|UFDC Home||myUFDC Home | Help|
This item has the following downloads:
1 FUNCTIONAL ANALYSIS OF THE CELLULOSE SYNTHASE GENE FAMILY IN MAIZE: FROM BIOINFORMATICS TO ASSESSMENTS AT WHOLE PLANT, TISSUE, CELL, AND PROTOPLAST LEVELS By A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2011
3 To my grandfather, S. Allen Poole, for his love, guidance, and undying support of my acade mic endeavours
4 ACKNOWLEDGMENTS I thank my advisor, Karen Koch, and supervising committee Drs., Ken Boote, Alice Harmon, Gary Peter, and Wilfred Vermerris for their guidance and scientific in sight throughout my graduate ca reer. I also thank my labmates (past and present) and our lab manager Wayne Avigne, for all of their help and support throughout my time in graduate school. Finally, I thank my family and friends for their support and encouragement.
5 TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................ ................................ ................................ ............... 4 LIST OF TABLES ................................ ................................ ................................ ........................... 7 LIST OF FIGURES ................................ ................................ ................................ ......................... 8 ABSTRACT ................................ ................................ ................................ ................................ ... 10 CHAPTER 1 INTRODUCTION ................................ ................................ ................................ .................. 12 2 ANALYSIS OF THE CELLULOSE SYNTHASE GENE FAMILY IN MAIZE ................. 24 Background ................................ ................................ ................................ ............................. 24 Results ................................ ................................ ................................ ................................ ..... 28 Identificat ion and Characterization of CesA Paralogs ................................ ..................... 28 Evolution of CesA Genes in Maize ................................ ................................ ................. 29 Micro RNA Targets in the CesA Gene Family ................................ ............................... 29 Expression of the CesA Gene Family During Development ................................ .......... 30 Discussion ................................ ................................ ................................ ............................... 32 Bioinformatic Analysis Reveals Additional CesA Paralogs ................................ ............ 32 Phylogenetic Analysis of the CesAs ................................ ................................ ................ 33 Evolution of the CesA Gene Family ................................ ................................ ................ 34 Potential for Micro RNAs to Affect Differential Regulation of CesA Genes ................. 35 Expression of CesAs Varies with Tissue and Development ................................ ............ 36 Cluster Analysis of Gene Expression Among the CesAs ................................ ................ 36 Methods ................................ ................................ ................................ ................................ .. 37 Bioinformatic Analysis of the Maize Genome for Identification of CesA Paralogs ....... 37 Phylogenetic Analysis of the CesA Family ................................ ................................ ..... 38 Evolutionary Analysis of the CesA Family ................................ ................................ ..... 38 Prediction of miRNA Target Sites in the CesAs ................................ ............................. 38 Plant Material ................................ ................................ ................................ .................. 39 Isolation of RNA and cDNA Synthesis ................................ ................................ ........... 39 Real Time Quantitative RT PCR ................................ ................................ ..................... 40 Ph loroglucinol Staining ................................ ................................ ................................ ... 40 3 EXPRESSION OF THE CELLULOSE SYNTHASE GENE FAMILY IN A MAIZE SUSPENSION CELL/PROTOPLAST SYSTEM ................................ ................................ .. 61 Background ................................ ................................ ................................ ............................. 61 Results ................................ ................................ ................................ ................................ ..... 65 Establishment of the Protoplast Regeneration System ................................ .................... 65 Expression of CesAs and Sucrose Synthases in Protoplasts Regenerating Cell Walls ... 66
6 CesA and Sucrose Synthase Expression During Cell Wall Regeneration With Perturbations in Sucrose Concentration ................................ ................................ ....... 67 Cellulose Synthase and Sucrose Synthase Expression in Response to Simulated Wounding/Pathogen Infection ................................ ................................ ..................... 68 Expression of the CesAs During Induction of Secondary Cell Wall Biosynthesis ......... 69 Discussion ................................ ................................ ................................ ............................... 70 Cellulose Synthase Expression During Cell Wall Regeneration ................................ ..... 70 Expression of CesAs During Cell Wall Regeneration Varied with Sucrose Supply ....... 71 Cellulose Synthase Expression in Response to Simulated Wounding/Pathogen Infection ................................ ................................ ................................ ....................... 72 Cellulose Synthase Expression During Induction of Secondary Cell Wall Synthesis .... 73 Methods ................................ ................................ ................................ ................................ .. 74 Generation of the Suspension Cell Line ................................ ................................ .......... 74 Protoplast Isolation, Regeneration, and Characterization ................................ ............... 75 Simulation of Wounding/Pathogen Infection ................................ ................................ .. 76 Induction of Secondary Cell Wall Biosynthesis ................................ .............................. 76 Quantitative Real Time RT PCR ................................ ................................ ..................... 77 4 CHARACTERIZATION OF MAIZE CESA MUTANTS FROM THE TRANSPOSON MUTAGENIC UNIFORMMU POPULATION ................................ ................................ .... 86 Background ................................ ................................ ................................ ............................. 86 Results ................................ ................................ ................................ ................................ ..... 89 Discussion ................................ ................................ ................................ ............................... 91 Methods ................................ ................................ ................................ ................................ .. 93 Plant Material ................................ ................................ ................................ .................. 93 Genotyping ................................ ................................ ................................ ...................... 93 5 SUMMARY AND FUTURE DIRECTIONS ................................ ................................ ......... 98 APPENDIX A FORWARD GENETICS ANALYSIS OF THREE MAIZE MUTANTS ........................... 104 B SEQUENCES OF GENE SPECIFIC PRIMERS USED FOR QRTPCR ANALYSIS ....... 110 LIST OF REFERENCES ................................ ................................ ................................ ............. 111 BIOGRAPHICAL SKETCH ................................ ................................ ................................ ....... 129
7 LIST OF TABLES Table page 2 1 Tissues sampled at key developmental stages ................................ ................................ ... 42 4 1 Cellulose synthase mutants identified in the UniformMu population for reverse genetics screening. ................................ ................................ ................................ ............. 94 4 2 Sites of maximum expression for the maize CesAs at each developmental stage. ............ 95
8 LIST OF FIGURES Figure page 2 1 Diagram of a canonical CesA protein. ................................ ................................ ............... 43 2 2 Unrooted, neighbor joining phylogenetic tree of the maize CesA family. ........................ 44 2 3 Predicted cDNA sequence of CesA7 a and its translated protein sequence. ..................... 45 2 4 Approximate distribution of the CesA family and close paralogs within the maize genome. ................................ ................................ ................................ .............................. 47 2 5 Predicted miRNAs targeting each of the maize CesAs .. ................................ .................... 49 2 6 Cluster analysis of potentially CesA targeting miRNAs.. ................................ ................. 50 2 7 Heat maps representing expression of CesA family members. ................................ .......... 51 2 8 Relative expression of the CesA family in all tissues samples. ................................ ........ 52 2 9 Expression of CesAs in tissues with low mRNA abundance.. ................................ ........... 54 2 10 Cluster analysis of the CesA family at the seedling and vegetative stages.. ................... 56 2 11 Cluster analysis of the CesA family at anthesis/reproductive maturity.. ........................... 58 2 12 Lignin and polyphenol deposition in tissues at the anthesis stage (72 DAG).. ................. 59 2 13 Expression of the CesaA gene family in kernel tissues ................................ ..................... 60 3 1 Diagram of the protoplast regeneration system.. ................................ ............................... 78 3 2 Relative expression of select CesA family members and sucrose synthases during cell wall regeneration by protoplasts.. ................................ ................................ ...................... 79 3 3 Sugar responsive expression of the CesA family during cell wall regeneration by protoplasts. ................................ ................................ ................................ ......................... 80 3 4 Expression of the systemic wound/pathogen response gene Maize Proteinase Inhibitor ( MPI ) ................................ ................................ ................................ ................... 81 3 5 Relative expression of CesA genes and the systemic wound response gene MPI (Maize Proteinase Inhibitor) in response to simulation of wounding and pathogen infection.. ................................ ................................ ................................ ........................... 83 3 6 Differentiation of suspension cells to tracheary element like cells after secondary cell wall induction.. ................................ ................................ ................................ ................... 84
9 3 7 Relative expression of CesA gene family members during secondary cell wall biosynthesis. ................................ ................................ ................................ ....................... 85 4 1 Unrooted, neighbor joining, interspecies phylogenetic tree with descriptions of cesA mutant phenotypes.. ................................ ................................ ................................ ........... 96 4 2 The mutant phenotype possibly associated with CesA11 ................................ ................. 97 A 1 The maize shredded mutant 05S 2500 under field conditions.. ................................ ...... 107 A 2 The shredded phenotype.. ................................ ................................ ................................ 108 A 3 Characteristics of the empty pericarp UF ( epuf1 ) mutant line 09S 3300 ....................... 109
10 Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy FUNCTIONAL ANALYSIS OF THE CELLULOSE SYNTHASE GENE FAMILY IN MAIZE: FROM BIOINFORMATICS TO ASSESSMENTS AT WHOLE PLANT, TISS UE, CELL, AND PROTOPLAST LEVELS By December 2011 Chair: Karen Koch Major: Plant Molecular and Cellular Biology primary cell wall of almost every plant cell. Cellulose is also central to formation of secondary cell walls in vascular tissues and wood. Since the function of cellulose in these walls varies with tissue and development, we hypothesized that these roles would be reflected in differential regulatio n of cellulose synthase genes. We utilized four approaches to test this hypothesis in the diverse cellulos e synthase gene family of maize, a species critical to human needs for food, fuel, and fiber. First, bioinformatic analysis of the cellulose synthase ( CesA ) gene family revealed several, potentially functional paralogs in this ancient tetraploid. In most i nstances, paralogs mapped to chromosomal segments associated with enhanced gene loss, supporting an emerging model for one subgenome remaining resistant to gene loss after a genome duplication event. Second, we quantified expression of 13 CesAs at cell p rotoplast and tissue levels. In suspension cultures and in planta three CesA family members (C esA10, CesA11, and CesA12) showed consistent, exclusive co expression. Each instance of their coordinate, elevated mRNA levels coincided with sites and/or timi ng of enhanced lignin deposition, consistent with
11 involvement of these genes in secondary cell wall biosynthesis. Other showed coordinate regulation among some family members, but composition of these subgroups changed during development. Third, we characterized CesA responses to induction of secondary cell wall biosynthesis in suspension cells. Expression of most CesAs was not altered, but the upregulation of C esA10, CesA11, and CesA12 with onset of secondary cell wall deposition supported their contribution to this process. For our fourth approach, we employed co segregational analysis to determine if cesA mutants were associated wi th visible phenotypes in maize. No such phenotypes wer e evident for homozygous mutations in any of the seven CesA family members or double mutant examined, indicating significant functional redundancy within the CesA family. Collectively, results support a model in which three CesAs ( CesA10, CesA11, and CesA12 ) predominate in deposition of secondary cell walls of maize, and where other C esAs function in primary wall formation through combinations of co expressed CesA subgroups that vary with given tissues and stages of development.
12 CHAPTER 1 INTRODUCTION The plant cell wall is a complex structure constructed predominantly from polysacchar ides such as cellulose, pectins, and hemicellulos es In addition to these various carbohydrate constituents, the cell w all also includes proteins, and in some instances, lignin, cutin and suberin (Fry, 2004) The cell wal l protects the cell from damage an d pathogens, provides mechanical support for the cell and plant and ultimately determines plant shape and size through its role in cell expansion. Furthermore, the endosperm cell walls in many species act as storage site s for re mobilizable, hemicellulosi c polysaccharides (Reid and Edwards, 1995 ; Olsen, 2007 ). Cell walls are classified as either primary or secondary, and primary wall s are further defined as type I or t ype II (Schaffner and Sheen, 1991) Primary cell walls are those that are initially laid down around the plasma membrane Their expansion allows plant cells to increase in size as they grow In some cell types, secondary layers of cell wall are deposited inside the primary wall once cell growth has stopped These layers can be distinguis hed by the pitch, or angle, of the cellulose microfibrils deposited within. S econdary cell walls have greater mechanical strength tha n primary cell walls, and are often associated with xylem vessels and woody tissue s rich in lignin ( Boerjan et al 2003 ) Secondary cell wall composition typically differs from that of the primary wall in that it often contain s substantial amounts of lignin (which adds strength) and waxy substances such as cutin and suberin. Among primary cell walls, the type I are more taxonomically widespread than the t ype II and are standard in gymnosperms, dicots, and many monocots. Type II cell walls occur only in the commeli noid monocots, which include grasses gingers, and bromeliads (Schaffner and Sheen, 1991 ; Carpita, 1996 ; Yokoyama and Nishitani, 2004 ) The main difference between these
13 two types of primary cell wall s is their chemical composition. Although cellulose is the most abundant component in both types, it is cross linked primarily with xyloglucans in t ype I walls and glucuronoarabinoxylans in t ype II walls (Carpita, 1996) In addition, t ype I walls contain considerably more pectin and structural protein than do type II walls (Carpita, 1996). Cellulose is the most abundant component in the cell wall and is largely responsible for its mechanical strength (Saxen a and Brown, 2005). Even in woody tissues and cells with abundant lignin, it is the cellulose infrastructure that provides a significant p ortion of the strength and flexi bility to the cell wall. The importance of cellulose in providing durability to cell walls is apparent in the brittle culm mutants of barley and rice, in which tissues are easily broken due to a deficiency of cellulose, even though lignin levels are unchanged or elevated (Tanaka et al 2003; Bu rton et al., 2010). This polysaccharide is present a s structural units comprised of 36 individual linear chains o f linked D Glucose that combine to form a single microfibril (Delmer and Amor, 1995) The individual glucan chains can range from 8 000 units long in primary cell walls to 15 000 units in secondary cell walls (Pear et al., 1996; Brett, 2000; Brown, 2004) In addition individual glucan chains overlap ( with regard to chain ends) within a microfibril (Carpita, 1996). The orientation of deposition for these microfibrils determines the direction that the cell can elongate (Saxen a and Brown, 2005) thus regulation of cellulose deposition is central to plant architecture. Cellulose is a semi crystalline molecule that can be polymorphic (Saxen a and Brown, 2005) The crystal state of cellulose depends on the positioning of the individual glucan chains relative to one another. Cellulose I, which is found in plants (as opposed to synthetic cellulose II) consists of parallel glucan c hains pa cked side by side. The two crystalline configurations of cellulose I are I and I The cellulose molecules in both configurations show the same skeletal conformation, but their hydrogen bonding patterns differ (Atalla and VanDerhart, 1989;
14 Nishiyama et al ., 2003) This results in each type having a different lattice structure (Atalla and VanDerhart, 1989). Plant cellulose is generally richer in cellulose I whereas the cellulose of more primitive organisms has more of the I configuration Chemical pertur bation experiments have shown that cellulose I is the more stable form, but microfibrils can contain both types and the ratio of each configuration can influence physical properties of cellulose (Atalla and VanDerhart, 1989; Saxen a and Brown, 2005). Cellulose crystallization is believed to occur at the same time the strands are synthesized in the rosette (Herth, 1983). This mode of action is supported by a correlation between the size of rosette arrays and the microfibrils they form Whereas individua l rosettes in plants yield 36 chain fibrils, the algae Micrasterias produce s much larger fibrils from arrays of up to 175 rosettes (Giddings et al., 1980). Additionally, the radial swelling 1 1 mutant in Arabidopsis is a temperature sensitive CesA mutant t hat, upon exposure to nonpermessive temperature, shows dissociation of individual rosettes and production of amorphous, noncrystalline cellulose (Arioli et al., 1998). Mechanisms underlying these configurations of cellulose in plants and their balance in a given microfibril have remained a subject of intensive interest for many years ( Vietor et al, 2002; Nishiyama et al., 2002 and 2008; Tanaka and Iwata, 2006) The first successful investigation into the biochemical mechanism of biological cellulose synthesis demonstrated that a lyophilized preparation from the cellulose producing bacteria Acetobacter xylinum could produce cellulose in the presence of glucose and oxygen (Hestrin and Schramm, 1954). A few years later the substrate required for cellul ose synthesis by this sam e system was specifically determined to be UDP glucose by providing different forms of C 14 glucose (Glaser, 1957). These results were also supported by the observation that A. xylinum mutants lacking capacity to produce UDP glucose were also cellulose deficient (Valla et. a l., 1989). Although much subsequent effort focused on elucidating mechanism s of cellulose
15 biosynthesis, progress has been slow due to the lability of membrane protein complexes the difficu lty associated with characterizing them and the extent of required cofactors. Furthermore, and perhaps more importantly, callose contamination of preparations to be used for study of cellulose synthesis has remained a problem for many years The basis for this difficulty included the similar ity in structures of callose and cellulose (callose being 1 D glucose and cellulose being 1 D glucose), production of both compounds at the plasma membrane, and synthesis of both from the same substrate (Pear et al., 1995). The enzyme responsible for providing this UDP glucose substrate remained elusive for some time, however sucrose synthase was considered a strong candidate due to its likelihood of contributing the same substrate for callose synthesis ( Fros t et al., 1990; Nolte and Koch, 1993). Further investigation by Amor et al. (1995) initially indicated that as much as 50% of sucrose synthase could be located at the plasma membrane, and that this immunolocaliz ed to helical arrays with patterns that paralleled cellulose deposition (Amor et al., 1995). Similar observations were reported for dense localization of sucrose synthase in the plasma membrane at sites of cellulose deposition in tips of rapidly growing cotton trichomes (Nolte et al., 1995) and at points of wall thickening in hypoxic wheat roots (Alberecht and Mustroph, 2003). The shrunken1 maize mutant, which is deficient in an endosperm specific sucrose synthase, lacks cell walls in its interior consistent with a possible role of sucrose synt hase in their formation (Chourey et al., 1991). Only recently has evidence emerged that directly links sucrose s ynthase with cellulose synthase machinery. Work by Fujii et al. (2010) has demonstrated through immunogold labeling with sucrose synthase antibo dies that sucrose synthase can bind to rosettes in vitro and synthesize cellulose, thus supporting previous observations that sucrose synthase is likely essential for cellulose production in planta
16 Cellulose synthase was first identified as an 83 kD poly peptide using a product entrapment approach in A cetobacter xylinum (Lin and Brown, 1989). This cellulose synthase gene was subsequently cloned (Saxena et al., 1990). Concurrent, yet independent experiments involving complementation of a cellulose deficient A. xylinum mutant led to the cloning of a four gene operon, with one gene ( BcsA ) being homologo us to the cellulose synthase cloned by Saxena et al. (1990) (Wong et al., 1990). Another cellulose synthase gene ( CelA ) was then cloned from Agrobacterium tumefaciens in 1995 (Matthysse et al., 1995), but identification of cellulose synthase genes in plants proved more challenging Most researchers sought BcsA/CelA homologs in plants using the bacterial BcsA gene as a probe. This approach was not successful because cellulose synthases are now known to have significant variation in sequence outside key domains conserved in processive glycosyltransferases (Saxena et al., 1994; Delmer and Amor, 1995). The first plant cellulose synthase ( CesA ) wa s cloned in cotton by sequencing a cDNA library from cotton fibers at a stage of maximum secondary cell wall deposition (Pear et al., 1996) Identification of t wo highly conserved genes having regions of high homology to the bacterial CelA genes led to the initial conclusion that these were indeed cellulose synthases. Since this discovery, the release of sequenced genomes and databases of gene expression during development al progression has led to identification of no fewer than 10 CesA genes in Arabidopsis and at least 12 in maize (Holland et al., 2000). T he presence of orthologous CesA genes ha s also been confirmed in other species with sequenced genomes, such as rice and poplar ( Burton et al., 2005) Structural d ata indicate that the CesA genes code for i ntegral membrane proteins that assemble into rosettes This association was first demonstrated in a temperature dependent Arabidopsis cesA mutant called radial swelling 1 1 that showed
17 disassembly of individual rosettes and production of noncrystalline cellulose at high temperatures ( Arioli et al 1998; Taylor, 2008) Cellulose synthesizing protein complexes were first observed at the ends of microfibrils in algae by imaging freeze fractured membranes (Giddings et al., 1980). This highly ordered, six these have since been observed in high concentration at sites of rapid cellulose synthesis in vascular plants (Herth, 1985). The cellulose synthase complex is only part ially exposed on the extracellular side of the plasma membrane, with a more substantial portion exposed to the cytoplasm (Nuhse et al., 2004; Taylor, 2008). Work also suggests that the cellulose synthase complex is associated with other essential molecular factors (Richmond and Somerville, 2000; Saxena and Brown, 2005; Taylor, 2008). These are hypothesized to include proteins tha t affect organization of the complex, as well as catalyze crystallization of glucan chains, and aid transfer of UDP glucose (the s ubstrate of cellulose synthase) to the catalytic sites ( Albersheim et al., 1999 ; Saxena and Brown, 2005; Fuji et al., 2010 ). Cellulose synthase has been observed, in vitro to processively (without stopping or detaching) catalyze the polymerization reactio n in one step (Richmond and Somervill e, 2000; Saxena and Brown, 2005; Taylor, 2008). Previous research has shown that different groups of CesA genes are expressed in cells synthesizing primary cell walls compared to cells synthesizing the cellulose in secondary cell walls (Taylor et al., 2003; Appenzeller et al., 2004 ; Saxena and Brown, 2005 ; Persson et al., 2007 ). Furthermore, mutant analysis in Arabidopsis has shown that expression of three different CesA genes is required to form a functional rosette in both primary and secondary cell walls (Burn et al, 2002; Zhong et al, 2003; Taylor et al., 2003; Tanaka et al., 2003). These observations ha ve led to the hypothesis that three distinct, non redundant cellulose synthases are required to form a functional rosette (Taylor et al., 2003; Saxena and Brown, 2005 ; Persson et
18 al., 2007 ; Jiang et al., 2008) ). Although a 3:2:1 stoichiometry of CesA isof orms has been depicted in models representing the heterohexameric CESA complex, there has been no direct evidence to support this specificity in balance of subunits. Studies of cellulose synthesis in plants suggest that initiation of this process may req uire a sitosterol glucoside primer to initiate glucan chain elongation (Peng et al., 2002 ; Endler and Persson, 2011 ). Data indicate that sitosterol glucoside s are synthesized on the cytosolic face of the pl asma membrane (Cantatore et al. 2000) which is consistent with work showing a plasma membrane association for the enzyme responsible for its synthesis UDP Glc:sterol glucosyltransferase (Elbein and Forsee, 1975) Furthermore, cotton fiber membranes can synthesize sitosterol cellodextrins in vivo when supplied with UDP glucose (Peng et al., 2002). Once cellulose synthesis has been initiated, current models suggest the sitosterol is cleaved from the nascent cellulose chain by a specific membrane bound cellulase calle d KORRIGAN This hypothes is is supported by the accumulation of lipid linked cellodextrins in Arabidopsis korrigan mutants (Sato et al., 2001). Furthermore, cellulose synthase activity is enhanced in naturally occurring detergent resistant membranes (Bessueille et al., 2009) The se sections of membrane have especially high levels of sitosterol glucoside, and are similar to lipid rafts in animals because they have altered lipid composition ( Lingwood and Simons, 2007 ). Although there has been much progress in understanding the mechanism of cellulose synthesis, several significant aspects of this process remain unclear One of these is the identity and stoichiometry of CesA isoforms that can assemble into CESA complexes. Another is the developmental and tissue specific regulation of CesA gene expression. It is also poorly understood how the identity and stoichiometry of CesA isoforms in a given CESA complex affects its function Still another challenge to the field is defining the presence and identity of unidentified cofactors required for assembly of complexes or their functioning These
19 unidentified cofactors are among suggested reasons for a consistent lack of success in achieving cellulose synthesis through transgenic approaches (using plant CesAs ) in non plant systems. Further research employing both genetic and biochemical approaches will be central to enhancing our understanding of how this seemingly simple polysaccharide is synthesized in plants. Work presented here employs t hree additional avenues of investigation. The first is a bioinformatic analysis of the maize cellulose syntha se family. The second combines molecular level analysis in isolated tissues, and in suspension culture and protoplast systems responding to perturbations The third extends the mutant analysis approach to the cellulose synth ase gene family in maize, a large, C4 panicoid species with immediate commercial value. S uspension cultures and protoplasts provide ideal platforms for experimental alterations Both systems can be readily perturbed and an investigator also has a choice of source tissue from which cultures or protoplasts are generated This provides a means of addressing properties and/or responses of specific cell types. These systems also offer the benefits of relative cellular uniform ity. In addition, s uspension cells an d protoplasts can be quickly regenerated, and easily perturbed Age of suspension cell line s is important, though, since these can become immortalized and perpetuated indefinitely When they do, cells dedifferentiate and their genomes undergo major epigenetic modifications (Tanurdzic et al., 2008). However when used and interpreted carefully, suspension cultures can provide a valuable avenue of inquiry. P rotoplasts are also useful and can be readily released from whole tissue s and cultured cells. They have successfully facilitated studies of diverse biological processes from cell wall regeneration (Shea et al., 1989) to C4 metabolism ( Shatil Cohen et al., 2011). Protoplasts are also ideal for genetic transformation, as the la ck of a cell wall facilitates gene transfer through electroporation and /or microinjection (Potrykus, 1991; Dong Yoo et al., 2007). Cautions in use
20 of protoplast systems include their mechanical delica cy which often result s in high mortality rates, and the ce ll wall removal process, which can activat e wounding and pathogen infection response s (Moreno et al., 2005; Walley et al., 2007). The latter can occur as a result of the mechanical stress of cell wall removal, exposure to fungal elicitors (cell wall degrading enzymes), or the presence of small oligosaccharides released from the cell wall during digestion (Cordero et al., 1994; Moreno et al., 2005; Walley et al., 2007). Changes in gene expression can also occur in respons e to sugar starvation while cells are incubating in cell wall digestion medium (Yu, 1999). The third, new avenue of investigation used here aids current research endeavors by bringing Zea m ays L. ssp mays into the realm of species with mutations altering genes for cellulose synthesis. Our current understanding of cellulose synthesis has come from characterization of mutants deficient in one or more of the cellulose synthase genes but has focused mainly on Arabidopsis rice and barley (Turner and Somerville, 1997: Tanaka et al. 2003; Taylor et al., 2003 Burton et al., 2010 ). Additional insights have also been gained through transgenic approaches in tobacco and aspen (Wu et al., 2009; Joshi et al., 2011). There are currentl y no reports associating a CesA mutation in maize with a phenotype Work here characterizes new mutations in most of the maize Ce s A gene family with the goal of determining which f amily members are essential to normal physiology, morphology, and development. The sour ce of these new maize mutants has been a transpo son mutagenic population that enables identification of insertion sites. Transposable elements provide a useful mechanism for naturally generating and characterizing mutants. These elements, or transposons, are small, mobile DNA sequences that can insert into genes often causing them to lose function. Although many transposons have the capacity to insert anywhere in the genome, the Mutator ( Mu ) class typically inserts in the regions of functional genes ( Dietrich et al., 2002 ) Since transposons
21 transposon families and subclasses have been defined. All transposable elements can be broadly Class Class (Type I), or DNA (TypeII) intermediate (Wicker, 2007) Furthermore, transposons are classified as either autonomous or nonautonomous. Autonomous transposon s are those with sequence s that code for a transposase enzyme having the capacity to mediate movement of a transposon sequence to a new location. In some instances the original sequence is excised for reinsertion, and in others, the original transposon remains in position while a copy is transposed to a new locale (Wicker et al., 2007). Nonautonomous transposons can contain as little as the essentia l cis elements needed for recogni tion and transposition by the transposase A utonomous transposons must be present in order for nonautonomous elements to transpose. Among some of the best characterized transposons in maize are the Mutator or Mu family of transposable elements. These transposons are particularly useful for genetic studies because they contain terminal inverted repeat (TIR) sequences from which primers can be designed, and sequences flanking insertion sites can be identified (Dietrich et al., 2002). This approach provides a convenient method for identifying disrupted genes and associating them with a phenotype. The Mu TIRs are generally about 215 base pairs long and are highly conserved within each of the 12 subclasses of Mu transposable e lements, termed Mu1 to Mu12. More recent work has revealed that these initial 12 sub classes account for over 300 separate Mu insertions in the B73 maize genome, and that these group into five overall Mu clades ( C. Hunter. Univ. of Florida, PhD dissertation 2010). Several research groups have used the Mutator family of transposons to generate and tag new mutations in maize (Walbot, 2000; Raizada et al., 2001; Meeley and Briggs, 1995; McCarty and Meeley, 2009; Williams Carrier et al., 2010). The
22 largest and most active at present is the UniformMu maize population developed at the University of Florida (McCarty et al., 2005; Settles et al., 2007) The UniformMu population was generated by introgressing a highly ( Mu ) transposon mutagenic line called into the maize W22 inbred Along with the active Mu transposon system, a Mu associated bz mum9 color marker was also in trogresse d into the population. This marker allows easy determin ation of whether a given plant has inh erited the transposase and can still generate new mutations. In the absence of transposase, kernels carrying bz mum9 are bronze, whereas wild type kernels are dark purple. Kernels carrying bz mum9 and an active transposase are bronze with a varying amount of small purple spots. Other benefits of the UniformMu population are its high mutation frequency, low mutant load, moderate total Mu TE copy number, a database of pedigreed insertions, and a uniform background of plant material that allows easy ide ntification of novel phenotypes (McCarty et al., 2005) Here, bioinformatic and evolutionary analyses have identified four new CesA paralogs (homologous genes arising directly f rom duplication within a genome) and shown that at least one copy of each uni que CesA is retained on chromosomal regions associated with resistance to gene degradation. Only redundant CesA paralogs were found on chromosomal regions associated with gene loss. This observation is consistent with a recent hypothesis that suggests gene loss a fter a genome duplication event such as occurred long ago in maize, will occur preferentially from that is more resistant to gene degradation (Schnable et al., 2010). Phylogenetic analysis has also revealed that one, and only one paralog was retained (and not subfunctionalized) from each of the three CesA subclades associated with primary cell wall synthesis in maize. A Gene Balance Hypothesis (Birchler and Veitia, 2010 ) suggests that this balance between gene dosage from CesA subclades may reflect a functional balance between
23 isomers in CESA heterohexamers. Gene retention and loss after polyploidization events in diverse species appears responsive to the stoichiometry o f isomers in multimeric complexes when this balance has been essential to functionality. Further support for such a relationship Arabidopsis primary cell wall biosynthesis depends on the presenc e of three different CesAs, one from each of the three sub clades associated with primary wall formation. We als o show that although expression of the CesA genes in maize is typically a coordinated process mRNA profiles show that co regulated groups of Ce sA transcripts vary across tissues and throughout development Three CesAs ( CesA10 CesA11 and CesA12 ) are consistently coexpressed at all stages of development, and are predominantly associated with tissues developing secondary cell wall s Also, these th ree genes are strong ly upregulated in suspension cells that have been hormonally induced to differentiate into tracheary element like cells rich in secondar y wall. The other, primary cell wall associated CesAs form discreet expression profiles that cluster together, but change at different stages of development. Additionally, we show that these co expression clusters observed in planta can change in response to perturbations of protoplasts and suspension cells, thus indicating plasticity in the co express ion modules and transcription level responses of CesAs Finally, mutant analysis has demonstra ted that several of the primary wall associated maize CesAs are functionally redundant, and do not cause visible phenotype s when mutated. This is likely a result of the proliferation of the maize CesA family, which would add to the high level of functional redundancy observed among the primary wall CesAs in Arabidopsis
24 CHAPTER 2 ANALYSIS OF THE CELL ULOSE SYNTHASE GENE FAMILY IN MAIZE Background Cellulose repre sents roughly 50% of total plant based biomass, making it one of the most abundant renewable resources on the planet. This biopolymer is also integral to plant structure, fiber quality, digestibility and carbon partitioning. As food and fossil fuels become limiting, a central importance emerges for better u tilizing cellulosic resources. Gaining greater understanding of the mechanisms underlying cellulose synthesis at the genetic and molecular level will provide potentially invaluable avenues for impro ving composition of cellulosic biomass (for use as bioreactor feedstock) and other cell wall based aspects of plant yield. The cellulose synthase ( CesA ) genes of plants derive from an ancient lineage, and are present in numerous taxonomically diverse orga nisms from vascular plants to algae, bacteria, protists, fungi, and urochordate animals (Richmon d, 1991; Hirose et al., 1999). Although sequences of CesAs in vascular plants do not have a high degree of overall homology, they do share several conserved reg ion s implicated in CesA function. The D,D,D,QxxRW domain (with conserved aspartate residues ) appear s essential for catalytic activity (Saxena et al., 1995; Delmer and Amor, 1995; Taylor, 2008) Also, an N terminal LIM (Lin11, Isl 1 & Mec 3) like Z inc binding domain/RING (or Cys3HisCys4 containing) domain is hypothesized to mediate protein protein interaction (Kawagoe and Delmer, 1997; Taylor, 2008 ). In addition eight transmembrane domains (two N terminal and four C terminal) allow orientation wi thin the plasma membrane (Pear et al, 1996; Taylor, 2008) ( Figure 2 1) The Zinc binding domain/RING domain and catalytic D,D,D,QxxRW domain are predicted to lie in cytoplasmic regions of the protein, and both of these domains are associated with a hypervariable region (HVR) not seen in the CesAs of bacteria or cyanobacteria (Delmer, 1999). One of these HRVs is N terminal,
2 5 positioned between the Zinc binding domain and the first two transmembrane domains. The other HVR is in the ce ntral region of the cyt oplasmic loop that contains the D,D,D,QxxRW domain ( Figure 2 1 and Taylor, 2008 ). This region of the c ytoplasmic loop is also termed t he Class Specific Region (CSR) because these sequences are conserved between related sub clades of CesAs from different species, but not among the diverse CesA family members from within the same species (Vergara an d Carpita, 2001; Taylor 2008). Mutation of residues affecting phosphorylation in the hypervariable regions has resulted in directionally asymmetric movement of rosettes, and discrepancy in the velocity of bi directional movement of these structures along cortical mic rotubules (Chen et al., 2010). This aberrant movement of rosettes leads to abnormal patterns of microfibril deposition and los s of anisotropic cell expansion (Chen et al., 2010). In contrast, rosettes in wild type plants move bidirectionally along cortical microtubules in roughly equal numbers (Paredez et al., 2006; Yoneda et al., 2007). Chen et al. (2010) thus suggest that the p hosphorylation status of CesA proteins can affect polar interaction with microtubules. Cellulose synthases are processive enzymes classified a s family 2 inverting nucleotide diphospho sugar glycosyltransfe rases (Campbell et al., 1997). Using UDP glucose a s a substrate, CesAs form (1 4) linked glucose chains estimated to range from as long as 8,000 residues in primary cell walls, up to 15,000 residues in secondary walls (Pear et al., 1996; Brett, 2000; Brown, 2004) Individual cellulo se chains can then as sociate th rough hydrogen bonds and form the highly structured, crystalline cellulose of t he microfibrils in cell walls. The CesAs function in vivo as membrane bound heterohexamers that require at least three different CesA isoforms to assemble (Taylor et a l, 2003). Research using an Arabidopsis (GFP) C es A3 fusion protein has shown that CesAs also localize to the G olgi, however this observation could be an artifact of overexpression (Crowell et al., 2009). Furthermore, insertion of CesAs into the plasma memb rane is regulated by the movement of microtubule associated CesA Golgi bodies
26 (Crowell et al., 2009). Currently, the mechanism of assembly and stoichiometry of isoforms within a complex remains unknown (Taylor, 2000, 2003; Tay lor, 2008). Visualization of CesA super complexes through freeze fracture microscopy has shown that the functional, microfib ril is a symmetric hexamer comprised of six (heterohexameric) CesA protein compl exes (Haigler and Brown, 1986). Dimensions of individual cellulose chains and the terminal rosette complexes indicate that each CesA isoform produces one cellulose chain, and each hexameric rosette i s made from six CesA hexamers. The microfibrils formed fro m each rosette are thus compri sed of 36 individual cellulose chains (per diameter) (Ha et al., 1998; Somerville et al., 2004). M aize has undergone two ma jor genome duplication events. The first occurred about 70 million years ago (mya), before the lineage of maize and rice diverged, and the second was relatively recent (5 to 12 mya) (Swigonova et al., 2004). Although the maize genome has returned to an essentially diploid state, there is still an abundance of g ene duplications (Schnable et al., 2011). The maize CesA gene family has 12 members with published cDNA sequences, however p seudo genes are also common within the CesA gene family. In addition, tandem duplications of genes are common in maize (Dooner and K ermicle, 1971; Veit et al., 1990), and may well have affected the CesA family. The prevalence tandem duplications poses a challenge for genome assembly, so their extent and locale may not yet be evident in the current version of the maize genome In additi on, there are small (presumably non coding) gene fragments with high sequence identity to several of the CesAs A complete appraisal of the maize CesA gene family is continuing to emerge and may include additional functional g enes and non coding sequences. All of these are relevant to our understanding of this complex family, and for genetic approaches to discern functional roles among them.
27 A central aspect of CesA function is learning which family members act together, possibly through assembly of heterohexamers. Research in Arabidopsis indicates that three specific CesAs (AtCesA4, AtCesA7, AtCesA8) are required for secondar y cell wall synthesis (their maize orth ologs being CesA10 CesA11 and CesA12 ) and that other family members are likely in volved in synthesis of the primary c ell wall (Taylor et al., 2003). Additional work shows that in A rabidopsis and rice each of the individual CesAs contributing to secondary cell w all biosynthesis is essential. Null mutations in any of these genes result in severe phenotypes that include reduced biomass, irregular xylem, and/or brittle tissu es (Turner and Somerville, 1997; Tanaka et al. 2003; Taylor et al., 2003). In contrast, synthesis of the primary wall involves a high level of functional redundancy am ong the CesAs that contribute predominantly to this process with null mutations in AtCesA1 and AtCesA3 being the only ones associated with phenotypes (Persson, 2 007; Taylor, 2008). These include an embryo lethality and temperature dependent radial swelling of cells for cesA1 mutants, and a gametophytic lethality associated with defective pollen formation for mutants of both cesA1 and cesA3 Here we present an in depth analysis of the maize cellulose synthase gene fami ly profile expression of its members under diverse conditions and identif y co regulated isoforms. Result s will be useful in future efforts to alter quality or quantity of cellul ose in plant tissues. In the current work, we show that ZmCesA10, ZmCesA11, and ZmCesA12 are expressed coordinately, and almost exclusively in tissues undergoing secondary cell wall synthesis. Furthermore, expression of ZmCesA10, ZmCesA11, and ZmCesA12 in these tissues is elevated relative to other tissue s where expression is detecte d. Previous work has indicated that the maize ZmCesA10, ZmCesA11, and ZmCesA12 belong to the same phylogenetic sub clade of CesAs as the Arabidopsis AtCesA4, AtCesA7, AtCesA8 (secondary wall genes), and are most highly expressed in stalk and root tissue (Appenzeller et al., 2004). Here we test the hypothesis that the cellulose
28 synthases are differentially regulated with specific isoforms being coordinately expressed as a resu lt of subfunctionalization to perform specific task s To this end, we quantify mRNA levels in diverse tissues at key stages of development Results Identification and C haracterization of CesA P aralogs A bioinformatic approach was used t o characterize the s tructure, phylogeny, and possible regulatory features of the CesA f amily in maize. The s ize of this family was determined by using p ublished cDNA sequence s from the 12 known CesA s to search the maize reference genome ( B73 RefGen_v2 .: maizesequence.org) with the basic local alignment search tool ( BLAST ) (Altschul et al. 1990) Results showed several additional sequences in the genome with high homology to CesA family members. In addition four of these sequences encoded potentially functional proteins. When predicted protein sequences from these putative family members were included in phylogenetic analysis of the 12 known Ces As four new sequences were found to group with respective paralogs of Ces A7 (one), Ces A11 (one) and Ces A12 (two ) ( Figure 2 2 ). These paralogs are hereafter referred to as CesA7 a, CesA11 a, CesA12 a and CesA12 b Together the published CesAs and paralogs in maize group into six subclades, consistent with organization of CesA families in other plant species (Carrol and Specht, 2011) A closer comparison of CesA7 and CesA7 a was undertaken to determine the types of differences that distinguish closely related family members. The cDNA sequence for CesA7 a was predicted by aligning its genomic sequence with the published cDNA of CesA7. Assuming that splice sites we re conserved, a 93.8% sequence homology was observed, along with numerous single nucleotide polymorphisms (SNPs), plus 11 small insertions (< 5 bp) and two deletions (relative to CesA7 ) ( Figure 2 3A ). When the predicted CesA7 a cDNA was translated in the correct frame, it encoded an open reading frame without premature stop codons, and
29 shared 98.5% amino acid sequence similarity with Ces A7 ( Figure 2 3 B ). Additionally, qRTPCR results show that CesA7 a is expressed at levels comparable, but not identical, to CesA7 Evolution of CesA Genes in M aize Previous work suggests that the maize genome arose from gradual diploidization of an ancient tetraploid giving rise to chimeral chromosomes with segments of both the original genome s (Swignova et al., 2004; Schnable et al., 2011). To determine the role this could have played in the current structuring of the CesA family, we mapped all family members, including paralogs, to their approximate chromosomal locati ons ( Figure 2 4 A ). One of the ancestral genomes dominated in retention of CesA genes, and included at least one gene copy of each unique CesA from each of the six subclades. Conversely, CesAs from the other genome were either lost, or retained only where detectable paralogs were present. ( Figure 2 4 B ). The maize CesA family thus provides a model for exploring the functional implications of an ev olutionary mechanism recently proposed by Schnable et al. (2011) to have affected the tetraploid to diploid conve rsion in Zea mays Micro RNA T argets in the CesA Gene Family Micro RNAs (miRNA) can contribute to regulation of protein expression through degr a dation of messenger RNA (mRNA), and/or inhibition of protein translation ( Chen and Rajewsky, 2007 ). To assess the potential for miRNA based regulation of the Ces As we identified putative target sites among the CesA family members using the miRNAFinder program from the Noble Foundation (bioinfo3.noble.org/mirna/). The number of miRNA target sites for each CesA varied greatly, ranging from two to 28 in CesA2 and CesA12 respectively (Figure 2 5 ). Cluster analysis based on miRNA sequence similarity showed little to no relationship with s ub clades of their putative CesA targets. However one cluster of miRNAs showed a significant enrichment of those potentially targeting CesA10 and CesA12 ( Figure 2 6 ).
30 Expression of the CesA Gene Family During Development To better understand how cellulose s ynthase s function in vivo, and the relationships between them, we quantified mRNA levels from diverse tissues at the seedling vegetative and anthesis (reproductive maturity) stages (Table 2 1 ). Tissue was harvested at 3 days after germination (DAG) for the seedling stage, 40 DAG for the vegetative stage and 72 DAG for the anthesis stage ( Figure 2 7 ). Results showed that expression of the maize cellulose synthases was highly dynamic, with mRNA levels from different CesAs varying many fold between different tissues and during development ( Figures 2 7 and 2 8). Effects of developmental stage were prominent and determined t he proportion of tissues in which a given family member was abundantly expressed ( mRNA levels greater than 50% of the gene specific maximum) ( Figure 2 7 ). M aximum and minimum mRNA levels from a given gene varied markedly and were especially apparent for CesA10 CesA11 and CesA12 during the transition from the vegetative to anthesis stage ( Figure 2 8 ). At the anthesis stage most of the CesA s mRNA levels were at or near their m inima in leaf blades and pollen. H owever CesA3 and CesA5 consistently had the most abundant mRNA s in these tissue s which raises the question of whether these genes have a ( Figure 2 9 ). To facil itate comparative analysis of the se expression profile s, and the inter relationship among CesA family members, we explored the extent to which responses of different CesAs could be clustered We used the Modulated Modularity Clustering (MMC) program (Stone and Ayroles, 2009) to test family member groupings based on similarity of expression patterns in all tissues examined a t a given developmental stage. Each c luster was defined based on the degree of correlation observed among expression profiles of i ts members The most highly corr elated group was designated luster I with successive numbering used for less strongly clustered group s. At the seedling and vegetative stages CesA10 CesA11 and CesA12 grouped clearly
31 into Cluster I. Furthermore, lev els of mRNA from these genes rose by several orders of magnitude in rapidly growing tissues that were developing vascular tissue ( Figure 2 10 ). Cluster II included CesA7 and CesA7 a With the exception of CesA8 which was essentially independent, the remaining family members showed similar enough patterns of expression at this stage to group collectively into Cluster III ( Figure 2 10 ). At anthesis Cluster I again included CesA10 CesA11 and CesA12 ( Figure 2 9). Here too, respective mRNA levels were n otably higher in tissues that were becoming lignified and /or developing vascul ar tissue (Figures 2 11 and 2 12 ) These results are consistent with a role for CesA10 CesA11 and CesA12 in secondary cell wall synthesis. Four CesA s grouped in C luster II w here responses of CesA4 and CesA6 at anthesis join ed those of the previously grouped CesA7 and CesA7 a ( Figure 2 11 ) Although expression patterns of CesA3 and CesA5 were independent of those from other family members at this stage, responses of all remaining family members grouped together in Cluster III ( Figure 2 11 ) Developing kernels warranted separate analysis, and were harvested at 15 days after pollination S amples were dissected into embryo, endosperm, pericarp, and pedicel fractions In these kernel tissues, CesA10, CesA11 and CesA 12 again showed a similar expression pattern, and here too, maximal mRNA levels coincided with deposition of secondary cell wall s in the pedicel ( Figure 2 13 B ). Phloroglucinol staining of a longitudinal kernel section show ed the abundance of lignin in the pedicel and indicate d the extent of iness that develops in this tissue of the kernel ( Figure 2 1 3 A ) These data are consistent with the hypothesis that CesA10, CesA11 and Cesa12 have a role in s econdary cell wall synthesis in maize
32 Discussion Bioinformatic Analysis Reveals Additional CesA Paralogs At the outset of this study, there were 12 named cellulose synthases with published cDNA sequences in maize The sequencing and updating of the maize genome ( B73 RefGen_v2 2010) facilitated further analysis presented here, including identification of four additional CesA genes by BLAST alignment of published CesA cDNAs with newly emerging sequences In most instances many unknown sequenc es aligned to each individual cDNA used as a query sequence and E values indicated high level s of homology. H owever, most of these sequences were considerably shorter tha n full length cDNAs, and many ha d deletions and stop codons thus indicating that the y were likely pseudogenes and not encoding functional protein s Nonetheless t his does not rule out their potential to serve as templates for generation of r egulatory RNA sequences ( possible roles of miRNA and siRNA are discussed below but they were not a nalyzed here ). Additionally the presence of ps e udogenes can complicate interpretation of results when using PCR based genetic approaches, depending on placement of primer pairs. Related issues of gene specificity are also involved for the full length, pot entially protein coding paralogs identified here for CesA7, CesA11 and CesA12 respectively designated CesA7 a CesA11 a CesA12 a and CesA12 b ( Figure 2 2 ). To investigate the extent of sequence divergence and protein coding potential of CesA paralogs, we aligned paralog genomic sequences to the cDNAs of corresponding family members. Assuming conservation of intron exon boundaries, the paralog CesA7 a had limited sequence divergence relative to CesA7 maintaining 93.8% cDNA sequence identity ( Figure 2 3 ) that translated to 98.5% identity at the protein level. These results raise questions regarding a possible evolutionary advantage to retention of some paralogs, especially where full length cDNAs are highly conserved Although there are three C esA12 paralogs in total these showed
33 somewhat less s trong sequence similarity and included possible premature stop codons. The CesA12 a and Ces A12 b genes may still en code functi onal proteins if nucleotide sequence divergence altered in t ron exon splice sites thus caus ing a frame shift. S equences of CesA11 and Ces A11 a unexpectedly aligned to one another with 100% identity F urther investigation showed that genomic sequences flanking both genes also aligned with 100% identity until roughly 3 kilobases ( kb ) upstream, and 5kb downstream of the CesA11 sequence. Outside the boundaries where 100% homology wa s lost the sequ ences diverged completely Initially assignment of these identical sequences to different chromosomal locales seemed the possible result of a genome assembly in progress. H owever further investigation traced these sequences to two fully independent bacterial artificial chromosomes (BACS) from which the maize genome was assembled thus supporting a valid assignment of these sequences to both genomic locations Because this duplication is large (roughly 13 kb), identical, and probably recent it is unlikely to have resulted from a long terminal repeat retrotransposon or a translocation. A remaining possibility is that this unusual duplication may have been mediated by a helitron, which can transpose while carrying genom ic fragments of 15 kb or greater (Kapitonov and Jurka, 2001 ; Yang and Bennetzen, 2009 ). Phylogenetic Analysis of the CesAs Phylogenetic analysis of the CesAs and paralogs noted above indicated that several of the previously named CesAs ( e.g. CesA1+CesA2 and CesA4+CesA9 ) are probable paralogs of one another ( Figure 2 2 ) At least one paralog (two genes total) can be found in each of the six subclades of the CesA family except for that represented by CesA10 alone. T his may indicate that CesA10 gene has a unique role in the assembly or functionality of the Ces A complex, and possibly one for which multiple copies of the gene are disadvantageous. O ther CesAs lacking paralog s are not alone in their clades (as is CesA10 ), and include CesA3, CesA5, CesA6 and
34 CesA8 In addition these family members are found in clades that include only CesAs thought to be involved in primary cell wall synthesis. Furthermore, many of these family members share more than 88% sequence identity, and the three subclades in which th ey are found all share a similar overall structure (Figure 2 4B ) This observation i ndicates that the original genes may have once been closely related paralogs but have since diverged without major change function. Evolution of the CesA Gene Family Work thus far suggests that t he modern maize genome has resulted from two ancient tetraploidization events followed by subsequent diploidization. Both of the genomes that arose from tetraploidization are represented in portions of the chimera l chromosomes of modern maize (Figure 2 4A ) as a result of numerous crossovers, tra nslocations between duplicated genomes, transposition events, and significant gene loss Chromosomal segments deriving from each of the two ancestral genomes were determined by alignment with the sorghum genome (Schnable et al., 2011). The ancestral genome s were then named Maize 1 and Maize 2 based on the amount of detected gene loss, with Maize 1 retaining more genes (Schnable et al., 2011). To better understand how gene duplication and loss could have affected evolution of the CesA family, we mapped each family member to the maize genome and determined its position based association with one or the other ancestral genomes ( Figure 2 4 A) We then appl ied this genome designation to the phylogenetic tree ( Figure 2 4 B) At least one CesA gene from each subclade was mapped to a chromosomal region designated as Maize 1 whereas CesAs mapping to components of the Maize 2 genome exist only as paralogs Results were consistent with the re cent suggestion that one genome, either in its ent iret y or segment by segment, dominates with respect to resisting gene deterioration (Schnable et al., 2011) Interestingly CesA11 and CesA12 have paralogs that appear to have originated in Maize 1 either before or after the tetraploidization event. We favor a relatively recent origin for these CesA11 and CesA12
35 gene duplication s due to their positioning and the extent of their sequence similarity. However the possibility remains that these genes may already have had two copies in the original genome be fore tetraploidization, and their paralogs have since deteriorated. Potential for Micro RNAs to Affect Differential Regulation of CesA Genes Short, 22 nucl eo tide miRNA sequences can regulate mRNA degradation and inhibit translation ( Chen and Rajewsky, 2007). Degradation of mRNAs occurs through miRNA recruitment of the RNA induced silencing complex, which then cleaves the mRNA and dissociates. An initial appraisal of predicted mRNA target sites showed a highly variable number of these among t he maize cellulose synthases, ranging from two in CesA2 to 28 in CesA12 ( Figure 2 5 ) The number of potential target sites could reflect the degree to which a given family member is miRNA regulated, since more target sites would increase opportunity for interaction with various miRNAs being expressed in different tissues. We initially hypothesized that CesAs sharing a high degree of homology would be targeted by the same miRNAs, however this was not observed. Instead, only two miRNA targe t sites appeared in more than one family member one was present in both in CesA4 and CesA9 and the other was in CesA10 and CesA12 Coordinate expression was observed to varying degrees within both gene pairs (see below). Results thus indicated that miRNA mediated regulation wa Additionally, when the predicted miRNAs that targeted CesAs were clustered based on sequence similarity, they group ed randomly with regard to which CesA family member they targeted. The only exception was in one clade of miRNAs, where approximately 30% of sequences targeted CesA10 another 36% targeted CesA12 and the remainder targeted five other family members. Taken together, these observations indicate that CesA10 and CesA12 may share some degree of miRNA dependant regulation.
36 Expression of CesAs Varies with Tissue and Development E xpression profiles of the CesA family members in diverse tissues sampled at each developmental time point, show a wide range in mRNA levels for any given gene These differences in expression vary markedly from tissue to tissue, and also over time in like tissues. The identity of maximally expressed family members at any given time or in any given tissue is variable ( Figures 2 7 and 2 8 ). We suggest that expression of the CesA gene family in maize, and consequently cellulose deposition, is under complex regulation involving diverse heterohexamers with different characteristics and functions depending on contributions from different CesA isoforms. Cluster Analysis of Gene Expression Among the CesAs Application of clustering analysis to the CesA expression data from each time point revealed groups of family members that shared a high degree of similarity in expression pattern, thus indicating some level coordinate regulation (Figures 2 10 and 2 11 ) With the exception of Cluster I ( CesA10, CesA11, and CesA12 ) whose members shared the highest degree of correlat ed expression the family members belonging to a given clus ter changed at different time points For example, C luster II consistently included CesA7 and CesA7 a but CesA4 and CesA6 also joined this group at the anthesis stage (Figures 2 10 and 2 11 ). One might initially expect that within a gene family, highly similar sequences such as CesA7 and CesA7 a would be similarly expressed. However, genes with clear, consistent differences in their sequences were also frequently coexpressed. This issue is especially important when the genes code for isoforms of a heterogenous protein complex like that of C ES A Our data show more support for the c onserved differences scenario, considering that Cluster I ( CesA10, CesA11, and CesA12 ) was the only group to consistently represent the same family members and each of these genes represents its own subclade on the CesA family tree. Furthermore, even though expression
37 profiles of CesA7 and CesA7 a group consistently together the same can not be said for other paralogs such as CesA4 and CesA9 (Figures 2 10 and 2 11 ) As noted above the only family members with expression patterns that clustered together exclusively and consistently throughout development were CesA10, CesA11, and CesA12 The variation in expression levels of these genes between tissues was also more extreme than for other family members, sometimes increasing coordinately by up to 60 fold ( Figure 2 6) At all stages of development, the tissues in which CesA10, CesA11, and CesA12 we re most highly expressed were also those where vascul ar tissue was developing or tissues we re becoming lignified (Fig ure s 2 10 2 11, and 2 12 ) Analysis of mRNA levels of CesA family members in the developing kernel are consisten t with whole plant observations, since mRNAs of CesA10, CesA11, and CesA12 were several fold more abundant in the highly lignified pedicel than in other kernel tissues ( Figure 2 13 ) Earlier studies also showed that maize CesA10, CesA11 and CesA12 had c losest homology to CesAs in Arabidopsis and rice that were necessary for secondary cell wall synthesis (Tanaka et al. 2003; Taylor et al., 2003; Appenzeller et al., 2004). Collectively, research presented here indicates that CesA10, CesA11, and CesA12 are primarily, although not exclusively, associated with secondary cell wall synthesis in maize Methods Bioinformatic Analysis of the Maize Genome for Identification of CesA Paralogs Paralogs of named CesA f amily members were identified by using BLAST (Altschul et al., 1990) to compare the cDNA sequence from each named CesA to the maize genome (B73 RefGen_v2 : maizesequence.org) O utput was screened to identify sequences most likely to encode functional CesA paralogs. Sequences were consi dered paralogs if they 1 ) had high homology to the query sequence (E value of 0), and 2 ) includ ed sufficient alignment to encode a potential protein of similar length to the CesAs Each newly identified, putative p aralog was
38 characterized by aligning its g enomic sequence with the cDNA of its respective, homologous CesA gene. For each paralog, cDNA sequence was predicted assuming a conservation of mRNA splice sites followed by removal of introns Each of these p redicted cDNA s was the n aligned with cDNA from its cor responding CesA family member ( multalin. toulouse. inra. fr ) to manually map SNPs, insertions and deletions ( Figure 2 3 ). Phylogenetic Analysis of the CesA Family Phylogen y of the CesA family, including paralogs, was analyzed using the MEGA4 program (Tamura et al., 2007 ). Sequences of p rotein s and predicted protein s (in the case of paralogs) were aligned and used to build an unrooted, neighbor joining, phylogenetic tree based on homology. The tree was constructed using a pairwise deletion option with 1000 bootstrap replications ( Figure 2 2 ). Evolutionary Analysis of the CesA Family The evolutionary origin of the mai ze CesAs and their paralogs was estimated by mapping each gene on the maize genetic map and ascertaining which portion of the ancient tetraploid genome was represented in the corresponding chromosomal segment This provided a m eans of designating which C esA genes resided on each of the two sub genomes generated by ancient tetraploidizatio n and subsequent diploidization ( Figure 2 4 A). T he positions of chromosomal regions derived from each subgenome have been described in previous work (Swignova et al., 2004; Wei et al., 2007; Schnable et al., 2011). Assigning a genome designation to each C es A protein in the phylogenetic tree facilitated interpretation o f the evolutionary relationship between CesA family members and their paralogs. Prediction of miRNA Target S ites in the CesAs Micro RNA target sites within the CesA genes were predicted by the miRNAFinder program (bi oinfo3.noble.org). G enomic sequence of each CesA family member was used to
39 query the r ice Expressed Sequence T ags (EST) collection, and the ric e genome w as then used to make target site predictions. Predicted miRNAs for each gene were compiled and redundant miRNAs, or those mapping to the same location in a gene ( i.e. identical pre miRNA sequences appearing multiple times in the output) were eliminated. All 154 predicted miRNAs were the n aligned and clustered using the MEGA4 program (Tamura et al., 2007) to determine if (predicted) miRNAs targeting co expressed CesAs were related The resulting tree was constructed using pairwise deletion with 1 000 bootstrap replications. Each CesA was assigned a uni que color that was shared by all miRNAs targ eting that gene ( Figure 2 6 ). Plant Material M aize inbred W22 was used for all expression analyse s. Plants were grown in the laboratory for samples dissected at the seedling stage, and under field conditions for samples harvested at the vegetative and anthesis stages. All material was frozen immediately in liquid nitrogen. Isolation of RNA and cDNA Synthesis Approximately 200 mg of tissue from each sample was finely ground in liquid nitrogen, and incubated in 1ml Trizol (Invitro gen Cat # 15596 018) for 5 min at 25C with frequent vortexing (15 s) Chloroform (200 mL) was added and the solution vortexed for 15 sec, allowed to incubate for 1 min at 25C, and vortexed again for 15 sec Samples were then centrifuged at 13,200 rpm for 10 min to separate phases and 200mL was transferred from the top phase to 700 mL Qiagen RLT buffer (RNeasy Plant Mini kit, Qiagen Cat # 74904). Ethanol (100%, 500 mL) was added before vortexting (15 sec) Total RNA was th en cleaned and eluted using the RNeasy Plant Mini kit (Qiagen Cat # 74904), a s per the manufacturer s protocol Any contaminating DNA was removed by treatment with DNase 1(Ambion Cat # AM1906). Total RNA concentration was quantified using a ( Bio Rad ) SmartSpec 3000 spectrophotometer, fol lowed by
40 dilution to 50 ng/L. Diluted RNA was used to synthesize cDNA with a SuperScript One Step kit (Invitrogen Cat # 10928 042) Resulting cDNA was diluted 10 fold Real Time Quantitative RT PCR Levels of mRNA in tissues sampled throughout development were quantified using a Step One Plus Real Time PCR System (ABI). Gene specific primers for each CesA were designed manually, and specificity was checked by ensuring that primers did not closely align to a ny other family member s paralogs or closely related genes (see appendix Table B 1 ) Three biological replicat es, and two technical replicates were quantified for each tissue sampled. Reaction s used the SYBR Green platform, with each 20 L reaction contai ning 10 L Fast SYBR Green Master Mix (ABI Lot # 1003024), 5 L cDNA, and 100 nM of both forward and reverse primers. Transcript abundance was normalized using 18 S ribosomal RNA (Taqman Ribosomal RNA Control Reagents, ABI Lot # 0804133) as a control. Optimization of the control reactions resulted in a final reaction volume of 20 L that contained 10 L Fast SYBR Green Master Mix, 1 L forward and reverse control primer s (diluted 1:18 from the ABI kit concentration), and 2.5L cDNA. Each 96 well PCR plate represented one of the tissues sampled, and contained the 84 reactions necessary for two technical replications of three biological replications (tissue samples) for ea ch of the 1 4 genes tested (13 CesAs and 18S rRNA). Amplification curves were closely monitored to ensure the kinetics of each run were comparable. Results were analyzed using the Modulated Modularity Clustering program (Stone and Ayroles, 2009) to assign c oregulated family members to modules based on similarity in expression patterns. This program was chosen because it was specifically designed to seek community structure in graphical data. Phloroglucinol Staining Whole kernels were harvested at 15 DAP, an d fresh tissue was longitudinally hand sectioned Other tissues were harvested at anthesis and sectioned in diverse planes
41 Samples were then immersed in saturated phloroglucinol solution containing 20% ethanol and 20% HCl. After a 2 min incubation at 25C, sections were washed with water and images obtained using a RT SPOT camera (Diagnostic Instruments Sterling Heights, MI ) mounted on a Leica MZ 12 5 dissection microscope.
42 Table 2 1. Tissues sampled at key developmental stages Seedling Vegetative Anthesis Coleoptile First leaf Scutellum Primary root Developing leaf Young leaf Node Stem Sheath Root Prop root initial Tassel init ial Ear initial Pith Leaf blade Leaf sheath Husk Ligule Midrib Stem (heavily lignified) Stem (moderately lignified) Mature prop root Developing prop root Pro p root ti p Lateral roo t Lat. Root initiation zone Spikelet Pollen Floret Silk Cob Seedlings were grown on a growth bench under lab conditions and sampled at 3 days after germination (DAG). Vegetative stage and anthesis stage plants were grown under field conditions (spring, 2008 at the UF Plant Science Research Unit, Citra, FL) and sampled at 40 and 72 DAG respectively.
43 Figure 2 1. Diagram of a canonical Ces A protein. The approximate locations of transmembrane domains are represented by grey cylinders. Cytopla smic domains are designated; Zn ( Zinc binding/RING ); HVR( hypervariable region ); and CSR ( class specific region ) The D,D,Q,X,XR,W glycosyl transferase mo tif is also shown.
44 Figure 2 2 Unrooted, neighbor joining phylogenetic tree of the maize CesA family determined by protein similarity The CesA1 through CesA12 proteins represent family members with published cDNA sequence s Family members in green are paralogs identified bioinformatically. The tree was constructed using pairwise deletion with 1,000 bootstrap replications in a MEGA4 analysis Bootstrap values less than 100 are shown.
45 Figure 2 3 Predicted cDNA sequence o f CesA7 a and its translated protein sequence A) Assuming conservation of intron exon boundaries, CesA7 a genomic sequence was aligned to CesA7 cDNA to remove introns. Polymorpmisms (relative to CesA7 ) were mapped, with SNPs highlighted in purple, and ins ertions in yellow. Deletions are denoted by white background with red text. Overall, the two cDNA sequences share 93.8% homology. B) When translated, the CesA7 a predicted cDNA does not introduce premature stop codons, and is 98.5% conserved with Ces A7. Met and Stop are written fully to clarify their status.
46 B Figure 2 3. Continued Met E A S A G L V A G S H N R N E L V V I R R D G D P G P K P P P R E Q N G Q V C Q I C G D D V G L A P G G E P F V A C N E C A F P V C R D C Y E Y E R R E G T Q N C P Q C R T R Y K R L K G C Q R V T G D E E E D G V D D L D N E F N W N G H D S R S V A D S Met L Y G H Met S Y G R G G D P N G A P Q P F Q L N P N V P L L T N G Q Met V D D I P P E Q H A L V P S F Met G G G G K R I H P L P Y A D P S L P V Q P R S Met D P S K D L A A Y G Y G S V A W K E R V E N W K Q R Q E R Met H Q T R N D G G G D D G D D A D L P L Met D E S R Q P L S R K I P L P S S Q I N P Y R Met I I I I R L V V L G F F F H Y R V Met H P V N D A F A L W L I S V I C E I W F A Met S W I L D Q F P K W F P I E R E T Y L D R L S L R F D K E G Q P S Q L A P I D F F V S T V D P L K E P P L V T A N T V L S I L S V D Y P V D K V S C Y V S D D G A A Met L T F E A L S E T S E F A K K W A P F C K R Y N I E P R A P E W Y F Q Q K I D Y L K D K V A A N F V R E R R A Met K R E Y E E F K V R I N A L V A K A Q K V P E E G W T Met Q D G T P W P G N N V R D H P G Met I Q V F L G Q S G G L D C E G N E L P R L V Y V S R E K R P G Y N H H K K A G A Met N A L V R V S A V L S N A P Y L L N L D C D H Y I N N S K A I K E A Met C F Met Met D P L L G K K V C Y V Q F P Q R F D G I D R H D R Y A N R N V V F F D I N Met K G L D G I Q G P I Y V G T G C V F R R Q A L Y G Y D A P K T K K P P S R T C N C W P K W C F C C C C C G N R K H K K K T T K P K T E K K K L L F F K K E E N Q S P A Y A L G E I D E A A P G A E N E K A G I V N Q Q K L E K K F G Q S S V F A T S T L L E N G G T L K S A S P A S L L K E A I H V I S C G Y E D K T D W G K E I G W I Y G S V T E D I L T G F K Met H C H G W R S I Y C I P K R P A F K G S A P L N L S D R L H Q V L R W A L G S I E I F F S N H C P L W Y G Y G G G L K F L E R F S Y I N S I V Y P W T S I P L L A Y C T L P A I C L L T G K F I T P E L N N V A S L W F Met S L F I C I F A T S I L E Met R W S G V G I D D W W R N E Q F W V I G G V S S H L F A V F Q G L L K V I A G V D T S F T V T S K G G D D D E F S E L Y T F K W T T L L I P P T T L L L L N F I G V V A G V S N A I N N G Y E S W G P L F G K L F F A F W V I V H L Y P F L K G L V G R Q N R T P T I V I V W S I L L A S I F S L L W V R I D P F L A K D D G P L L E E C G L D C N Stop G Met S A H Q L P Q S A Y D Stop S I F A G V C P H I Y S A P S V G K R Q E Met S P V P F D P W Stop T S T Stop Y L G Y T G G K Met E A A A I L V Q Met G R G I Q H Met Q V F D C A A F F I T W A Q N Stop S S E P S S K V F Stop S C T A P V Y K L G S Q Stop G R Q E C A S A S G T E E P A Q Y L C T N V H W R A C S L H V R L Y Stop E K Q N I C T N L Y L I K V C K G V P F F F L C T V I V G V G F V
47 Figure 2 4 Approximate distribution of the CesA family and close paralogs within the maize genome. A) R epresentation of CesA positions alongside the chimeral chromosomes of the modern maize genome that result ed from an ancient tetraploidization event and subsequent diploidization. Genome copies ( result ing from tetraploidy) are termed Maize 1 and Maize 2 and are represented by blue and orange, respectively. Chromosomal segments that could not be assigned to either genome are white. Note that several CesA genes are paralogs of others (see Figure 2 4B). N ewly identified paralogs are named after the family member they are paralagous to, followed by a letter (e.g. CesA7a ). In some instances ( CesA12a and CesA12b for example) paralogs map to different genomic copies. Chromosomal domains a re shown as per Schnab le et al. ( 2011 ) B) A phylogenetic tree showing the relationship between Ces A and Ces A paralog proteins relative to the genome copy in which they reside The tree was constructed using pairwise deletion with 1,000 bootstrap replications in a MEGA4 analysis ( Tamura et al., 2007 ).
48 Figure 2 4 Continued
49 Figure 2 5. Predicted miRNAs targeting each of the maize CesAs Potential target sites and cooresponding miRNAs were predicted using the miRNAFinder program (bioinfo3.noble.org/mirna/). Results were based on similarity of maize CesA genomic sequence to the rice EST database and the rice genome. Potential miRNAs that t argeted more than one CesA are highlighted.
50 Figure 2 6. Cluster analysis of potentially CesA targeting miRNAs. MicroRNAs were grouped based on sequence similarity, using pairwise deletion with 1,000 bootstrap replications in a MEGA4 analysis ( Tamura et al., 2007 ) The CesA family member that each miRNA targets is designated by color. A clade enriched for miRNAs potentially targeting CesA10 and CesA12 is circled in orange.
51 Figure 2 7 Heat maps representing expression of CesA family members in all tissues sampled at each time point. Diagrams show relative developmental state of plants sampled at seedling (3DAG), vegetative (40DAG) and reproductive stages of development. M aps show the relative expression of each gene at a given time point as a percentage of its maximum
52 Figure 2 8 Relative expression of the CesA family in all tissues samples A) The seedling (3DAG) and vegetative stages (40DAG). B) Anthesi s ( 72 DAG ) Expression levels are relative to the amount 18SrRNA measured in each tissue. Data represent three biological replications.
53 Figure 2 8 Continued
54 Figure 2 9 Expression of CesAs in tissues with low mRNA abundance. In three of the tissues with overall lowest transcript levels, CesA3 and then CesA5 are the dominantly expressed family members. Expression levels are relative to the amount 18SrRNA measured in each tissue. Data represent three biological replicates.
56 Figure 2 10. Cluster analysis of the CesA family at the see dling and vegetative stages. A correlation value of 0.8 was used as the cutoff for cluster designation. Genes having insufficient similarity in expression pattern to other family members were considered unclustered. Cluster number indicates strength of correlation between its members. The mean correlation values for cluster I, cluster II, and Cluster III were 0.99, 0.98, and 0.87, respectively. Colored bars along the X axis indicate tissue type, with green = above ground brown = below ground yellow = male reproductive and red = female reproductive tissues. The Modulated Modularity Clustering program (Stone and Ayroles, 2009) was used to define clusters. Expression levels are relative to the amount 18SrRNA measured in each tissue. Data represent thre e biological replicates.
58 Figure 2 11. Cluster analysis of the CesA family at a nthesis/reproductive maturity. A correlation value of 0.8 was used as the cutoff for cluster designation. Genes having insufficient similarity in expression pattern to other family members were considered unclustered. Cluster number indicates strength of correlation between its members The mean correlation values for Cluster I, Cluster II, and C luster III were 0.96, 0.90, and 0.89, respectively. The Modulated Modularity Clustering program (Stone and Ayroles, 2009) was used to define clusters. Expression levels are relative to the amoun t 18SrRNA measured in each tissue. Data represent three biological replicates.
59 Figure 2 12. Lignin and polyphenol deposition in tissues a t the anthesis stage (72 DAG). Cross sectioned t issues were stained (pink/red) with phlorogluc inol/HCl soluti on to detect the presence of lignin/secondary cell wall deposition. Only tissues showing visible staining are shown
60 Figure 2 13 Expression of the CesaA gene family in kernel tissues 15 days after pollination. A) A longitudinal kernel section stained with phloroglucinol. Pink staining indicates presence of lignin/secondary c ell wall. B) Expression of CesA s in 4 kernel tissues. Expression levels are relative to the amount 18SrRNA measured in each tissue. Data re present three biological replicates. Photo by
61 CHAPTER 3 EXPRESSION OF THE CE LLULOSE SYNTHASE GENE FAMILY IN A MAIZE SUSPENSION CELL/PROT OPLAST SYSTEM Background The plant cell wall is a complex, cellulose rich stru cture that serves several essential functions. The cell wall not only provides the cell and whole plant with mechanical strength, but als o aids in defense against pathogens determines cell shape and participates in signaling (Aziz et al., 2007; Galletti et al., 2009). Because cellulose is the ma jor load bearing component of the cell wall, considerable effort has been directed towards understanding its synthesis and regulation at the whole plant and tissue levels. Most studies have investigated CesA ex pre ssion in tissues of wild type plants ( Pear et al., 1996; Delmer, 1999 ; Appenzeller et al., 2004; Wang, 2010), or analyzed effects of CesA mutations (Tanaka et al. 2003; Taylor et al., 2003; Taylor, 2008). In contrast, comparatively little work has utilize d cell culture or protoplast approaches to investigate cellulose synthase expression in these readily perturbed systems. A useful approach to question s of how cellulose synthesis is r egulated at the cellular level can be addressed by investigating gene expression in response to perturbations of undifferentiated suspension cells and during cell wall regeneration by protoplasts Release of protoplasts from cell s of various plant tissues is a common practice, and a widely used approach for diverse f undament al studies For example, o rganelles and other cellular constituents can be isolated more readily from protoplasts than from whole tissues (Tallman and Reeck 1980) and removal of the cell wall facilitates investigations into physical and chemical properties of the plasma membrane (Stafford and Warren, 1991 ; Yamazaki et al., 2008 ). Protoplasts can be obtained from specific cell types where they have been invaluable in studies of processes such as the roles of bundle sheath cel ls in C4 metabolism (Edwards et al. 1979 ; Shatil Cohen et al.,
62 2011 ) and light effects on guard cell functions (Zeiger and Hepler, 1979 ; Pandey et al., 2002 ). Protoplasts have also proven versatile for analysis of transient gene expression because the ac cessible plasma membrane makes them ideal for genetic transformation through ele ctroporation or microinjection (Potrykus, 1991 ; Dong Yoo et al., 2007 ). Protoplasts are stressed during the isolation process, however, and resulting changes in gene expression and metabolism necessitate ample controls and cautious interpretation of results. T he cell wall can be effectively removed by use of enzyme treatments that degrade its polysaccharide constituents. The enzymes are typically extracted from fungal sources ( parasitic and/or necrotrophic ) (Wang et al., 1991 ; Ling et al., 2010 ) L iquid medium used to culture cellulytic bacteria has also been shown to degrade cell walls (Tamaru, 2002 ). W oun ding and pathogen response pathways can be induced by exposure of cells t o fungal elicitors such as cell wall deg rading enzymes and contaminants and/or oligosaccharides released from the cell wall during d igestion (Cordero et al., 1994; Moreno et al., 2005) Changes in metabolism and gene exp ression can also occur as cells and protoplasts respond to 8 h ours of incubation in the sugar free environment of cell wall digestion medium (Yu, 1999). Furthermore, biotic and abiotic stress responses may result from the mechanical damage and/or stres ses associated with the lack of a cell wall (Walley et al., 2007) Cell wall composition of regenerated protoplasts typically differs from that of walls from either the cell cultures or plant tissues from which protoplasts were obtained (Blaschek et al. 1981; Pilet et al. 1984; Gould et al. 1986; Wang et al., 1991). During de novo cell wall synthesis, protoplasts can produce callose D glucan ) instead of cellulose D glucan) and synthes ize different polysaccharides than those in cell walls of the s ource tissue. However, work by Shea et al. (1989) demonstrated that protoplasts derived from carrot suspension cells c ould regenerate compositionally similar cell wall to that of normal suspension cells An except ion was
63 that crystalline cellulose was slower to form, and took 3 days for full regene ration (Shea et al., 1989). The authors suggest ed that purity of wall degrading enzymes may be important to prevention of wound response s that could cause abnormal synthesis of callose and other cell wall polysaccharides. Work with maize has demonstrated that protoplasts derived from suspension cells regenerate cell wall s more rapidly than those derived from mesophyll (Wang et al., 1991). Furthermore, the regenerated cell walls of protoplasts der ived from suspension cells are more evenly distributed, more compact, and better organized than those from mesophyll (Wang et al., 1991). Additionally, when g ene expression in potato leaf protoplasts was examined during cell wall regeneration, expression profiles showed no significant upregulation of cell wall biosynthetic genes relative to shoot tissue in planta (Oomen et al., 2003). Suspension cells provide yet another approach for f undamental studies of cell wall biosynthesis Like protoplasts, suspension cells can be derived from diverse tissues, but the tissue from which the suspension cells are initiated must be able to form callus. Suspension cells can also be derived from both d ifferentiated and undifferentiated tissues, thus allowing a choice in the characteristics ( i.e. cell wall composition, genetic pre programming, metabolic properties) of the cell type to be studied. It should be noted that cultured plant cells are often sub ject to somaclonal variation, and cell lines derived from these variants may have different properties than the source tissue (Lee et al., 1988 ; Bruneau and Qu, 2010) Application of growth mod ulators such as phytohormones, phosphates, micronutrients, and sugars has also been shown to control cell differentiation and organ formation (Skoog and Miller, 1957; Oda et al., 2005). Furthermore, research in Arabidopsis indicates that in immortalized suspension cell cultures, where cells are continuously proliferating, dedifferentiation occurs concomitantly with clear epigenetic changes (Tanurdzic et al., 2008). Chromatin immunoprecipitation analysis has shown these chang es include hypermethylation of euchromatic regions, and hypomethylation of
64 heterochromatic regions corresponding to transposable elements (TEs). The resulting activation of TEs was accompanied by an increase in 21 nucleotide small interfering RNAs, thus i mplicating RNA interference along with chromatin remodeling in epigenetic restructuring of immortalized suspension cell lines (Tanurdzic et al., 2008). Plant suspension cell cultures are a highly versatile research platform because one can subject the sys tem to diverse, targeted perturbations. Not surprisingly, they have been used to study a wide variety of cellular processes including specific responses to abcisic acid and gibberelic acid (Xin and Li, 1992; Huang and Lloyd, 1999), aquaporin function (Cave z et al., 2009), mitochondrial release of calcium under anoxia (Subbaiah et al., 1998), and tracheary element formation (Fukuda, 1997). Suspension cells ha ve been pa rticularly useful for studying cell wall biosynthesis because they can be induced to produce secondary cell wall s Depending on the species and source tissue, different hormone treatments and/or sugar concentrations can induce differentiation of suspension cells to tracheary element like (TE L ) cells that are similar to those found in xyle m ( Falconer and Seagull, 1985; Fukuda, 1997; Oda et al., 2005; Ohashi Ito et al., 2010). Indu ction of TEL formation was optimized in Zinnia elegans L. to study xylogenesis, and the cha racteristics of TELs were remarkably similar to those of intact xylem (F alconer and Seagull, 1985). Tracheary element like cells had diverse patterns of secondary cell wall deposition typical of xylem such as annular, reticulate, spiral, sc a lariform and pitted thickenings Subsequently, TEL induction in suspension cells has b een used to study the relationship between microtubules and secondary cell wall deposition (Oda et al., 2005), expression and regulation of cell wall biosynthetic genes ( Ohashi Ito et al., 2010) and programmed cell death (McCabe and Leaver, 2005; Ohashi I to et al., 2010)
65 Here we employ both protoplast and suspension cell systems to experimentally perturb regulation of gene expression in the maize CesA gene family. Relative gene expression was quantified during de novo cell wall biosynthesis by protoplasts for up to 60 h of regeneration. Expression of the CesA family was als o assayed in suspension cells undergoing hormone induced differentiation to TELs. In addition, CesA expression wa s analyzed in both the protoplast and suspension cell systems after specific perturbations. During de novo cell wall synthesis by protoplasts, CesAs from the primary and secondary cell wall synthesizing subclades were highly upregulated relative to suspension cells. Also, reducing the sucrose content o f protoplast regeneration media showed that the timing and identity of CesAs expressed is responsive to sugar availability. Futhermore, induction of secondary cell wall deposition in suspension cells resulted in concurrent increase in levels of CesA10 Ces A11 and CesA12 mRNAs, consistent with previously described observations in planta (see chapter 2) Results Establishment of the Protoplast Regeneration System Multiple p arameters including digestion media, digestion time, agitation speed, harvest methods, purification, and regeneration media were optimized to establish an efficient system for harvest of protoplasts, and develop a suitable environment for cell wall regeneration. The concentr ations of cell wall degrading enzymes in the digestion medium was adjusted to release the greatest number of protoplasts. Digestion time and agitation speed were adjusted to maximize yield of viable undamaged protoplasts. Protoplast harvest (centrifugatio n speed) purification (filtering and flotation methods) and regeneration me dia (sucrose and 2 4 D concentration) were also optimized, with the end result being a reliable system with reproducible results from a given time course for cell wall regeneratio n (Figure 3 1). The most essential aspect of this process was
66 adjusting the digestion media, digestion time, and agitation speed to maximize effective release of protoplasts from suspension cells without compromising viability. Embryo derived suspension ce lls from maize inbred B104 were used as the source of protoplasts. After 8 h ours of incubation in digestion media, prot oplasts were harvested, purified, and transferred to regeneration media for observation of cell wall biogenesis during the subsequent 60 h ours Prior to protoplast release, light microscopy showed s uspension cells were typically present as aggregations of translucent to slightly opaque, irregularly shaped, ovoid cells T hese cells also fluoresce d bright bluish white under fluorescent l ight when stained with calcofluo r white ( Figure 3 1), indicating presence of abundant cellulose. After the 8 h our period of protoplast release in digestion media, n ascent protoplasts (0 h ours ) we re transparent, completely spherical, and show ed no fluorescence ( Figure 3 1) As protoplasts regenerate d cell wall s, the cells bega n to aggregate and at 12 h ours of regeneration, slight, patchy calcofluo r white fluorescence became evident ( Figure 3 1). After 20 h ours most viable cells had aggregated become translucent, taken on an irregular shape, and show ed fluorescence The fluo rescence was not as strong as that of suspension cells however, indicating that cell wall regeneration was likely still in progress ( Figure 3 1). Expression of CesAs and Sucrose Synthases in Protoplasts Regenerating Cell Walls To determine responses of CesA family members to the extreme demands for cell wall biosynthesis by protoplasts mRNA levels of CesAs from the primary and secondary cell wall synthesizing subclades ( Figure 2 1) were quantified Additionally, transcript abundance was determined for three sucrose synthases since these are hypothesized to associate loosely with the C ES A complex and provide UDP glucose as substrate for cellulose synthesis (Amor et al., 1995; F ujii et al., 2010). The CesAs did not show significant upregulation until about 4 h ours after induction of cell wall regeneration, and expression of the sucrose synthases remained low until
67 20 h ours after transfer to induction medium ( Figure 3 2 ) With the exception of CesA1, CesA2 and CesA11 relative expression levels of all other genes analyz ed peaked at 44 h ours after initiation of cell wall regeneration and dropp ed sharply thereafter In addition, a transient and repeatable peak in mRNA levels occurred at 12 hours for CesA7 CesA7 a CesA10 and CesA11 ( Figure 3 2 ) An additional transient peak for CesA11 was observe d at 28 h ours Relative expression of most other genes increase d with each progressive time point, up to 44 h ours With the exception of Sus1 and Sus2 expression profiles were highly variable, thus indicating a lack of coordinate regulation under these conditions ( Figure 3 2 ). CesA and Sucrose Synthase Expression During Cell Wall Reg eneration With Perturbations in Sucrose Concentration Sugar substrates are necessary for cell wall biosynthesis and s ucrose metabolism is hypothesized to provide substrates to the CESA complex ( the reversible sucrose synthase reaction can generate fructose and UDP glucose). In addition, sugar availability can profound ly affect m etabolism and reprogram expression of related genes (Su, 1999; Koch 2004). We thus investigated the effects of sucrose starvation on CesA and sucrose synthase expression during cell wall regeneration The sucrose conce ntration in regeneration media used for both pr otoplasts and suspension cells wa s normally 2% w/v, so we compared results under these conditions to responses under 0.5% and 0% sucrose. Under reduced sucrose conditions (0.5%) sufficient RNA for analysis could be recovered for the first 36 h ours after induction of cell wall regeneration ( Figure 3 3 ) E xpression of the CesAs w as comparable (to one another) at each time point between 12 h ours and 28 h ours of regeneration, with the exception of CesA12 ( Figure 3 3 ) At 4 h ours, mRNAs of all CesAs from CesA1 through CesA11 were at their lowest levels, whereas expression of CesA12 was 8 fold higher than its minimum, which did not occur until 20 h ours Transcript levels of CesA12 peaked
68 at 28 hours increasing 32 fold relative to expression at 20 h ours ( Figure 3 3 ). The most prominent change in expression pattern was at 36 h ours, where transcript levels of most CesAs roughly doubled their previous maxima ( Figure 3 3 ) The sucrose synthases did not share any of these changes in expression. L evels of mRNAs were consistently similar for all three genes ( Sh runken 1 Sucrose synthase 1 and Sucrose synthase 2 ), with transcript abundance increasing slightly throughout the experi ment Overall, expression of the sucrose synthases was relatively low, with mRNA levels at 36 h ours roughly double those at 4 h ours The expression patterns of the CesAs and sucrose synthases changed markedly when protoplasts were allowed to regenerate in media without sucrose. Under these conditions, sufficient RNA could only be recovered for the first 28 h ours after induction of cell wall regeneration. Expression of the cellulose synthases was most highly upregulated and variable at 12 h ours instead of at the last time point available (28 h ours ), as was observed under reduced sucrose ( Figure 3 3B ) The observation that expression levels of CesA12 fluctuated more than other family members may indicate that some aspect of its regulation is more sensitive to sugar limiting conditions. The sucrose synthases had expression pattern s nearly identical to th ose observed when protoplasts were grown in reduced sucrose, i llustrating that these genes were less responsive to variation in sugar availability than the CesAs ( Figure 3 3B ). Cellulose Synthase and Sucrose Synthase Expression in Response to Simulated Wounding/Pathogen Infection Incubation in cell wall digestion media expose s cells to molecules that stimulate expression of defense response genes. Examples include contaminating fungal toxins cell wall degrading enzymes, and oligosaccharides released from the cell wall s during digestion. These can all elicit defense respons e s to wounding and/or pathogen infection (Cordero et al., 1994; Seifert and Blaukopf, 2010) Additionally, removal of the cell wall may initiate signals of
69 mechanical damage in protoplast s further enhancing response s to wounding and altering normal gene expression patterns. To determine the extent of influence potentially exerted on CesA expression by defense response s we simulated wounding and/or pathogen infection in suspension cells, then quantif ied mRNAs from selected CesAs ( CesA1, CesA7 ,CesA12 ) and the systemic wound/pathogen response gene MPI (maize proteinase inhibitor) (Cordero et al., 1994). Expression of MPI was also assayed in regenerating protoplasts. The expression profile of MPI in regenerating protoplast s was similar to that o bserved for the CesAs and sucrose synthases, with slight upregulation beginning at 4 h ours and increasing rapidly through 44 h ours ( Figure 3 4). Expression levels of test genes were measured at four time points under conditions in which regeneration media contained either 1) no treatment ( control ) 2) ground whole cells, 3) denatured cell wall degrading enzymes, or 4) purified partially digested cell wall. In the control media, MPI levels were greatest in freshly transferred suspension cells, but dropped rapidly during a 12 hour incubation. Cells in media containing ground whole cells showed a peak in MPI expression at 2 h ours and slight downregulation of CesA7 and CesA12 ( Figure 3 5 ). In solution containing denatured cell wall digestion enzy mes, all genes were slightly upregulated with peak expression at 2 h ours ( Figure 3 5 ) These data show that expression of the CesAs is most responsive to simulated pathogen infection. However, this response is transient, with expression levels returning to those of pre treatment conditions by 12 h ours (Figure 3 5 ). Expression of the CesAs During Induction of Secondary Cell Wall Biosynthesis With the goal of defining which cellulose synthases may be involved in secondary cell wall biosynthesis, we treated m aize suspension cells with phytohor mones that were previously shown to induce secondary cell wall formation in Zinnia (Falconer and Seagull, 1985; Endo et al., 2009) After 7 days of incubation in secondary cell wall induction media, suspension cells
70 began to differentiat e into tracheary element like (TEL) cells ( Figure 3 6 ) consistent wi th results from previous work (Falconer and Seagull, 1985; Fukuda, 1997; Oda et al., 2005; Ohashi Ito et al., 2010) Concurrent with the appearance of TELs in culture, mRNA levels of CesA10 CesA11 and CesA12 were highly elevated relative to the control (Figures 3 6 and 3 7 ). Upregulation of these genes was maintained, although at lower levels, until 25 days after transfer (DAT) to induction medium Taken together, thes e results are consistent with our in vivo observations indicating that CesA10 CesA11 and C esA12 play a role in secondary cell wall biosynthesis Discussion Cellulose S ynthase Expression During Cell Wall Regeneration E xpression profiles of the CesAs anal yzed indicate that family members associated with both primary and seco ndary cell wall formation were involved in de novo wall bio synthesis by protoplasts ( Figure 3 2). This result was contrary to expectations because the suspension cells used as a source of protoplasts had only primary cell walls, and were derived from undifferentiated embryonic cells that also lacked detectable secondary cell wall s Involvement of both CesA classes in cell wall regeneration may be due to concurrent upregulation of all cell wall biosynthetic genes in response to the severity of complete cell wall removal and signals for its regeneration Other stresses associated with cell wall digestion may also have been involved, as well as elicitation of defense mechanisms The i mpact of stress signals would be consistent with the duration and pattern of observed gene expression although cell wall regeneration also followed a similar time course. Sucrose synthase mRNA levels decreased concurrently with those of most CesA genes, indicating a potential reduction in capacity for direct conversion of sucrose to precursors of cellulose biosynthesis. The proposed role of sucrose synthase in cellulose formation may have
71 been especially important early in cell wall regeneration, when biosynthesis of cellulose was a major sink for carbon metabolism. Later, however, the carbon demands for cell wall formation would be expected to lessen, even though mRNAs for some biosynthetic genes remain abundant. Also, mRNA levels of all genes tes ted did not rise detectably until 4 h ours after transfer to regeneration media This 4 h our delay may have been due to a period of cellular recovery from the combination of stress and starvation during cell wall removal (digestion medium lacks metabolizabl e carbohydrate). Expression of three CesAs was distinctive during cell wall regeneration, with mRNA levels of CesA1, CesA7 and CesA11 continu ing to rise through out a 60 hour period, well after transcript levels from other family members had decreased ( Fig ure 3 2) This degree of coordinate regulation was not observed for these selected CesA genes under any of the developmental stages or tissues examined in planta (Figures 2 10, 2 11, and 2 13 ). However, their different regulat ion under these conditions remains consistent with the flexibility observed in vivo for the shifting combinations of expression for members of the CesA gene family The possibility also exists that these CesAs may contribute to formation of functional comp lexes not typically abundant in planta Expression of CesAs During Cell Wall Regeneration Varied with Sucrose Supply Limiting the concentration of sucrose in protoplast regeneration media changed the expression profile s for CesA family members and also the duration of cell viability ( Figure 3 3). Under reduced (0.5% w/v) sucrose sufficient RNA for analysis could be obtained for 36 h ours but viability was apparently compromised thereafter The protoplasts may have been unable to form a cell wall, and/or lack ed a sufficient energy source Prior to this point, two distinctive responses were evident among the CesA mRNAs First transcript levels of CesA12 fluctuated markedly and independently of the other family members. The basis of this is unknown, but
72 indicates a clear contrast to the consistent co regulation of CesA10 CesA11 and CesA12 in planta With the exception of CesA12 e xpression of other family members did not increase markedly until 36 hours when mRNA levels of CesA3 CesA6 and CesA10 rose appreciably ( Figure 3 3A) Again, these results were unexpected since these genes were not co expressed in planta It is possible that extreme stress and/or starvation causes co regulation of specific family members to for m CESA complexes with specialized functions. Since these genes were upregulated shortly before loss of detectable mRNAs the expression of these CesAs may have resulted from one or more stress signals and/ or been part of a cell death program. Previous work has provided evidence that programmed cell death is controlled by a signal that concomitantly induces secondary cell wall synthesis (and presumably upregulation of associated genes) (Groover and Jones, 1999). Similar results were observed during cell wall regeneration in media wi thout sucrose. Under complete starvation, s ufficient levels of RNA could not be recovered after 28 hours of cell wall regeneration S trong upregulation of any CesA gene was observed at only 12 hours ( Figure 3 3B) which we attribute to duration of cell viability. With the exception of CesA3 the identity of the CesAs upregulated at 12 hours with 0% sucrose differed from those upregulated at 36 hours under with 0.5% sucrose. This may indicate that the CesAs are diffe rentially regulated depending on the availability of nutrients Also, under both reduced and no sucrose conditions, CesA12 transcript levels varied more often and with greater magnitude than other family members. This may indicate that regulation of this gene is more sensitive to the sugar status of the environment, and could have a specialized function when carbohydrates become limiting. Cellulose Synthase Expression in Response to Simulated Wounding/Pathogen Infection Since cell wall removal and protopl ast formation may involve multiple stresses, we sought to determine the extent of their contribution by testing treatments intended to simulate
73 exposure to fungal elicitors, wounding, and/ or oligosaccharides released from the native wall as a result of pathogen infecti on. Plants exposed to the aforementioned situations and/or molecules typically undergo a defense response (Cordero et al., 1994; Seifert and Blaukopf, 2010) thus altering gene expression. Transcript levels of all genes increased in the den atured enzyme treatment, having expression peaks at 2 hours and then rap idly declining (Figure 3 5 ) Although different treatments ha d different effect s on gene expression of the cellulose synthase s and MPI most changes were either modest, transient, or b oth indicating that the CesAs respond weakly to the treatments applied Complete removal of the cell wall could have a more severe effect on expression of the CesA s than was evident in these cell culture experiments, since treatments did not include active cell wall degrading enzymes. Cellulose Synthase Expression During Induction of Secondary Cell Wall Synthesis Secondary cell wall deposition was induced w hen suspension cells were transferred to medium optimal for this pu rpose in maize. After 10 days of incubation, c ells began to differentiate into tracheary element like cells ( TELs ) ( Figure 3 6). Direct quantification of differentiat ed cells was complicated by cell aggregation, but visual quantifications indicated that approximately 20% of cells had become TELs Transfer to induction media slightly increased mRNA levels of nearly all CesA but by 7 days after transfer (DAT), levels of CesA10, CesA11 and CesA12 mRNAs rose strongly ( Figure 3 7) This upregulation persist ed until 25 DAT when cells appear ed to have entered a stationary phase Visible differentiation of cells into TELs was not evident until 10 to 14 DAT, but had presumably begun considerably earlier. T he strong upregulation of CesA10 CesA11 and CesA12 at 7 DAT, followed by a lesser degree of upregulation at subsequent time points, may indicate that transcript and/or protein turnover rates for these genes may be low under the given conditions. In addition, many of the cells capab le of differentiating in to TELs may have done so relatively early and subsequently underwent
74 programmed cell death, as has been previously reported (McCabe and Leaver, 2000; Ohashi Ito et al., 2010). Considering that t he concomitant induction of secondary cell wall synthesis wit h the increase in transcript levels of CesA10 CesA11 and CesA12 is consistent with in vivo observations indicating that these genes are primarily, al though not exclusively involved in secondary cell wall biosynthesis. Methods Generation of the Suspension Cell Line To initiate a fresh, maize suspension cell line for these experiments kernels from inbred B104 inbred plants were harvested at 10 to 13 days after pollina tion and surface sterilized in 3% NaClO for 10 min. Embryos were dissected from the kernel under sterile conditions and those between 1 mm and 2 mm were transferred to 100 x 15 mm petri plates An N6 callus initiatio n medium (pH 5.8) was prepared by combining 4 g/L N6 salts (Chu et al., 1975), 1 mL/L (1000X) N6 vitamin stock 2 mg/L 2,4 D, 100 mg/L myo inositol, 2. 76 g/L proline, 30 g/L sucrose, 100 mg/L casein hydrolysate, and 2.5 g/L phytagel. Media w ere autoclaved before adding filter sterilized (0.2 m filter) 1 M silver nitrate to a final concentration of 25 M. Samples were subcultu red onto N6 medium as necessary (depending on growth), and Type II, friable calli were selected for additional subcultures After three months of sub culture, Type II, friable calli were transferred to autoclaved liquid regeneration med ia containing 20 g/L sucrose, 4.4 g/L Murashige & Skoog MS Medium with Vitamins (RPI Corp. Cat# M10400), and 2 mg/L 2,4 D at pH 5.8. Suspension cells were grown in darkness at 25C in 250 mL flasks with 50 mL medium agitating at 120 rpm. Cell lines were s ubcultured (using 10 mL of c ells condensed by sedimentation) every 10 days, selecting for cells that f ormed the small est aggregates by drawing cells from the top layer after sedimentation After three months of subculture, approximately
75 10% of cell lines h ad beco me acclimated to the liquid environment. A rapidly dividing cell line that did not form large aggregations was selected for subsequent studies. Protoplast Isolation Regeneration and Characterization M aize suspension cells derived from B104 embryos were used as the source for generation of protoplasts. Suspension cells (10 DAT) were briefly dried on sterile Whatman paper to remove excess moisture (Schleicher and Schuell CAT# 057145) and 3 g were transferred to sterile Erlen meyer flasks containing 200 mL ( cell wall ) digestion media (pH 5.8) containing 1.5% Onazuka RS cellula se (Yakult, Tokyo) 0.3% R 10 M acerozyme (Yakult, mercaptoethanol, and 1 g/L BSA Cells were incubated at 25C for 8 hours in the dark, shaking at 40 rpm. Protoplasts were released by agitating at 80 rpm for 5 min followed by filtering through 35 Nitex nylon mesh (Sefar Prod. Ref. 03 35/16). Protoplasts were col lected by centrifu g ation at 150 x g, retrieving the pellet and resuspending for washing with regeneration media (described above for suspension cells) This washing process was repeated three times. For subsequent p urification protoplasts were resuspended in 2 mL of 0.6 M mannitol, then layered on 5 mL 0.6 M sucrose in a 10 mL falcon tube. After centrifugation at 225x g for 10 min, floating protoplasts were collected and transferred to regeneration media. The experiments involving cell wall regeneration in reduced sucrose used the same regeneration media described above f or suspension cells, except that only 5 g/L or 0 g/L of sucrose were used instead of 20 g/L Cell wall regenera tion was monitored at 4, 12, 20, 28, 36, 44, and 60 hours after trans fer by o bservation under bright field and fluorescent microscopes. Samples were also harvested at these time points for qRT PCR analysis of cellulose synthase m RNA levels Cellulose deposition was visualized under UV (365 nm) after staining with one drop of calcof luor white (Sigma Aldrich Prod.# 18909) then adding one drop of 10% potassium hydroxide. Bright field i mages were
76 obtained using an RT SPOT camera (Diagnostic Instruments Sterling Heights, MI ) mounted on an Olympus BH2 light microscope and fluorescent microscopy i mages were captured with an Evo lutionMP camera (Media Cybernetics Bethesda, MD ) mounted on an Olympus BX51 fluorescent microscope. Simulation of Wounding/Pathogen Infection Suspension cells (10 DAT) were transferred to three different solutions inten ded to simulate wounding and/or pathogen infection. S uspension cells (described above) were weighed after blotting to remove excess media, and 1 g samples were transferred to 50 mL of experimental media Each of these treatments included reg eneration media with either 1) boiled cell wall degrading enzymes ( 0.25 g of boiled (5 min) RS cellulase (Yakult, Tokyo) and 0.25 g of boiled (5 min) R 10 macerozyme (Yakult, Tokyo) 2) ground cells ( 0.5 g of whole cells ground in liquid nitrogen ) or 3) c ell walls ( 0.5 g purified cell wall s ground in liquid nitrogen ) Suspension cells were incubated in darkness at 25C shaking at 120 rpm. S amples were harvested at 2, 6, and 12 hours for qRT PCR analysis. Cell walls were isolated by fine ly grinding 0.5 g samples of leaf tissue in liquid nitrogen, the n adding 9 mL 1% SDS. Samples were the n heated to 80C for 15 min and centrifuged at 3 500 rpm for 5 min. Pellet s were washed with 80C water and recentrifuged a total of three times Pellets were ground again in 2 mL water before washing three times in 70C water, three times in 50% ethanol at 70C, and then three more times in 70C water. Induction of Secondary Cell Wall Biosynthesis S uspension cells (10 DAT) were weighed after briefly blotting away excess m oisture, and 1 gram aliquots were transferred to 250 mL flasks with 50 mL of induction media (pH 5.8) containing 20 g/L sucrose, 4.4 g/L Murashige & Skoog MS Medium with Vitamins (RPI Corp. Cat# M10400), 0.2 mg/l 2,4 D, 125 mL 8 M brassinolide, 1 mg/L BAP (cytokinin), and 10 mL
77 of 1 M boric acid. Flasks were incubated in darkness at 25C sh aking at 120 rpm. Samples were harvested at 7, 12, 20 and 25 DAT for qRT PCR analysis of CesA mRNA levels Images were captured with an RT SPOT camera (Diagnostic Instruments) mounted on an Olympus BH2 light microscope. Quantitative R eal Time RT PCR Levels of mRNA were quantified as described in the methods section of chapter 2.
78 Figure 3 1. Diagram of the protoplast regeneration system. Top; diagram representing the protoplast regeneration system Grey ovals represent organelles in illustrations corresponding to protoplasts, and cell walls are depicted in green. From left to right; Suspension cells a re stripped of their cell walls during 8 h i ncubation in digestion media ; As protoplasts regenerate they begin to stain for cellulose with calcofluor white at 12 h. By 20 h cells ass ume non spherical shape s form aggregations and fluoresce strongly, indicating continued cellulose deposition Photos by Brent
79 Figure 3 2. Relative expression of select CesA family members and sucrose synthases during cell wall regeneration by protoplasts The x axis represents time since transfering nacent pr otoplasts to regeneration media. Suspension cell s represent mRNA sampled (from 10 day old cell cultures) immediately before an 8 hour cell wall digestion (indicated by the break) Expression levels are relative to the amount 18SrRNA measured in each tissue.
80 Figure 3 3. Sugar responsive expression of t he CesA family during cell wall regeneration by protoplasts. A) Expression in media containing reduced sucrose (0.5% w/v). B) Expres sion in media without sucrose. Protoplasts were sampled at 8 h our intervals Expression levels are relative to the amount 18SrRNA measured in each tissue. Data represent three biological replicates.
81 Figure 3 4 Expression of the systemic wound/pathogen response gene Maize Proteinase Inhibitor ( MPI ) in suspension cells (SC) and through 44 hours of cell wall regeneration in protoplasts (after an 8 hour digestion) Expression levels are relative to the amount 18SrRNA measured.
83 Figure 3 5. Relative e xpression of CesA genes and the sys temic wound response gene MPI (Maize Proteinase I nhibitor) in response to simulation of wounding and pathogen infection. The x axis represents time after transfer of suspension cells (at 7 days after transfer) to fresh regeneration media with the specified treatment. Ground whole cells were in tended t o mimic mechanical damage whereas denatured digestion enzymes and purified, partially digested cell wall treatments were intended to simulate pathogen infection. Graphs were not designed for comparison of genes, but to show changes in expression ov er time for each gene tested. Expression levels are relative to the amount 18SrRNA measured in each tissue. Data represent three biological replicates.
84 Figure 3 6 Differentiation of suspension cells to tracheary element like cells after secondary cell wall induction. Maize suspension cells were transferred from regeneration media to either control media or induction media. A) The control group was transferred to normal regeneration media. Scale bar equals 0.15 mm. B) The experimental group was tra nsferred to inductio n media containing 1% sucrose, 1 M brassinolide, 10 mM boric acid, 0.1 mg/l 2 4 D, and 0.5 mg/l cytokinin (BAP) Scale bars equal 0.15 mm Photos by
85 Figure 3 7 Relative e xpression of CesA gene family members during secondary cell wall biosynthesis. Expression of the CesA family after maize (embryo derived B104) suspension cells are transferred to control (untreated) culture m edium. In the induction medium (1% sucrose, 1 M brassinolide, 10 mM boric acid, 0.1 mg/l 2 4 D, and 0.5 mg/l cytokinin [BAP] ) expression levels of CesA10, CesA11 and CesA12 are highly upregulated relative to the control. Expression levels are relative to the amount 18SrRNA measured in each tissue. Data represent three biological replicates.
86 CHAPTER 4 CHARACTERIZATION OF MAIZE CESA MUTANTS FROM THE TRA NSPOSON MUTAGENIC UNIFORMMU POPULATION Background Transposable elements (TEs) or transposons, are mobile genetic elements of varying size (pieces of DNA or RNA) that have the capacity to move to new sites within the genome. The Mutator ( Mu ) class of these elements typically insert s in the coding sequence of genes rather than in the intergenic regions and thus provide a natural mechanism for mutagenesis. Furthermore, transposable elemen ts can play major role s in genome evolution, because in addition to direct effects of insertion on loss of gene function, transposon action can alter promoters, change intron exon splice junctions, and modify terminators (Bennetzen, 2000). Tran s posons fro m diverse families (especially retrotransposons) are estimated to make up 85% of the maize genome, and evidence indicates t hat activity of functional transposon systems is variable in different tissues ( Vicient, 2010). Since the discovery of transposons in the 1940 s (McClintock, 1947), many types have been characterized, and the y can be designa ted as either class I or c lass II depending on whether they use RNA (class I) or DNA (class II) as a transposition intermediate (Wicker et al., 2007). Trans posons f rom both classes can also be categorized as either autono mous which contain sequence that encodes protein( s ) necessary to for transposition, or non autonomous which contain only the cis elements required for recogniti on by the transposition protein( s) T he transposition protein(s), or transposases, of autonomous elements must be present in a system for non autonomous elements to transpose. Many transposons have b een well characterized in maize, particularly because of their usefulness to genetic studies. Especially prominent among them are the Mu tra nsposable elements which include 12 sub families designated Mu1 to Mu12
87 designated based on differences between characteristic s e quences at their and ends Most transposons are define d, in part, by the presence of terminal inverted repeat (TIR) sequences at both of their ends ( Robertson, 1978; Dietrich et al., 2002). Within the Mu family, distinctive TIRs are highly conserved especially within each of the Mu subtypes (Brutnell, 2002). The TIRs are roughly 215 bp in length and contain the recognition sequence for the Mu transposase (Lisch, 2002 ; Diao and Lisch, 2007 ). The vast majority of Mu TEs are non autonomous, but the Mu9 or MuDr TEs encode a transposase, and are thus autonomous. In addition to the transposase gene ( Mu rA ), Mu9 also includes a MurB gene Although MurB function is unknown transgenic approaches and experiments with deletion derivatives have provided evidence that MurB is necessary for Mu transposition, possibly aiding stability of transposase function (Walbot and Rudenko, 2002 ; Diao and Lisch, 2007 ). Another characteristic of Mu TEs that make them useful in genetic studies is the high r ate of forward mutation that they confer. They are m ore active than any other class II TE in plants (Brutnell, 2002; Lisch, 2002). P otentially high mutation rates can thus result, depending on the number of autonomous and non autonomous that are present In addition, Mu elements can insert virtually anywhere in the genome, although evidence indicates a propensity for insert ion in the region of genic sequences ( Dietrich et al., 2002 ; Liu et al., 2009; Vollbrecht et al., 2010). Considering these characteristics, as well as the conservation of Mu TIRs, it is not surprising th at Mu has been the TE family of choice for mutagenesis and transposon tagging research Such efforts have included the Trait Utility System for Corn (TUSC) developed by Pioneer Hi Bred Int (Meeley and Briggs, 1995; McCarty and Meeley, 2009), the Maize Targeted Mutagenisis (MTM) population developed at Cold Springs Harbor (May et al., 2003), RescueMu project (Walbot, 2000; Raizada et al., 2001), the,
88 Photosynthetic Mu Carrier et al., 2010), and UniformMu population ( McCarty et al., 2005; Settles et al., 2007) The UniformMu maize population was created by introgressing an act ive Mu line called maize inbred a highly mutagenic line containing a bz mum9 color marker. Kernels with the bz mum9 gene, in the absence of MuDR, are bronze whereas W22 kerne ls are dark purple. When MuDR is present with bz mum9 kernels are bronze with varying amounts of small purple spots Having this selectable marker provides a convenient visible means of determining whether a specific line contains an active MuDR. This is important because in the absence of a functional MuDR mutations in a line will be stable and their effects can be readily studied. A lso, a ny (non somatic) mutations in this family will be heritable (Chomet et al., 1991). For this reason, bronze kernels are chosen to allow selection for both mutagenic and stable lines in each new generation of the UniformMu population A key issue has been the exclusion of non heritable somatic mutations in the database of m utations contained within any given line. Maintenance of the UniformMu population also includes backcrossing new lines to the W22 inbred. This ensures a steady st ate mutagenesis, prevents build up of ancestral mutations, and maintains a uniform background (McCarty et al., 2005). Sites of Mu insertions, which allow identification and mapping of mutations within the UniformMu population were originally determined by sequencing clones generated through thermal asymmetric interlaced (TAIL) PCR. This method used primers designed to anneal to the conserved TIRs, coupled with arbitrary degenerate primers to amplify sequences flanking inserti ons (McCarty et al., 2005). This method was successful in identifying numerous insertion sites, but it did not capture all insertions caused by Mu The basis of this was unclear, but adaptation of next generation sequencing methods for Mu flanks has proven effective.
89 Sequencing platforms such as 454 (long read) ( Margulies et al., 2005 ), and Illumina (Illumina Inc., San Diego CA) have since been used, and have provided higher throughput as well as greater coverage of insertion sites. Methods are currently b eing refined for additional sub groups of that had not been previously recognized ( C. Hunter, unpublished data). The defining features of the UniformMu maize population are 1) high mutation frequencies, 2) low mutant load 3) a selectable color marker for MuDR transposase activity, 4) moderate total Mu TE copy number, 5 ) a database of pedigreed insertions, and 6 ) a unifor m background in plant material (McCarty et al., 2005). The uniform background has been especially valuable in that it provid es uniform controls for comparative analyses of newly generated mutants. Here, we utilize d the UniformMu population to characterize mutations in the cellulose synthase gene family ( CesAs ) We employed a reverse genetics appr oach in which the identity of a disrupted gene is known, and analysis of resulting mutant plants allows the functional role of this gene to be addressed. Forward genetics was also used to determine the causal mutations for lines having phenotypes possibly based on defective cell wall biosynthesis. Results thus far have not associated these phenotypes with known genes, but their study is ongoing (Appendix A). Results Reverse Genetics and the UniformMu CesAs To date there are no reports of phenotypes arising from mutations of CesA genes in maize To address the question of whether maize CesA mutants can cause a visible phenotype, we searched the UniformMu population for lines carrying insertions in or near Ces A genes. In total, 16 alleles wer e analyzed, as well as a double mutant of CesA7 and CesA7 a (Table 4 1) The only family members in which no insertions have yet been identified were CesA1, CesA3 and CesA5 If mutations in these genes condition a male let hal phenotype (i.e. defective pollen and/or pollen tube), the absence of associated insertions in the
90 UniformMu population can be explained by the strategy used for generating and maintaining (UniformMu) lines, where pollen from mutagenic parents was applied to W22 females. In most other instances there were two alleles available and examined for each family member, although CesA7 a, CesA10 and CesA11 were represented by a single family member each (Table 4 1) The location of insertions included exo ns (5 to s At least 30 plants were examined from lines carrying each of these CesA insertions Plants were grown under field conditions at the UF Plant Science Research Unit (Citra, FL) during spring and fall planting seasons begi nning in spring 2008, and continuing through fall 2011. New mutants were acquired during the course of this period and each was added to ongoing investigations. Phenotypic analyses were focused using two overall criteria. First, for a given CesA mutant, we looked at the plant tissues and developmental stages where maximum expression was observed for the CesA in wild type plants o ur hypothesis being that loss of function mutants would show visible phenotypes at these locations (Table 4 2) Second, we watc hed for responses similar to those observed in other species (mainly Arabidopsis and rice) if any loss of func tion mutations had been identified in similar CesAs (Figure 4 1) Ultimately, no visible phenotypes were clearly associated with any of these muta nts. However, a recently obtained cesA11 line segregated for a phenotype that has been previously described by Postlethwait and Nelson (1957). Characteristics of this mutant are similar to the brittle culm mutants of rice and include; severely reduced biomass, brittle stalk and leaf tissue, delayed development, and chronically wilted leaf tips (Figure 4 2 ) Preliminary PCR genotyping has indicated that these plants are heterozygous for cesA11 but this result may be due to the maize genome havi ng two identical copies of this gene.
91 Discussion Reverse genetics and the UniformMu CesAs The lack of visible phenotypes associated with the 16 Mu insertions in maize CesA s examined thus far may be due to several factors. First, the extent of gene duplications in the maize genome, and especially the CesA family may buffer the species from s evere consequences of otherwise deleterious mutations P aralogs, or even more distantly related family members may be functionally re dundant, in which case mult iple null mutants might need to be created before a visible phenotype is apparent This is a strong possibility considering a similar situation has been observed with the Arabidopsis CesAs (Desprez et al 2007). T he lack of phenotype observed here for ma ize cesA12 mutants, and complexities associated with co segregational analysis of c esA11 may both be due to the presence of nearly identical paralogs Such paralogs may compensate to varying degrees for the loss of function mutation. The high degree of sequence similarity also poses considerable Another possible explanation for the lack of phenotype in certain lines could be the location of insertions. Transposons residi ng in introns may simply be spliced out of the gene without causing any disruption particularly if the Mu insertion is not a large one In addition, insertions in the untranslated region can sometimes only partially disrupt transcription resulting in function sufficiently, and therefore do not condition a phenotype. In several instances, no heritable homozygous insertions could be obtained for the alleles examined which could be due to wild type copies of near identical paralogs or lethality of the homozygous state, as wa s observed for some alleles in A rabidopsis ( At cesA1 At cesA3 ) Considering that CesA mutants in other species are recessive, the possibility exists that maize phenotypes may be evident in those homozygous mutants not yet accessible at the time of the present study.
92 Although we are unable to determine (through PCR) if cesA11 is the causal insertion for the mutants segregating in that line, similarity of the mutant phenotype to b rittle culm mutants in rice indicates that cesA11 is a good candidate. In addition, three of the brittle culm mutants result from disruption of the rice CesAs ( OsCesA4 OsCesA7, and OsCesA9 ) involved in secondary wall synthesis (Tanaka et al., 2003) one of which ( OsCesA4 ) is homologous to ZmCesA11 Phenotypic characteristics shared between brittle culm mutants and those in the cesA11 line include severely reduced biomass, and brittle stem and leaf tissue. In addition, the mutants segregating in the cesA11 line examined here had chronically wilted leaf tips. If cesA11 is the causal mutation, wilting may be a result of irregular xylem preventing adequate water transport. This hypothesis seems reasonable considering that maize is a panicoid, C4 grass wi th a larger architecture, more rapid growth rate, and a higher rate of transpiration than rice. Considering that the maize genome contains two, identical copies of CesA11 (both from the Maize I genome) additional approaches will be needed for future work, including restriction enzyme digestion followed by Southern blotting quantification of CesA11 mRNAs in mutants and possible re sequencing of selected BAC regions will be necessary to determine the relationship between the two CesA11s and whether one of them is associated with the mutant phenotype Mutations in any of the secondary wall associated CesAs in Arabidopsis also lead to reduced biomass and irregular xylem formation. Other phenotypic features of CesA mutations in Arabidopsis CesAs that could em erge in maize homologs include 1) the embryo lethality and radial root swelling (that occurs under high temperature ) in AtcesA1 2) defective pollen formation in both AtcesA1 and AtcesA3 and reduced root elongation in AtcesA6 (Figure 4 1) The other Arabidopsis CesAs are functionally redundant, with phenotypes arising only in double or triple mutants This scenario is likely to apply to maize as well, and may explain the lack of phenotypes in several of the mutants reported here.
93 Methods Plant Materia l All mutant lines were obtained from the UniformMu population at the University of Florida. Plants were grown under field conditions at the UF Plant Science Research Unit (Citra, FL) during spring and fall planting seasons beginning in spring 2008, and c ontinuing through fall 2011. Lines have been, or are in the process of being backcross ed to W22 to separate the mutation and/or phenotype of interest from other i nsertions within a given line. Map l ocations of insertion sites associated with the CesAs wer e determined by aligning flanking sequences to gene models through the MaizeGDB website (maizegdb.org). Genotyping Presence of insertions in lines potentially harboring CesA mutants was determined by PCR, using primers that target the TIR sequences of Mu transposons in combination with forward and reverse orientated, gene specific primers.
94 Table 4 1. Cellulose synthase mutants identified in the Unif ormMu population for reverse genetics screening. Homozygous mutation (+/ ) Phenotype (+/ ) Allele/s Insertion location CesA2 CesA4 CesA6 CesA7 CesA7 a CesA7xCesA7 a CesA8 CesA9 CesA10 CesA11 CesA12 + + + + + + + + + + + ? ? ? ? +/? ? ? mu1005736 mu1015515 mu1016007 mu1041689 mu1040612 mu1037352 mu1016043 454AC190931 mu1016043 454AC190931 mu1011353 mu1039702 mu 1019324 mu1031574 mu00193 mu1007966 mu1039500 mu1041598 exon exon intron intron exon exon exon exon exon intron exon intron At least 30 plants were grown and characterized from each of the segregating families carrying a given insertion. Material was grown to maturity at the UF Plant Research Unit (Citra, FL) and examined closely for visible phenotypes. Presence of the insertions was confirmed by PCR with TIR specific and gene specific primers, as was the zygosity of insertions. In lines heterozygous for the Mu insertion the possibility of a mutant phenotype could not be determined. Lines that clearly do not have a phenotype are highlighted.
95 Table 4 2. Sites of maximum expression for the maize CesAs at each developmental stage. Gene Seedling Maximum Expression Vegetative Maximum Expression Anthesis Maximum Expression CesA1 CesA2 CesA3 CesA4 CesA5 CesA6 CesA7 CesA7 a CesA8 CesA9 CesA10 CesA11 CesA12 Coleoptile Coleoptile Coleoptile Coleoptile Coleoptile Coleoptile Primary root Primary root Coleoptile Coleoptile and primary root Primary root Primary root Primary root Stem Stem Stem Stem Stem Stem Stem Stem Stem, node, and root Stem Stem Stem Stem Stem Developing prop root Developing prop root Ligule and midrib Husk and developing prop root Lat. root initiation zone Husk, developing prop root, and cob Husk and cob Husk and developing prop root Developing prop root Developing prop root Mature prop root, husk, and stem Mature prop root, husk, and stem Mature prop root, husk, and stem Sites of maximum expression were focused on when appraising mutants for phenotypes.
96 Figure 4 1 .Unrooted, neighbor joining, interspecies phylogenetic tree with descriptions of cesA mutant phenotypes. Related cellulose synthases from maize (red), rice (blue), and Arabidopsis (green) are represented by six clades, with four Arabidopsis genes forming a sub clade. Phenotypes of characterized cesA mutants are described in the boxes. The radial swelling 1 1 isoxaben resistant 1 and procuste mutants are represented by AtcesA1 AtcesA3 and AtcesA6 respectively. The tree was constructed using pairwise de letion with 1,000 bootstrap replications in a MEGA4 analysis.
97 Figure 4 2. The mutant phenotype possibly associated with CesA11 Mutants are phenotypically identical to the wilty1 mutant described in 1957 by Nelson and Postlethwait, having chronical ly wilted leaves, reduced bi omass ( relative to the wild type), and brittle tissues.
98 CHAPTER 5 SUMMARY AND FUTURE D IRECTIONS A two fold rationale motivated this investigation of the evolution and regulation of genes for cellulose biosynthesis in maize ( Zea mays L.). The first was the importance of cellulose to both plant biology and human needs. The prominence of cellulose in the cell wall of virtually every plant cell is fundamental to the strength and architecture of plants, and also makes cellu lose the most abundant polymer on earth. As such, its uses by humankind h as extended from food and fiber, to the rapidly increasing demand for bio fuels. Recent research has thus focused intensely on the genetics, and regulation of cellulose formation yet this area has remained notoriously challenging and complex (Taylor et al., 2003 and 2008; Tanaka et al., 2003, Persson et al., 2007; Burton et al., 2010; Fujii et al., 2010). The second motivation was that maize provides not only a valuable model for the grain and grass type cell wall, but also constitutes the largest single crop in the nation and world. Here we 1) investi gate the structure and evolution of the maize CesA gene family, 2) analyze CesA expression in planta and use suspension cell and protop last systems to test responses to experimental perturbations, and 3) characterize cesA mutants in maize Bioinformatic analyses included the identification of four, previously undescribed CesA paralogs This indicated that the maize CesA family may be more functionally redundant than that in many other species possibly leading to enhanced expression. In addition (or alternatively) the prevalence of paralogs could also increase the robustness of the maize CesAs by al lowing subfunctionalization of dupli cated genes. We also show that the maize CesAs group into six distinct subclades, consistent with phylogen et ic analyses of CesAs from other plant species (Carrol and Specht, 2011). These subclades can be further divided, with three containing primary wall associated CesAs, and three having members that are predominantly involved in
99 secondary cell wall synthesis. This division of subclades may reflect the necessity for three different CesAs to be present for assembly of functional CESA complexes, regardless of whether these complexes are associated with primary or secondary wall synthesis ( Taylor et al, 2003; Persson et al, 2007 ). Our results also show that at least one, each unique CesA gene maps to chromosomal regions previously determined to predominate in gene retention. Conversely, genomic segments associated with enhanced gene degradation contain only CesAs that are paralogs. These observations are consistent with a recent hypothesis (Sc hnable et al., 2011) proposin g that after a genome duplication event (such as occurred in maize) resistant to gene degradation. The mechanisms behind this observed bias in gene retention remain unclear, however some f orm of epigenetic labeling of one genome copy is likely. We also observed that one paralog was retained from each of the subclades associated with primary cell wall synthesis, and expression analysis of these paralagous pairs showed there was little or no subfunctionalization These results are consistent with the Gene Balance hypothesis proposed by Birchler and Veitia (2010 ) that suggests gene retention and loss will be constrained by the stoichiometry of isoforms in multi subunit complexes where this balance affects the function of the whole It thus follows that if essential members from each of these subclades were duplicated, then essential paralogs from each subclade would be retained due to the selective advantage of preserving the bala nce of potentially interacting isoforms required for proper CESA complex assembly. I n planta analyses revealed a high level of coordinate expression among members of the maize CesA gene family However, clusters of coordinately expressed genes varied in di fferent tissues and at different stages of development. Three groups of family members that clustered together based on similarity of overall expression profile s indicated a co regulation of genes that
100 contributed to cellulose synthesis at a given developmental stage. Clustering analysis showed that CesA10 CesA11 and CesA12 were th e only family members to exclusively and consistently group together based on expression pattern. In addition, th is group of genes invariably showed the tightes t correlation among any family members tested, and they were most highly expressed in tissues undergoing secondary cell wall biosynthesis. Furthermore, when suspension cells were induced to differentiate into (secondary wall rich) tracheary element like ce lls, only these genes were highly upregulated relative to untreated control s Together these data indicate that CesA10 CesA11 and CesA12 function predominantly in secondary cell wall deposition, supporting the subdivision of CesA clades. Expression patt erns of diverse maize CesAs during cell wall regeneration by protoplasts generally followed comparable trends, maintaining relatively high mRNA levels that peaked at 44 h. Unexpectedly, three family members that did not appear to be co regulated in planta had similar profiles during wall regeneration indicating that atypical clustering modules are possible under extreme conditions. Additionally, the responses of CesA gene family members during wall regeneration with little (0.5%) to no sucrose further su pport ed plasticity in the co expression modules observe d for the CesA genes Also, transcript levels of CesA10 CesA11 and CesA12 were comparable to other family members even though protoplasts do not form secondary cell walls. These results could be expl ained by two scenarios. First protoplasts may initially form CesA10 CesA11 and CesA12. S econd removal of cell walls could elicit stress and/or defense response s, which in turn induce CesA10 CesA11 and CesA12 To investigate how altering normal CesA expression affects plant morphology and development, a reverse genetics approach was used. Homozygous mutations in seven primary wall associated maize CesAs a nd one double mutant did not show a readily visible
101 phenotype. This may have been due to the prevalence of paralogs in the maize genome, which would add to the high level of functional redundan cy reported for some of the primary wall CesAs in Arab idopsis ( Burton et al., 2004; Persson et al, 2007) In addition, four of the possibly leaving gene function at least partially intact in some of these instances Overall, research described here has shown that groups of CesA gene family members are co expressed, with subgroups being fine tuned to perform specific functions at different developmental stages. Also, the evolution and structure of the CesA family indicates that duplicat ions of certain CesAs associated with primary wall synthesis were retained without subfunctionalization, and this proliferation of CesA family members in maize is reflected by the high level of functional redundancy observed in cesA mutants lacking a visib le phenotype. Work presented here, together with the genetic lines and molecular materials developed during this research will provide a valuable foundation for future studies. These future directions can range from short to long term objectives as foll ows. First, combinations of double and even triple mutants can be generated from primary wall associated cesA mutants produced during these studies. Such materials would provide a means to determine which of the CesA family members we re functionally redundant. Second, further analysis of single and double mutants generated here may also be productive. L ines having homozygous insertions in CesAs associated with secondary wall formation have yet to show a clear association with a mutant phe notype in maize however w e have obser ved a phenotype showing Mendelian segregation in a line carrying an insertion in CesA11 Although we have validated the presence of a cesA11 insertion, the identical CesA11 paralogs appear to have prevented accurate genotyping of segregating families by obscuring the presence of homozygous mutants The nature of the phenotype is promising, however, with
102 characteristics similar to homologous mutations in rice and Arabidopsis ( Taylor et al., 2003; Tanaka et al 2003). These shared features include severely reduced biomass, brittle tissues, and stunted development. In addition, the maize mutant also has chronically wilted leaves, which would be an expected result of reduced water transport accompanying malform ed xylem tissue A phenotypically similar mutant with abnormal xylem structure was observed in the maize wilty1 mutant (Postlethwait and Nelson, 1957) however wilty1 does not map near either of the Cesa11 locales Future work will require alternate approa ches to determine if the cesA11 mutation is indeed causal for the wilty phenotype observed These approaches would i nclude back crossing the mutant line to the W22 inbred ( recently completed ) to reduce numbers of other possible mutations segregating in subsequent F 2 progeny (in progress). Another approach will be to quantif y cesA11 mRNA levels of the mutant relative to a wild type sibling to determine if there is a correlation between reduced cesA11 expression and the mutant phenotype. Finally, restrict ion enzyme digestion followed by Southern blot analysis can be used to determine if there is a Mu insertion in either of the cesA11s. If successful, this work will test the hypothesis that the role of a maize CesA11 is essential to secondary cell wall form ation and stalk integrity, as well as showing that regulation of the CesAs extends beyond the identical coding and upstream sequence of these two genes. Third, future work can further develop the forward genetics approach initially taken to identify gene s with critical roles in cell wall biosynthesis by prioritizing lines based on relevant phenotypes Three lines were selected that had readily visible phenotype s as well as insertion/s in genes that could affect cell wall biosynthesis. These lines were nam ed shredded (for its shredded leaves) (Figures A 1 and A2), flecked (for its anomalous leaf pigmentation), and epuf1 ( for a kernel phenotype that results when the fruit wall (pericarp) is left empty looking by minimal development of the single seed within it ) (Figure A3) We have yet to identify the causal genes
103 for these phenotypes through conventional PCR, however high throughput sequencing technology has provided an alternative approach. Future efforts can effectively use Illumina based sequenci ng to identify Mu insertions in mutants and wild type siblings from each line. Flanking sequences from wild type siblings can be subtracted ( in silico ) from those of mutants, thus providing a list of potential candidate genes. Association of these candidat es with the mutant phenotype of interest will then need to be validated by PCR and co segregational analysis If causal genes for any of these three mutants can be identified, then essential roles can be ascribed to their sequences, and further defined by more in depth analysis of their phenotypes. Longer term directions that would be facilitated by this work include testing functional associations between isoforms that may form complexes as indicated by co expression modules revealed here. Such work would likely require concomi tant expression of maize CesAs in either an orthologous system, or in a plant system containing an inducible mechanism to silence expression of its own CesA family members. In addition, the Gene Balance hypothesis, which we suggest may be governing the retention of specific CesA paralogs, can be tested by transforming maize with additional CesA result.
104 APPENDIX A FORWARD GENETICS ANA LYSIS OF THREE MAIZE MUTANTS Forward Genetics Mutants with phenotypes possibly caused by defective cell wall biosynthetic genes were selected from segregating lines in the UniformMu population for further investigation using a forward genetics approach. Lines segregating for mutant phenotypes were ide ntified in the field or from ears after harvest, and prioritized for further analysis depending on whether they contained insertion s in cell wall associated genes. Lines selected include a shredded line (05S 2500), identified by Don McCarty a flecked line (11S and an empty pericarp UF 1 ( epuf1 ) line (09S 3300), identified by Karen Koch and initially characterized by Stephanie Marunich and Gregorio Hueros The shredded line was named for its shredded leaves, the flecked line was named for its anomalous leaf pigmentation, and the epuf1 line for a kernel phenotype that results when the fruit wall (pericarp) is left empty looking by minimal development of the single seed within it. These mutant phenotypes were characterized to an initial level, and mutant sequences within them were tested for potential causality. The candidate insertions were identified by in silico subtraction of ancestral insertions from the total of those identified in parental lines. Characterization of the shredded Mutant The shredded phenotype was selected for analysis because the characteristic lesions that formed between major vascular bundles and eventually caused separation of tissue seemed likely to add insights into cell wall synthesis. These l esions extended to varying lengths along the longitudinal axis of a leaf, parallel to the midrib (Figure A 1). Leaves of shredded mutants thus had alternating vertical stripes of normal and affected tissue that were separated by vascular bundles (Figure A 2). Affected tissue progressed through three stages. First, previously unaffected, green tissue became chlorotic, then lost all color, turning white and eventually translucent (Figures A 1 and A 2). At the second stage of lesion formation, cell shape, cell size, and cell organization became abnormal (Figure A 2B), and affected vertical stripes showed wave like undulations (Figure 4 2A). In addition, aberrant stomatal distribution, as well as underdeveloped stomata were observed (Figure A 2B). At the third s tage of lesion formation, cells within the affected tissue separated, usually parting along a single vertical axis in a given region, opening splits in the leaf that ranged from a few centimeters long to the full length of the leaf blade. Leaves were often split severely enough to A 2A). When these leaves were stained with acetocarmine for contrast, unaffected areas stained pink and had a normal distribution of vascular bundles, whereas first stage lesion tissue did not take up the stain and had a reduced number of vascular bundles (Figure A 2C). Also, boundaries between normal and lesion tissues were clearly defined by vascular bundles (Figures A 2B and C). Other characteristics of the shredded mutant included a reduced plant size relative to WT (Figure A 1), and variable intensity of the phenotype that appeared dependent on environmental conditions. Families segregating for the shredded phenotype were genotyped to test for co segregation with Mu insertions in genes cons idered best candidates for causal mutations (17
105 genes with Mu insertions were tested), but co segregation analyses did not identify a causal insertion among these. This mutant has been further back crossed to the W22 inbred, and is being tested for presenc e of other Mu insertions not identified by earlier methods. The phenotype of the shredded mutant raised several questions about the developmental progression of stripe type lesions and the demarcation of boundaries between affected and unaffected tissues. The observation that cell size and shape were aberrant in affected tissue, indicates that the lesion phenotype probably results from defects in cell division and expan sion that arise early in leaf development. This hypothesis is supported by the observation that cells in affected tissue can be either smaller or larger than normal cells in unaffected, neighboring tissue, and that the cell files of affected tissue are hig hly disorganized (Figure A 2B). Additionally, distribution and development of stomata in pre lesion tissue was atypical, with fewer total stomata per unit area and several stomata being severely underdeveloped. Another interesting feature of shredded plan ts is that the borders between lesion tissue and normal tissue were defined by the large vascular bundles. This may indicate that the defective, early development of lesion tissue is the result of some mobile signal that does not pass readily beyond the la rger vascular bundles that delimit a given region (Figures A 2B and C). Another possibility is that the sugar status of affected tissues is responsible for abnormal development. The lack of minor vascular bundles and apparent degradation or incomplete form ation of large vascular bundles in pre lesion tissue (Figure A 2C) raises the question of whether the bundles failed to form, or formed correctly and were subsequently degraded. Alternatively, the stain used to show contrast between normal tissue and affec ted tissue may not have moved effectively into pre lesion tissue, thus giving a false negative result for the presence of vascular bundles. The causal insertion for this mutation has yet to be identified, and may be more readily accessed by methods curren tly under development. Initially, insertions in this line were identified by a TAIL PCR method, which identifies less than a f ull complement of insertions. A high throughput sequencing approach can now be used, although this too is PCR based and thus potentially sensitive to amplification biases that could favor some flanking sequences at the expense of others. Characterization of the bz linked flecked Mutant The flecked mutant was selected for analysis because several aspects of its phenotype suggested cell wall physiology might be altered. Its short stature is typical of mutants such as those in CesA mutants of rice and Arabidopsis (Tanaka et al., 2003; Taylor et al., 2003). Cell wall components can also affect development of lesions such as t hose seen here. Aside from growing to only half the size of WT plants, the flecked mutant is also characterized by slightly yellowish, drab olive colored leaves with a dull appearance on the adaxial side. As mutants matured, numerous, small (0.5 1 mm) yell ow spots developed on the leaves, similar to the phenotypes of lesion mimic mutants, but without the clearly defined lesion boundaries. Additionally, varying degrees of male sterility were evident in flecked plants. At anthesis, visible appearance of anth ers indicated that they were fully developed, but collapsed, and also devoid of pollen. In addition, genetic analysis of the flecked mutant indicated that the causal gene was linked to the bz color marker. Plants growing from bronze and purple kernels of a single ear from a self pollinated flecked parent showed a 100% correspondence between bronze kernels and the flecked phenotype (25 plants tested, 5 bronze). Thus far, genotyping and co segregation analysis
106 of candidate insertions has yet to identify the causal gene. However, candidate genes remain to be tested including those for a calmodulin binding protein ( GRMZM2G429807) and an MSF (mitochondrial stimulation factor) type transporter ( GRMZM2G104942). In addition, linkage of this mutation to the Bz gene should facilitate future analyses. The flecked mutant is particularly interesting because it is one of the few mutations to come out of the UniformMu population that is known to be linked to the bz color marker locus. This is a relatively recent obse rvation that will facilitate our search for the causal gene by allowing us eliminate candidates that are not on chromosome 9, or are too distant from the bz locus to be in linkage. Characterization of the empty pericarp uf 1 ( epuf1 ) Mutant (line 09S 3300) The epuf1 mutant was selected because empty pericarp (ep) phenotypes often have underdeveloped and/or aborted kernel tissues, which in turn may arise from defects in cell wall formation. As the name implies, epuf1 had kernels with empty appearing perica rps that, in this case, surrounded an underdeveloped endosperm and apparently intact aleurone (Figure A 3 A). Presence and location of starch in the epuf1 endosperm was solution. As in WT kernels, the epuf1 endosperm stained deep purple, indicating the presence of abundant starch. A major difference was observed in starch distribution of the mutant relative to WT (Figure A 3 B), with pro portionately less starch in the basal area of the epuf1 endosperm (Figure A 3 B). Embryo formation was also affected in the epuf1 mutant, however the severity of defective embryo development was variable. Kernels were stained with acetocarmine for contrast in cell type since the small cells of the embryo are more resistant to uptake of stain than the endosperm and surrounding tissues. Results showed a clearly defined embryo in WT kernels, whereas mutant kernels had severely reduced embryos or completely lac ked visibly detectable embryo s (Figure A 3 C). The causal insertion for this mutant has yet to be identified through genotyping for candidate genes and co segregational analysis. Identification of Mu Flanking Sequence Tags Sequences flanking Mu insertion were identified by the UniformMu group as described by Settles et al. (Settles et al., 2004; 2007). of the shredded and empty pericarp uf 1 mutants Acetocarmine solution (1%) was prepared by dissolvin g 10 g carmine (Fisher CAT# C579 25) in 100 mL glacial acetic acid. Boileezers (Fischer Scientific, Fair Lawn, NJ) were added, and the solution was refluxed for 24 h. Tissues were stained by submerging in 1% acetocarmine solution for 2 to 3 min, then rinsi dissolving 10 g potassium iodide in 100 mL of water, then adding 5 g iodine crystals and mixing with water. Images were captured with an RT SPOT camera (Diagnostic Instruments) mounted on a Leica MZ 12 5 dissection microscope or an Olympus BH2 light microscope.
107 Figure A 1 The maize shredded mutant 05 S 2500 under field conditions. Visible phenotypic features of the shredded mutant include longitudinal strips of tissue that turn between vascular bundles (left), and small plant stature with lighter green leaves than the wild type (right). Wild type plants are visible in the adjacent row. Photos by
108 Figu re A 2 The shredded phenotype. A) The three stages of the shredded leaf phenotype viewed under a dissection microscope. Longitudinal strips between major vascular bundles first became chlorotic, eventuall y turning white and translucent ( white arrow). Affected tissue then ( black arrow) before tissue finally separat ed ( grey arrow). B) Cyano acrylate impression of the boundary between affected and unaffected leaf tissue. Cell files and stomatal distribution appeared normal (right side of B ), however tissue to the left of the vascular bundle showed aberrant cell shape, size and arrangement with disorganized cell files and under developed stomata. C) A portion of leaf stained with acetocarmine for contrast. Affected (white) and unaffected (red) areas were clearly defined, with absence and/or deterioration of vascular bundles in chlorotic zones Photos by
109 Figure A 3 Characteristics of the empty pericarp UF ( epuf1 ) mutant line 09S 3300. A) Longitudinal sections of WT and epuf1 kernels at 20 day s after pol lination. The mutant phenotype wa s characterized by small kernels, and underdeveloped endosperm surro unded by expanded pericarp. B) Wild type and epuf1 kernels stained d starch in the endosperm, however the (starchless) basal endosperm in the mutant comprised a greater portion of the whole relative to WT (arrows) C) Mutant and WT kernels stained with acetocarmine for contrast. Embryo tissue can be seen in the WT as a prominent area of unstained tissue (arrow), however epuf1 kernels had either an underdeveloped or aborted embryo as shown (a rrow), or no visible embryo Photos by
110 APPENDIX B SEQUENCES OF GENE SPECIFIC PRIMERS USE D FOR QRTPCR ANALYSI S Table B 1. Sequence of primers used for qRTPCR analysis. Primer Sequence RTCESA1 F GCCTCGTCGGTGTTGGTGT RTCESA1 R CTC CCA CCG AGA CGG CT RTCESA2 F GCCTCGTCGGTGTCGGT RTCESA2 R CAT CTC CGC GCT CCT CCT A RTCESA3 F TAGTGCCTGTTTCATGTTGACTGTCGT RTCESA3 R GTT ACA TCA CGA CAG TCC AGA GCC RTCESA4 F ATACCCAGACGTGTGGCATCAACT RTCESA4 R CACG TGG TAT TCC TCC GTT TCT ACA AAG RTCESA5 F AGCGAGCTCCACCACTTGC RTCESA5 R AGG CAG AAG CAG AGG CAG C RTCESA6 F GATCTTCTCGCTGCTTTGGGTCC RTCESA6 R CAG GGG ACC ATC ATC CTT CGC RTCESA7 F CGG TCT GTG GCC GAC TC RTCESA7 R CCA CGG CCG TAG CTC ATG T RTCESA7 Paralog F ATG TGC ATC TGC CAG TGG AAC AGA RTCESA7 Paralog R TTC CTA GTA TAG ACG AAC ATG TAA TGA AGT TTG T RTCESA8 F GCTGTAGATAGAAACCACATGTCCACGG RTCESA8 R GGC ACC TCT CTC CTG CTC C RTCESA9 F GAAACAGAGAGATACCACGAATGTGCCG RTCESA9 R ACG CCA CCT GCC TAT ATA ACT TAC TAA CAG RTCESA10 F CGTTTGGACATACAGGCACTTTTGGG RTCESA10 R GAG TGA ATT CCA TCA CAG TTC TTA CAC CC RTCESA11 F ATCTCGCATCTGGGCTTTTGCC RTCESA11 R CCG AAT TTT AAC ATT TCA GGT TTC ACC ACC A RTCESA12 F TCAGGCAGTGTGGCATCAATTGC RTCESA12 R CCG ACA ATT CTG GGT ACC ATA ACA TTA CAG AC RT MPI F TAG CCG CTA TTT CCT TTC CTT GCC RT MPI R TGA GAA TTC ACA CAT CCA TTA TTC GGC ATG C
111 LIST OF REFERENCES Abedon, B.G., Hatfield, R.D., and Tracy, W.F (2006). Cell wall composition in juvenile and adult leaves of maize ( Zea mays L.) J. Agric Food Chem 54 : 3896 3900 Albrecht, G., and Mustroph, A. (2003). Localization of sucrose synthase in wheat roots: increased in situ activity of sucrose synthase correlates with cell wall thickening by cellulose deposition under hypoxia. Planta 217: 252 260. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., and Lipma n, D.J. (1990). Basic local alignment search tool. J. Mol. Biol. 215: 403 10. Amano, Y., and Kanda, T. (2002). New insights into cellulos e degradation by cellulases and related enzymes. Trends Glycosci. Glycotechnol. 14 : 27 34. Amor, Y., Haigler, C.H., Johnson, S., Wainscott, M., and Delmer, D.P. (1995).A membrane associated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants. Proc. Natl Acad. Sci. USA 92: 9353 9357. Andrew Carroll, A., and Specht, C.D. (2011). Understanding plant cellulose synthases through a comprehensive investigation of the cellulose synthase family sequences. Front. Plant Sci. 2: doi:10.3389/fpls.2011.00005 Appenzeller, L., Doblin, M., Barreiro, R., Wang, H.Y., N iu, X.M., Kollipara, K., Carrigan, L., Tomes, D., Chapman, M., and Dhugga, K.S., (2004). Cellulose synthesis in maize: isolation and expression analysis of the cellulose synthase ( CesA ) gene family. Cellulose 11: 287 299. Arioli, T. et al (1998). Molecular analysis of cellulose biosynthesis in Arabidopsis Science 279 : 717 720. Atalla, R. H. and VanderHart, D. L. (1989). Studies on the structure of cellulose using Raman spectroscopy and solid state 13C NMR. Cellulose Wood Chem. Tech. 169 188. A ziz, A., Gauthier, A., Bezier, A., Poinssot, B., Joubert, J.M., Pugin, A., Heyraud, A., and Baillieul, F. (2007). Elicitor and resistance inducing activities of beta 1,4 cellodextrins in grapevine, comparison with beta 1,3 glucans and alpha 1,4 oligogalact uronides. J. Exp. Bot. 58: 1463 1472. Bacic, A. (2006). Breaking an impasse in pectin biosynthesis. Proc. Natl. Acad. Sci. 103: 5639 5640. Bacic, A., and Stone, B.A. (1981). Chemistry and organization of aleurone cell wall components from wheat and barley. Aust. J. Plant Physiol. 8: 475 495.
112 Beeckman, T., Przemeck, G.K.H., Stamatiou, G., Lau, R., Terryn, N., De Rycke. R., Inze, D., and Berleth, T. (2002). Genetic complexit y of cellulose synthase a gene function in Arabidopsis embryogenesis. Plant Physiol. 130 : 1883 1893. Bennetzen, J.L. (2000). Transposable element contributions to plant genome evolution. Plant Mol. Biol. 42: 251 269. Bessueille, L., Sindt, N., Guichardant, M., Djerbi, S., Teeri, T.T., and Bulone, V. (2009). Plasma membrane microdomains from hybrid aspen cells are involved in cell wall polysaccharide biosynthesis. Biochem. J. 420: 93 103. Betancur, L., Singh, B., Rapp, R.A., Wendel, J.F., Marks, M.D., Roberts, A.W., and Haigler, C.H. (2010). Phylogenetically distinct cellulose synthase genes support secondary wall thickening in Arabidopsis shoot trichomes and cotton fiber. J. Integrative Plant Biol. 52: 205 220. Bewley, J.D. (1997). Breaking down the walls: a role for endo mannanase in release from seed dormancy. Trends Plant Sci. 2: 464 469. Birchler, J.A., and Veitia, R.A. (2010). The Gene Balance Hypothesis: implications for gene regulation, quantitative traits and evolution. New Phytol. 186: 54 62. Blaschek, W., Haass, D., Koehler, H., and Franz, G. (1981). Cell wall regeneration by Nicotiana tabacum protoplasts: chemical and b iochemical aspects. Plant Sci. Lett. 22: 47 57. Boerjan, W., Ralph, J., and Baucher, M. (2003). Lignin biosynthesis. Ann. Rev. Plant. Biol. 54: 519 546. Bouton, S., Leboeuf, E., Mouille, G., Leydecker, M. T., Talbotec, J., Granier, F., Lahaye, M., Hfte H., and Truong, H. N. (2002). QUASIMODO1 encodes a putative membrane bound glycosyltransferase required for normal pectin synthesis and cell adhesion in Arabidopsis Plant Cell 14: 2577 2590. Bradley, D.J., Kjellbom, P., and Lamb, C.J. (1992). Elicitor and wound induced oxidative cross linking of a proline rich plant cell wall protein: A novel, rapid defense response. Cell 70: 21 30. Brett, CT. ( 2000). Cellulose microfibrils in plants: biosynthesis, deposition and integration into the cell wa ll. Int. Rev. Cyt. 199 : 161 199. Briggs, C. L. (1996). An ultrastructural study of the embryo/endosperm interface in the developing seeds of Solanum nigrum L. zygote to mid torpedo stage. Ann. Bot 78 295 304. Brown, R. M., and Saxena, I. M. (2000). Cellulose biosynthesis: A model for understanding the assembly of biopolymers. Plant Physiol. Biochem. 38 : 57 67.
113 Brown, R.M. Jr. (2004). Cellulose structure and biosynthesis: what is in store for the 21st century? J. Polymer Sci. Part A: Polymer Chem. 42 : 487 495. Bruneau, A.H., and Qu, R. (2010). Tissue culture induced morphological somaclonal variation in St. Augustinegrass [ Stenotaphrum secundatum (Walt.) Kuntze]. Plant Breeding 129: 96 99. Brutnell, T.P. (2002). Transposon tagging in maize. Funct. Integr. Genom. 2: 4 12. Burn, J.E., Hocart, C.H., Birch, R.J., Cork, A.C., and Williamson RE. (2002). Functional analysis of the cellulose synthase genes CesA1 CesA2 and CesA3 in Arabidopsis Plant Physiol. 129 797 807. Burton, R. A., Shirley, N. J., King, B. J., Harvey, A. J., and Fincher, G. B. (2004). The CesA gene family of barley. Quantitative analysis of transcripts reveals two groups of coexpressed genes. Plant Physiol. 134: 224 236. Burton, R.A., Farrokhi, N., Bacic, A., and Fincher, G.B. (20 05). Plant cell wall polysaccharide biosynthesis: real progress in the identification of participating genes. Planta 221: 309 312. Burton, R.A., Wilson, S.M., Hrmova, M., Harvey, A.J., and Shirley, N.J., Medhurst, A., Stone, B.A., Newbigin, E.J., Bacic, A., and Fincher, G.B. (2006). Cellulose Synthase Like CslF genes mediate the synthesis of cell wall (1,3;1,4) D Glucans. Science 311: 1940 1942. Burton, R.A., Ma, G., Baumann, U., Harvey, A.J., and Shirley, N.J. et al. (2010). A customized gene expression microarray reveals that the brittle stem phenotype fs2 of barley Is attributable to a retroelement in the HvCesA4 cellulose synthase gene. Plant Phys. 153: 1716 1728. Campbell, J.A., Davies, G.J., Bulone, V., and Henrissat, B. (1997). A classification of nucleotide diphospho sugar glycosyltransferases based on amino acid sequence similarities. Biochem J. 326: 929 939. Cano Delgado, A., Penfield, S., Smith, C., Catley, and M., Bevan, M. (2003). Reduced cellulose synthesis invokes lignification and defense responses in Arabidopsis thaliana Plant J. 34 : 351 362. Cantatore, J.L. Murphy, S.M. and Lynch, D.V. Compartmentation and topology of glucosylceramide synthesis. Biochem. Soc. Trans 28 : 748 750. Carpita, N.C. (1984). Fractionation of hemicelluloses from maize cell walls with increasing concentrations of alkali. Phytochem. 23: 1089 1093. Carpita, N.C. (1986). Incorporation of pro line and aromatic amino acids into cell walls of maize coleoptiles. Plant Physiol 80: 660 666.
114 Carpita, N.C. (1996). Structure and biogenesis of the cell walls of grasses. Ann. Rev. Plant Physiol. Plant Molec. Biol. 47: 445 476. Carpita, N.C., and Gibeaut, D.M. (1993). Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant J. 3 : 1 30. Cavez, D., Hachez, C., and Chaumont, F. (2009). Maize black Mexican sweet suspensio n cultured cells are a convenient tool for studying aquaporin activity and regulation. Plant Signal Behav. 4: 890 892. Chen, K., and Rajewsky, N. (2007). The evolution of gene regulation by transcription factors and microRNAs. Nat. Rev. Genet. 8: 93 103. Chen, L.M., Carpita, N.C., Reiter, W.D., Wilson, R.W., Jeffries, C., and McCann, M.C. (1998). A rapid method to screen for cell wall mutants using discriminant analysis of Fourier transform infrared spectra. Plant J. 8 : 375 382. Chen, S.L., Ehrhardt, D. W., and Somerville, C.R. (2010). Mutations of cellulose synthase (CESA1) phosphorylation sites modulate anisotropic cell expansion and bidirectional mobility of cellulose synthase. Proc. Natl. Acad. Sci. USA 107: 17188 17193. Chomet, P., Lisch, D., Hardem an, D.J., Chandler, V.L., and Freeling, M. (1991). Identification of a regulatory transposon that controls the Mutator transposable element system in maize. Genetics 129 : 261 270. Chourey, P.S., Chen, Y.C., and Miller, M.E. (1991). Early cell degeneration in developing endosperm is unique to the shrunken mutation in maize. Maydica 36: 141 146. Chu, C.C., et al. (1975). Establishment of an efficient medium for anther culture of rice through comparative experiments on the nitrogen so urces. Sci. Sinica 18: 659 668. Cocuron, J.C., Lerouxel, O., Drakakaki, G., Alonso, A.P., Leipman, A.H., Keegstra, K., Raikhek, N., and Wilkerson, C.G. (2007). A gene from the cellulose synthase like C family encodes a 1,4 glucan synthase. Proc. Natl. A cad. Sci. 104: 8550 8555. Cordero, M.J., Ravents, D., and San Segundo, B. (1994). Expression of a maize proteinase inhibitor gene is induced in response to wounding and fungal infection: systemic wound response of a monocot gene. Plant J. 6: 141 150. Cosgrove, D.J. (1999). Enzymes and other agents that enhance cell wall extensibility. Ann. Rev. Plant Physiol. Plant Mol. Biol 50 : 391 417. Cosgrove, D.J. (2003). Expansion of the plant cell wall The Plant Cell Wall (JKC Rose ed). 237 263. Delmer, D.P. and Amor, Y. (1995). Cellulose biosynthesis. Plant Cell 7: 987 1000.
115 Delmer, D. P. ( 1999). Cellulose biosynthesis: exciting times for a difficult field of study. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50: 245 276. Desprez, T., Juraniec, M., Crowel ,l E.F., Jouy, H., Pochylova, Z., Parcy. F., Hofte, H., Gonneau, M., and Vernhettes, S. (2007). Organization of cellulose synthase complexes involved in primary cell wall synthesis in Arabidopsis thaliana Proc. Natl. Acad. Sci. USA 104 : 15572 15577. Dhu gga, K. S. (2001). Building the wall: genes and enzyme complexes for polysaccharide synthases. Curr. Opin. Plant Biol 4 : 488 493. Dhugga, K. S. (2005). Plant G olgi cell wall synthesis: From genes to enzyme activities. Proc. Natl. Acad. Sci. USA 102: 1815 1816. Diao, X.M., and Lisch, D. (2006). Mutator transposon in maize and MULEs in the plant genome. Acta Genetica Sinica 33: 477 487. Dietrich, C.R., Cui, F., Packila, M.L., Li, J., Ashlock, D.A., Nikolau, B.J., and Schnable, P.S. (2002). Maize Mu transposons are targeted to the 5` untranslated region of the gl8 gene and sequences flanking Mu target site duplications exhibit nonrandom nucleotide composition throughout the genome. Genetics 160: 697 716. Ding, S.Y., and Himmel, M.E. (2006). The maiz e primary cell wall microfibril: A new model derived from direct visualization. J. Agric. Food Chem. 54 : 597 606. Doblin, M.S., Durek, I., Jacob Wild, D., and Delmer, D. P. (2002). Cellulose biosynthesis in plants: from genes to rosettes. Plant Cell Physi ol 43 : 1407 1420. Dubois, M., Gilles, D.A., Hamilton, J.K., Rebers, P.A., and Smith, F. (1956). Colorimetric method for the determination of sugars and related substances. Anal. Chem. 28 : 350 356. Edwards, G.E., Lilley, R.M., Craig, S., and Hatch, M.D. (1979). Isolation of intact and functional chloroplasts from mesophyll and bundle sheath protoplasts of the C4 plant Panic um miliaceum Plant Physiol. 63: 821 827. Elbein, A.D. Forsee, W.T., Shultz, J.C., and Laine, R.A. (1975). Biosynthesis and str ucture of glycosyl diglycerides, steryl glucosides, and acylated steryl glucosides. Lipids 10 : 427 436. Endo, S., Pesquet, E., Yamaguchi, M., Tashiro, G., Sato, M., Toyooka, M., Nishikubo, N., Udagawa Motose, M., Kubo, M., Fukuda, H., and Demura, T. (2009). Identifying new components participating in the secondary cell wall formation of vessel elements in Zinnia and Arabidopsis. Plant Cell 21 : 1155 1165. Engels, F. M., and Jung, H. G. ( 1998). Alfalfa stem tissues: Cell wall development and lignification. Ann. Bot 82 : 561 568.
116 Faik, A., Price, N.F., Raikhel, N.V., and Keegstra, K. (2002). Arabidopsis gene encoding an xylosyltransferase involved in xyloglucan biosynthesis. Proc. Natl. Acad. Sci. 99: 7797 7802. Falconer, M.M., and Seagull, R.W (1985). Immunofluorescent and calcofluor white staining of developing tracheary elements in Zinnia elegans L. suspension cultures. Protoplasma 125: 190 198. Farrokhi, N., Burton, R.A., Brownfield, L., Hrmova, M., and Wilson, S.M., Bacic, A, and Fincher, G.B. (2006). Plant cell wall biosynthesis: genetic, biochemical and functional genomics approaches to the identification of key genes. Plant Biotech. J. 4: 145 167. Fincher, G.B. (2009). Revolutionary times in our understanding of cell wall biosynthesis and remodeling in the grasses. Plant P hysiol. 149: 27 37. Fleischer, A., Titel, C., and Ehwald, R. (1998). The boron requirement and cell wall properties of growing and stationary suspension cultured Chenopodium album L cells. Plant Physiol. 117: 1401 1410. Fowler, J.E., and Quatrano, R.S. (1997). Plant cell morphogenesis: Plasma membrane interactions with the cytoskeleton and cell wall. Annu. Rev. Cell Dev. Biol. 13 : 697 743. Freshour, G., Clay, R.P., Fuller, M.S., Albershein, P., Darvill, A.G., and Hahn, M.C. (1996). Developmental and tissue specific structural alterations of the cell wall polysaccharides of Arabidopsis thaliana roots Plant Physiol. 110: 1413 1429. Frost, D.J., Read, S.M., Drake, R.R., Haley, B.E., and Wasserman, B.P. (1990). Identification of the UDP glucose binding polypeptide of callose synthase from Beta vulgaris L. by photoaffinity labeling with 5 azido UDP glucose. J. Biol. Chem. 265: 2162 2167. Fry S.C. (2004). Primary cell wall metabolism: tracking the careers of wall polymers in living plant cells. New physiol. 161: 641 675. Fuj i i, S., Hayashi, T., and Mizuno, K. (2010). Sucrose synthase is an integral component of the cellulose synthesis machinery. Plant Cell Physiol 51: 294 301. Fukuda, H. (1997). Tracheary element differentiation. Plant Cell 9: 1147 1156. Galletti, R., De Lorenzo, G., and Ferrari, S (2009). Host derived signals activate plant innate immunity. Plant Signal Behav. 4: 33 34. Giddings, T.H., Brower, D.L ., and Staehelin, L.A. (1980). Visualization of particle complexes in the plasma membrane of Micrasterias denticulata associated with the formation of cellulose fibrils in primary and secondary cell walls. J. Cell Biol. 84 : 327 339.
117 Gomez, J., Sanchez Martnez, D., Stiefel, V., Rigau, J., Puigdomenech, P., and Page s, M. (1988). A gene induced by the plant hormone abscisic acid in response to water stress encodes a glycine rich protein. Nature 334: 262 264. Gmez, L.D., Baud, S., Gild ay, A., Li, Y., and Graham, I.A. (2006). Delayed embryo development in the Arabidopsis Trehalose 6 Phosphate Synthase 1 mutant is associated with altered cell wall structure, decreased cell division and starch accumulation. Plant J. 46: 69 84. Gorshkova, T.A., Salnikov, V.V., Pogodina, N.M., Chemikosova, S.B., Yablokova, E.V., Ulanov, A.V., Ageeva, M.V., Van Darn, J.E.G., and Lozovaya, V.V. (2000). Composition and distribution of cell wall phenolic compounds in flax ( Linum usitatissimum L ) st em tissues. Annal. Bot. 85: 477 486. Gould, J.H., Palmer, R.L., and Dugger, W.M. (1986). Isolation and culture of cotton ovule epidermal protoplasts (prefiber cells) and analysis of the regenerated wail. Plant Cell Tiss. 6: 47 59. Grabber, J. H. (2005). How do lignin composition, structure, and cross linking affect degradability? A review of cell wall model studies. Crop Sci. 45: 820 831. Gu, Y ., Kaplinsky, N Bringmann, M Cobb, A ., Carroll, A ., Sampathkumar, A Baskin, T.I Persson, S and Somerville, C.R (2010). Identification of a cellulose synthase associated protein required for cellulose biosynthesis. Proc. Natl. Acad. Sc i. USA 29: 12866 12871. Gutierrez, R., Lindeboom, J.J., Paredez, A.R, Mie, A.M.C., Ehrhardt, D.W. (2009). Arabidopsis cortical microtubules position cellulose synthase delivery to the plasma membrane and interact with cellulose synthase trafficking compartments. Nat. Cell Biol. 11: 797 806. Ha, M.A., Apperley, D.C., Evans, B.W., Huxham, M., Jardine, W.G., Vietor, R.J., Reis, D., Vian, B., and Jarvis, M.C. (1998). Fine structure in cellulose microfibrils: NMR evidence from onion and quince. Plant J. 16: 183 190. Hall, Q. and Cannon, M.C. (2002). The cell wall hydroxyproline rich glycoprotein RSH is essential for normal embryo development in arabidopsis. Plant Cell 14: 1161 1172. Hansen, K.M ., Truesen, A.B ., and Soderberg, J.R (2001). Enzyme assay for identification of pectin and pectin derivatives, based on recombinant pectate lyase. J.A.O.A.C. Int. 84: 185 1 1854. Harris, P.J., and Hartley, R.D. (1980). Phenolic constituents of the cell walls of monocotyledons. Biochem. System. Ecol. 8: 153 160. Hatfield R., and Vermerris W. (2001). Lignin formation in plants. The dilemma of linkage specificity.Plant Physio l. 126: 1351 1357.
118 Hernandez Blanco, C., Feng, D.X., Hu, J. et al. (2007). Impairment of cellulose synthases required for Arabidopsis secondary cell wall formation enhances disease resistance. Plant Cell 19: 890 903. Herth, W. (1983) Arrays of plasma formation in spirogyra. Planta 159 : 347 356. Herth, W. (1985). Plasma membrane rosettes involved in localized wall thickening during xylem vessel formation of Lepidium sativum L. Planta 164: 12 21. Hirose, E., Kimura, S., Itoh, T., and Nishikawa, J. (1999). Tunic of pyrosomas, doliolids and salps (Thaliacea, Urochordata): morphology and cellulosic components. Biol. Bull. 196: 113 120. Holland, N., Holland, D., Helentjaris, T., Dhugga, K. S., Xoconostle Cazares, B., and Delmer, D. P. (2000). A comparative analysis of the plant cellulose synthase ( CesA ) gene family. Plant Physiol. 123 : 1313 1323. Horine, R K., and Ruesnik, A.W. (1972). Cell wall regeneration around protoplasts isolated from Convolvulus tissue culture. Plant Physiol. 50 : 438 445. Huang, R.F., and Lloyd, C.W. (1999). Gibberellic acid stabilises microtubules in maize tubulin1. FEBS Lett. 443: 317 320. Hughs, R., and Street, H.E. (1974). Galactose as an inhibitor of the expansion of root cells. Annals Bot. 38 : 555 564. Iwai, H., Masaoka, N., Ishii, T., and Satoh, S. (2002). A pectin glucuronyltransferase gene is essential for intercellular attachment in the plant meri stem Proc. Natl. Acad. Sci. USA 99 : 16319 16324. Jackson, P., Galinha, C., Pereira, C., Fortunato, A., Soares, N., Amncio, S. and Ricardo, C.P. (2001). Rapid deposition of extensin during the elicitation of grapevine callus cultures is specifically catal ised by a 40kDa peroxidase. Plant Physiol. 127: 1065 1076. Jang, J. C., and Sheen, J. (1994). Sugar sensing in higher plants. Plant Cell 6 1665 1679. Joshi, C.P., Thammannagowda, S., Fujino, T., Gou, J., and Avci, U. et al. (2011). Perturbation of Wood Cellulose Synthesis Causes Pleiotropic Effects in Transgenic Aspen. Mol. Plant 4: 331 345. Juge, N. (2006). Plant protein inhibitors of cell wall degrading enzymes. Trends Plant Sci. 11 : 359 367. Jung, H.G., and Casler, M. D. (2006). Maize stem tissues: Cell wall concentration and composition during development. Crop Sci. 46: 1793 1800.
119 Kapitonov, V.V., and Jurka, J. (2001). Rolling circle transposons in eukaryotes. Proc. Natl. Acad. Sci. USA 98: 8714 8719. Kawagoe, Y., Delmer, D.P. (1997). Cotton CelA1 has a LIM like Zn binding domain in the N terminal cytoplasmic region. Plant Physiol 114: S 85. Kawano Y Saotome T Ochiai Y Katayama M Narikawa R Ikeuchi M (20 11). Cellulose accumulation and a cellulose synthase gene are responsible for cell aggregation in the cyanobacterium Thermosynechococcus vulcanus RKN. Plant Cell Physiol. 52: 957 966. Keegstra, K., and Walton, J. Glucans -brewer's bane, dieticia n's delight. Science 311 : 1872 1873. Koch, K.E. (2004). Sucrose metabolism: regulatory mechanisms and pivotal roles in sugar sensing and plant development. Curr. Opin. Plant Biol 7: 235 246. Lampe L. (1931). A microchemical and morphological study of the developing endosperm in maize. Bot. Gaz. 91: 337 376. Lee, M., and Phillips, R.L. (1988). The chromosomal basis of somaclonal variation. Annu. Rev. Plant Physiol. Plant Mol. Biol. 39: 413 437. Lerouxel, O., Cavalier, D.M., Liepman, A.H., and Keegstr a, K. (2006). Biosynthesis of plant cell wall polysaccharides a complex process. Cur. Opin. Plant Biol. 9 : 621 630. Liepman, A. H., Wilkerson, C. G., and Keegstra, K. (2005). Expression of cellulose synthase like ( Csl ) genes in insect cells reveals that CslA family members encode mannan synthases. Proc. Natl. Acad. Sci. USA 102 : 2221 2226. Ling, A.P.K., Phua, G.A.T., Tee, C.S., and Hussein, S. (2010). Optimization of protoplast isolation protocols from callus of Eurycoma longifolia J. Med. Plants Res. 4: 1778 1785. Lingwood, D., and Simons, K. (2007). Detergent resistance as a tool in membrane research. Nat. Protocols 2: 2159 2165. Lisch, D. (2002). Mutator transposons. Trends Plant Sci. 7: 498 504. Liu, S. et al. (2009). Mu transposon insertion sites and meiotic recombination events co localize with epigenetic marks for open chromatin across the maize genome. PLoS Genet. 5: e1000733. doi:10.1371/journal.pgen.1000733 Lozovaya, V., Gorshkova, T., Yablokova, E., Zabotina,O., Ageeva, M., Rumyantseva, N., Kolesnichenko, E., Waranuwat, A., and Widholm, J. (1996). Callus cell wall phenolics and plant regeneration ability. Plant Physiol. 148 : 711 717. Ludwig, S. R., Somers, D.A., and Peterson, W.L. (1985). High frequency callus formation from maize protoplasm. Theor. Appl. Genet. 71 : 344 350.
120 Margulies M., Egholm1, M., Altman1, W.E., Attiya1, S, and Bader, J.S et al. (2005). Genome sequencing in microfabricated high density picolitre reactors. Nature. 437: 376 80. Mathew, S., and Abraham, T.E. (2004). Ferulic acid: an antioxidant found naturally in plant cell walls and feruloyl esterases involved in its release and their applications. Crit. Rev. Biotechnol 24 : 59 83. Matthysse, A.G., White, S., and Lightfoot, R. (1995). Genes required for cellulose synthesis in Agrobacterium tumefaciens J. Bacteriol. 177: 1069 1075. May, B.P. Liu, H., Vollbrecht, E., Senior, L., Rabinowicz, P.D., Roh, D., Pan, X., Stein, L., Freeling, M., Alexander, D., and Martienssen, R. (2003). Maize targeted mutagenesis: A knockout resource for maize. P roc. Natl. Acad. Sci 100: 11541 11546. McCabe P.F., and Leaver C.J. (2000). Programmed cell death in cell cultures Plant mol. biol. 44: 359 368. McCarty, D.R. Settles, A.M., Suzuki, M., Tan, B.C., and Latshaw, S. et al. (2005). Steady state transposon mutagenesis in inbred maize. Plant J. 44: 52 61. McCarty, D.R., and Meeley, R.B. (2009). Trans poson r esources for forward and reverse genetics in maize. Handbook of Maize. J.L. Bennetzen, and S. Hake eds (New York, NY: Springer) pp. 561 584. McClintock, B. (1947). Chromosome organization and gene expression. Cold Spring Harb. Symp. Quant. Biol. 16: 13 47. Meeley, R.B., Briggs, S.P. (1995). Reverse genetics for maize. Maize Genet Coop Newsl 69: 67 82. Moreno, A.B., Peas, G., Rufat, M., Bravo, J.M., Estop, M., Messeguer, J., and San Segu ndo, B. (2005). Pathogen induced production of the antifungal AFP protein from Aspergillus giganteus confers resistance to the blast fungus Magnaporthe grisea in transgenic rice. Mol. Plant Microbe Interactions 18: 960 972. Murashige T., and Skoog, F ( 1962) A revised medium for rapid growth and bioassays with tobacco cultures. Physiol. Plant 15: 473 497. Nieuwland, J. Feron, R., Huisman, B.A.H., Fasolino, A., Hilbers, C.W., Derksen, J., and Mariani, C. (2005). Lipid transfer proteins enhance cell wa ll extension in tobacco. Plant Cell. 17 : 2009 2019. Nishiyama, Y., Sugiyama, J., Chanzy, H., and Langan, P. (2003). Crystal structure and hydrogen bonding system in cellulose I from synchrotron X ray and neutron fiber diffraction. J. American Chem. Soc. 125: 14300 14306.
121 Nishiyama, Y., Johnson, G..P, French, A.D., Forsyth, V.T., and Langan, P. (2008). Neutron crystallography, molecular dynamics, and quantum mechanics studies of the nature of hydrogen bonding in cellulose I b Biomacromolecules 9: 3133 314 0. Nobles, D.R., Romanovicz, D.K., and Brown, R.M. Jr (2001). Cellulose in cyanobacteria. Origin of vascular plant cellulose synthase? Plant P hysiol. 127: 529 542. Nolte, K.D., Hendrix, D.L., Radin, J.W., and Koch, K.E. (1995). Sucrose synthase localization during initiation of seed development and trichome differentiation in cotton ovules. Plant Physiol. 109: 1285 1293. Oda,Y., Mimura, T., and Hasezawa, S. (2005). Regulation of secondary cell wall development by cortical microtubules during t racheary element differentiation in Arabidopsis cell suspensions. Plant Physiol. 137: 1027 1036. Offler, C.E., McCurdy, D.W., Patric k, J.W., and Talbot M.J. (2003). Transfer cells: Cells specialized for a special purpose. Ann. Rev. Plant Biol. 54: 431 454. Ohashi Ito, K., Oda, Y., and Fukuda, H. (2010). Arabidopsis VASCULAR RELATED NAC DOMAIN6 directly regulates the genes that govern programmed cell death and secondary wall formation during xylem differentiation. Plant Cell 22: 346 1 3473. Oomen, R.J.F.J., Bergervoet, M. J.E.M., Bachem, C.W.B., Visser, R.J.F., and Vincken, J.P. (2003). Exploring the use of cDNA AFLP with leaf protoplasts as a tool to study primary cell wall biosynthesis in potato. Plant Physiol. Biochem. 41: 965 971. Palmer, E., and Freeman, T. (2004). Investigation into the use of C and N terminal GFP fusion proteins for subcellular localization studies using reve rse transfection microarrays. Comp. Funct. Genom. 5: 342 353. Pandey, S., Wang, X Q., Coursol, S.A., and Assmann, S.M. (2002). Preparation and applications of Arabidopsis thaliana guard cell protoplasts. New Phytol 153: 517 526. Park, Y.W., Tominaga, R., Sugiyama, J., Furuta, Y., Tanimoto, E., Samejima, M., Sakai, F., and Hayashi, T. (2003). Enhancement of growth by expression of poplar cellulase in Arabidopsis thaliana Plant J. 33: 1099 1106. Paredez, A.R., Somerville, C.R., an d Ehrhardt, D.W. (2006). Visualization of cellulose synthase demonstrates functional association with microtubules. Science 312: 1491 1495. Pauly, M., Albersheim, P., Darvill, A., and York, W.S. (1999). Molecular domains of the cellulose/xyloglucan netwo rk in the cell walls of higher plants. Plant J. 20: 629 639. Pauly, M., and Keegstra, K. (2009). Cell wall carbohydrates and their modification as a resource for biofuels. Plant J. 54: 559 568.
122 Pear, J.R., Kawagoe, Y., Schreckengost, W.E., Delmer, D.P., and Stalker, D.M. (1996). Higher plants contain homologs of the bacterial celA genes encoding the catalytic subunit of cellulose synthase. Proc. Natl. Acad. Sci. USA. 93 (22) : 12637 12642. Peng, L., Kawagoe, Y., Hogan, P., and Delmer, D. (2002). Sitosterol glucoside as primer for cellulose synthesis in plants. Science 295: 147 150. Perrin, R., Wilkerson, C., and Keegstra, K. (2001). Golgi enzymes that synthesize plant cell wall polysaccharides: finding and evaluating candidates in the genomic era. Plant Molec. Biol. 47 : 115 130. Perrin, R.M., DeRocher, A.E., Bar Peled, M., Zeng, W. and Norambuena, L. (1999). Xyloglucan f ucosyltransferase, an enzyme involved in plant cell wall biosynthesis. Science 18 : 1976 1979. Persson, S., Paredez, A., Carroll, A., Palsdottir, H., Doblin, M., Poindexter, P., Khitrov, N., Auer, M., and Somerville, C.R. (2007). Genetic evidence for three unique components in primary wall cell wall cellulose synthase complexes in Arabidopsis Proc. Natl. Acad. Sci. USA 104: 15566 15571. Philippe, S., Barron, C., Robert, P., Devaux, M.F., Saulnier, L., and Guillon, F. (2006). Characterization using R aman microspectroscopy of arabinoxylans in the walls of different cell types during the development of wheat endosperm. J. Agric. Food Chem. 54 : 5113 5119. Pilet, P.E., Blaschek, W., Senn, A., and Franz, G. (1984). Comparison between maize root cells and their respective regenerating protoplasts: wall polysaccharides. Planta 161: 465 469. Postlethwait, S.N., and Nelson, O.E. (1957). A chronically wilted mutant of maize. Am. J. Bot. 44: 628 633. Potrykus, I. (1991). Gene transfer to plants: Assessment of published approached and results. Annu. Rev. Plant Physiol. Plant Mo l Biol. 42: 205 225. Priefert, H ., Rabenhorst, J ., and Steinbuchel, A (2001). Biotechnological production of vanillin. Appl. Microbiol. Biotechnol. 56 : 296 314. Raizada, M.N., and Walbot, V. (2000). The late developmental pattern of Mu transposon excision is conferred by a cauliflower mosaic virus 35S driven MURA cDNA in transgenic maize. Plant Cell 12: 5 22. Ralph, J., Quideau, S., Grabber, J.H., and Hatfield, R.D. (1994). Identification and synthesis of new ferulic acid dehydrodimers present in grass cell walls. J. Chem. Soc. Perkin Transactions1 23 : 3485 3498. Ramakrishna, P., and Amritphale, D. (2005). The perisperm endosperm envelope in Cucumis : Structure, proton diffusion and cell wall hydrolysing activity Ann. Bot. 96: 769 778.
123 Ramsden, M. J., and Blake, F. S. R. (1997). A kinetic study of the acetylation of cellulose, hemicellulose and lignin components in wood. Wood Sci. Tech. 31: 45 50. R eid, J.S.G., and Edwards, M.E. (1995). Galactomannans and other cell wall storage polysaccharides in seeds. Food polysaccharides and their applications A.M. Stephen, ed.(Boca Raton, FL: CRC Press) pp. 155 186. Richmond, P.A. (1991). Occurrence and functions of native cellulose. Biosynthesis and biodegradation of cellulose. C.H. Haigler, and P.J. Weimer, eds (New York, NY: Marcel Dekker) pp. 5 23. Richmond T .A., and Somerville, C.R. (2000). The cellulose synthase superfamily Plant Physiol. 124: 49 5 498. (2001). Pectins: structure, biosynthesis, and oligogalacturonide related signaling. Phytochem. 57 : 929 967. Roberts, E., and Roberts, A.W. (2009). A cellulose synthase ( CesA ) gene from the red alga Porphyra Yezoensis (Rhodophyta). J. Phycol. 45: 203 212. Robertson, D.S. (1978). Characterization of a mutator system in maize. Mutat. Res. 51: 21 28. Rose, J.K.C., and Bennett, A.B. (1999). Cooperative disassembly of the cellulose xyloglucan network of plant cell walls: parallels between cell expansion and fruit ripening. Trends Plant Sci. 4 : 176 183. Rose, J.K.C., Saladi, M., and Catal, C. (2004). The plot thickens: new perspectives of primary cell wall modification. Curr. Opin. Plant Biol. 7 : 296 301. Ross, P., Mayer, R. and Benziman, M. (1991). Cellulose biosynthesis and function in bacteria Microbiol. Rev. 55: 35 58. Ruan, Y.L., Llewellyn, D.J., and Furbank, R.T (2003). Suppression of sucrose synthase gene expression represses cotton fi ber cell initiation, elongation and seed development. Plant Cell 15: 952 964. Sampedro, J., and Cosgrove, D. J. (2005). The expansin superfamily. Genome Biol. 6: 242.1 242.11. Sato S, et al. (2001). Role of the putative membrane bound endo 1,4 beta glucanase KORRIGAN in cell elongation and cellulose synthesis in Arabidopsis thaliana Plant Cell Physiol. 42: 251 63. Saxena, I.M., Brown, R.M. Jr., Fevre, M., Geremia, R.A., and Henrissat, B. (1995). glycosyltransfer ases: implications for mechanism of action. J Bacteriol. 177: 1419 1424.
124 Saxena, I.M., Brown, R.M. Jr.,and Dandekar, T. (2001). Structure function characterization of cellulose synthase: relationship to other glycosyltransferases. Phytochem. 57: 1135 1148. Saxena, I. M., and Brown, R. M (2005). Cellulose biosynthesis: Current views and evolving concepts. Ann. Bot. 96: 9 21. Schaffner, A.R., and Sheen, J. (1991). Maize rbcS promoter activity depends on sequence elements not found in dicot rbcS promoters. Plant Cell 3: 997 1012. Scheible, W.R., and Pauly M. (2004). Glycosyltransferases and cell wall biosynthesis: novel players and insights. Curr. Opin. Plant Biol. 7 : 285 295. Schnable, J.C., Springer, N.M., and Freeling, M. (2011). Differentiation of the maize subgenomes by genome dominance and both ancient and ongoing gene loss. Proc. Natl. Acad. Sci. doi/10.1073/pnas.1101368108 Schnable P.S., Ware, D., Fulton, R.S., Stein, J.C., and Wei, F. et al. (2 009). The B73 maize genome: complexity, diversity, and dynamics. Science 326: 1112 1115. Schrick, K., Fujioka, S., Takatsuto, S., Stierhof, Y.D., and Stransky, H., Yoshida, S., and Jurgens, G. (2004). A link between sterol biosynthesis, the cell wall and cellulose in Arabidopsis. Plant J. 38: 227 243. Seifert, G.J. (2004). Nucleotide sugar interconversions and cell wall biosynthesis: how to bring the inside to the outside. Curr. Opin. Plant Biol. 7 : 277 284. Seifert, G.J., and Blaukopf, C. (2010). Irritable walls: The plant extracellular matrix and signaling. Plant Physiol. 153: 467 478. Settles, A.M., Latshaw, S., and McCarty, D.R. (2004). Molecular analysis of high copy insertion sites in maize. Nucl. Acids Res. 32: e54. doi: 10.1093/nar/gnh052 Settles, A.M. et al. (2007). Sequence indexed mutations in maize using the UniformMu transposon tagging population. BMC Gen omics. 8: 116. Shatil Cohen, A., and Menachem Z.A. (2011). Bundle sheath cell regulation of xylem mesophyll water transport via aquaporins under drought stress: a target of xylem borne ABA? Plant J. 72 80. Shea, E.M., Gibeaut, D.M., and Carpita, N.C. (1989). Structural analysis of the cell walls regenerated by carrot protoplasts. Planta 179: 293 308. Singh, B., Cheek, H.D., and Haigler, C.H. (2009). A synthetic auxin (NAA) suppresses secondary wall cellulose synthesis and enhances elongation in cultur ed cotton fiber. Plant Cell Rep. 28: 1023 1032.
125 Skoog, F., and Miller, C.O. (1957). Chemical regulation of growth and organ formation in plant tissues cultured in vitro. Symp. Soc. Exp. Biol. 54: 118 130. Somerville,C., Bauer, S., Brininstool, G., Facett e, M., and Hamann,, T., Milne, J., Osborne, E., Paredez, A., Persson, S., Raab, T., Vorwerk, S., and Youngs, H. ( 2004). Toward a systems approach to understanding plant cell walls. Science 306: 2206 2211. Stafford, A., and Warren, G. (1991). Plant Cell and Tissue Culture. (Buckingham, UK: Open University Press) pp. 48 81. Sterling, J. D., Atmodjo, M. A., Inwood, S. E., Kumar Kolli, V. S., Quigley, H. F., Hahn, M. G., and Mohnen, D. (2006) From the cover: Functional identification of an Arabidopsis pect in biosynthetic homogalacturonan galacturonosyltransferase. Proc. Natl. Acad. Sci. USA 103 : 5236 5241. Stone, E.A., and Ayroles, J.F. (2009) Modulated Modularity Clustering as an exploratory tool for functional genomic inference. PLoS Genet 5(5): e1000479. doi:10.1371/journal.pgen.1000479 Subbaiah, C.C., Bush, S.D., and Sachs, M.M. (1998). Mitochondrial contribution to the anoxic Ca2 + signal in maize suspension cultured cells. Plant Physiol. 118: 759 771. Swigonova, Z., Lai, J., Ma, J., Ramakrishna, W.,Llaca, V., Bennetzen, J., and Messing, J. (2004). Close split of sorghum and maize genome progenitors. Genome Res. 14: 1916 1923. Szymanska Chargot, M ., Cybulska, J ., and Zdunek, A (2011). Sensing the structural differences in cellulose from apple and bacterial cell wall materials by Raman and FT IR spectroscopy. S ensors 11: 5543 5560. Tamaru, Y., Ui, S., Murashima, K., Kosugi, A., Chan, H., Doi, R. H., Liu, B. (2002). Formation of protoplasts from cultured tobacco cells and Arabidopsis thaliana by the action of cellulos omes and pectate lyase from C lostridium cellulovorans Appl. Environ. Microbiol. 68: 2614 2618. Tamura, K., Dudley, J., Nei, M., and Kumar, S. (2007). MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24 : 1596 1599. Tanaka, F., and Iwata, T. (2006). Estimation of the elastic modulus of cellulose crystal by molecular mechanics simulation. Cellulose 13: 509 517. Tanaka, K., Murata, K., Yamazaki, M., Onosato, K., and Miyao, A. et al. (2003).Three distinct ri ce cellulose synthase catalytic subunit genes required for cellulose synthesis in the secondary wall. Plant Physiol. 133: 73 83. Tanurdzic, M. et al. (2008). Epigenomic consequences of immortalized plant cell suspension culture. PLoS Biol. 6: e302. doi:10 .1371/journal.pbio.0060302
126 Taylor, N.G., Laurie, S., and Turner, S.R. (2000). Multiple cellulose synthase catalytic subunits are required for cellulose synthesis in Arabidopsis Plant Cell 12: 2529 2539. Taylor, N.G., Howells, R.M., Huttly, A.K., Vickers, K., and Turner, S.R. (2003). Interactions among three distinct CesA proteins essential for cellulose synthesis. Proc. Natl. Acad. Sci. USA 100: 1450 1455. Taylor, N. G. (2008). Cellulose biosynthesis and deposition in higher plants. New Phytol. 17 8 : 239 252. Thompson, J.E., and Fry, S.C. (2001). Restructuring of wall bound xyloglucan by transglycosylation in living plant cells. The Plant J. 26 : 23 34. Thorpe, M.R., Macrae, E.A., Minchin, P.E.H., and Edwards, C.M. (1999). Galactose stimulation of carbon import into roots is confined to the Poaceae. J. Exp. Bot. 50: 1613 1618. Turner, S., Gallois, P., and Brown, D. (2007). Tracheary element differentiation Annu. Rev. Plant Biol. 58: 407 433. Valla, S., Coucheron, D.H., Fjaervlk, E., Kjosbakken, J., and Welnhouse, H. et al. (1989). Cloning of a gene involved in cellulose biosynthesis in Acetobacter xylinum : Complementation of cellulose negative mutants by the UDPG pyrophosphorylase structural gene. Mol. Gen. Genet. 217: 26 30. Varner, J.E., and Lin, L. (1989). Plant cell wall architecture. Cell 56: 231 239. Veit, B., Vollbrecht, E., Mathern, J., and Hake, S. (1990). A tandem duplication causes the Knl O allele of Knotted, a dominant morphological mutant of maize. Genet ics 125: 623 631. Vicient, C.M. (2010). Transcriptional activity of transposable elements in maize. BMC Genom. 11: 601doi:10.1186/1471 2164 11 601 Vietor, R.J., Newman, R.H., Ha, M.A., Apperley, D.C., and Jarvis, M.C. (2002). Conformational features of crystal surface cellulose from higher plants. Plant J. 30: 721 731. Vollbrecht, E. Duvicka, J., Scharesa, J.P., Ahernb, K.R., and Deewatthanawong P. et al. (2010). Genome wide distribution of transposed Dissociation elements in maize. Plant Cell 22: 1667 1685. Walley, J.W., Coughlan, S., Hudson, M.E., Covington, M.F., and Kaspi R, et al. (2007) Mechanical stress induces biotic and abiotic stress responses via a novel cis element. PLoS Genet. 3: e172. doi:10.1371/journal.pge n.0030172 Walbot, V. (2000). Saturation mutagenesis using maize transposons. Curr. Opin. Plant Biol. 3: 103 107.
127 Walbot, V., and Rudenko, G.N. (2002). MuDR/Mu transposable elements of maize. In Mobile DNA II N.L. Craig, R. Craigie, M. Gellert, and A. Lambowitz, eds (Washington, DC: American Society of Microbiology), pp. 533 564. Wang, H., Slater, G.P., Fowke, L.C., Saleem, M., Cutler, A.J. (1991). Comparison of cell wall regeneration on maize pr otoplasts iso lated from leaf tissue and suspension cultured cells In Vitro Cell. Dev. Biol. 27 : 70 77. Wang, J., Elliott, J.E., and Williamson, R.E. (2008). Features of the primary wall CESA complex in wild type and cellulose deficient mutants of Arabidopsis thaliana J. Exp. Bot. 59: 2627 2637. Wang, L., Guo, K., Li, Y., Tu, Y., Hu, H., Wang, B., Cui, X., and Peng, L. ( 2010). Expression profiling and integrative analysis of the CESA / CSL superfamily in rice. BMC Plant Biol. doi/10.1186/1471 2229 10 282 Wang, Q.H. Zhang, X ., Li, F.G ., Hou, Y.X ., Liu, X.L ., and Zhang, X.Y ( 2011). Identification of a UDP glucose pyrophosphorylase from cotton ( Gossypium hirsutum L. ) involved in cellulose biosynthesis in Arabidopsis thaliana. Plant Cell Rep. 30: 1303 1312. Wei, F., Coe, E., Nelson, W., Bharti, A.K., Engler, F. et al. (2007) Physical and genetic structure of the maize genome reflects its complex evolutionary history. PLoS Genet. 7: e123. doi:10.1371/journal.pgen.0030123 Whitney, S.E.C., Gothard, M.G.E., Mitchell, J.T., and Gidley, M.J. (1999). Roles of cellulose and xyloglu can in determining the mechanical properties of primary plant cell walls. Plant Physiol 121 : 657 663. Wicker, T. et al. (2007). A unified classification system for eukaryotic transposable elements. Nature Rev. Genet. 8: 973 982. Wilkie, K.C.B. (1979). The hemicelluloses of grasses and cereals. Adv. Carbohydrate Chem. Biochem 36: 215 64. Williams Carrier, R., Stiffler, N., Belcher, S., Kroeger, T., Stern, D.B., Monde, R.A., Coalter, R., and Barkan, A. (2010). Use of Illumina sequencing to identify tran sposon insertions underlying mutant phenotypes in high copy Mutator lines of maize. Plant J. 63: 167 77. Wu, A., Hu, J.S., and Liu, J.Y (2009). Functional analysis of a cotton cellulose synthase A4 gene promoter in transgenic tobacco plants. Plant Cell Reports 28: 1539 1548. Wu, C.T., Leubner Metzger, G., Meins, F., and Bradford, K.J. 1,3 glucanase and chitinase are expressed in the micropylar endosperm of tomato seeds prior to radicle emergence. Plant Physiol. 126: 1299 13 13. Xin, Z., and Li, P.H (1992). Abscisic acid induced chilling tolerance in maize suspension cultured cells. Plant Physiol. 99: 707 711.
128 Yamamoto, R. (1987). Effect of galactose on auxin induced cell elongation in oat coleoptile segments in mannitol solutions. J. Plant Res. 100: 43 49. Yamazaki, T., Kawamura, Y., Minami, A., and Uemura, M. (2008). Calcium dependent freezing tolerance in Arabidopsis involves membrane resealing via synaptotagmin SYT1. Plant Cell 20: 3389 3404. Ye, H., Song, Y.R., Marcus, A., and Varner, J.E. (1991). Comparative localization of three classes of cell wall proteins. Plant J. 1: 175 183. Yim, K.O. and Bradford, K.J. (1998). Callose deposition is responsible for apoplastic semipermeability of the endosperm envelope of muskmelon seeds. Plant Physiol. 118 : 83 90. Yokoyama R and Nishitani K (2001). A comprehensive expression analysis of all members of a gene family encoding cell wall enzymes allowed us to predict cis regulatory regions involved in ce ll wall construction in specific organs of Arabidopsis Plant Cell Physiol. 42: 1025 33. Yong W. P. et al. (2003). Enhancement of growth by expression of poplar cellulose in Arabidopsis thaliana. Plant J. 33 : 1099 1106. Yoo, S. D., Cho, Y. H., and Sheen, J. (2007). Arabidopsis mesophyll protoplasts: a versatile cell system for transient gene expression analysis. Nature Protocols 2: 1565 1572. Yu, S.M. (1999). Cellular and genetic responses of plants to sugar starvation. Plant Physiol. 121: 687 693. Zeiger, E., Hepler, P.K. (1979). Blue light induced intrinsic vacuolar fluorescence in onion guard cells. J. Cell Sci 37: 1 10. Zhong et al. (2003). Expression of a mutant form of cellulose synthase AtCesA7 causes dominant negative effect on cellulose biosynthesis. Plant Physiol 132 : 786 795.
129 BIOGRAPHICAL SKETCH 1982 in Davenport Iowa. The oldest child in a family of five, in Florida, he had spent 4 years in Jamaica, 8 years in Okinawa, Japan, and 7 years in Jacksonville, North Carolina. Brent began his post hi gh school academic career in 2000 at Embry Riddle Aeronautical University with a focus on aeronautical engineering. His interests soon shifted, however, and he transferred to th e University of Florida in 2002. He graduated cum e in Plant Science, (with a focus in biotechnology) in 2004, and remained at the University of Florida for graduate school. Brent was awarded a n Alumni Fellowship to enroll in the Plant Molecular and Cellular Biology program, and received his PhD in Decemb er, 2011.