then mounted onto membrane-coated slides (Palm Micro-laser Technology, Bernried, Germany). Just before dissection, the sections were gradually dehydrated in increasing concentrations of ethanol (50, 70, 95, 100%, 15 min at each step), washed for 15 min in toluene and dried overnight at room temperature.

For laser microdissection, a Palm MicroBeam System (Palm Micro-laser Technology) was employed as described previously (Angeles et al., 2006). The target cells were cut using the UV laser, and then collected by catapulting with a single laser pulse into the cap of a 0.5 ml microcentrifuge tube containing a drop of distilled water. Pure ifs and vbs were separated. About 100 catapulted tissue fragments were sufficient for thioacidolysis and GC-MS analyses of the lignin-derived monomers.

Thioacidolysis of the LCM isolated materials was run by carefully transferring the tissue fragments into a Teflon-lined, screw-capped glass tube with water, and then freeze–drying the sample before the addition of 5 ml of thioacidolysis reagent, together with 1840 ng of C19 and 1980 ng of C21 internal standards (added in CH2Cl2 solution). Thioacidolysis was run as described previously (Lapierre et al., 1995) and the lignin-derived products were solubilized in 50 µl of CH2Cl2. The whole extract was then silylated by 50 µl of N,O-bis-trimethylsilyl-trifluoroacetamide and 5 µl of pyridine before injection onto a GC-MS (Varian 4000 (Palo Alto, CA, USA), ion trap, 1 µl injected). GC separation was performed with a Supelco (Saint-Germain-en-Laye, France) SPB1 column (25 m × 0.25 mm, 0.25 µm film thickness) operated in the temperature program mode (from 40 to 180°C at +30°C min−1; then 180 to 260°C at +2°C min−1), with helium as the carrier gas (constant flow rate of 1 ml min−1), an injector at 270°C operating in the split/splitless mode and a transfer line to MS at 280°C. The mass spectral analyses were run with an ion trap (electronic impact, 70 eV). All GC-MS analyses were carried out as duplicate injection analyses, whereas only one thioacidolysis was run per sample as sample collection was very time-consuming (100 collections per tissue type). The GC-MS determinations of the p-hydroxyphenyl (H), G and S lignin-derived monomers were carried out on ion chromatograms reconstructed at m/z 239, 269 and 299 (base peaks of their silylated derivatives). Thioacidolysis blank experiments were carried out in the same way, but without any tissue sample.

For polysaccharide hydrolysis, the tissues were transferred to conical centrifuge tubes with ethanol. Lipids were extracted with a chloroform–methanol mixture (1 : 1, v/v), and the tissues were soaked in an ethanol–toluene mixture (1 : 1, v/v, 50°C) to extract pigments. At each step, the solvents were carefully removed with a glass Pasteur pipette whose tip had been made thinner by drawing above the flame of a burner. The extraction was repeated at 50°C. Starch and cytoplasmic soluble sugars were extracted with hot water (95°C, 30 min). The resulting crude cell wall material was then submitted to sulfuric acid hydrolysis, as in Angeles et al. (2006).

Scanning electron microscopy (SEM)

Stems were sectioned with a razor blade (sections cut to c. 3 mm thick) and dehydrated through an ascending series of ethanol, ending in 100% ethanol. Then, the ethanol inside the stems was replaced by CO2 making a critical point. Before SEM examination, the stems were coated with a very thin film of gold–palladium and then observed with a JEOL (Tokyo, Japan) JSM 6100.

Transmission electron microscopy (TEM) procedures

Fixation and embedding  Small slices (1 mm thick), taken from the base of the stem by freehand sectioning with a razor blade, were fixed in a freshly prepared mixture of 0.2% glutaraldehyde (v/v), 2% paraformaldehyde (w/v) in 0.05 m phosphate buffer (pH 7–7.4), and processed as discussed in Ruel et al. (1994), before embedding in LR White resin.

Cytochemical staining for polysaccharides  The polysaccharide moiety of the walls was contrasted on ultrathin sections by the periodic acid–thiocarbohydrazide–silver proteinate method (Thiery, 1967), modified by Ruel et al. (1981). Briefly, periodate oxidation used a 5% solution of periodic acid in water for 90 min, followed by soaking in thiocarbohydrazide for 48 h and then silver proteinate (30 min).

Immunocytochemistry analysis  Five polyclonal antibodies directed against specific lignin structures were used: S/CZt directed against noncondensed mixed GS lignin structures; SZt directed against noncondensed homo-syringyl lignin structures (S) (Joseleau et al., 2004a); GZl directed against condensed lignin homo-guaiacyl structures (G); GSZl directed against condensed mixed GS lignin structures (Ruel et al., 1994; Joseleau & Ruel, 1997); and an antibody directed against dibenzodioxocin structures (Kukkola et al., 2003). The samples were immunolabeled on ultrathin transverse sections (50 nm) floating on plastic rings, as described previously (Joseleau & Ruel, 1997). Briefly, the sections were incubated on 50 µl droplets of diluted antisera (1 : 50 to 1 : 120). The secondary marker was Protein A-gold (pA G5, TEBU, Le Perray en Yvelines, Fance). The gold particles were further enhanced using a silver-enhancement kit (Amersham Bioscience). Finally, the thin sections were transferred to carbon-coated copper grids, post-stained with 2.5% aqueous uranyl acetate and examined with a Philips CM 200 Cryo-TEM at an accelerating voltage of 80 kV.

All comparative immunolabeling experiments were carried out in parallel in order to maintain the same experimental conditions. Pre-immune serum for each antibody was assayed as for the immunogold labeling. For quantitative estimation of the labeling, sections of the control plant were used as reference for each antiserum. Counting results reflect the means obtained from a minimum of 10 images. The number of gold particles per square micrometer was counted, and the relative labeling index was calculated. A value of 1.00 was assigned to the highest labeled sublayer in the control, and that for each antiserum, as it is not possible to compare the values given by two different antisera (Ruel et al., 2001; Mayhew et al., 2004).

X-Ray diffraction experiment

Small segments of fresh floral stem were used. X-Ray diffraction experiments were carried out using a point focus beam from an X-ray tube operated at 30 kV and 20 mA. The CuKa line filtered with nickel foil was collimated with two pinholes before the sample. The diffraction was recorded on an imaging plate (Fujifilm type FDL), which was subsequently read using an imaging plate reader BAS 1800II. The wet stem samples were positioned vertical to the incident beam, and the diffraction was shown with the fiber axis vertical.

Results and Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results and Discussion
  6. Acknowledgements
  7. References

Cell wall compositional analysis of the mature floral stems of the wild-type (Col-0) and mutant (ccr1g) lines

In agreement with previous results obtained on CCR1-deficient A. thaliana lines (Jones et al., 2001; Goujon et al., 2003a; Mir Derikvand et al., 2008), the extract-free stems from the ccr1g mutant displayed a substantially lower Klason lignin (KL) level than the control sample (Table 1).

Table 1.  Lignin content and structure in the extract-free mature stems of Arabidopsis thaliana wild-type Col-0 and mutant ccr1g
LineKlason lignin (KL) (wt%)Total yield H + G + SFerulic acidG-CHR-CHR2H (%)G (%)S (%)
  1. The KL content is expressed as the weight percentage of the extract-free stems. The lignin structure was evaluated by thioacidolysis. The yields of lignin-derived p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) monomers, from thioacidolysis-released ferulic acid, and of the G marker compound for ferulic acid incorporation into lignins (G-CHR-CHR2 with R = SEt) are expressed in micromoles per gram of KL. Standard errors between duplicate analyses are in the 5–10% range.

Col-015.0611271.81.00.878.121.3
ccr1g13.76 8257.14.70.671.627.8

The lignin structure was evaluated by thioacidolysis (Lapierre et al., 1995). The H, G and S lignin units only involved in β-O-4 bonds give rise to thioethylated H, G and S monomers, respectively. Therefore, when calculated on the basis of the KL content, the recovery yield of these diagnostic monomers is a reflection of the frequency of β-O-4 bonds in lignins. The total yield of thioacidolysis monomers released by the lignins of the extract-free ccr1g stem was lower than that of the control (Table 1). This lower thioacidolysis yield (by 50%) is diagnostic for a higher frequency of resistant interunit bonds (referred to as condensed bonds in the following). Such a higher condensation degree has been reported previously in ASCCR lines (Goujon et al., 2003a). In both the mutant and wild-type, these β-O-4-linked units are mainly G, accompanied by a smaller amount of S and a trace amount of H. In addition, CCR deficiency systematically induced a larger amount of the thioacidolysis marker for ferulic acid incorporation into lignins (Table 1) (Ralph et al., 2008).

Impact of the ccr1g mutation on the carbohydrate status of the cell walls

To evaluate the possible change in the hemicellulose fraction, two types of hydrolysis were performed. TFA, which hydrolyzes only the noncellulosic polysaccharides (Albersheim et al., 1967), revealed minor variations of composition within the hemicellulose moiety, with a slightly higher proportion of xylose (Table 2). This result is in line with that reported for the irx4 mutant (Jones et al., 2001).

Table 2.  Noncellulosic polysaccharide composition in the extract-free mature stems of Arabidopsis thaliana wild-type (WT) Col-0 and ccr1g mutant lines
LineFucoseArabinoseXyloseMannoseGalactoseGlucose
  1. Values are relative molar proportions (standard deviation).

WT Col-08.5 (0.6)7.8 (0.6)58.5 (0.1)5.5 (0.1)11.3 (0.3)8.1 (0.4)
ccr1g9.0 (0.8)9.2 (0.3)61.5 (1.0)4.4 (0.1) 9.5 (0.4)6.0 (0.4)

The total polysaccharide composition, including the resistant cellulose fraction, was obtained by concentrated sulfuric acid hydrolysis (Saeman et al., 1954). Considering that glucose and xylose reflect cellulose and hemicellulose proportions, respectively, the results reported in Table 3 reveal a higher proportion of hemicelluloses in the walls of ccr1g and a lower proportion of cellulose. These results are reminiscent of the data reported for irx4 plants, in which the cellulose content in mature stems was slightly lower and the hemicellulose content was higher (Jones et al., 2001), but differ from those observed in CCR-down-regulated poplar, in which the proportion of hemicellulose was reduced and that of cellulose was increased (Lepléet al., 2007).

Table 3.  Total cell wall carbohydrate composition in the extract-free mature stems of Arabidopsis thaliana wild-type (WT) Col-0 and ccr1g mutant lines
LineArabinoseXyloseMannoseGalactoseGlucoseGlucose/xylose
  1. Values are relative molar proportions (standard deviation).

WT Col-04.3 (0.3)31.3 (2.0)3.9 (0.2)3.4 (0.3)57.6 (1.8)1.8
ccr1g6.5 (0.2)41.5 (2.0)3.0 (0.1)2.9 (0.2)45.7 (1.3)1.1

Cell wall ultrastructure in the floral stems of the ccr1g mutant

As initially described in tobacco (Piquemal et al., 1998), CCR down-regulation induced changes in general growth. Our ccr1g mutant displayed a dwarf phenotype, with a floral stem height decreased by c. 50% (Mir Derikvand et al., 2008). The micromorphological repercussions of this mutation were first studied by SEM. Basal segments taken from control stems showed good internal cohesion of the walls of ifs (Fig. 1a) and vessels in vbs (Fig. 1c). By contrast, ccr1g displayed slightly collapsed ifs (Fig. 1b) and strongly collapsed xylem vessels (Fig. 1d). It should be noted that the dehydration step involved in the sample preparation for SEM caused a separation within the secondary wall (arrows) in the mutant. The phenomenon affected mostly ifs, but also some of the vessels from vbs. This loss of cohesion constitutes a first indication of the mechanical weakness of the cell walls of the mutant plants, and was also evidenced in longitudinal sections of ifs, which exhibited a loosened internal network of fibrils, contrary to the control plant (Fig. 1e,f). The collapsed vessel phenotype has been repeatedly observed in transgenic or mutant plants with decreased CCR activity (Piquemal et al., 1998; Chabannes et al., 2001b; Jones et al., 2001; Goujon et al., 2003a). In addition, the mutation induced an apparent diminution of the cell wall thickness compared to the wild-type. There is therefore a clear correlation between morphological alterations and modifications of composition of the secondary cell walls.

image

Figure 1. Scanning electron microscopic images of the main tissues of the mature floral stems of control Col-0 (a, c, e) and ccr1g (b, d, f) lines of Arabidopsis thaliana. (a–d) Transverse sections: (a, b) interfascicular fibers (ifs); note, in ccr1g, the detachment of the secondary wall resulting from shrinkage during dehydration; (c, d) vascular bundle (vb); collapse of the vessel wall (v) is evident in ccr1g (d). (e, f) Longitudinal sections; the fibrillar organization of the secondary wall induced in ccr1g is apparent.

Download figure to PowerPoint

To obtain a more precise picture of the impact of the mutation on the organization of the cell wall, investigation at the scale of resolution of TEM is required. The global architecture of the cell wall was examined using the periodic acid–thiocarbohydrazide–silver proteinate cytochemical method adapted to secondary walls (Ruel et al., 1981), which stains the majority of polysaccharides, including cellulose, and therefore provides images of the micromorphological arrangement. At low magnification, the secondary walls of the control stems (Fig. 2a) are organized and compact, and display good cohesion. By contrast, the mutant exhibited dramatic disorganization and loosening of the secondary walls (Fig. 2b). Higher magnification evidenced the loss of the ability of CMFs to assemble into a coherent framework in ifs and vessels from the mutant plant. Although the collapse appears to be most pronounced in the vessels, it also affects the supporting fibers (Fig. 2b–d). It is also noteworthy that the loosening mostly concerns the S2 layer, particularly its inner part, which exhibits an important delamination of the CMFs, accompanied by an apparent loss of the orientation of the cellulose network (Fig. 2d,f). By contrast, the S1 layer appears to be normal. The juxtaposition of the compact S1, against the loosened S2, may explain the shrinkage observed in SEM. It is interesting to note that similar shrinkage of the gelatinous layer of fibers from tension wood, in which the lignin content is very low, also occurs in the same conditions of dehydration (Clair & Thibaut, 2001; Joseleau et al., 2004b). The fact that the impact of the mutation affects primarily S2, leaving S1 apparently unmodified, suggests that the ccr1g gene is more specifically involved in the lignification of S2.

image

Figure 2. Ultrastructural organization of the interfascicular fibers (ifs) and vascular bundles (vbs) of the basal part of the mature floral stems of Arabidopsis thaliana Col-0 (a, c, e) and ccr1g (b, d, f). Periodic acid–thiocarbohydrazide–silver proteinate staining in transmission electron microscopy (TEM). (a, b) Low magnification showing the collapse of vessels and fibers in ccr1g vbs. Note that the three ‘classical’ sublayers of the secondary walls, S1, S2 and S3, are present. A loosening is evident in both fibers (f) and vessels (v). (c, d) Details of if walls; in ccr1g, the modification of the walls is restricted to S2, and S1 remains compact. (e, f) Vessels walls are strongly collapsed in ccr1g; S1 remains unchanged. S1, first deposited secondary wall sublayer; S2, secondary wall medium sublayer; S3, last deposited. Bars, 1 µm.

Download figure to PowerPoint

Combining the results from the structural and ultrastructural analyses, it appears that the mutation of CCR1 in A. thaliana has the same type of effect as that shown in the ASCCR lines of tobacco (Chabannes et al., 2001b) and Arabidopsis (Goujon et al., 2003a), although somewhat more pronounced.

X-Ray analysis

The perturbation in the orientation of cellulose observed in S2 was confirmed by the X-ray diffraction diagrams of control and ccr1g stems (Fig. 3). Because X-ray diffraction was recorded from the whole plant stem sections, the contribution of the different cell types with their diversity in cellulose and noncellulose polymer composition somewhat blurs the diagrams and the modifications that affect specific cells. Notwithstanding, the results show the typical patterns of cellulose with the clearly distinct spots corresponding to the 1-10/110 (5.8 Å) and 200 (3.9 Å) crystal planes of cellulose Ib (Nishiyama et al., 2003). The diagrams underscore the lower resolution of ccr1g compared with the Col-0 control. This suggests a loose aggregation and a lower degree of orientation of the CMFs in ccr1g, corresponding to the inability to properly associate CMFs in the S2 layer of fibers and vessels. Similar manifestation of disordered CMF orientation has been observed previously in the electron diffraction pattern from antisense CCR-down-regulated tobacco (Ruel 2007). This is coherent with the suggestion that Arabidopsis mutants show a link between cellulose synthesis and lignification (Cano-Delgado et al., 2003), and with the requirement of cellulose to nucleate lignin polymerization (Taylor et al., 1992). Such a relationship between the patterns of cellulose deposition and lignin assembly was originally suggested by the study of irregular xylem (irx) mutations (Turner & Somerville, 1997). Our results support the involvement of the CCR gene in the supramolecular arrangement of CMFs suggested by quantitative trait loci (QTL) mapping analysis (Thumma et al. 2005).

image

Figure 3. X-Ray diagrams of the floral stems of control Col-0 (a) and ccr1g mutant (b) Arabidopsis thaliana. Arrows indicate typical crystalline cellulose positions.

Download figure to PowerPoint

CP/MAS 13C-NMR spectra

The solid-state CP/MAS 13C-NMR spectra of Col-0 and ccr1g (Fig. 4) did not show significant differences between Col-0 and ccr1g corresponding to compositional variations. Similar observations were made in the study of the irx4 Arabidopsis mutant (Patten et al., 2005). Indeed, solid-state 13C-NMR gave a global picture of the wall constituents of the plant stems, but could not provide sufficient resolution to detect minor differences in lignin and carbohydrate composition, nor in cellulose crystallinity and ordering, because of the cell type specificity of the mutation, as shown by the TEM results (see below).

image

Figure 4. Cross-polarization/magic angle spinning (CP/MAS) 13C-NMR spectra of the floral stem tissues of control Col-0 and ccr1g mutant Arabidopsis thaliana.

Download figure to PowerPoint

Distribution of lignin structures in the floral stems of Col-0 and ccr1g lines by immunolabeling and TEM analyses

To analyze the patterns of lignification in the walls of the different cell types of Col-0 and ccr1g, we carried out immunolabeling in TEM using antibodies directed against noncondensed (Joseleau et al., 2004a) and condensed (Ruel et al., 1994; Joseleau & Ruel, 1997) lignin structures. Manual counting of the number of gold particles per square micrometer (Ruel et al., 2001; Mayhew et al., 2004) provided a semi-quantitative comparative estimation of the respective distribution of the epitopes within the ifs and vessels from vbs in the control and mutant A. thaliana plants, allowing a relative labeling index to be calculated for each antiserum (Table 4). It must be emphasized that, to ensure semi-quantitative comparisons, labeling experiments were carried out in parallel for the control and ccr1g samples using the same solution of the antibody. Comparing quantitative estimations of labeling provided by different antibodies is meaningless.

Table 4.  Semi-quantitative immunocytochemical analysis of the topochemical distribution of lignin substructure epitopes in the cell walls of Arabidopsis thaliana Col-0 and ccr1g mature stems
EpitopeCol-0 Fiber S1S2Col-0 Vessel S1S2S3CCR1 Fiber S1S2CCR1 Vessel S1S2S3
  1. nd, sublayer, nondifferentiated.

  2. The intensity of labeling was evaluated manually by the number of gold grains per square micrometer.

  3. Values in italics represent the relative labeling indices and were calculated by giving, for each antibody, the value 1.00 to the highest labeled sublayer in the wild-type sample Col-0. In this way, the variation in the intensity of labeling by each antiserum in each cell type was clearly apparent. Absolute values cannot be compared between different antisera. Only their variation is significant.

GZl7.54.55.03.13.26.310.211.27.9nd
Condensed1.000.600.660.410.430.85 1.361.501.05nd
GSZl3.54.14.92.03.61.2 1.11.20.80.8
Condensed0.710.841.000.820.730.25 0.220.250.160.16
S : G2.16.31.53.51.50 1.20.42.10.6
Noncondensed0.331.000.240.550.240 0.190.060.330.10
S3.410.70.93.22.00 0.3 000
Noncondensed0.321.000.080.300.190 0.03 000
Dibenzodioxocin2.53.31.81.40.30 0.7 000
 0.761.000.300.420.090 0.21 000

Labeling of noncondensed (β-O-4-linked) lignin structures

The noncondensed β-O-4 units were localized using the anti-S/C (Joseleau & Ruel, 2006) antibody, which specifically labels mixed GS, and anti-homosyringyl S (Joseleau et al., 2004a), with specificity for S units. These two probes were prepared from synthetic lignin polymers. The third probe, directed against the dibenzodioxocin structure, was prepared with the appropriate model compound (Kukkola et al., 2003). Labeling with the anti-S/C antibody revealed that these epitopes in Col-0 were particularly abundant in the S2 sublayers of ifs and present in smaller amounts in S1 (Fig. 5a). In the control vessels, the three sublayers S1, S2 and S3 were equivalently labeled (Fig. 5c). Clearly, middle lamellae and cell corner junctions were less labeled. In the mutant, the absence of CCR1 expression substantially reduced the general frequency of these epitopes in both ifs and vessels (Table 4). At the spatial level, the noncondensed GS units were restricted to the S2 sublayer in ifs (Fig. 5b), and to the S2 and S3 sublayers in the vessels, with a very low density in the cell corners (Fig. 5d). These results reveal that, not unexpectedly, the CCR1 null mutation has similar, but more pronounced, effects than those previously observed in CCR1-down-regulated plants (Chabannes et al., 2001b; Ruel et al., 2002; Goujon et al., 2003a).

image

Figure 5. Topochemical distribution of noncondensed lignin structures in the basal part of the mature floral stems of Col-0 (a, c, e, g) and ccr1g (b, d, f, h) Arabidopsis thaliana. (a–d) Noncondensed mixed guaiacyl–syringyl (GS) structures. In ccr1g, there is a significant reduction in mixed GS epitopes in both interfascicular fibers (ifs) (b-arrowheads) and vessel walls (d). Note that, in the normal plant Col-0, the noncondensed GS epitopes are abundant in S1 and S2. They are equally distributed in the vessel walls (v) of vascular bundles (vbs) and predominate in S2 of ifs. (e–h) Syringyl (S) epitopes. S epitopes have almost disappeared in ccr1g in both ifs, f (fibers), vessel walls and cc (cell corners). Bars, 1 µm.

Download figure to PowerPoint

Anti-S showed that, in Col-0, the distribution of gold particles corresponding to noncondensed homo-syringyl units varied between tissues and cell wall sublayers. In ifs, the highest concentration was in S2 (Fig. 5e). In vessels, the labeling was positive, but weaker than that observed in ifs, particularly in adjacent cell walls (not seen in the photographs). The noncondensed S epitopes appear to be concentrated in the S2 sublayer (Fig. 5g). By contrast, in the mutant plant, the immunolabeling showed that these lignin structures could hardly be detected in ifs (Fig. 5f) and vessels (Fig. 5h). These observations, together with the semi-quantitative immunolabeling analysis (Table 4), agree with the significant decrease in lignin content and with the thioacidolysis-revealed reduction in noncondensed lignin structures (Table 1). They further indicate which areas of the cell walls are more specifically concerned with the reduction in noncondensed lignin structures.

The topochemical distribution of the dibenzodioxocin epitope structures was carried out with the specific antibody described previously by Kukkola et al. (2003). In the control Col-0, the dibenzodioxocin structures were well represented in the secondary wall of ifs, particularly in S2 (Fig. 6a), which confirms that the end-wise lignin polymerization mode prevails in S2. In the vessels, the distribution was weak, but homogeneous in the three sublayers, with few gold particles in cell corners (Fig. 6c). The labeling in ifs of the mutant was very low (Fig. 6b), and almost not detected in the vessels (Fig. 6d). The dibenzodioxocin structures, which can be formed by end-wise oxidative coupling of coniferyl alcohol with a 5,5-biphenyl structure of the growing lignin polymer (Karhunen et al., 1995), can lead to branch points in the chain (Brunow et al., 1998; Ralph et al., 2004). The fact that these structures were considerably decreased in the ccr1g mutant is an indication that the mutation impacted the mode of polymerization of the monolignols. According to Brunow et al. (1998), the dibenzodioxocin structures are produced in the late phases of cell wall lignification. The low level of these structures in fibers, and their near absence in vessels, in the ccr1g mutant can be viewed as a manifestation of the occurrence of lignins more similar to early developmental stage lignins.

image

Figure 6. Topochemical distribution of noncondensed dibenzodioxocin dimers in the basal part of the mature floral stems of Col-0 and ccr1g Arabidopsis thaliana. (a, b) Interfascicular fibers (ifs); less gold particles in the loosened walls of ccr1g fibers. (c, d) Disappearance of the gold particles in vessels walls and fibers (f) of ccr1g. cc, cell corners. Bars, 1 µm.

Download figure to PowerPoint

The results obtained with the three preceding antibodies directed against structures formed by the end-wise coupling mechanism (i.e. β-O-4 and dibenzodioxocin bonding patterns) lead to the following conclusions. In the control A. thaliana Col-0, the particularly low intensity of immunolabeling in the S1 layer suggests that noncondensed lignin structures are deposited at the later stages of secondary wall thickening. This result is in agreement with literature data (Terashima et al., 1993) on the spatiotemporal deposition of lignins. The suppression of CCR1 expression produced the spatial distribution of lignin structures, indicating that, in addition to a reduced lignin content, the plant did not synthesize a normal proportion of noncondensed units, particularly S units.

Labeling of condensed lignin structures

Two antibodies were used for the labeling of condensed lignin structures: the anti-homoguaiacyl (GZl) and anti-mixed GS lignin (GSZl). The two probes revealed different patterns in ifs and vbs, respectively. In Col-0, condensed G units are abundant and regularly distributed in the different sublayers of ifs and vessels of vbs (Fig. 7a,c; Table 4). By contrast, condensed mixed GS lignins (Fig. 7e,g) are poorly represented (or accessible) in S1 and S2 of ifs. In vbs, the localization of these epitopes is restricted to S1 in fibers and to S1 and S3 in vessels, which appears to be a trait common to various plants (Ruel et al., 1999; Chabannes et al., 2001b; Goujon et al., 2003a).

image

Figure 7. Topochemical distribution of condensed lignin structures in the basal part of the mature floral stems of Col-0 and ccr1g Arabidopsis thaliana. (a–d) Homoguaiacyl lignin structures. (a, b) In interfascicular fibers (ifs), the gold particles are strongly distributed in both Col-0 and ccr1g, but their relative intensity in S2 and S2 are inverted between Col-0 and ccr1g (see Table 4). (c, d) Vascular bundles (vbs) show high concentrations of gold particles in S1 and S2 of Col-0 and ccr1g. These epitopes appear to be more accessible in the loosened wall of the mutant. (e–h) Condensed mixed guaiacyl–syringyl structures. (e, f) The labeling of ifs in the control Col-0 is reduced in ccr1g. (g, h) In vessel walls, condensed GS structures, which are more concentrated in S1 and S3, are reduced in ccr1g. cc, cell corner; f, fiber of the vbs. Bars, 1 µm.

Download figure to PowerPoint

In the ccr1g mutant, the density of condensed G epitopes appeared higher (see also Table 4), suggesting that the G residues became major units, contributing in the main to condensed lignins. From this, it can be deduced that the absence of CCR1 expression induced a mechanism of monolignol polymerization that favored the formation of condensed lignin structures. This confirms previous suggestions reported in Arabidopsis ASCCR lines (Ruel et al., 2002). In the case of the mixed condensed GS epitopes, CCR1 mutation induced a reduction in these lignin structures in ifs (Fig. 7f) as well as in the vessels (Fig. 7h; Table 4). This is a clear indication that the mutation modified the distribution patterns of the mixed condensed GS and G epitopes, suggesting that the mode of coupling of these units was differentially influenced by the mutation.

LCM and microanalysis of different tissues from floral stems of Col-0 and ccr1g lines

The results obtained in TEM, with the immunological probes, demonstrated that the alterations in the deposition of the corresponding lignin epitopes were largely tissue specific. However, immuno-TEM only allowed a semi-quantitative evaluation of the modifications. To obtain more precise quantitative information, and to overcome the averaging of the chemical analysis performed on the whole stem tissues, we implemented LCM (Matsunaga et al., 1999; Asano et al., 2002; Nakashima et al., 2008) to separate and isolate the interfascicular area and vbs. In this technique, the laser beam selectively cuts out the ifs (Fig. 8a–c) and vbs (Fig. 8d–f), allowing separate pure ifs and vbs to be collected (Fig. 8c,f).

image

Figure 8. Laser capture microdissection (LCM) of different tissues from the basal part of the mature stem of Arabidopsis thaliana ccr1g. (a–c) Isolation of interfascicular fibers (ifs). (d–f) Isolation of vascular bundles (vbs).

Download figure to PowerPoint

Polysaccharide microanalysis

Polysaccharide analyses using the whole stem tissues reflected global changes, regardless of the tissue-specific action of the genes involved in cell wall differentiation (Campbell & Sederoff, 1996; Demura & Fukuda, 2007). Here, the ifs and vbs of Col-0 and the ccr1g mutant were subjected to selective and total polysaccharide analyses (Angeles et al., 2006). As a result of the limited amount of collected material, only the relative proportions of the main cell wall sugars, glucose and xylose, could be established, allowing the evaluation of the relative proportions of cellulose and xylans in the walls (Fig. 9). The variation of the glucose to xylose ratio showed that the ccr1g mutation differentially affected vbs and ifs, but did not reveal in what way the respective cellulose and xylan syntheses had been affected. The TEM images showing the loosened microfibril organization and low compactness of the fiber wall support a defect in cellulose synthesis. This also correlates with the X-ray diffraction results of altered cellulose deposition. However, the increased glucose to xylose ratio of vbs, whose collapse indicates a decreased mechanical resistance, would better correlate with a diminution of xylans than with an augmentation of cellulose, in agreement with the contribution of xylans to cell wall cohesion and mechanical resistance known to be closely related to their arrangement relative to CMFs (Salmén, 2004; Lawoko et al., 2005; Zhong et al., 2005; Joseleau & Ruel, 2006).

image

Figure 9. Variation of glucose to xylose ratio in the walls of vascular bundles (VBs) and interfascicular fibers (iFs) from Arabidopsis thaliana Col-0 and the ccr1g mutant.

Download figure to PowerPoint

Lignin microanalysis

The lignin structure of the microdissected samples was evaluated by thioacidolysis from 100 collections per tissue type (ifs and vbs) recovered from the mature floral stems of A. thaliana. A satisfactory signal-to-noise ratio was obtained, as evidenced from the GC traces employed for the quantitative determination of the S and G lignin-derived monomers (Fig. 10). As the analyses of these monomers are in the range of trace analyses, we checked that blank experiments did not yield any lignin-derived monomers.

image

Figure 10. Partial GC-MS chromatograms showing the separation of the guaiacyl (G) and syringyl (S) thioacidolysis lignin-derived monomers recovered from vascular bundles (vbs) (a) and interfascicular fibers (ifs) (b) collected from 100 microdissections of the basal part of the mature floral stems of Arabidopsis thaliana Col-0.

Download figure to PowerPoint

Importantly, in the control samples, the relevance of the LCM–thioacidolysis combination was confirmed by the higher S : G molar ratio of ifs relative to vbs (Fig. 10; Table 5). Consistent with the results of Mäule histochemical staining (data not shown) and of immunolabeling (Fig. 5; Table 4) with the anti-S antibody, the frequency of thioacidolysis S monomers recovered from control samples was found to be twice as high in ifs relative to vbs. Contrary to the classical assertion that vessel walls are primarily formed from G units, S units were also present in vbs (= vessels + fibers), confirming the localization detected with the anti-S antibody (Fig. 5e,g). The total thioacidolysis yields obtained from the control Col-0 samples, and expressed in nanomoles per 100 microdissections, are given as indicative values, as we cannot certify that all 100 samples were actually recovered in the sampling tube. The yield (Table 5) and the surface area of the collected samples (data not shown) were found to be similar for ifs and vbs, suggesting that the lignin content of these two regions are not very different.

Table 5.  Molar ratio (S : G) and relative frequency (molar%) of the p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) lignin-derived monomers recovered by laser capture microdissection from the interfascicular fibers (ifs) and vascular bundles (vbs) of Arabidopsis thaliana wild-type (Col-0) and ccr1g mutant
Line and tissueS : GH (%)G (%)S (%)H + G + S
  1. The total yield (H + G + S) is expressed in nanomoles per 100 microdissections. Standard errors between two injections are indicated in parentheses.

Col-0 ifs0.64 (0.01)0.72 (0.11)60.7 (0.1)38.6 (0.1)3.02 (0.01)
Col-0 vbs0.27 (0.01)0.88 (0.03)78.3 (0.2)20.8 (0.2)2.53 (0.05)
ccr1g ifs0.51 (0.00)1.6 (0.2)64.8 (0.2)33.6 (0.5)1.98 (0.04)
ccr1g vbs0.39 (0.00)1.5071.6 (0.3)27.9 (0.3)0.51 (0.03)

In the mutant plant, the S : G ratio was also found to be higher in ifs than in vbs (Table 5). However, and compared with Col-0, the S : G ratio in ccr1g was higher in vbs and lower in ifs. This tissue-specific impact of the mutation could not be detected from the global analyses of the stem lignins. The thioacidolysis yields of the ccr1g ifs and vbs were substantially lower than the control levels. This reduction could be related to several parameters: the smaller surface area of the collected samples (data not shown), the lower lignin level (revealed by the global analyses) and/or the higher condensation degrees of ccr1g lignins. In agreement with the analysis of whole mature stems (Table 1), thioacidolysis of microdissected samples revealed that the ccr1g lignins released more H monomers than the control, regardless of the tissue (Table 5).

Conclusions

The impact of the ccr1g mutation in A. thaliana was analyzed by a series of methods and approaches adapted to the characterization of the cell wall microstructure of individual cell types. LCM coupled with microanalysis has proven its efficiency in the investigation of the heterogeneous distribution of lignins and polysaccharides in A. thaliana stems. Moreover, it allowed the quantification of the effects of the mutation affecting different cell types. The results provide evidence that caution must be exerted in the interpretation of global analyses of plants which average the contribution of heterogeneous tissues. This new methodology appears to be of particular interest for genetically modified plants in which the impact of the mutation is tissue specific (Nakashima et al., 2008). Although some improvements need to be made in the tedious sample recovery, the results reported herein confirm the potential of such a microstrategy for cell wall investigation.

Taken together, our results demonstrate that the CCR1 mutation in A. thaliana has a strong impact on the assembly of the secondary walls of fibers and vessels. The defect in CCR activity, which slightly reduces the level of lignin content, clearly modifies the ultrastructural distribution of the G and S units, with a severe alteration of the assembly of the lignified secondary cell walls. TEM observations demonstrate that the overall result of the mutation is to impair the capacity of the lignified walls (ifs and vessels) to achieve the correct orientation of the CMF network in the S2 layer, as confirmed by the altered X-ray diffraction pattern. The defect in the orientation and packing of the CMFs in S2 seems to be the indirect consequence of lignin and, perhaps, hemicellulose modifications in quantity and structure. We found a correlation between the defect in lignin synthesis and the capacity of organization of CMFs. In this respect, our results substantiate genetic analysis documented by the use of linkage disequilibrium mapping (Thumma et al., 2005), which suggested an influence of the CCR gene on CMF angle orientation. They also correlate with a developmental study in wheat, showing that CCR contributes to stem strength support (Ma, 2007), a mechanical property which is closely related to the value of the microfibril angle (Evans & Ilic, 2001). As the perturbation of the microfibril angle denotes a disorder in microtubule orientation, the latter would also correlate with an alteration in cellulose synthesis. Such a change in general carbohydrate metabolism was observed in CCR-down-regulated transformants and was suggested to be associated with a stress response (Lepléet al., 2007). The topochemical investigation points to the particular participation of noncondensed lignin structures in CMF assembly in S2. It is remarkable that, in ccr1g, the capacity of the S1 layer to assemble properly is hardly affected, particularly in ifs, contrary to S2. Our results indicate that the absence of CCR1 influences the mode of coupling, leading to noncondensed units, whether they involve G units only or mixed SG units. Thioacidolysis shows that 80% of the lignin in the ccr1g mutant is of the condensed type, and immunolabeling completes these results in showing that this condensed moiety, which cannot be fully characterized by chemical analysis, is predominantly made up of G units. It is noteworthy that, despite their differences in tissue type and in lignin topochemistry, ifs and vessels in the CCR1-depleted mutant are affected in a similar manner, indicating that the implication of ccr1 is essentially directed to S2 assembly regardless of the tissue. Beyond the description of the precise impact of the ccr1g mutation on the wall structure of A. thaliana, the present results underscore the fundamental difference in the process of assembly of S1 and S2 in the lignified if wall. Thus, in A. thaliana, S1 can form in a coherent manner in spite of a minimum proportion of noncondensed S lignin units, whereas the participation of these units seems fundamental for S2 to organize a normal spatial pattern of its cellulose framework. This suggests that, in the process of lignified secondary wall thickening, the CCR1 gene is more specifically involved in the assembly of S2.

All of these results point to the role of noncondensed lignin structures in the interaction between cellulose and lignin, in which the orientation of the lignin rings is guided by the orientation of cellulose, shown by Raman spectroscopy (Houtman & Atalla, 1995) and further demonstrated by molecular modeling (Besombes & Mazeau, 2005). Reminiscent of the inhibition of cellulose synthesis causing the abnormal deposition of lignin in zinnia cells (Taylor et al., 1992), we witness here, with the ccr1g mutant, the reverse situation, in which the alteration of lignin synthesis impedes cellulose organization. Together, our observations support the view that, during secondary wall assembly, the synthesis of one component mediates the patterning of the others (Taylor et al., 1992).

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results and Discussion
  6. Acknowledgements
  7. References

The authors express their thanks to the Department of Anatomy, Cytology and Pathology, CHU Grenoble, France, to Dr Ingo Burgert (Max-Planck-Institute of Colloids and Interfaces, Department of Biomaterials, Potsdam, Germany) and COST E50 (CEMARE) for financing the ‘Short Term Missions for LCM Facilities’. The authors are indebted to Dr Yoshiaru Nishiyama (CERMAV, Grenoble, France) for the X-ray diffraction analyses and helpful discussions, and to Michel Trierweiler (CERMAV, Grenoble, France) for recording the solid-state NMR spectra. They thank Danielle Dupeyre (CERMAV) for technical assistance in SEM studies. They are also very grateful to Bruno Letarnec (Biologie Cellulaire, INRA-Versailles, France) for his work in the glasshouse and to Frédéric Legée (UMR 206, AgroParisTech-INRA, Grignon, France) and Laurent Cézard (UMR 206, AgroParisTech-INRA, Grignon, France) for running the Klason determinations and some thioacidolysis experiments. This research was performed in part during J.B.-S.'s PhD thesis supported by a Marie-Curie Grant and by the Institut National de la Recherche Agronomique (INRA)-CEPIA Department, France.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results and Discussion
  6. Acknowledgements
  7. References
  • Albersheim P, Nevins DJ, English PD, Karr A. 1967. A method for the analysis of sugars in plant cell-wall polysaccharides by gas–liquid chromatography. Carbohydrate Research 5: 340345.
  • Angeles G, Berrio-Sierra J, Joseleau JP, Lorimier P, Lefebvre A, Ruel K. 2006. Preparative laser capture microdissection and single-pot cell wall material preparation: a novel method for tissue-specific analysis. Planta 224: 228232.
  • Anterola AM, Lewis NG. 2002. Trends in lignin modification: a comprehensive analysis of the effects of genetic manipulations/mutations on lignification and vascular integrity. Phytochemistry 61: 221294.
  • Asano T, Masumura T, Kusano H, Kikuchi S, Kurita A, Shimada H, Kadowaki K. 2002. Construction of a specialized cDNA library from plant cells isolated by laser capture microdissection: toward comprehensive analysis of the genes expressed in the rice phloem. Plant Journal 32: 401408.
  • Baucher M, Monties B, Van Montagu M, Boerjan W. 1998. Biosynthesis and genetic engineering of lignin. Critical Reviews in Plant Sciences 17: 125197.
  • Besombes S, Mazeau K. 2005. The cellulose/lignin assembly assessed by molecular modeling. Part 2: Seeking for evidence of organization of lignin molecules at the interface with cellulose. Plant Physiology and Biochemistry 43: 277286.
  • Boudet A-M. 2000. Lignins and lignification: selected issues. Plant Physiology and Biochemistry 38: 8196.
  • Boyes DC, Zayed AM, Ascenzi R, McCaskill AJ, Hoffman NE, Davis KR, Gorlach J. 2001. Growth stage-based phenotypic analysis of Arabidopsis: a model for high throughput functional genomics in plants. Plant Cell 13: 14991510.
  • Brunow G, Kilpelainen I, Sipila J, Syrjanen K, Karhunen P, Setala H, Rummakko P. 1998. Oxidative coupling of phenols and the biosynthesis of lignin. In: LewisNG, SarkanenS, eds. Lignin and lignan biosynthesis. American Chemical Society Symposium Series 697. Washington, DC, USA: American Chemical Society, 131147.
  • Campbell MM, Sederoff RR. 1996. Variation in lignin content and composition – mechanism of control and implications for the genetic improvement of plants. Plant Physiology 110: 313.
  • Cano-Delgado A, Penfield S, Smith C, Catley M, Bevan M. 2003. Reduced cellulose synthesis invokes lignification and defense responses in Arabidopsis thaliana. Plant Journal 34: 351362.
  • Chabannes M, Barakate A, Lapierre C, Marita JM, Ralph J, Pean M, Danoun S, Halpin C, Grima-Pettenati J, Boudet AM. 2001a. Strong decrease in lignin content without significant alteration of plant development is induced by simultaneous down-regulation of cinnamoyl CoA reductase (CCR) and cinnamyl alcohol dehydrogenase (CAD) in tobacco plants. Plant Journal 28: 257270.
  • Chabannes M, Ruel K, Yoshinaga A, Chabbert B, Jauneau A, Joseleau JP, Boudet AM. 2001b. In situ analysis of lignins in transgenic tobacco reveals a differential impact of individual transformations on the spatial patterns of lignin deposition at the cellular and subcellular levels. Plant Journal 28: 271282.
  • Chaffey N, Cholewa E, Regan S, Sundberg B. 2002. Secondary xylem development in Arabidopsis: a model for wood formation. Physiologia Plantarum 114: 594600.
  • Chambat G, Cartier N, Lefebvre A, Marais MF, Joseleau JP. 1997. Changes in cell wall and extracellular polysaccharides during the culture cycle of Rubus fruticosus cells in suspension culture. Plant Physiology and Biochemistry 35: 655664.
  • Clair B, Thibaut B. 2001. Shrinkage of the gelatinous layer of poplar and beech tension wood. Iawa Journal 22: 121131.
  • Demura T, Fukuda H. 2007. Transcriptional regulation in wood formation. Trends in Plant Science 12: 6470.
  • Dence CW. 1992. The determination of lignin. In: LinSY, DenceCW, eds. Methods in lignin chemistry. Heidelberg, Germany: Springer-Verlag, 3361.
  • Dixon RA, Chen F, Guo D, Parvathi K. 2001. The biosynthesis of monolignols: a ‘metabolic grid’, or independent pathways to guaiacyl and syringyl units? Phytochemistry 57: 10691084.
  • Evans R, Ilic J. 2001. Rapid prediction of wood stiffness from microfibril angle and density. Forest Products Journal 51: 5357.
  • Goujon T, Ferret V, Mila I, Pollet B, Ruel K, Burlat V, Joseleau JP, Barriere Y, Lapierre C, Jouanin L. 2003a. Down-regulation of the AtCCR1 gene in Arabidopsis thaliana: effects on phenotype, lignins and cell wall degradability. Planta 217: 218228.
  • Goujon T, Sibout R, Eudes A, MacKay J, Joulanin L. 2003b. Genes involved in the biosynthesis of lignin precursors in Arabidopsis thaliana. Plant Physiology and Biochemistry 41: 677687.
  • Heux L, Dinand E, Vignon MR. 1999. Structural aspects in ultrathin cellulose microfibrils followed by 13C CP-MAS NMR. Carbohydrate Polymers 40: 115124.
  • Houtman CJ, Atalla RH. 1995. Cellulose–lignin interactions. A computational study. Plant Physiology 107: 977984.
  • Jones L, Ennos AR, Turner SR. 2001. Cloning and characterization of irregular xylem4 (irx4): a severely lignin-deficient mutant of Arabidopsis. Plant Journal 26: 205216.
  • Joseleau JP, Ruel K. 1997. Study of lignification by noninvasive techniques in growing maize internodes. An investigation by Fourier transform infrared cross-polarization-magic angle spinning 13C-nuclear magnetic resonance spectroscopy and immunocytochemical transmission electron microscopy. Plant Physiology 114: 11231133.
  • Joseleau JP, Ruel K. 2006. New insights on the occurrence and role of lignin in the secondary wall assembly. In: HayashiT, ed. The science and lore of the plant cell wall; biosynthesis, structure and function. Boca Raton, FL, USA: Brown Walker Press, 294302.
  • Joseleau JP, Faix O, Kuroda KI, Ruel K. 2004a. A polyclonal antibody directed against syringylpropane epitopes of native lignins. Comptes Rendus Biologies 327: 809815.
  • Joseleau JP, Imai T, Kuroda K, Ruel K. 2004b. Detection in situ and characterization of lignin in the G-layer of tension wood fibres of Populus deltoides. Planta 219: 338345.
  • Karhunen P, Rummakko P, Sipila J, Brunow G, Kilpelainen I. 1995. Dibenzodioxocins – a novel type of linkage in softwood lignins. Tetrahedron Letters 36: 169170.
  • Kukkola EM, Koutaniemi S, Gustafsson M, Karhunen P, Ruel K, Lundell TK, Saranpaa P, Brunow G, Teeri TH, Fagerstedt KV. 2003. Localization of dibenzodioxocin substructures in lignifying Norway spruce xylem by transmission electron microscopy-immunogold labeling. Planta 217: 229237.
  • Lacombe E, Hawkins S, Van Doorsselaere J, Piquemal J, Goffner D, Poeydomenge O, Boudet AM, Grima-Pettenati J. 1997. Cinnamoyl CoA reductase, the first committed enzyme of the lignin branch biosynthetic pathway: cloning, expression and phylogenetic relationships. Plant Journal 11: 429441.
  • Lapierre C, Pollet B, Rolando C. 1995. New insights into the molecular architecture of hardwood lignins by chemical degradative methods. Research on Chemical Intermediates 21: 397412.
  • Laskar DD, Jourdes M, Patten AM, Helms GL, Davin LB, Lewis NG. 2006. The Arabidopsis cinnamoyl CoA reductase irx4 mutant has a delayed but coherent (normal) program of lignification. Plant Journal 48: 674686.
  • Lawoko M, Henriksson G, Gellerstedt G. 2005. Structural differences between the lignin–carbohydrate complexes present in wood and in chemical pulps. Biomacromolecules 6: 34673473.
  • Leplé JC, Dauwe R, Morreel K, Storme V, Lapierre C, Pollet B, Naumann A, Kang KY, Kim H, Ruel K et al . 2007. Downregulation of cinnamoyl-coenzyme a reductase in poplar: Multiple-level phenotyping reveals effects on cell wall polymer metabolism and structure. Plant Cell 19: 36693691.
  • Ma QH. 2007. Characterization of a cinnamoyl-CoA reductase that is associated with stem development in wheat. Journal of Experimental Botany 58: 20112021.
  • Matsunaga S, Schutze K, Donnison IS, Grant SR, Kuroiwa T, Kawano S. 1999. Single pollen typing combined with laser-mediated manipulation. Plant Journal 20: 371378.
  • Mayhew TM, Griffiths G, Lucocq JM. 2004. Applications of an efficient method for comparing immunogold labelling patterns in the same sets of compartments in different groups of cells. Histochemistry and Cell Biology 122: 171177.
  • Mir Derikvand M, Berrio Sierra J, Ruel K, Pollet B, Do C-T, Thévenin J, Buffard D, Jouanin L, Lapierre C. 2008. Redirection of the phenylpropanoid pathway to feruloyl malate in Arabidopsis mutants deficient for cinnamoyl-CoA reductase 1. Planta 227: 943956.
  • Nakashima J, Chen F, Jackson L, Shadle G, Dixon RA. 2008. Multi-site genetic modification of monolignol biosynthesis in alfalfa (Medicago sativa): effects on lignin composition in specific cell types. New Phytologist 179: 738750.
  • Nishiyama Y, Sugiyama J, Chanzy H, Langan P. 2003. Crystal structure and hydrogen bonding system in cellulose 1(alpha), from synchrotron X-ray and neutron fiber diffraction. Journal of the American Chemical Society 125: 1430014306.
  • O’Connell A, Holt K, Piquemal J, Grima-Pettenati J, Boudet A, Pollet B, Lapierre C, Petit-Conil M, Schuch W, Halpin C. 2002. Improved paper pulp from plants with suppressed cinnamoyl-CoA reductase or cinnamyl alcohol dehydrogenase. Transgenic Research 11: 495503.
  • Patten AM, Cardenas CL, Cochrane FC, Laskar DD, Bedgar DL, Davin LB, Lewis NG. 2005. Reassessment of effects on lignification and vascular development in the irx4 Arabidopsis mutant. Phytochemistry 66: 20922107.
  • Pinçon G, Chabannes M, Lapierre C, Pollet B, Ruel K, Joseleau JP, Boudet AM, Legrand M. 2001. Simultaneous down-regulation of caffeic/5-hydroxy ferulic acid-O-methyltransferase I and cinnamoyl-coenzyme A reductase in the progeny from a cross between tobacco lines homozygous for each transgene. Consequences for plant development and lignin synthesis. Plant Physiology 126: 145155.
  • Piquemal J, Lapierre C, Myton K, O’Connel A, Schuch W, Grima-Pettenati J, Boudet AM. 1998. Down-regulation of cinnamoyl-CoA reductase induces significant changes of lignin profiles in transgenic tobacco plants. Plant Journal 13: 7183.
  • Ralph J, Hatfield RD, Piquemal J, Yahiaoui N, Pean M, Lapierre C, Boudet AM. 1998. NMR characterization of altered lignins extracted from tobacco plants down-regulated for lignification enzymes cinnamylalcohol dehydrogenase and cinnamoyl-CoA reductase. Proceedings of the National Academy of Sciences, USA 95: 12 80312 808.
  • Ralph J, Kim H, Lu F, Grabber JH, Leple J-C, Berrio-Sierra J, Derikvand MM, Jouanin L, Boerjan W, Lapierre C. 2008. Identification of the structure and origin of a thioacidolysis marker compound for ferulic acid incorporation into angiosperm lignins (and an indicator for cinnamoyl CoA reductase deficiency). Plant Journal 53: 368379.
  • Ralph J, Lundquist K, Brunow G, Lu F, Kim H, Schatz PF, Marita JM, Hatfield RD, Ralph SA, Christensen JH et al . 2004. Lignins: natural polymers from oxidative coupling of 4-hydroxyphenyl-propanoids. Phytochemistry Reviews 3: 2960.
  • Ruel K. 2007. Genetically transformed plants to evaluate the function of constituent polymers in the assembly of wood cell walls. In: EntwistleK, HarrisP, WalkerJ, eds. The compromised wood workshop 2007. Christchurch, New Zealand: Wood Quality Initiative, 125141.
  • Ruel K, Barnoud F, Eriksson KE. 1981. Micromorphological and ultrastructural aspects of spruce wood degradation by wild type Sporotrichum pulverulentum and its cellulose-less mutant Cel 44. Holzforschung 35: 157171.
  • Ruel K, Burlat V, Joseleau JP. 1999. Relationship between ultrastructural topochemistry of lignin and wood properties. Iawa Journal 20: 203211.
  • Ruel K, Chabannes M, Boudet A, Legrand M, Joseleau J. 2001. Reassessment of qualitative changes in lignification of transgenic tobacco plants and their impact on cell wall assembly. Phytochemistry 57: 875882.
  • Ruel K, Faix O, Joseleau JP. 1994. New immunogold probes for studying the distribution of the different lignin types during plant-cell wall biogenesis. Journal of Trace and Microprobe Techniques 12: 247265.
  • Ruel K, Montiel MD, Goujon T, Jouanin L, Burlat V, Joseleau JP. 2002. Interrelation between lignin deposition and polysaccharide matrices during the assembly of plant cell walls. Plant Biology 4: 28.
  • Saeman JF, Moore JE, Mitchell RL, Millet MA. 1954. Techniques for the determination of pulp constituents by quantitative paper chromatography. Tappi 37: 336343.
  • Salmen L. 2004. Micromechanical understanding of the cell-wall structure. Comptes Rendus Biologies 327: 873880.
  • Taylor JG, Owen TP, Koonce LT, Haigler CH. 1992. Dispersed lignin in tracheary elements treated with cellulose synthesis inhibitors provides evidence that molecules of the secondary cell-wall mediate wall patterning. Plant Journal 2: 959970.
  • Terashima N, Fukushima K, He L-F, Takabe K. 1993. Comprehensive model of the lignified plant cell wall. In: JungHG, BuxtonDR, HatfieldRD, RalphR, eds. Forage cell wall structure and digestibility. Madison, WI, USA: American Society of Agronomy, Crop Science Society of America, Soil Science Society of America, 247270.
  • Thiery J. 1967. Mise en évidence de polysaccharides sur coupes fines en microscopie. Journal de Microscopie 6: 9871017.
  • Thumma BR, Nolan MF, Evans R, Moran GF. 2005. Polymorphisms in cinnamoyl CoA reductase (CCR) are associated with variation in microfibril angle in Eucalyptus spp. Genetics 171: 12571265.
  • Turner SR, Somerville CR. 1997. Collapsed xylem phenotype of Arabidopsis identifies mutants deficient in cellulose deposition in the secondary cell wall. The Plant Cell 9: 689701.
  • Zhong RQ, Pena MJ, Zhou GK, Nairn CJ, Wood-Jones A, Richardson EA, Morrison WH, Darvill AG, York WS, Ye ZH. 2005. Arabidopsis fragile fiber8, which encodes a putative glucuronyltransferase, is essential for normal secondary wall synthesis. The Plant Cell 17: 33903408.