Knockout of the AtCESA2 Gene Affects Microtubule ... - Plant Physiology

Knockout of the AtCESA2 Gene Affects Microtubule Orientation and Causes Abnormal Cell Expansion in Arabidopsis1[C][OA] Zhaoqing Chu2,3, Hao Chen2, Yiyue Zhang, Zhonghui Zhang, Nouyan Zheng, Bojiao Yin, Hongyan Yan, Lei Zhu, Xiangyu Zhao, Ming Yuan, Xiansheng Zhang, and Qi Xie* Temasek Life Sciences Laboratory, National University of Singapore, Singapore 117604 (Z.C., H.Y.); State Key Laboratory of Plant Genomics, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing 100101, China (H.C., Y.Z., Z.Z., N.Z., B.Y., Q.X.); State Key Laboratory of Plant Physiology and Biochemistry, Department of Plant Sciences, College of Biological Sciences, China Agricultural University, Beijing 100094, China (L.Z., M.Y.); College of Life Sciences, Shandong Agricultural University, Taian City, Shandong 271018, China (X. Zhao, X. Zhang); and Graduate University of the Chinese Academy of Sciences, Beijing 100101, China (H.C., Y.Z., Z.Z., B.Y.)

Complete cellulose synthesis is required to form functional cell walls and to facilitate proper cell expansion during plant growth. AtCESA2 is a member of the cellulose synthase A family in Arabidopsis (Arabidopsis thaliana) that participates in cell wall formation. By analysis of transgenic seedlings, we demonstrated that AtCESA2 was expressed in all organs, except root hairs. The atcesa2 mutant was devoid of AtCESA2 expression, leading to the stunted growth of hypocotyls in seedlings and greatly reduced seed production in mature plants. These observations were attributed to alterations in cell size as a result of reduced cellulose synthesis in the mutant. The orientation of microtubules was also altered in the atcesa2 mutant, which was clearly observed in hypocotyls and petioles. Complementary expression of AtCESA2 in atcesa2 could rescue the mutant phenotypes. Together, we conclude that disruption of cellulose synthesis results in altered orientation of microtubules and eventually leads to abnormal plant growth. We also demonstrated that the zinc finger-like domain of AtCESA2 could homodimerize, possibly contributing to rosette assemblies of cellulose synthase A within plasma membranes.

Plant cell shape is a key determinant of plant morphogenesis, which is strongly influenced by the organization of the cell wall (Beeckman et al., 2002). Cellulose, the predominant fiber of higher plant cell walls, is made of Glc subunits linked by b(1,4)glycosidic bonds in unbranched, linear chains. In cell walls, individual cellulose molecules combine in ordered parallel rows to form cellulose microfibrils. 1 This work was supported by the Chinese Ministry of Science and Technology (grant no. 863–2002AA224111/MST 973– 2003CB114304), by the Outstanding Youth Project from the Chinese Natural Science Foundation (grant no. 30325030), and by the Chinese Academy of Sciences (grant nos. KSCX2–YW–N–010 and CXTD– S2005–2 to Q.X.). 2 These authors contributed equally to the paper. 3 Present address: Genome Institute of Singapore, 60 Biopolis Street, Singapore 138672. * Corresponding author; e-mail [email protected]; fax 86–10– 64889351. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Qi Xie ([email protected]). [C] Some figures in this article are displayed in color online but in black and white in the print edition. [OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.106.088393

Primary cell walls contain cellulose microfibrils in a loose, mesh-like array embedded in a relatively soft, gel-like matrix. As cells mature, they may elongate to as much as 10 to 100 times their embryonic lengths. During this elongation, the cellulose meshwork is first loosened and then stretched. Throughout the period of stretching and elongation, new cellulose microfibrils are deposited just at the outer plasma membrane, gradually thickening the wall as it extends and forms the secondary cell wall. Field emission scanning electron microscopy revealed that Arabidopsis (Arabidopsis thaliana) root epidermal cells have typical dicot primary cell wall structure with prominent transverse cellulose microfibrils embedded in pectic substances. Sugimoto et al. (2000) reported that the innermost layer of the Arabidopsis root epidermal cell wall was mainly composed of a transversely aligned fibrous structure that is most likely made of cellulose microfibrils. The catalytic subunit responsible for elongation of glucan chains, cellulose synthase A (CESA), is believed to be a plasma membrane glycosyltransferase. These enzymes are thought to form complexes of rosettes embedded within the membrane (Taylor et al., 2000; Scheible et al., 2001). Each rosette of enzymes, consisting of a circle of six protein subunits arranged in a hexagon, is thought to cast a microfibril of about 5 nm in diameter. The cellulose microfibrils assembled by the enzyme complexes are supposedly

Plant Physiology, January 2007, Vol. 143, pp. 213–224, www.plantphysiol.org Ó 2006 American Society of Plant Biologists

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stationary as they elongate. As a result, the enzyme complexes are thought to be pushed through the fluid membrane bilayer by the growing end of the cellulose microfibril. Genes encoding the cellulose synthase catalytic subunits have been identified in bacteria and plants (Pear et al., 1996). The Arabidopsis genome encodes 10 isoforms of CESA (http://cellwall.stanford. edu). The temperature-sensitive rsw1 mutant encodes AtCESA1 (Arioli et al., 1998). It has reduced cellulose synthesis, accumulation of noncrystalline b-1,4-glucan, and disassembly of cellulose synthase, resulting in widespread morphological abnormalities. In many higher plants, cellulose synthesis is inhibited by isoxaben and thiazolidinone herbicides. The semidominant Arabidopsis mutants ixr1 and ixr2 encode AtCESA3 and AtCESA6, respectively, and conferred isoxaben and thiazolidinone resistance (Scheible et al., 2001; Desprez et al., 2002). The irx1 and irx3 mutants of Arabidopsis have defective AtCESA8 and AtCESA7, respectively (Taylor et al., 1999, 2000), and have severe deficiency in the deposition of cellulose in secondary cell walls, resulting in collapsed xylem cells. Pulldown experiments suggested that CESA7 and CESA8 physically interact and may operate as a heterodimer or multimer (Taylor et al., 2000). Later on, the same group reported that IRX5, IRX3, and IRX1 were coexpressed within cells and all three proteins interacted in detergent-soluble extracts, suggesting that they were required for cellulose synthesis in secondary cell walls (Taylor et al., 2003). Mutants of CESA7 and CESA8 specifically exhibit a cellulose defect in the secondary wall of the xylem, whereas mutants of CESA1 and CESA6 have defects in the primary cell wall (Arioli et al., 1998; Fagard et al., 2000). Two different allelic mutants of CESA8, lew2-1 and lew2-2, were more tolerant to drought stress as well as to NaCl, mannitol, and other osmotic stresses (Chen et al., 2005). Mutations in the endo-1,4-b-glucanase gene, which is highly conserved between mono- and dicotyledonous plants, both in Arabidopsis and rice (Oryza sativa), resulted in a reduction in cell elongation and a decrease in cellulose content, but an increase in pectin content (Sato et al., 2001; Zhou et al., 2006). Although quite a few studies addressed the biological functions of several AtCESA genes, the function of the remaining Arabidopsis CESA genes (i.e. AtCESA2, AtCESA5, AtCESA9, and AtCESA10) are less clear (Scheible and Pauly, 2004). A minor phenotype reported for CESA2 antisense lines in Columbia (Col) background was a small reduction in the stem elongation rate, but not the final plant height, at elevated temperature (Burn et al., 2002). By analysis of tobacco (Nicotiana tabacum) protoplasts and suspension-cultured cells treated with the cellulose synthesis inhibitor isoxaben, Fisher and Cyr suggested that cellulose biosynthesis was required for microtubule organization and stabilization and the cellulose microfibrils and cortical microtubules become coordinated during cell wall formation (Fisher and Cyr, 1998). The same group developed a very powerful method by fusing the microtubule-binding 214

domain of the mammalian microtubule-associated protein 4 (MAP4) gene with the green fluorescent protein (GFP) gene and transient expression of the recombinant protein in epidermal cells to directly view microtubule movement (Marc et al., 1998). A consensus model predicts that cellulose synthase moves along cortical microtubules and uses them as a template for the oriented deposition of microfibrils in the cell wall (Giddings and Staehelin, 1991). Hypothetically, the cortical microtubules control the orientation of cellulose microfibril deposition. Burk and Ye (2002) proposed that the aberrant microtubule orientation caused by a mutation in katanin, a microtubule-severing protein, results in the distorted deposition of cellulose microfibrils, which, in turn, leads to a defect in cell elongation. But the reports from Wasteneys’ lab led to serious questioning of the consensus model on microtubule regulation of microfibril orientation. Wasteneys (2004) and Himmelspach et al. (2003) reported in their studies that organized cortical microtubules were not essential for maintaining or reestablishing transversely oriented cellulose microfibrils in expanding cells. Their results demonstrated that cellulose microfibrils could recover in well-ordered, transverse patterns without a preexisting transverse microfibril template, lending further support to the idea that cellulose microfibril orientation was largely generated by self-assembly mechanisms that had little reliance on cortical microtubule organization or preexisting microfibrils. Wasteneys proposed that microtubule-dependent microfibril length may contribute, along with microfibril orientation, to the mechanical properties of the plant cell wall. Recently, expression of a functional yellow fluorescent protein fusion to cellulose synthase (CESA) in transgenic Arabidopsis plants and spinning-disc confocal microscopy revealed that CESA complexes moved at constant rates in linear tracks that were aligned and coincident with cortical microtubules (Paredez et al., 2006). Furthermore, whether defects in cellulose synthesis affect the orientation of microtubules remains uncertain. All cellulose synthases described to date have a number of conserved structural features. It was discovered that phosphorylation of the catalytic subunit plays a key role in regulation of cellulose synthesis (Taylor, 2005). The amino terminus of the CESA protein contains a domain that bears some resemblance to a zinc finger or LIM transcription factor. Within this domain is a strongly conserved CxxC motif beginning 10 to 40 amino acids from the amino terminus: Cx2Cx12FxACx2PxCx2Cx-Ex5Gx3Cx2C (C 5 Cys, F 5 phenylananine, etc; X 5 any amino acid; Richmond, 2000). This aspect, together with other recent developments in the cellulose synthase field, was recently well reviewed by Somerville (2006). In this study, we report that the AtCESA2 knockout mutant (ecotype Landsberg erecta [Ler]), atcesa2, has severe defects in cell wall formation and microtubule orientation, resulting in abnormal plant growth and development. The function of the zinc finger-like domain of the AtCESA2 gene was also analyzed. Plant Physiol. Vol. 143, 2007

The AtCESA2 Gene Affects Cell Expansion

Mutant plants were almost sterile, although they could produce some seeds when grown directly under fluent air. Root hairs were of normal length and morphology, indicating that atcesa2 did not affect tipgrowing cells. This was confirmed by the absence of atcesa2 expression in root hairs and root tips (Fig. 3B). The atcesa2 phenotype was not due to brassinosteroid, auxin, or gibberellin deficiency because the addition of each hormone separately to the growth medium could not rescue the dwarf phenotype (data not shown). Genetic Analysis and Molecular Identification of the atcesa2 Mutant

Figure 1. Phenotype of the atcesa2 mutant. A and B, Ten-day-old seedlings grown on Murashige and Skoog medium. A, Wild type. B, atcesa2 (Ler background). C, Six-day-old dark-grown wild-type (left) and atcesa2 (right) seedlings (two seedlings each) grown in the dark. D and E, Seven-day-old wild-type (D) and atcesa2 (E) seedlings grown vertically on Murashige and Skoog medium in 1.5% agar. F and G, Adult plants grown for 6 weeks (F) and 8 weeks (G) in a greenhouse. F, Wild type (left), atcesa2 (Ler background; bottom right), details of atcesa2 (top right). G, Mature wild type (left) and atcesa2 (right) grown under fluent air conditions. Scale bars 5 2 mm (A–E), 1 cm (F and G).

RESULTS General Phenotypic Characterization of the atcesa2 Mutant

A screen for RING finger mutants led to the identification of atcesa2. The mutant was isolated from an activator (Ac)/dissociation (Ds)-mutagenized population in the Ler background generated as described by Sundaresan et al. (1995). When grown in light, the atcesa2 seedling phenotype deviated from wild type in that the leaves were rolled out and shrunken (Fig. 1, A and B) and the hypocotyls were much shorter than those of the wild type (Fig. 1, D and E). Hypocotyls of the mutant grown in the dark for 6 d displayed a similar phenotype, but they were 2 times shorter than those of the wild type (Fig. 1C). They also had a reduced apical hook and opening of the cotyledon compared to wild-type seedlings. Mature greenhouse-grown homozygous plants of atcesa2 showed a strong dwarf phenotype with small inflorescence stems with shorter internodes (Fig. 1, F and G). Plant Physiol. Vol. 143, 2007

The atcesa2 mutant was isolated from a pool of Ac/ Ds-transformed lines in the Ler background. The F1 progeny of the mutant backcrossed to wild type were grown on Murashige and Skoog medium and scored for segregation: None of the F1 plants had the mutant phenotype. In the F2 generation, by analysis of a total of 3,069 seedlings germinated on a Murashige and Skoog plate, 2,307 seedlings showed a wild-type phenotype, whereas 762 seedlings appeared as mutant phenotypes, which segregated approximately in a 3:1 ratio. No mild phenotype was observed. The segregation ratio of the original heterozygotes was also 3:1 (data not shown). The mutant phenotype was also linked to the kanamycin-resistant phenotype, which had no segregation in the next generation. Together, these data suggest that atcesa2 was a recessive mutation in a single Mendelian locus. Thermal asymmetric interlaced-PCR analysis demonstrated that the Ds was inserted 143 bp upstream of the start codon of the AtCESA2 gene (Fig. 2A). Sequencing of the PCR products using primers specific to the Ds and to the AtCESA2 gene further confirmed that the Ds was linked to the AtCESA2 gene and no chromosome fragment deletion was found in the insertion region (data not shown). By checking genome sequence information of the insertion region, no coding sequence was found within 3 kb upstream from the insertion site. Northern-blot analysis (Fig. 2B), using the first specific region (236–676 bp from the ATG) of the AtCESA2 gene as the probe, showed no expression of AtCESA2 in the atcesa2 homomutant and reduced expression in the heteromutant, confirming that the AtCESA2 gene was knocked out in the atcesa2 mutant. Analysis of the cellulose content of the atcesa2 mutant revealed that it had about 61% of wild-type cellulose levels in leaves. However, the cellulose content in roots did not differ from wild type (Fig. 2D). To further confirm that the atcesa2 mutant phenotype was due to mutation of the AtCESA2 gene, the 7.7-kb genomic sequence, including a 2-kb fragment upstream of ATG and a 500-bp fragment downstream of the stop codon of the AtCESA2 wild-type gene, was amplified and cloned into pCambia 1301, which was transformed into atcesa2 homozygotes by Agrobacterium-mediated methods. Twelve T1 progeny with hygromycin resistance were obtained, which all exhibited the wild-type 215

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Correlation of the AtCESA2 Gene Expression Profile with the atcesa2 Mutant Phenotype

Northern blotting showed that AtCESA2 was expressed in all tissues, such as roots, stem, flowers, and cauline leaves (Burn et al., 2002). To investigate the AtCESA2 gene expression profile in detail, a 2-kb fragment upstream of the ATG of the AtCESA2 gene was amplified by PCR and fused to the b-glucuronidase (GUS) reporter gene by inserting the promoter sequence into pCambia 1300-221 and transformed into Ler wild-type plants. The transgenic plant was then grown in soil and 30 stable GUS transgenic lines were obtained. GUS staining results indicated that the AtCESA2 gene was expressed in all tissues in 10-dold seedlings; low levels of expression were even detected in root hairs. High levels of expression were also detected in mature leaves, stamen filaments, and siliques (Fig. 3). By analysis, the AtCESA2 gene expression profiles in the public Web site Genevestigator indicated that the expression pattern of AtCESA2 (At4G39350) is similar to promoter-GUS results where high expression was resumed in the hypocotyl, stem, petiole, and leaves (https://www.genevestigator. ethz.ch/at/index). The expression pattern of AtCESA2 was only correlated with the phenotype of the atcesa2 mutants in the aerial part. The reasons are discussed further in the ‘‘Discussion’’ section. The atCESA2 Mutation Causes Abnormal Cell Expansion

The size of a plant organ is determined by the rate of cell division and cell expansion and/or elongation.

Figure 2. AtCESA2 was knocked out in the atcesa2 mutant. A, The Ds element was inserted 143 bp upstream of the ATG of AtCESA2. B, Northern blot detected AtCESA2 gene expression in the atcesa2 mutant, as well as in wild type using an AtCESA2 gene-specific probe. C, Complementation of the atcesa2 mutant. a, atcesa2 mutant. b, atcesa2 mutant (pCambia 1301) transgenic vector control. c, atcesa2 mutant (pCambia 1301-AtCESA2). d, Wild-type control. D, Cellulose content of the atcesa2 mutant and wild-type control. Cellulose content was reduced in the mutant leaves rather than in wild-type leaves, but there was almost no change in roots. [See online article for color version of this figure.]

phenotype (Fig. 2C). T2 progeny were grown on Murashige and Skoog medium plus kanamycin or hygromycin. On the kanamycin-containing medium, all seedlings had resistance, including wild-type and mutant seedlings, whereas on the hygromycin-containing medium, resistant seedlings all exhibited the wildtype phenotype. Northern-blot analysis revealed that rescue of the phenotype was due to expression of the AtCESA2 gene by the complementary construct in transgenic plants (Fig. 2C). 216

Figure 3. Histochemical analysis of GUS expression in transgenic Arabidopsis plants carrying the AtCESA2 promoterTGUS fusion constructs. A, Seedling showing GUS expression in the cotyledon and vascular bundles as well as roots. B, GUS expression in roots. C, Details of GUS expression in root tip region. D, No clear GUS expression was detected in root hairs. E, Flower showing GUS expression in the stamen and stigma. F, Leaf of mature plant showing GUS expression in the same part of vascular bundles. G, Silique showing GUS expression at abscission zone. Plant Physiol. Vol. 143, 2007


The hypocotyls and stamen filaments of the atcesa2 were clearly shorter than the wild type (Fig. 4, A, B, E, F, and I). To investigate whether this shorter phenotype was caused by defects in cell expansion or cell division, the epidermal cell number and length of hypocotyls and stamen filaments were measured and counted. Statistical analysis of these data (Fig. 4J) demonstrated that the number of epidermal cells in hypocotyls and stamen filaments of the mutants did not differ significantly from those of wild type. Furthermore, the epidermal cells of the mutant hypocotyls and stamen filaments were much shorter than the wild type (Fig. 4, C, D, G, and H), suggesting that the altered phenotypes were caused by an alteration in cell size rather than cell number. This is consistent with the notion that the mutant phenotype was due to a defective cellulose synthase gene. Knocking out this gene may cause a defect in cellulose synthesis and lead to a disruption in the deposition of microfibrils, which results in alterations in cell wall extensibility and cell expansion. The atCESA2 Mutation Causes Cell Wall Defects Mainly in Leaves

The above results demonstrated that the atcesa2 phenotypes were caused by the loss of function of the AtCESA2 gene. The AtCESA2 gene is a member of the CESA family, which is responsible for cell wall synthesis. To investigate whether the cell wall defects occurred in atcesa2, we first studied transverse sections of the roots, leaves, and stems of wild-type and mutant plants. The overall anatomical features of the mutant roots and vascular bundles were unaltered and the diametric size of the roots of the mutant was almost the same as the wild type (Fig. 5, A and B). However, the stems of atcesa2 showed a decrease in girth as a result of decreased cell diameter. The whole vascular bundle of the mutant was unaltered relative to wild type (Fig. 5, G and H). The most affected parts of the mutant were the leaves. The overall anatomy of the mutant leaves differed from wild type, curling much more than wild-type leaves (Fig. 5, C and D). The cell morphology of mutant leaves, especially the cell walls, was also significantly altered (Fig. 5F) compared to the wild type (Fig. 5E). To further investigate the cell and cell wall aberrances, the ultrastructure of mutant and wild-type leaves was observed by transmission electron microscopy. The palisade cells of wild-type leaves were evenly arrayed perpendicularly to the epidermal cells (Fig. 6A), but the palisade cells in the mutant were arranged parallel to the epidermal cells with the cell shape altered (Fig. 6C). These results were consistent with the outward rolling of atcesa2 leaves. Additionally, the cell walls of the wild-type cells were smooth and complete (Fig. 6B), whereas those of the mutant were shrunken (Fig. 6D) and incomplete (Fig. 6E). No obvious differences in callose levels were observed between the cell walls of wild-type and mutant cells Plant Physiol. Vol. 143, 2007

(data not shown). Quantitative analysis revealed that pectin content in the aerial part of mature atcesa2 mutant plants was higher than that in the wild-type control (Fig. 6G). The atCESA2 Mutation Affects Microtuble Orientation

Microtuble orientation is a key factor controlling cell expansion and elongation. Himmelspach et al. (2003) observed that cellulose synthesis inhibitor 2,6-dichlorobenzonitrile treatment partially disorders cortical microtubules, supporting the possibility that microtubule orientation is influenced by cellulose microfibril organization. To observe whether microtubule orientation was affected in atcesa2, we generated a transgenic line expressing MAP4 and GFP fusion proteins (MAP4-GFP) by crossing a MAP4-GFP transgenic line (Mathur and Chua, 2000) with the atcesa2 line. As its name suggests, MAP4 decorates the microtubule and its fusion to GFP allows the visualization of microtubule organization and orientation. Using confocal microscopy, we observed microtubule orientation in the epidermal cells of hypocotyls (Fig. 7, A–D) and petioles (Fig. 7, E–H) in mutant and wild-type plants. The orientation of microtubules to the direction of cell expansion was observed in epidermal cells of wildtype hypocotyls. In contrast, disordered microtubules were observed in mutant hypocotyls. Similar results were found in the epidermal cells of petioles. In roots, there was no obvious difference of microtubule orientation observed between wild type and mutant (Fig. 7, I–L). Consideration that overexpressing MAP4-GFP fusion proteins produce a developmental phenotype that may affect normal microtubule orientation, microtubule orientations were further tested in hypocotyls by immunofluorescence microscopy with a mouse anti-b-tubulin monoclonal antibody (see details in ‘‘Materials and Methods’’). Disordered microtubules were observed in mutant hypocotyls compared to those in wild-type hypocotyls (Fig. 7, M and N). Together, our experiments indicated that microtubule orientation was affected in the atcesa2 mutant aboveground parts. AtCESA2 Interacts with Itself through the N-Terminal Zinc-Binding Domain

The Arabidopsis AtCESA zinc finger shows high similarity to RING finger domains, a small zinc-binding domain found in many functionally distinct proteins (Freemont, 2000). Recent works have also suggested that more than one type of CESA may be required to make up a functional rosette (Taylor et al., 2000; Scheible et al., 2001). All CESA genes encode a zinc finger domain at the N-terminal region of the protein (Fig. 8A). Kurek et al. (2002) confirmed that GhCESA1 could form homodimers as well as heterodimers with GhCESA2 through the zinc-binding domain. To investigate whether AtCESA2 can homodimerize and to 217

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Figure 4. The atcesa2 mutation caused defective cell expansion and elongation. Scanning electron microscopy (SEM; A–H) of 6-d-old wild-type (A) and atcesa2 (B) seedlings. C and D, Epidermal cells of hypocotyl of wild type (C) and atcesa2 (D). E and F, Flower of wild type (E) and atcesa2 (F). G and H, Epidermal cells of stamen filament of wild type (G) and atcesa2 (H). Scale bars 5 500 mm (A and B, E, and F), 100 mm (C and D, G and H). Bar chart of the length (I) and epidermal cell number (J) of hypocotyl and stamen filament of wild type and atcesa2. Data represent mean 6 SE for measurements of 10 seedlings and stamen filaments. Hypocotyls and stamen filaments of wild type are longer than the atcesa2 mutants (I), whereas the epidermal cell numbers of hypocotyls and stamen filaments of wild type are almost the same as atcesa2 mutants (J).

identify the domain responsible, we performed a yeast (Saccharomyces cerevisiae) two-hybrid assay with different truncations of AtCESA2 protein. Our results demonstrated that AtCESA2 could homodimerize and that the zinc finger domain of AtCESA2 was important for the interaction (Fig. 8B), consistent with previously reported results (Kurek et al., 2002). In contrast, no E3 ligase activity was detected in the in vitro assay for detecting the E3 ligase activity of RING finger proteins (data not shown). DISCUSSION

The atcesa2 mutant was obtained during our screening of insertion lines for RING finger-containing mutants. Genetic analysis, thermal asymmetric interlaced-PCR, northern-blot, cellulose content assay, and complementation experiments demonstrated that the mutation caused a loss of expression of the AtCESA2 gene. Reduced expression of the AtCESA2 gene by an antisense method showed no obvious phenotype, except reductions in stem length under certain growth conditions (Burn et al., 2002). One possibility is that reduced expression of the AtCESA2 gene by antisense gave rise to a mild phenotype compared to the complete knockout of the gene. These results highlight the differences between the two reverse-genetics approaches, reduction of RNA level (gene silencing) versus gene deletion. Scheible and Pauly (2004) have reported in their review that an atCesA2 insertion line has no obvious phenotypes in plant growth and development. But 218

recently, the same author found some phenotypes with more insertion lines and it is believed that different CesA2 insertion mutants probably have very different effects on plant growth (W.R. Scheible, personal communication). We also analyzed two T-DNA knockout lines, SALK_091570 and SALK_125303, of the AtCesA2 gene in the Col background from the Arabidopsis Biological Resource Center. No obvious phenotypes were detected at all stages of plant growth and regeneration (data not shown). Disruption or mutation of the same gene in different backgrounds (ecotypes) of Arabidopsis also could give different phenotypes or no phenotypes due to gene redundancy in different ecotypes (Maloof et al., 2001; Yang and Hua, 2004; Werner et al., 2005). We used the Ler background in our study and previous reports were all from the Col background (Burn et al., 2002; Scheible and Pauly, 2004). Diversity and Overlap of CESA Functions

The AtCESA2 gene is one of about 10 currently known members of the cellulose synthase family. Each member may be one catalytic subunit of the rosettes responsible for cellulose synthesis. Several questions regarding CESA function remain to be answered. Are the various CESA isoforms functionally unique or do some isoforms overlap in function? Available data suggest that significant functional specialization has occurred, including differences in gene expression, regulation, and possibly catalytic function. Various CESA expression analyses have been conducted in Plant Physiol. Vol. 143, 2007


code the catalytic subunit of the mixed-linkage glucan synthase or callose synthase. Expression studies of CESA genes facilitate functional studies using loss-of-function mutants because only genes that are expressed at the same stage or in the same tissue need to be examined for potentially redundant functions. Using AtCESA-promoterTGUS fusions, CESA2 and CESA5, like CESA1, CESA3, and CESA6, were shown to be expressed at the sites of primary wall synthesis (Scheible and Pauly, 2004). Our findings demonstrated that AtCESA2 was expressed both in the aerial and root part of the seedling, that the cellulose content was nearly the same in mutant and wild-type roots, and that no obvious phenotypes were observed in the root compared to the aerial part, suggesting that some genes expressed in the root have redundant functions in atCESA2.

Figure 5. Toluidine blue-stained sections of Arabidopsis 14-d-old seedling. Roots (A and B), leaves (C–F), and vascular bundles (G and H). Wild type (A, C, E, and G) and atcesa2 mutant (B, D, F, and H). Scale bars 5 25 mm (A and B), 50 mm (C and D), 5 mm (E and F), and 50 mm (G and H). [See online article for color version of this figure.]

both monocot and dicot species. Arabidopsis CESA family members have different expression levels, particularly when multiple CESA genes are expressed in the same cell type. AtCESA1, AtCESA3, and AtCESA6 all exhibit very similar expression patterns, being expressed in cells undergoing expansion in tissues such as roots and hypocotyls (Arioli et al., 1998; Fagard et al., 2000; Scheible et al., 2001). AtCESA4, AtCESA7, and AtCESA8 also have overlapping expression patterns in xylem cells (Holland et al., 2000; Taylor et al., 2000). Phylogenetic analysis suggested that the former group was involved in primary cell wall formation, whereas the latter group participated in secondary cell wall formation. In this study, AtCESA2 was expressed in cells undergoing expansion in tissues such as roots, hypocotyls, and, especially, in leaves. Disruption of the gene caused defects in primary cell wall formation. Phylogenetic analysis also demonstrated that AtCESA2 is highly homologous to AtCESA9 and AtCESA5. Current data suggest that some CESA isoforms are potentially responsible for initiation or elongation of the recently identified sterol b-glucoside primer within different cell types, including cells undergoing primary and secondary cell wall biosyntheses (Read and Bacic, 2002). Different CESA isoforms may also play distinct roles within the rosette and there is some circumstantial evidence that the CESA gene may enPlant Physiol. Vol. 143, 2007

Figure 6. Ultrastructure analysis of leaves by transmission electron microscopy. Leaves were taken from 14-d-old light-grown seedlings. Wild type (A and B) and atcesa2 mutant (C–F). Black arrowheads, normal cell walls; white arrowheads, defective cell walls. Scale bars 5 5 mm (A and C) and 1 mm (B, D, E, and F). Quantitative analysis of pectin content (G). Each analysis was conducted by three repeat reactions (n 5 3). 219

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Figure 7. Visualization of cortical microtubules in hypocotyls (A–D), petioles (E–H), and roots (I–L) of epidermal cells by confocal microscopy. A, C, E, G, I, and K, View of whole hypocotyls (A and C), petioles (E and G), and root (I and K) with MAP4-GFP expression. A, E, and I, Wild type. C, G, and K, atcesa2 mutants. B, D, F, H, J, and L, View of microtubule orientation in epidermal cells of hypocotyls (B and D), petioles (F and H), and roots ( J and L) with MAP4-GFP expression. B, F, and J, Wild type. D, H, and L, atcesa2 mutants. M and N, View of microtubule orientation in epidermal cells of wild-type (M) and mutant (N) hypocotyls growing in the dark for 3 d with immunofluorescence microscopy with a mouse anti-b-tubulin monoclonal antibody. Scale bars 5 100 mm (A, C, E, and G), 50 mm (I and K), 10 mm (B, D, F, H, M, and N), and 5 mm ( J and E).

Components and Formation of CESA Rosettes

Cellulose synthase proteins are components of CESA complexes (rosettes) and catalyze glucan polymerization. Little is understood about rosette assembly, including how CESAs interact with each other or with other components within the complexes. How many types of CESA are present in one rosette and are components of all rosettes the same in different tissues and organs? AtCESA7 and AtCESA8 were the first to 220

be found to interact with each other in vivo (Taylor et al., 2000). Taylor et al. (2003) reported that IRX5, IRX3, and IRX1 were coexpressed within cells and all three proteins interacted in detergent-soluble extracts, suggesting that they were required for cellulose synthesis in secondary cell walls. The conserved regions at the N terminus of plant CESA proteins contain two putative zinc fingers that show high homology to the RING finger motif and are thought to facilitate protein-protein interactions. Plant Physiol. Vol. 143, 2007


Figure 8. The dimerizing domain of AtCESA2. A, Alignment of the CESA members containing zinc finger domains. Black triangle indicates the conserved Cys within the zinc finger domain. B, Interactions between AtCESA2 and itself in yeast. Schematic representation of the AtCESA2 N-terminal fragment, zinc finger domain, and whole protein (top). Yeast transformed with plasmids containing the GAL-4 binding domain (pGBKT7) and the GAL-4 activating domain (pGADT7); fusion proteins were grown on synthetic complete medium minus Leu and Trp (His1) and minus Leu, Trp, and His, plus 5 mM 3-amino 1,2,4-triazole (5 mM 3-AT). b-Galactosidase activity was assayed according to the standard protocol provided by CLONTECH. [See online article for color version of this figure.]

Kurek et al. (2002) reported that cotton fiber cellulose synthase catalytic subunits dimerized via the N-terminal zinc finger domains. Our yeast two-hybrid experiment suggested that AtCESA2 could homodimerize via the zinc finger domains. Taken together, it is likely that CESA zinc finger domains are essential for rosette formation. Cell Wall Composition and Extensibility

Plant cell expansion is driven by internal turgor pressure and restricted by the ability of the cell wall to extend under this pressure. The extensibility of cell walls depends on at least two related events, including the composition of the wall and how its components are bound to one another and the modification of existing wall structures (Darley et al., 2001). Initially, primary cell wall material must be synthesized in a form that is competent to undergo extension. The primary cell wall is essentially a composite structure comprising a framework of cellulose microfibrils embedded in a matrix of other polymers. Most of the polymers are pectin or cross-linking glycan polysaccharides. Alterations in cellulose synthesis can affect Plant Physiol. Vol. 143, 2007

cell wall extensibility, which impacts on plant growth. Studies of the effects of cellulose biosynthesis inhibitors revealed that cultured plant cells can survive in the absence of cellulose (Shedletzky et al., 1990). The walls of these cells are effectively devoid of cellulose, and much of this absence appears to be compensated by increased levels of pectin. How pectin contributes to cell wall extensibility remains unclear. However, as pectin is mainly found in primary cell walls, it very likely fulfills an important function during wall extension. The high pectin concentration can provide a high hydration potential to the cell wall, which in turn may endow the wall with compressive strength. Cellulose microfibrils, which are deposited in a transverse direction along the axis of elongation, have been proposed to control cellular morphogenesis. Disruption of the normal deposition of cellulose microfibrils by mutation of a cellulose synthase gene in the temperaturesensitive rsw1 mutant correlated with the alteration of cell elongation (Sugimoto et al., 2001). In atcesa2, the AtCESA2 gene was completely knocked out, which resulted in reduced cellulose synthesis and increased levels of pectin in the cell wall (Fig. 6), altering the cell shape and extensibility. 221

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Microtubule Orientation and Cell Expansion

In the microtubule parallelism hypothesis, transverse deposition of cellulose microfibrils along the axes of elongating cells is proposed to be controlled by the transversely oriented cortical microtubules lying beneath the plasma membrane (Giddings and Staehelin, 1991; Baskin, 2001). Alternatively, the coalignment of microtubules and cellulose microfibrils may be determined by cellular geometry (Emons and Mulder, 1998). Disruption in cell growth induced by pharmacological drugs can influence cell morphology, resulting in simultaneous alteration in the orientations of microtubules and cellulose mirofibrils (Emons et al., 1992). Furthermore, Fisher and Cyr (1998) suggested that cellulose biosynthesis is required for microtubule organization and stabilization and the cellulose microfibrils and cortical microtubules become coordinated during cell wall formation (Fisher and Cyr, 1998). Our studies demonstrated that cellulose synthesis and microfibril deposition were disrupted in the primary cell wall in the atcesa2 mutant. As such, we conclude that cellulose synthesis and microfibril deposition can affect microtubule orientation. It is noteworthy that the requirements in microtubule organization for cell wall formation may vary with primary and secondary cell walls (Eckardt, 2003). In the irx3 and irx5 null mutants, microtubules in the developing xylem appeared normal, despite the lack of association of the microtubules with the CESA complexes and the lack of cellulose deposition in the secondary wall (Gardiner et al., 2003). Together, we conclude that the cell shape and cell wall extensibility was altered in the atcesa2 mutant due to disruption of microfibril deposition, which affected microtubule orientation and eventually caused abnormal cell expansion and elongation.

of these lines was analyzed on Murashige and Skoog, Murashige and Skoog plus kanamycin, and Murashige and Skoog plus hygromycin media.

Northern Blot RNA was isolated using the Qiagen RNeasy kit and 10 mg of total RNA were loaded per lane. To detect the AtCESA2 transcript, blots were probed with an N-terminal fragment (with HindIII digestion [236–676 bp from ATG]) labeled with [a-32P]dCTP using a Ready-primed labeling kit (Amersham International).

Cellulose Content Measurement Seedlings were grown on one-half-strength Murashige and Skoog for 2 weeks. Some of the seedlings were transferred into B5 liquid medium and cultured in suspension at 22°C; root parts were harvested after 3 weeks. The other set of seedlings were potted in soil and placed in a growth chamber at 22°C with 75% humidity under a 16-h light/8-h dark photoperiod; aerial parts were harvested after 3 weeks. Samples were ground into fine powder in liquid nitrogen, washed in phosphate buffer (50 mM, pH 7.2) three times. After centrifugation, pellets were extracted three times in 70% ethanol at 70°C for 1 h and dried at 80°C overnight. Dried materials were extracted with an acetic/ nitric acid reagent at 100°C, followed by digestion with 67% sulfuric acid. Cellulose content was determined by the phenol-sulfuric acid method (Dubois et al., 1956), using cellulose as a standard.

GUS Staining For histochemical localization of GUS activity, seedlings were incubated in a solution containing 50 mM sodium phosphate buffer, pH 7.0, 5 mM K3Fe(CN)6, 0.1% Triton X-100, and 1 mM 5-bromo-4-chloro-3-indolyl-b-glucuronic acid and incubated at 37°C for several hours as described (Xie et al., 2000).

Preparation of Histological Samples Seedlings were collected and fixed in 2.5% glutaraldehyde in 50 mM NaPO4 buffer, pH 7.4, for 2.5 h at 4°C. Subsequently, they were washed four times for 15 min in 50 mM NaPO4 buffer, pH 7.4, then twice for 15 min in water. The samples were then dehydrated in increasing concentrations of ethanol and embedded in historesin using a kit according to the manufacturer’s instructions (Leica). Sections (2-mm-thick) were prepared and stained with 0.05% toluidine blue O in sodium citrate buffer, pH 4.4. Samples were eventually mounted in water for microscopic observation.

Callose Staining and Pectin Quantitative Analysis MATERIALS AND METHODS Plant Growth and Transformation Seeds of Arabidopsis (Arabidopsis thaliana; ecotype Ler) were surface sterilized with 30% bleach and 0.001% Triton X-100 for 10 min and washed three times with sterile water. Sterile seeds were suspended in 0.2% agarose and plated on Murashige and Skoog medium or Murashige and Skoog plus 1.5% Suc. Plants were vernalized in darkness for 2 d at 4°C and then transferred to a tissue culture room under white fluorescent light (45 mE m22 s21, 16-h light/8-h dark cycle) at 22°C. After 2 to 3 weeks, seedlings were potted in soil and placed in a growth chamber at 22°C with 75% humidity under a 16-h light/8-h dark photoperiod. Transgenic plants were generated by Agrobacteriummediated methods (Bent and Clough, 1998).

Complementation The whole genomic sequence, including a 2-kb fragment upstream of the ATG and a 500-bp fragment downstream of the stop codon of the AtCESA2 wild-type gene, was amplified by the Expand Long Template PCR system (Roche Diagnostics GmbH; catalog no. 1 681 842). The forward and reverse primers were 5#-TTTCCAgATTTgTTggTggAgCCTTgTAgg-3# and 5#-TTAATAAACCATgCCTgATTCAATggAACCgAg-3#, respectively. The PCR product was inserted into the pGEM-T easy vector at the AatII and PstI restriction sites. The insert was subsequently cloned into pCAMBIA 1301 at the SmaI and PstI restriction sites. After confirming sequence accuracy, the construct was transformed into atcesa2 and 12 T2 progeny were obtained. The segregation

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Callose was stained in a solution of 0.005% aniline blue in 50% ethanol and observed under confocal microscopy (Zeiss LSM510) at 405 nm with a bandpass 420- to 480-nm emission filter. For pectin quantitative analysis, samples were harvested from the aerial part of 3-week-old soil-grown plants. Measurement of pectin content was performed according to Zhou et al. (2006). Briefly, samples were first incubated for 1 h in a boiling water bath. After cooling to room temperature, samples were filtrated by crucible. One hundred milliliters of 0.1 M NaOH were added to one-tenth of filtrate and then incubated for 30 min. Then 50-mL 1 M acetic acid was added, following 5-min incubation, 50 mL 1 M CaCl2 was supplied, and the incubation was continued for 1 h. After boiling for 5 min, the mixture was filtered and the pellet was washed. The pelleted pectin (Ca salt) on the filter was dried at 105°C and measured. Each experiment was repeated by three reactions.

Scanning Electron Microscopy Fresh tissue samples were placed on a specimen stub and secured with Tissue-Tek (Sakura Finetake Europe B.V.) before cryofixing in liquid nitrogen slush. Samples were then sputter coated with gold and viewed in high vacuum mode on a cryostage maintained at 190°C (CT1500 Cryo transfer system; Oxford Instruments) using a JEOL-JSM 5310-LV scanning electron microscope.

Transmission Electron Microscopy Leaf samples were fixed in 4% glutaraldehyde in 100-mM cacodylate buffer for 3 h and then postfixed with 2% osmium tetroxide in 100 mM sodium

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cacodylate buffer for 1 h at 4°C. Samples were then dehydrated through a series of 30%, 50%, 70%, 90%, and 100% ethanol and finally immersed in propylene oxide prior to infiltration with Spurr’s resin (Spurr, 1969). Samples were embedded in 100% Spurr’s resin and polymerized at 65°C overnight. Ultrathin sections were cut on a Jung Reichert ultramicrotome and examined with a transmission electron microscope (JEM1010; JEOL) at 100 kV.

Microtubule Observation by Confocal Microscopy Three-day-old wild-type and atcesa2 seedlings with Map4-GFP expression of wild type and mutants were mounted in water. Samples were imaged using a dip-in long working distance 633 (NA0.9) water-immersion lens (Zeiss). GFP fluorescence was excited with the 488-nm line of an argon/krypton laser. Emission was detected through a 510-nm beam splitter and a 515 to 565 bandpass filter.

Immunofluorescent Staining For visualization of microtubules within cells of the root elongation zone and the middle region of the hypocotyl, we used the protocol for fixing and immunofluorescent staining of 5-d-old seedling roots and etiolated hypocotyls as described (Sugimoto et al., 2000), with modifications. Whole seedlings were fixed in 4% (v/v) paraformaldehyde and 0.1% (v/v) glutaraldehyde made up in PEMT buffer (100 mM PIPES, 4 mM EGTA, 4 mM MgSO4, 0.05% [v/v] Triton X-100) for 1 h. Cell wall digestion was performed with 1% (w/v) pectinase, 1% cellulose (w/v), and 0.4 M mannitol dissolved in PEM (100 mM PIPES, 4 mM EGTA, 4 mM MgSO4) buffer for 1 h at 37°C. A mouse anti-b-tubulin monoclonal antibody (Sigma T-4026; 1:800 dilution) was used as a primary antibody and rabbit anti-mouse IgG/fluoresceinisothiocyanate (DingGuo; 1:500 dilution) was used as a secondary antibody. Fluorescent images were collected with a Zeiss Axioplan2 LSM 510 META confocal laser-scanning microscope.

Protein Interaction Assay by the Yeast Two-Hybrid System The AtCESA2 zinc finger domain, the N-terminal AtCESA2 fragment, and full-length proteins were inserted into pGADT7 and pGBKT7. The yeast (Saccharomyces cerevisiae) strain HF7c (MATa ura3-52, his3-200, ade2-101, lys2801, trp1-901, leu2-3,112, gal4-542, gal80-538 LYS2TGAL1UAS-GAL1TATAHIS3, URA3TGAL4, 17 mers [33]-CyC1TATA-LacZ), containing the two reporter genes LacZ and HIS3, was used. Both pGBKT7 and pGADT7 bearing different lengths of AtCESA2 were transformed into yeast cells. Sequence data from this article can be found in the GenBank/EMBL data libraries under accession number At4g39350.

ACKNOWLEDGMENTS We thank Dr. Venkatesan Sundaresan for Ac/Ds insertion lines, the Arabidopsis Biological Resource Center, Ohio State University, for providing the T-DNA insertion lines, and Ms. Yang Sun Chan for excellent scanning electron microscopy/transmission electron microscopy support. Received August 16, 2006; accepted October 31, 2006; published November 3, 2006.

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