Logo of pnasPNASInfo for AuthorsSubscriptionsAboutThis Article
Proc Natl Acad Sci U S A. Sep 25, 2007; 104(39): 15566–15571.
Published online Sep 18, 2007. doi:  10.1073/pnas.0706592104
PMCID: PMC2000526
Plant Biology

Genetic evidence for three unique components in primary cell-wall cellulose synthase complexes in Arabidopsis


In higher plants, cellulose is synthesized at the plasma membrane by the cellulose synthase (CESA) complex. The catalytic core of the complex is believed to be composed of three types of CESA subunits. Indirect evidence suggests that the complex associated with primary wall cellulose deposition consists of CESA1, -3, and -6 in Arabidopsis thaliana. However, phenotypes associated with mutations in two of these genes, CESA1 and -6, suggest unequal contribution by the different CESAs to overall enzymatic activity of the complex. We present evidence that the primary complex requires three unique types of components, CESA1-, CESA3-, and CESA6-related, for activity. Removal of any of these components results in gametophytic lethality due to pollen defects, demonstrating that primary-wall cellulose synthesis is necessary for pollen development. We also show that the CESA6-related CESAs are partially functionally redundant.

Keywords: gametophytic lethal, isoforms, pollen, cellulose synthesis, mutant

Cellulose, a composite of paracrystalline β-1,4-glucan chains, is one of the major components of the cell wall and provides major tensile strength for the wall matrix (1, 2). The deposition of cellulose is catalyzed by cellulose synthase complexes (CSCs) located within the plasma membrane (3). These CSCs are believed to hold as many as 36 individual cellulose synthase (CESA) proteins, which are proposed to be the catalytic subunits of the CSCs. The CESAs were originally identified through sequence similarities to bacterial cellulose synthases (4) and have been shown by immunological techniques to be components of the terminal rosette structures found at the ends of cellulose strands (5). The Arabidopsis thaliana genome harbors 10 CESA genes (6). Immunoprecipitation and mutant analyses showed that three of the CESA genes, CESA4, -7, and -8, directly interact and are necessary for cellulose deposition during secondary cell wall formation (7, 8). These observations demonstrated that at least three CESA proteins are necessary for secondary cell wall cellulose deposition.

The constitution of the primary-wall CESA complex is less clear, as are the roles for most of the other seven CESA genes in Arabidopsis. CESA1 is inferred to be required for primary-wall synthesis because a temperature-sensitive allele, rsw1-1 (9), results in temperature-dependent disappearance of rosettes from the plasma membrane, and strong alleles are embryo lethals (10, 11). The isoxaben-resistant mutants ixr1-1, 1-2, and 2-1 correspond to mutations in CESA3 and CESA6, respectively (12, 13). Because these mutations confer resistance to the cellulose synthesis-inhibiting herbicide isoxaben, both proteins are inferred to be components of CSCs involved in primary cell wall synthesis. CESA3 has also been shown to be coexpressed with CESA1, which is consistent with the idea that the corresponding proteins are in the same complex (14).

If the CESA complex requires CESA1, -3, and -6 to synthesize cellulose, mutations in the genes corresponding to the individual CESAs should result in similar phenotypic traits. Strong mutant alleles of the CESA1 gene are embryo lethal (10, 11). However, null mutations affecting CESA6, i.e., the procuste mutants prc1-1 to 1-12, exhibit only slight deficiencies in cell elongation and are viable (15). No null mutant for CESA3 has been reported. The difference in phenotypes between the cesa1 and cesa6 mutations suggests either that there is an unequal contribution of the CESAs or that several CESAs are functionally redundant. Two studies concerning expression of the remaining CESA genes (i.e., CESA2, -5, -9, and -10) have been reported (16, 17). Reduced expression of CESA2 using an antisense construct resulted in a weak temperature-sensitive phenotype affecting internode lengths in A. thaliana Col-0 (16). A cesa2 null mutation in A. thaliana Ler was reported to result in a severely dwarfed phenotype (17). No mutant analyses have been reported for CESA5, -9, and -10.

Here, we present genetic evidence that the CSC involved in primary-wall synthesis has two unique components, CESA1 and -3, and that a small family of CESA6-related proteins provides the third essential component of the primary CSC. Null mutations in these essential CESAs result in aberrant pollen morphology and male sterility. We also show that the weak phenotypes exhibited by the prc mutants are a result of functional redundancy by other CESAs such as CESA2. However, complementation of prc1-1 with CESA2 reveals only partial functional conservation between the CESA6-related proteins.


cesa1 and cesa3 Null Mutants Are Gametophytic Lethals.

Previously described mutations in CESA1 and -6 cause growth defects and cellulose deficiencies (10, 11, 15). However, the strong mutant phenotypes associated with CESA1 (rsw1-2, -1-20, and -1-45) are due to missense mutations and may not be nulls (10, 11). In addition, no null mutants have been reported for CESA3. To assess the phenotypes of cesa1 and cesa3 null mutants, we sought to identify homozygous mutant lines from two and three populations carrying T-DNA insertions in the genes, respectively [supporting information (SI) Table 2]. We did not obtain any homozygous plants for any of the lines, indicating that the mutations affected either embryo development or fertility. Genotyping of progeny from heterozygous T-DNA plants revealed that ≈50% of the progeny were wild type and 50% were heterozygous for the T-DNA inserts for both CESA1 and -3, suggesting that disruption of the CESA1 and -3 genes cause fertility problems. Structural features of female and male reproductive organs were, therefore, examined by using light microscopy and SEM. Fig. 1 B–D shows that ≈50% of the pollen grains were significantly deformed (Table 1). To assess whether these deformations led to sterile pollen grains, we used anthers from the heterozygous cesa1 and cesa3 plants for fertilization of wild-type plants. None of 30 resulting F1 plants analyzed contained a T-DNA insertion, corroborating that the pollen grains containing the T-DNA insert were defective. Furthermore, pollen-tube growth assays revealed that none of the deformed pollen grains produced pollen tubes (SI Fig. 5 A and B). Mutant pollen was also found to contain little, if any, cellulose as determined by Calcofluor white staining (SI Fig. 5 A and B).

Fig. 1.
Mutant phenotypes. (A) Seven-week-old plants showing shorter inflorescence stems in prc1-1 cesa2 mutants (plant 4) compared with wild-type (plant 1) and mutant parent lines, cesa2 (plant 2), and prc1-1 (plant 3). (Scale bar, 5 cm.) (B–D) Environmental ...
Table 1.
Pollen-grain deformation in cesa mutants

CESA2 Is Functionally Redundant with CESA6.

Whereas CESA6 null mutants result in only subtle growth phenotypes, null mutants for CESA1 and CESA3 result in pollen defects. This observation suggests that other CESAs may be functionally redundant with CESA6. Three CESAs, CESA2, -5, and -9, have high sequence similarity to CESA6 in Arabidopsis (SI Fig. 6 and SI Table 3), which indicates that these proteins may be functionally related to CESA6. Only one of these CESAs, CESA2, is expressed in tissues corresponding to rapidly dividing cells as assessed by promoter–reporter gene fusions (SI Fig. 7), consistent with data from various microarray analyses (i.e., Genvestigator, https://www.genevestigator.ethz.ch/; ref.18) and a previous study (17). CESA2 is, therefore, a likely candidate to be at least partially functionally redundant with CESA6 in the prc1-1 background.

To investigate whether CESA2 affects root or hypocotyl elongation, we identified homozygotes for two independent CESA2 T-DNA insertion mutations (SI Table 2). The homozygous cesa2 seedlings did not show any phenotype compared with wild type when grown in light (Fig. 2A). However, etiolated cesa2 seedlings were significantly shorter (13.9% shorter; P < 0.001) compared with wild type (Fig. 2B). These observations correlate well with the CESA2 promoter β-glucuronidase (GUS) activity (SI Fig. 7) and suggest that CESA2 has a direct role in elongation of etiolated hypocotyls in Arabidopsis Col-0. Our results differ from a previous study of an activator (Ac)-induced cesa2 mutation in the Landsberg erecta background, in which a strong decrease in growth was observed (17). We infer that there is an important genetic background difference between the Col ecotype, used here, and the Landsberg erecta ecotype with respect to some aspect of cellulose synthase function.

Fig. 2.
cesa2 and prc1-1 cesa2 mutant phenotypes. (A and B) Five-day-old light-grown (A) and etiolated (B) mutant seedlings. From left to right: wild type, cesa2-1, prc1-1, and prc1-1 cesa2-1. (C and D) Differential interference contrast microscopy images of ...

To explore possible functional redundancy of CESA2 and CESA6, we generated reciprocal crosses of prc1-1 and cesa2-1. Double-mutant seedlings had significantly shorter and swollen roots, and etiolated hypocotyls were extremely short and swollen compared with the single-mutant parent lines, respectively (Fig. 2 A and B). In addition, mutant plants had stunted growth and reduced organ sizes compared with wild-type and mutant parent lines (Fig. 1A).

To assess the cellular organization of the root in the prc1-1 cesa2 double mutants, we analyzed cross-sections of seedling roots by using light microscopy. Although the lateral organization of the root seemed to be preserved in the double mutant, all cells appeared swollen, which also seemed to affect cellular adhesion (Fig. 2 C and D). To investigate this observation in more detail, sections from light-grown seedling roots were analyzed by using transmission electron microscopy (TEM). SI Fig. 8 A and B shows that the prc1-1 cesa2 double mutant had wider spacing of the tripartite cell corners, which mainly consists of pectic cell-wall polymers. To investigate whether prc1-1 cesa2 double-mutant seedlings contained altered levels of such cell-wall polymers compared with control seedlings, we measured neutral sugars and uronic acids (SI Fig. 8C). The prc1-1 cesa2 double mutants had significantly more arabinose, and possibly more uronic acids, compared with wild type, indicating a compensatory increase in cell-wall polymers containing these sugars.

The loss of both CESA2 and CESA6 would be expected to cause a reduction in cellulose content. Interestingly, we did not observe a significant decrease in cellulose for the prc1-1 cesa2 double mutants compared with prc1-1 [90% ± 2% (cesa2-1), 70% ± 5% (prc1-1), and 69% ± 2% (prc1-1 cesa2) as compared with wild-type control, respectively]. However, the lateral swelling of root cells and additive phenotype observed in prc1-1 cesa2 compared with prc1-1 suggest additive cellulose-deposition defects in the double mutant. To investigate if the microfibril orientations were altered in the mutants, we analyzed cell walls by using field emission SEM of longitudinal sections from seedling roots (Fig. 2 E and F). The orientation of microfibrils in the prc1-1 cesa2 double mutants appeared to be more randomized compared with prc1-1 and also appeared to be aggregated to a higher degree than microfibrils in prc1-1 (compare Fig. 2 E with F), possibly explaining the additive phenotype in prc1-1 cesa2 compared with prc1-1.

cesa2 cesa6 cesa9 Triple Mutants Are Gametophytic Lethals.

The additive phenotypes for prc1-1 cesa2 double mutants suggest functional redundancy between CESA2 and CESA6. However, the prc1-1 cesa2 double mutant did not exhibit comparable phenotypes to cesa1 and cesa3 mutants (i.e., gametophytic lethality). Based on sequence similarity, CESA5 and CESA9 are likely candidates to also be functionally redundant to CESA2 and CESA6 (SI Fig. 6). No reporter gene activity was, however, evident for CESA5 and -9 in elongating tissues where primary cellulose synthesis is prevalent (SI Fig. 7). To assess whether CESA5 and CESA9 are expressed at low levels in these tissues, we isolated mRNA from dark-grown hypocotyls and light-grown seedling roots and performed semiquantitative RT-PCR (Fig. 3A). Both CESA5 and CESA9 transcripts were detected, suggesting that both genes are active in these tissues. To determine if one or both of these CESA genes plays a significant role during pollen maturation, we investigated the GUS expression of promoter:GUS fusions of CESA5 and CESA9 in floral tissues. No GUS activity was observed for CESA5 in these tissues (data not shown). However, CESA9 showed strong expression in pollen (Fig. 3B). Two independent homozygous cesa9 TDNA mutant lines (SI Table 2) did not exhibit any apparent phenotype (data not shown). To further examine functional redundancy among CESA2, CESA6, and CESA9 we generated different combinations of prc1-1, cesa2-1, and cesa9-1 double and triple mutants. The prc1-1 cesa9 and cesa2 cesa9 double mutants appeared indistinguishable from prc1-1 and cesa2 parent plants, respectively (data not shown). We were, however, unable to generate homozygous prc1-1 cesa2 cesa9 triple mutants. When we assessed the progeny from the heterozygous triple mutants (cesa2+/− prc1-1 cesa9), we found that 49% of plants were heterozygous for the T-DNA insertion affecting CESA2 and 51% were wild type. This observation indicates that the triple mutant has fertility problems similar to cesa1 and cesa3.

Fig. 3.
cesa2 cesa6 cesa9 triple mutants are pollen deficient. (A) Expression of CESA5, -9, and -10 in 5-day-old light-grown seedling roots and etiolated seedlings assessed by RT-PCR. ACTIN1 was used as control. (B) Expression analysis of CESA9 in floral tissues ...

To analyze structural features of the female and male reproductive organs of the cesa2+/− prc1-1 cesa9 line, we examined the tissues by using electron microscopy and SEM. Fig. 3 C and D illustrates that approximately half of the pollen grains were deformed in the cesa2+/− prc1-1 cesa9 triple mutants (Table 1). Analogous to cesa1 and cesa3 back-crosses, no transmission (30 plants investigated) of the cesa2 T-DNA insert was observed when anthers from cesa2+/− prc1-1 cesa9 plants were used for fertilization of wild-type plants. These results show that CESA2, -6, and -9 exhibit functional redundancy, and we refer to these three genes as the “CESA6 family.”

To assess structural features in the cellulose-deficient pollen grains produced by cesa2+/− cesa6 cesa9 plants, high-pressure freezing in combination with TEM was used to examine the pollen cell wall. As expected from the environmental SEM analysis, the prc1-1 cesa2 cesa9 triple-mutant pollen grains were severely deformed (compare Fig. 1 E with F). Interestingly, the cell walls appeared slightly thicker in the mutant pollen compared with wild type (Fig. 3 E and F), possibly because of compensation by other cell-wall polymers in the mutant pollen. Furthermore, the cell walls were unevenly deposited in the mutant compared with wild type (compare Fig. 3 E with F), compatible with the idea that the cellulose microfibrils constitute a framework for deposition of other cell-wall polymers (1).

Functional Redundancy Between CESAs Is Only Partial.

Although the CESA2 and -6 expression profiles largely overlap, close examination reveals that the overlap is not complete (SI Fig. 9 A–C). Whereas CESA6 is expressed in rapidly elongating tissues (i.e., at the top of the hypocotyl directly below the cotyledons and in the root elongation zone), CESA2 expression is slightly shifted toward zones that are less rapidly expanding. To investigate whether CESA2 may take over the function of CESA6 in rapidly elongating cells, we generated a construct where a CESA2 cDNA was placed in frame with a yellow fluorescent protein (YFP) cDNA, under the control of the CESA6 promoter (A6:Y-A2; SI Fig. 9D). We then transformed the construct into prc1-1 mutant plants. To assess whether the A6:Y-A2 protein was incorporated into CSCs, we then examined the elongating hypocotyls in etiolated seedlings by using a spinning-disk confocal microscope (19). The fluorescently labeled particles associated with the plasma membrane were indistinguishable from the previously reported particles encoded by the YFP-CESA6 construct (compare SI Fig. 9 E with F; any apparent differences are due to cell–cell variations), suggesting that the YFP-CESA2 protein was functionally incorporated into the CESA complexes. In addition, YFP-CESA2 complex velocities were determined to be similar to YFP-CESA6 velocities (Fig. 4 A–C). These data provide additional evidence that CESA2 and 6 are functionally redundant, and we speculate that similar results would be obtained with a CESA9 fusion. However, although the A6:Y-A2 construct rescued the prc1-1 phenotype (Fig. 4 D and E), the complementation was not complete (compare Fig. 4 F with I). Whereas the A6:Y-A6-complemented seedlings very closely represented the wild-type control (19), the A6:Y-A2-complemented seedlings exhibited swollen roots and did not elongate to the same extent as wild-type seedlings (Fig. 4 F–I). To ensure that the YFP tag did not affect the complementation efficiency, we also generated a construct in which the YFP was excluded (A6:A2). The A6:A2 construct was subsequently transformed into prc1-1 and analyzed. Similar to the A6:Y-A2-transformed prc1-1, the A6:A2-transformed prc1-1 seedlings also exhibited tissue swelling and elongation deficiencies (data not shown). These data suggest incomplete functional redundancy between CESA2 and CESA6. In addition, cesa2 prc1-1 double mutants transformed with the A6:A2 construct exhibited more-severe phenotypes compared with cesa2 single mutants (SI Fig. 9 G and H), corroborating that CESA2 and CESA6 are not completely functionally equivalent.

Fig. 4.
Incomplete functional redundancy between CESA2 and CESA6. (A and B) Kymographs based on time-average images from prc1-1 seedlings complemented by A6:Y-A6 (A) or A6:Y-A2 (B). (C) Velocity measurements of CESA particles established by analyzing multiple ...


Here, we show that the primary wall CSC requires three unique CESA components, corresponding to CESA1, CESA3, and CESA2 CESA6 CESA9 functions, to produce cellulose. This requirement is similar to the secondary CESA complex, in which CESA4, -7, and -8 are necessary (7, 8). Interestingly, it is apparent that one component of the primary wall CSC can be provided by multiple CESA isoforms represented by the CESA6 family members. In view of subtle differences in the expression pattern of CESA2, -6, and -9, the apparent redundancy is possibly connected to developmental differences in cellulose deposition. CESA2 expression corresponds closely with CESA1, 3, and 6 (14), but the overlap does not appear to be complete (SI Fig. 9 A and B). CESA9 is mainly expressed in pollen and anthers, indicating that this CESA, together with CESA1 and -3, may constitute the CSCs in these organs. Additionally, we speculate that CESA5 is a member of the CESA6 family. The expression of CESA5 is mostly observed in light-grown tissues, such as the hypocotyl and leaves, and is therefore likely to participate in cellulose production in such tissues. Consistent with this idea, we have observed additive phenotypic behaviors in hypocotyl and aerial tissues in cesa5 prc1-1 double mutants (data not shown). Since completing this work, it has also come to our attention that Desprez et al. (23) have come to similar conclusions. These authors found that cesa5 prc1-1 double mutants are deficient in hypocotyl growth, and that growth of plants on soil is arrested after rosette leaves are formed. The differences in expression of the CESA2, -5, -6, and -9 genes, therefore, suggest that different compositions of the primary CESA complex may be present depending on developmental stage and tissue types. Whether these differences have a direct effect on the properties of cellulose microfibrils remains uncertain because of the lack of suitable methods for determining the properties of cellulose (i.e., glucan chain, fibril length, or diameter) from different cell types.

Complete cessation of primary wall cellulose production results in pollen sterility. Therefore, it appears that cellulose is an essential wall polymer for sustaining primary wall integrity during pollen development. A number of other genes have also been implicated in wall integrity during pollen formation and for pollen-tube growth (20). Deficiencies in UDP-sugar pyrophosphorylase (usp-2) and in reversibly autoglycosylated proteins (RGPs) result in phenotypic features very similar to the cellulose deficiencies (21, 22). Interestingly, the intine also has an uneven distribution in these cellulose-deficient pollen grains, suggesting that the cellulose microfibrils provide a framework for deposition of other cell-wall polymers. Removal of the microfibrils may in that case result in random incorporation and aggregation of other wall polymers. This feature is also present in the irregular xylem (irx) mutants that are deficient in secondary cell-wall cellulose (7).

Functional redundancy among the CESA2, -6, and -9 genes suggests that one CESA protein may substitute for another. As expected, CESA6 complemented prc1-1 when expressed under a CESA6 promoter. However, CESA2 did not fully complement the prc1-1 mutant when expressed under the same promoter, suggesting that there are functional differences between the CESA6 family members. Indeed, it is evident from a comparison of the CESA sequences that CESA2, -5, -6, and -9 share very high sequence identity at the protein level. However, CESA2 and -9 are clearly different from CESA5 and -6 in two short regions near the N terminus, corresponding to GRGSNDDD and GRGDGFIVD for CESA2 and -9, respectively. These regions are not found in CESA5 and CESA6, indicating functional diversification of CESA2 and -9. More-subtle assays of CESA function will be required to understand why plants carry at least three and probably four functionally redundant CESAs for primary wall synthesis but not for secondary wall synthesis. One possibility is that functional diversity of the CESA6 family is associated with aspects of cellulose deposition that are unique to dividing or expanding cells.

Materials and Methods

Plant Material and Genetic Analysis.

A. thaliana (Col-0) plants were germinated and grown as described in ref. 14. Insertion lines (24) for CESA2, CESA3, and CESA9 listed in SI Table 2 were obtained from the Arabidopsis Biological Resource Center (http://arabidopsis.org). Primers used for PCR and RT-PCR to obtain homozygous insertion lines are listed in SI Table 2.

Light Microscopy.

Lignin was visualized by staining seedlings with phloroglucinol (Matheson, Norwood, OH) (7). Cellulose was visualized by staining pollen and pollen tubes with Calcofluor white (Polysciences, Warrington, PA) and was viewed with an epifluorescence microscope (Leitz DMRB; Leica, Wetzlar, Germany). Roots from 5-day-old seedlings were fixed and embedded (as described below) and viewed by using differential interference contrast settings with an epifluorescence microscope (Leitz DMRB; Leica).

Electron Microscopy.

Environmental SEM was performed on anthers from 5-week-old plants by using a Quanta 200 microscope (FEI, Hillsboro, OR) under a pressure of 50 Pa and a voltage of 12.5 kV with a low-vacuum detector. TEM was performed as described in ref. 11 on roots from 5-day-old seedlings. Field emission SEM was performed as described in ref. 25 by using a Hitachi S-5000 microscope (Hitachi, Yokohama, Japan) at 10 kV.

Anthers from budding and open flowers were immersed in 10% glycerol (wt/vol) as cryoprotectant, placed in 200-μm-deep aluminum planchettes, and cryoimmobilized by using a Wohlwend HPF 01 high-pressure freezer (Sennwald, Switzerland) at 2,100 bar (210 MPa) for <10 ms and stored in liquid nitrogen. Samples were then freeze-substituted over 4 days and thawed under a controlled temperature gradient in 1% osmium tetroxide and 0.1% uranyl acetate in acetone by using a Leica automated freeze substitution system (AFS) as described in ref. 25. Stepwise infiltration with Epon–Araldite was followed by overnight incubation in pure resin and polymerized at 60°C over 1–2 days. One hundred-nanometer-thick sections perpendicular to the long axis were poststained with 2% uranyl acetate in methanol followed by Sato's lead citrate and imaged with a Tecnai 12 TEM (FEI).

Pollen-Tube Growth.

Pollen was dotted onto microscope slides covered with solid pollen-tube growth medium [1 mM CaCl2/1 mM MgSO4/0.01% boric acid/18% (wt/vol) sucrose/0.5% select agar]. Pollen tubes were allowed to grow for ≈6 h at 30°C then visualized by light and fluorescence microscopy (Leitz DMRB; Leica).

Expression of CESAs.

RNA was isolated by using the RNeasy plant mini kit (Qiagen, Valencia, CA). The Qiagen Onestep RT-PCR kit was used for first-strand cDNA synthesis and for subsequent PCR steps using gene-specific primers (SI Table 2). The expression of the ACTIN1 gene (At2g37620) was used as control.

The expression pattern of the CESAs was studied by using the GUS reporter gene. Genomic DNA fragments 978–2,634 bp upstream of the CESA initiating AUG codon (SI Table 2) were amplified by PCR, cloned into pRITA upstream of the GUS gene, and sequenced to confirm that errors had not been introduced. The promoter-GUS cassettes were then cloned as NotI fragments into the Ti plasmid pMLBART in the same orientation as the BAR gene, introduced into Agrobacterium tumefaciens strain ASE by electroporation, and used to transform Col-0 Arabidopsis (27). Transgenic plants were selected with a 1:1,000 dilution of Finale (Hoechst-Roussel Agri-Vet, Somerville, New Jersey) twice a week for 3 weeks. Tissues were stained for GUS activity as described in ref. 13.

Glycosyl Residue Composition Analysis of Total Cell-Wall Material.

Alditol acetate derivatives of the neutral sugars were measured on ball-milled (1–2 h) 5-day-old dark-grown seedlings (27) by using 1 mg (dry weight) of material. The neutral sugars were analyzed by gas chromatography with myo-inositol (Sigma, St. Louis, MO) as internal standard. Cellulose contents were measured (29) on ball-milled (1–2 h) 5-day-old dark-grown seedlings by using 1 mg (dry weight) of material. Uronic acid content was determined (30) by using 500 μg (dry weight) of ball-milled material from 5-day-old etiolated seedlings. Galacturonic acid was used as standard.

DNA Constructs.

The coding region of CESA6 from pBlue-CESA6 (19), was removed by cleavage with SgrAI and XbaI. A CESA2 cDNA was ligated into the cut vector after nested PCR amplification (CESA2UTR-For and -Rev outer, and CESA2-For and -Rev inner, respectively; SI Table 2) and digestion using SgrAI and XbaI. The CESA2-For primer introduced two silent mutations creating a SgrAI restriction site beginning in the second amino acid of CESA2. A CITRINE (YFP) cDNA was amplified (SI Table 2) from a CITRINE donor vector, cut with AgeI, and inserted into the AgeI site between the CESA6 promoter and CESA2 cDNA for the pBlue-A6::Y-A6 construct. The two fragments containing either A6:A2 or A6:Y-A2 were then cut out of pBlueSK+ by using BspEI and PmlI and inserted into pCAMBIA1301 cut with XmaI and PmlI to create binary A6:A2 and A6:Y-A2 constructs, respectively. The verified construct was introduced into the A. thaliana Col-0 cesa6 null mutant (prc1-1) by Agrobacterium-mediated transformation (27). Transgenic plants were identified by rescue of the short-hypocotyl phenotype of the prc1-1 mutant. Lines were established through self-pollination of rescued seedlings.

Confocal Microscopy.

The velocities of individual CESA complexes were acquired from 10-min time-lapse movies of 3- and 4-day-old etiolated seedlings (19).

Supplementary Material

Supporting Information:


We thank Ms. Lisa Thelin for assistance with the phylogenetic analyses, and Kent McDonald and Jessie K. Lee at the Electron Microscope Laboratory (Berkeley, CA) for assistance with TEM experiments. This work was supported in part by Grant DOE-FG02-03ER20133 from the United States Department of Energy (to C.R.S.) and by Contract DE-AC03-76SF00098 from the United States Department of Energy to Lawrence Berkeley National Laboratory.


cellulose synthase
cellulose synthase complex
transmission electron microscopy
yellow fluorescent protein


The authors declare no conflict of interest.

This article contains supporting information online at www.pnas.org/cgi/content/full/0706592104/DC1.


1. Somerville C, Bauer S, Brininstool G, Facette M, Hamann T, Milne J, Osborne E, Paredez A, Persson S, Raab T, et al. Science. 2004;306:2206–2211. [PubMed]
2. Somerville CR. Annu Rev Cell Dev Biol. 2006;22:53–78. [PubMed]
3. Mueller SC, Brown RM., Jr J Cell Biol. 1980;84:315–326. [PMC free article] [PubMed]
4. Pear JR, Kawagoe Y, Schreckengost WE, Delmer DP, Stalker DM. Proc Natl Acad Sci USA. 1996;93:12637–12642. [PMC free article] [PubMed]
5. Kimura S, Laosinchai W, Itoh T, Cui X, Linder CR, Brown RM., Jr Plant Cell. 1999;11:2075–2086. [PMC free article] [PubMed]
6. Richmond T. Genome Biol. 2000;1 reviews3001. [PMC free article] [PubMed]
7. Turner SR, Somerville CR. Plant Cell. 1997;9:689–701. [PMC free article] [PubMed]
8. Taylor NG, Howells RM, Huttly AK, Vickers K, Turner SR. Proc Natl Acad Sci USA. 2003;100:1450–1455. [PMC free article] [PubMed]
9. Arioli T, Peng L, Betzner AS, Burn J, Wittke W, Herth W, Camilleri C, Höfte H, Plazinski J, Birch R, et al. Science. 1998;279:717–720. [PubMed]
10. Beeckman T, Przemeck GK, Stamatiou G, Lau R, Terryn N, De Rycke R, Inze D, Berleth T. Plant Physiol. 2002;130:1883–1893. [PMC free article] [PubMed]
11. Gillmor CS, Poindexter P, Lorieau J, Palcic MM, Somerville C. J Cell Biol. 2002;156:1003–1013. [PMC free article] [PubMed]
12. Desprez T, Vernhettes S, Fagard M, Refregier G, Desnos T, Aletti E, Py N, Pelletier S, Hofte H. Plant Physiol. 2002;128:482–490. [PMC free article] [PubMed]
13. Scheible WR, Eshed R, Richmond T, Delmer D, Somerville C. Proc Natl Acad Sci USA. 2001;98:10079–10084. [PMC free article] [PubMed]
14. Persson S, Wei H, Milne J, Page GP, Somerville CR. Proc Natl Acad Sci USA. 2005;102:8633–8638. [PMC free article] [PubMed]
15. Fagard M, Desnos T, Desprez T, Goubet F, Refregier G, Mouille G, McCann M, Rayon C, Vernhettes S, Höfte H. Plant Cell. 2000;12:2409–2424. [PMC free article] [PubMed]
16. Burn JE, Hocart CH, Birch RJ, Cork AC, Williamson RE. Plant Physiol. 2002;129:797–807. [PMC free article] [PubMed]
17. Chu Z, Chen H, Zhang Y, Zhang Z, Zheng N, Yin B, Yan H, Zhu L, Zhao X, Yuan M, et al. Plant Physiol. 2007;143:213–224. [PMC free article] [PubMed]
18. Zimmermann P, Hirsch-Hoffmann M, Hennig L, Gruissem W. Plant Physiol. 2004;136:2621–2632. [PMC free article] [PubMed]
19. Paredez AR, Somerville CR, Ehrhardt DW. Science. 2006;312:1491–1495. [PubMed]
20. Ma H. Annu Rev Plant Biol. 2005;56:393–434. [PubMed]
21. Drakakaki G, Zabotina O, Delgado I, Robert S, Keegstra K, Raikhel N. Plant Physiol. 2006;142:1480–1492. [PMC free article] [PubMed]
22. Schnurr JA, Storey KK, Jung HJ, Somers DA, Gronwald JW. Planta. 2006;224:520–532. [PubMed]
23. Desprez T, Juraniec M, Crowell E, Jouy H, Pochylova Z, Parcy F, Höfte H, Gonneau M, Vernhettes S. Proc Natl Acad Sci USA. 2007;104:15572–15577. [PMC free article] [PubMed]
24. Alonso JM, Stepanova AN, Leisse TJ, Kim CJ, Chen H, Shinn P, Stevenson DK, Zimmerman J, Barajas P, Cheuk R, et al. Science. 2003;301:653–657. [PubMed]
25. MacKinnon IM, Sturcova A, Sugimoto-Shirasu K, His I, McCann MC, Jarvis MC. Planta. 2006;224:438–448. [PubMed]
26. McDonald K. Methods Cell Biol. 2007;79:23–56. [PubMed]
27. Clough SJ, Bent AF. Plant J. 1998;16:735–743. [PubMed]
28. Blakeney AB, Harris PJ, Henry RJ, Stone BA. Carbohydr Res. 1983;113:291–299.
29. Updegraff DM. Anal Biochem. 1969;32:420–424. [PubMed]
30. Blumenkrantz N, Asboe-Hansen G. Anal Biochem. 1973;54:484–489. [PubMed]

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...