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Ruiqin Zhong and Zheng-Hua Ye* Department of Plant Biology, University of Georgia, Athens, GA 30602, USA *Corresponding author: E-mail, [email protected]; Fax, +1-706-542-1805. (Received September 18, 2014; Accepted September 29, 2014) Secondary walls are mainly composed of cellulose, hemicelluloses (xylan and glucomannan) and lignin, and are deposited in some specialized cells, such as tracheary elements, fibers and other sclerenchymatous cells. Secondary walls provide strength to these cells, which lend mechanical support and protection to the plant body and, in the case of tracheary elements, enable them to function as conduits for transporting water. Formation of secondary walls is a complex process that requires the co-ordinated expression of secondary wall biosynthetic genes, biosynthesis and targeted secretion of secondary wall components, and patterned deposition and assembly of secondary walls. Here, we provide a comprehensive review of genes involved in secondary wall biosynthesis and deposition. Most of the genes involved in the biosynthesis of secondary wall components, including cellulose, xylan, glucomannan and lignin, have been identified and their co-ordinated activation has been shown to be mediated by a transcriptional network encompassing the secondary wall NAC and MYB master switches and their downstream transcription factors. It has been demonstrated that cortical microtubules and microtubule-associated proteins play important roles in the targeted secretion of cellulose synthase complexes, the oriented deposition of cellulose microfibrils and the patterned deposition of secondary walls. Further investigation of many secondary wall-associated genes with unknown functions will provide new insights into the mechanisms controlling the formation of secondary walls that constitute the bulk of plant biomass. Keywords: Cellulose Cortical microtubule Lignin Microtubule-associated protein Secondary wall Secondary wall NAC Xylan. Abbreviations: Ara, arabinose; BC, brittle culm; CesA, cellulose synthase A; COBL, cobra-like; CSI, cellulose synthaseinteracting protein; CslA, cellulose synthase-like A; CTL, chitinase-like protein; DUF, domain of unknown function; ESK, eskimo; FRA, fragile fiber; GalA, galacturonic acid; GlcA, glucuronic acid; GT, glycosyltransferase; GUX, glucuronic acid substitution of xylan; GXM, glucuronoxylan methyltransferase; IRX, irregular xylem; KOR, korrigan; LAC, laccase; MAP, microtubule-associated protein; MeGlcA, methylglucuronic acid; MIDD, microtubule depletion domain; NST, NAC secondary wall thickening-promoting factor; PRX, peroxidase; Rha, rhamnose; RWA, reduced wall acetylation; SMRE, secondary wall MYB-responsive element; SNBE, secondary wall NAC-binding element; SND, secondary wall-associated NAC domain protein; SWN, secondary wall NAC; VND, vascularrelated NAC-domain; XAT, xylan arabinosyltransferase; XAX, xylosyl arabinosyl substitution of xylan; Xyl, xylose. Introduction Secondary walls are mainly found in tracheary elements (tracheids in seedless vascular plants and gymnosperms and vessels in angiosperms) and fibers in the primary xylem and the secondary xylem (wood) (Fig. 1). They provide mechanical strength to these cell types, which serve as mechanical tissues to enable vascular plants to grow to a great height. Deposition of lignified secondary walls in tracheary elements not only reinforces these water conduits to resist the negative pressure generated during transpiration, but also renders them waterproof for efficient water transport. The ability of plants to deposit secondary walls in water-conducting elements was evolved during the Silurian period (about 430 million years ago), which is considered to be one of the pivotal steps for vascular plans to conquer the terrestrial habitats (Raven et al. 1999). Secondary wall-containing cell types are also present in tissues other than xylem for protection and structural support, such as phloem fibers in stems of many dicot plants, extraxylary fibers beneath the epidermis in grass stems, interfascicular fibers between the vascular bundles in Arabidopsis stems (Fig. 1A), and sclereids in pear fruits (also called stone cells) and seed coats of jojoba (Mauseth 1988). In addition, secondary wall deposition in anther endothecium is required for the generation of the tensile force necessary for stomium rupture to release pollen grains (Yang et al. 2007). Similarly, secondary wall deposition in extraxylary fibers of dehiscent seed pods is necessary for seed pod dehiscence leading to seed dispersal (Liljegren et al. 2000). The major components of secondary walls are cellulose, hemicelluloses (xylan and glucomannan) and lignin (Fig. 1D). Cellulose is the load-bearing unit, in which cellulose microfibrils cross-link with hemicelluloses to form the framework of secondary walls. Hemicelluloses, such as xylan, are required for the normal assembly and mechanical strength of secondary walls; reduction in xylan content causes a severe decrease in cellulose deposition, secondary wall thickening and mechanical strength Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140, Advance Access publication on 7 October 2014, available online at www.pcp.oxfordjournals.org ! The Author 2014. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: [email protected] Special Focus Issue – Review Secondary Cell Walls: Biosynthesis, Patterned Deposition and Transcriptional Regulation R. Zhong and Z.-H. Ye | Secondary cell walls Fig. 1 Deposition of secondary walls in xylem and fibers. (A) Cross-section of an Arabidopsis stem showing lignified secondary walls (red; stained with phloroglucinol-HCl) in vessels (ve), xylary fibers (xf), interfascicular fibers (if) and phloem fibers (pf). co, cortex. (B) Transmission electron micrograph showing three layers (S1, S2 and S3) of secondary walls in interfascicular fibers of an Arabidopsis stem. (C) Cellulose microfibrils in the S2 layer of a developing interfascicular fiber cell of an Arabidopsis stem showing their nearly parallel orientations to the cell elongation axis (double arrows). Scale bars in (A), (B) and (C) = 180, 1.3 and 0.28 mm, respectively. (D) Structural formula of secondary wall components, including cellulose, hemicelluloses (glucuronoxylan, glucuronoarabinoxylan and glucomannan) and monolignols. of vessels and fibers (Zhong et al. 2005b, Brown et al. 2007). The essential role of xylan in secondary wall assembly is in sharp contrast to primary wall xyloglucan, a complete loss of which has no discernible effect on primary wall assembly (Cavalier et al. 2008). Lignin, being hydrophobic and inert, impregnates the cellulose and hemicellulose network to provide additional mechanical strength, rigidity and hydrophobicity to secondary walls. The composition of secondary walls, i.e. the proportion of cellulose, hemicelluloses and lignin, may vary among different plant species and even in different secondary wall-containing cell types of the same species. For example, wood from a gymnosperm species (Pinus strobus) consists of 41% cellulose, 9% arabinoglucuronoxylan, 18% galactoglucomannan and 29% 196 lignin, whereas wood from an angiosperm species (Populus tremuloides) contains 48% cellulose, 24% glucuronoxylan, 3% glucomannan and 21% lignin (Timell 1967). Secondary wall composition may also change in response to different developmental and environmental stimuli. For example, compression wood, which forms on the lower side of a leaning gymnosperm stem, contains a higher lignin content and a lower cellulose content compared with normal wood, whereas tension wood, which forms on the upper side of a leaning angiosperm stem, has an additional gelatinous layer in secondary walls that are rich in cellulose but low in lignin content (Timell et al. 1967). In addition, some specialized secondary walls may be composed predominantly of one or two polymers. For example, secondary Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 walls of cotton fibers contain >90% cellulose with little xylan or lignin (Haigler et al. 2012), and secondary walls of phloem fibers from flax and hemp consist of cellulose and hemicelluloses without lignin (Mauseth et al. 1988). Because secondary walls in the form of wood and fibers are important raw materials for our daily uses and a potential source for biofuel production, there have been extensive studies on the molecular mechanisms controlling secondary wall composition, biosynthesis and assembly. In this review, we provide an update on the biosynthesis of secondary wall components, patterned deposition of secondary walls, and transcriptional regulation of secondary wall biosynthesis. Much of the current knowledge on genes involved in secondary wall biosynthesis stems from research in the model flowering plant, Arabidopsis thaliana, and therefore we will mainly focus on discussions of Arabidopsis secondary wall genes and only briefly remark on their homologs in other species. Due to the evolutionary conservation of secondary wall biosynthesis, it is anticipated that knowledge gained from the study of Arabidopsis secondary wall genes could be extrapolated into identifying genes involved in secondary wall biosynthesis in other species. Cellulose Biosynthesis Cellulose, being the load-bearing polymer in secondary walls, constitutes about 40–50% of wood components. It is made of linear chains of b-1,4-linked glucosyl residues with a degree of polymerization of approximately 10,000 (Timell et al. 1967) (Fig. 1D). It has long been held that 36 glucan chains are coalesced via hydrogen bonding into a highly insoluble crystalline microfibril. However, recent studies using X-ray and nuclear magnetic resonance techniques give an estimate of the diameter of a primary wall microfibril to be about 3 nm, which fits well with a model of 18 glucan chains per microfibril (Newman et al. 2013). The biosynthesis of cellulose is mediated by cellulose synthase complexes located at the plasma membrane (McFarlane et al. 2014). The complexes are visualized via transmission electron microscopy as rosettes made of six particles (McFarlane et al. 2014), and each particle is proposed to be an aggregate of three cellulose synthase A (CesA) subunits in the 18-CesA model (Newman et al. 2013). The 18-CesA model predicts that the 18 CesA subunits in each rosette synthesize 18 glucan chains that aggregate into a microfibril, which is consistent with the structural analysis of bacterial CesA revealing that each CesA polypeptide synthesizes and translocates one glucan chain (Morgan et al. 2013). The Arabidopsis genome harbors 10 genes encoding CesA proteins, which belong to glycosyltransferase family GT2 (McFarlane et al. 2014). Biosynthesis of secondary wall cellulose has been shown to involve three of them, CesA4/IRX5, CesA7/ IRX3 and CesA8/IRX1, which function non-redundantly (Taylor et al. 1999, Taylor et al. 2000, Taylor et al. 2003) (Table 1). Mutation of any of them causes a severe reduction in cellulose content and secondary wall thickening, and concomitantly a collapsed xylem phenotype. The three secondary wall CesAs interact with each other within the same cellulose synthase complex, and the presence of all of them is required for the normal assembly and targeting of the complex (Taylor et al. 2004, Timmers et al. 2009). In the 18-CesA model, each particle in the six particle rosette is proposed to contain one of each of the three interacting secondary wall CesAs (Newman et al. 2013). Close homologs of CesA4, CesA7 and CesA8 are present in other vascular plants and their disruption has been shown to result in a reduction in cellulose content and secondary wall thickening in rice, Brachypodium and Populus (Tanaka et al. 2003, Joshi et al. 2011, Handakumbura et al. 2013). In addition to CesA genes, a number of other genes have been implicated in the biosynthesis and assembly of secondary wall cellulose (Table 1). Mutation of the Arabidopsis KORRIGAN (KOR) gene, which encodes an endo-b-1,4-glucanase, causes a reduction in cellulose content in both primary and secondary walls (Nicol et al. 1998, Szyjanowicz et al. 2004). Although KOR does not associate with the secondary wall CesA complex (Szyjanowicz et al. 2004), it has been shown to be an integral part of the primary wall CesA complex (Vain et al. 2014). KOR is proposed to function as a cellulase in releasing the growing microfibrils from the cellulose synthase complex and/or in increasing the amount of non-crystalline cellulose microfibrils (Szyjanowicz et al. 2004, Takahashi et al. 2009). Arabidopsis COBRA-like 4 (COBL4), a glycosylphosphatidylinositol-anchored protein, and its rice ortholog, brittle culm 1 (BC1), are also implicated in cellulose biosynthesis since their mutations result in a reduction in cellulose content and secondary wall thickening (Li et al. 2003, Brown et al. 2005, Sato et al. 2010). BC1 has a carbohydrate-binding module that interacts with crystalline cellulose, and this interaction is proposed to modulate cellulose assembly and cellulose microfibril crystallinity (Liu et al. 2013). The Arabidopsis chitinase-like protein CTL2, whose expression is associated with secondary wall CesA genes, is proposed to modulate cellulose crystallization and cellulose interaction with hemicelluloses via binding to growing cellulose microfibrils (Sanchez-Rodriguez et al. 2012). A role for CTLs in modulating cellulose biosynthesis is consistent with the reduced cellulose content and mechanical strength of the rice brittle culm mutant bc15, which harbors a mutation in a CTL gene (Wu et al. 2012). The xylem-associated TED6 protein has been shown to interact with secondary wall CesA7, and RNA interference of its expression results in a reduction in secondary wall thickening, indicating its involvement in cellulose biosynthesis (Endo et al. 2009). Several additional genes are also implicated in secondary wall cellulose biosynthesis, including two Arabidopsis genes encoding fasciclin-like arabinogalactan proteins, AtFLA11 and AtFLA12, whose mutations reduce cellulose content and alter cellulose microfibril angles (MacMillan et al. 2010), a rice gene encoding a dynamin-related protein (BC3/ OsDRP2B), whose mutation causes a brittle culm phenotype with a decreased cellulose content and an altered secondary wall structure (Hirano et al. 2010, Xiong et al. 2010), and another rice gene encoding a DUF266 domain protein, whose mutation results in a brittle culm phenotype with a reduced cellulose content (Y. Zhou et al. 2009). In addition, a number of proteins with unknown functions have been shown to copurify with cellulose synthase complexes in Populus xylem, 197 R. Zhong and Z.-H. Ye | Secondary cell walls Table 1 Arabidopsis genes that are involved in secondary wall biosynthesis, patterned deposition and transcriptional regulation Gene AGI code Cellulose biosynthesis and assembly CesA4/IRX5 At5g44030 CesA7/IRX3 At5g17420 Functions References Cellulose synthase catalytic subunits; functioning redundantly in secondary wall cellulose biosynthesis Taylor et al. (1999, 2000, 2003) CesA8/IRX1 At4g18780 KORRIGAN At5g49720 Endo-b-1,4-glucanase; required for normal cellulose microfibril deposition Nicol et al. (1998), Szyjanowicz et al. (2004) COBL4 At5g15630 A glycosylphosphatidylinositol-anchored protein; modulating cellulose microfibril deposition Brown et al. (2005) CTL2 At3g16920 A chitinase-like protein; modulateing cellulose microfibril deposition Sanchez-Rodriguez et al. (2012) TED6 At1g43790 Interacting with CesA7 Endo et al. (2009) AtFLA11 At5g03170 MacMillan et al. (2010) AtFLA12 At5g60490 Fasciclin-like arabinogalactan proteins; involved in cellulose microfibril deposition Two functionally non-redundant groups of GT43 proteins (IRX9/ I9H vs. IRX14/I14H) required for xylan backbone elongation Brown et al. (2007), Pena et al. (2007), Lee et al. (2010), Wu et al. (2010) Xylan xylosyltransferases (GT47); catalyzing the elongation of xylan backbone Brown et al. (2009), Wu et al. (2009), Jensen et al. (2014), Urbanowicz et al. (2014) Xylan glucuronyltransferases (GT8); catalyzing the addition of GlcA side chains in xylan Mortimer et al. (2100), Lee et al. (2012a) Glucuronoxylan methylatransferases (DUF579); catalyzing GlcA methylation of xylan Lee et al. (2012d), Urbanowicz et al. (2012) DUF579 proteins required for normal xylan biosynthesis Brown et al. (2011), Jensen et al. (2011) Involved in acetylation of xylan; homologs of fungal CAS1 protein required for acetylation of glucuronoxylomannan in fungi Lee et al. (2011) Xylan biosynthesis IRX9 At2g37090 I9H/IRX9L At1g27600 IRX14 At4g36890 I14H/IRX14L At5g67230 IRX10/GUT2 At1g27440 IRX10L/GUT1 At5g61840 GUX1 At3g18660 GUX2 At4g33330 GUX3 At1g77130 GXM1 At1g09610 GXM2 At4g09990 GXM3/GXMT1 At1g09610 IRX15 At3g50220 IRX15L At5g67210 RWA1 At5g46340 RWA2 At3g06550 RWA3 At2g34410 RWA4 At1g29890 ESK1/TBL29 At3g55990 Xylan acetyltransferase catalyzing 2-O- and 3O-monoacetylation of xylan (DUF231) Xiong et al. (2013), Yuan et al. (2013), Urbanowicz et al. (2014) FRA8 At2g28110 At5g22940 Functional paralogs (GT47); required for the biosynthesis of xylan reducing end sequence Zhong et al. (2005b), Lee et al. (2009) F8H IRX8 At5g54690 A GT8 protein; required for the biosynthesis of xylan reducing end sequence Brown et al. (2007, Pena et al. (2007, Persson et al. (2007 PARVUS At1g19300 A GT8 protein; required for the biosynthesis of xylan reducing end sequence Brown et al. (2007), Lee et al. (2007b) Glucomannan synthases; cellulose synthaselike A family Liepman et al. (2005), Goubet et al. (2009) Glucomannan biosynthesis CslA2 At5g22740 CslA3 At1g23480 CslA9 At5g03760 (continued) 198 Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 Table 1 Continued Gene AGI code Lignin biosynthesis and polymerization PAL1 At2g37040 Functions References Phenylalanine ammonia lyases; highly expressed in stems Raes et al. (2003) PAL2 At3g53260 PAL4 At3g10340 C4H At2g30490 Cinnamate 4-hydroxylase; highly expressed in stems Raes et al. (2003) 4CL1 At1g51680 At3g21240 4-Commarate CoA ligases; highly expressed in stems Raes et al. (2003) 4CL2 HCT At5g48930 Hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase; highly expressed in stems Raes et al. (2003) C3H1 At2g40890 p-Coumaroyl shikimate 30 -hydroxylase; highly expressed in stems Raes et al. (2003) CSE At1g52760 Caffeoyl shikimate esterase; converting caffeoyl shikimate into caffeate Vanholme et al. (2013) Caffeoyl CoA O-methyltransferases; highly expressed in stems Raes et al. (2003 CCoAOMT1 At4g34050 CCoAOMT7 At4g26220 CCR1 At1g15950 Cinnamoyl CoA reductase; highly expressed in stems Raes et al. (2003) F5H1 At4g36220 Ferulate 5-hydroxylase; highly expressed in stems Raes et al. (2003) COMT At5g54160 Caffeic acid O-methyltransferase; highly expressed in stems Raes et al. (2003) Cinnamyl alcohol dehydrogenases; highly expressed in stems Raes et al. (2003) Laccases; catalyzing the polymerization of monolignols Berthet et al. (2011), Zhao et al. (2013) Peroxidases; catalyzing the polymerization of monolignols Herrero et al. (2013b), Shigeto et al. (2013, 2014) CAD3 At4g34230 CAD6 At4g37970 LAC4 At2g38080 LAC11 At5g03260 LAC17 At5g60020 AtPRX2 At1g05250 AtPRX25 At2g41480 AtPRX71 At5g64120 AtPRX72 At5g66390 PDR1/AtABCG29 At3g16340 ATP-binding cassette-like transporter; involved in the transport of p-coumaryl alcohol Alejandro et al. (2012) MED5a/REF4 At2g48110 At3g23590 Transcriptional co-regulatory complex subunits; required for homeostatic repression of phenylpropanoid biosynthesis Bonawitz et al. (2014) MED5b/RFR1 Interacting with cortical microtubules and CesAs; functioning as a scaffold protein to guide the movement of CesA complexes along the track of cortical microtubules Gu et al. (2010), Bringmann et al. (2012), S. Li et al. (2012) Patterned deposition of secondary walls CSI1 At2g22125 AtKTN1/FRA2 At1g80350 Katanin microtubule-severing protein; essential for cortical microtubule organization and oriented deposition of cellulose microfibrils Burk et al. (2001, 2002) FRA1 At5g47820 Kinesin motor protein; required for cellulose microfibril deposition and secondary wall strength Zhong et al. (2002) MAP65–8 At1g27920 Microtubule-associated protein whose zinnia ortholog is involved in bundling of microtubules Mao et al. (2006) (continued) 199 R. Zhong and Z.-H. Ye | Secondary cell walls Table 1 Continued Gene AGI code Functions References MAP70–1 At1g68060 Pesquet et al. (2010) MAP70–5 At4g17220 Plant-specific microtubule-associated proteins; regulating the organization of cortical microtubule bundles MIDD1 At3g53350 Interacting with microtubules; acting as a scaffold protein to recruit Kinesin-13A for microtubule depolymeirzation Oda et al. (2010), Oda and Fukuda (2013a) Kinesin-13A At3g16630 Microtubule depolymerization protein; depolymerizing cortical microtubules underneath the pit-forming areas Oda and Fukuda (2013a) ROP11 At5g62880 GTPase; recruiting MIDD1/Kinesin-13A to the microtubule-depleting plasma membrane domains Oda and Fukuda (2012) ROPGAP3 At2g46710 ROP GTPase-activating protein; inactivating ROP11 Oda and Fukuda (2012) ROPGEF4 At2g45890 Guanine nucleotide exchange factor; activating ROP11 Oda and Fukuda (2012) Vessel-specific secondary wall NAC master switches; functioning redundantly in regulating secondary wall biosynthesis in vessels Kubo et al. (2005), Yamaguchi et al. (2008), Zhou et al. (2014) Secondary NAC master switches regulating secondary wall biosynthesis in fibers (NST1, NST2 and SND1) and anther endothecium (NST1 and NST2) Mitsuda et al. (2005), Zhong et al. (2006, 2007b), Mitsuda et al. (2007, 2008), Zhong and Ye (2014) Secondary wall-associated NACs required for normal secondary wall biosynthesis Zhong et al. (2008) Secondary wall MYB master switches activating secondary wall biosynthesis Zhong et al. (2007a), McCarthy et al. (2009) Secondary wall-associated MYBs induced by SND1, NSTs and VNDs; involved in regulating secondary wall biosynthesis Zhong et al. (2008) Lignin-specific transcriptional activators Zhong et al. (2008), J. Zhou et al. (2009) Transcriptional regulation VND1 At2g18060 200 VND2 At4g36160 VND3 At5g66300 VND4 At1g12260 VND5 At1g62700 VND6 At5g62380 VND7 At1g71930 NST1 At2g46770 NST2 At3g61910 SND1 At1g32770 SND2 At4g28500 SND3 At1g28470 MYB46 At5g12870 MYB83 At3g08500 MYB20 At1g66230 MYB42 At4g12350 MYB43 At5g16600 MYB52 At1g17950 MYB54 At1g73410 MYB69 At4g33450 MYB103 At1g63910 MYB58 At1g16490 MYB63 At1g79180 MYB85 At4g22680 KNAT7 At1g62990 A homeodomain transcription factor proposed to repress secondary wall biosynthesis E. Li et al. (2012) LBD18 At2g45420 Soyano et al. (2008) LBD30 At4g00220 LOB domain proteins involved in a positive feedback loop to regulate VND7 expression XND1 At5g64530 A NAC domain protein shown to repress secondary wall biosynthesis and programmed cell death in vessels Zhao et al. (2008) MYB4 At4g38620 MYB7 At2g16720 Sonbol et al. (2009), Fornale et al. (2010), Legay et al. (2010), Shen et al. (2012) MYB32 At4g34990 Induced by MYB46 and their orthologs in other species known to be transcriptional repressors of lignin biosynthesis (continued) Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 Table 1 Continued Gene AGI code Functions References BLH6 At4g34610 A BEL1-like homeodomain transcription factor involved in regulating lignin biosynthesis Cassan-Wang et al. (2013) MYB26 At3g13890 A MYB specifically regulating secondary wall biosynthesis in anther endothecium Yang et al. (2007) Others genes important for secondary wall deposition IFL1/REV At5g60690 A homeodomain leucine-zipper protein essential for fiber differentiation and secondary wall thickening Zhong and Ye (1999) WAT1 At1g75500 A vacuolar auxin transporter essential for fiber differentiation and secondary wall thickening Ranocha et al. (2013) EXO70A1 At5g03540 A subunit of exocyst complex proposed to function in vesicle trafficking in vessels Li et al. (2013) FRA3 At1g65580 At1g22620 Phosphoinositide phosphatases essential for actin organization and secondary wall deposition Zhong et al. (2004, 2005a) AtSAC1/FRA7 RHD3 At3g13870 A GTPase essential for actin organization and secondary wall deposition Hu et al. (2003) suggesting their potential roles in cellulose biosynthesis (Song et al. 2010). Xylan Biosynthesis Xylan, a major hemicellulosic component in secondary walls of angiosperm species, is a polymer consisting of a backbone of b1,4-linked xylosyl residues with the degree of polymerization around 100 (Fig. 1D). The xylosyl residues are often substituted with monosaccharide residues, including a-1,2-linked glucuronic acid (GlcA), a-1,2-linked 4-O-methylglucuronic acid (MeGlcA), and a-1,2 and/or a-1,3-linked arabinose (Ara) (Ebringerova and Heinze 2000). Although monosaccharide substituents are dominant, xylans from grasses may contain disaccharide side chains composed of a-1,3-linked Ara substituted at O-2 with Ara or xylose (Xyl) (Ebringerova and Heinze 2000). Xylans from Eucalyptus globulus and Arabidopsis also contain disaccharide side chains composed of a-1,2-linked GlcA/ MeGlcA substituted at O-2 with galactose (Shatalov et al. 1999, Zhong et al. 2014). Based on the nature of their substitutions, xylans are named glucuronoxylan, arabinoxylan and glucuronoarabinoxylan. Glucuronoxylan is the dominant form of xylan in dicot secondary walls, and glucuronoarabinoxylan is the dominant form in the wood of gymnosperms and cell walls of grass species. In addition to the sugar substitutions, xylans may be acetylated at C-2 and/or C-3, with the degree of acetylation varying from 40% to 70% (Teleman et al. 2002, Evtuguin et al. 2003, Goncalves et al. 2008). In grasses, Ara substituents of xylan may be covalently cross-linked with lignin via ferulic acid residues (Ebringerova and Heinze 2000). Xylans from gymnosperms and dicots contain a distinct tetrasaccharide sequence, b-D-Xyl-(1!3)-a-L-Rha-(1!2)-a-D-GalA-(1!4)-D-Xyl, at its reducing end (Shimizu et al. 1976, Johansson and Samuelson 1977, Andersson et al. 1983, Pena et al. 2007, Lee et al. 2009a). The biosynthesis of xylan requires a number of enzymes, including xylosyltransferases, glucuronyltransferases, methyltransferases, acetyltransferases and arabinosyltransferases, that are responsible for xylan backbone elongation and substitutions (Table 1). Genetic and biochemical analysis in Arabidopsis has uncovered the essential roles of glycosyltransferases from GT43 and GT47 families in xylan backbone elongation. There exist four GT43 genes in the Arabidopsis genome, IRX9, I9H/IRX9-L, IRX14 and I14H/IRX14-L, all of which are expressed in secondary wall-forming cells, including vessels and fibers, and their encoded proteins are located in the Golgi (Brown et al. 2007, Pena et al. 2007, Lee et al. 2010, Wu et al. 2010). Mutation of IRX9 or IRX14 alone is sufficient to cause a drastic reduction in xylan xylosyltransferase activity, xylan chain length and xylan content (Brown et al. 2007, Lee et al. 2007a, Pena et al. 2007). As a result, the secondary wall thickness in fibers and vessels of the mutants is severely reduced and the vessels are collapsed due to their inability to resist the negative pressure generated during transpiration. Genetic complementation studies have revealed that the four GT43 genes form two functionally non-redundant groups, IRX9 and I9H being in one and IRX14 and I14H being in another, both of which are required for xylan backbone elongation (Lee et al. 2010). Coexpression of IRX9 and IRX14 in tobacco BY2 cells leads to a significant increase in xylosyltransferase activity that is able to add xylosyl residues successively from the UDP-Xyl donor onto the xylooligomer acceptor (Lee et al. 2012b). GT43 genes from Populus and rice also form two functionally non-redundant groups (Lee et al. 2012c, Lee et al. 2014), indicating that the recruitment of two functionally non-redundant groups of GT43 201 R. Zhong and Z.-H. Ye | Secondary cell walls genes in xylan backbone elongation is conserved in vascular plants. Mutations of two GT47 genes, IRX10 and IRX10-L, cause similar phenotypes to the mutation of IRX9 or IRX14, such as a reduction in xylan xylosyltransferase activity, xylan chain length, xylan content and secondary wall thickness (Brown et al. 2009, Wu et al. 2009). Activity assays of recombinant IRX10 and IRX10L proteins and their close homologs from moss and psyllium revealed that they exhibit xylan xylosyltransferase activity that is capable of adding xylosyl residues onto the xylooligomer acceptor (Jensen et al. 2014, Urbanowicz et al. 2014). In the developing mucilaginous tissues of psyllium where xylan is predominant, IRX10 homologs are highly expressed but GT43 homologs are not, suggesting the possibility that IRX10 proteins may function without GT43 proteins in xylan backbone elongation in the mucilaginous tissues of psyllium (Jensen et al. 2013). Although IRX9, IRX14 and IRX10 are all required for the xylan xylosyltransferase activity, IRX9 and IRX14 are proposed to play a structural instead of a catalytic role in xylan backbone biosynthesis (Ren et al. 2014). The addition of GlcA side chains in Arabidopsis xylan is mediated by three glucuronyltransferase genes (GUX1, GUX2 and GUX3) that belong to family GT8 (Mortimer et al. 2010, Lee et al. 2012a). These GUX genes are expressed in secondary wall-forming cells and their encoded proteins are targeted to the Golgi. Mutation of a single GUX gene causes a partial reduction in the amount of GlcA, but no effects on plant growth and secondary wall thickness. In contrast, simultaneous mutations of all three GUX genes lead to a complete loss of GlcA side chains in xylan, a reduction in secondary wall thickness and a deformation of vessels (Lee et al. 2012a). The finding that GUXs exhibit xylan glucuronyltransferase activities when heterologously expressed in tobacco cells (Lee et al. 2012a, Rennie et al. 2012) further supports that GUX1/2/3 are glucuronyltransferases responsible for GlcA substitution of xylan during secondary wall biosynthesis. About 60% of the GlcA side chains in Arabidopsis xylan are methylated at O-4, and it has been shown that the GlcA methylation of xylan is catalyzed by three Golgi-localized glucuronoxylan methyltransferases (GXM1, GXM2 and GXM3/GXMT1) that belong to a family of DUF579 domain-containing proteins (Lee et al. 2012d, Urbanowicz et al. 2012). The three GXM genes are expressed in secondary wall-forming cells and their recombinant proteins exhibit methyltransferase activity that is capable of transferring the methyl group from S-adenosylmethionine to GlcA-substituted xylooligomers. Although mutation of individual GXM genes causes a partial reduction in the degree of GlcA methylation, simultaneous mutations of all three GXM genes lead to a complete loss of GlcA methylation in xylan (Yuan et al. 2014a). No reduction in xylan content and secondary wall thickening was detected in the gxm1/2/3 triple mutant, indicating that loss of GlcA methylation has no negative impact on xylan biosynthesis and secondary wall assembly. The GlcA side chains in Populus wood xylan are completely methylated, which has been shown to be mediated by four wood-associated GXM homologs, PtrGXM1/2/3/4 (Yuan et al. 202 2014b). Recombinant proteins of PtrGXM3 and PtrGXM4 exhibit 10 times lower Km values than GXM3/GXMT1 that is responsible for the bulk of GlcA methylation of Arabidopsis xylan, indicating that PtrGXM3 and PtrGXM4 have a much higher substrate affinity than Arabidopsis GXM, which may enable a complete methylation of GlcA side chains in Populus xylan during wood formation. In Arabidopsis, the GXM activity could be rate limiting so that it may not be able to catch up with the rate of xylan biosynthesis to methylate all the GlcA side chains during secondary wall biosynthesis. This hypothesis is supported by overexpression analysis showing that an increase in GXM activity is sufficient to methylate all GlcA side chains in Arabidopsis xylan (Yuan et al. 2014a). Two additional DUF579 domain-containing proteins, IRX15 and IRX15L, are involved in xylan biosynthesis, although their exact functions remain to be investigated (Brown et al. 2011, Jensen et al. 2011). The addition of Ara side chains in grass xylan has been shown to be mediated by glycosyltransferases from family GT61 (Anders et al. 2012). RNA interference inhibition of a wheat GT61 gene, TaXAT1, causes a specific decrease in a-1,3-linked Ara substitution of xylan in wheat endosperm. Heterologous expression of TaXAT2 and its rice close homologs, OsXAT2 and OsXAT3, in Arabidopsis results in a-1,3linked Ara substitution of xylan, which normally does not occur in Arabidopsis. These findings provide genetic evidence demonstrating that XATs are probably arabinosyltransferases that catalyze the a-1,3-linked Ara substitution of grass xylan. Another GT61 gene, XAX1, is proposed to mediate the transfer of b-1,2-linked Xyl onto the Ara side chains that are attached to the xylan backbone at O-3 (Chiniquy et al. 2012). O-Acetylation of xylan involves the RWA genes and the ESK1/TBL29 gene. The Arabidopsis genome contains four RWA genes, all which are expressed in secondary wall-forming cells. Simultaneous mutations of the four RWA genes result in a 40% reduction in xylan acetylation, reduced secondary wall thickening and a collapsed vessel morphology, indicating their roles in xylan acetylation (Lee et al. 2011). RWAs are close homologs of fungal CAS1 that is involved in acetylation of glucuronoxylomannan, a main capsule polysaccharide (Janbon et al. 2001). Because RWAs contain only multiple transmembrane domains but lack the putative acetyltransferase domain found in CAS1, they are proposed to be putative transporters for channeling the acetyl donor, acetyl-CoA, from the cytoplasm to the Golgi lumen (Gille and Pauly 2012). ESK1 belongs to a large, plant-specific DUF231 domain protein family, several other members of which have been shown to be involved in O-acetylation of plant cell wall polysaccharides (Gille and Pauly 2012). ESK1 is specifically expressed in secondary wall-forming cells and its encoded protein is located in the Golgi (Yuan et al. 2013). The esk1 mutation causes a reduction in secondary wall thickening and a deformation of vessel morphology, but no decrease in xylan content. Analysis of xylan acetylation and acetyltransferase activity revealed that the esk1 mutant had a specific reduction in 2-O- and 3-Omonoacetylation of xylan and a drastic decrease in xylan acetyltransferase activity compared with the wild type, indicating that ESK1 is a putative acetyltransferase mediating 2-O- and Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 3-O-monoactylation of xylan (Xiong et al. 2013, Yuan et al. 2013). Activity assay of recombinant ESK1 confirmed that it exhibits acetyltransferase activity catalyzing 2-O- and 3-Omonoacetylation of xylan (Urbanowicz et al. 2014). A number of genes involved in the biosynthesis of xylan reducing end tetrasaccharide sequence have been identified (Zhong et al. 2005b, Brown et al. 2007, Lee et al. 2007b, Pena et al. 2007, Persson et al. 2007, Lee et al. 2009b). These genes include FRA8, F8H, IRX8 and PARVUS, all of which are expressed in secondary wall-forming cells, and their encoded proteins are located in the Golgi, except for PARVUS that is predominantly located in the endoplasmic reticulum. FRA8 and it close homolog, F8H belong to family GT47, and IRX8 and PARVUS belong to family GT8. Mutation of FRA8, IRX8 or PARVUS alone causes a reduction in xylan content and secondary wall thickness, and a loss of the xylan reducing end tetrasaccharide sequence, b-DXyl-(1!3)-a-L-Rha-(1!2)-a-D-GalA-(1!4)-D-Xyl (Brown et al. 2007, Lee et al. 2007b, Pena et al. 2007, Lee et al. 2009b). F8H is a functional paralog of FRA8, and simultaneous mutations of FRA8 and F8H result in a severe deformation of vessels and an extreme retardation in plant growth (Lee et al. 2009b). No reduction in xylan xylosyltransferase or glucuronyltransferase activities occurs in these mutants (Lee et al. 2007a). The available evidence indicates that these glycosyltransferases mediate the biosynthesis of the xylan reducing end sequence. Based on their catalytic mechanisms, it was proposed that FRA8 might catalyze the transfer of Xyl to Rha or of Rha to GalA, and IRX8 might mediate the transfer of GalA to the reducing Xyl residue at the reducing end of xylan (Pena et al. 2007). PARVUS might be involved in the initiation of the reducing end sequence biosynthesis by transferring the reducing end Xyl residue to an unknown acceptor (Lee et al. 2007b). Glucomannan Biosynthesis Glucomannan, a minor hemicellulosic component of dicot wood, consists of a linear chain of b-1,4-linked mannosyl residues interspersed with glucosyl residues with a ratio of 2 : 1 (Timell 1967) (Fig. 1D). Galactoglucomannan, a major hemicellulosic component of gymnosperm wood, is composed of the glucomannan backbone substituted with a-1,6-linked galactosyl residues. Glucomannan from aspen and birch wood has been shown to be O-acetylated at C-2 or C-3 of mannosyl residues, with a degree of acetylation of about 0.3 (Teleman et al. 2003). The biosynthesis of glucomannan is catalyzed by members of the cellulose synthase-like A (CslA) family, which belong to family GT2 (Dhugga et al. 2004, Liepman et al. 2005). Recombinant Arabidopsis CslA proteins are able to catalyze the transfer of both mannosyl and glucosyl residues from GDP-mannose and GDP-glucose, respectively, to produce b-linked glucomannan polymers, demonstrating that CslA family members are glucomannan synthases. There are nine CslA genes in the Arabidopsis genome, and three of them, CslA2, CslA3 and CslA9, are responsible for the synthesis of glucomannan in secondary wall-containing cells of stems (Goubet et al. 2009) (Table 1). CslA genes are present in all taxa of land plants, and those from tree species, including pine and Populus, have been demonstrated to exhibit glucomannan synthase activities (Suzuki et al. 2006, Liepman et al. 2007). The a-1,6-linked galactosyl substitution in galactoglucomannan is catalyzed by UDP-galactose-dependent a-1,6-D-glucomannan galactosyltransferase, whose corresponding gene was identified in fenugreek (Edwards et al. 1999). Lignin Biosynthesis Lignin is a complex three-dimensional polyphenolic polymer that is generated through oxidative polymerization in muro of three monolignols, p-coumaryl, coniferyl and sinapyl alcohols (Fig. 1D). The lignin polymer is divided into three types, p-hydroxyphenyl (H) lignin that is polymerized from p-coumaryl alcohol, guaiacyl (G) lignin polymerized from coniferyl alcohol, and syringyl (S) lignin polymerized from sinapyl alcohol. Lignin composition, i.e. the proportion of each of three polymerized monolignols, may differ significantly among different plant species and even among different cell types in the same species. For example, secondary walls in gymnosperms contain guaiacyl lignin but lack syringyl lignin, whereas those from dicot species are typically rich in both guaiacyl lignin and syringyl lignin. In secondary walls of grass species, a significant proportion of p-hydroxyphenyl lignin is present in addition to guaiacyl lignin and syringyl lignin (Bonawitz and Chapple 2010). Monolignol biosynthesis is carried out through the common phenylpropanoid pathway starting from phenylalanine to produce hydroxycinnamoyl-CoA esters. These esters are then routed into the lignin branch pathway to generate monolignols. At least 10 enzymes are involved in catalyzing monolignol biosynthesis; they are phenylalanine ammonia lyase (PAL), cinnamate 4-hydroxylase (C4H), 4-coumarate CoA ligase (4CL), hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase (HCT), p-coumaroyl shikimate 30 -hydroxylase (C3H), caffeoyl CoA O-methyltransferase (CCoAOMT), ferulate 5hydroxylase (F5H), caffeic acid O-methyltransferase (COMT), cinnamoyl CoA reductase (CCR) and cinnamyl alcohol dehydrogenase (CAD) (Bonawitz and Chapple 2010). It has recently been demonstrated that caffeoyl shikimate esterase (CSE) that catalyzes the conversion of caffeoyl shikimate into caffeic acid is another enzyme in the lignin biosynthetic pathway (Vanholme et al. 2013). Genes encoding these monolignol biosynthetic enzymes have been identified and functionally characterized in a number of species (Bonawitz and Chapple 2010). In Arabidopsis, 35 monolignol biosynthetic genes have been identified, and the expression of 16 of them, i.e. PAL1, PAL2, PAL4, C4H, 4CL1, 4CL2, HCT, C3H1, CSE, CCoAOMT1, CCoAOMT7, CCR1, F5H, COMT, CAD3 and CAD6, is closely associated with the development of lignified xylem and interfascicular fibers in stems, indicating that they are the core monolignol biosynthetic genes responsible for lignin synthesis during secondary wall formation in stems (Raes et al. 2003, Vanholme et al. 2013) (Table 1). Similarly, expression analysis identified 18 monolignol biosynthetic genes that are abundant and specific in differentiating xylem in Populus and are likely to 203 R. Zhong and Z.-H. Ye | Secondary cell walls be responsible for lignin synthesis during wood formation (Shi et al. 2010). Mutants with mutation of each of the monolignol biosynthetic genes in Arabidopsis have been generated and analyzed by transcriptomics and metabolomics. Although not all of the mutants have reduced lignin content and altered lignin composition, the expression of many genes and the level of various metabolites are altered in these mutants (Vanholme et al. 2012). Monolignols are synthesized in the cytosol and then transported into cell walls, where they are polymerized into lignin via oxidative reactions catalyzed by oxidases, including laccases and peroxidases (Bonawitz and Chapple 2010). The transport of monolignols into cell walls is proposed to be mediated by plasma membrane-localized transporters (Kaneda et al. 2008, Liu et al. 2011). Using isolated Arabidopsis plasma membrane vesicles, it has been demonstrated that plasma membranes contain ATP-binding cassette-like transporter activities that are able to transport monolignols (Miao and Liu. 2010). Heterologous expression of one of the Arabidopsis ATP-binding cassette-like transporter genes, AtABCG29, in yeast cells results in an increased activity of p-coumaryl alcohol transport, suggesting that AtABCG29 is involved in the transport of p-coumaryl alcohol (Alejandro et al. 2012). Transporters responsible for the transport of coniferyl and sinapyl alcohols remain to be identified. Several genes encoding laccases and peroxidases have been shown to be involved in lignin polymerization during secondary wall biosynthesis in Arabidopsis and Populus (Table 1). Simultaneous mutation of Laccase 4 (LAC4) and LAC17 results in a reduction in lignin content in Arabidopsis stems (Berthet et al. 2011) and simultaneous mutation of LAC4 and LAC17 together with LAC11 leads to plant growth arrest and a loss of lignification in root vessels (Zhao et al. 2013). Down-regulation of laccase genes in Populus by overexpression of a microRNA that targets a number of laccase genes leads to a reduction in laccase activity and lignin content in transgenic wood (Lu et al. 2013). Several Arabidopsis peroxidase genes have been implicated in lignin polymerization (Herrero et al., 2013a); mutation of AtPRX72 causes a reduction in lignin content (Herrero et al. 2013b) and mutations of AtPRX2, AtPRX25 and AtPRX71 result in reduced lignin content and altered lignin structure in secondary walls (Shigeto et al. 2013, Shigeto et al. 2014). Mutants with a reduction in lignin content often lead to impaired secondary wall deposition, deformed vessels and reduced plant growth (Zhong et al. 1998, Jones et al. 2001, Ruel et al. 2009), which may hinder the effort of generating low-lignin crops for biofuel production. The recent finding that the reduced growth and abnormal secondary wall deposition seen in the ref8 mutant, which is due to a mutation in the C3H1 gene, could be partially rescued by mutations of two genes encoding transcriptional co-regulatory complex subunits MED5a and MED5b may provide a new tool to overcome the plant growth defect caused by a reduced lignin content (Bonawitz et al. 2014). An alteration in lignin structure without a change in lignin content and plant growth has been achieved by introduction into Populus of a monolignol ferulate transferase that generates monolignol ferulate conjugates (Wilkerson 204 et al. 2014). Lignin in transgenic Populus wood with incorporation of ester-linked monolignol ferulates is easier to remove, which represents a novel approach to engineer biofuel crops with easy removal of lignin. Patterned Deposition of Secondary Walls Secondary walls in tracheary elements and fibers of vascular plants are deposited in elaborate patterns. Tracheary elements (protoxylem) in elongating organs typically have secondary walls deposited in a helical or annular pattern, which provides the necessary mechanical support to the xylem conduits and at the same time allows organs to continue to elongate. Tracheary elements (metaxylem) in non-elongating organs typically have secondary walls organized in a reticulated or pitted pattern (Fig. 2A), which provides maximum mechanical support to the xylem conduits and the plant body. Fiber cells generally have secondary walls deposited uniformly between the primary wall and the plasma membrane, except for the pit areas that are used for solute transport (Mauseth, 1988) (Fig. 2B). Secondary walls typically contain three distinct layers, called S1, S2 and S3, when observed using transmission electron microscopy (Fig. 1B). The layered deposition of secondary walls is caused by changes in the orientations of cellulose microfibrils. Both the S1 layer and the S3 layer are typically thin, and the cellulose microfibrils in these two layers are oriented in a crossed organization and a flat helix relative to the cell elongation axis, respectively. The S2 layer is the thickest, with cellulose microfibrils oriented almost parallel to the cell elongation axis (Fig. 1C), and the angles of these microfibrils largely determine the mechanical strength of fibers in wood (Timell 1967). The number of layers in secondary walls may change in response to environmental stimuli, and secondary walls in some plant species may have more than three layers. For example, secondary walls in compression wood lack the S3 layer but contain an extra layer of lignin located between S1 and S2, whereas those in tension wood may lack one or two of the three typical layers but often have an additional cellulose-rich gelatinous layer next to the lumen (Timell et al. 1967). In addition, up to six distinct layers are observed in the secondary walls of bamboo fibers (Gritsch and Murphy 2005). It has long been known that cortical microtubules play a crucial role in both the oriented deposition of cellulose microfibrils and the patterned deposition of secondary walls. At the subcellular level, the orientation of cortical microtubules coaligns with that of cellulose microfibrils, and disruption of the cortical microtubule organization by pharmacological drugs results in an altered orientation of cellulose microfibrils, which led to the alignment hypothesis that cortical microtubules guide the oriented deposition of cellulose microfibrils (Baskin 2001). This hypothesis is supported by the study of fluorescent protein-tagged CesAs showing that CesA complexes move along the track of cortical microtubules located underneath the plasma membrane, and disruption of the cortical microtubule organization alters the movement of CesA complexes (Paradez et al. 2006). Primary wall CesAs have been found to interact Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 Fig. 2 Patterned deposition of secondary walls. (A) Ectopic deposition of secondary walls in a reticulated pattern in Arabidopsis leaf epidermal cells overexpressing SND1. Note the gap areas between thickened walls. (B) Transmission electron micrograph showing a pair of bordered pits in secondary walls of interfascicular fibers from an Arabidopsis stem. Scale bar = 2.6 mm. (C) Diagram of regulation of patterned deposition of secondary walls. Cortical microtubules are bundled beneath the plasma membrane where secondary wall thickening occurs. Bundling of cortical microtubules and delimitation of the borders of bundled cortical microtubules are facilitated by microtubule-associated proteins (MAP65 and MAP70). Cortical microtubules, actin cytoskeleton and the exocyst complex (including a subunit EXO70A1) are involved in targeted transport of CesA complexes and vesicles carrying secondary wall components. Formation of the pit area with no secondary wall thickening is controlled by the ROP11/ MIDD1/Kinesin-13A complex that mediates local depolymerization of cortical microtubules. with cortical microtubules via cellulose synthase-interacting protein 1 (CSI1) (Gu et al. 2010) (Table 1). CSI1 directly interacts with microtubules and the cytoplasmic domain of CesAs, and csi1 mutation affects the distribution and the movement of CesA complexes. CSI1 is suggested to function as a scaffold protein to guide the movement of CesA complexes along the track of cortical microtubules (Bringmann et al. 2012, S. Li et al. 2012). Although the functional roles of CSI1 in secondary wall cellulose biosynthesis have not been investigated, it is likely that CSI1 or CSI homologs may also guide the movement of secondary wall CesA complexes along cortical microtubules, and thereby control the oriented deposition of cellulose microfibrils in secondary walls. Since the orientation of cortical microtubules is correlated with the direction of CesA movement and the subsequent orientation of cellulose microfibrils, it is likely that the different orientations of cellulose microfibrils seen in the S1, S2 and S3 layers of secondary walls reflect the dynamic re-organization of cortical microtubules during the transition of formation of different secondary wall layers. An important role for cortical microtubules in controlling the oriented deposition of cellulose microfibrils during secondary wall biosynthesis is exemplified by mutation of the AtKTN1 gene in the fra2 mutant (Burk et al. 2007) (Table 1). AtKTN1 encodes a katanin microtubule-severing protein, and its mutation in the fra2 mutant causes a disorganization of cortical microtubules, an aberrant orientation of cellulose microfibrils and a loss of layers in the secondary walls of fibers. These findings provide genetic evidence supporting a role for cortical microtubules in directing the formation of layered secondary walls (Burk et al. 2001, Burk and Ye. 2002). It has been shown that the kinesin motor proteins, Arabidopsis FRA1 and its rice ortholog BC12, also play a significant role in the oriented deposition of cellulose microfibrils and the mechanical strength of secondary walls (Zhong et al. 2002, Zhang et al. 2010) (Table 1). Mutation of FRA1 or BC12 has no effect on cortical microtubule organization and cellulose content, but it drastically reduces the mechanical strength of fibers, probably due to the disorganization of cellulose microfibrils (Zhong et al. 2002, Zhang et al. 2010). FRA1 overexpression results in a reduction in cellulose content and secondary wall thickness in fibers, a collapsed vessel phenotype and an increase in the number of layers in secondary walls (Zhou et al. 2007). FRA1 and BC12 belong to the kinesin-4 subfamily, and kinesin-4 proteins in animal cells are implicated in regulating the movement of multiple intracellular components, such as vesicle transport (Martinez et al. 2008). It is likely that FRA1 and BC12 are involved in the transport of vesicles carrying cell wall materials along microtubules and thereby influence cellulose microfibril deposition and secondary wall assembly. It has been demonstrated that cortical microtubules in tracheary elements form bundles beneath specific plasma membrane domains where secondary wall thickening occurs (Oda et al. 2005). Disruption of the cortical microtubule organization by pharmacological drugs alters the patterned deposition of secondary walls in tracheary elements, suggesting that cortical microtubules determine the patterned deposition of secondary walls. It has been shown that CesA complexes 205 R. Zhong and Z.-H. Ye | Secondary cell walls responsible for cellulose microfibril deposition and laccases involved in monolignol polymerization are concentrated as bands at the plasma membrane domains where secondary wall thickening occurs, and the underlying cortical microtubule bundles are essential for CesA localization (Wightman and Turner 2008, Schuetz et al. 2014). It is evident that the cortical microtubule bundles not only control the trajectory of CesA movement in the plasma membrane but also direct the targeted transport of vesicles carrying the CesA complexes, hemicelluloses and monolignol polymerization enzymes to specific plasma membrane domains, and thereby control the patterned deposition of secondary walls. The bundling of cortical microtubules is mediated by MAP65 microtubule-associated proteins. ZeMAP65-1 has been shown to co-localize with cortical microtubule bundles in the Zinnia tracheary elements and, when overexpressed in Arabidopsis suspension cells, it induces the normally evenly distributed cortical microtubules to form thick helical or transverse bundles that resemble cortical microtubule bundles seen in the tracheary elements (Mao et al. 2006). In addition, two plant-specific MAPs, the Arabidopsis MAP70-5 and MAP70-1, are proposed to play an essential role in defining the locations of secondary wall thickening through regulating the organization of cortical microtubule bundles (Pesquet et al. 2010). It has been shown that MAP70-5 lines the borders of each cortical microtubule bundle. MAP70-5 expression is induced during tracheary element differentiation in Arabidopsis suspension cells, and manipulating the level of MAP70-5 or MAP70-1 by their down-regulation or overexpression leads to a loss of patterned secondary wall deposition or an increase in the population of tracheary elements with the spiral secondary wall pattern, respectively (Pesquet et al. 2010). These findings indicate that MAP65 and MAP70 proteins play crucial roles in bundling cortical microtubules and defining the borders of cortical microtubule bundles, respectively, and thereby regulate the pattered secondary wall deposition (Table 1; Fig. 2C). Cortical microtubule bundles are separated by distinctive gaps that are low in microtubules. It was proposed that the maintenance of low levels of microtubules in the gap areas is mediated by the ROP11/MIDD1/Kinesin-13A complexes, which are located underneath the microtubule-depleting plasma membrane domains that coincide with regions where no secondary wall thickening occurs (Oda and Fukuda 2013b) (Table 1; Fig. 2C). In these complexes, the microtubule depletion domain 1 protein, MIDD1, directly interacts with microtubules (Oda et al. 2010) and acts as a scaffold protein to recruit the microtubule depolymerization protein, Kinesin-13A, to depolymerize cortical microtubules locally, leading to formation of microtubule-deleting plasma membrane domains (Oda and Fukuda 2013a). RNA interference inhibition of MIDD1 results in a failure of localized depolymerization of cortical microtubules and concomitantly a uniform deposition of secondary walls (Oda et al. 2010). Similarly, RNA interference inhibition of Kinesin-13A causes formation of secondary walls with smaller pit areas. In contrast, overexpression of Kinesin-13A leads to increased depolymerization of cortical microtubules and formation of secondary walls with larger pit areas (Oda and 206 Fukuda 2013a). The recruitment of MIDD1/Kinesin-13A to the microtubule-depleting plasma membrane domains is mediated by the active form of ROP11 GTPase. The inactivation and activation of ROP11 are controlled by a ROP GTPase-activating protein (ROPGAP3) and a ROP guanine nucleotide exchange factor (ROPGEF4), respectively, both of which have been shown to be localized at the microtubule-depleting plasma membrane domains (Oda and Fukuda 2012). These findings suggest that the spacing between cortical microtubule bundles is delimited by the microtubule-depleting plasma membrane domains containing the ROP11/MIDD1/Kinesin13A complexes, which control the patterns of cortical microtubule bundles and the subsequent patterns of secondary wall deposition (Fig. 2C). Transcriptional Regulation of Secondary Wall Biosynthesis and Deposition Because secondary walls are deposited after cessation of cell elongation in secondary wall-forming cells, genes encoding the suite of catalytic enzymes required to produce secondary wall polymers need to be co-ordinately turned on temporaly and spatially. This co-ordinate activation of secondary wall biosynthetic genes is controlled by a transcriptional network encompassing secondary wall NAC and MYB master switches and their downstream transcription factors (Zhong et al. 2010a) (Table 1; Fig. 3). In this transcriptional network, secondary wall NAC master switches (SWNs) are the top-level master switches that are capable of turning on the entire secondary wall biosynthetic program. In Arabidopsis, 10 SWNs (VND1– VND7, SND1, NST1 and NST2), which belong to the same subgroup of the NAC family, regulate secondary wall biosynthesis in various secondary wall-forming cell types. VND1–VND7 are specifically expressed in vessels, and their dominant repression causes a loss of secondary wall thickening in vessels (Kubo et al. 2005, Yamaguchi et al. 2008, Zhou et al. 2014). SND1, NST1 and NST2 are expressed specifically in xylary fibers and extraxylary fibers in stems, roots and siliques. Simultaneous mutations of SND1, NST1 and NST2 cause a complete loss of secondary wall thickening in fibers (Zhong et al. 2006, Mitsuda et al. 2007, Zhong et al. 2007b, Mitsuda and Ohme-Takagi 2008, Zhong and Ye 2014). NST1 and NST2 are also expressed in anther endothecium cells, and simultaneous mutations of NST1 and NST2 led to a loss of secondary wall thickening in endothecium cells and an anther indehiscence phenotype (Mitsuda et al. 2005). Overexpression of any of the SWNs induces the expression of the biosynthetic genes for cellulose, xylan and lignin, and concomitantly results in ectopic deposition of secondary walls in normally parenchymatous cells. Furthermore, expression of any of the SWNs driven by the SND1 promoter in the snd1 nst1 double mutant rescues the secondary wall defect in fibers (Zhong et al. 2010c, Yamaguchi et al. 2011, Zhou et al. 2014). SWN homologs exist in all taxa of vascular plants, ranging from seedless vascular plants to gymnosperms and angiosperms (Zhong et al. 2010a). Functional characterization of SWN homologs from Populus, Eucalyptus, maize, rice, Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 Brachypodium, Medicago and spruce has revealed that similarly to Arabidopsis SWNs, they are capable of activating secondary wall biosynthetic genes leading to ectopic secondary wall deposition when overexpressed, causing a reduction in secondary wall thickening when dominantly repressed, and/or restoring secondary wall thickening in fibers of the snd1 nst1 double mutant when expressed under the SND1 promoter (Zhong et al. 2010a, Zhong et al. 2010b, Zhao et al. 2010, Zhong et al. 2011a, Zhong et al. 2011b, Valdivia et al. 2013, Yoshida et al. 2013). In Populus and rice, some of the SWN homologs have alternative splicing forms, resulting in truncated SWNs without the carboxyl activation domain that may serve as negative regulators of SWN functions (Q. Li et al. 2012, Zhao et al. 2014). These findings indicate that vascular plants have evolved to employ SWNs as master switches to regulate secondary wall biosynthesis (Fig. 3). SWNs have been shown to activate a battery of downstream transcription factors, including SND2, SND3, MYB20, MYB42, MYB43, MYB46, MYB52, MYB54, MYB58, MYB63, MYB69, MYB83, MYB85, MYB103 and KNAT7 (Zhong et al. 2008) (Table 1; Fig. 3). Among them, MYB46 and its close homolog, MYB83, act as the second-level master switches regulating secondary wall biosynthesis (Zhong et al. 2007a, McCarthy et al. 2009). MYB46 and MYB83 are expressed in both vessels and fibers, and their dominant repression or RNA interference inhibition results in a severe reduction in secondary wall thickening in fibers and vessels and a concomitant collapsed vessel phenotype. Simultaneous mutations of MYB46 and MYB83 lead to a loss of secondary wall thickening in vessels and an arrest of plant growth at the seedling stage (McCarthy et al. 2009). Similar to SWNs, MYB46 and MYB83 are also able to induce the expression of the biosynthetic genes for cellulose, xylan and lignin, thus causing ectopic deposition of secondary walls (Zhong et al. 2007a, Ko et al., 2009, McCarthy et al. 2009). MYB46/83 close homologs from gymnosperms and angiosperms have been demonstrated to be functional orthologs of Arabidopsis MYB46/83 (Patzlaff et al. 2003, Goicoechea et al. 2005, McCarthy et al. 2010, Zhong et al. 2010a, Zhong et al. 2013), indicating that the recruitment of MYB46/83-like transcription factors as second-level wall MYB master switches is also evolutionarily conserved at least in gymnosperms and angiosperms. Another MYB transcription factor, MYB26, may function as a master switch regulating secondary wall biosynthesis in anther endothecium because its mutation causes a loss of secondary wall thickening in anther endothecium and an anther indehiscence phenotype, whereas its overexpression results in ectopic deposition of secondary walls. It is suggested that MYB26 acts upstream of NST1 and NST2 in regulating secondary wall biosynthesis in endothecium (Yang et al. 2007). A number of SWN- and MYB46/83-regulated downstream transcription factors have been shown to play important roles in regulating secondary wall biosynthesis (Table 1; Fig. 3). Arabidopsis MYB58, MYB63, MYB85, BLH6 and their close homologs in other species are specific activators of the lignin biosynthetic pathway (Zhong et al. 2008, J. Zhou et al. 2009, Cassan-Wang et al. 2013, Hirano et al. 2013b), LBD18 and Fig. 3 Diagram of transcriptional regulation of the secondary wall biosynthesis program in Arabidopsis. Secondary wall NAC and MYB master switches bind to and activate the SNBE and SMRE sites, respectively, in the promoters of their targets, and, together with their downstream transcription factors, activate the expression of genes involved in secondary wall biosynthesis and deposition, leading to secondary wall formation. VNDs also control the programmed cell death in vessels. LBD30 are involved in a positive feedback loop to regulate VND7 expression (Soyano et al. 2008), XND1 is probably a negative regulator of secondary wall biosynthesis and programmed cell death (Zhao et al. 2008), KNAT7 is proposed to be a negative regulator of secondary wall biosynthesis (E. Li et al. 2012), and MYB4, MYB7, MYB32 and their close homologs in other species are probably negative regulators of lignin biosynthesis (Sonbol et al. 2009, Fornale et al. 2010, Legay et al. 2010, Shen et al. 2012). Other transcription factors, such as SND2, SND3, MYB52, MYB54, MYB69 and MYB103, might be involved in fine-tuning the transcriptional regulation of secondary wall biosynthesis because their dominant repression causes a reduction in secondary wall thickening but their overexpression does not result in ectopic deposition of secondary wall components (Zhong et al., 2008). SWNs bind to a 19 bp imperfect palindromic consensus sequence, (T/A)NN(C/T)(T/C/G)TNNNNNNNA(A/C)GN(A/C/ T)(A/T), which is named secondary wall NAC-binding element (SNBE), and thereby activate their target gene expression (Zhong et al. 2010c). MYB46/MYB83 bind to a 7 bp secondary wall MYB-responsive element (SMRE) that has a consensus 207 R. Zhong and Z.-H. Ye | Secondary cell walls sequence of ACC(A/T)A(A/C)(T/C) (Zhong and Ye 2012). The SMRE consensus sequence has eight variants, and three of them are identical to the AC elements, which is congruent with the early findings that pine and Eucalyptus MYB46 orthologs bind to the AC elements (Patzlaff et al. 2003, Goicoechea et al. 2005). Another study showed that the MYB46-binding element (namely M46R) is an 8 bp core motif, (T/C)ACC (A/T)A(A/C)(T/C) (Kim et al. 2012a), which is identical to SMRE except for having an additional nucleotide at the 50 end. SWN orthologs and MYB46/MYB83 orthologs from other plant species also bind to SNBE and SMRE, respectively (Zhong et al. 2011a, Zhong et al. 2011b, Valdivia et al. 2013, Zhong et al. 2013, Yoshida et al. 2013), further demonstrating the evolutionary conservation of SWNs and MYB46/MYB83like secondary wall MYBs in regulating secondary wall biosynthesis in vascular plants. Direct target analyses have revealed that SWNs and MYB46/MYB83 are able to activate directly the expression of not only a suite of downstream transcription factors but also a number of secondary wall biosynthetic genes (OhashiIto et al. 2010, Yamaguchi et al. 2011, Zhong and Ye 2012, Zhong et al. 2010c, Kim et al. 2012b, Kim et al. 2014). In addition, SWNs directly activate many genes involved in cell wall modification and programmed cell death. These findings provide important insights into the underlying mechanism of the transcriptional network regulating secondary wall biosynthesis, suggesting that it does not utilize a simple linear regulatory cascade; instead it employs a feed-forward loop regulatory structure in which one transcription factor regulates another and they together regulate downstream targets (Fig. 3). In this network, SWNs directly activate MYB46/ MYB83 and they together directly activate other transcription factors, such as KNAT7, and some biosynthetic genes for xylan and lignin. Similarly, MYB46/MYB83 directly activate MYB58/MYB63 and they together directly regulate the expression of lignin biosynthetic genes. This multileveled feed-forward loop regulatory structure is adopted by plants probably to ensure the efficient activation and regulation of secondary wall biosynthetic genes during the development of secondary wall-forming cells. The ectopically deposited secondary walls induced by overexpression of SWNs and secondary wall MYB master switches form specific patterns similar to those of tracheary elements (Kubo et al. 2005, Mitsuda et al. 2005, Zhong et al. 2006, Zhong et al. 2007a) (Fig. 2A), indicating that they also regulate patterned deposition of secondary walls. Indeed, several genes known to be involved in cortical microtubule organization, oriented deposition of cellulose microfibrils and patterned deposition of secondary walls, including MAP65-8 (ZeMAP65-1 ortholog), FRA1, MIDD1 and ROP11, are induced by SWNs (Zhong et al. 2010c, Yamaguchi et al. 2011, Oda and Fukuda 2012). ROP11 is also a direct target of MYB46 (Zhong and Ye 2012). It is apparent that SWNs and secondary wall MYB master switches activate genes involved in both biosynthesis and patterned deposition of secondary walls. Furthermore, VNDs also regulate the expression of genes involved in programmed cell death in tracheary 208 elements (Ohashi-Ito et al. 2010, Zhong et al. 2010c, Yamaguchi et al. 2011). Concluding Remarks Secondary wall formation is a complex process involving signaling that triggers the secondary wall biosynthetic program, transcriptional activation of secondary wall biosynthetic genes, biosynthesis of secondary wall components, targeted transport of enzymes and secondary wall materials, and patterned deposition and assembly of secondary walls. Despite the tremendous progress made toward identification and functional characterization of genes involved in secondary wall biosynthesis, a number of aspects regarding secondary wall formation remain to be investigated and a few of them are highlighted below. First, secondary wall deposition is initiated in secondary wall-forming cells after cessation of elongation, but signals that trigger the secondary wall biosynthetic program have not been well characterized. Identifying such signals will be of significance in dissecting the molecular mechanisms underlying wood formation and have important implications in tree biotechnology. Auxin, an important signal for initiation of differentiation of secondary wall-forming cells, may serve as a signal for initiation and/or maintenance of the secondary wall biosynthetic program since a reduction in auxin transport results in a defect in secondary wall thickening in fibers (Zhong and Ye 1999, Zhong and Ye 2001, Ranocha et al. 2013). Secondly, cortical microtubules and the actin cytoskeleton are important for trafficking of vesicles carrying CesA complexes (Wightman and Turner 2008) and hemicelluloses to the plasma membrane regions where secondary wall thickening occurs, but proteins involved in vesicle trafficking for secondary wall formation are largely unknown. The EXO70A1 gene, which encodes a subunit of the exocyst complex, is proposed to function in vesicle trafficking during vessel differentiation (Li et al. 2013). The exo70a1 mutant has irregular secondary wall thickening, incomplete perforation plates and large vesicular compartments in vessels. Phosphoinositides, which are important for vesicle trafficking and actin cytoskeleton organization in yeast and animals (Takenawa and Itoh 2001), may also play crucial regulatory roles in vesicle trafficking of secondary wall materials. Mutations of two phosphoinositide phosphatase genes, FRA3 and AtSAC1 (FRA7), result in altered actin organization and aberrant deposition of secondary walls (Zhong et al. 2004, Zhong et al. 2005a). RHD3, a GTPase required for tubular endoplasmic reticulum generation and Golgi distribution (Chen et al. 2011), is essential for normal actin organization and secondary wall deposition, probably also through regulating vesicle trafficking of secondary wall materials (Hu et al. 2003). Further deciphering the exocytosis machineries involved in the targeted transport of secondary wall materials will help dissect the mechanisms controlling the patterned deposition of secondary walls. Thirdly, secondary wall composition varies among different plant species, in different cell types within the same species and may change in response to developmental and environmental stimuli. With the increased understanding of genes Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140 involved in the biosynthesis of secondary wall components and its transcriptional regulation, tools are now available to investigate the molecular mechanisms controlling secondary wall heterogeneity. Fourthly, secondary walls are deposited in specific patterns in xylem cells during different developmental stages, e.g. an annular or helical pattern in protoxylem and a reticulated or pitted pattern in metaxylem. Signals that trigger the developmental changes of the patterned secondary wall deposition and of the underlying cortical microtubule organization are yet to be identified. Similarly, little is known about the molecular switches directing the oriented deposition of cellulose microfibrils in the S1, S2 and S3 layers of secondary walls. Finally, many additional secondary wall-associated genes have been identified through genomic and co-expression analyses (Ko et al. 2009, Wilkins et al. 2009, Ohashi-Ito et al. 2010, Yamaguchi et al. 2011, Zhong et al. 2010c, Zhong and Ye 2012, Hirano et al. 2013a) and the functions of most of them are unknown. Functional characterization of these genes may provide new insights into the molecular mechanisms of secondary wall formation. Identification and functional characterization of all players participating in the formation of secondary walls, the major constituents of plant biomass, is not only important for basic plant biology but also provides a knowledge foundation for better utilizing plant biomass for various human needs. Funding The work in the authors’ lab was funded by the Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the US.Department of Energy [DEFG02-03ER15415]; the US Department of Agriculture National Institute of Food and Agriculture [AFRI Plant Biology program (#2010-65116-20468)]; the National Science Foundation [ISO-1051900]. Acknowledgments We regret that many original articles on secondary wall biosynthesis could not be cited due to space limitation. Disclosures The authors have no conflicts of interest to declare. References Alejandro, S., Lee, Y., Tohge, T., Sudre, D., Osorio, S., Park, J. et al. 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