Characterization of Cellulose Synthesis in Plant Cells

Maleki, Samaneh Sadat; Mohammadi, Kourosh; Ji, Kong-shu

doi:https://doi.org/10.1155/2016/8641373

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Review Article | Open Access

Volume 2016 | Article ID 8641373 | https://doi.org/10.1155/2016/8641373

Characterization of Cellulose Synthesis in Plant Cells

Samaneh Sadat Maleki,¹Kourosh Mohammadi,¹and Kong-shu Ji¹

Academic Editor: Charles Hocart

Received06 Jan 2016

Revised02 Apr 2016

Accepted28 Apr 2016

Published25 May 2016

Abstract

Cellulose is the most significant structural component of plant cell wall. Cellulose, polysaccharide containing repeated unbranched β (1-4) D-glucose units, is synthesized at the plasma membrane by the cellulose synthase complex (CSC) from bacteria to plants. The CSC is involved in biosynthesis of cellulose microfibrils containing 18 cellulose synthase (CesA) proteins. Macrofibrils can be formed with side by side arrangement of microfibrils. In addition, beside CesA, various proteins like the KORRIGAN, sucrose synthase, cytoskeletal components, and COBRA-like proteins have been involved in cellulose biosynthesis. Understanding the mechanisms of cellulose biosynthesis is of great importance not only for improving wood production in economically important forest trees to mankind but also for plant development. This review article covers the current knowledge about the cellulose biosynthesis-related gene family.

1. Introduction

Plant cell wall is required not only for the structure to determine the actual shape of cells and functional properties to control inner and outer cellular communication but also for overall growth and tree expansion. Cellulose is the most significant component of plant cell walls. Due to the enormous economic beneficial aspect of tree cellulose for paper, lumber, pulp, and industrial products, understanding the mechanism of cellulose biosynthesis is valuable research objective.

2. Molecular Features of Cellulose

Cellulose is a polymer of glucose (C6H12O6), rotation 180° of one glucose molecule in relation to the next glucose to form β (1-4)-linked residues, called cellobiose (C12H22O11). Cellulose chain is made up of a repeating unit of cellobiose [1]. Cellulose is synthesized by cellulose synthase enzymes (CesAs) [2]. Cell walls consist of three types of layers. The middle lamella is formed during cell division as a first layer. The cell wall is microfibril-based; the primary cell wall (Pcw) is formed after the middle lamella but the secondary cell wall (Scw) is formed after cell enlargement completion. The secondary wall is often layered to S1, S2, and S3 (outer, middle, and inner layers, resp.) which vary in microfibrils orientation. S2 is the thickest layer with steep helices of microfibrils, while S1 and S3 are arranged in flat fibrillar slope [3]. All the cell wall layers consist of microfibrillar and matrix phases. The microfibrils have a crystalline core and less crystalline outer side but the matrix is noncrystalline phase that contains pectins and hemicelluloses, lignin, and other polymers [4]. The six crystalline polymorphs of cellulose, namely, I, II, , , , and , have been known. Cellulose I and cellulose II are the most common forms of cellulose, whereas others are not yet known to exist freely in nature. Cellulose I or native cellulose has two suballomorphs Iα and Iβ. Both of them are found in higher plants [1]. The intramolecular hydrogen bonds are responsible for rigidity and stability of cellulose. Cellulose microfibrils I that have strong intra- and interchain hydrogen bonds which make the cellulose structures I and II run parallel and antiparallel to the long axis, respectively [5]. The degree of polymerization (DP) is different in Pcw and Scw which represent the monomer units in each cellulose chain which shows low DP, 2000–6000 for the primary cell wall, and high DP, 14000 in secondary cell wall [6].

3. History and Chromosomal Location of CesA

The CesA was initially identified in the Gram-negative bacterium Acetobacter xylinus [7]. In 1996, the plant CesA proteins were first recognized in ESTs isolated from cotton based on sequence homogeny to bacterial CesA [8]. CesA proteins have been localized to the plasma membrane. The Arabidopsis genome hosts a large family of cellulose biosynthesis genes, ten CesA genes (CesA1–10) and 29 Csl encoding cellulose synthase-like genes [9]. Expression analysis of Arabidopsis revealed that CesA genes are expressed in most plant organs with some differences at the tissue level [10].

4. Structural Features of CesA

Different range of size CesA genes and amino acids in Arabidopsis is from 3.5 to 5.5 kb and 985 to 1088, respectively [11]. CesA is relevant to membrane-bound glycosyltransferase family 2 (GT-2) enzyme [12]. Glycosyltransferase is situated on the cytoplasmic domain between two sets of transmembrane domain. Eight TMDs are identified in plant CesA protein, TM1-2 towards N-terminus and TM3–8 towards C-terminus. These TMDs are suggested to form a pore across the inner membrane to embed cellulose chain secretion through the cell wall [13, 14]. In the conserved region, D,D,D,QXXRW, the first two ASP(D) residues help coordinate the uridine diphosphate, while the third D is the catalytic base and the QXXRW motif (Q is glutamine, R is arginine, W is tryptophan, and X is any amino acid) [10] (Figure 1) helps form a binding site for the terminal disaccharide of the growing glucan chain [15]. There is specific extra protein structure only in plants that consist of two (A, B) regions in GT domain: (A) highly conserved domains which have been named plant-specific conserved region (P-CR) and (B) specific conserved region and hypervariable region (HVR) which have been called class specific region (CSR) together [16] which sit in the mid part of GT (Figure 1). Sethaphong et al. [12] noted the CesA hexamer and tetramers assemblies are possible role of plant-specific subdomains and the CSR has highlighted role for dimers and trimmers assembly.

Figure 1

Model for structure of CesA proteins (right) shows a conserved zinc finger domain (ZN) and a hyper variable region (HVRΙ) near N-ter of TM1-2 and short C ter of TM3–8 and central hypervariable region (HVRΙΙ), plant-specific conserved region (P-CR), and class specific region (CSR); position of the processive glycosyltransferase motif D,D,D,QXXRW. Plant’s cellulose biosynthesis (left). The plasma membrane-associated sucrose synthase (SuSy) channels uridine diphosphate-glucose (UDP-G) substrate to form rosette and glucan chain formation, the UDP formed can be recycled back to SuSy, and Korrigan cellulase (Kor) has been involved in monitoring of cellulose synthesis. Microtubules (MT) play role to regulate CesA proteins trafficking [2, 54, 72].

A comparison between bacterial and plant CesAs shows that a short N-terminal region contains 12 amino acids and 208 amino acids long in C-terminus of BcsA, while plant CesAs have short 17–21 amino acids long in C-terminus and 160–260 amino acids long in N-terminal region with zinc finger; 50 amino acids long is settled in this region [16]. Kurek et al. in 2002 proposed that the N-terminal zinc-binding domain partakes in protein interaction for rosette assembly in cotton; two GhCesA1 and GhCesA2 zinc finger domains were shown to interact with each other to regulate CesA assembly through dimerization via intermolecular disulfide bonds under oxidative conditions [14].

5. The CSC Models

As early as 1972, cellulose synthase complexes were visualized by electron microscopy [17]. Since CSCs were attached to the end of microfibrils and were observed in three rows ordered particles in alga Oocystis apiculata, so Brown Jr. and Montezinos [18] called them the linear terminal complex (TC) for the first time. Four years later, not only the connection between the rosette and the TC to synthesize microfibrils in higher plants was described by freeze fracture for first time, but also a different form of cellulose synthesizing sites was found as hexameric rosette TCs. Measurements suggest that rosette is in diameter, containing six particles with each of them having six cellulose synthase polypeptides to polymerize six glucan chains [19]. Cellulose synthase utilizes sitosterol glucoside which is synthesized by UDP-glucose as substrate to synthesis microfibril [20]. Ding and Himmel [21] proposed the cellulose microfibril model containing 36 glucose chains which is composed of both crystalline and noncrystalline chains by using atomic force microscopy (AFM) of direct visualization of the maize stem. In studies of the primary cell wall, except 36-chain model, two alternative models which apply to structures of CSCs containing the 24-chain and 18-chain models have been described. The 24-chain (eight three-chain sheets) models, three CesA polypeptides, make a particle and eight particles make a rosette formation with conformational disorder surfaces rather than packing disorders. The 18-chain twinned microfibrils (six three-chain sheets) models described the rosette with six particles of three cellulose synthase polypeptides. As the cross-sectional area of 36 chains microfibrils were evidently larger than the primary wall microfibril, so this model was a poor fit to experimental data. But 18-chain microfibrils model showed good fits to experimental data [22–24]. The rosette TCs are believed to be assembled by multiple CesA in the Golgi and then transported to plasma membrane in active form to cellulose synthesis by cytoplasmic vesicles [2, 25] which are termed SmaCCs (Small CesA Compartments) or MASCs (Microtubule Associated Cellulose Synthase Compartments), but then these small compartments have an operation in recycling CesA proteins from the plasma membrane [16, 26]. Multiple glucan chains can be synthesized by multiple cellulose synthase genes in each TC [27]. The rosette takes part in both glucan chain polymerization and crystallization [13].

6. Bacterial and Arabidopsis Genes That Encode Proteins of the Cellulose Synthase Complex

Bacterial cellulose synthase (Bcs) complex operon encodes four genes, BcsA, B, C, and Z. The Bcs A, B, and C activity is required for the synthesis and translocation of the polysaccharide; BcsZ encodes a cellulase to cellulose production [28–30]. The recent study on cellulose synthase operon genes (bcsABZC) of Cronobacter species confirmed the particular role of bcsA and bcsB mutants in cellulose production and showed involvement in biofilm formation and cell aggregation [31]. The cellulose synthesis in plants takes place in the context of rosettes more than TCs row [2] containing multiple steps, β-1,4-glucan chain initiation, elongation, and termination. Omadjela et al. [30] described that there is no requirement for a primer for chain initiation and there is no need to add other energy sources to the assembly of individual chains into a higher-order structure for cellulose synthesis in plants because the polymerization of UDP-glucose (DP range: 200–300) provides energy for growing cellulose chain through the membrane pore. Expression of different genes in Arabidopsis demonstrated that AtCesA4, AtCesA7, and AtCesA8 are required to make the secondary cell wall, while AtCesA1, AtCesA3, and AtCesA6 take part in cellulose biosynthesis of primary cell wall [32, 33]. AtCesA2, AtCesA5, and AtCesA9 appear to be partially redundant with AtCesA6 [27]. No precise role has been assigned to AtCesA10 [16]. First experimental evidence for the CesA function involvement in cellulose synthesis came from Arabidopsis mutant radial analyses swelling 1 (rsw1) which exhibited deficiency in cellulose content and number of rosette TCs at higher temperature in the primary cell wall due to changing valine amino acid to alanine in AtCesA1 [34] and irregular xylem (irx) which represented defect in secondary cell wall formation in tissue xylem; in fact the irx1 (CesA8), irx3 (CesA7), and irx5 (CesA4) mutants exhibit collapsed or irregular xylem cells and reduced mechanical strength in fiber cells due to changing aspartic acid to asparagine, displacing tryptophan with a stop codon at position 859 and stop codon at position 263 replacing a glutamine, respectively [32, 35, 36] (see Table 1); observation of collapsed xylem cells resulting from mutation in Exigua (exi) genes, which were mapped to three cellulose synthase subunits CesA4, CesA7, and CesA8, leads to blocking water transport and reduced cell enlargement subsequently enhanced tolerance to osmotic stress which affects secondary cell wall deposition [37]; Mur10 mutants altered primary cell wall carbohydrate composition in response to secondary cell wall defects due to a mutation in CesA7 locus [38]. Mutations in CesA8 (lew2) enhance drought stress and accumulate ABA in secondary cell wall [39]. Stork et al. [40] reported the definite role for CesA9 in Arabidopsis seed coats; CesA9 mutant seeds contained 25% cellulose reduction and no changes in other tissues. Carroll et al. [41] in an analysis of transgenic lines in Arabidopsis demonstrated that CesA7 and CesA1 can rescue sort of deficiency in SCW biosynthesis in cesa3 and in PCW biosynthesis in cesa8ko mutant, respectively. IXR1 and IXR2 mutant alleles are point mutations in the CesA3 and CesA6 genes that confer isoxaben resistance [42, 43]; CesA1aegeus and CesA3ixr1-2 mutants showed considerably reduced crystallinity and increased CesA velocity in the PM and resistance to quinoxyphen was conferred by CesA1aegeus A903V [44]; conversely, reduction in CSC velocity was observed in anisotropy1 D604N missense mutation in CesA1 [45]; prc1-1 null CesA6 mutant caused cellulose deficiency resulting in reduced cell elongation which was examined in Arabidopsis [33]. Surprisingly, both cellulose content reduction and constitutive stress response are due to accumulation of JA and ethylene in the cev1 mutant in CesA3 [46]. Lignification in nonlignified cells in eli1-1 and eli1-2 (CesA3) inhibits cellulose synthesis that invokes overproduction of jasmonate and ethylene [47]. Reduction in primary and secondary cell wall thickness and cellulose content are affected by a missense mutation which occurred in fragile fiber 5 (fra5), a dominant mutant of AtCesA7, while neither cell wall thickness nor cellulose content is affected in fra6 mutant form of AtCesA8 [48].

7. Non-CesA Genes Involved in Cellulose Biosynthesis

Interestingly, apart from CesA proteins, the KORRIGAN, sucrose synthase (SuSy), microtubules and actin cytoskeletons, and COBRA-like proteins are involved in cellulose biosynthesis indirectly (Figure 1). It is thought that the membrane-bound endo-1,4-β-D-glucanase (KOR) has editing and monitoring role in conversion of glucan chain to release of newly synthesized cellulose microfibrils and eliminating defective glucan chains from the microfibril assembly [49, 50]. In theory, mutation in KOR may alter crystallization of the cellulose microfibrils [51]. Liebminger et al. [52] mentioned that activation of A. thaliana KOR1 depends on utilization of eight N-glycosylation sites in the extracellular domain. PtrKOR1 and GhKOR1 get involved in secondary cell wall cellulose formation in Populus tremuloides, endosperm cellularization, and embryo development in cotton (Gossypium hirsutum) through RNAi suppression [53].

The strategic role of plasma membrane-associated sucrose synthase (P-SUSY) of developing cotton fibers (Gossypium hirsutum) in channeling UDP-glucose to cellulose synthase from sucrose was first illustrated in 1995 [54]; the UDP formed from UDP-G can be recycled back to SUSY (Figure 1). Mutation in SUS (1–4) in Arabidopsis shows less SuSy activity in all cells not in phloem and amazingly no cellulose deficiency was found, while the sus5/sus6 mutants showed callose reduction in plates screening, but the plant growth was severely affected by mutant invertase (INV), recommending that catalysis of sucrose may need cytosolic invertase rather than SuSy [55]. But after two years Baroja-Fernández et al. [56] evidenced the possible role of sucrose synthase activity in cellulose biosynthesis under optimum pH 7.0 conditions.

The relation between the cytoskeleton (cortical microtubules and actin) and CSC localization and movement has been the topic in the majority of studies. MTs are another key player in plant cell morphogenesis. Gardiner et al. [57] showed the colocalization of all three AtCesA4 (IRX5), AtCesA7 (IRX3), and AtCesA8 (IRX1) proteins with cortical microtubule bands in older developing xylem vessel with GFP. But actin microfilaments localize with the CesA proteins to regions of cell wall thickening. Intracellular trafficking and plasma membrane localization of the CSC during secondary cell wall formation were supported by live-cell imaging of fluorescently labeled proteins fusion [58–63]. The alignment hypothesis describes that microtubules can control the alignment of cellulose microfibril deposition in the plasma membrane [59, 61]; however, the examination results of polymerization and depolymerization of MTs with short treatment by taxol and oryzalin, respectively, do not agree with the alignment hypothesis because no changes were observed in cellulose microfibril orientation [60]. Li et al. [61] investigated the role of CesA interactive protein 1 (CSI1) as a linker protein in association between CesA complexes and cortical microtubules in vivo. Based on result of Zhong et al. [62] the FRA1, kinesin-like microtubule binding protein, mutant made alteration deposition of cellulose microfibrils in fiber cell walls in Arabidopsis. Intracellular trafficking of CSC by actin filaments has been suggested by Wightman and Turner [63] for controlling delivery of CSC to the PM to maintain proper patterned deposition of the Scw. Newly, dynamic coordination between AF and MT was investigated using dual labeled probe in interphase plant cell [64].

A role of COBRA (cob) mutant in regulating the orientation of cell expansion was associated with a screen for Arabidopsis with defect expanded roots for the first time in 1993 [65]. Short and swollen roots resulted from mutations in the COB. It encodes a putative GPI-anchored protein that is necessary for oriented cell expansion in Arabidopsis. COBRA causes the cell elongation and reduction in crystalline cellulose content. Also it has been identified in association with deposition of cellulose microfibrils in the root tissue [66, 67]. COBRA-like gene family members COBL2, COBL6, COBL9, COBL10, and COBL11 are required for oriented crystalline cellulose deposition during seed development, root hairs, and pollen tube elongation in PCW, respectively, while COBL4 was identified during the formation of the vascular system in xylem cells [68, 69]. As irx6 is a member of the COBRA gene family (COBL4), it also regulates continuous growth cell and manifests decreased crystalline cellulose content in root cell walls [70, 71].

8. Conclusion

Despite the numerous studies and progress towards understanding the mechanism of cellulose biosynthesis in higher plants, numbers of questions are still pending. How many proteins are necessary for cellulose synthesis process? How can every plant regulate the synthesis of cellulose? How is the CesA position in cellulose synthesis complex? How does a plant cell control the crystallization of microfibrils? Investigation of cellulose structure and key genes engaged in cellulose synthesis can be important because of widespread utilization of cellulose product in daily life.

Abbreviations

CesA:	Cellulose synthase catalytic subunit
Csl:	Cellulose synthase-like
Pcw:	Primary cell wall
Scw:	Secondary cell wall
GT:	Glycosyltransferase
TMD:	Transmembrane domain
HVR:	Hypervariable region
TC:	Terminal complex
CMF:	Cellulose microfibril
Kor:	Korrigan cellulase
MT:	Microtubule
SuSy:	Sucrose synthase
UDP-G:	Uridine diphosphate-glucose
GFP:	Green fluorescent protein.

Competing Interests

The authors declare that there are no competing interests regarding the publication of this paper.

Acknowledgments

This work was supported by the Priority Academic Program Development of Jiangsu Higher Educational Institutions, China.

References

R. A. Festucci-Buselli, W. C. Otoni, and C. P. Joshi, “Structure, organization, and functions of cellulose synthase complexes in higher plants,” Brazilian Journal of Plant Physiology, vol. 19, no. 1, pp. 1–13, 2007.
View at: Publisher Site | Google Scholar
M. S. Doblin, I. Kurek, D. Jacob-Wilk, and D. P. Delmer, “Cellulose biosynthesis in plants: from genes to rosettes,” Plant and Cell Physiology, vol. 43, no. 12, pp. 1407–1420, 2002.
View at: Publisher Site | Google Scholar
C. B. Beck, An Introduction to Plant Structure and Development. Plant Anatomy for the Twenty-First Century, Cambridge University Press, Cambridge, UK, 2nd edition, 2010.
C. Brett and K. Waldron, “Physiology and biochemistry of plant cell walls,” in Topics in Plant Physiology, M. Black and J. Chapman, Eds., vol. 2, pp. 4–57, Unwin Hyman, London, UK, 1990.
View at: Google Scholar
I. Simon, L. Glasser, H. A. Scheraga, and R. S. T. John Manley, “Structure of cellulose. 2. Low-energy crystalline arrangements,” Macromolecules, vol. 21, no. 4, pp. 990–998, 1988.
View at: Google Scholar
J. M. Stewart, D. Oosterhuis, J. J. Heitholt, and J. R. Mauney, Eds., Physiology of Cotton, Springer, 2009.
H. C. Wong, A. L. Fear, R. D. Calhoon et al., “Genetic organization of the cellulose synthase operon in Acetobacter xylinum,” Proceedings of the National Academy of Sciences of the United States of America, vol. 87, no. 20, pp. 8130–8134, 1990.
View at: Publisher Site | Google Scholar
J. R. Pear, Y. Kawagoe, W. E. Schreckengost, D. P. Delmer, and D. M. Stalker, “Higher plants contain homologs of the bacterial celA genes encoding the catalytic subunit of cellulose synthase,” Proceedings of the National Academy of Sciences of the United States of America, vol. 93, no. 22, pp. 12637–12642, 1996.
View at: Publisher Site | Google Scholar
N. C. Carpita, “Update on mechanisms of plant cell wall biosynthesis: how plants make cellulose and other (1→4)-β-D-Glycans,” Plant Physiology, vol. 155, no. 1, pp. 171–184, 2011.
View at: Publisher Site | Google Scholar
R. E. Williamson, J. E. Burn, and C. H. Hocart, “Towards the mechanism of cellulose synthesis,” Trends in Plant Science, vol. 7, no. 10, pp. 461–467, 2002.
View at: Publisher Site | Google Scholar
T. Richmond, “Higher plant cellulose synthases,” Genome Biology, vol. 1, no. 4, pp. 3001-1–3001-6, 2000.
View at: Google Scholar
L. Sethaphong, C. H. Haigler, J. D. Kubicki et al., “Tertiary model of a plant cellulose synthase,” Proceedings of the National Academy of Sciences of the United States of America, vol. 110, no. 18, pp. 7512–7517, 2013.
View at: Publisher Site | Google Scholar
S. Li, L. Bashline, L. Lei, and Y. Gu, “Cellulose Synthesis and Its Regulation,” The Arabidopsis Book, vol. 12, article e0169, 2014.
View at: Publisher Site | Google Scholar
I. Kurek, Y. Kawagoe, D. Jacob-Wilk, M. Doblin, and D. Delmer, “Dimerization of cotton fiber cellulose synthase catalytic subunits occurs via oxidation of the zinc-binding domains,” Proceedings of the National Academy of Sciences of the United States of America, vol. 99, no. 17, pp. 11109–11114, 2002.
View at: Publisher Site | Google Scholar
J. L. W. Morgan, J. Strumillo, and J. Zimmer, “Crystallographic snapshot of cellulose synthesis and membrane translocation,” Nature, vol. 493, no. 7431, pp. 181–186, 2013.
View at: Publisher Site | Google Scholar
M. Kumar and S. Turner, “Plant cellulose synthesis: CESA proteins crossing kingdoms,” Phytochemistry, vol. 112, no. 1, pp. 91–99, 2015.
View at: Publisher Site | Google Scholar
S. Kimura, W. Laosinchai, T. Itoh, X. Cui, C. R. Linder, and R. M. Brown Jr., “Immunogold labeling of rosette terminal cellulose-synthesizing complexes in the vascular plant Vigna angularis,” Plant Cell, vol. 11, no. 11, pp. 2075–2085, 1999.
View at: Publisher Site | Google Scholar
R. M. Brown Jr. and D. Montezinos, “Cellulose microfibrils: visualization of biosynthetic and orienting complexes in association with the plasma membrane,” Proceedings of the National Academy of Sciences of the United States of America, vol. 73, no. 1, pp. 143–147, 1976.
View at: Publisher Site | Google Scholar
S. C. Mueller and R. M. Brown, “Evidence for an intra membrane component associated with a cellulose microfibril synthesizing complex in higher plants,” The Journal of Cell Biology, vol. 84, no. 2, pp. 315–326, 1980.
View at: Google Scholar
L. Peng, Y. Kawagoe, P. Hogan, and D. Delmer, “Sitosterol-β-glucoside as primer for cellulose synthesis in plants,” Science, vol. 295, no. 5552, pp. 147–150, 2002.
View at: Publisher Site | Google Scholar
S.-Y. Ding and M. E. Himmel, “The maize primary cell wall microfibril: a new model derived from direct visualization,” Journal of Agricultural and Food Chemistry, vol. 54, no. 3, pp. 597–606, 2006.
View at: Publisher Site | Google Scholar
R. H. Newman, S. J. Hill, and P. J. Harris, “Wide-angle X-ray scattering and solid-state nuclear magnetic resonance data combined to test models for cellulose microfibrils in mung bean cell walls,” Plant Physiology, vol. 163, no. 4, pp. 1558–1567, 2013.
View at: Publisher Site | Google Scholar
L. H. Thomas, V. Trevor Forsyth, A. Šturcová et al., “Structure of cellulose microfibrils in primary cell walls from collenchyma,” Plant Physiology, vol. 161, no. 1, pp. 465–476, 2013.
View at: Publisher Site | Google Scholar
V. G. Vandavasi, D. K. Putnam, Q. Zhang et al., “A structural study of CESA1 catalytic domain of arabidopsis cellulose synthesis complex: evidence for CESA trimers,” Plant Physiology, vol. 170, no. 1, pp. 123–135, 2015.
View at: Publisher Site | Google Scholar
U. Neumann, F. Brandizzi, and C. Hawes, “Protein transport in plant cells: in and out of the Golgi,” Annals of Botany, vol. 92, no. 2, pp. 167–180, 2003.
View at: Publisher Site | Google Scholar
E. F. Crowell, V. Bischoff, T. Desprez et al., “Pausing of Golgi bodies on microtubules regulates secretion of cellulose synthase complexes in Arabidopsis,” Plant Cell, vol. 21, no. 4, pp. 1141–1154, 2009.
View at: Publisher Site | Google Scholar
S. Persson, A. Paredez, A. Carroll et al., “Genetic evidence for three unique components in primary cell-wall cellulose synthase complexes in Arabidopsis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 39, pp. 15566–15571, 2007.
View at: Publisher Site | Google Scholar
C.-L. Keiski, M. Harwich, S. Jain et al., “AlgK Is a TPR-Containing protein and the periplasmic component of a novel exopolysaccharide secretin,” Structure, vol. 18, no. 2, pp. 265–273, 2010.
View at: Publisher Site | Google Scholar
O. Mazur and J. Zimmer, “Apo- and cellopentaose-bound structures of the bacterial cellulose synthase subunit BcsZ,” The Journal of Biological Chemistry, vol. 286, no. 20, pp. 17601–17606, 2011.
View at: Publisher Site | Google Scholar
O. Omadjela, A. Narahari, J. Strumillo et al., “BcsA and BcsB form the catalytically active core of bacterial cellulose synthase sufficient for in vitro cellulose synthesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 110, no. 44, pp. 17856–17861, 2013.
View at: Publisher Site | Google Scholar
L. Hu, C. J. Grim, A. A. Franco et al., “Analysis of the cellulose synthase operon genes, bcsA, bcsB, and bcsC in Cronobacter species: prevalence among species and their roles in biofilm formation and cell-cell aggregation,” Food Microbiology, vol. 52, pp. 97–105, 2015.
View at: Publisher Site | Google Scholar
N. G. Taylor, R. M. Howells, A. K. Huttly, K. Vickers, and S. R. Turner, “Interactions among three distinct CesA proteins essential for cellulose synthesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 100, no. 3, pp. 1450–1455, 2003.
View at: Publisher Site | Google Scholar
M. Fagard, T. Desnos, T. Desprez et al., “Procuste1 encodes a cellulose synthase required for normal cell elongation specifically in roots and dark-grown hypocotyls of arabidopsis,” Plant Cell, vol. 12, no. 12, pp. 2409–2423, 2000.
View at: Publisher Site | Google Scholar
T. Arioli, L. C. Peng, A. S. Betzner et al., “Molecular analysis of cellulose biosynthesis in Arabidopsis,” Science, vol. 279, no. 5351, pp. 717–720, 1998.
View at: Publisher Site | Google Scholar
N. G. Taylor, W.-R. Scheible, S. Cutler, C. R. Somerville, and S. R. Turner, “The irregular xylem3 locus of Arabidopsisencodes a cellulose synthase required for secondary cell wall synthesis,” Plant Cell, vol. 11, no. 5, pp. 769–779, 1999.
View at: Publisher Site | Google Scholar
N. G. Taylor, S. Laurie, and S. R. Turner, “Multiple cellulose synthase catalytic subunits are required for cellulose synthesis in Arabidopsis,” Plant Cell, vol. 12, no. 12, pp. 2529–2539, 2000.
View at: Publisher Site | Google Scholar
S. Rubio-Díaz, J. M. Pérez-Pérez, R. González-Bayón et al., “Cell expansion-mediated organ growth is affected by mutations in three EXIGUA genes,” PloS one, vol. 7, no. 5, Article ID e36500, 2012.
View at: Publisher Site | Google Scholar
S. Bosca, C. J. Barton, N. G. Taylor et al., “Interactions between MUR10/CesA7-dependent secondary cellulose biosynthesis and primary cell wall structure,” Plant Physiology, vol. 142, no. 4, pp. 1353–1363, 2006.
View at: Publisher Site | Google Scholar
Z. Chen, X. Hong, H. Zhang et al., “Disruption of the cellulose synthase gene, AtCesA8/IRX1, enhances drought and osmotic stress tolerance in Arabidopsis,” The Plant Journal, vol. 43, no. 2, pp. 273–283, 2005.
View at: Publisher Site | Google Scholar
J. Stork, D. Harris, J. Griffiths et al., “CELLULOSE SYNTHASE9 serves a nonredundant role in secondary cell wall synthesis in Arabidopsis epidermal testa cells,” Plant Physiology, vol. 153, no. 2, pp. 580–589, 2010.
View at: Publisher Site | Google Scholar
A. Carroll, N. Mansoori, S. Li et al., “Complexes with mixed primary and secondary cellulose synthases are functional in Arabidopsis plants,” Plant Physiology, vol. 160, no. 2, pp. 726–737, 2012.
View at: Publisher Site | Google Scholar
W.-R. Scheible, R. Eshed, T. Richmond, D. Delmer, and C. Somerville, “Modifications of cellulose synthase confer resistance to isoxaben and thiazolidinone herbicides in Arabidopsis Ixr1 mutants,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 18, pp. 10079–10084, 2001.
View at: Publisher Site | Google Scholar
T. Desprez, S. Vernhettes, M. Fagard et al., “Resistance against herbicide isoxaben and cellulose deficiency caused by distinct mutations in same cellulose synthase isoform CESA6,” Plant Physiology, vol. 128, no. 2, pp. 482–490, 2002.
View at: Publisher Site | Google Scholar
D. M. Harris, K. Corbin, T. Wang et al., “Cellulose microfibril crystallinity is reduced by mutating C-terminal transmembrane region residues CESA1 A903V and CESA3 T942I of cellulose synthase,” Proceedings of the National Academy of Sciences of the United States of America, vol. 109, no. 11, pp. 4098–4103, 2012.
View at: Publisher Site | Google Scholar
M. Fujita, R. Himmelspach, J. Ward et al., “The anisotropy1 D604N mutation in the Arabidopsis cellulose synthase1 catalytic domain reduces cell wall crystallinity and the velocity of cellulose synthase complexes,” Plant Physiology, vol. 162, no. 1, pp. 74–85, 2013.
View at: Publisher Site | Google Scholar
C. Ellis, I. Karafyllidis, C. Wasternack, and J. G. Turner, “The Arabidopsis mutant cev1 links cell wall signaling to jasmonate and ethylene responses,” Plant Cell, vol. 14, no. 7, pp. 1557–1566, 2002.
View at: Publisher Site | Google Scholar
A. Caño-Delgado, S. Penfield, C. Smith, M. Catley, and M. Bevan, “Reduced cellulose synthesis invokes lignification and defense responses in Arabidopsis thaliana,” Plant Journal, vol. 34, no. 3, pp. 351–362, 2003.
View at: Publisher Site | Google Scholar
R. Zhong, W. H. Morrison III, G. D. Freshour, M. G. Hahn, and Z.-H. Ye, “Expression of a mutant form of cellulose synthase AtCesA7 causes dominant negative effect on cellulose biosynthesis,” Plant Physiology, vol. 132, no. 2, pp. 786–795, 2003.
View at: Publisher Site | Google Scholar
F. Nicol, I. His, A. Jauneau, S. Vernhettes, H. Canut, and H. Höfte, “A plasma membrane-bound putative endo-1,4-β-D-glucanase is required for normal wall assembly and cell elongation in Arabidopsis,” The EMBO Journal, vol. 17, no. 19, pp. 5563–5576, 1998.
View at: Publisher Site | Google Scholar
M. Mølhøj, S. Pagant, and H. Höfte, “Towards understanding the role of membrane-bound endo-β-1,4-glucanases in cellulose biosynthesis,” Plant and Cell Physiology, vol. 43, no. 12, pp. 1399–1406, 2002.
View at: Publisher Site | Google Scholar
J. Takahashi, U. J. Rudsander, M. Hedenström et al., “KORRIGAN1 and its aspen homolog PttCel9A1 decrease cellulose crystallinity in Arabidopsis stems,” Plant and Cell Physiology, vol. 50, no. 6, pp. 1099–1115, 2009.
View at: Publisher Site | Google Scholar
E. Liebminger, J. Grass, F. Altmann, L. Mach, and R. Strasser, “Characterizing the link between glycosylation state and enzymatic activity of the endo-β1,4-glucanase KORRIGAN1 from Arabidopsis thaliana,” The Journal of Biological Chemistry, vol. 288, no. 31, pp. 22270–22280, 2013.
View at: Publisher Site | Google Scholar
L. Yu, H. Chen, J. Sun, and L. Li, “PtrKOR1 is required for secondary cell wall cellulose biosynthesis in populus,” Tree Physiology, vol. 34, no. 11, pp. 1289–1300, 2014.
View at: Publisher Site | Google Scholar
Y. Amor, C. H. Haigler, S. Johnson, M. Wainscott, and D. P. Delmer, “A membrane-associated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants,” Proceedings of the National Academy of Sciences of the United States of America, vol. 92, no. 20, pp. 9353–9357, 1995.
View at: Publisher Site | Google Scholar
D. H. P. Barratt, P. Derbyshire, K. Findlay et al., “Normal growth of Arabidopsis requires cytosolic invertase but not sucrose synthase,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 31, pp. 13124–13129, 2009.
View at: Publisher Site | Google Scholar
E. Baroja-Fernández, F. J. Muñoz, J. Li et al., “Sucrose synthase activity in the sus1/sus2/sus3/sus4 Arabidopsis mutant is sufficient to support normal cellulose and starch production,” Proceedings of the National Academy of Sciences of the United States of America, vol. 109, no. 1, pp. 321–326, 2012.
View at: Publisher Site | Google Scholar
J. C. Gardiner, N. G. Taylor, and S. R. Turner, “Control of cellulose synthase complex localization in developing xylem,” Plant Cell, vol. 15, no. 8, pp. 1740–1748, 2003.
View at: Publisher Site | Google Scholar
X. Shang, Q. Chai, Q. Zhang et al., “Down-regulation of the cotton endo-1,4-β-glucanase gene KOR1 disrupts endosperm cellularization, delays embryo development, and reduces early seedling vigour,” Journal of Experimental Botany, vol. 66, no. 11, pp. 3071–3083, 2015.
View at: Publisher Site | Google Scholar
P. B. Green, “Mechanism for plant cellular morphogenesis,” Science, vol. 138, no. 3548, pp. 1404–1405, 1962.
View at: Publisher Site | Google Scholar
K. Sugimoto, R. Himmelspach, R. E. Williamson, and G. O. Wasteneys, “Mutation or drug-dependent microtubule disruption causes radial swelling without altering parallel cellulose microfibril deposition in Arabidopsis root cells,” Plant Cell, vol. 15, no. 6, pp. 1414–1429, 2003.
View at: Publisher Site | Google Scholar
S. Li, L. Leia, C. R. Somervill, and Y. Gua, “CSI1 links microtubules and cellulose synthase complexes,” Proceedings of the National Academy of Sciences, vol. 109, no. 1, pp. 185–190, 2011.
View at: Google Scholar
R. Zhong, D. H. Burk, W. H. Morrison III, and Z.-H. Ye, “A kinesin-like protein is essential for oriented deposition of cellulose microfibrils and cell wall strength,” Plant Cell, vol. 14, no. 12, pp. 3101–3117, 2002.
View at: Publisher Site | Google Scholar
R. Wightman and S. R. Turner, “The roles of the cytoskeleton during cellulose deposition at the secondary cell wall,” The Plant Journal, vol. 54, no. 5, pp. 794–805, 2008.
View at: Publisher Site | Google Scholar
A. Sampathkumar, J. J. Lindeboom, S. Debolt et al., “Live cell imaging reveals structural associations between the actin and microtubule cytoskeleton in arabidopsis,” Plant Cell, vol. 23, no. 6, pp. 2302–2313, 2011.
View at: Publisher Site | Google Scholar
P. N. Benfey, P. J. Linstead, K. Roberts, J. W. Schiefelbein, M.-T. Hauser, and R. A. Aeschbacher, “Root development in Arabidopsis: four mutants with dramatically altered root morphogenesis,” Development, vol. 119, no. 1, pp. 57–70, 1993.
View at: Google Scholar
G. Schindelman, A. Morikami, J. Jung et al., “COBRA encodes a putative GPI-anchored protein, which is polarly localized and necessary for oriented cell expansion in arabidopsis,” Genes and Development, vol. 15, no. 9, pp. 1115–1127, 2001.
View at: Publisher Site | Google Scholar
F. Roudier, A. G. Fernandez, M. Fujita et al., “COBRA, an Arabidopsis extracellular glycosyl-phosphatidyl inositol-anchored protein, specifically controls highly anisotropic expansion through its involvement in cellulose microfibril orientation,” Plant Cell, vol. 17, no. 6, pp. 1749–1763, 2005.
View at: Publisher Site | Google Scholar
S. Li, F.-R. Ge, M. Xu et al., “Arabidopsis COBRA-LIKE 10, a GPI-anchored protein, mediates directional growth of pollen tubes,” Plant Journal, vol. 74, no. 3, pp. 486–497, 2013.
View at: Publisher Site | Google Scholar
D. Ben-Tov, Y. Abraham, S. H. Stav et al., “COBRA-LIKE2, a member of the glycosylphosphatidylinositol-anchored COBRA-LIKE family, plays a role in cellulose deposition in Arabidopsis seed coat mucilage secretory cells,” Plant Physiology, vol. 167, no. 3, pp. 711–724, 2015.
View at: Publisher Site | Google Scholar
D. M. Brown, L. A. H. Zeef, J. Ellis, R. Goodacre, and S. R. Turner, “Identification of novel genes in Arabidopsis involved in secondary cell wall formation using expression profiling and reverse genetics,” Plant Cell, vol. 17, no. 8, pp. 2281–2295, 2005.
View at: Publisher Site | Google Scholar
P. Derbyshire, D. Ménard, P. Green et al., “Proteomic analysis of microtubule interacting proteins over the course of xylem tracheary element formation in Arabidopsis,” The Plant Cell, vol. 27, no. 10, pp. 2709–2726, 2015.
View at: Publisher Site | Google Scholar
A. Sampathkumar, R. Gutierrez, H. E. McFarlane et al., “Patterning and lifetime of plasma membrane-localized cellulose synthase is dependent on actin organization in Arabidopsis interphase cells,” Plant Physiology, vol. 162, no. 2, pp. 675–688, 2013.
View at: Publisher Site | Google Scholar

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Copyright © 2016 Samaneh Sadat Maleki et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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