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Virus-Induced Silencing of a Plant Cellulose Synthase GeneRachel A. Burtona, David M. Gibeauta, Antony Bacicb, Kim Findlayc, Keith Robertsc, Andrew Hamiltond, David C. Baulcombed, and Geoffrey B. Fincheraa Department of Plant Science, University of Adelaide, Waite Campus, Glen Osmond, South Australia 5064, Australia b School of Botany, University of Melbourne, Parkville, Victoria 3052, Australia c Department of Cell Biology, John Innes Centre, Norwich Research Park, Colney, Norwich NR4 7UH, United Kingdom d Sainsbury Laboratory, John Innes Centre, Norwich Research Park, Colney, Norwich NR4 7UH, United Kingdom Correspondence to: Geoffrey B. Fincher, gfincher{at}waite.adelaide.edu.au (E-mail), 61-8-8303-7109 (fax)
Specific cDNA fragments corresponding to putative cellulose synthase genes (CesA) were inserted into potato virus X vectors for functional analysis in Nicotiana benthamiana by using virus-induced gene silencing. Plants infected with one group of cDNAs had much shorter internode lengths, small leaves, and a "dwarf" phenotype. Consistent with a loss of cell wall cellulose, abnormally large and in many cases spherical cells ballooned from the undersurfaces of leaves, particularly in regions adjacent to vascular tissues. Linkage analyses of wall polysaccharides prepared from infected leaves revealed a 25% decrease in cellulose content. Transcript levels for at least one member of the CesA cellulose synthase gene family were lower in infected plants. The decrease in cellulose content in cell walls was offset by an increase in homogalacturonan, in which the degree of esterification of carboxyl groups decreased from ~50 to ~33%. The results suggest that feedback loops interconnect the cellular machinery controlling cellulose and pectin biosynthesis. On the basis of the phenotypic features of the infected plants, changes in wall composition, and the reduced abundance of CesA mRNA, we concluded that the cDNA fragments silenced one or more cellulose synthase genes.
Primary cell walls of higher plants are dynamic, extracytoplasmic structures that typically are deposited in dividing and growing cells. They provide strength and flexibility for the plant as a whole but also allow intercellular exchange of water, nutrients, phytohormones, and other small molecules. After the cessation of cell growth, wall deposition may continue, but the thickened wall is referred to as a secondary wall. Wall composition varies widely across the plant kingdom and also between tissues and cell types within a particular species. In dicotyledons, cellulosic microfibrils, variously associated with xyloglucans, glucomannans, and heteroxylans, are embedded in a matrix consisting primarily of pectic polysaccharides. Additional networks of structural proteins or glycoproteins are also present. Numerous models have been developed to depict the possible interactions of cellulose and other molecular networks within the wall (
Despite the fundamental importance of cell walls in plant growth and development, a complete description of enzymes involved in their biosynthesis has not been obtained. Biochemical approaches have been frustrated by difficulties associated with the purification of membrane-bound polysaccharide synthases, by the inherent instability of many such enzymes, by possible changes in specificity during extraction, by potential losses of critical cofactors, and by probable requirements for a multienzyme complex and ancillary proteins (
An important breakthrough in understanding wall synthesis in plants was made by
It therefore remains difficult to identify with confidence cellulose synthase or other polysaccharide synthase genes on the basis of their sequence similarity with the cotton GhCesA genes. At this early stage in the characterization of genes involved in wall synthesis in higher plants, a gene knockout system, which could be used to evaluate rapidly the effects of candidate genes on the synthesis of cell wall polysaccharides, would prove valuable in assigning functions to the wide array of cellulose synthase–like genes. One such system is virus-induced gene silencing (VIGS), which can be used to examine gene function in Nicotiana spp. In this system, genes or gene fragments of interest are inserted into a modified potato virus X (PVX; potexviral group) cDNA, and RNA transcripts are prepared in vitro for infection of N. benthamiana seedlings (
The precise mechanism of silencing has not been defined, but it may involve the formation of double-stranded RNA ( Here, we have used VIGS to show that one CesA homolog from Nicotiana spp silences an endogenous cellulose synthase gene and very probably encodes a cellulose synthase, whereas another Nicotiana spp cDNA, which is 80% identical with the first, produces a completely different phenotype. This emphasizes the need for development of discriminating systems for the functional analysis of polysaccharide synthase genes in wall synthesis in higher plants.
Isolation of cDNAs from N. tabacum
To isolate the N. tabacum homologs of the cotton GhCesA gene, PCR primers were designed so that the PCR products would start at the same point at their 3' ends, just 3' to the encoded QXXRW motif found in the homologous region (HR3) of all CesA genes. The positioning of the 5' primers allowed the inclusion of different lengths of the adjacent plant-specific insertion regions (the conserved plant-specific CRP4 region and the hypervariable HVR2 regions) and, in the case of the longest cDNA, inclusion of the homologous region HR2 (Fig 1A; The three N. tabacum cDNAs so obtained were designated NtCesA-1a, NtCesA-1b, and NtCesA-2. They are 670, 377, and 485 bp long, respectively, and their nucleotide sequences have been lodged in the EMBL and GenBank databases with accession numbers AF233892 for NtCesA-1a and -1b and AF233893 for NtCesA-2. The nucleotide sequences of cDNAs NtCesA-1a and NtCesA-1b are identical where they overlap, and the cDNAs clearly represent fragments of the same gene. The 485-bp NtCesA-2 cDNA corresponds to a related but distinct gene; it shares 80% identity with the NtCesA-1a cDNA at the nucleotide level. The sequences of the N. tabacum cDNAs are compared with the corresponding sequences of the cotton GhCesA-1 gene and the Arabidopsis AtCesA-1 (rsw1) gene in Fig 1B. Sequence alignments show that the NtCesA-1 and NtCesA-2 cDNAs fall into the CesA group of the CesA superfamily (C. Somerville and T. Richmond, http://cellwall.stanford.edu/tree.html).
CesA Gene Family of Nicotiana spp
Phenotypes of Infected Plants
Anatomic Changes
Transcriptional Activity of N. benthamiana CesA Genes Examining the effects of VIGS on CesA-1 mRNA levels therefore required a cDNA encoding a region outside the NtCesA-1a sequence. Using anchor-ligated PCR, we amplified a 399-bp cDNA from N. benthamiana leaf RNA preparations. The 399-bp fragment included a 99-bp sequence at its 3' end that was 98% identical with the 99-bp sequence at the 5' end of the NtCesA-1a cDNA and 100% identical at the amino acid sequence level. This overlap confirmed that the amplified 399-bp fragment corresponded exactly to the NtCesA-1a gene. The 399-bp cDNA fragment extended beyond the 5' end of NtCesA-1a by 300 bp, and the sequence of this 5' region of the cDNA fragment could therefore be used to design primers for RT-PCR (Fig 1A). Quantitative RT-PCR was realized by adjusting the number of cycles during the PCR reaction until easily detectable but submaximal amounts of DNA were amplified. The amounts of amplified DNA were subsequently quantitated from a digital camera image of the gel. Care was taken to ensure that any apparent decreases in CesA-1 mRNA abundance did not result simply from an overall decrease in nuclear transcription attributable to the PVX infection in VIGS plants. Control RT-PCR amplifications were therefore performed using primers for mRNA encoding glyceraldehyde-3-phosphate dehydrogenase (GAPDH), an enzyme of the glycolytic pathway, because GAPDH mRNA abundance should be a reasonable measure of the relative metabolic activity of the cells. Reductions in RT-PCR products amplified with both GAPDH and NtCesA-1 primers were observed in the PVX–NtCesA-1a and PVX–NtCesA-1b plants (Fig 4A and Fig 4B). For the PVX–NtCesA-1 plants, the RT-PCR experiments indicated that GAPDH mRNA abundance was ~66% of that observed in PVX control plants and that the NbCesA-1 mRNA decreased to ~43% of that observed in the PVX control plants (Table 1).
Linkage Analysis and Major Polysaccharides in the Cell Walls
The most abundant polysaccharide types in the wall preparations were subsequently deduced from the linkage compositions shown in Table 2, basing calculations on the totals for individual glycosyl residues that are characteristic of well-defined wall polysaccharides. These calculations embody certain assumptions about polysaccharide structures but are widely used as good indicators of the contents of specific polysaccharide types in plant cell walls (
To confirm the decreases in cellulose in walls of PVX–NtCesA-1a and PVX–NtCesA-1b plants indicated by methylation analyses (Table 2), we applied the acetic acid/nitric acid procedure for estimating crystalline cellulose (
Degree of Esterification of Pectic Polysaccharides
To further examine the lesser esterification of pectic polysaccharides in walls of the PVX–NtCesA-1a plants as well as to locate the Ca2+-pectate, we stained leaf sections of those plants with NiCl2/Na2S, which binds to deesterified polygalacturonates, presumably in regions of Ca2+ cross-linking (
Protein and Amino Acid Composition
The functions of three cDNAs corresponding to putative cellulose synthase genes from Nicotiana spp were analyzed with the VIGS system. In this system, endogenous plant genes can be silenced by high expression of homologous DNA fragments carried in the genome of the infecting virus ( Although the sequences of NtCesA-1 and NtCesA-2 were 80% identical where they overlapped, infection of N. benthamiana seedlings with RNA carrying the NtCesA-1 or NtCesA-2 sequences produced dramatically different effects (Fig 2). Thus, growth of the PVX–NtCesA-1a and PVX–NtCesA-1b plants was severely inhibited after infection compared with those plants infected with PVX–NtCesA-2 and the PVX control. The stunted growth patterns of plants infected with PVX–NtCesA-1a and PVX–NtCesA-1b were highly reproducible and were characterized not only by much shorter internode lengths but also by the presence of smaller leaves, which were both "lumpy" in form (Fig 2B and Fig 2C) and "crisp" in texture.
Examination of the leaves from the PVX–NtCesA-1a and PVX–NtCesA-1b plants by light and electron microscopy showed extensive disruption of the surfaces of infected leaves, particularly on their undersurfaces and in the vicinity of vascular bundles. The latter effect presumably reflected the spread of infection across the leaves as the virus moved through the vascular system ( To further investigate the possibility that the swelling of cells in plants infected with PVX–NtCesA-1a or PVX–NtCesA-1b was attributable to a decrease in cellulose content, we isolated cell walls from infected leaves for analysis. The cellulose content of walls isolated from PVX–NtCesA-1a and PVX–NtCesA-1b leaves was ~25% less than that in walls of the PVX control plants, as measured by methylation analysis (Fig 5). The lower cellulose content of walls from PVX–NtCesA-1a and PVX–NtCesA-1b plants was confirmed by the acetic acid/nitric acid procedure for estimating crystalline cellulose (Table 3). The loss of cellulose in walls of the PVX–NtCesA-1a and PVX–NtCesA-1b plants was accompanied by a 45% increase in homogalacturonan (Fig 5). Furthermore, the degree of esterification of pectic polysaccharides decreased from ~50% in walls of control plants to ~33% in walls of plants infected with PVX–NtCesA-1a or PVX–NtCesA-1b. That pectin esterification was less in the infected plants than in the control plants was confirmed by NiCl2/Na2S staining of tissue sections (Fig 6). Cell walls were also isolated by microdissection from the characteristic lumps that were observed on leaves of the PVX–NtCesA-1a and PVX–NtCesA-1b plants, although yields of these walls were low and replication of experiments was difficult. Nevertheless, their cellulose content was reduced by 50 to 75% compared with walls from the PVX control plants (data not shown).
The increase in pectin content of walls, together with much less esterification of the pectic polysaccharides (Fig 5), suggested that plants infected with the PVX–NtCesA-1a and PVX–NtCesA-1b constructs specifically compensated for the decreased cellulose content of walls through the deposition of additional pectic polysaccharides. Furthermore, the presence of longer sections of pectic polysaccharides containing deesterified galacturonosyl residues would allow the formation of more extensive Ca2+-bridged junction zones ( Associated with the reduction in cellulose content of the cell walls in the PVX–NtCesA-1 plants was an apparent decrease in NtCesA-1 mRNA in the infected leaves (Fig 4 and Table 1). Although these RT-PCR results must be interpreted cautiously (Table 1), they provide prima facie evidence that infection with the PVX–NtCesA-1 constructs results in a marked decrease in CesA-1 mRNA in PVX–NtCesA-1 plants and that the decrease is greater than that observed for mRNA encoding the cellular "housekeeping" enzyme, GAPDH (Fig 4 and Table 1). On the basis of these results, we conclude that the NtCesA-1a and NtCesA-1b cDNAs from N. tabacum have silenced a cellulose synthase gene or genes and, moreover, that the data demonstrate the usefulness of VIGS for functional analysis of unknown genes. We further conclude that on the basis of differences in phenotypes (Fig 2) and cell wall compositions (Fig 4) of the PVX–NtCesA-1a and PVX–NtCesA-2 plants, the VIGS system can discriminate between DNA fragments from closely related genes. The NtCesA-2 cDNA, which shares 80% overall sequence identity with the NtCesA-1a cDNA (Fig 1B), does not appear to inhibit cellulose synthesis, although RT-PCR showed that CesA-2 mRNA was detectable in all tissues of control plants examined, albeit in lesser amounts than CesA-1 mRNA (data not shown). Because the NtCesA-1a and NtCesA-2 cDNAs are 85% identical in the conserved HR3 region of the gene (Fig 1A), the differences in their abilities to silence the cellulose synthase gene therefore appear to lie in the HVR2 (72% identity) and CRP4 (77% identity) regions of the genes (Fig 1A). We note that the CesA-1 cDNA fragments might silence, in addition to the NbCesA-1 gene itself, other very closely related genes. Although we can conclude that the NtCesA-1 cDNA corresponds to a cellulose synthase gene, at this stage the data do not allow us to rule out a role for NtCesA-2 in cellulose synthesis. Five or six genes in Nicotiana spp have a high degree of sequence similarity with the NtCesA-1 gene. The NtCesA-2 gene could indeed encode a cellulose synthase, but if the NtCesA-2 isoenzyme expressed in leaf tissue at very low amounts at the time of VIGS was only one of several cellulose synthases involved in wall synthesis, no obvious phenotypic effect might be observed. This raises another point about the use of VIGS and the interpretation of VIGS data with genes that are members of multigene families. The optimal strategy for VIGS as a means of assigning function to a particular member of a gene family would have two stages. First, use of cDNA fragments corresponding to the most highly conserved domains of the gene would reveal the collective functions of the multigene family. Having established a role for the gene family in the trait of interest, VIGS vectors based on the least-conserved regions of the gene could subsequently be used to assign a function to the individual gene or to subsets of the gene family.
The phenotypic differences observed here in plants infected with the closely related (80% sequence identity) PVX–NtCesA-1 and PVX–NtCesA-2 constructs serve to emphasize that sequence comparisons alone do not allow an unequivocal identification of cellulose synthase genes. Neither does it allow identification of genes encoding other polysaccharide synthases that participate in wall synthesis in higher plants. This point can be illustrated by comparing nucleotide sequence identities of the cellulose synthase genes for which proof-of-function is available. The N. tabacum NtCesA-1a sequence used here is 69 and 72% identical with corresponding regions of the CesA cellulose synthase genes of cotton (GhCesA;
More likely, proof-of-function for the many Csl genes that have been identified in plants (
Another question that should be addressed relates to the relative reduction of cellulose content in plants infected with the PVX–NtCesA-1a and PVX–NtCesA-1b constructs. Why is cellulose content of the walls reduced by only 25%, and why is NbCesA mRNA apparently reduced by a similar amount? First, preexisting walls would contain normal amounts of cellulose; however, if cellular activity, including wall synthesis, were stopped very rapidly after infection, then large decreases in final cellulose content of the walls would not be expected. An additional explanation is that the magnitude of the effect on wall composition depends on the developmental age of the cell at the time of its infection. The virus is transported through the phloem from the point of infection to unloading points in the vascular tissue and hence moves between cells through plasmodesmata ( In support of these suggestions is the observation that cell structure appears to be most affected in the vicinity of vascular bundles (Fig 3), where the surrounding cells are likely to be infected at an earlier stage of their development than are cells that are farther away. Furthermore, not all epidermal cells balloon from the surface, which suggests some variation in the strength of walls. The considerable variation in pectate staining of walls might indicate that the degree of compensation of cellulose loss through alterations to pectic polysaccharides also varies between cells (Fig 6). A further possible explanation for the observation that VIGS does not lead to a greater loss of cellulose in walls would be that N. benthamiana has other cellulose synthase genes that are not silenced by the PVX–NtCesA-1a or PVX–NtCesA-1b constructs, as discussed earlier. When a whole-plant cDNA library was screened with the NtCesA-1a fragment, four additional, nonidentical cDNAs were isolated (data not shown), an indication that N. benthamiana contains a number of homologous genes. Whether any of these other genes encodes additional cellulose synthase isoforms remains to be demonstrated. Answers to the specific questions regarding viral infection patterns and VIGS in infected cells will require the availability of antibodies or stains that would allow changes in specific components of the cell wall to be monitored in situ.
Finally, the silencing of a cellulose synthase gene by VIGS has revealed homeostasis in the wall, through which decreased amounts of cellulose are compensated for by an increase not only in the amount of pectic polysaccharides but also in the abundance of deesterified regions in those pectic polysaccharides. These quantitative and qualitative changes imply the existence of feedback loops interconnecting the cellulose synthesis system with the cellular and enzymic machinery that controls pectin biosynthesis. Given that pectic polysaccharides are generally deposited in the wall in a highly esterified form (
Isolation of cDNAs
The PCR program involved 35 cycles at 94°C for 40 sec, 49°C for 40 sec, and 72°C for 90 sec in a reaction mixture containing a standard PCR buffer with 200 µM deoxynucleotide triphosphates and 10% DMSO. Products were cloned into the EcoRV site of pBluescript SK+ (Stratagene, La Jolla, CA), and their identity was verified by nucleotide sequence analysis (
DNA Constructs and Seedling Infection
Infectious RNA molecules were produced by in vitro transcription of the DNA constructs, as described previously (
Transcript Levels and Reverse Transcription–PCR For reverse transcription (RT)–PCR, total RNA was extracted from N. benthamiana leaves, as described above. The quality and concentration of RNA preparations were accurately determined with a UV-visible spectrophotometer (model Cary 50BIO; Varian, Walnut Creek, CA) and using the Cary WinUV RNA–DNA estimation software. Samples of total RNA (1 µg) were used in a first-strand cDNA synthesis reaction with the reagents supplied in the Thermoscript RT-PCR System (Gibco BRL), according to the manufacturer's instructions. The RNA samples were primed with oligo(dT)20, and the reverse transcriptase reaction was performed at 52°C for 1 hr in a final volume of 50 µL. Samples from each reaction (2 µL) were used in a 50-µL PCR mixture containing the single-stranded cDNA template, Taq polymerase buffer (Gibco BRL), 0.2 mM dNTPs, 1.5 units of Taq polymerase (Gibco BRL), and each glyceraldehyde-3-phosphate dehydrogenase (GAPDH) oligonucleotide at 1 µM or each N. benthamiana oligonucleotide at 2 µM. Amplification of the extended NbCesA-1 sequence was performed for 30 cycles with the oligonucleotides 5'-TGCCATGAGTGCACTGGTTCGAGTG-3' and 5'-TACGGTTGGCATATCGATCATTCC-3' at 94°C for 30 sec, 56°C for 30 sec, and 72°C for 30 sec to yield a 211-bp product. This product was purified from agarose gels by using the BRESAclean DNA purification kit (Geneworks, Adelaide, Australia) and sequenced on the DNA sequencer to confirm its identity. GAPDH cDNA was amplified by using the two oligonucleotides GAPDH5 (5'-CAGGAACCCTGAAGATATCCC-3') and GAPDH3 (5'-GCAGTTGGTACTCTGAAGGCC-3'), which were based on conserved regions in the published sequences for the potato and tomato GAPDH genes (GenBank accession numbers U17005 and U97257, respectively). The PCR cycles each consisted of 94°C for 30 sec, 50°C for 30 sec, and 72°C for 1 min. The 550-bp product was purified from an agarose gel and sequenced to confirm its identity. To quantitate PCR band intensities, we scanned digital camera images of the gels and assigned individual bands a relative intensity value by using Gel-Pro Analyzer version 2.0 software (Media Cybernetics, Atlanta, GA). Preliminary experiments in which PCR amplifications were performed for 27, 30, 32, and 35 cycles showed that easily detectable DNA bands were visible after 30 cycles and that the intensities of these bands remained well below the maximal intensities, which were observed after 35 cycles.
Microscopy
To locate polygalacturonate in cell walls, we fixed sections of leaves in 80% ethanol, rinsed them in water, stained them with 15 mM NiCl2 for 30 min, and rinsed them in water. Color was developed in a solution of 1% Na2S (
Preparation of Cell Walls
Carboxyl Reduction of Uronic Acids
Methylation Analysis of Cell Walls Acetylation of the partially methylated alditols was performed in 0.5 mL of acetic anhydride at 100°C for 2.5 hr. The anhydride was destroyed with 2.0 mL of H2O, and the partially methylated alditol acetate derivatives were partitioned into 1.0 mL of dichloromethane, washed three times with water, and dried.
The derivatives were separated and analyzed in a gas chromatograph (model 6890; Hewlett-Packard) linked to a mass spectrometer (model 5973), using a 25 m x 0.22 mm (i.d.) BPX70 column (SGE, Melbourne, Australia). Identification of the derivatives and deduction of the glycosidic linkages were based on published mass spectra (
Calculation of the Polysaccharide Composition of Cell Walls Rhamnogalacturonan I (RGI) = 2 (2-Rha + 2,4-Rha); this assumes a repeating motif with 4-GalA; the branched residues are not included. Galacturonan = 4-GalA - (2-Rha + 2,4-Rha) + 3,4-GalA; the 4-GalA not accounted for in RGI is assigned to galacturonan; all GalA esterifications are assigned to galacturonan; the branched residues are not included. Arabinan = 5-Ara + 3,5-Ara + t-Arap + (t-Araf - 2-Xyl); the t-Araf remaining after accounting for t-Araf in arabinoxyloglucan is assigned to arabinan. 4-Galactan = 4-Gal + 2 (2,4-Gal); this assumes that the branched residues are t-Gal. Xylan = 4-Xyl + 2 (2,4-Xyl); this assumes that the branches comprise t-GlcA and t-Araf. Xyloglucan = 1.5 (4,6-Glc) + 2 (4,6-Glc) + 2-Xyl; this assumes three 4-Glc units for every two 4,6-Glc branched units, each 2-Xyl branch terminates with t-Araf, and the remaining 4,6-Glc units are branched with t-Xyl. Galactoglucomannan = 2 (4-Man + 4,6-Man) + 4,6-Man; this assumes a repeating motif of the mannosyl units with 4-Glc, and branches terminate with t-Gal; traces of 2-Gal indicate some disaccharide branches. Cellulose = 4-Glc - 1.5 (4,6-Glc) - (4-Man + 4,6-Man); the 4-Glc not assigned to arabinoxyloglucan or galactoglucomannan is assigned to cellulose. Arabinogalactan = 6-Gal + 3-Gal; the trace amounts of 3,6-Gal indicate the presence of a relatively unbranched type II arabinogalactan. Other = the sum of linkages not assigned to specific polysaccharides.
Determination of Crystalline Cellulose
This work was supported by grants from the Grains Research and Development Corporation of Australia (to G.B.F.) and from the Australian Research Council (to A.B.). The Sainsbury Laboratory is supported by the Gatsby Charitable Foundation. We thank Monika Doblin, Zofia Felton, Maureen McCann, Bruce Stone, and Brian Wells for their valuable contributions to aspects of this work.
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-amylase (Sigma; type 1-A) in 0.5 mL of 0.5 M sodium acetate buffer, pH 6.5, were added. The samples were kept at 22°C for 3 hr with constant shaking. Starch degradation was monitored by iodine/iodide staining. After this treatment, the preparations still contained adherent cytoplasmic material, which was probably proteinaceous in nature. Preparations were therefore extracted twice with 1.5% sodium dodecyl sulfate at 60°C, washed three times in H2O, and freeze-dried. Although water-soluble components of the walls would have been lost during isolation, we expected that little if any cellulose, pectin, or other wall polysaccharides would be lost under these conditions. 
