A family 15 carbohydrate esterase (CE15) from the white-rot basidiomycete, Phanerochaete carnosa (PcGCE), was transformed into Arabidopsis thaliana Col-0 and was expressed from the constitutive cauliflower mosaic virus 35S promoter. Like other CE15 enzymes, PcGCE hydrolyzed methyl-4-O-methyl-d-glucopyranuronate and could target ester linkages that contribute to lignin–carbohydrate complexes that form in plant cell walls. Three independently transformed Arabidopsis lines were evaluated in terms of nine morphometric parameters, total sugar and lignin composition, cell wall anatomy, enzymatic saccharification and xylan extractability. The transgenic lines consistently displayed a leaf-yellowing phenotype, as well as reduced glucose and xylose content by as much as 30% and 35%, respectively. Histological analysis revealed 50% reduction in cell wall thickness in the interfascicular fibres of transgenic plants, and FT-IR microspectroscopy of interfascicular fibre walls indicated reduction in lignin cross-linking in plants overexpressing PcGCE. Notably, these characteristics could be correlated with improved xylose recovery in transgenic plants, up to 15%. The current analysis represents the first example whereby a fungal glucuronoyl esterase is expressed in Arabidopsis and shows that the promotion of glucuronoyl esterase activity in plants can alter the extent of intermolecular cross-linking within plant cell walls.
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Glucuronoxylan is characterized by a β-1,4 linked xylose backbone that can be substituted by α-(1,2)-glucopyranosyl uronic acid (GlcA) or 4-O-methyl-α-d-glucopyranosyl uronic acid (MeGlcA) side groups. In gymnosperms, glucuronoxylan can be substituted with α-(1,2)-l-arabinose, while glucuronoxylans in angiosperms is acetylated (Ishii and Shimizu, 2001). Arabinose and MeGlcA side groups often participate in ester linkages as pendants of lignin. For instance, approximately 30% and 40% of MeGlcA are esterified to lignin in beechwood and birchwood, respectively (Takahashi and Koshijima, 1988). Such lignin–carbohydrate complexes (LCCs) can complicate the removal of lignin from cellulosic pulp, and free MeGlcA typically transform to hexenuronic acid during alkaline pulping processes, which increases the consumption of pulping and bleaching chemicals (Chakar et al., 2000; Koshijima and Watanabe, 2003; Watanabe and Koshijima, 1988). Recent efforts to develop economic bioprocesses for the conversion of lignocellulosic biomass to fermentable sugars also reveal the inhibitory effect of LCCs on the hydrolytic activities of cellulases and hemicellulases (Berlin et al., 2006; Várnai et al., 2010).
Physical and chemical pretreatments have been developed to optimize the separation of lignin and cell wall polysaccharides from different feedstocks (Kumar et al., 2009; Taherzadeh and Karimi, 2008). Plant fibre engineering is also seen as a route to improving feedstock characteristics and further reducing the energy and cost of refining biomass (Abramson et al., 2010; Boerjan, 2005; Taylor et al., 2008). For example, many tree-breeding programmes aim to reduce the lignin content and increase the production of cellulose (Chen and Dixon, 2007; Halpin and Boerjan, 2003; Merkle and Dean, 2000; Pilate et al., 2002). Several publications also describe the potential of modifying plant cell wall composition by altering the expression of endogenous enzymes involved in cell wall synthesis (Mellerowicz and Sundberg, 2008; Stewart et al., 2009). For instance, the Arabidopsis fra8 mutant encodes a defective glycosyltransferase from family GT47 and displays reduced cellulose and xylan content as well as reduced plant height and fibre cell wall thickness (Zhong et al., 2005). In addition, the Arabidopsis gux1 and gux2 mutants, which are defective in xylan glucuronyltransferase activity, display a wild-type phenotype and almost unaltered cell wall composition despite differential glucuronoxylan branching pattern and extractability (Mortimer et al., 2010).
Similarly, cell wall composition, architecture and susceptibility to downstream processing can be improved through transgenic expression of exogenous carbohydrate active enzymes (CAZymes) in planta. Examples here include the transgenic expression of a secreted cellulase from Populus alba in Arabidopsis, leading to increased cellulose content and fibre elongation (Park et al., 2003). By comparison, the membrane bound KORRIGAN1 cellulase from Populus tremula × tremuloides expressed in Arabidopsis decreased cellulose crystallinity in transgenic plants (Takahashi et al., 2009). The effect of microbial CAZyme expression in plants has also been investigated. Borkhardt et al. (2010) recently demonstrated that transgenic expression of thermostable xylanases in Arabidopsis increased the solubilization of cell wall xylan at elevated temperature. Similarly, the transgenic expression of a family GH5 endoglucanase in Nicotiana tabacum and Zea mays improves the pretreatment efficacy and enzyme digestibility of cells walls from these plants (Brunecky et al., 2011). The transgenic expression of accessory hemicellulases, including fungal feruloyl esterases in Festuca arundinacea, and feruloyl esterases and arabinofuranosidases in Arabidopsis, also increases the degradability of resulting biomass (Buanafina et al., 2010; Pogorelko et al., 2011). Because the activity of accessory hemicellulases is limited to branching groups along the xylan backbone, it is anticipated that additional examples of accessory hemicellulase expression in plants could reveal options for recovering high-quality hemicellulose as well as fermentable sugars.
Glucuronoyl esterase (EC 3.1.1.-) is an accessory enzyme that hydrolyzes the ester linkage between lignin alcohols and MeGlcA and was originally isolated from Schizophyllum commune (Li et al., 2007; Spániková and Biely, 2006). The CAZy classification has grouped glucuronoyl esterase in Carbohydrate Esterase (CE) family 15 (Li et al., 2007). At the time of writing, CE15 comprised 89 members mostly from wood-degrading micro-organisms, with only six characterized enzymes (Ďuranováet al., 2009b; Li et al., 2008; Vafiadi et al., 2009). The structure of the Hypocrea jecorina 4-O-methyl-glucuronoyl methylesterase Cip2 (PDB ID: 3PIC) was solved and revealed the Ser-His-Glu as the putative catalytic triad (Pokkuluri et al., 2011).
Recently, a novel glucuronoyl esterase (PcGCE) was identified in the Basidiomycete, Phanerochaete carnosa (MacDonald et al., 2011), based on sequence homology to the Phanerochaete chrysosporium esterase ge2 (Ďuranováet al., 2009b). Herein, the activity of recombinant PcGCE was evaluated, and PcGCE was constitutively expressed in Arabidopsis to investigate the impact of this carbohydrate esterase on plant cell wall chemistry and architecture.
Results and discussion
Recombinant expression and purification of a glucuronoyl esterase from P. carnosa
The glucuronoyl esterase gene cloned from Phanerochaete carnosa (PcGCE; NCBI accession: JQ972915) encoded a mature protein containing 407 amino acids, which shared 91.2% sequence identity with the previously identified glucuronoyl esterase from P. chrysosporium (GE2; Ďuranováet al., 2009b). The recombinant expression of PcGCE in Pichia pastoris GS115 was highest after 4 days of induction with 0.5% methanol, and approximately 5.3 mg/L of protein was purified to over 95% homogeneity by affinity chromatography. Mass spectrometry was used to confirm the identity of the purified enzyme, and eight trypsin fragments were matched to the parent sequence (approximately 23% protein coverage). The deduced molecular mass of the mature protein with a c-myc tag epitope and polyhistidine tag was approximately 42 kDa, which was less than the electrophoretic molecular weight of the purified enzyme (72 kDa) (Figure S1). NetNGlyc (http://www.cbs.dtu.dk/services/NetNGlyc/) predicted eight putative N-glycosylation sites in PcGCE, and treatment of PcGCE with N-glycosidase F (PNGase F) reduced the molecular weight of the enzyme to approximately 50 kDa (Figure S1). PcGCE activity was measured using methyl 4-O-methyl-d-glucopyranuronate as previously described (Ďuranováet al., 2009a). The maximum activity of PcGCE was observed at pH 6 and 40 °C (data not shown), which was similar to the pH and temperature optima of the glucuronoyl esterase (GE2) from P. chrysosporium (pH 5.0–6.0 and 45–55 °C, respectively; Ďuranováet al., 2009a) as well as other glucuronoyl esterases from the same family (Li et al., 2007; Spániková and Biely, 2006; Vafiadi et al., 2009).
Transgenic expression of PcGCE in Arabidopsis
PcGCE was constitutively expressed in Arabidopsis thaliana Col-0 using the cauliflower mosaic virus (CaMV) 35S promoter from the p35SMYC binary vector (see Experimental Procedures). Twenty independent Col-0 transformant lines (PcGCE) were isolated based on growth in the presence of 15 μg/mL hygromycin, and three randomly selected lines (PcGCE6, PcGCE7 and PcGCE13) were cultivated to the T3 stage for analysis.
Transcript and functional expression of PcGCE in Arabidopsis was confirmed using semiquantitative RT-PCR and the glucuronoyl esterase activity assay (Figure 1) (Spániková and Biely, 2006). Band intensities from RT-PCR analyses were positively correlated with activity measurements, and among the three lines analysed, PcGCE activity was the highest in PcGCE7 and absent in wild-type Arabidopsis.
Assessments of nine morphometric parameters were performed to evaluate the impact of PcGCE expression on Arabidopsis phenotype. Transgenic plants had smaller leaf size and shorter plant height, and their flowering was delayed (Table S1). Approximately 4 weeks after germination, yellowing rosette leaves resembling early senescence were also observed in all of the transgenic lines (Figure 2). Leaf yellowing has been observed in other transgenic plants expressing fungal enzymes in the cell wall, including Arabidopsis plants expressing a yeast-derived invertase (von Schaewen et al., 1990). In the current analysis, leaf yellowing was highest in PcGCE7 and PcGCE13 lines, which also produced highest levels of functional PcGCE. Of the other eight morphometric parameters analysed, leaf yellowing could be directly correlated with reduced above-ground biomass. Notably, however, differences in stem length and stem biomass contributed more to the differences in total above-ground biomass than did leaf size.
Effect of PcGCE transgenic expression on the anatomy and composition of Arabidopsis cell walls
Stem cross-sections sampled from the basal 25% of plant height were prepared for microscopic analyses and then stained using toluidine blue. The cell wall thickness of all schlerenchyma cells was significantly reduced in all lines, particularly PcGCE7 and PcGCE13, which also had the highest level of PcGCE expression. Notably, PcGCE expression did not appear to affect the thickness of parenchyma cells, which is consistent with PcGCE acting on intermolecular bonds between xylan and lignin that are absent in parenchyma tissue (Peña et al., 2007; Persson et al., 2007; Zhou et al., 2006). Within the schlerenchyma tissue, cell wall thinning was particularly evident in the interfascicular fibre region, where the cell wall thickness was reduced by nearly 55% in PcGCE7 and PcGCE13 lines compared with wild-type plants (Figure 3, Table 1).
Table 1. Summary of analyses by immunolocalization and electron microscopy
*Statistically significant difference relative to Col-0 as determined using a 2-way ANOVA followed by Bonferroni post-test (P < 0.05). Between 8 and 12 measurements were collected to calculate cell wall thickness; 60 measurements were collected to calculate LM10 labelling density. Two independent plants were analysed for each line.
Cell wall thickness (μm)
1.8 ± 0.8
1.1 ± 0.4*
0.8 ± 0.1*
0.8 ± 0.2*
1.0 ± 0.2
1.0 ± 0.2
0.8 ± 0.1
0.8 ± 0.3
0.9 ± 0.2
0.6 ± 0.2
0.9 ± 0.2
0.8 ± 0.2
LM10 Label (per μm2)
375.2 ± 52.0
259.6 ± 65.9
581.7 ± 115.2*
515.6 ± 51.1*
541.7 ± 152.8
360.4 ± 80.1*
549.4 ± 103.5
529.3 ± 90.6
434.2 ± 104.8
326.4 ± 164.3
418.3 ± 63.8
434.4 ± 53.2
Stem material was collected from the three transgenic lines and the wild-type plant to determine whether cell wall thinning was correlated with changes in total sugar and lignin composition. Total sugar and lignin was assessed using plant stems that were harvested after the first silique shattered (developmental stage 8.0; Boyes et al., 2001). The siliques, leaves and roots were removed from the stems, and stems were air-dried, milled and extracted before complete hydrolysis of plant polysaccharides in 72% sulphuric acid. The transgenic lines contained higher soluble lignin content as compared to wild-type plants (Figure 4a), which was mainly attributed to comparatively high levels of acid soluble 4-hydroxybenzoic acid and 4-hydroxybenzaldehyde released through the acid hydrolysis (Figure 4b). Compared with wild-type plants, the 4-hydroxybenzoic acid and 4-hydroxybenaldehyde fraction of acid hydrolyzed PcGCE7 stem increased by 81% and 45%, respectively. The same compounds were increased by 61% and 56% in PcGCE13. All transgenic lines, including PcGCE6, contained comparatively low glucose and xylose content (Figure 5). In particular, glucose and xylose contents in PcGCE7 lines were reduced by over 20% and 25%, respectively, while these same sugars in PcGCE13 lines were reduced by 30% and 35%, respectively.
Although consistent with cell wall thinning, it was initially surprising to observe reduced glucose and xylose contents given that PcGCE does not target these particular sugars. However, similar phenomena were previously observed in Arabidopsis mutants of glucuronoxylan biosynthesis genes. In particular, the Arabidopsis mutants fra8 and irx7 are defective in allelic genes from glycosyltransferase (GT) family 47 that are predicted to synthesize the reducing end sequence of the glucuronoxylan backbone (Brown et al., 2007; Zhong et al., 2005). In addition to reduced xylose content, Arabidopsis fra8/irx7 mutants show reduced cell wall thickness and glucose content. Moreover, while the frequency of 4-O-methyl glucuronic acid (MeGlcA) side branches is similar to wild-type plants, glucuronoxylans in the fra8/irx7 mutants show dramatic reduction in unmethylated glucuronic acid (GlcA). Other mutants in glucuronoxylan biosynthesis include Arabidopsis irx9 (Brown et al., 2005), irx10 (Brown et al., 2009; Wu et al., 2009) and irx14 (Brown et al., 2007), which are mutated in genes from families GT43 and GT47 that are predicted to participate in xylan chain elongation. In addition to reduced xylose content, these mutations also lead to reduced GlcA side groups and reduced cellulose content in late stages of stem development. It is conceivable that similar to the predicted effect of PcGCE transgene expression, cross-linkages between xylan and lignin in these Arabidopsis mutants might be altered by consequence of reduced xylan or GlcA content.
In situ compositional analysis of interfascicular fibres
The predicted activity of PcGCE, along with reduced cell wall thickness and xylose contents in the transgenic plants, provoked an interest to characterize the distribution of xylan within plant cell walls. Immunolocalizations were performed using antibodies towards unsubstituted xylan (LM10; McCartney et al., 2005) and the glucuronoxylan MeGlcA side group (UX; Koutaniemi et al., 2012). Statistically significant differences in label density between transgenic and wild-type samples were not observed using the UX antibody (Table S3). By contrast, decreased cell wall thickness of interfascicular fibres in PcGCE7 and PcGCE13 could be correlated with increased LM10 immunolabel density in cell walls in the same region (Table 1, Figure S2).
Increased LM10 immunolabel density suggests that xylan was more tightly assembled within the interfascicular fibre region of PcGCE7 and PcGCE13 transgenic lines. Alternatively, the accessibility or frequency of LM10 epitopes was higher within corresponding fibres. In an effort to resolve these possibilities, FT-IR microspectroscopy was applied using a 64 × 64 focal plane array detector to directly analyse intermolecular and intramolecular bonding within stem sections of PcGCE7, which also expressed highest PcGCE activity. The spectra were collected specifically from cell walls of interfascicular fibres; orthogonal projections to latent structures discriminant analysis (OPLS-DA) was then performed and revealed good separation of the PcGCE7 plants and the control (Figure 6a) (Bylesjöet al., 2006; Gorzsás et al., 2011; Trygg and Wold, 2002).
Positive loadings correspond to bands that were more intense in PcGCE7 compared with control samples (Figure 6b). The loadings plot shows that PcGCE7 was enriched in cell wall components that absorb at 1595 cm−1 and 1650 cm−1 and was reduced in components that absorb at 1130 cm−1 and 1510 cm−1. While bands at 1595 cm−1 and 1510 cm−1 are correlated with aromatic carbon–carbon double-bond vibrations originating from lignin, the band at 1510 cm−1 is typically more intensive in more cross-linked lignin structures (Stewart et al., 1997; Zhong et al., 2000). Notably then, the lower 1510 cm−1 per 1595 cm−1 ratio in the PcGCE7 plants compared with the control, suggests a reduction in lignin cross-linking within the transgenic line. The band at 1130 cm−1 is attributed to the occurrence of asymmetrical –C-O-C– stretching in polysaccharides, while the band near 1650 cm−1 is assigned to absorbed water as well as conjugated carbonyl (C=O) groups in lignin (Faix, 1991; Kacurákováet al., 2002; Pandey and Pitman, 2003). While the correct interpretation of reduced band intensity at 1650 cm−1 is uncertain, reduced absorbance at 1130 cm−1 together with a reduced 1510 cm−1 per 1595 cm−1 ratio is consistent with an altered intermolecular linkage structure in transgenic PcGCE7 plants.
Taken together, LM10 immunolabeling and FT-IR microspectroscopy of PcGCE7 lines suggest that PcGCE overexpression in Arabidopsis could increase xylan accessibility through altered intermolecular linkages, at least within the interfascicular fibre region. Enzymatic digestibility and xylan extractability of transgenic stem was evaluated next to assess the practical implications of these alternations to plant cell walls.
Effect of PcGCE transgenic expression on enzymatic saccharification and xylan extractability of plant cell walls
To investigate the effect of changes to stem cell wall composition and architecture on lignocellulose processing, stem sections from PcGCE transgenic lines and wild-type Arabidopsis were compared in terms of digestibility using a cellulase cocktail and in terms of xylan extractability through alkali treatment.
For the cellulase treatment, air-dried Arabidopsis stems were hand-sectioned and hydrolyzed using a Celluclast and Novozyme 188 cellulase cocktail. On the basis of total glucose in respective stem samples, cellulase treatment for 24 h at 50 °C released similar amounts of total reducing sugars from PcGCE7 and wild-type plants, and slightly higher amounts of total reducing sugar from PcGCE6 and PcGCE13 lines (Figure S3). While higher PcGCE activity in PcGCE7 compared with PcGCE6 and PcGCE13 might be expected to enhance cellulase hydrolysis, it is possible that stress responses to higher levels of esterase activity counteract the possible benefits of altering cross-linkages between hemicellulose and lignin. To address this possibility, future analyses will investigate the impact of PcGCE activity on the expression of stress response genes and production of corresponding metabolites in Arabidopsis.
The influence of PcGCE activity on the interaction between xylan and other cell wall components was investigated by extracting xylan from equal quantities of alcohol insoluble residue (AIR) prepared from stems of PcGCE lines and wild-type Arabidopsis. In all cases, the majority of solubilized xylose was contained in 1 m KOH fractions of AIR (Figure 7); the amount of xylose extracted per milligram of AIR was also similar between the transgenic lines and the wild-type plants (Figure S4). However, when considering differences in starting xylose content, xylan extractability using 1 m KOH had clearly increased by approximately 15% in two of the three PcGCE transgenic lines (Figure 7). This result is consistent with higher xylan extraction from Arabidopsis gux1/2 mutants, which are defective in genes encoding GT8 enzymes involved in the formation of xylan glucuronate side chains that can form cross-linkages to lignin (Mortimer et al., 2010). Moreover, in the case of the PcGCE7 line, this result supports the functional interpretation of immunolocalization and FT-IR microspectroscopy analyses, which predict increased xylan accessibility in this transgenic line. Notably, improved xylan extractability from PcGCE6 suggests that through modulating PcGCE expression, it might be possible to enhance both cellulase digestability and xylan extractability.
In summary, while most efforts to improve bioprocesses for lignocellulose utilization focus on reducing the amount of lignin and altering cellulose content and crystallinity, the effect of branching sugars that participate in LCCs has been less well characterized. This study demonstrates that transgenic expression of a fungal glucuronoyl esterase from family CE15 can alter plant cell wall composition and architecture, and thereby improve the recovery of xylan from resulting plant biomass. The comparison of three independently transformed lines also suggests that highest improvement to cell wall fractionation might be achieved through regulated expression of the PcGCE enzyme. Analyses have been initiated to test this possibility.
Recombinant expression of PcGCE in Pichia pastoris
The Phanerochaete carnosa glucuronoyl esterase gene (PcGCE, NCBI accession: JQ972915) was amplified from P. carnosa cDNA and cloned into the pPICZαB expression vector (Invitrogen, Carlsbad, CA) using the restriction sites PstI and NotI. Resulting plasmids were transformed into Pichia pastoris GS115 by electroporation and selected at 30 °C on YPDS plates (1% yeast extract, 2% peptone, 2% dextrose, 1 m sorbitol, 2% agar and 100 μg/mL Zeocin™) as described by the manufacturer’s instructions (Invitrogen). Transformants were transferred to buffered methanol complex medium (BMMY: 1% yeast extract; 2% peptone; 100 mm potassium phosphate (pH 6.0); 1.34% yeast nitrogen base without amino acids (YNB); 4 × 10−5% biotin; 0.5% methanol) and then screened for protein expression by immuno-colony blot using nitrocellulose membranes (0.45 μm; Bio-Rad, Hercules, CA), anti-myc antibodies (Invitrogen), alkaline phosphatase-linked anti-rabbit IgG conjugates (Sigma-Aldrich, St. Louis, MO) and 5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium solution (BCIP/NBT; Sigma-Aldrich). Positive transformants were grown overnight in buffered glycerol complex medium (BMGY: 1% yeast extract; 2% peptone; 100 mm potassium phosphate (pH 6.0); 1.34% YNB without amino acids (YNB); 4 × 10−5% biotin; 1% glycerol) at 28 °C with continuous shaking at 265 rpm. The cells were harvested by centrifugation and suspended in BMMY medium to OD600∼1. Cultures were grown at 28 °C and 265 rpm for 4 days, and 0.5% methanol was added every 24 h to induce recombinant protein expression.
Supernatants from methanol-induced cultures were filtered through 0.44 μm Acrodisc filters (Pall, Port Washington, NY) and then concentrated approximately 20 times using Vivaspin 20 concentration units (GE Healthcare, Chalfont St. Giles, UK). The concentrated culture medium was replaced by 20 mm sodium phosphate (pH 7.4), 500 mm NaCl and 30 mm imidazole using HiPrep™ 26/10 Desalting column (GE Healthcare) and a BioLogic DuoFlow™ FPLC (Bio-Rad), before being applied to a HisTrapTM HP column (GE Healthcare). Fractions were eluted with 20 mm sodium phosphate (pH 7.4), 500 mm NaCl and 250 mm imidazole, and the buffer was replaced by 50 mm sodium phosphate (pH 6.0) using Vivaspin 20 concentration units (GE Healthcare).
In-gel trypsin digestion with sequencing-grade trypsin (Promega, Madison, WI) followed by tandem mass spectrometry was performed to confirm the identity of the purified protein. Tryptic fragments were analysed using an Autoflex III Maldi-TOF-TOF (Bruker, Billerica, MA) located at the Centre for the Analyses of Genome Evolution and Function (CAGEF), University of Toronto. Protein deglycosylation was performed using PNGase F (BioLabs, Ipswich, MA) as per the manufacturer’s instructions.
Activity measurement of purified PcGCE
Glucuronoyl esterase activity was measured using methyl 4-O-methyl-d-glucopyranuronate as previously described (Spániková and Biely, 2006). Briefly, the enzyme reaction contained 50 mm Britton and Robinson’s universal buffer (pH 6.0), 2 mm substrate, 1 μg enzyme and was incubated at 30 °C for up to 20 h. Remaining substrate was detected by adding two volumes of alkaline hydroxylamine hydrochloride (prepared by mixing an equal volume of 2 m hydroxylamine hydrochloride and 3.5 N sodium hydroxide) followed by acidification with one volume of hydrochloric acid and addition of one volume of 0.37 m ferric chloride before absorbance measurement at 540 nm (Hestrin, 1949). Acetylcholine chloride was used to generate the standard curve. The pH optimum of PcGCE was determined by measuring activity at 30 °C from pH 4 to pH 10, and the optimal temperature for PcGCE activity was determined by measuring activity at pH 6.0 from 30 °C to 60 °C.
Vector construction for transgenic expression of PcGCE in Arabidopsis
A plant expression vector (p35SMYC) was constructed by amplifying the CaMV 35S promoter from pCAMBIA2201 (CAMBIA, Canberra, Australia) and ligating a HindIII digested fragment to pCAMBIA1390 (CAMBIA) using the In-FusionTM Advantage PCR Cloning Kit (Clontech, Mountain View, CA). The NOS terminator from pCAMBIA1302 (CAMBIA) was similarly cloned as an EcoRI fragment downstream of the 35S promoter, and a myc tag (EQKLISEEDL) was generated de novo between the 35S promoter and NOS terminator, leaving a unique SalI site upstream and in frame. Fusion-Blue™ chemically competent cells (Clontech) were used to propagate resulting plasmids in all cases. After sequencing, plasmids containing the gene of interest along with the native secretion signal peptide were transformed into chemically competent Agrobacterium tumefaciens strain GV3101 (GV3101) using a standard heat-shock method.
Stable plant transformation
Arabidopsis thaliana Col-0 were transformed using a modified floral-dip method (Clough and Bent, 1998). Briefly, YEP (2% peptone, 2% yeast extract, 0.5% NaCl) containing 30 μg/mL of kanamycin was inoculated with GV3101 containing a plasmid of interest and allowed to grow for 16 h at 28 °C and 250 rpm. The suspension was then centrifuged at 1810 g for 10 min before suspending the pelleted cells in 40 mL of 5% sucrose containing 0.05% Silwet L-77. After excising mature flowers and immature siliques, the remaining stems and immature flowers were dipped in the sucrose solution for approximately 5 s. Treated plants were left on their side and covered with a semitransparent plastic dome for approximately 16 h before being returned to a 16/8 light/dark regime at 21 °C.
Transformant selection and growth
Mature seeds were collected from fully senesced transformed plants and sterilized using chlorine gas. Approximately 1000 T0 seeds were then plated on ½ Murashige and Skoog (pH 5.8) medium containing 15 μg/mL of hygromycin before being cold stratified at 4 °C for 48 h. The plates were then incubated at 21 °C under a 16/8 light/dark regime until transformed seedlings developed roots and at least one set of true leaves. PCR and RT-PCR using gene-specific primers were performed to confirm transgene insertion into host chromosomal DNA and transgene expression, respectively (Table S2). Positive transformants were then moved to soil and allowed to mature. The resulting T1 seeds were gas sterilized and plated as before. This process was repeated until the resulting T3 seedlings were moved to soil for growth and phenotype analyses. Methods were the same for nontransformed Arabidopsis thaliana Col-0 except hygromycin was omitted from the growth medium. Inflorescence stems for subsequent analyses were collected at developmental stage 8.00 corresponding to the first shattered silique (Boyes et al., 2001).
T3 plants were continuously measured after seedlings were transferred to soil with respect to morphometric characteristics such as plant height, number of leaves, rosette and leaf size, as well as observing for growth traits including bolting, flowering and leaf yellowing. At least 24 plants were measured for each line. Three plants from each line were used to measure the above ground and stem biomass after 40 days postgermination. Two-way analyses of variance (ANOVA) followed by Bonferroni post-test were performed using GraphPad Prism (GraphPad Software, La Jolla, CA).
Total protein isolation for glucuronoyl esterase activity assay
Total protein extract was isolated from transgenic Arabidopsis seedlings and was quantified with the BCA Protein Assay Kit (Thermo Scientific, Waltham, MA). Three-week-old Arabidopsis seedlings were homogenized and suspended in protein extraction buffer [50 mm HEPES-NaOH (pH 7.0); 10 mm MgCl2; 1 mm Na2EDTA; 2.6 mm DTT; 10% ethylene glycol; and 0.02% Triton X-100. The mixture was centrifuged at 16 200 g for 10 min at 4 °C, and the supernatant was recovered to measure protein concentration. An aliquot representing 100 μg of protein was used in the glucuronoyl esterase activity assay as described above for the Pichia pastoris expressed PcGCE.
Inflorescence cell wall compositional analyses
For each line, stems from 50 to 60 plants were pooled and ground. Ground stem samples were divided into three technical 100 mg replicates, which were extracted, hydrolyzed and analysed for cell wall composition using an modified acid hydrolysis procedure (Sluiter et al., 2008). Briefly, the siliques and leaves were removed from the Arabidopsis stems, which were air-dried for 1 week. The stems were milled with a Wiley mill and sieved with an 80-mesh sieve. The milled stems were refluxed with acetone in a Soxhlet apparatus for 24 h (Cullis et al., 2004). The stems were then oven-dried at 105 °C for 18 h before being hydrolyzed with 72% (w/w) sulphuric acid for 2 h at room temperature with stirring every 10 min. The hydrolysate was diluted to 4% (w/w) sulphuric acid and autoclaved at 121 °C for 1 h in crimped-top serum bottles. The hydrolysate was filtered with a glass medium-fritted crucible, and the insoluble residue (insoluble lignin) was quantified by measuring the oven-dried mass. For the carbohydrate analyses, the hydrolysate was diluted 10-fold with 75% acetonitrile/25% methanol. The mixture was analysed with an Acquity ultra performance liquid chromatography (UPLC) equipped with a photodiode array and a mass spectrometer (Waters, Milford, MA) fitted with a 2.1 × 50 mm BEH Amide column with a pore size of 1.7 μm (Waters). The mass to charge (m/z) ratios used to quantify the individual carbohydrates were 149.1 (xylose, arabinose) and 179.2 (mannose, glucose and galactose) according to Canam et al., 2011. N-acetylglucosamine served as an internal standard (m/z 220.1). For UPLC analysis of lignin moieties, the hydrolysate was diluted 10-fold with 50% methanol before separation with a 2.1 × 50 mm BEH C18 column with a pore size of 1.7 μm (Waters). Acid moieties were detected with UV absorbance at 260 nm, while aldehyde moieties and the internal standard (o-anisic acid) were detected at 285 nm (Canam et al., 2011).
Immunolocalization of xylan distribution in plant cell walls
The Arabidopsis was harvested at developmental stage 8.00 (Boyes et al., 2001). Stem segments (approximately 5 mm) were collected and fixed in 4% paraformaldehyde and 0.05% glutaraldehyde in 25 mm phosphate buffer (pH 7.2) overnight at 4 °C. The stems were dehydrated with ethanol series (30%, 50%, 70%, 80%, 90%, 95% and 99.5%) for three 10-min incubations. The stems were infiltrated with 10%, 50% and 100% LR White (Electron Microscopy Sciences, Hatfield, PA) in ethanol overnight. The stems were cured in gelatine capsule #4 (Electron Microscopy Sciences) with LR White at 60 °C for 16 h and subsequently sectioned at 2 μm. The sections were incubated in 0.05% toluidine blue for 5 min. The sections were washed twice with distilled water, mounted in glycerol and viewed with an Axiovision microscope (Carl Zeiss, Oberkochen, Germany).
For the α-glucuronoyl xylan antibody labelling, the stem sections were incubated in 50 mm NaOH for 2 h at room temperature followed by three 5-min 0.1-m phosphate saline buffer (pH 7.2) (PBS). The sections were treated with 50 mm glycine for 15 min, blocked with the blocking solution (BTP: 1% bovine serum albumin and 0.05% Tween-20 in PBS) for 30 min, followed by incubation with primary antibody in 1 : 10 dilution in BTP for 1 h. The sections were washed with six 5-min BTP washes and subsequently incubated with the 5-nm gold particles conjugated goat α-mouse IgG (L + M) or α-rat IgG antibodies (BBInternational, Cardiff, UK) diluted in BTP in 1 : 100 dilutions for 1 h. The sections were washed with two 5-min BTP washes, two 5-min PBS washes and two 5-min ddH2O washes. The sections were viewed directly with a JEM 1230 electron microscope (JEOL Ltd., Tokyo, Japan). The antibodies used were rat monoclonal α-xylan antibody (LM10) (PlantProbes, Leeds, UK), and mouse monoclonal α-glucuronoyl xylan antibody (UX) generously provided by Luc Saulnier (INRA, France) and Maija Tenkanen (University of Helsinki, Finland). Two primary inflorescence stem cross-sections from the basal part (at 25% total plant height) of two plants were analysed for each line. The label density and cell wall thickness were measured with ImageJ software (Abràmoff et al., 2004). At least eight cell wall thickness and 60 immunolabel density measurements were made from six cells for each line. Two-way analyses of variance (ANOVA) followed by Bonferroni post-test were performed using GraphPad Prism (GraphPad Software).
Fourier-transform infrared microspectroscopy
The basal stem segments were fixed in 70% ethanol overnight at 4 °C then sectioned with a vibratome to the thickness of 30 μm. The sections were dried in a desiccator between glass slides for at least 48 h. FT-IR microspectroscopy followed the procedure described in Gorzsás et al. (2011). Briefly, spectra were recorded using a Bruker Tensor 27 spectrometer equipped with a Hyperion 3000 microscopy accessory including a 64 × 64 FPA detector (Bruker Optik GmbH, Ettlingen, Germany), using 32 scans and 4 cm−1 spectral resolution. Before multivariate analysis, cell-specific spectra were extracted, baseline corrected and area normalized in the region 900–1850 cm−1, using custom-built MATLAB scripts (Stenlund et al., 2008). Spectra were converted to data point tables using OPUS (version 5.0.53; Bruker Optik GmbH). Data processing was performed by custom scripts programmed within the MATLAB software 7.0 (Mathworks, Natick, MA). Multivariate analysis [initial principal component analysis (PCA) followed by OPLS-DA] (Bylesjöet al., 2006; Trygg and Wold, 2002) was performed using SIMCA®-P+ (12.0; Umetrics, Umeå, Sweden).
Enzyme hydrolyzability assay
The assay was modified from the NREL enzyme saccharification protocol (Selig et al., 2008). For each line, stems from 10 plants were pooled, air-dried, hand-sectioned to 5-mm segments and then conditioned to 65% humidity for 3 days. Conditioned stem sections were divided into three technical replicates, which were then incubated at 50 °C and 160 rpm 50 mm sodium citrate buffer (pH 4.8) containing 0.2% sodium azide, 0.1 filter paper units of Celluclast 1.5 L (Sigma-Aldrich) and 0.2 cellobiose units of Novozyme 188 (Sigma-Aldrich). Aliquots were removed at 0, 1, 2, 4, 8 and 24 h for quantification using the dinitrosalicylic acid assay (Adney and Baker, 1996; Xiao et al., 2004).
Alcohol insoluble residue preparation
Cell wall AIR was prepared as described in Goubet et al. (2009). For each line, inflorescence stems from 30 plants were harvested, pooled and then incubated in 95% (v/v) ethanol at 65 °C for 30 min. The tissues were cryo-milled using the Mini-Beadbeater-16 (Biospec Products, Bartlesville, OK), and the residues were collected by centrifuging at 4000 g for 15 min. The pellet was washed with 100% ethanol, chloroform/methanol (2 : 1) twice, 65%, 85% and 100% ethanol. The resulting AIR was oven-dried at 70 °C overnight.
The cell wall fractionation was modified from Coimbra et al. (1996). Briefly, triplicate 5-mg samples of AIR were incubated in 0.25 mL of 0.05 m 1,2-cyclohexanediaminetetraacetic acid (CDTA, pH 6.5) for 24 h at room temperature. The residue was collected by centrifuging at 16 200 g for 30 min. The pellet was washed once with water and was subsequently incubated with 0.05 m Na2CO3 (24 h, 4 °C), 1 m KOH (24 h, room temperature), followed by 4 m KOH (24 h, room temperature). All reagents contained 0.01 m NaBH4. The fractionated cell wall material was hydrolyzed with sulphuric acid, and the xylose content was determined by UPLC as described above. All samples were processed as three technical replicates.
We thank Dr Peter Biely (Slovak Academy of Sciences, Slovakia) for generously providing methyl 4-O-methyl-d-glucopyranuronate to measure glucuronoyl esterase activity, and Dr. Luc Saulnier (INRA, Nantes, France) and Dr Maija Tenkanen (University of Helsinki, Helsinki, Finland) for providing UX antibodies for immunolocalizations. This work is financially supported by grants from the Natural Sciences and Engineering Research Council in Canada and FORMAS in Sweden. The authors declare that they have no competing interests.