Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Elucidation of secondary cell wall secretion mechanisms of Arabidopsis thaliana, Poplar (Populus deltoides… Kaneda, Minako 2008

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Notice for Google Chrome users:
If you are having trouble viewing or searching the PDF with Google Chrome, please download it here instead.

Item Metadata

Download

Media
24-ubc_2009_spring_kaneda_minako.pdf [ 9.49MB ]
Metadata
JSON: 24-1.0066871.json
JSON-LD: 24-1.0066871-ld.json
RDF/XML (Pretty): 24-1.0066871-rdf.xml
RDF/JSON: 24-1.0066871-rdf.json
Turtle: 24-1.0066871-turtle.txt
N-Triples: 24-1.0066871-rdf-ntriples.txt
Original Record: 24-1.0066871-source.json
Full Text
24-1.0066871-fulltext.txt
Citation
24-1.0066871.ris

Full Text

 ELUCIDATION OF SECONDARY CELL WALL SECRETION MECHANISMS OF ARABIDOPSIS THALIANA, POPLAR (POPULUS DELTOIDES X P. TRICHOCARPA) AND PINE (PINUS CONTORTA)  by MINAKO KANEDA M.Sc. Yamagata University 2001  A THESIS SUBMITTED IN PARTIAL FUFILLMENT OF THE REQUIREMENT FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Botany) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December 2008   © Minako Kaneda, 2008  ii Abstract Lignin is a key component of plant secondary cell walls, providing strength to the plant and allowing water transport. Lignin is a polymer of monolignols that are synthesized in the cell and transported into the cellulose rich cell wall. The primary goal of this thesis is to understand the mechanism(s) of monolignol deposition during xylogenesis. The currently accepted theory is that monolignols are exported by Golgi-mediated vesicle delivery to the secondary cell wall.  When this theory was re-examined using cryofixed developing pine, quantitative autoradiography showed that monolignols did not accumulate in Golgi but were rapidly translocated from cytosol to cell wall.  This suggests alternative mechanisms, such as membrane transporters, work in monolignol export. ATP binding cassette (ABC) transporters were chosen because they transport other secondary metabolites and some ABC transporter encoding genes are highly expressed in lignifying cells.  Four candidate ABC transporters were selected in Arabidopsis (ABCB11, ABCB14, ABCB15 from the ABCB/MDR subfamily and ABCG33 from the ABCG/PDR subfamily) and shown to have overlapping, high vasculature expression patterns. Mutants with T- DNA insertions in single ABC transporter genes had no change in lignification of inflorescence stems. However, a reduced polar auxin transport phenotype was detected in mutants of ABCB11, ABCB14 and ABCB15. An additional approach was the use of inhibitors of ABC transporters.  A new assay, which was developed to quantify lignification in primary xylem of Arabidopsis roots, demonstrated that ABC inhibitors did not change lignin deposition.  Monolignols are exported and polymerized in the polysaccharide matrix of the cell wall, which includes hemicelluloses that may organize monolignols during polymerization. Since diverse lignified cell types are enriched in either G- or S-lignin, I hypothesized that this pattern could reflect different hemicellulose distributions, which was examined using antibody labeling  iii of xylans or mannans in hybrid poplar xylem.  While xylans were generally distributed in all secondary cell walls, mannans were enriched in fibers but not in the ray and vessel walls. In summary, during secondary cell wall deposition, monolignols are exported by unknown transporter(s) rather than Golgi vesicles. In developing poplar wood, the monolignols are deposited into diverse hemicellulose domains in different cell types.  iv  TABLE OF CONTENTS Abstract ………………………...………………………………..….…….……… ii Table of contents ………………..............……………….……..…….....…..…… iv List of tables...…….………….......……………………………….……..…..…...viii List of figures….……………….......…………………....…………..…..………...ix Acknowledgements...……..………..………………..………………..…..…....… xi Co-Authorship statement……………………....…………..…..………...……..……......….xii  CHAPTER 1: GENERAL INTRODUCTION……… .........................................1 1.1 Xylogenesis ....................................................................................................2 1.1.1 Primary xylem vs. secondary xylem..........................................................................2 1.1.2 Stages of cell differentiation in the secondary cell wall .............................................3 1.2 Lignin biosynthesis pathway in secondary cell wall formation ................12 1.2.1 Lignin.....................................................................................................................12 1.2.2 The enzymes of phenylpropanoid metabolism in angiosperms and gymnosperms. ..13 1.2.3 Subcellular localization of monolignol synthesis enzymes ......................................16 1.3 Monolignol export models: vesicle fusion or membrane transport..........17 1.3.1 Historic theory and early autoradiography methods.................................................17 1.3.2 Alternative hypotheses: Plasma membrane monolignol transporters........................18 1.3.3 ATP binding cassette (ABC) transporters................................................................19 1.4 Objectives ....................................................................................................21 1.5 Bibliography................................................................................................24 CHAPTER 2: TRACKING MONOLIGNOLS DURING SECONDARY CELL WALL DEVELOPMENT IN PINUS CONTORTA VAR. LATIFOLIA AND POPULUS DELTOIDES X P. TRICHOCARPA 1 ...............................................30 2.1 Introduction ................................................................................................31 2.1.1 Preliminary data from Samuels lab .........................................................................32  v 2.2 Results .........................................................................................................42 2.2.1 3H-phenylalanine is incorporated into the lignin precursors and polymers. ..............42 2.2.2 3H-phenylalanine was intensively incorporated into developing xylem as lignin and its precursors. ....................................................................................................................48 2.2.3 Quantitative decay event analysis showed localization of lignin precursors/lignin in the cytoplasm and in secondary cell wall. ..........................................................................54 2.3 Discussion ....................................................................................................60 2.4 Materials and methods ...............................................................................66 2.4.1 Plants material and growth condition ......................................................................66 2.4.2 Autoradiography.....................................................................................................66 2.4.3 High pressure freezing (HPF) .................................................................................67 2.4.4 Light microscopy....................................................................................................68 2.4.5 Electron microscopy ...............................................................................................69 2.4.6 Detection of radiolabel in tissues and solutions. ......................................................69 2.5 Bibliography................................................................................................72 CHAPTER 3: ABC TRANSPORTERS COORDINATELY EXPRESSED DURING LIGNIFICATION IN ARABIDOPSIS STEMS INCLUDE A SET OF ABCB’S ASSOCIATED WITH AUXIN TRANSPORT 1 ...........................75 3.1 Introduction ................................................................................................76 3.2 Results .........................................................................................................78 3.2.1 Selection of Arabidopsis stem ABC transporter genes correlated with lignification.78 3.2.2 Promoter::GUS fusions indicate that ABCB15, ABCB14, ABCB11 and ABCG33 have high vasculature expression and overlapping expression patterns.......................................86 3.2.3 Reverse genetics of candidate ABC transporter genes in Arabidopsis......................95 3.2.4 Reduced polar auxin transporter in ABC transporter mutants ..................................99 3.3 Discussion ..................................................................................................108 3.4 Materials and methods .............................................................................114 3.4.1 In silico promoter cis-element analysis .................................................................114 3.4.2 Plant materials and growth conditions...................................................................114 3.4.3 Generating promoter::GUS constructs...................................................................115  vi 3.4.4 Reverse transcription PCR and T-DNA.................................................................117 3.4.5 Light microscopy and lignin histochemistry..........................................................117 3.4.6 Polar auxin transport analysis in ABC transporter mutants. ...................................117 3.4.7 DR5::GUS in abcb14............................................................................................118 3.4.8 Phylogenetic relationship of ABCB subfamily genes ............................................118 3.5 Bibliography..............................................................................................119 CHAPTER 4: TESTING THE ROLE OF ABC TRANSPORTERS ON LIGNIN DEPOSITION DURING XYLOGENESIS USING INHIBITORS 1 .....    .................................................................................................122 4.1 Introduction ..............................................................................................123 4.1.1 Plant PM H+-ATPase is not inhibited with ABC transporter inhibitors..................125 4.2 Results .......................................................................................................130 4.3 Discussion ..................................................................................................138 4.4 Materials and methods .............................................................................140 4.4.1 ABC transporter inhibitor treatment in poplar with autoradiography .....................140 4.4.2 ABC transporter inhibitor treatment in Arabidopsis seedling root. ........................140 4.4.3 Barley growth test with inhibitors .........................................................................141 4.5 Bibliography..............................................................................................142 CHAPTER 5: CELL SPECIFIC DEPOSITION OF HEMICELLULOSE IN DEVELOPING SECONDARY CELL WALL OF HYBRID POPLAR (POPULUS DELTOIDES X P. TRICHOCARPA) VESSELS AND FIBERS 1 145 5.1 Introduction ..............................................................................................146 5.2 Results .......................................................................................................148 5.2.1 Secondary xylem and ultrastructure during xylogenesis ........................................148 5.2.2 Xylan and mannan distribution in poplar secondary xylem....................................153 5.3 Discussion ..................................................................................................158 5.4 Materials and methods .............................................................................160 5.4.1 Plant materials and growth conditions...................................................................160 5.4.2 Immunofluorescent localization in xylem tissue by LM10 antibody and anti mannan antibody. .........................................................................................................................160 5.4.3 Immuno-gold labeling for xylem tissue by monoclonal mannan antibody. ............161  vii 5.5 Bibliography..............................................................................................162 CHAPTER 6: CONCLUSION AND FUTURE DIRECTIONS......................164 6.1 Summary of the results and conclusion ...................................................165 6.2 Bibliography..............................................................................................173   viii List of tables   Table 3.1 ABC transporter genes whose expression is correlated with phenylpropanoid biosynthetic genes in Arabidopsis inflorescence stems……….……………………………....79  Table 3.2 Candidate ABC transporter genes and their expression in the Arabidopsis stem epidermis and whole stem, based on microarray data. In addition, in silico predictions of their subcellular targeting are shown……………………….…………………….……….84  Table 3.3 Candidate ABC transporter mutant list ……………….…….………………96   ix  List of figures  Figure 1.1 Xylogenesis –an example of tracheary element development in Zinnia. ............5 Figure 1.2 Xylogenesis –xylem development in poplar wood. ..............................................6 Figure 1.3 Primary and secondary cell wall models .............................................................8 Figure 1.4 Phenylpropanoid pathway in angiosperm and gymnosperm (Humphreys and Chapple, 2002). ................................................................................................................ 14 Figure 2.1 Light autoradiographs of pine stem sections through the cambium  and secondary xylem following incorporation of 3 H-phenylalanine..................................... 35 Figure 2.2 Quantification of grey levels in light microscopy autoradiographs of pine developing secondary xylem............................................................................................ 38 Figure 2.3 TEM autoradiographs of pine tracheids treated with 3H-phenylalanine and either an inhibitor of protein synthesis, cycloheximide (CH), or an inhibitor of phenylpropanoid metabolism, piperonylic acid (PA)..................................................... 40 Figure 2.4 3H-phenylalanine incorporation into lignin demonstrated by radioactive thioacidolysis products. ................................................................................................... 43 Figure 2.5 Analysis of radioactive compounds in methanolic extracts of dissected cambium and associated developing secondary xylem from P. contorta following 4 h incubation in 3 H-phenylalanine....................................................................................... 45 Figure 2.6 Analysis of radioactive compounds in methanolic extracts of dissected cambium and associated developing secondary xylem from hybrid poplar (H11-11) following 4 h incubation in 3 H-phenylalanine. ............................................................... 47 Figure 2.7 Light microscopy autoradiography of poplar developing xylem tissue............ 50 Figure 2.8 Quantification of grey levels in light microscopy autoradiographs of poplar secondary growth tissue. ................................................................................................. 51 Figure 2.9 TEM autoradiographs of poplar xylem tissue treated with 3H-phenylalanine.53 Figure 2.10 TEM autoradiography quantification example (control, 3H-phenylalanine only) in developing tracheids of P. contorta.................................................................... 56 Figure 2.11 Quantification of radioactive label in developing tracheids of P. contorta following cryofixation, autoradiography and TEM. ...................................................... 59 Figure 3.1 Sequence logos visualizing the consensus of boxes P, A and L boxes and their nucleotide matrix............................................................................................................. 81 Figure 3.2 In silico promoter analysis of candidate ABC transporters whose expression is correlated with lignin biosynthetic pathway enzyme genes. .......................................... 82 Figure 3.3 Gene expression profiles of ABCB15 and ABCB11 in young stem segments.... 88 Figure 3.4 ABC transporter expression in mature (within 1 cm of rosette) stem segments. ............................................................................................................................. 90 Figure 3.5  pABCB14::GUS activity detected by fluorogenic substrate. ............................ 91 Figure 3.6 Gene expression profiles of candidate ABC transporter genes, pABCG33::GUS in stem vasculatures. ....................................................................................................... 93 Figure 3.7 GUS expression profiles of candidate ABC transporter gene in seedling root tissue ............................................................................................................................. 94 Figure 3.8 The inflorescence stems of abcb14 knockout mutant showing vasculature development deficient phenotype.................................................................................... 98  x Figure 3.9 Neighbour-Joining (NJ) tree of Arabidopsis ABCB/MDR subfamily subfamily genes. ........................................................................................................................... 100 Figure 3.10 The measurement of polar auxin transport in inflorescence stems of abcb14 mutants using radioactive IAA. .................................................................................... 101 Figure 3.11 The measurement of polar auxin transport in inflorescence stems using DR5::GUS activity......................................................................................................... 103 Figure 3.12 Transcriptional level by RT-PCR in ABC transporter candidate gene mutants. ......................................................................................................................... 106 Figure 3.13 Polar auxin transport of inflorescence stems in ABC transporter mutants. 107 Figure 4.1 Average proton fluxes for potassium (K+) starved barley roots when exogenous potassium was added. .................................................................................................... 126 Figure 4.2 Quantification of grey levels in light microscopy autoradiographs of pine developing secondary xylem.......................................................................................... 128 Figure 4.3 Quantification of grey levels in light microscopy autoradiographs of poplar developing secondary xylem following ABC transporter inhibitor treatment............ 131 Figure 4.4 The procambium length from the root tip to the first lignified TE in Arabidopsis seedling root is increased by auxin transport inhibition. ........................ 134 Figure 4.5  Procambium length after variety of inhibitor treatments in seedling roots .. 135 Figure 4.6 FK506 inhibited sclareol detoxification in barley root.................................... 137 Figure 5.1 Ultrastructure during xylogenesis in poplar.................................................... 150 Figure 5.2 Golgi morphology during xylogenesis in poplar xylem cells. .......................... 152 Figure 5.3 Poplar secondary xylem and immunolabeling of LM10 (xylan) localization . 154 Figure 5.4 Immunolabeling of anti-mannan in poplar secondary xylem ......................... 155 Figure 5.5 Anti-mannan immunogold labeling in poplar developing xylem.................... 157   xi Acknowledgements  It is a pleasure to thank the many people who made this thesis possible. Foremost, I would like to thank my Ph.D. supervisor, Dr. Lacey Samuels, who shared with me a lot of her expertise and research skills. This work would not have been possible without her support and courages.  I would like to thank Dr. Yoshiaki Hara, my supervisor for Master’s degree, at Yamagata University, Japan and Dr. Ken-Ichiro Ishida, a mentor during master program.  Dr. Kim Rensing is a co-author of my manuscript published in Plant Physiology, who was a postdoctoral fellow in Samuels Lab. I’m very grateful to Kim trained me the major experimental skills such as cryofixation, freeze substitution, high pressure freezing and autoradiography.  I’m grateful to my supervisory committee members, Dr. Brian Ellis, Dr. Carl Douglas, Dr. Shawn Mansfield for helpful suggestions on my research and thesis editing.  I would like to thank Garnet Martens, Kevin Hodgson, and Derrick Horne for technical support for TEM and HPF.  I’m very thankful to Dr. Kyu-Young Kang and Russell Chedgy and Shawn Mansfield for helping with HPLC, GC and thioacidolysis work in the department of wood science.  Special thanks to lab members, Robin Young, Allan DeBono, David Bird, Jamie Pighin, Billy Lin, Sarah Ravn, John Wong, Hardy Hall.  Prof. Tamara Western gave me many advices for Arabidopsis promoter::GUS construct design and transformation.  I would like to thank you to Dr. Paul Knox kindly provided xylan antibodies (LM10 and LM11).   xii  Co-Authorship Statement  Chapter 2  Cryofixation and freeze substitution of pine xylem samples were prepared by Dr. Kim Rensing. The measurement of 3H-phenylalanine incorporation into secondary cell wall was done by Brian Banno and John Wong. All other experiments in this chapter were designed and executed by Minako K.   Chapter 3  Billy Lin and Colin McLeod contributed to phenotype observation of T-DNA insertion mutants. For promoter analysis, Dr. Bjoern Hamberger designed P, A and L DNA sequences elements. All other experiments in this chapter were designed and executed by Minako K.   Chapter 4  The root assay of potassium transport with inhibitors in barley was done by Brian Banno. A part of root procambium feeding with inhibitors was executed by Sarah Raven. All other experiments in this chapter were designed and tested by Minako K.   Chapter 5  All ultrastructure study and immunolocalization assays of anti-xylan and anti-mannan were done by Minako K.             1                 CHAPTER 1:  GENERAL INTRODUCTION  2  CHAPTER 1: GENERAL INTRODUCTION  1.1  Xylogenesis Xylogenesis can be defined as the series of common developmental steps found in differentiation of both primary xylem (produced via procambium by apical meristems) and secondary xylem (produced by the vascular cambium). 1.1.1 Primary xylem vs. secondary xylem Xylem is the water conduction tissue in plants that is produced in its primary xylem form at apical meristems as procambium or in its secondary xylem form in the vascular cambium of the secondary vascular system (Esau, 1965). Both primary and secondary xylem are water and mineral conducting tissues in plants. Xylem is made up of both lifeless tubes with thick secondary cell walls and living cells such as parenchyma. The general function of xylem requires strong and lignified secondary cell walls in the tracheary elements and fibers. In addition, the shapes of secondary cell walls in xylem are distinctive.  Primary xylem develops unique annular, spiral or reticular cell wall thickenings in the tracheary elements (McCann, 1997), and they are reinforced in some, but not all, of its surface.  On the other hand, secondary xylem tracheary elements have heavily pitted secondary cell walls throughout the cell surface.  Secondary xylem of angiosperms and gymnosperms each have unique modifications to their walls (Esau, 1965). Vessels, large tracheary elements characteristic of angiosperms, have perforated end walls with simple pits. Around them, there are fibers, also lifeless at maturity, which lend mechanical support, and xylem parenchyma, some of which are long-lived, to give metabolic support (Chaffey et al,  3 2001). In contrast, gymnosperm secondary xylem consists predominantly of one cell type, tracheids, with bordered pits and relatively little xylem parenchyma (Esau, 1965).  Wood is secondary xylem, a tissue in which there are both lifeless lignified cells providing support and water conduits, as well as living cells with diverse metabolic activities. The majority of the axial cells are lifeless tracheary elements and fibers.  In angiosperms, living axial parenchyma also make up the secondary xylem. In both gymnosperms and angiosperms, ray parenchyma cells are vitally active extending through the dead axial cells of the secondary xylem to the pith or heartwood.  These cells can form a lignified secondary cell wall very late in development, long after vessels/tracheids and fibers have matured. Their function is mainly transporting substances radially and storage but complete details of their function are still unknown. However, multiple phenylpropanoid pathway genes such as phenylalanine ammonia- lyase (PAL) and 4-coumarate CoA-ligase (4CL) are highly expressed in rays (Takabe et al. 2001). The transport of phenolics such as lignans into heartwood from the sapwood along the rays is one example of the active function of rays (Davin and Lewis, 2000). The extent of metabolic exchange between lignifying cells and rays has been speculated upon (Davin and Lewis, 2000) but is unknown. 1.1.2  Stages of cell differentiation in the secondary cell wall All xylem cells that lignify undergo common developmental steps: differentiation begins with cell expansion, secondary cell wall deposition of polysaccharides and lignin, and finally programmed cell death (Roberts and McCann 2000; Fukuda, 1996; Samuels et al., 2006). Zinnia elegans tracheary elements grown in cell culture medium have been used as a model system to study xylem differentiation steps such as secondary wall deposition, lignification and programmed cell death (Fukuda 1997, Stacey et al. 1995). The morphological shifts seen in  4 Zinnia culture system resemble the cell differentiation sequence seen in planta. This system has influenced research on xylem development by defining a common sequence of biochemical and morphological events that are necessary to build a tracheary element, as shown in Figure 1.1. Although they are derived from different cell types, lignifying cells from both primary and secondary xylem undergo these common developmental steps (see Fig. 1.1 and Fig.1.2). The first step in xylem differentiation is cell expansion. Angiosperm vessels and gymnosperm tracheids, the cells that will conduct water, undergo radial expansion, while supportive fibers of angiosperms undergo intrusive elongation (reviewed by Mellerowicz et al., 2001).  Expansins, proteins which are reported to aid in wall loosening (Im et al., 2000), are co- expressed with cell-cycle genes in xylem mother cells, indicating that completion of mitosis and cell expansion are closely linked (Schrader et al., 2004).  While several studies have reported that expansin genes are present during xylogenesis (Zhao et al., 2005), a recent study not only identified which hybrid poplar expansin genes family members are present in the expanding cambial derivatives (-expansin subfamily A), but also demonstrated by elegant in situ RT-PCR that the PttEXP1 mRNA is found in the tips of fusiform initials as well as in intrusively growing fibers (Gray-Mitsumune et al., 2004).   5   Figure 1.1 Xylogenesis –an example of tracheary element development in Zinnia.  Xylogenesis in Zinnia elegans in vitro. Transdifferentiated mesophyll cells are cultured under conditions that induce them to differentiate to tracheary elements.  The steps of differentiation are expansion, secondary cell wall polysaccharide deposition, followed by lignification and programmed cell death.  6  Figure 1.2 Xylogenesis –xylem development in poplar wood. Xylogenesis processes are very similar in primary xylem lignified cells and secondary xylem, although secondary xylem arises from the secondary vascular cambium. For vessels of angiosperms, and to a lesser extent for gymnosperm tracheids, expansion is strong in the radial direction, followed by secondary wall production and protoplast lysis.  For angiosperm fibers, expansion is axial as the cells elongate by intrusive growth, followed by secondary cell wall synthesis and protoplast lysis.  7 Xyloglucan endotransglucosylases/hydrolases (XTHs) are proposed to be important in wall- loosening and rearrangement of primary cell walls by cutting and reforming xyloglucan chains (Hyodo et al. 2003). In developing xylem of poplar, however, XET activity was correlated with the end of radial expansion and beginning of secondary cell wall synthesis (Bourquin et al. 2002, Mellerowicz et al. 2001), leading the authors to suggest that the role of this enzyme is to remodel the primary wall for integration with the secondary cell wall. At the end of the cell expansion phase, the protoplast begins to produce the thickened secondary cell wall, a three-layered structure (S1, S2, S3 : Fig.1.3) made of cellulose microfibrils (Wardrop and Harada, 1965; Donaldson, 2001).  The microfibril orientations of secondary cell wall layers during development have been examined using field emission- scanning electron microscopy (FESEM), which confirmed earlier studies using light microscopy with birefringence and transmission electron microscopy (reviewed by Abe and Funada, 2005).  8  Figure 1.3 Primary and secondary cell wall models Model of the cell wall structure of a softwood tracheid.    9 A generally accepted model of cellulose synthesis is that the functional cellulose synthase complex (called a rosette or terminal complex) consists of ß-glucosyltransferase (cellulose synthase) subunits encoded by CESA genes (Doblin et al. 2002).  In a variety of taxa, recent studies support the view that cellulose synthesis enzymes for secondary cell wall formation are different from the cellulose synthases used in primary cell wall formation.  In Arabidopsis, three CesA genes (AtCesA1, AtCesA3 and AtCesA6) are required for primary cell wall deposition, while other CesA genes (AtCesA4, AtCesA7, AtCesA8) work together to produce secondary cell wall (Turner 2007). The cellulose synthases responsible for secondary cell wall deposition in hybrid poplar (Joshi et al. 2004) and pine (Nairn and Haselkorn, 2005) have been identified. For example, in situ mRNA hybridization results indicated that 3 poplar CesA genes (PtrCes1, PtrCes2, and PtrCes3) were expressed in developing xylem and phloem fibers undergoing secondary cell wall formation (Joshi et al. 2004). In loblolly pine, semi-quantitative RT-PCR was used to examine developing wood and these data suggest three CesA genes (PtCesA1, PtCesA2 and PtCesA3) are highly expressed in wood but not in the needle and lateral shoot (Nairn and Haselkorn, 2005).  A multiple alignment of full-length CesA protein sequences showed that secondary CesAs in pine are homologues of Arabidopsis and poplar secondary wall CesAs (Nairn and Haselkorn, 2005).  Primary cell wall CesAs and secondary cell wall CesAs might be specialized to form cellulose microfibrils in different environments, e.g. the pectin rich primary cell wall during expansive growth or the relatively low pectin, ordered wall layers of secondary cell wall. Biased wall thickenings in the tracheary element are correlated with parallel cortical microtubule arrangement under the plasma membrane (Hepler and Newcomb, 1964; Giddings et al. 1991). In protoxylem tracheary elements of Arabidopsis roots, when microtubules were  10 disrupted with oryzalin, CesA complexes depositing spiral wall thickenings lost their normal spiral pattern (Gardiner et al. 2003). In the Zinnia system, secondary cell walls are formed along microtubules arranged in spiral patterns (Ingold et al. 1988).  In developing wood of both angiosperms and gymnosperms, secondary cell wall polysaccharide deposition is correlated with parallel arrays of cortical microtubules (Chaffey et al., 1997; Abe et al., 1995). It is suggested that microtubules play a role in microfibril organization but how the cortical microtubules guide cellulose microfibrils alignment is still not known. Expression of one of the primary cell wall cellulose synthases, CesA6, fused with a yellow fluorescent protein (YFP) in Arabidopsis plants showed that microtubules were not required for CESA complexes to move in “highly coordinated” trajectories (Paredez et al. 2006).  However, partial disruption of the microtubules with oryzalin disorganized CESA complex distribution (Paredez et al. 2006).  Disorganization of microtubule organization and dynamics in the Arabidopsis mor1 mutant leads to no change in the cellulose microfibril orientation but the cellulose microfibril function is altered, as shown by loss of growth anisotropy in the root and stem (Himmelspach et al., 2003; Kawamura, 2006; Fujita, 2008). Current cell wall models have the cellulose microfibrils coated with hemicelluloses via hydrogen bonding.  In the secondary cell wall, hemicellulose is a predominant component, which cross-links cellulose microfibrils, and provides a complex matrix for embedding the lignin polymer.  Xylans are the major secondary cell wall hemicellulose components by mass in wood and grasses (Ebringerova and Heinze, 2000). The backbone of xylans is 1,4-linked ß- D-xylosyl residues with short side chains. To produce such a complex molecules, numerous numbers of enzymes are involved in the backbone elongation and side chain residue addition. Awano’s group (2002) showed that, in fibers of Fagus crenata wood, xylan was continuously  11 accumulating around the cellulose microfibrils before the lignification.  Genetic screens of collapsed xylem phenotype identified an Arabidopsis glycosyltransferase related mutant, irregular xylem8 (irx8).  The irx8 is deficient in normal xylan production leading to secondary cell wall disruption (Persson et al. 2007), suggesting that xylan is required for secondary cell wall integrity. Mannan hemicelluloses are also wide spread in higher plants. The cellulose microfibrils can crosslink to mannans, indicating that mannan plays some structural role in secondary cell walls.  A lignin/xylan matrix is formed after cellulose microfibril and lignin deposition, which develops the water resistance of the lignified wall, and protects against microbial invasion (Lawoko et al. 2003).  However, which types of lignin polymers such as H-, G- or S-lignin have preference to interact with xylan or mannan hemicelluloses have not been studied. Lignin is accumulated in the middle lamella, primary and secondary cell walls in xylem. Initiation of lignification starts in the middle lamella (Terashima et al. 1986). Then, lignin is deposited throughout the primary and secondary cell wall layers including S1, S2, and S3 (Donaldson et al. 2001, Fig. 1.3). Polymerization of lignin precursors occurs through coupling of monolignols to the growing lignin polymer following activation to superoxide radicals, by peroxidases and laccases (Olson and Varner 1993, Sato et al., 2006). However, regulation of the growth of the lignin polymer is not well understood. There are two models that have been suggested: the random coupling model (Hatfield and Vermerris, 2001), where the cell wall polysaccharide matrix as well as the “type and quantity of monolignols at the lignification site control lignin formation” or the dirigent protein model (Davin and Lewis, 2005).  Dirigent proteins mediate stereoselective coupling of coniferyl alcohol to pinoresinol (Davin et al., 1997) but the evidence for their role in lignification is circumstantial.  The current view of this  12 debate seems to be that the control of lignin polymerization is not protein mediated (Ralph et al., 2004, 2008). The last step in xylogenesis is programmed cell death (PCD) (McCann 1997). The vacuole breaks down, and then finally all cell contents are degraded except secondary cell wall (Beers et al. 2001). The mechanism of PCD has been studied in the Zinnia system. The transcripts for cell death enzymes and enzymes involved in secondary cell wall lignification are up-regulated at the same time (Demura et al. 2002). This implies that a common signal may induce both cell death and in secondary cell wall lignification (Fukuda 1997).  More recently, gene expression studies of PCD in poplar have described genes up-regulated during cell death in secondary xylem (Courtois-Moreau et al. 2008). 1.2 Lignin biosynthesis pathway in secondary cell wall formation 1.2.1  Lignin Lignin is a complex polymer of monolignols, which is produced through the phenylpropanoid pathway in a series of enzymatic reactions from phenylalanine (Boerjan et al., 2003). This amino acid is produced by the shikimate pathway, although the enzymes catalyzing all steps of carbon flow through the phenylalanine branch of this pathway have not been elucidated (Ehlting et al., 2005).  The general phenylpropanoid pathway supplies carbon skeletons to both monolignol biosynthesis and flavonoid production (Besseau et al., 2007).  Because lignin plays such essential roles in plants, disturbing normal lignification has a large impact on plant growth.  Forward genetics approaches in Arabidopsis have identified irregular xylem mutants, some of which have defects in lignin biosynthesis (Turner 2007).  For example, a severe lignin mutant, irx4, in Arabidopsis thaliana has 50 % reduced lignin comparing to a wild type. In this mutant, which is deficient in cinnamoyl-CoA reductase, the  13 reduced lignin phenotype causes collapse of the xylem cells, as well as reduced fertility and growth rate in the Arabidopsis stem. Secondary cell walls in irx4 stems have diffusely expanded and uneven secondary cell wall thickenings (Jones et al. 2001).  Another phenotype associated with secondary cell wall defects is fragile or floppy inflorescence stems. The ifl1 mutant is not able to initiate normal interfascicular fiber differentiation in stem of Arabidopsis and has a severe phenotype where the stem of ifl1 mutant doesn’t function as physical support for the plant (Zhong et al. 1997). In trees, almost all of tissue is xylem, which contains abundant lignin.  Downregulation of phenylpropanoid genes in trees leads to a variety of phenotypes including changes in metabolic phenolic pools, irregular vessels and decreased plant growth and development (Coleman et al, 2008; Vanholme et al., 2008). 1.2.2  The enzymes of phenylpropanoid metabolism in angiosperms and gymnosperms.  Lignin has three different types of units, p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units derived from the polymerization of three monolignols: p-coumaryl alcohol, coniferyl alcohol and sinapyl alcohol, respectively (Fig.1.4). The first part of monolignol biosynthesis is called the general phenylpropanoid pathway. Phenylalanine, following de-amination to cinnamate by phenylalanine ammonia-lyase (PAL), under goes a hydroxylation reaction to become p-coumaric acid by cinnamate 4- hydroxylase (C4H). Then p-coumaric acid is conjugated to the co-enzyme A (CoA) to form an ester, p-coumaryl CoA, by the enzyme 4-(hydroxy) cinnamoyl CoA ligase (4CL) and p- coumaryl CoA represents the branchpoint for flavonoids and monolignol biosynthesis.  14  Figure 1.4 Phenylpropanoid pathway in angiosperm and gymnosperm (Humphreys and Chapple, 2002). 4CL, 4-(hydroxy)cinnamoyl CoA ligase; C3H, p-coumarate 3-hydroxylase; C4H, cinnamate 4-hydroxylase; CAD, cinnamyl alcohol dehydrogenase; CCoAOMT, caffeoyl CoA O-methyltransferase; CCR, cinnamoyl CoA reductase; COMT, caffeic acid/5- hydroxyferulic acid O-methyltransferase; CQT, hydroxycinnamoyl CoA:quinate hydroxycinnamoyltransferase; CST, hydroxycinnamoyl CoA:shikimate hydroxycinnamoyltransferase; F5H, ferulate 5-hydroxylase; PAL, phenylalanine ammonia-lyase; pCCoA3H, p-coumaryl CoA 3-hydroxylase; SAD, sinapyl alcohol dehydrogenase.  15 The monolignol biosynthesis pathway starts from here to the three monolignols p-coumaryl, coniferyl and sinapyl alcohol (reviewed by Humphrey and Chapple, 2002, Boerjan et al., 2003). The simplest but minor monolignol, p-coumaryl alcohol, is synthesized by conversion of p-coumaryl CoA to the corresponding aldehyde by cinnamoyl coenzymeA  reductase (CCR) and then reduction of the aldehyde to the alcohol by cinnamoyl alcohol dehydrogenase (CAD). The other two monolignols require additions of methoxy groups at the 3’ position (coniferyl) or 3’ and 5’positions (sinapyl) of the aromatic ring. For production of methoxylated monolignols, p-coumaryl-CoA is esterified to shikimate or quinate by hydroxycinnamoyltransferase (CQT/CST).  Then the p-coumaryl ester is the substrate for p-coumarate 3-hydroxylase (C3H), which adds the hydroxy-group to 3 position of the phenolic ring. The phenolic portion of the ester is then conjugated to CoA again and caffeoyl-CoA O-methyltransferase (CCoAOMT) adds the methyl group to complete the methoxy at the 3 position, forming feruloyl-CoA. Finally coniferyl alcohol (G-lignin unit) is formed from this branch following reduction by CCR and CAD. Sinapyl alcohol synthesis continues the modification of the phenolic ring to add another methoxy group with ferulate 5-hydroxylase (F5H) and caffeic acid/5- hydroxyconiferaldehyde O-methyltransferase (COMT) although it is not clear if this methoxylation happens at the level of the aldehyde or alcohol, i.e. before reduction by CCR alone or CCR and CAD. Interesting phenotypes of genetic mis-regulation of many of these enzymes in this pathway has been reported. The mis-regulation of genes encoding these biosynthetic enzymes leads to different type of monomers incorporated into the lignin polymer (Vanholme et al., 2008).  For example, downregulated HCT leads to high levels of the usually minor H-lignin (Besseau et al., 2007), while downregulation of F5H leads to high G lignin in normally S-rich cells such as fibers (Franke et al., 2002).  The incorporation of these different  16 types of units in mutant plant lines suggests a “non-specific transport route” is able to export a variety of substrates out of the cytoplasm during lignification (VanHolme et al., 2008). 1.2.3 Subcellular localization of monolignol synthesis enzymes The location of monolignol biosynthesis is relevant to the understanding of lignification mechanisms including monolignol transport into the secondary cell wall. More than 10 enzymes are involved in the monolignol biosynthesis pathway. All of the genes encoding these enzymes have been isolated in Arabidopsis (Raes et al. 2003) and the locations of the gene products, i.e. the monolignol biosynthetic enzymes, have been determined.  Most of the enzyme localization studies have used immuno-gold labeling with electron microscopy or green fluorescent protein (GFP)-enzyme fusions with confocal laser microscopy. Takabe’s group showed that the PAL enzyme localizes in the cytosol in poplar secondary xylem using immuno-gold transmission electron microscopy (TEM) (Takabe et. al 2001).  The next enzyme of hydroxylation in the pathway, C4H enzyme, was localized to the endoplasmic reticulum (ER) using GFP fused to C4H and visualized with confocal laser microscopic analysis (Ro et al., 2001). COMT enzyme and CCoAOMT enzyme are thought to be responsible for methylation steps in monolignol synthesis and these enzymes were localized in the cytosol, ER and Golgi apparatus in poplar developing xylem (Takeuchi et al. 2001). Two cytochrome P450 hydroxylation enzymes, C3H and F5H, have been described as low abundance and are predicted to be localized to the ER (Raes et al. 2003).  CAD catalyzes the last step of monolignol biosynthesis. Under electron microscopic observation of poplar, antibodies specific for CAD label both polysomes and cytosol, with low label detected in ER, Golgi apparatus and cell wall (Takabe et al. 2001). In addition, the subcellular localization of  17 the CAD protein was to the cytosol and ER in poplar developing xylem by immuno-gold cytochemistry (Samaj et al 1998). Even though the sensitivity of immunolocalization is limited to show clearly the localization of enzymes, these results suggested that monolignols are synthesized in cytoplasm with some steps associated with the ER. This raises the question of what is the mechanism of transport of monolignols from the cytosol to the site of polymerization in the cell wall. 1.3 Monolignol export models: vesicle fusion or membrane transport 1.3.1  Historic theory and early autoradiography methods. The mechanism of monolignol export is unknown. The current theory of Golgi-mediated monolignol export is primarily based on autoradiography studies. The value of the early autoradiography studies at the light microscope level was to show the sequence of lignification. A number of studies feeding different intermediates showed that cell corners and middle lamella lignified first and that early lignin is mostly H-lignin, followed by lignification of the S2 layers and lastly the S3 layers with G-lignin in Cryptomeria japonica (Terashima and Fukushima, 1989) and Pinus thunbergii (Takabe et al., 1981). Autoradiography at the TEM level showed the indistinct localization of monolignol precursors in chemically fixed primary xylem (Pickett-Heaps, 1968) and in secondary xylem (Fujita et al., 1979; Wooding, 1968). When developing xylem of wheat coleoptiles was treated with 3H-cinnamic acid, then localized with autoradiography, phenylpropanoid products were found in the ER and Golgi body (Pickett-Heaps, 1967).  Similar results were obtained when Takabe et al. (1985) fed 3H- phenylalanine to developing wood of Cryptomeria japonica, then prepared the samples for TEM autoradiography. Since samples fed 3H-lignin precursors for autoradiography were chemically fixed, the quality of preservation of cell structures was low. In addition to that, a  18 lack of quantification meant that data in autoradiography studies at the electron microscopic level were not precise for lignin precursor distribution. In cases where phenylalanine was used, it was not possible to distinguish between protein incorporation and monolignol incorporation.   Another study that contributed to the theory of Golgi-mediated export of monolignol precursors used immuno-labeling of developing secondary xylem cells in chemically fixed samples. Antibodies specific for CAD, one of the late stage monolignol biosynthetic enzymes, localized on Golgi vesicles in poplar, leading the authors to suggest a possible vesicle secretory pathway of monolignols (Samaj et al 1998). Together, these studies did not strongly support the Golgi-mediated export hypothesis, so one can consider possible alternative hypotheses. 1.3.2 Alternative hypotheses: Plasma membrane monolignol transporters A possible alternative mechanism is that membrane transporters pump monolignols across the plasma membrane to the secondary cell wall.  Normally, molecules diffuse in plants following energetically favorable paths determined by their concentration gradient. Some of the transporters/pumps can transport substrates against their concentration gradient by using ATP, which is called active transport.  Transporting mechanisms can either depend directly on ATP hydrolysis or indirectly use the energy of ATP hydrolysis by using the proton concentration gradient at the plasma membrane established by the proton-ATPase (Pardo and Serrano, 1989; Sussman and Harper, 1989). Transporters can be classified into superfamilies (Transport Classification data base, http://www.tcdb.org/tcdb) based on their energetics and conserved structure.  Within the larger transporter classification, transporters that confer multidrug resistance and export of xenobiotics or secondary metabolites consist of five superfamilies: ATP binding cassettes (ABC) superfamily, the major facilitator superfamily, the small multidrug  19 resistance superfamily, the resistance-nodulation-cell division superfamily and the multidrug and toxic compound extrusion (MATE) family of transporters (Yazaki 2005; Omote et al. 2006). 1.3.3 ATP binding cassette (ABC) transporters ATP binding cassette (ABC) transporters are the largest transporter family, and have been found in bacteria, fungi, plant and animals (Martinoia et al. 2002, Rea, 2007). In diverse systems like humans and fungi, ABC transporters can translocate a broad range of polar and non-polar substrates across biological membranes (Dean et al. 2001). In Arabidopsis, 129 putative ABC transporters have been identified based on conserved ATP binding sequences (Sanchez-Fernandez et al. 2001). Most of the functions are unknown in plants yet (Rea, 2007), although ABC transporters have been implicated in auxin transport (reviewed by Geisler and Murphy, 2006), heavy metal detoxification (Rea, 1999), and secondary metabolite transport (Jasinski et al. 2001; Shitan et al. 2003, Yazaki, 2006). Wax deposition to the cuticle in Arabidopsis stem involves two ABC transporters found in the plasma membrane of epidermal cells (ABCG12/WBC12/CER5) (Pighin et al., 2005) and ABCG11/WBC11 (Bird et al., 2007). Specific ABC transporters are expressed in lignin rich tissues. Ehlting et al. (2005) determined using microarrays, that multiple ABC transporter genes are differentially expressed along the axis of bolting stems in Arabidopsis. The gene expression analyses showed lignin biosynthesis pathway enzyme genes and multiple ABC transporter genes are co-expressed in developing Arabidopsis stems. In several studies of gene expression during wood formation, expressed sequence tags (ESTs) for ABC transporters have been reported in loblolly pine and Zinnia culture (Allona et al., 1998; Kirst et al. 2003; Hertzberg et al., 2001), suggesting ABC transporter genes might be involved in secondary cell wall formation including lignification. If  20 ABC transporters can export diverse substrates like auxin and alkaloids, then perhaps they can also be responsible for monolignol export.  21 1.4 Objectives  The primary goal of this thesis is to understand the mechanism(s) of monolignol deposition during xylogenesis. The currently accepted theory is that monolignols are exported by Golgi- mediated vesicle delivery to the secondary cell wall. Since the current theory was tested by microscopy methods that were outdated, I have re-examined the Golgi-mediated vesicle delivery hypothesis using new technology. In order to test this hypothesis, it was necessary to localize lignin precursors within the cell during the lignification. Monolignols were localized by feeding 3H-phenylalanine to developing wood and then cryofixation was used to preserve the cells and their contents. The localization of incorporated lignin precursors could then accurately be visualized with autoradiography. Since phenylalanine can be incorporated into both lignin precursors and proteins, inhibitors were used to distinguish the two pathways. Incorporated products were quantified in the different subcellular compartments with inhibitor treatments. At the same time, incorporated compounds were analyzed biochemically by HPLC and scintillation counting.  These studies have been published in Plant Physiology and are described in Chapter 2, entitled Tracking monolignols during secondary cell wall Development in Pinus contorta var. latifolia and Populus deltoides x P. trichocarpa. An alternative hypothesis is that instead of Golgi vesicles transporting the monolignols to the cell wall, plasma membrane transporter proteins would pump the monolignols out of the cell. I hypothesized that ABC transporters are involved in lignin precursor export to the cell wall in developing xylem. While other transporters could be involved in monolignol export, I have chosen to test the ABC transporters for several reasons. ABC transporters are required for transport of hydrophobic compounds and secondary metabolites. Some ABC transporter  22 encoding genes are up-regulated in xylem tissue in pine, poplar and Zinnia (Whetten et al. 2001, Hertzberg et al. 2001, Demura et al. 2002). In the developing stem of Arabidopsis, ABC transporter genes were coordinately expressed with phenylpropanoid biosynthesis enzyme genes (Ehlting et al. 2005). The ABC transporter hypothesis was tested in two ways. In chapter 3, entitled ABC transporters coordinately expressed during lignification of Arabidopsis stems include a set of ABCB's associated with auxin transport, reverse genetics approaches are described to test the role of specific ABC transporters. These ABC transporter genes were chosen based on the expression data of Ehlting et al. (2005) and bioinformatic analysis. During this study, T- DNA insertion mutants with disruption of single ABC transporter genes were tested, not only for normal lignin deposition, but also for polar auxin transport and normal development of lignified cells. In chapter 4, entitled Testing the role of ABC transporters on lignin deposition during xylogenesis using inhibitors, lignifying cells were treated with ABC transporter inhibitors to examine if lignification was disrupted. Secondary xylem of pine and poplar was used but gave contradictory results and the experiments reuired many months, so I developed a new simple assay using Arabidopsis root primary xylem development. Using this assay, a wide range of inhibitors was tested on roots, and the effect on single lignifying cells could be monitored. Experiments in chapter 2, 3 and 4 all addressed how the monolignols are exported to the cell wall where they are polymerized. In chapter 5, entitled Cell specific deposition of hemicellulose in developing secondary cell walls of hybrid poplar (Populus deltoides x P. trichocarpa) vessels and fibers, I studied the secretion and distribution of hemicelluloses, cell wall polysaccharides that may influence the organization of monolignols prior to/during polymerization.  23 Based on the observation that diverse lignified cell types are enriched in either G-lignin, in tracheary elements, or S-lignin, in fibers, I hypothesized that different cell types have different hemicellulose distributions. Hemicelluloses are important components in secondary cell walls where they cross link cellulose microfibrils with lignin. I tracked the xylan and mannan (hemicelluloses) distribution and deposition during secondary xylem development in poplar. Using cryofixation and immuno-labeling techniques, I evaluated whether deposition occurred in a cell type specific manner. In summary, I performed the following studies, each of which is presented as one chapter of this thesis.  1. Tracking Monolignols during secondary cell wall Development in Pinus contorta var. latifolia and Populus deltoides x P. trichocarpa. 2. ABC transporters coordinately expressed during lignification of Arabidopsis stems include a set of ABCB's associated with auxin transport 3. Testing the role of ABC transporters on lignin deposition during xylogenesis using inhibitors 4. Cell specific deposition of hemicellulose in developing secondary cell walls of hybrid poplar (Populus deltoides x P. trichocarpa) vessels and fibers    24  1.5 Bibliography Abe, H., Funada R, Ohtani J, Fukazawa K. (1995) Changes in the arrangement of microtubules and microfibrils in differentiating conifer tracheids during the expansion of cells. Ann Bot 75, 305-310. Abe, H., and Funada, R. (2005). The orientation of cellulose microfibrils in the cell walls of tracheids in conifers. A model based on observations by field  emission-scanning electron microscopy. IAWA J. 26, 161-174. Allona, I., Quinn M, Shoop E, Swope K, Carlis J, Riedl J, Retzel E, Campbell MM, Sederoff R, Whetten RW. (1998) Analysis of xylem formation in pine by cDNA sequencing. Proc Natl Acad Sci USA 95, 9693-9698. Awano, T., Takabe K, Fujita M. (2002) Xylan deposition on secondary wall of Fagus crenata fiber. Protoplasma 219, 106-115. Beers, EP., McDowell JM. (2001) Regulation and execution of programmed cell death in response to pathogens, stress and developmental cues. Curr Opin Plant Biol. 4, 561- 567 Besseau, S., Hoffmann L, Geoffroy P, Lapierre C, Pollet B, Legrand M. (2007) Flavonoid accumulation in Arabidopsis repressed in lignin synthesis affects auxin transport and plant growth. Plant Cell 19, 148-162. Bird, D., Beisson F, Brigham A, Shin J, Greer S, Jetter R, Kunst L, Wu X, Yephremov A, Samuels L. (2007) Characterization of Arabidopsis ABCG11/WBC11, an ATP binding cassette (ABC) transporter that is required for cuticular lipid secretion.  Plant J 52,485-98 Boerjan, W., Ralph J, Baucher M. (2003). Lignin biosynthesis. Annu Rev Plant Biol 54, 519- 546. Bourquin, V., Nishikubo, N., Abe, H., Brumer, H., Denman, S., Eklund, M., Christiernin, M., Teeri, T.T., Sundberg, B. and Mellerowicz, E.J. (2002) Xyloglucan endotransglycosylases have a function during the formation of secondary cell walls of vascular tissues. Plant Cell, 14: 3073-3088 Chaffey, NJ., Barnett JR, Barlow PW. (1997) Visualization of the cytoskeleton within the secondary vascular system of hardwood species. J Microsc  2, 77-84. Chaffey, N., Barlow P, Barnett J (2001) The cytoskeleton facilitates a three-dimensional symplasmic continuum in the long-lived ray and axial parenchyma cells of angiosperm trees. Planta 214, 330-1. Coleman, HD., Park JY, Nair R, Chapple C, Mansfield SD. (2008) RNAi-mediated suppression of p-coumaroyl-CoA 3'-hydroxylase in hybrid poplar impacts lignin deposition and soluble secondary metabolism. Proc Natl Acad Sci USA 105, 4501-4506. Courtois-Moreau, CP., Edouard; Sjodin, Andreas; Muñiz, Luis; Bollhöner, Benjamin; Kaneda, Minako; Samuels, Lacey; Jansson, Stefan; Tuominen, Hannele. (2008) A unique program for cell death in xylem fibers of Populus stem. Plant J, in press. Davin, LB., Lewis, N. G. (2000). Dirigent Proteins and Dirigent Sites Explain the Mystery of Specificity of Radical Precursor Coupling in Lignan and Lignin Biosynthesis. Plant Physiol. 123, 453-462 Dean, M., Hamon Y, Chimini G. Human (2001) The human ATP-binding cassette (ABC) transporter superfamily. Genome Res. 11: 1156-66  25 Demura, T., Tashiro G, Horiguchi G, Kishimoto N, Kubo M, Matsuoka N, Minami A, Nagata- Hiwatashi M, Nakamura K, Okamura Y, Sassa N, Suzuki S, Yazaki J, Kikuchi S, Fukuda H. (2002) Visualization by comprehensive microarray analysis of gene expression programs during transdifferentiation of mesophyll cells into xylem cells. Proc Natl Acad Sci USA. 99, 15794-9 Doblin, MS., Kurek I, Jacob-Wilk D, Delmer DP. (2002) Cellulose biosynthesis in plants: from genes to rosettes. Plant Cell Physiol 43, 1407-1420. Donaldson, LA (2001) Lignification and lignin topochemistry-an ultrastructural view. Phytochem 57, 859-873. Ebringerová, A., and Heinze, T. (2000). Xylan and xylan derivatives - Biopolymers with valuable properties. I. Naturally occurring xylans structures, isolation procedures and properties. Macromol. Rapid Commun. 21, 542–556. Ehlting, J., Mattheus N, Aeschliman DS, Li E, Hamberger B, Cullis IF, Zhuang J, Kaneda M, Mansfield SD, Samuels L, Ritland K, Ellis BE, Bohlmann J, Douglas CJ. (2005) Global transcript profiling of primary stems from Arabidopsis thaliana identifies candidate genes for missing links in lignin biosynthesis and transcriptional regulators of fiber differentiation. Plant J 42, 618-640. Esau, K. (1965) Plant Anatomy 2nd. Ed.,  John Wiley and Sons, Inc., New York. Franke, R., Hemm MR,  Denault JW, Ruegger MO, Humphreys JM, Chapple C (2002) Changes in secondary metabolism and deposition of an unusual lignin in the ref8 mutant of Arabidopsis.  Plant J 30, 47-59 Fujita, M., Keiji Takabe, Hiroshi Harada (1979) Autoradiographic Investigations of Cell Wall Development II. Tritiated phenylalanine adn ferulic acid assimilation in relation to lignification. Mokuzai Gakkaishi. 25, 89-94 Fujita, M., (2008) Cortical microtubules and physical properties of cellulose microfibrils during primary cell wall formation in Arabidopsis thaliana. Ph.D thesis, University of British Columbia, Canada Fukuda, H., (1996) Xylogenesis: Initiation progression and cell death. Plant Mol. Biol. 47, 299- 325 Fukuda, H. (1997) Tracheary element differentiation Plant Cell 9, 1147-1156 Gardiner, JC., Neil G.Taylor , and Simon R. Turner (2003) Control of cellulose synthase complex localization in developing xylem. Plant Cell 15, 1740-1748 Geisler, M., Murphy AS. (2006) The ABC of auxin transport: the role of p-glycoproteins in plant development. FEBS Lett 580, 1094-1102. Giddings, TH., Jr, Staehelin LA (1991). Microtubule-mediated control of microfibril deposition: a re-examination of the hypothesis. In Lloyd CW, ed, The Cytoskeletal Basis of Plant Growth and Form. Academic Press, San Diego, CA, - Gray-Mitsumune, M., Mellerowicz EJ, Abe H, Schrader J, Winzell A, Sterky F, Blomqvist K, McQueen-Mason S, Teeri TT, Sundberg B. (2004) Expansins abundant in secondary xylem belong to subgroup A of the alpha-expansin gene family. Plant Physiol. 135, 1552-1564. Hatfield, R., Vermerris W.( 2001) Lignin formation in plants. The dilemma of linkage specificity. Plant Physiol. 126, 1351-1357. Hepler, PK. and E. H. Newcomb (1964) Microtubules and Fibrils in the Cytoplasm of Coleus Cells Undergoing Secondary Wall Deposition. J. Cell Biol. 20, 529-533  26 Hertzberg, H. A., J. Schrader, A. Andersson, R. Erlandsson, K. Blomqvist, R. Bhalerao, M. Uhlén, T. Teeri, J. Lundeberg, B. Sundberg, P.Nilsson, and G.Sandberg.(2001) A transcriptional roadmap to wood formation Proc Natl Acad Sci USA 98, 14732- 14737 Himmelspach, R., Williamson RE, Wasteneys GO.( 2003) Cellulose microfibril alignment recovers from DCB-induced disruption despite microtubule disorganization. Plant J 36, 565-575. Humphreys, J., M.  M.R. Hemm, and C.Chapple (1999) New routes for lignin biosynthesis defined by biochemical characterization of recombinant ferulate 5-hydroxylase, a multifunctional cytochrome P450-dependent monooxygenase Proc Natl Acad Sci USA 96, 10045–10050 Humphrey, J., M., Chapple. (2002) Rewriting the lignin roadmap. Curr Opin Plant Biol 5, 224- 229 Hyodo, H., S. Yamakawa, Y. Takeda, M. Tsuduki, A. Yokota, K. Nishitani and T. Kohchi. (2003) Active gene expression of a xyloglucan endotransglucosylase/ hydrolase gene, XET9, in inflorescence apices is related to cell to cell elongation in Arabidopsis thaliana.  Plant Mole Biol 52, 472-483. Im, K., D. Cosgrove, and A. Jones (2000) Subcellular localization of expansin mRNA in xylem cells. Plant Physiol 123, 463-470 Ingold, E., Sugiyama M, Komamine A  (1988). Secondary cell wall formation: changes in cell wall constituents during the differentiation of isolated mesophyll cells of Zinnia elegans to tracheary elements. Plant Cell Physiol 29, 295-303 Jasinski, M., Yvan Stukkens,a Hervé Degand,a Bénédicte Purnelle,a Jacqueline Marchand- Brynaert,b and Marc Boutry (2001) A Plant Plasma Membrane ATP Binding Cassette–Type Transporter Is Involved in Antifungal Terpenoid Secretion Plant Cell 5, 1095–1108 Jones, L.,  A.R. Ennos and S.R. Turner (2001) Cloning and characterization of irregular xylem 4 (irx4): a severely lignin-deficient mutant of Arabidopsis. Plant J 26, 205-216 Joshi, CP., Suchita Bhandari, Priya Ranjan, Udaya C. Kalluri,  Xiaoe Liang, Takeshi Fujino and Anita Samuga (2004) Genomics of cellulose biosynthesis  in poplars New Phytol 164, 53–61. Kawamura, E., Himmelspach R, Rashbrooke MC, Whittington AT, Gale KR, Collings DA, Wasteneys GO. (2006) MICROTUBULE ORGANIZATION 1 regulates structure and function of microtubule arrays during mitosis and cytokinesis in the Arabidopsis root. Plant Physiol 140, 102-114. Kirst, M., Johnson AF, Baucom C, Ulrich E, Hubbard K, Staggs R, Paule C, Retzel E, Whetten R, Sederoff R. (2003) Apparent homology of expressed genes from wood-forming tissues of loblolly pine (Pinus taeda L.) with Arabidopsis thaliana. Proc Natl Acad Sci USA 100, 7383-7388. Lawoko, M., Henriksson G., Gellerstedt G. (2003) New method for quantitative preparation of lignin-carbohydrate complex from unbleached softwood kraft pulp: Lignin- polysaccharide networks Holzforschung 57, 69-74 Martinoia, E., Klein, M., Geisler, M., Bovet, B., Forestier, C., Kolukisaoglu,U., Müller-Röber, B. and Schulz,B. (2002) Multifunctionality of plant ABC transporters – more than just detoxifiers. Planta 214, 345 – 355  27 McCann, MC. (1997). Tracheary element formation: building up to a dead end. Trends Plant Sci 2, 333-338 Mellerowicz, Ewa J. Marie Baucher, Bjorn Sundberg, (2001) Unravelling cell wall formation in the woody dicot stem. Plant Mole Biol 47, 239 Nairn, CJ. and Haselkorn TH. (2005) Three loblolly pine CesA genes expressed in developing xylem are orthologous to secondary cell wall CesA genes of angiosperms. New Phytol 166, 907-917. Olson, PD., Varner JE. (1993) Hydrogen peroxide and lignification. Plant J 4, 887-892. Omote, H., Hiasa M, Matsumoto T, Otsuka M, Moriyama Y. 2006. The MATE proteins as fundamental transporters of metabolic and xenobiotic organic cations. Trends Pharmacol Sci 27, 587-593. Pardo, JM., Serrano R. 1989. Structure of a plasma membrane H+-ATPase gene from the plant Arabidopsis thaliana. J Biol Chem 264, 8557-8562. Paredez, AR., Somerville CR, Ehrhardt DW. (2006) Visualization of cellulose synthase demonstrates functional association with microtubules. Science 312, 1491-1495. Persson, S., Caffall KH, Freshour G, Hilley MT, Bauer S, Poindexter P, Hahn MG, Mohnen D, Somerville C. (2007) The Arabidopsis irregular xylem8 mutant is deficient in glucuronoxylan and homogalacturonan, which are essential for secondary cell wall integrity. Plant Cell 19, 237-255. Pickett-Heaps, JD. (1967) Further observations on the Golgi apparatus and its functions in cells of the wheat seedling. J Ultrastruct Res 18, 287-303. Pighin, JA., Zheng H, Balakshin LJ, Goodman IP, Western TL, Jetter R, Kunst L, Samuels AL. (2004) Plant Cuticular Lipid Export Requires an ABC Transporter. Science 306, 702- 704. Raes, J., A. Rohde, J. H. Christensen, Y. Van de Peer, and W. Boerjan (2003) Genome-Wide Characterization of the Lignification Toolbox in Arabidopsis. Plant Physiol 133, 1051–1071 Ralph, J., Knut Lundquist, Gosta Brunow, Fachuang Lu, Hoon Kim, Paul F. (2004) Lignins: Natural polymers from oxidative coupling of 4-hydroxyphenyl- Propanoids. Phytochemreviews 3, 29-60 Ralph, J., Kim H, Lu F, Grabber JH, Leple JC, Berrio-Sierra J, Derikvand MM, Jouanin L, Boerjan W, Lapierre C. (2008) Identification of the structure and origin of a thioacidolysis marker compound for ferulic acid incorporation into angiosperm lignins (and an indicator for cinnamoyl CoA reductase deficiency). Plant J 53, 368- 379. Rea, PA. (1999) MRP subfamily ABC transporters from plants and yeast. J Exp Bot 50, 895- 913 Rea, PA. (2007) Plant ATP-Binding Cassette Transporters. Annu Rev Plant Biol 58, 347-375. Ro, D.K., Mah N., Ellis B.E., Douglas C.J. (2001)  Functional characterization and subcellular localization of poplar (Populus trichocarpa x Populus deltoides) cinnamate 4- hydroxylase. Plant Physiol 126, 317-29. Roberts, K., McCann MC. (2000) Xylogenesis: the birth of a corpse. Curr Opin Plant Biol 6, 517-22 Samaj, J., Hawkins S, Lauvergeat V, Grima-Pettenati J, Boudet A, (1998) Immunolocalization of cinnamyl alcohol dehydrogenase 2 (CAD 2) indicates a good correlation with cell- specific activity of CAD 2 promoter in transgenic poplar shoots. Planta 204, 437-43.  28 Samuels, AL., Kaneda, M., Rensing, K. H. 2006. The cell biology of wood formation: from cambial divisions to mature secondary xylem. Can J Bot 84, 631-639. Sanchez-Fernandez, R., Davies TG, Coleman JO, Rea PA. (2001) The Arabidopsis thaliana ABC protein superfamily, a complete inventory. J Biol Chem 276, 30231-44. Sato, Y., T. Demura, K. Yamawaki, Y. Inoue, S. Sato, M. Sugiyama, and H. Fukuda (2006) Isolation and Characterization of a Novel Peroxidase Gene ZPO-C Whose Expression and Function are Closely Associated with Lignification during Tracheary Element Differentiation. Plant Cell Physiol 47, 493 - 503. Schrader, J., Moyle R, Bhalerao R, Hertzberg M, Lundeberg J, Nilsson P, Bhalerao RP. (2004) Cambial meristem dormancy in trees involves extensive remodelling of the transcriptome. Plant J 40, 173-187. Shitan, N., Bazin, Kazuyuki Dan, Kazuaki Obata, Koji Kigawa, Kazumitsu Ueda, Fumihiko Sato, Cyrille Forestier, and Kazufumi Yazak. (2003) Involvement of CjMDR1, a plant multidrug-resistance-type ATP-binding cassette protein, in alkaloid transport in Coptis japonica. Proc Natl Acad Sci USA 100, 751-756 Stacey, N. J., K Roberts NC Carpita, B Wells and MC McCann (1995) Dynamic changes in cell surface molecules are very early events in the differentiation of mesophyll cells from Zinnia elegans into tracheary elements Plant J 8, 891-906 Sussman, MR., Harper JF. 1989. Molecular biology of the plasma membrane of higher plants. Plant Cell 1, 953-960. Takabe, K.,Fujita, M.,Harada,H.,Saiki,H.(1981) Lignification process of Japanese black pine (Pinus thunbergii Parl.) tracheids. Mokuzai Gakkaishi 27, 813-820. Takabe, K, Fujita M, Harada H, Saiki H (1985) Autoradiographic investigations of lignification in the cell walls of cryptomeria (Cryptomeria japonica d. Don). Mokuzai Gakkaishi 31, 613-619 Takabe, K., Takeuchi M, Sato T, Ito M, Fujita M (2001) Immunocytochemical localization of enzymes involved in lignification of the cell wall. J Plant Res 114, 509-515 Takeuchi, M., Takabe K, Fujita M (2001) Immunolocalization of O-methyltransferase and peroxidase in differentiating xylem of poplar. Holzforschung 55, 146-150, Terashima, N., Fukushima, K., Takabe, K. (1986). Heterogeneity in formation of lignin. VIII. An autoradiographic study on the formation of guaiacyl and syringyl lignin in Magnolia  kobus DC. Holzforschung 40, 101-105. Turner, S., Gallois P, Brown D, (2007) Tracheary element differentiation. Annu Rev Plant Biol 58, 407-33. Vanholme, R., Morreel K, Ralph J, Boerjan W. (2008) Lignin engineering. Curr Opin Plant Biol 11, 278-285. Wardrop, A., and Harada,. H. (1965) The formation and structure of the cell wall in fibers and. tracheids. J Exp Bot 16, 350-356. Whetten, R., Sun YH, Zhang Y, Sederoff R.(2001) Functional genomics and cell wall biosynthesis in loblolly pine. Plant Mole Biol 47, 275-291. Wooding, FDP (1968) Radioautographic and chemical studies of incorporation into sycamore vascular tissue walls. J Cell Sci 3, 71-80 Yazaki, K., Transporters of secondary metabolites (2005) Curr Opin Plant Biol 8, 301–307. Zhao C., JohannaC.Craig,H.EarlPetzold,AllanW.Dickerman,andEricP.Beers. (2005) The Xylem and Phloem Transcriptomes from Secondary Tissues of the Arabidopsis Root-Hypocotyl. Plant Physiol 138, 803-818.  29 Zhong, R., Taylor, J.T., and Ye, Z.-H. (1997). Disruption of interfascicular fiber differentiation in an Arabidopsis mutant. Plant Cell  9, 2159-2170.  30              CHAPTER 2:  TRACKING MONOLIGNOLS DURING SECONDARY CELL WALL DEVELOPMENT IN PINUS CONTORTA VAR. LATIFOLIA AND POPULUS DELTOIDES X P. TRICHOCARPA1                   1. A version of a manuscript this has been published M. Kaneda, K.H. Rensing , J.C.T. Wong , B. Banno , S.D. Mansfield , and A.L. Samuels. Tracking Monolignols During Wood Development in Pinus contorta var. latifolia. Plant Physiol. August 2008, volume 147, pp. 1750-1760.  31 CHAPTER 2: TRACKING MONOLIGNOLS DURING SECONDARY CELL WALL DEVELOPMENT IN PINUS CONTORTA VAR. LATIFOLIA AND POPULUS DELTOIDES X P. TRICHOCARPA 1   2.1 Introduction This chapter presents a study of monolignol export during lignification of secondary cell walls in pine and poplar during xylem development. This introduction describes the rationale for the study as well as the preliminary work done in the Samuels lab on this project. The results detail my work on the chemical analysis and autoradiography to localize phenylpropanoids during xylem development. The processes of secondary xylem development, from cambial divisions to maturation of tracheary elements and fibers by programmed cell death, have been defined using ultrastructural (Samuels et al., 2002), molecular (Kirst et al., 2003; Allona et al., 1998), and biochemical approaches (Savidge, 1989).  The most prominent feature of xylogenesis is the development of the lignified secondary cell wall that takes place in two sequential steps: polysaccharide synthesis followed by lignification. Lignification results from the dehydrogenative polymerization of monolignols, which are synthesized from phenylalanine via the phenylpropanoid pathway.  Detailed characterization of properties of the phenylpropanoid biosynthetic enzymes has led to revisions of the proposed pathway for monolignol biosynthesis (Humphreys and Chapple, 2002; Goujon et al., 2003; Boerjan et al., 2003).  In gymnosperms, the major monolignol, coniferyl alcohol, accumulates in its glucosylated form as coniferin and its concentration in the inner bark has been positively correlated with active growth (Savidge, 1988).  The abundance of coniferin led to the hypothesis that first, coniferin is exported during xylem development, and then coniferin  32 -glucosidases in the secondary cell wall cleave the glucose, generating coniferyl alcohol for lignification in the apoplast (Freudenberg, 1959).  In angiosperms, the monolignol glucosides have yet to be shown to accumulate and sinapyl alcohol is produced in addition to coniferyl and p-coumaryl alcohols for export and incorporation into lignin.  However, the mechanism and form of monolignols being exported from their site of synthesis in the cytoplasm to their site of polymerization in the apoplast is still poorly understood (Whetten et al., 1998; Whetten and Sederoff, 1995).  During secondary cell wall deposition in lodgepole pine [Pinus contorta var. latifolia], the protoplasts of the developing tracheids have prominent Golgi with grape-like clusters of vesicles being shed from the trans-Golgi network (Samuels et al., 2002).  The predominant hemicelluloses of Pinus secondary cell wall, galactoglucomannans, have been localized to these vesicles using enzyme-gold probes (glucomannan-specific mannanase linked to colloidal gold) (Samuels et al., 2002).  This work led to the following question: do these abundant Golgi vesicles also carry monolignols to the developing secondary cell wall? 2.1.1 Preliminary data from Samuels lab The objective of this study was to localize monolignols in intact developing tracheids of pine, as well as vessels and fibers of poplar, during lignification using light and transmission electron microscopy (TEM).  In order to do this, I relied on preliminary data on these systems that are summarized below. Since antibodies specific to lignin detect only polymeric forms with varying degrees of condensation (Muesel et al., 1997), they are unable to detect monolignols prior to polymerization.  Therefore, we employed a novel autoradiography approach to localize monolignols.  Autoradiography has been used in the past to localize soluble radioactive  33 phenylpropanoids and it has been reported that monolignols are secreted via Golgi-mediated vesicle fusion with the plasma membrane (Wooding, 1968; Pickett-Heaps, 1968).  In these studies, following the uptake of radiolabeled precursors, developing xylem samples for autoradiography were prepared for light and transmission electron microscopy using chemical fixatives.  The slow penetration of chemical fixatives during this process has been shown to be particularly detrimental to developing wood cells (Rensing, 2002) and, during fixation, solute migration and membrane vesiculation can occur (Gilkey and Staehelin, 1986).  In contrast, cryofixation, such as high-pressure freezing, fixes cellular constituents within milliseconds, and consequently the cell contents remain immobile during freeze-substitution at –80oC, while frozen cellular water is substituted with solvent such as acetone prior to embedding in plastic resin for sectioning (Kiss and Staehelin, 1995). Prior to cryofixation, 3H-phenylalanine was fed to dissected lodgepole pine (Pinus contorta var. latifolia) or hybrid poplar (Populus deltoides x P. trichocarpa) cambium and developing secondary xylem, and then localization of the incorporated phenylpropanoids was established by autoradiography. This technique was developed by a post-doctoral fellow, Dr. Kim Rensing, in the Samuels lab using the pine samples.  Dr. Rensing first used light microscopy to establish a general overview of uptake and incorporation of radiolabeled phenylpropanoids in developing secondary xylem as dark deposits in the tracheids, rays and phloem of cryofixed samples (Fig. 2.1).  In the control samples when only 3H-phenylalanine was in the incubation solution, the radioactivity associated with developing secondary xylem was concentrated in the secondary cell walls of tracheids including the bordered pits.  It is difficult to distinguish the secondary cell wall from the cytoplasm using a light microscope, because the developing tracheids have such large central vacuoles that only a thin layer of  34 cytoplasm is pressed against the longitudinal walls.  When the plane of section passed through this cortical cytoplasm and secondary cell wall, the label pattern was most intense (bracketed arrows, Fig. 2.1A).  In the cytoplasm-rich rays, radioactivity was found throughout the cytoplasm, including the nucleus.  Radioactivity was relatively low in the vacuoles of all cell types.  35  Figure 2.1 Light autoradiographs of pine stem sections through the cambium  and secondary xylem following incorporation of 3 H-phenylalanine. In each panel, the cambium ( c ) and the region of developing xylem (x) are shown, with areas where cells are lignifying bracketed by arrows.  The black areas are the result of silver grain deposition over areas of radioactivity. A: Controls treated with 3H-phenylalanine alone have heavy incorporation. B: Addition of cycloheximide (CH), an inhibitor of protein synthesis, diminished 3H-phenylalanine labeling in rays. C: Inhibitors of the phenylpropanoid pathway such as C4H inhibitor piperonylic acid (PA) had the opposite effect: Labeling over the developing xylem was strongly reduced while that in the rays and cortical parenchyma remained.  D: Inhibition of phenylpropanoid metabolism with the PAL inhibitor, L-a- aminooxy-ß-phenylpropionic acid (AOPP). Data from Kaneda et al. (2008), collected by Kim Rensing.  36 To evaluate the extent of 3H-phenylalanine incorporation resulting from protein synthesis, developing wood samples were incubated with both 3H-phenylalanine and cycloheximide to block protein translation (Fig. 2.1B).  Cycloheximide treatment decreased the radiolabel incorporated into rays but developing tracheids were still labeled, indicating that 3H- phenylalanine incorporation in these cells represents metabolic pathways other than translation, i.e. phenylpropanoid metabolism.  In complementary experiments, phenylpropanoid biosynthesis was decreased either by inhibition of cinnamate-4-hydroxylase (C4H) using piperonylic acid (PA) (Chong et al., 2001) or by inhibition of phenylalanine ammonia-lyase (PAL) using L--aminooxy-ß-phenylpropionic acid (AOPP) (Appert et al., 2003).  When inhibitors were applied with 3H-phenylalanine, the label in the developing tracheids’ secondary cell wall was diminished (Fig. 2.1C, D).  The ray cells were still labeled, but the intensity was lower.  This effect was difficult to judge because there was strong variability in label density between sections, even within one treatment. Therefore, a protocol was developed (by NSERC undergraduate student John C.T. Wong) to quantify the incorporated radioactivity by measuring grey levels on light microscope sections. Levels were corrected to the mean white of the background, to get average density values in defined cell types (secondary cell wall/cortical cytoplasm of developing tracheids; cytoplasm of rays) under different inhibitory conditions (Fig. 2.2).  In developing tracheids, inhibition of C4H by 10 μM PA decreased the average label to 19% of the control density, suggesting a large component of 3H-phenylalanine incorporation into these cells was indeed phenylpropanoid in nature.  Since treatment with 100 μM PA led to changes in Golgi ultrastructure (Kaneda et al., 2008), an additional enzyme in the phenylpropanoid pathway, PAL, was inhibited with either 50 μM AIP or AOPP (Appert et al., 2003).  As with PA  37 inhibition of C4H, treatment with AOPP or AIP led to decreases in labeling of tracheid secondary cell wall.  Label levels from samples treated with either AOPP or AIP were not statistically different from each other, so all PAL inhibitors data were pooled.  As with PA inhibition of C4H, treatment with PAL inhibitors led to decreases in labeling of tracheid secondary cell walls to 31% of the control levels (Fig. 2.2).  In comparison, after treatment with CH, tracheid walls were still labeled at 75% of control levels, suggesting that inhibition of protein biosynthesis had little effect on the lignification of tracheids.  The parenchymous rays of conifers remain active in phenylpropanoid metabolism after tracheids finish their maturation (Davin and Lewis, 2000), and a role for rays in contributing monolignols for tracheid lignification can be envisioned.  When inhibitors of phenylpropanoid metabolism were applied with 3H-phenylalanine, the radioactivity incorporated into the rays was decreased.  For AOPP inhibition of PAL, the levels were decreased to 36% of controls, while for PA inhibition of C4H, the levels were decreased to 60% of controls, suggesting that phenylpropanoid metabolism is active in these cells.  Cycloheximide inhibition of protein synthesis, which did not stop lignification of tracheid cell walls, strongly decreased incorporation of radiolabeled phenylpropanoids into rays to 16% of control levels.  38     Figure 2.2 Quantification of grey levels in light microscopy autoradiographs of pine developing secondary xylem. Dissected cambium and associated tissue samples were fed 3H-phenylalanine and various inhibitors for 4 hours.  To inhibit the phenylpropanoid pathway, 2-aminoindan-2-phosphonic acid (AIP) and L-  -aminooxy-ß-phenylpropionic acid (AOPP) were used as inhibitors of phenylalanine ammonia lyase (PAL) and piperonylic acid (PA) as an inhibitor of cinnamate-4- hydroxylase.  Cycloheximide (CH) was used to block translation and inhibit incorporation of phenylalanine into proteins. Grey levels, where 0 is black and 1024 is white, were inverted and scaled to mean white background to give positive mean+SE values.  Data from Kaneda et al. (2008), collected by John C.T. Wong.   39 In addition, Dr. Rensing developed a protocol to employ the cryofixed samples for TEM so the subcellular distribution of phenylpropanoid derivatives could be determined (Fig. 2.3). When radiolabeled developing tracheid cells were examined with TEM, the most heavily labeled area was the developing secondary cell wall with relatively low label inside the protoplasts (Fig. 2.3).  Both 3H-phenylalanine alone (Fig. 2.3A) and 3H-phenylalanine plus CH treatments (Fig. 2.3B, C) produced similar patterns of heavy label deposition in the secondary cell wall while PA treatment lead to decreased secondary wall label (Fig. 2.3D). Increasing the time of incubation (4 hours, 6 hours, or overnight) or exposure time of sections to the emulsion did not increase the amount of radioactivity observed in the protoplast, only in the secondary cell wall. This suggests that the flux of phenylpropanoids through these cells to secondary cell wall is very rapid and incorporated phenylpropanoids do not pool inside the tracheid cells.  40   Figure 2.3 TEM autoradiographs of pine tracheids treated with 3 H-phenylalanine and either an inhibitor of protein synthesis, cycloheximide (CH), or an inhibitor of phenylpropanoid metabolism, piperonylic acid (PA). Silver grains indicating sites of incorporation of radioactive phenylpropanoid are most abundant over the secondary cell wall (CW) and are also associated with Golgi (G) in control (A) and PA treated tissue (D) but not in CH treated tissue (B and C).  Label over vacuole (V) is relatively low. Data from Kaneda et al. (2008), collected by Kim Rensing.  41  Dr. Rensing’s work established that this technique could be used to detect individual radioactive decay events in the cells but there were two outstanding questions remaining: 1. What is the chemical nature of the compounds into which the 3H-phenylalanine was incorporated? 2. Are the radioactive decay events in the cells more likely to localize in the Golgi vesicles or the cytoplasm?  Further, can the incorporation of 3H-phenylalanine into protein vs. phenylpropanoid in the Golgi signal be distinguished?  To verify that 3H-phenylalanine was incorporated into the phenylpropanoid pathway, I performed chemical analyses to test 3H-phenylalanine incorporation into lignin as well as methanol soluble precursors in both pine and poplar samples. To test if monolignols occur in the Golgi, I developed a mechanism to quantify the decays in TEM autoradiography of 3H-phenylpropanoid incorporation under conditions involving inhibition of protein synthesis or phenylpropanoid metabolism.   42 2.2 Results 2.2.1 3 H-phenylalanine is incorporated into the lignin precursors and polymers. In order to interpret autoradiography results showing where 3H-phenylalanine and its derivatives are found in developing wood after feeding, it was necessary to determine the biochemical fate of the 3H-phenylalanine, i.e. did 3H-phenylalanine remain as unincorporated amino acid or did it enter the phenylpropanoid pathway and ultimately get incorporated into lignin?  Dissected pine cambium and associated developing xylem were incubated for 4 hours in media containing 3H-phenylalanine, which were also the conditions used for cryofixation and microscopy.  Samples were then processed to detect radiolabel in both polymerized lignin and in the soluble components. Following depolymerization and solubilization of lignin by thioacidiolysis, lignin polymer breakdown products were consistently shown to be radioactive, indicating incorporation of 3H-phenylalanine into the lignin polymer (Fig. 2.4).  43   Thioacidolysis experiment 1 7230 cpm Thioacidolysis experiment 2 13124 cpm Thioacidolysis experiment 3 19783 cpm Thioacidolysis experiment 4 6772 cpm Thioacidolysis experiment 5 8121 cpm Thioacidolysis experiment 6 7201 cpm  Figure 2.4 3 H-phenylalanine incorporation into lignin demonstrated by radioactive thioacidolysis products. Under similar conditions to those used for autoradiography, 220-440 mg tissue samples was incubated in 3H-phenylalanine solution for 4 hours, followed by thioacidolysis as described in Rolando and Lapierre (1992). Thioacidolysis breakdown products (100 l) were added to 3 ml scintillation fluid.   44 To investigate the nature of the soluble metabolites that were labeled during the four hour incubation with 3H-phenylalanine, methanol soluble phenolic compounds were separated and identified using high performance liquid chromatography (HPLC) and mass spectroscopy (LC- MS).  In pine samples, coniferin, coniferyl alcohol and p-coumaryl alcohol peaks were prominent, as detected with absorption of ultraviolet light at 280 nm (Fig. 2.5A).  Identification of the peaks was first by co-elution with standard compounds and then confirmed by LC-MS. Minor peaks included the coniferyl alcohol dimer, pinoresinol, and phenylalanine. To test which of these peaks were radioactive, fractions were collected every 30 seconds for 75 min and tested for radioactivity using scintillation counting. Phenylalanine represented only a small fraction of the total soluble radiolabel, suggesting that most of the 3H-phenylalanine was metabolized (Fig. 2.5B). The expected monolignols, p-coumaryl alcohol and coniferyl alcohol, had retention times of 7.8 and 8.7 min.  All fractions during these time periods were radiolabeled (Fig. 2.5B).  Coniferin, the glucoside of coniferyl alcohol, although abundantly detected on the HPLC, was not radioactive after the four hour incubation period.  There was, however, a strong radioactive signal correlated with pinoresinol.  In addition, there was a large radioactive peak that eluted at 11 min.  There were three peaks detected by UV absorption from 10-11 min that gave complex MS fragmentation patterns, including coniferyl alcohol and p- coumaryl alcohol fragments. These peaks did not co-elute with the 8-O-4 coniferyl alcohol dimer (elution time 12.0 min), the 8-5 coniferyl alcohol dimer (elution time 12.4 min), or the 8- 5 dimer of p-coumaryl alcohol or p-coumarylresinol. Overall, this data indicates that treatment of pine developing secondary xylem with 3H-phenylalanine leads to incorporation of radioactivity into lignin, monolignols, lignans, as well as additional compounds.  45  Figure 2.5 Analysis of radioactive compounds in methanolic extracts of dissected cambium and associated developing secondary xylem from P. contorta following 4 h incubation in 3 H-phenylalanine. A) HPLC chromatogram shows separation of phenolic compounds, detected by their ultraviolet absorption at 280 nm and identified by co-elution with standards and LC-MS.  B) Radioactivity from each HPLC fraction above was collected every 30 sec and the amount of radiolabel in the fraction measured by scintillation counting.  Peaks correspond to the monolignols, p-coumaryl alcohol and coniferyl alcohol, at 8 min elution; pinoresinol at 14.5 min; and a large peak at 11 min corresponding to three peaks at 10-11 min that gave complex MS fragmentation patterns, including coniferyl alcohol and p-coumaryl alcohol fragments.   46 In parallel study work of poplar, a similar protocol of 4 hours of feeding 3H-phenylalanine to dissected cambium and associated derivative tissues was used. To analyze the metabolites that were radiolabeled by 3H-phenylalanine incubation, methanolic extracts were separated by HPLC.  More than 20 peaks with absorption of ultraviolet light at 280 nm were detected (Fig. 2.6). Compared to pine xylem methanolic extracts, more major peaks of phenolic compounds were separated from the poplar methanolic extract. Based on standard compounds elution time, p-coumaryl alcohol (retention time of 7.8 min), sinapyl alcohol (retention time of 8.4 min), 8-5 dimer of p-coumaryl alcohol (retention time of 12.2 min), p-coumarylresinol (retention time of 13.3 min), syringaresinol (retention time of 14 min) and pinoresinol (retention time of 14.8 min) were present in poplar methanol extracts (Fig. 2.6A). At longer elution times, several peaks were observed that probably correspond to the oligolignols characterized by Morreel et al., (2004) in poplar methanolic extracts. Their identity is under further investigation (S.D. Mansfield et al., unpublished data).                      47         Figure 2.6 Analysis of radioactive compounds in methanolic extracts of dissected cambium and associated developing secondary xylem from hybrid poplar (H11-11) following 4 h incubation in 3 H-phenylalanine. A) HPLC chromatogram shows separation of phenolic compounds, detected by their ultraviolet absorption at 280 nm and identified by co-elution with standards and LC-MS.  B) Radioactivity from each HPLC fraction above was collected every 30 sec and the amount of radiolabel in the fraction measured by scintillation counting.   48 To test if any phenolic compounds from poplar wood were radioactive, fractions were collected every 30 seconds for 75 min and tested for radioactivity levels by scintillation counting. There were four very high radioactive fractions (9 min, 11.5 min, 13 min and 15.5 min) seen (Fig.2.6B).  Radiolabel was only slightly above background in the 2-3 minute fractions where unincorporated 3H-phenylalanine would elute, suggesting free 3H-phenylalanine was again incorporated.  All fractions between 6 min to 21 min corresponding to phenolic peaks were relatively highly radiolabeled, suggesting that 3H-phenylalanine incorporated into diverse phenolic compounds including signals correlated with monolignols/dilignols in the poplar methanolic extract. Knowing that the 3H-phenylalanine was entering the monolignol biosynthetic pathway, we could then use cryofixation to immobilize these metabolites in situ and visualize them using light and electron microscopy. 2.2.2 3 H-phenylalanine was intensively incorporated into developing xylem as lignin and its precursors.  To compare the distribution of 3H-phenylalanine derivatives in developing hybrid poplar with the previous results from pine, poplar dissected cambium and associated developing xylem were incubated in 3H-phenylalanine for autoradiography. Light microscopy autoradiography of radiolabeled phenylpropanoids in developing secondary xylem showed dark deposits in the phloem, developing xylem and ray parenchyma cells (Fig. 2.7). To visualize the anatomy of the cambial region and developing xylem, toluidine blue stained light microscope sections are shown (Fig. 2.7A, C).  Incorporation of 3H- phenylalanine into the tissue is shown in adjacent panels (Fig. 2.7B, D).  Strong incorporation of label into developing xylem as well as rays and phloem was observed (Fig. 2.7B).  In  49 contrast, label incorporated into mature xylem vessel or fiber secondary cell walls was very low indicating that little polymerization of lignin occurs in mature xylem.  Cytoplasm of living ray parenchyma cells in the mature xylem was labeled, indicating these cells continuing metabolic activity even after programmed cell death of many of the surrounding axial cells.  Incorporation of radioactivity was relatively low in the vacuoles of all cell types (Fig. 2.7B, D).  Comparing Figs. 2.7B and D, the variation between samples is quite apparent, therefore the grey levels of multiple samples were measured. In the controls, the strong label in developing xylem, rays and phloem, compared to cambium and mature xylem, was clear (Fig. 2.8). To distinguish protein synthesis from lignin precursor distribution, CH was used as a protein synthesis inhibitor or piperonylic acid (PA) was used to inhibit the phenylpropanoid pathway (Chong et al., 2001). In the developing xylem, inhibition by CH decreased the average to 67% of control levels, and inhibition of PA decreased incorporation of label to 22% of control level (Fig. 2.8), suggesting that while there was some coexistence of protein and lignin biosynthetic activity in these cells, most activity was due to lignin biosynthesis.  In contrast, CH treatment decreased the radiolabel incorporated into phloem and ray by 65% and 50%, while PA treatment did not change the extent of radiolabel incorporation in ray and phloem (Fig. 2.8), indicating that these incorporated compounds were primarily protein, not lignin precursors.  As with pine, inhibition of protein synthesis could potentially inhibit translation of monolignol biosynthetic enzymes in developing xylem, but, following CH treatment, radioactive label was still strongly incorporated into the secondary cell walls of the fibers and vessels, suggesting that monolignol production was not impaired.  50  Figure 2.7 Light microscopy autoradiography of poplar developing xylem tissue. A) and C) Toluidine blue stained light microscope section with primary walls stained dark purple and lignified walls stained bright blue. B) and D) LM autoradiographs showing deposition of phenylpropanoid in secondary cell walls of poplar.  51  Figure 2.8 Quantification of grey levels in light microscopy autoradiographs of poplar secondary growth tissue. Dissected cambium and associated tissue samples in poplar wood were fed 3H-phenylalanine and various inhibitors for 4 hours.  To inhibit the phenylpropanoid synthesis, piperonylic acid (PA) as an inhibitor of cinnamate-4-hydroxylase (C4H), was employed.  Cycloheximide (CH) was used to block translation and inhibit incorporation of phenylalanine into proteins. Grey levels, where 0 is black and 1024 is white, were scaled to mean white background and inverted that 1 is black and 0 is white, to give positive mean+SE values.  52  At the subcellular level, electron microscopy was used to observe the distribution of incorporated 3H-phenylalanine derivatives in developing poplar cells. In the developing xylem tissue, decay events were preferentially localized to the secondary cell wall, in the middle lamella and the corners of adjacent cells (Fig. 2.9). Whereas electron micrographs showed high decay events in the secondary cell wall, relatively lower signals were localized in the cytoplasm (Fig. 2.9).  In addition, there was no signal in the vacuole, indicating that at least under these experimental conditions, vacuoles did not accumulate phenylpropanoids during lignification. There was, however, a fairly low density of decay events in cytoplasm as well as Golgi in the protoplasts of the poplar vessels (Fig. 2.9B) and fibers (Fig. 2.9C).  53   Figure 2.9 TEM autoradiographs of poplar xylem tissue treated with 3 H- phenylalanine. Decay events indicating sites of incorporation of radioactive phenypropanoid were abundant in developing xylem.  A) At low magnification, high density of decay events was seen in secondary cell wall. B) A developing fiber with strong secondary cell wall label, as well as low cytosolic label. The vacuole (V) labels were relatively less than cytoplasm and cell wall (CW). C) High magnification of a fiber with decay events associated with Golgi (G) and cytoplasm. White bar, 1 m.   54 These experiments provide verification that 3H-phenylalanine is incorporated into lignin and monolignols in the samples under these experimental conditions and that the decay events could be visualized.  However, due to the low numbers of decay events in the cytoplasm and uncertainty about the radioactive source geometry, these experiments were not able to pinpoint where, in the lignifying cells, monolignols are found, i.e. if they are localized in Golgi. For these reasons, subsequent experiments were undertaken on the subcellular distribution of monolignols in lignifying cells using TEM quantification techniques. Getting large numbers of TEM autoradiography samples with good contrast and resolution was challenging.  Due to the homogeneity of the pine tracheids, compared to the diverse cell types of poplar, pine was used for subsequent quantification. 2.2.3  Quantitative decay event analysis showed localization of lignin precursors/lignin in the cytoplasm and in secondary cell wall. Radiolabeling was observed in the protoplast of developing pine tracheids, in the cytoplasm, Golgi, and vacuole in both 3H-phenylalanine alone (Fig. 2.3A, Fig. 2.10) and CH or PA treated samples (Fig. 2.3B,C).  The radioactive decay pathway into the autoradiographic emulsion is not necessarily perpendicular to the plane of section, but can radiate out from a point source in all directions. Thus, the source-detector geometry means that the reduced silver grains in the emulsion are located in a pattern decreasing exponentially from the radioactive source.  For 3H detection by Ilford L4 emulsion, the half-distance was determined to be 145 nm, e.g. 50% of the radioactivity radiating from a point source was detected experimentally within a circle of this radius (Salpeter and Bachmann, 1972).  By counting a ‘probability circle’ of a 500 nm circle around a silver grain, we estimated that 90% of decay events exposing that grain would come from within that circle.  Therefore, all organelles within the circle were scored as  55 potential sources (Fig. 2.10).  The area of each organelle was outlined as illustrated in Fig. 2.10 and where the probability circle covered a portion of this area, a percentage of the probability circle was attributed to that organelle.  For example, for a decay event where one third of the probability circle covers the Golgi area, 0.33 of a probability circle/decay event would be attributed to Golgi.  The sum of decay events divided by total organelle area measured was used to establish how much radioactivity can be attributed to that organelle.  56  Figure 2.10 TEM autoradiography quantification example (control, 3 H-phenylalanine only) in developing tracheids of P. contorta. The density of the decay events was measured by drawing probability circles, with radius of 500 nm, which would capture 90% of possible sources around a decay event.  The area of the underlying organelle that intersected the probability circle was calculated.  Bar=2 m.  CW, cell wall; G, Golgi; V, vacuole.   57  Quantification of decay events detected in TEM thin sections showed that the secondary cell wall had the highest number of decay events/m2 (Fig. 2.11).  As with the coarser light microscopy measurements, cell wall deposition was strongly inhibited with PA but not by CH (Fig. 2.11).  This is important because it demonstrates that during the incubation period, CH treatment did not lead to inhibition of phenylpropanoid metabolism due to decreased biosynthetic enzyme production.  In the cytoplasm of control samples treated with 3H- phenylalanine alone, the majority of label was found in the cytoplasmic compartment, which would include the endoplasmic reticulum (ER) in our samples, as well as in the Golgi and the vacuole (Fig. 2.11).  By comparing control with samples in which phenylpropanoid metabolism or protein incorporation were inhibited, label in the Golgi could be attributed to either primarily phenylpropanoid or primarily protein synthesis. Inhibition of protein synthesis with CH decreased the Golgi label, while inhibition of phenylpropanoid metabolism with PA did not (Fig. 2.11).  This implies that the signal in the Golgi can be attributed to protein rather than phenylpropanoids and makes Golgi-mediated exocytosis an unlikely route for export of monolignols. Since the vacuoles were not strongly labeled and the cytoplasm of the developing xylem had low label levels, I was concerned that soluble metabolites were being extracted during processing (i.e. by the freeze substitution medium and embedding resin).  The substitution medium and resins were found to contain measurable, but low, radioactivity (Kaneda et al. 2008).  In particular, the extraction, as assessed by scintillation counts, increased with concentration of epoxy resin during embedding.  When utilized low temperature embedding at –70° C with Lowicryl HM20 to reduce the extent of extraction, scintillation counts showed no measurable extraction by the resin.  In low temperature embedded samples, preservation of ER  58 was improved but abundant labeling was still not observed in the cytoplasm of developing xylem.  Consistent with the earlier experiments, the radioactivity was found throughout the cytoplasm, with lower label in the Golgi and vacuoles (Kaneda et al., 2008).   59  Figure 2.11 Quantification of radioactive label in developing tracheids of P. contorta following cryofixation, autoradiography and TEM. Mean+standard error indicates the density of decay events associated with A) radiolabel in the lignifying secondary cell wall of tracheids and B) radiolabel associated with intracellular structures of tracheids.  Total decays divided by the summed area for each contributing organelle gave a density metric of where radioactivity was located.  Inhibition of protein synthesis with cycloheximide (CH) did not alter lignin deposition in the cell wall but produced significantly decreased incorporation of radioactivity in the Golgi (Mann-Whitney test, p<0.05).  In contrast, inhibition of phenylpropanoid metabolism by C4H inhibitor, piperonylic acid (PA) significantly decreased label in cell wall but Golgi label was not different from control.  60 2.3 Discussion The combination of autoradiography and cryofixation has allowed us to re-examine the question of how monolignols are exported during tracheid lignification in pine.  In diverse species, phenylpropanoid biosynthetic enzymes, such as PAL have been localized in the cytosol (Smith et al., 1994; Takabe et al., 2001) while cytochrome P450 enzymes such as C3H and C4H are predicted to be targeted to the endoplasmic reticulum (Emanuelsson et al., 2000). The location of these enzymes suggests that monolignols are synthesized in the cytosolic compartment and, to be transported to the apoplast, must cross the plasma membrane.  In a Golgi-vesicle model of transport, monolignols would have to accumulate in the endomembrane system, either at the ER or the Golgi.  Such accumulation was not observed in this study where 3H-phenylalanine incorporated into the Golgi of lignifying tracheids was due to protein synthesis, rather than phenylpropanoid metabolism.  This result does not support the Golgi- mediated export of monolignol hypothesis and suggests that other mechanism(s) are used by the plant to export monolignols into the secondary cell wall. If monolignols were carried within the endomembrane system, they would move from the cell with kinetics determined by the rate of vesicle budding and fusion.  The time of transit of a cargo protein from synthesis on the rough ER to secretion at the plasma membrane in classical autoradiographic studies of mammalian cell protein secretion is around 120 minutes (Palade, 1975), a figure that has been supported more recently in live mammalian cell studies using green fluorescent protein (GFP) (Hirschberg et al., 1998). Assuming phenylpropanoid- containing vesicles moved with the same kinetics, they would be easily ‘trapped’ using cryofixation during their transit.  While abundant Golgi and Golgi vesicles clusters were  61 observed in this and earlier (Samuels et al., 2002) studies, they did not appear to be loaded with phenylpropanoids. My results are consistent with an early study of autoradiography in sycamore wood where 3H-glucose labeled the Golgi of the developing xylem, presumably due to polysaccharide biosynthesis, but 3H-phenylalanine treatment gave sparse, random distribution of label over the cytoplasm and strong label of the secondary cell wall (Wooding, 1968).  A review of the literature supporting the Golgi-vesicle model of monolignol export indicates that there is no experimental evidence that directly contradicts our findings.  The classic autoradiography study by Pickett-Heaps (1968) showed incorporation of 3H-cinnamic acid into developing primary xylem of wheat coleoptiles, with label often occurring at the cell periphery where “Golgi-derived” vesicles were fusing with the cell wall “…particularly when the plane of section approached the wall thickenings tangentially”.  Other autoradiography studies that employed 3H-phenylalanine to label developing xylem found the label associated with rough endoplasmic reticulum (ER), Golgi and vesicles fusing with the plasma membrane (Pickett- Heaps, 1968; Takabe et al., 1985).  In these studies, vesicles close to or fusing with the plasma membrane were assumed to be Golgi vesicles, which is a fair assumption given the prominent Golgi structures in developing xylem cells with high polysaccharide production.  Unlike cryo- fixed cells, where the plasma membrane retained a smooth profile typical of cells under turgor, cells that are chemically fixed show wavy, vesiculated membrane profiles.  If the region of cytoplasm adjacent to the plasma membrane contained label, and if it were vesiculated, it is easy to see why previous studies concluded that  “Golgi” vesicles were labeled as they fused with the plasma membrane. In addition, these studies lacked controls for protein incorporation  62 and our current data show that phenylalanine treatment results in radiolabel in the Golgi that is more likely to be protein than monolignol in nature. The demonstration that 3H-phenylalanine fed to dissected developing xylem is incorporated into lignin is not surprising.  Similar results were found in earlier studies where 3H-phenylalanine or 3H-cinnamic acid, fed to developing xylem, were recovered as lignin constituents following alkaline nitrobenzene oxidation (Pickett-Heaps, 1968; Wooding, 1968). This pattern of lignification was described previously using fluorescent detection of the lignin polymer (Terashima and Fukushima 1988, Donaldson et al. 1991).  Given that radioactivity was found in the thioacidolysis breakdown products and in the coniferyl and p-coumaryl alcohol fractions, the simplest interpretation is that both monomeric lignin precursors and lignin polymer were radioactively labeled during the experimental incubation period. What is more surprising is that no radiolabel was shown to be associated with coniferin in the pine samples.  However, this result is consistent with the low phenylpropanoid label found inside the central vacuole where coniferin would be expected to be stored.  Using HPLC, it was clear that coniferin was present in our samples and qualitatively the levels were consistent with reports of coniferin levels present in gymnosperms (Marcinowski and Grisebach, 1977; Savidge, 1988; Leinhos and Savidge, 1994).  Historically, feeding experiments using radiolabeled coniferin showed incorporation of radiolabel into lignin and introducing radiolabeled phenylalanine into spruce seedlings led to labeled coniferin being detected in “one to two days” (Freudenberg, 1959).  Pulse-chase experiments showed that when spruce seedlings were fed a pulse of radiolabeled phenylalanine from 0-22 hours, coniferin levels peaked during the chase period at about 100 hours (Marcinowski and Grisebach, 1977). Using microtome sections of Pinus thunbergii, Fukushima and coworkers (1997) determined  63 that the spatial and temporal distributions of coniferin were highest early in the xylem differentiation and early in the growing season.  The relatively slow turnover of coniferin and the discrepancy between coniferin levels and the amount of lignin deposited led to the suggestion that the direct pathway from coniferyl alcohol to lignin makes a larger contribution than flow via coniferin (Fukushima et al., 1997).  It is possible that distinct spatially and temporally regulated pools of coniferin exist: one that turns over rapidly and is directly associated with metabolite channeling towards lignification, and a slowly exchanging storage pool of coniferin, associated with other phenylpropanoid metabolism such as pinoresinol production in the rays.  If coniferin is not the form of lignin precursor transported during tracheid development then the predicted transport substrate would be p-coumaryl alcohol in the early stages of lignification and coniferyl alcohol subsequently (Terashima and Fukushima, 1988). If Golgi-mediated export of monolignols is not operating, then alternative mechanisms are diffusion or transporter-mediated export.  Studies of model membranes using liposomes or lipid bilayer disks demonstrate partitioning of model phenolic substrates into the membrane (Boija and Johansson, 2006; Boija et al., 2007).  It is not clear how the desorption of these compounds from that hydrophobic environment into the apoplast would occur in this model. Therefore, a transporter model of monolignol export is attractive but the nature of this transporter remains elusive.  Such a transporter could be powered either by ATP hydrolysis directly, e.g. ATP binding cassette (ABC) transporters (Verrier et al., 2008) or indirectly via the proton gradient, as is the case for aromatic amino acid importers of the amino acid permease class (Yazaki, 2006).  ABC transporters have been proposed to play a role exporting monolignols across the plasma membrane into the developing secondary cell wall, but there is  64 no direct evidence for this model (Ehlting et al. 2005).  In several studies of gene expression during wood formation, expressed sequence tags (ESTs) for ABC transporters have been reported (Allona et al., 1998; Kirst et al. 2003; Hertzberg et al., 2001).  A microarray study of gene expression during lignification of the bolting Arabidopsis primary inflorescence stem also clearly identified several ABC transporter genes that were coordinately expressed with known phenylpropanoid metabolic genes (Ehlting et al., 2005), but inhibiting the most highly coordinately regulated ABC transporter genes produced only altered auxin transport and not lignin deficient phenotypes (Chapter 3).  In poplar, 3H-phenylalanine fed to dissected developing xylem was incorporated into the lignifying secondary cell walls, which is considered reasonable and proper based on the similar results found in pine xylem autoradiography. However, the number of phenolic compounds extracted from dissected xylem tissue showed high complexity of intermediates.  These results are not surprising because angiosperms contain specific enzymes for the extra lignin monomer: sinapyl alcohol.  A recent analysis of wild-type and caffeoyl-CoA O-methyltransferase (CCoAOMT) deficient poplar identified a similarly complex pattern of phenolics in the methanolic fraction including dimers and oligolignols from the three monomers (Morreel et al. 2004). Gymnosperm wood provides a simple system to study wood formation and lignification, since unlike angiosperms secondary xylem which contains multiple cell types, the conifer secondary xylem consists primarily of axial tracheids and parenchymous rays. Inhibition of radiolabel into rays of both pine and poplar by the protein translation inhibitor, cycloheximide, suggests that rays are metabolically active throughout tracheid differentiation and that proteins are actively turning over in ray cells.  During CH treatment, lignification in  65 the secondary cell wall in tracheary elements was not strongly inhibited, but there was decrease label in the ray cells.  It is hard to reconcile that result with the idea that the rays are major sources of monolignol.  It could be that either monolignols move through the ray cells very quickly or that the phenylpropanoids produced in the rays are not a major source of monolignols for lignification.  This would be surprising since it has often been suggested that neighboring cells could contribute to developing tracheary element lignification, as shown in the model Zinnia tracheary element culture system where neighboring cells contribute to lignification of tracheary elements (Hosokawa et al., 2001; McCann et al., 2001).  This paradigm is also based on reports of phenylpropanoid biosynthetic enzymes being produced in xylem parenchyma adjacent to lignifying cells as detected by GUS assay (Hauffe et al., 1991), and immunolocalization of phenylpropanoid enzymes (Samaj et al., 1998; Takabe et al., 2001). It was therefore also surprising to observe that, in contrast with the 50% inhibition of phenylpropanoid incorporation into the rays seen with PA treatment in pine, poplar rays inhibited with PA incorporated radiolabel at levels identical to controls.  It is possible that poplar rays are still active in monolignol metabolism, just downstream of the C4H block by PA, an idea that would invoke extensive exchange of metabolites between ray cell and lignified cells. In pine, the drop in phenylpropanoid metabolism in the rays could be due to enzymes involved in the production of diverse phenylpropanoids such as lignans.  The presence of radiolabeled pinoresinol, the coupling product of coniferyl alcohol, in the methanol soluble extractive would be consistent with lignan production in rays.  This production has been linked to both sapwood and heartwood formation and suggested to be the result of dirigent protein activity (Davin and Lewis, 2000).  66  2.4  Materials and methods 2.4.1  Plants material and growth condition Dormant Pinus contorta var. latifolia seedlings were transplanted, 3 per 15 cm square planters, and grown in a growth chamber at 24°C under 24 hour light.  When hand sections showed evidence of wood formation, typically after 7-10 days, seedlings were destructively sampled. To sample wood later in the growing season, seedlings were maintained in the growth chamber for up to three months.  Five-centimeter-long stem segments, cut from the seedlings 2 cm below the base of the terminal bud, were radially bisected. Approximately 0.5 mm-thick radial longitudinal slices containing portions of xylem, cambium and phloem were hand-cut from each half and immediately immersed in 0.2 M sucrose after which the cortex was removed.  In January, dormant 1-2 year old branches from a large poplar tree: Populus deltoides x P. trichocarpa hybrid (H11-11) at UBC south campus field were transplanted into 15 cm square pots, and grown in a greenhouse. Around April, the planted poplar branches broke dormancy and actively grew with many green leaves, which indicated that they were in the growing season.  This was confirmed by checking the cambium anatomy. Wood stems were radially dissected and sliced to approximately 0.3-0.5 mm thickness. These slices contains phloem, cambium and xylem tissues. 2.4.2  Autoradiography For autoradiography, the slices were transferred to 0.25 ml of 0.2 M sucrose containing 25 Ci of L-[2,6-3H]phenylalanine (1.8 M phenylalanine) in 2 ml Eppendorf tubes. Separate solutions containing labeled phenylalanine were prepared with various inhibitors. PA (Sigma) and CH (Sigma) were used at 10 M.  For inhibition of phenylalanine ammonia lyase, both 2-  67 aminoindan-2-phosphonic acid (AIP) and L--aminooxy-ß-phenylpropionic acid (AOPP) (gifts from Dr. N. Amrhein, Institute of Plant Sciences, Swiss Federal Institute of Technology, Zürich, Switzerland) were used at 50 M.  The tissues were incubated for the prescribed time at room temperature after which they were processed either for autoradiography by high-pressure freezing or for biochemical analysis.  Results from AOPP (4 h) and AIP (4 or 6 h) were not significantly different so results for all PAL inhibitors were pooled. 2.4.3  High pressure freezing (HPF) Slices were high-pressure-frozen in 0.2 M sucrose (as cryoprotectant) using a Bal-Tec HPM 010 (Bal-Tec AG, Balzers, Liechtenstein). For standard processing, frozen samples were freeze-substituted with 2% osmium tetroxide (Electron Microscopy Sciences) and 8% dimethoxypropane (Aldrich) in acetone for 120 hours, using a dry ice-acetone bath which equilibrated at -80°C.  The tissues were then warmed to –20°C in a freezer for 4 hours, and to 4°C in a refrigerator for 4 hours, after which time they were brought to room temperature. The tissues were transferred to HPLC-grade acetone and then Spurr’s resin was gradually added over 2 hours to bring the concentration to approximately 25%. The slices were then transferred to 50% resin for 2 hours, to 75% resin in open vials for 12 hours, and finally to 100% resin with changes two times per day, for 3 days.  The infiltrated samples were polymerized in fresh Spurr’s resin at 60°C. Low-temperature embedded samples were transferred from high-pressure freezing to uranyl acetate-saturated acetone in a Leica AFS (Leica Microsystems, Bannockburn IL, USA) pre-cooled to –80°C and allowed to substitute for 120 hours. They were subsequently warmed to –70° and gradually infiltrated with Lowicryl HM20. Samples in 100% Lowicryl were polymerized by UV light at –50°C and then allowed to warm to room temperature. Extraction of radiolabeled compounds during fixation and embedding was assessed by adding  68 10 ul from each mixture to 3 ml of Fisher ScintiVerse scintillation cocktail and counting the decays on a Beckman LS600IC liquid scintillation counter. 2.4.4  Light microscopy Sections 0.3 m were mounted on glass slides by heating.  All the blocks from each treatment were present on each slide to control for variability of emulsion thickness between slides and facilitate easier comparisons. One slide was stained with toluidine blue for structural reference. Slides for autoradiography were coated in Ilford L4 emulsion mixed with an equal amount of distilled water at 40°C under sodium safelight with closed filters (Thomas Duplex Super Safelight). The slides were dipped vertically into the emulsion and placed vertically on a paper towel to dry. The slides were stored in a black plastic slide box, wrapped in two photographic storage bags and stored at 4° C for 2 weeks. Emulsions were developed for 2 minutes in Kodak D19 developer diluted with 1 part distilled water, rinsed in distilled water then fixed for one minute in Ilford Multigrade Paper Fixer mixed with 4 parts water. They were then dried after a 20 minute rinse in cold water. The autoradiographs were observed with differential interference contrast using a Zeiss Axioplan light microscope (Carl Zeiss AG, Germany).  Images were captured using a Q-CAM digital camera (Q-Imaging, Burnaby, BC, Canada) and photographed unstained. To determine labeling intensity of different treatments, the grey levels of the tissues were quantified using Kohler illumination only (Openlab, Improvision). The darkness level, expressed as the inverse of the pixel value, was measured as an indication of labeling intensity after normalization against the mean background white level for each image. Deposits were measured as grey levels of a monochrome image, using unstained sections of cryofixed material for 977 observations from 5 independent experiments.   69 2.4.5  Electron microscopy 70 nm thick sections containing the labeled material were mounted on formvar-coated grids, stained for 30 min. with uranyl acetate and 10 min. with lead citrate and then carbon coated. The grids were observed and photographed prior to coating with emulsion for observation of cell structure at high resolution.  Grids were coated with Ilford L4 emulsion under sodium safelight with closed filters. Seven millimetre diameter wire loops were dipped in Ilford L4 emulsion at 40°C diluted with 2 parts water and the excess allowed to drain off. Loops were placed in horizontal position and allowed to dry only until the emulsion just started to get shiny. The grid was then placed onto film, sections side down and the emulsion was allowed to dry completely. Coated grids were exposed in a standard grid storage box inside 2 black photographic storage bags for 3 weeks at 4°C.  Emulsions were developed as above for light microscopy.  For quantification, the density of the decay events was measured by drawing probability circles, with radius of 500 nm, which would capture 90% of possible sources around a decay event (Salpeter and Bachmann, 1972).  The area of each organelle was outlined and where the probability circle covered a portion of this area, a percentage of the probability circle was attributed to that organelle.  The total area of the underlying organelles was calculated to express the radiolabel as decay events/m2.  Heavy deposits in the secondary cell wall were not quantified. 9-12 cells from 2-3 independent experiments were quantified. 2.4.6 Detection of radiolabel in tissues and solutions. After incorporation of 3H-phenylalanine, tissues were removed from the radiolabeling solutions, rinsed twice in 0.2 M sucrose and placed on several layers of filter paper to absorb excess apoplastic fluid.  Tissues were then frozen in liquid nitrogen and ground in a mortar and pestle. The ground tissue was extracted 3 times with 1 mL room temperature HPLC-grade  70 methanol. The methanol fractions were combined, centrifuged at 13000 rpm for 2 min, and supernatant collected and evaporated to dryness under a stream of nitrogen. To the methanolic fraction, an equal volume of ethyl acetate was added, and mixed thoroughly and allowed to phase partition.  The ether phase was then removed and retained, while the extract was again extracted with a second volume of ethyl acetate, and removed and pooled.   The ether phase was concentrated to dryness in a speedvac, and resuspended in 100 ul methanol and analysed by HPLC via a Summit HPLC system (Dionex) fit with a C18 Luna column (150  2.1 mm, 3um) (Phenomenex, Torrance, CA), an autosampler and a photodiode array detector was employed for all HPLC analysis.  Fractions were collected continuously every 0.5 min. For measurements of radioactivity, 100 ul from each fraction was mixed in 3 ml of Fisher ScintiVerse scintillation cocktail and decays counted on a Beckman LS600IC liquid scintillation counter.  LC/MS detection was achieved by injecting 10 ul onto a C18 Luna column (150  2.1 mm, 3 m) using a Waters 2695 Separations module (Waters, Milford, MA, USA).  Separation was performed with a mobile phase gradually changing from 83% solvent A [H2O:acetonitrile (ACN):formic acid (FA), (100:1:0.1, v/v/v), pH2.5] to 77% solvent B [ACN:H2O:FA, (100:1:0.1, v/v/v), pH2.5] within 21 minutes, at a flow rate of 0.3 ml/min and a column temperature of 40 ˚C. Detection was done using negative ionization on a Micromass Quattro Micro API triple quadrupole mass spectrometer with an APCI source (Micromass, Inc., Manchester, UK). The instrument was operated under the following conditions: source temperature, 130 ˚C; APCI Probe temperature, 500 ˚C; corona current, 5.0 μA; cone voltage, 25 V; extractor voltage, 5 V; radio frequency lens, 0.0V. Nitrogen from a nitrogen generator (Domnick Hunter, Ltd., Tyne and Wear, United Kingdom) was used as both the cone gas (50  71 l/h) and the desolvation gas (200 l/h). Quadrupole-1 parameters were as follows: low mass (LM) resolution, 14; high mass (HM) resolution, 14; ion energy, 0.5 V. Quadrupole-2 parameters were as follows: LM resolution, 14; HM resolution, 14; ion energy, 3.0. Collision cell entrance and exit potential were set at 50 V. Multipliers were set at 650 V. Scan time was 1 s and interscan delay 0.02 s. Data were acquired in continuous mode. Data acquisition and instrument control was performed using Masslynx 4.0.  2.4.7 Thioacidolysis Residual wood following methanolic extraction was subject to lignin degradation and solubilization by thioacidolysis (Rolando et al., 1992).  An aliquot solution 100 ul containing degradation products was sampled and mixed in 3 ml of Fisher ScintiVerse scintillation cocktail. Decays counted on a Beckman LS600IC liquid scintillation counter.  72  2.5 Bibliography Allona I, Quinn M, Shoop E, Swope K, Cyr SS, Carlis J, Riedl J, Retzel E, Campbell MM, Sederoff R, Whetten RW (1998) Analysis of xylem formation in pine by cDNA sequencing. Proc Natl Acad Sci USA 95, 9693-9698. Appert C, Zon J, Amrhein N (2003) Kinetic analysis of the inhibition of phenylalanine ammonia-lyase by 2-aminoindan-2-phosphonic acid and other phenylalanine analogues. Phytochem 62, 415-422. Boerjan W, Ralph J, Baucher M (2003) Lignin biosynthesis. Physiol Plant 54, 519-546. Boija E and Johansson G (2006) Interactions between model membranes and lignin-related compounds studied by immobilized liposome chromatography Biochim Biophys Acta 1758, 620-626. Boija E, Lundquist A, Edwards K, Johansson G (2007) Evaluation of bilayer disks as plant cell membrane models in partition studies Anal Biochem 364, 145-152. Chong J, Pierrel M-A, Atanassova R, Werck-Reichhart D, Fritig B, Saindrenan P (2001) Free and conjugated benzoic acid in tobacco plants and cell cultures. Induced accumulation upon elicitation of defense responses and role as salicylic acid precursors. Plant Physiol 125, 318-328. Davin LB, Lewis NG (2000) Dirigent proteins and dirigent sites explain the mystery of specificity of radical precursor coupling in lignan and lignin biosynthesis. Plant Physiol 123, 453–462. Donaldson LA. (1991) Seasonal changes in lignin distribution during tracheid development in Pinus radiata D. Don. Wood Sci Technol 25, 15-24. Ehlting J, Mattheus N, Aeschliman DS, Li E, Hamberger B, Cullis IF, Zhuang J, Kaneda M, Mansfield SD, Samuels L, Ritland K, Ellis BE, Bohlmann J, Douglas CJ. (2005) Global transcript profiling of primary stems from Arabidopsis thaliana identifies candidate genes for missing links in lignin biosynthesis and transcriptional regulators of fiber differentiation. Plant J 42, 618-640. Emanuelsson O, Nielsen H, Brunak S, von Heijne G (2000) Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J Mol Biol 300, 1005– 1016. Fukushima K, Taguuchi S, Matsui N, Yasuda S (1997) Distribution and seasonal changes of monolignol glucosides in Pinus thunbergii. Mokuzai Gakkaishi 43, 254-259. Freudenberg K (1959) Biosynthesis and constitution of lignin. Nature 163, 1152-1155. Goujon T, Sibout R, Eudes A, Mackay J, Jouanin L. (2003) Genes involved in the biosynthesis of lignin precursors in Arabidopsis thaliana. Plant Physiol Biochem 41, 677-687. Gilkey JC, Staehelin LA. (1986) Advances in ultrarapid freezing for the preservation of cellular ultrastructure. J Electron Microsc Tech 3, 177-210. Hauffe K, Paszkowski U, Schulze-Lefert P, Hahlbrock K, Dangl J, Douglas C (1991) A parsley 4CL-1 promoter fragment specifies complex expression patterns in transgenic tobacco. Plant Cell 3, 435-443. Hertzberg M, Aspeborg H, Schrader J, Andersson A, Erlandsson R, Blomqvist K, Bhalerao R, Uhlen M, Teeri T, Lundeberg J, Sundberg B, Nilsson P, Sandberg G (2001) A transcriptional roadmap to wood formation. Proc Natl Acad Sci USA 98, 14732-14737.  73 Hirschberg K, Miller CM, Ellenberg J, Presley JF, Siggia ED, Phair RD, Lippincott-Schwartz J (1998) Kinetic analysis of secretory protein traffic and characterization of Golgi to plasma membrane transport intermediates in living cells. J Cell Biol 143, 1485-1503. Hosokawa M, Suzuki S, Umezawa T, Sato Y. (2001) Progress of lignification mediated by intercellular transportation of monolignols during tracheary element differentiation of isolated Zinnia mesophyll cells. Plant Cell Physiol 42, 959-968. Humphreys JM, Chapple C (2002) Rewriting the lignin roadmap. Curr Opin Plant Biol 5, 224- 229. Kaneda M, Rensing KH, Wong JCT, Banno B, Mansfield SD, Samuels AL. 2008. Tracking monolignols during wood development in lodgepole pine. Plant  Physiol 147, 1-11. Kiss JZ, Staehelin LA. (1995) High pressure freezing. In: Severs NJ, Shotton DM, eds. Rapid Freezing, Freeze Fracture and Deep Etching. New York: Wiley-Liss Inc. Kirst M, Johnson AF, Baucom C, ulrich E, Hubbard K, Staggs R, Paule C, Retzel E, Whetten R, Sederoff R. (2003) Apparent homology of expressed genes from wood-forming tissues of loblolly pine (Pinus taeda L ) with Arabidopsis thaliana. Proc Natl Acad Sci USA 100, 7383-7388. Leinhos V, Savidge R (1994) Investigation of coniferin compartmentation in developing xylem of conifers during lignification. Acta Horticul 381, 97-102 Marcinowski S, Grisebach H (1977) Turnover of coniferin in pine seedlings. Phytochem 16, 1665-1667 Morreel K, Ralph J, Kim H, Lu F, Goeminne G, Ralph S, Messens E, Boerjan W. (2004) Profiling of oligolignols reveals monolignol coupling conditions in lignifying poplar xylem. Plant Physiol 136, 3537-3549. Muesel G, Schindler T, Bergfeld R, Ruel K, Jacquet G, Lapierre C, Speth V, Schopfer P. (1997) Structure and distribution of lignin in primary and secondary cell walls of maize coleoptiles analyzed by chemical and immunological probes. Planta 201, 146- 159. McCann MC, Stacey NJ, Dahiya P, Milioni D, Sado P-E, Roberts K. (2001) Zinnia. Everybody Needs Good Neighbors. Plant Physiol. 127, 1380-1382. Pickett-Heaps JD (1968) Xylem wall deposition: Radioautographic investigations using lignin precursors. Protoplasma 65, 181-205 Palade G. (1975) Intracellular aspects of the process of protein synthesis. Science 189, 347-358. Rensing KH (2002) Chemical and cryo-fixation for transmission electron microscopy of gymnosperm cambial cells. In J CN, ed, Wood formation in trees: Cell and molecular biology techniques. Taylor and Francis, New York, pp 65-81 Rolando C, Monties B, Lapierre C. (1992. Thioacidolysis. In: Lin S, Dence C, eds. Methods in Lignin Chemistry. Berlin, Germany: Springer-Verlag, 334-349. Salpeter M, Bachmann L. (1972) Autoradiography. In: Hayat M, ed. Principles and techniques of electron microscopy, Vol. 2. New York: Van Nostrand Reinhold Co., 221-278. Samaj J, Hawkins S, Lauvergeat V, Grima-Pettenati J, Boudet A. (1998) Immunolocalization of cinnamyl alcohol dehydrogenase 2 (CAD 2) indicates a good correlation with cell- specific activity of CAD 2 promoter in transgenic poplar shoots. Planta 204, 437-441. Samuels AL, Rensing KH, Douglas CJ, Mansfield SD, Dharmawardhana DP, Ellis BE. (2002) Cellular machinery of wood production: Differentiation of secondary xylem in Pinus contorta var latifolia. Planta 216, 72-82.  74 Savidge RA. (1989) Coniferin a biochemical indicator of commitment to tracheid differentiation in conifers. Can J Bot 67, 2663-2668. Savidge RA. (1988) A biochemical indicator of commitment to tracheid differentiation in Pinus contorta. Can J Bot 66, 2009-2102. Smith CG, Rodgers MW, Zimmerlin A, Ferdinando D, Bolwell GP (1994) Tissue and subcellular immunolocalisation of enzymes of lignin synthesis in differentiating and wounded hypocotyl tissue of french bean (Phaseolus vulgaris l.). Planta 192, 155-164 Takabe K, Fujita M, Harada H, Saiki H. (1985) Autoradiographic investigations of lignification in the cell walls of cryptomeria (Cryptomeria japonica D. Don). Mokuzai Gakkaishi 31, 613-619. Takabe K, Takeuchi M, Sato T, Ito M, Fujita M. (2001) Immunocytochemical Localization of Enzymes Involved in Lignification of the Cell Wall. J Plant Res 114, 509. Terashima N and Fukushima K (1988) Heterogeneity in formation of lignin - XI: An autoradiographic study of the heterogeneous formation and structure of pine lignin Wood Sci Technol 22, 259-270 Verrier PJ, Bird D, Burla B, Dassa E, Forestier C, Geisler M, Klein M, Kolukisaoglu U, Lee Y, Martinoia E, Murphy A, Rea PA, Samuels L, Schulz B, Spalding EJ, Yazaki K, Theodoulou FL (2008) Plant ABC proteins - a unified nomenclature and updated inventory Trends Plant Sci 13, 151-9 Whetten R, Sederoff R. (1995) Lignin biosynthesis. Plant Cell 7, 1001-1013. Whetten RW, Mackay JJ, Sederoff RR. 1998. Recent advances in understanding lignin biosynthesis. In: Jones RLE, Somerville, C. R., Walbot, V., ed. Annu Rev Plant Physiol. Plant Mol Biol 49, 585-609. Wooding FDP. (1968) Radioautographic and chemical studies of incorporation into sycamore vascular tissue walls. J Cell Sci 3, 71-80. Yazaki K (2006) ABC transporters involved in the transport of plant secondary metabolites. FEBS Lett. 580, 1183–1191  75           CHAPTER 3:  ABC TRANSPORTERS COORDINATELY EXPRESSED DURING LIGNIFICATION IN ARABIDOPSIS STEMS INCLUDE A SET OF ABCB’S ASSOCIATED WITH AUXIN TRANSPORT 1                    1. A version of a manuscript this will be published M. Kaneda, B.S.P. Lin, C. Chanis, B. Hamberger, T. Western, J. Ehlting, A.L. Samuels. ABC transporters coordinately expressed during lignification of Arabidopsis stems include a set of ABCB's associated with auxin transport   76 CHAPTER 3: ABC TRANSPORTERS COORDINATELY EXPRESSED DURING LIGNIFICATION OF ARABIDOPSIS STEMS INCLUDE A SET OF ABCB’S ASSOCIATED WITH AUXIN TRANSPORT  3.1 Introduction Lignified secondary cell walls are essential components for plants, contributing to water conduction in the vasculature system and providing mechanical strength to the plant body. Lignin is the major strengthening component in the secondary cell wall and consists of a complex polymer of monolignols, produced by the phenylpropanoid pathway, and deposited into the thickened secondary polysaccharide wall.  Intensive research using genetic and biochemical approaches have identified the genes that are involved in developmental monolignol biosynthesis (Boerjan et al., 2004).  However, the mechanisms of monolignol export from the cell to the wall are not clear.  In developing pine wood, autoradiographic evidence suggests that a Golgi mediated vesicle export route is unlikely (Kaneda et al., 2008), and implicats either membrane transporters or diffusion routes of monolignol export. The inflorescence stem of Arabidopsis has been used to study the lignification process (Turner et al., 2001), due to its abundant lignin-rich cell types such as primary xylem tracheary elements, and interfascicular fibers (IFF) between vascular bundles (Altamura et al., 2001; Lev-Yadun 1997; Lev-Yadun et al. 2005). Moreover, vascular tissues and IFF differentiate at predictable positions in the stem and display maturation gradients, with fiber differentiation following vascular bundle differentiation toward the bottom of stems. These features of ontogenesis indicate that stem patterning is under the strict control.  Using microarrays, Ehlting et al. (2005) determined which genes are differentially expressed along the axis of bolting  77 stems in Arabidopsis.  The co-expression pattern analyses of genes whose expression is correlated with phenylpropanoid pathway enzyme genes suggested that multiple ATP binding cassette (ABC) transporter genes are correlated with lignification. The ABC transporter superfamily is one of the largest transporter protein families in plants (Rea, 2007; Verrier, 2008). More than 129 genes have been annotated in Arabidopsis based on the specific nucleotide binding motif and multiple transmembrane domains in the sequence (Sanchez-Fernandez et al., 2001,Verrier et al., 2008). Plant ABC transporters move diverse substrates including fatty acids, cuticular lipids, auxin, heavy metals, xenobiotics and secondary metabolites. Multiple members of ABCB/MDR subfamily are involved in auxin transport (Geisler and Murphy, 2006), one of the most important hormones for plant differentiation and response to environmental signals. Auxin gradients and polar transport are important factors determining vascular differentiation and are critical for xylogenesis (Woodward and Bartel, 2005; Demura and Fukuda, 2007).  In this study, I examined the ABC transporter candidate genes whose expression is correlated with phenylalanine biosynthetic gene expression in Arabidopsis inflorescence stems. The goal was to test if these genes function in secondary cell wall deposition, specifically lignification. Four ABC transporter genes, which were strongly expressed in the stem vasculature, were characterized. I tested 10 mutant lines for changes in lignin deposition in the xylem or interfascicular fibers or changes in auxin transport.  None of the mutant lines had altered lignification patterns based on the measurement of thickness of secondary cell wall and colour density of lignin staining, although three ABCB/MDR candidate genes showed reduced auxin transport in the stem and one mutant line, abcb14, had disorganized vascular bundles. My data suggests that experimentally testing a monolignol export role for ABC transporters,  78 especially those of the ABCB subfamily, will be a complex undertaking, given their overlapping expression patterns and involvement in auxin transport, a hormone which is critical for normal development of cells with secondary cell walls. 3.2 Results 3.2.1 Selection of Arabidopsis stem ABC transporter genes correlated with lignification. Selecting candidate ABC transporter genes that are correlated with vascular and interfascicular fiber maturation was possible by observing the changes in gene expression levels along the axial stem (Ehlting et al., 2005).  The Arabidopsis full genome microarray data set showed that 14 ABC transporter genes (Table. 3.1) were differentially expressed along the axis of developing primary stems. Some genes, ABCB14, ABCG29 and ABCG33, were up-regulated in the stem segment 3-5 cm from the shoot apex, a region of the stem where vascular bundle development is pronounced but interfascicular fibers have not yet begun to lignify.  Others showed increasing expression towards the base of the stem where interfascicular fibers are lignifying, in a pattern that closely matched that of the phenylpropanoid biosynthetic genes (ABCB11; ABCB15; ABCG22; ABCG36/ PEN8; ABCG39).  79   MIPS locus Synonymsa ABC transporter nomenclatureb Pattern of expression in stem c 1 At3g28345 MDR13 ABCB15 High from 3cm to base 2 At1g02520 MDR8 ABCB11 High from 3cm to base 3 At5g06530 WBC23 ABCG22 Increasing towards base 4 At1g59870 PDR8/PEN3 ABCG36 Increase towards base 5 At5g61690 ATH15 ABCA11 High in 7-9cm segment 6 At1g66950 PDR13 ABCG39 Stem apex only 7 At3g16340 PDR1 ABCG29 High at 3-5 cm segment 8 At4g15230 PDR2 ABCG30 Negative in 3-5cm stem 9 At5g61730 ATH11 ABCA9 High between 2-5 cm 10 At2g37280 PDR5 ABCG33 Stem apex only 11 At4g25960 MDR2 ABCB2 Negative at base 12 At2g26910 PDR4 ABCG32 Decreasing towards base 13 At1g28010 MDR12 ABCB14 Stem apex only 14 At2g13610 WBC5 ABCG5 Negative at base a Arabidopsis ABC protein system by Sanchez-Fernandez et al. 2001 b  Plant ABC proteins updated nomenclature by Verrier et al. 2008 c  data from Ehlting et al. (2005) Operon 'longmer' microarray study  Table 3.1 ABC transporter genes whose expression is correlated with phenylpropanoid biosynthetic genes in Arabidopsis inflorescence stems.   80 Genes encoding phenylpropanoid biosynthetic enzymes contain short conserved sequence motifs in the promoter region called the A box, L box and P box (Hahlbrook et al., 1995).  If these elements were present in selected ABC transporter gene promoters, there is an increased probability that these selected ABC transporter genes would be co-regulated with lignin biosynthesis pathway genes and could be involved in development of lignified cells.  The 1000 base pairs upstream from each ATG of candidate ABC transporter gene were analyzed to identify A box, P box and L box (Fig. 3.1) by “MotifViz” (http://biowulf.bu.edu/MotifViz). The nucleotide element matrices were kindly provided by Dr. Bjorn Hamberger from University of British Columbia. Phenylpropanoid pathway enzyme genes contained all three elements, as illustrated in Fig. 3.2. Five of the top candidate ABC transporters whose gene expression was correlated with lignification also contained P boxes in their upstream sequence (Fig. 3.2).  Of all the promoter regions examined, the phenylpropanoid genes were also likely to contain L and A motifs, while ABC transporter gene promoters did in some, but not all, cases (Fig. 3.2).  As a negative control, the promoter regions of ABC transporter genes, whose expression is higher in the epidermis than the stem as a whole (Suh et al., 2005), were also examined for these cis-elements.  Epidermal ABC transporters’ promoters contained no P or A motifs and only rarely L (Fig. 3.2).  One of the ABC candidates, ABCG22, lacked any of these motifs and its pattern of gene expression in the microarray, coming on relatively late in development, made it an unlikely candidate to be involved in lignified cell differentiation. The presence of the motifs in the remaining candidate ABC transporter genes’ promoters is consistent with the co-regulation of these genes with lignin biosynthetic pathway genes.  81  Figure 3.1 Sequence logos visualizing the consensus of boxes P, A and L boxes and their nucleotide matrix. All published putative cis-acting elements common to phenylpropanoid biosynthetic gene promoters and with similarity to the previously established in vivo DNA footprint of parsley (Petroselinum crispum PcPAL1) were used to create the consensus matrices for the boxes P, A and L.  These matrices were provided by Dr. B. Hamberger.  82  Figure 3.2 In silico promoter analysis of candidate ABC transporters whose expression is correlated with lignin biosynthetic pathway enzyme genes. A) The presence of P, A and L boxes, motifs commonly found upstream from genes encoding phenylalanine biosynthetic enzymes as illustrated for phenylalanine ammonia-lyase (PAL) and p-coumarate-3-hydroxylase (C3H).  The presence of these motifs was mapped upstream from the translational start for each candidate ABC transporter gene. The coding region is shown in black. B) The proportion of genes that contain a P, L, or A motif within each gene group is shown for phenylpropanoid biosynthetic enzyme genes, stem ABC transporters and, as a negative control, epidermally-expressed ABC transporters of the Arabidopsis stem.  83 One of the problems with microarray data sets is that often the extracted mRNA arises from multiple tissues and cell types.  Cross-referencing multiple data sets is one way to narrow gene expression to a specific cell type or developmental stage of interest, in this case, lignification.  The list of candidate genes was narrowed by the elimination of ABC transporters with high expression in the epidermis, where ABC transporters are known to be involved in lipid export to the cuticle (reviewed by Bird, 2007).  ABC transporters whose expression is high in the epidermis were reported in a study of cuticle development across the developmental gradient from inflorescence apex to base (Suh et al., 2005).  By comparing the epidermal gene expression levels of ABC transporters, candidate ABC transporter genes whose expression was correlated with phenylpropanoid biosynthetic genes but whose epidermal expression makes them unlikely candidates for developmental lignification-related processes in the xylem and interfascicular bundles were eliminated (Table 3.2).  In addition to ABCG41, another ABCG candidate that fell into this category was ABCG36/ PEN3, an ABC transporter that was identified in a forward genetic screen for mutants deficient in resistance to powdery mildew haustoria (Stein et al., 2006).  84    MIPS locus  Gene namea New Nomenclatureb Epidermal expression patternc Targeting predictiond 1 At1g02520 MDR8 ABCB11 ND Other 2 At1g28010 MDR12 ABCB14 Stem >Epi Other 3 At3g28345 MDR13 ABCB15 Stem >Epi Other 4 At2g37280 PDR5 ABCG33 Stem >Epi Other 5 At3g16340 PDR1 ABCG29 Stem > Epi Chl aArabidopsis ABC protein system by Sanchez-Fernandez et al. 2001 bPlant ABC proteins updated nomenclature by Verrier et al. 2008 cData represented on Affymetrix "full genome" microarray study of Arabidopsis Stem epidermal peel with reference to whole stems by Suh et al. 2005. ND= no data dSubcellular targeting predictions based on sequence data were made using Aramemnon.   Table 3.2 Candidate ABC transporter genes and their expression in the Arabidopsis stem epidermis and whole stem, based on microarray data. In addition, in silico predictions of their subcellular targeting are shown.  85  After eliminating epidermis-enriched ABC transporter genes, sequence information from the remaining candidates was examined for predicted subcellular targeting information.  If a transporter were involved in monolignol export, it would be predicted to be located at the plasma membrane (PM), requiring it to be trafficked via ER and Golgi during its biosynthesis. N-terminal hydrophobic signal sequences predict targeting to the ER and endomembrane system, while an internal hydrophobic sequence of amino acids can act as a ‘start transfer’ signal to target a membrane protein with a cytoplasmic N-terminal to the ER.  Several of these ABC transporter candidates have been detected in the PM by either proteomics (ABCB11; Dunkley et al., 2006) or green fluorescent protein (GFP) fusion localization (ABCG39/ PEN3; Stein et al., 2006).  The predicted protein architecture of ABC transporters includes ATP binding cassettes, also called nucleotide binding domain (NBD), as well as clusters of transmembrane alpha-helices (TMD).  The architecture of (TMD-NBD)2 is conserved for the ABCB/MDR subfamily of ABC transporters and (NBD-TMD)2 for ABCG/PDR  subfamily of ABC transporters (Verrier et al., 2008).  Using Aramemnon, a compendium of web-based targeting prediction programs (http://aramemnon.botanik.uni-koeln.de/), eight ABC transporters are predicted to have “Other” as a protein destination, indicating that they could have internal start transfer sequences and cytoplasmic N-terminal sequences, the targeting prediction expected for the protein architecture of both the ABCB/MDR and ABCG/PDR subfamilics (Table. 3.2). In one case, C-terminal targeting sequences predict targeting of the ABC transporter ABCG29 to the chloroplast so this ABC transporter was not investigated in further detail.  86 After predicted targeting information was considered, the remaining four candidate ABC transporters (ABCB11; ABCB14; ABCB15; ABCG33) were further investigated by promoter activity assays and analysis of T-DNA insertional mutants. 3.2.2 Promoter::GUS fusions indicate that ABCB15, ABCB14, ABCB11 and ABCG33 have high vasculature expression and overlapping expression patterns. The previously described gene expression data indicated that these ABC transporter genes have differential developmental expression along the inflorescence stem.  In particular, ABCB11 and ABCB15 appear to be expressed in a pattern co-incident with interfascicular fiber lignification, while ABCB14 and ABCG33 were more highly expressed towards the stem apex where vascular bundles are differentiating.  In this experiment, promoter::GUS constructs were made to report where in the stem the candidate ABC transporters would be expressed during development and, more specifically, to distinguish if these genes are actively expressed in the vasculature or interfascicular regions.  The 1000 base pairs upstream from translational start of ABCB15, fused to GUS, led to the highest expression of any of the candidate ABC transporters in the stem.  In transverse hand sections cut from the top of the stem, there was strong pABCB15::GUS activity in the vascular bundles and in the parenchyma cells between vascular bundles that were differentiating into interfascicular fibers (Fig. 3.3C).  The upstream sequence of ABCB11 fused to GUS led to GUS activity in an overlapping pattern with pABCB15::GUS, i.e.  GUS activity was found in vascular bundles including differentiating xylem tracheary elements and parenchyma cells and phloem of the young stem (Fig. 3.3A,B).  Again, early in interfascicular fiber differentiation, the pABCB11 was active in the fiber precursor cells, i.e. parenchymous cells between vascular bundles, as shown in resin sections of young pABCB11::GUS stem where the interfascicular  87 fiber precursors arise (Fig. 3.3A). Additionally, pABCB11::GUS activity was seen in the epidermis of the young stem.  88           Figure 3.3 Gene expression profiles of ABCB15 and ABCB11 in young stem segments. A) Resin section of pABCB11::GUS transgenic plants with stain in interfascicular fiber precursors, xylem parenchyma and tracheary elements and phloem. Arrows indicate interfascicular region. B) hand section of the young stem showing strong vascular bundle expression and interfascicular expression. C) hand section of pABCB15::GUS transgenic plants with stain showing overlapping activity with pABCB11 in the vascular bundles.   89  In the more mature base of the stem, both pABCB15::GUS and pABCB11::GUS activity was more restricted to phloem (Fig. 3.4), in contrast to the immature stem that showed more general expression in vascular bundles and interfascicular regions. As the stem matured, there was a subtle shift in the expression pattern: the vascular bundles still stained but there was additional GUS activity in the photosynthetic cortex and the epidermis, including guard cells (Fig. 3.4). The increasing promoter activity in the cortex and epidermis and continuing expression in the phloem (Fig. 3.4), rather than interfascicular fiber expressions, may explain why the pattern of gene expression found by microarray shows a steady increase in intensity towards the base of the stem. The upstream sequence that produced the weakest GUS activity was that of ABCB14. Conventional chromogenic substrate produced weak blue signal that could barely be detected in the vascular bundles.  Using a fluorogenic substrate, ABCB14 promoter activity was observed in procambium and vascular bundles of the elongation region of the stem (Fig. 3.5).  90       Figure 3.4 ABC transporter expression in mature (within 1 cm of rosette) stem segments. A) and C) pABCB11::GUS activity in vascular bundles, both in phloem (phl) and xylem parenchyma (arrow), cortex (c) and epidermis (epi). B) and D) pABCB15::GUS activity in vascular bundles, both phloem and xylem, as well as cortex.  No activity is seen in interfascicular fibers (IFF). Bars= 50 m   91  Figure 3.5  pABCB14::GUS activity detected by fluorogenic substrate. A) WT control, apical 1 cm lacked yellow-green fluorescence and displayed only red chlorophyll autofluorescence. B) ABCB14::GUS transgenic plants stained in the vascular bundles of apical 1 cm of stem.  C) Apex of stem with clusters of floral buds and procambium stained by GUS activity. D) An anther containing pollens showed high GUS activity.    92 As with the ABCB subfamily members described above, the upstream sequence from ABCG33 fused to GUS produced reporter activity in the vascular bundles, both in xylem and phloem, and differentiating fibers/interfascicular parenchyma cells of the young stem (Fig. 3.6). However, promoter activity of ABCG33 decreased in intensity towards the base of the stem, as predicted by the microarray data.  In the stem base, the epidermis and the phloem contained GUS product, with weak staining in pith and cortex. In addition, GUS activity was observed in developing vascular cylinder above the differentiation zone in seedling roots of pABCG33::GUS, pABCB15::GUS and pABCB11::GUS (Fig. 3.7).  93  Figure 3.6 Gene expression profiles of candidate ABC transporter genes, pABCG33::GUS in stem vasculatures. A) Young stem sections, and  B) and C) mature stem sections showed constant vasculature (V) specific GUS expression. cortex, epi: epidermal cells, IFF: interfascicular fiber, phl: phloem tissue.  94  Figure 3.7 GUS expression profiles of candidate ABC transporter gene in seedling root tissue A) to F) Seedling root of promoter::GUS showed vascular cylinder specific GUS activity but no root tip in all three transporters. Bar = 250 m  95  3.2.3 Reverse genetics of candidate ABC transporter genes in Arabidopsis  For further analysis of the function of these selected genes, T-DNA insertional mutant lines were obtained from the SIGnal database (http://signal.salk.edu).  Lines were selected depending on the position of the insertion in the gene to increase the likelihood of disrupting function, i.e. in an exon or promoter.  The Salk lines tested for the candidate ABC transporters are shown in Table 3.3.  For each Salk line, T-DNA homozygous mutant lines were selected by PCR, based on the presence or absence of the T-DNA insertion using T-DNA specific primers and gene specific primers.  A T-DNA insertion specific primer and a gene specific primer produced a PCR product in the mutant line but not in wild-type. Gene specific primers did not produce a PCR product in the mutant but did give a PCR product in wild-type.  96  MIPS locus Synonyms  Synonyms  mutant name  T-DNA insertions Polymorphism site At1g02520 ABCB11 MDR8/PGP11 abcb11 Salk_094249 exon At1g28010 ABCB14 MDR12/PGP14 abcb14 Salk_016005 exon       abcb14-1 Salk_026876 exon At3g28345 ABCB15 MDR13/PGP15  Salk_121099 promoter    abcb15-1 WiscDsLox501E11 exon    abcb15-2 SAIL1187-C04  exon        (CS843941)     Salk_036868 intron         Salk_034562 intron At3g16340 ABCG29 PDR1 abcg29 Salk_113825 exon At2g37280 ABCG33 PDR5 abcg33 Salk_002380 exon  Table 3.3 T-DNA insertion lines list and their polymorphism sites  97  Based on the correlations between phenylpropanoid biosynthetic gene expression and the candidate ABC transporters, we predicted that the loss of function of a gene might lead to abnormal vascular bundle development. If so, it suggests that the gene is required for vasculature formation.  Transverse hand sections were cut at various developmental stages and stained with toluidine blue and phloroglucinol.  The thickness of lignified secondary cell wall and the density of lignin staining were compared between wild-type and mutants at early stages of development, i.e. 3 cm from the top of a 10 cm stem, and in mature vascular bundles at the base of the stem. There were no changes of lignin quantity or cell wall thickness, or significant differences from wild-type vasculature in all 12 lines, with the exception of the abcb14 T-DNA insertion line. The abcb14 mutant stem had slightly disorganized vascular bundles, with phloem occupying a smaller cross-sectional area than in wild-type stems (Fig. 3.8, compare A and B). In transverse sections along the developmental axis of the stem for abcb14, the phloem area was significantly smaller than wild-type phloem through the stem (Fig 3.8C).  In addition, the vessel elements of the abcb14 metaxylem in the vascular bundles were reduced in diameter compared to wild-type xylem (Fig. 3.8D). None of the mutants had changes in the interfascicular fibers development, morphology, or lignification. Unlike mutants like abcb1/pgp1 and abcb19/pgp19 (Geisler and Murphy, 2006), the inflorescence stem development of all the mutants tested was identical to wild-type except abcb14, which showed slow stem development. The number of rosette leaves at the time of bolting was larger in the abcb14 and the stem growth rate compared to the wild-type stem was slower.  At the end of growth, however, the final stem height was the same as wild-type, as found in pgp19 and pgp1 (Noh et al. 2001; Geisler et al 2003).  98  Figure 3.8 The inflorescence stems of abcb14 knockout mutant showing vasculature development deficient phenotype. The vascular bundle of A) Wild type stem and B) abcb14 stem by phloroglucinol staining shows xylem with smaller vessel diameters (asterisks). Bars = 50 m.  C: cortex, IFF: interfascicular fiber, phl: phloem C) The phloem areas of abcb14 stems were significantly smaller than the phloem areas in wild-type stems, measured by length and width of cross sectioned phloem tissue. Means + standard error. D) The frequency distribution of metaxylem vessel diameters when 163 vessels in each genotype were measured. The sample size was >900 cells and from these, the largest 163 cells in mutant and wild-type were binned into size classes. More abcb xylem cells were in the smallest size class and only wild-type were in the largest size classes.  99 3.2.4 Reduced polar auxin transporter in ABC transporter mutants  There is extensive evidence linking auxin to vascular bundle development (Turner et al. 2007; Woodward and Bartel, 2005), which led us to perform experiments to test whether auxin transport is altered in the abcb14 mutant plants.  The morphological changes in the stem in the mutants indicated that there was a disruption in cell differentiation or cell development in the vasculature.  In addition, ABCB14 belongs to same ABC transporter subfamily (Fig.3.9) as AtMDR1/PGP1/ABCB1 (At2g36910) and AtABCB19/AtMDR11/AtPGP19 (At3g28860), which have both been identified as polar auxin transporters (Geisler and Murphy, 2006).  Therefore, polar auxin transport (PAT) assays were performed using the inflorescence stem of wild-type Arabidopsis and abcb14 mutants (Fig. 3.10).  The apical portion of the stem was immersed in a 3H-IAA solution for 24 hours to allow auxin transport. The amount of 3H-IAA that was transported to the distal portion of the stem was measured by excising the basal 1.5 cm of stem, which had never been in contact with the radiolabel, and immersing it in scintillation fluid.  If the stem segments were placed with the apical side away from the 3H-IAA, or the polar auxin transport inhibitor 1-N-naphthylphthalamic acid (NPA) was included with the 3H-IAA, then transport was diminished to 20 % of the wild-type control (Fig.3.10).  abcb14 mutants had reduced polar auxin transport that was 60% of the wild-type transport levels.  100  Figure 3.9 Distance-based guide tree of Arabidopsis ABCB/MDR subfamily subfamily genes Full genomic DNA sequence of Arabidopsis thaliana ABCB subfamily genes were aligned using CLUSTALX and displayed as a guide tree by TREEVIEW.   101  Figure 3.10 The measurement of polar auxin transport in inflorescence stems of abcb14 mutants using radioactive IAA. A) Schematic diagram of polar auxin transport assay for inflorescence stem.  Top 2.5 cm was dissected, inverted into 3H-IAA (centre) and incubated for 24 h.  The uppermost “basal” 1 cm was removed and radioactivity measured.  Pink arrow indicates directional IAA transport from the apical side of the stem. B) The amount of transported IAA in abcb14 inflorescence stems was significantly reduced compared to WT. Mean + SD.  Controls were treated with polar auxin inhibitor NPA or placement of the stem segments with the basal zone in 3H-IAA and apical zone facing up (Up-Right) to show that auxin transport was polar. The difference between wild type and abcb14 3H-IAA polar auxin transport is significant using the non- parametric Mann-Whitney test to compare means (p <0.05).  102 In order to observe altered auxin distributions in the abcb14 plants, the synthetic auxin- responsive promoter DR5 (pDR5), fused to GUS gene (Ulmasov et al., 1997), was crossed into the abcb14 mutant background and wild-type background.  When the inflorescence stem was tested for GUS activity, there were no differences detected when compared to pDR5 wild-type background, even in the apical region.  However, when exogenous IAA was applied in the polar auxin transport assay, and the stems exposed to the chromogenic GUS substrate, wild- type stems showed the blue product in a band pattern near the cut end of the stem (Fig. 3.11A). This could be quantified as density of pixel grey scale values in digital images and it was apparent that pDR5 wild-type accumulated significantly more product than mutant lines (Fig. 3.11B), suggesting pDR5 wild-type stems contained higher auxin levels than mutant. The difference of GUS activity between pDR5::GUS wild-type and pDR5::GUS abcb14 is considered significant (p < 0.01) by using Student’s t test.  103    Figure 3.11 The measurement of polar auxin transport in inflorescence stems using DR5::GUS activity. A)  DR5::GUS activity was induced when exogenous IAA was added to wild-type (WT) and abcb14 stems using the experimental setup described in Fig. 6A. B) Quantification of GUS activity by measuring the intensity of the blue bands in the basal side of the stem for mutant background and wild-type background after the 24 hours of treatment. Bar = 5 mm, mean+SD. Three times of independent experiments are performed. Asterisk indicates significant differences from wild-type by using Student’s t test (p < 0.01 n=40).  104 When reverse-transcriptase PCR was used to test the mRNA levels of the candidate gene of interest in the various mutant lines, there was no transcript detected in the abcb14 mutant (Salk_016005) but transcript was detected in the other Salk line for ABCB14, Salk_026876) (Fig. 3.12).  An alternative to T-DNA insertion to decrease the level of mRNA transcripts of the gene of interest is post-transcriptional gene silencing using interference RNA (RNAi), where a hairpin-type RNA is expressed in a plant.  The plant’s response to the double stranded RNA is to decrease transcript levels of the endogenous transcript as well as introduced RNA (Hilson et al., 2004).  To test the phenotype of abcb14 in independent mutant lines, plant lines transformed with RNAi specific for abcb14 lines were obtained from the Arabidopsis Genomic RNAi Knock-out Line Analysis (AGRIKOLA; www.agrikola.org).  When these lines were tested, the lines with reduced transcript (Fig. 3.12) also had reduced polar auxin transport (Fig. 3.13) with similar reductions to the T-DNA insertional mutant.  Since several of the other candidate ABC transporters were from the ABCB subfamily, all mutant lines (abcb11, abcb14, abcb15, abcg29, abcg33) were tested for altered polar auxin transport (Fig. 3.13).  Mutants in every candidate member of the ABCB subfamily tested had reduced polar auxin transport, with reductions ranging from 20% - 40%.  Polar auxin transport was significantly reduced in both abcb14 and abc14 RNAi [Mann-Whitney U=543.5 (P < 0.001) in abcb14, Mann-Whitney U= 9 (P < 0.02) in abcb14RNAi].  Polar auxin transport was also significantly reduced in abcb11 [Mann-Whitney U = 0 (P<0.002) in abcb11]. For the abcb15-2 mutants, mean polar auxin transport was always lower than wild-type but this trend was not consistently statistically significant.  In two experiments, abcb15-2 had significantly decreased polar auxin transport, while in the third experiment using abcb15-2, differences were  105 not significant.  Similarly, abcb15-1 had lower mean polar auxin transport that was not statistically significantly different than wild-type, however this mutant line also displayed detectable transcript in RT-PCR (Fig. 3-12).  In contrast to the abcb mutants, there were no lines of abcg subfamily mutants with reduced polar auxin transport.  Two lines with independent mutant alleles of the ABCG subfamily members, abcg29 and abcg33, did not statistically differ from wild-type [Mann- Whitney U=184 (P>0.05) in abcb29 and Mann-Whitney U=42.5 (P>0.05) in abcb33].  These results suggest that the function of the ABCB transporters, at least ABCB14, ABCB11 and conditionally ABCB15, in the stem could be auxin transport but, in the absence of phenotypes, functions of the ABCG subfamily are not known.  106    Figure 3.12 Transcriptional level by RT-PCR in ABC transporter candidate gene mutants. Transcriptional level of ABCB14, ABCB15 and ABCG33 were very low or not detected either in abcb14, abcb14 RNAi, abcb15-1, abcb15-2 and abcg33 compare to wild-type Columbia. Actin (ACT) and ubiqitin (UBQ10) transcriptions were used as control.    107       Figure 3.13 Polar auxin transport of inflorescence stems in ABC transporter mutants. (A) Average polar auxin transport in mutant stems in the ABCB subfamily showed reduction in mean, while ABCG subfamily mutants were the same as wild-type. For this analysis, two to five independent experiments were repeated, resulting in eight to 43 stems examined. Black stars indicate that the differences between the mutant and wild type were considered significant (p < 0.05), and a white star indicated that the difference is considered significant in two out of three time experiments (p < 0.05) by using Mann-Whitney test. No mark means there are no significant differences between mutants and wild type. Error bars represent the range of percentages obtained in different replicates.      108  3.3 Discussion The diversity of roles played by ABC transporters can be illustrated in all organisms from all kingdoms.  In plants, ABC transporters are required for transport of a broad range of compounds including wax, auxin, terpenoids and alkaloids. ABC transporters whose expression is correlated with secondary xylem development belong to the Multidrug Resistance (ABCB/MDR) subfamily and the Pleiotropic Drug Resistance (ABCG/PDR) subfamilies. These subfamilies were originally identified in mammalian and yeast cells, for their ability to export toxic compounds such as anti-cancer or anti-fungal drugs from cells.  Plant ABCG/PDRs have been shown to transport other secondary metabolites such as alkaloids (Shitan et al., 2003) and diterpenoids (Jasinski et al., 2001). In this study, we hypothesized that ABC transporters were exporting monolignols during lignification, based on their coordinate expression with phenylpropanoid metabolic genes in a microarray co-expression analysis (Ehlting et al., 2005). Testing this hypothesis has proven challenging on several fronts: first, there were an abundance of ABC transporters so the list of candidate genes had to be narrowed using a combination of in silico prediction tools. Secondly, the promoter::GUS assays suggested that all four of the ABC transporters characterized were expressed in overlapping patterns, both in lignifying tissues such as interfascicular fibers and xylem but also in non-lignifying tissues like phloem.  The abcb and abcg mutants appeared phenotypically normal, with regard to lignin, a finding that was not unexpected when overlapping expression patterns were considered. One anomaly that pointed to other potential roles for these transporters was the subtle altered vasculature phenotype of the abcb14 mutant.  Thus, our third challenge was that, although we cannot draw conclusions on a  109 role in monolignol export, it does appear that auxin transport is disrupted in the abcb/mdr mutants, a hypothesis supported by their polar auxin transport phenotypes.  So a large challenge remains: to test if these ABC transporters are involved in monolignol transport, in addition to auxin export, then making double, triple or quadruple mutant combinations will be required to overcome potential redundancy.  However making such mutants may change auxin transport, which is required for normal tissue differentiation and since lignification is a late event in differentiation of lignified cells, blocking early differentiation by altered auxin transport would lead to pleiotropic effects.  In addition, ABC transporters can be induced by xenobiotics or some secondary metabolites (Jasinski et al., 2001) so blocking transport, which would be expected to lead to a build-up of transport substrate, might induce other ABC transporters for detoxification.  These exported compounds might still be incorporated into the secondary cell wall, giving a wild-type appearance to the mutants when tested with phloroglucinol or toluidine blue. When lignin biosynthetic genes are altered, lignin monomer composition is often altered, e.g. increased H-lignin at the expense of S- and G-lignin in C3H downregulated plants (Franke et al., 2002; Coleman et al., 2008), suggesting that the export and polymerization processes are somewhat flexible. However, when the S/G ratio of lignin monomers was tested in abcb14 mutants (data not shown), it was identical to wild-type. The ABCB/MDR/PGP subfamily is the largest subfamily of full-length ABC transporters in Arabidopsis.  Our gene sequence analysis (Fig. 3.9) as well as analyses by Geisler and Murphy (2006) and Verrier et al. (2008) indicated that three clades of related genes make up the ABCB subfamily.  Our candidate genes came from each of the clades: ABCB14 from clade I whose prototype is ABCB1; ABCB11 from clade II whose prototype is ABCB4/PGP4 (the only putative auxin importer); ABCB15 from clade III whose prototype was  110 originally annotated as AtPGP8 (Geisler and Murphy, 2006) although that was more recently considered a protogene by Verrier et al. (2008).  Clade III is interesting as every member gene is found clustered on chromosome 3 in a tandem repeat pattern (At3g28345-At3g28415). Mutants of abcb14 had the vascular phenotype and the strongest polar auxin transport defect and this gene is a member of clade I, which also contains the most well characterized auxin transporting ABC transporters ABCB1/PGP1 and ABCB19/PGP19.  It is interesting that the other mutants exhibiting an auxin transport phenotype are members of the previously uncharacterized clade III.  111  NPA treatment in this study strongly reduced the polar auxin transport levels in wild-type to about 20% of control, similar to the results of 10-20% of WT found by Noh et al. (2001).  The abcb14 polar auxin transport phenotype was about 60% of wild-type transport, suggesting that other transporters contribute to the residual activity. It is likely that PGP19 is the other key player in auxin transport in the Arabidopsis stem, based on the mutant phenotype described by Noh et al. (2001) where their mdr1 mutant, now called abcb19/pgp19 had polar auxin transport levels at about 40% of wild-type. In contrast, the mutant (abcb1/pgp1) of the other major stem transporter, ABCB1, did not have a stem polar auxin transport phenotype. In their study and ours, NPA treatment of abcb mutants resulted in inhibition of transport that was similar to NPA treated wild-type.  This suggests that these ABCB transporters are NPA sensitive. Overall, GUS staining in the vasculature confirmed that these ABC transporters were being expressed in temporal association with lignifying cells, suggesting that the microarray data sets were accurate.  However, their expression was not restricted to lignifying cells suggesting that they may play a variety of functions.  The overlapping gene expression patterns in vascular tissues indicate that the developing xylem tracheary elements with their surrounding parenchyma, as well as phloem could contain multiple ABC transporters including ABCB11; ABCB14,  ABCB15 and ABCG33.  Similarly, all genes were expressed early in development of interfascicular fibers but not in the interfascicular fibers of the stem base.  Of course, mature fibers are dead but the increasing number of layers of interfascicular fibers in the stem base suggest that continuing recruitment of fibers from parenchymous pith occurs.  The lack of expression in this region was not consistent with a role for these ABC transporters in lignifying interfascicular fibers.  112 It is possible that our candidate ABC transporters selection was premature and one of these eliminated ABC transporters could still be active in monolignol export.  In some cases, the in silico targeting predictions are ambiguous or wrong, e.g. PDR8/ABCG36/PEN8 is predicted to be targeted to chloroplast but experimentally has been shown to be localized in the plasma membrane (Stein et al., 2006). The phenotype of abcb14 was changes in the vascular tissue organization. The size of the phloem area was smaller than wild-type phloem and metaxylem vessels had smaller diameters than wild-type. This suggests that this mutation is affecting vascular development generally and/or radial cell expansion of vessels during procambium differentiation.  Given the many reports link auxin transport to vascular development (Mattson et al., 1999; Little et al., 2001; Turner et al., 2007), it is a reasonable hypothesis that polar auxin transport defects in the young developing stem where ABCB14 is expressed, are responsible for the changes to the vascular system seen in abcb14.  The roles of vascular-related NAC-domain transcription factors, VND6 and VND7, in controlling metaxylem and protoxylem differentiation have been dramatically demonstrated using overexpression studies (Demura and Fukuda, 2007) and normal expression of VND6 and VND7 requires auxin as well as cytokinin (Kubo et al., 2005).  A recent paper tested the same Salk lines with T-DNA insertions in ABCB14 for defects in guard cell responses to altered carbon dioxide levels and guard cell transport of malate (Lee et al., 2008).  These authors found that ABCB14 was highly expressed in guard cells but they did not detect in procambium.  Both studies showed pABCB14 activity in pollen. My GUS assay results did not show a guard cell pattern for ABCB14, which might reflect that they used a larger 2 kb upstream promoter fragment than the 1 kb upstream sequence used in this study.  113  In conclusion, the role of ABC transporters in monolignol export remains a plausible hypothesis. The expression of multiple ABC transporters in lignifying cells and the role of ABCB subfamily members in auxin transport mean that alternative approaches will be required to test this hypothesis.  In addition, alternatives to ABC transporter mediated monolignol export are still possible (Yazaki, 2006).  There is a large gene family of transporters, called ‘Multi- drug and toxin exporter’ (MATE), which could use the energy of the proton gradient to export monolignols, although MATE mutants have not been tested for reduced lignin phenotypes.  114  3.4 Materials and methods 3.4.1 In silico promoter cis-element analysis The up-stream of 1000 base pair promoter regions of candidate ABC transporter genes, previously identified by Ehlting et al. (2005), and phenylpropanoid pathway genes, were tested for the presence of the cis-elements called A box (CCGTCC), P box (CTTCAACCAACCCC) and L box (TCTCACCTACC) by “MotifViz” (http://biowulf.bu.edu/MotifViz).  Upstream sequences were obtained from the TAIR website (www.tair.com) and the weighting element matrix was kindly provided by Dr. B.Hamberger (UBC). 3.4.2 Plant materials and growth conditions Arabidopsis thaliana (ecotype; Col-0) was used in this research. Wild-type and transgenic seeds were surface-sterilized with 3 % H2O2 in 50 % ethanol. Sterile seeds were plated on Arabidopsis thaliana (AT) medium containing 0.9% agar in plastic Petri dishes. They were incubated at 4oC in the dark for 2 days. Then plants were grown at 21 oC under 24 h light condition for 7 days. Plants were transferred to soil (MetroMix#3, SunGro, USA) and grown at 21oC long-day conditions (16 h light/ 8 h dark) or 24 h light conditions in a Conviron E7-2 growth chamber.  T-DNA insertional mutant lines for candidate ABC transporters were obtained from the Arabidopsis Biological Resource Center (http:// arabidopsis.org).  The presence of the T-DNA insertion was detected by PCR using primers listed below. Lines were genotyped to ensure homozygosity at the insertion site or transcriptional levels were tested by semi-quantitative RT- PCR (Fig. 3.12). Homozygous mutants of T-DNA insertion were obtained by PCR amplification using a combination of gene specific primers and T-DNA primer.  115   The RNA interference (RNAi) lines for ABCB14, ABCB14 RNAi N213345, were obtained from the Nottingham Arabidopsis Stock Center (http://www.agrikola.org). 3.4.3 Generating promoter::GUS constructs To generate GUS:: promoter fusion lines of 4 ABC transporter genes, the promoter regions of PDR5/ABCG33 (At2g37280), ABCB14 (At1g28010), ABCB11 (At1g02520) and ABCB15 (At3g28345) were amplified by PCR using the following primers. For ABCG33, forward primers, 5’-TTTGAATTCGAATTGGACACAGTTTGGCTA-3’; reverse primer, 5’-CCAGGATCCTTTCCAATCTCTGCAAGCTC-3’ (underlined are Bam and EcoRI cut sites, respectively). For ABCB11, forward primers, 5’-CGGGAATTCGGGAATCGCAAGAAGGTAAG-3’; reverse primer, 5’-CGTGGATCCTTCGGCGCTGACAAAAAT-3’ (underlined are Bam and EcoRI cut sites, respectively). For ABCB14, Forward primers, 5’-TGTGGATCCTTATCACTTTAGCACTGTG-3’; reverse primer, 5’-TTAGGATCC ATGGATGGGAATTGATGC-3’. For ABCB15, forward primers, 5’-TGTGGATCCGCCTATAAATAAAGTACGTAAGAC-3’; reverse primer, 5’-CCCGGATCCTTGTGTGTGTTTAAAGAGACAG-3’ line (underlined are Bam HI cut sites).  PCR products were cloned into pUC10 and checked by sequencing and enzyme digestion. After digestion of the plasmid with BamH1/BamH1, the fragments were purified and cloned into the BamH1 site of pCAMBIA1281Z binary vector.  The constructs (pCAMBIA- ABCG33pro::GUS, pCAMBIA-ABCB11pro::GUS, pCAMBIA-ABCB14pro::GUS and pCAMBIA-ABCB15pro::GUS) were introduced into Agrobacterium tumefaciens strain  116 LB4404. Arabidopsis were transformed with these constructs by the floral dip or spraying method. TransgenicT1 plants were selected on AT media plates that contained 30 ug/ml of hygromycin.  In situ GUS staining was performed by incubation of inflorescence stems and seedlings of transformants in 90% acetone for 30 min under the vacuum. Then, tissues were rinsed 3 times in a solution of 50 mM Na phosphate buffer, pH 7.0, 500 nM Fe(CN), 0.1% Triton X- 100. The tissues were exposed to X-Gal (1 mM 5-bromo-4-chloro-3-indolyl-ß-D-glucuronic acid) at 37oC overnight. Stained tissues were cleared with 50% ethanol.  Arabidopsis tissues which had been GUS stained were taken into 100% acetone, then the inflorescences were infiltrated in Spurr’s resin through a graded resin series using low temperature microwave PELCO 3450 LABORATORY MICROWAVE, USA). The samples, under vacuum, were microwaved for 3 min each in a graded resin series at 28oC and polymerized at 70oC. The polymerized samples were sliced into 1-2 um sections.  The sections viewed with a Leica DMR compound microscope (Wetzlar, Germany) and images captured with a Q-Cam (Burnaby, BC) camera.  For histochemical staining of pABCB14::GUS expression in stems, the longitudinally dissected young transgenic and wild-type stems were incubated in 50 m C12 FDGlcU (ImaGene Green, Molecular Probes I-2908) at room temperature for 3 hours. Then, the dissected surface of stem was observed under the fluorescent light by Leica light microscope with fluorescein filter to detect the yellow-green fluorescent GUS products.   117 3.4.4 Reverse transcription PCR and T-DNA The extracted mRNA of T-DNA insertion mutant and RNAi mutant using TRIzol reagent (Invitrogen) were used as template for cDNA synthesis with oligo(dt) primers. About 2 ug of cDNA was obtained from RT reaction and used for PCR amplification with target gene specific primers. 3.4.5 Light microscopy and lignin histochemistry Hand sections were placed on glass slides. One slide was stained with toluidine blue for structural reference. Phloroglucinol (10% in 90% ethanol for 5 min) stained to see the general lignin distribution. Maule staining (3% potassium permanganate for 5 min at room temperature) was used to distinguish G-lignin (yellowish) and S-lignin (reddish) in secondary cell walls. The sections were mounted with concentrated HCl and observed with differential interference contrast using a Zeiss Axioplan light microscope (Carl Zeiss AG, Germany). Images were captured using a Q-CAM digital camera (Q-Imaging, Burnaby, BC, Canada). 3.4.6 Polar auxin transport analysis in ABC transporter mutants. Basipetal auxin transport in inflorescence stems was measured by using 3H-indole acetic acid (3H-IAA), using the protocol of Dai et al. 2006.  When Arabidopsis inflorescence stems were about 10-15 cm, the top 3 cm stems were cut and floral parts removed.  Only the apical 2-3 mm of the stems were submerged in 30 ul of solution (5 mM MES buffer containing 1.45 m 3H- IAA with 1% sucrose [W/V], pH 5.5) containing 100 nM 3H-IAA for 24 hrs at room temperature in darkness.  The basal 1.5 cm region of the stem, which had been upended in the vial was carefully removed so it never contacted the radioactive solution, then placed into scintillation vials with 3 ml of scintillation cocktail (Sigma-give exact kind) for 24 hrs.  The  118 radioactive counts per minute (CPM) was measured by scintillation counting (Beckman LS600IC) for 2 min in 5 to 10 stems per genotype in 2 to 5 replicate experiments.  3.4.7 DR5::GUS in abcb14 DR5::GUS was crossed with the abcb14 mutant. The presence of the DR5::GUS construct and the T-DNA insertions in the mutants were identified by PCR, as described above. Stems from transformed plants were treated with 1.45 m IAA for 12 hours. Then, the whole dissected stems were stained by X-Gal as above. 3.4.8 Relationship analysis of ABCB subfamily genes Full genomic DNA sequences of 21 ABCB subfamily genes were aligned, and the distance was estimated using neighbour-joining method based on pairwise score produced by CLUSTALW version 2 (Larkin et al. 2007). The NJ tree of ABCB subfamily genes was displayed by TREEVIEW X version 0.5.0 (http://darwin.zoology.gla.ac.uk/~rpage/treeviewx/index.html)  119  3.5 Bibliography Altamura, M., Possenti M, Matteucci A, Baima S, Ruberti I, Morelli G. (2001) Development of the vascular system in the inflorescence stem of Arabidopsis. New Phytol 151, 381- 389. Bird, D.A.  (2008) The role of ABC transporters in cuticular lipid secretion. Plant Sci, 174, 563-569 Boerjan, W., Ralph J, Baucher M. (2003) Lignin biosynthesis. Plant Physiol. 54, 519-546. Coleman, HD., J.-Y. Park, R. Nair, C. Chapple, and S. D. Mansfield (2008) RNAi-mediated suppression of p-coumaroyl-CoA 3'-hydroxylase in hybrid poplar impacts lignin deposition and soluble secondary metabolism. Proc Natl Acad Sci USA, 105, 4501 - 4506. Demura, T., Fukuda. (2007) Transcriptional regulation in wood formation. Trends In Plant Sci 12, 65-70. Dunkley, TPJ., HS, Shadforth IP, Runions J, Weimar T, Hanton SL, Griffin JL, Bessant C, Brandizzi F, Hawes C, Watson RB, Dupree P, Lilley KS (2006) Mapping the Arabidopsis organelle proteome. Proc Natl Acad Sci USA 103, 6518-6523. Ehlting, J., Mattheus N, Aeschliman DS, Li E, Hamberger B, Cullis IF, Zhuang J, Kaneda M, Mansfield SD, Samuels L, Ritland K, Ellis BE, Bohlmann J, Douglas CJ. (2005) Global transcript profiling of primary stems from Arabidopsis thaliana identifies candidate genes for missing links in lignin biosynthesis and transcriptional regulators of fiber differentiation Plant J 42, 618-640. Franke, R., Humphreys JM, Hemm MR, Denault JW, Ruegger MO, Cusumano JC, Chapple C. (2002) The Arabidopsis REF8 gene encodes the 3-hydroxylase of phenylpropanoid metabolism. Plant J 30, 33-45. Geisler, M., H. Üner Kolukisaoglu, Rodolphe Bouchard, Karla Billion, Joachim Berger, Beate Saal, Nathalie Frangne, Zsuzsanna Koncz-Kálmán, Csaba Koncz, Robert Dudler, Joshua J. Blakeslee, Angus S. Murphy, Enrico Martinoia and Burkhard Schulz. (2003) TWISTED DWARF1, a Unique Plasma Membrane-anchored Immunophilin- like Protein, Interacts with Arabidopsis Multidrug Resistance-like Transporters AtPGP1 and AtPGP19. Mol Biol Cell 14, 4238-4249. Geisler, M., Murphy A. (2006) The ABC of auxin transport: the role of p-glycoproteins in plant development. FEBS Letters 580, 1094-1102. Hahlbrock, H., Scheel D, Logemann E, Nürnberger T, Parniske M, Reinold S, Sacks WR, Schmelzer E. (1995) Oligopeptide elicitor-mediated defense gene activation in cultured parsley cells. Proc Natl Acad Sci USA 92,4150-4157. Hilson, P., Allemeersch J, Altmann T, Aubourg S, Avon A, Beynon J, Bhalerao RP, Bitton F, Caboche M, Cannoot B, Chardakov V, Cognet-Holliger C, Colot V, Crowe M, Darimont C, Durinck S, Eickhoff H, Falcon de Longevialle A, Farmer EE, Grant M, Kuiper MTR, Lehrach H, Léon C, Leyva A, Lundeberg J, Lurin C, Moreau Y, Nietfeld W, Paz-Ares J, Reymond P, Rouzé P, Sandberg G, Dolores Segura M, Serizet C, Tabrett A, Taconnat L, Thareau V, Van Hummelen P, Vercruysse S, Vuylsteke M, Weingartner M, Weisbeek P J, Wirta V, Wittink FRA, Zabeau M and Small I (2004) Versatile gene-specific  120 sequence tags for Arabidopsis functional genomics: transcript profiling and reverse genetics applications, Genome Res 14, 2176-2189. Jasinski, M., Stukkens Y, Degand H, Purnelle B, Marchand-Brynaert, J, Boutry M. (2001) A plant plasma membrane ATP binding cassette-type transporter is involved in antifungal terpenoid secretion. Plant Cell 13,1095-1107. Kaneda, M., Rensing KH, Wong JCT, Banno B, Mansfield SD, Samuels AL. (2008) Tracking monolignols during wood development in lodgepole pine. Plant Physiol 147, 1750- 1760. Kubo, M., Makiko Udagawa, Nobuyuki Nishikubo, Gorou Horiguchi,, Masatoshi Yamaguchi JI, Tetsuro Mimura HFa, Demura T. (2005) Transcription switches for protoxylem and metaxylem vessel formationsion Genes & Devel 19, 1855–1860. Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ and Higgins DG (2007) ClustalW and ClustalX version 2.Bioinformatics 23(21): 2947-2948. Lee, M., Y Choi, B Burla, Y Kim, B Jeon, M Maeshima, J-Y Yoo, E Martinoia, Y Lee (2008) The ABC transporter AtABCB14 is a malate importer and modulates stomatal response to CO2. Nature Cell Biol 10, 1217-1223. Lev-Yadun, S. (1997) Fibers and fiber-sclereids in wild-type Arabidopsis thaliana. Ann Bot 80, 135-129. Lev-Yadun, S., Wyatt SE, Flaishman MA. (2005) The inflorescence stem fibers of Arabidopsis thaliana Revoluta (ifl1) mutant. J Plant Growth Reg 23, 301-306. Little, C., MacDonald JE, Olsson O. (2002) Involvement of indole-3-acetic acid in fascicular and interfascicular cambial growth and interfascicular extraxylary fiber differentiation in Arabidopsis thaliana inflorescence stems. Int J Plant Sci 163, 519- 529. Mattsson, J., Sung ZR, Berluth T. (1999) Responses of plant vascular systems to auxin transport inhibition. Development 126, 2979-2991. Noh, B., Murphy A, Spalding EP. (2001) Multidrug resistance-like genes of Arabidopsis required for auxin transport and auxin-mediated development. Plant Cell 13, 2441- 2454. Rea, PA. (2007) Plant ATP-Binding Cassette Transporters. Ann Rev Plant Biol 58, 347-375. Sanchez-Fernandez, R., T. G. Emyr Davies. (2001) The Arabidopsis thaliana ABC Protein Superfamily, a Complete Inventory. J Biol Chem 276, 30231–30244. Shitan, N., Bazin I, Dan K, Obata K, Kigawa K, Ueda K, Sato F, Forestier C, Yazaki K. (2003) Involvement of CjMDR1, a plant MDR-type ABC protein, in alkaloid transport in Coptis japonica. Proc Natl Acad Sci USA, 100, 751-756. Stein, M., Dittgen J, Sanchez-Rodrıguez C, Hou B-H, Molina A, Schulze-Lefert P, Lipka V, Somerville S. (2006) Arabidopsis PEN3/PDR8, an ATP Binding Cassette transporter, contributes to nonhost resistance to inappropriate pathogens that enter by direct penetration. Plant Cell 18, 731-746. Suh, MC., Samuels AL, Jetter R, Kunst L, Pollard M, Ohlrogge J, Beisson F. (2005) Cuticular Lipid Composition, Surface Structure, and Gene Expression in Arabidopsis Stem Epidermis. Plant Physiol 139, 1649-1665. Turner, S., Tayler N, Jones L  (2001) Mutation of the secondary cell wall. J. Plant Mol Biol 47, 209-219.  121 Turner, S., Gallois P, Brown D. (2007) Tracheary Element Differentiation. Annu Rev Plant Biol 58, 407-433. Ulmasov, T., Hagen G, Guilfoyle TJ. (1997) ARF1, a transcription factor that binds to auxin response elements. Science 276, 1865-1868. Verrier, P., Bird D, Burla B, Dassa E, Forestier C, Geisler M, Klein M, Kolukisaoglu U, Lee Y, Martinoia E, Murphy A, Rea PA, Samuels L, Schulz B, Spalding EJ, Yazaki K, Theodoulou FL. (2008) Plant ABC proteins - a unified nomenclature and updated inventory. Trends Plant Sci 13, 151-159. Woodward, AW., Bartel B. (2005) A receptor for auxin. Plant Cell 17, 2425-2429. Yazaki, K. 2006. ABC transporters involved in the transport of plant secondary metabolites. FEBS Letters 580, 1183-1191.  122                 CHAPTER 4:  TESTING THE ROLE OF ABC TRANSPORTERS ON LIGNIN DEPOSITION DURING XYLOGENESIS USING INHIBITORS 1                    1. A version of this chapter will be published: M. Kaneda, B. Banno, S. Raven, A.L. Samuels. Tracheary element lignification in primary root of Arabidopsis: a novel assay to test mechanisms of monolignol export.  123 CHAPTER 4: TESTING THE ROLE OF ABC TRANSPORTERS ON LIGNIN DEPOSITION DURING XYLOGENESIS USING INHIBITORS 4.1 Introduction Autoradiography data from Kaneda et al. (2008) indicated that monolignols are transported to the extracellular matrix by non-Golgi mediated mechanisms. One such mechanism of monolignol deposition could be membrane transporter-mediated export of monolignols. Due to their ability to transport secondary metabolites and wide variety of substrates, ABC transporters are good candidates to transport monolignols.  However, as the previous chapter demonstrated, using molecular genetics approaches to knock out single ABC transporters of the ABCB/MDR subfamily is problematic because multiple ABC transporter genes show overlapping gene expression patterns, and they are also required for auxin transport.  In addition, neighbors of lignifying cell types contain multiple ABC transporters.  In order to inactivate a broader spectrum of ABC transporters, chemical inhibitors were chosen, which have been shown to halt ABC transporter functions in other cell types.  ABC transporters have been found to be responsible for multi-drug resistance in mammalian cancer cells (Gottesman and Pastan 1993, Dean et al., 2001).  Inhibition of ABC transporters is a major target for chemotherapy or cancer treatments, and thus there are many different chemical compounds with high affinity to ABC transporters that act as inhibitors. Vanadate, fumitremorgin C and reversin were conventionally used as inhibitors for ABC type drug pumps (al-Shawi & Senior, 1993). Vanadate inhibits all P-type ATPase activities such as Ca2+ ATPase and Na+/K+ ATPase including ABC transporters and, in plant cells, the plasma membrane H+-ATPase (Palmgren, 2001).  Reversin 121 has been characterized as an inhibitor of multidrug resistance in cancer tissues by halting the efflux of anticancer drugs carried out by  124 ABCB/MDR subfamily ABC transporters (Cardenas et al. 1999, Lowes et al. 2002). Fumitremorgin C (FTC) is a fungal toxin that binds specifically to the ABCG/WBC subfamily of ABC transporters such as the human breast cancer resistance protein (Robey et al., 2001; Ozvegy et al. 2002). Relatively few inhibitors have been tested on plant ABC transporter function. Vanadate was found to have high affinity for CjMDR1 and could clearly inhibit CjMDR1-dependent berberine uptake (Shitan et al., 2003, Terasaka et al., 2003).  The transport of flavonoids into the tonoplast by ABCC/MRP-type ABC transporters was tested, and vanadate was shown to inhibit uptake of flavonoids in isolated vacuoles from barley and broccoli mesophyll cells (Klein et al. 2001).  Auxin efflux by ABCB1/PGP1 and ABCB19/PGP19 from mesophyll protoplasts was reduced by (1-N-naphthylphthalamic acid) NPA and Cyclosporin A (Geisler et al., 2005).  In mammalian HeLa cells expressing ABCB4/PGP4, NPA inhibited auxin transport but cyclosporin A did not (Terasaka et al., 2005). Tests of ABC transporter function, particularly for the ABCG/PDR subfamily, have used a transport substrate called sclareol.  A member of the ABCG/PDR subfamily genes had increased gene expression in the presence of sclareolide, a compound with similar structure to the antifungal diterpenoid sclareol in tobacco (Jasinski et al 2001). While sclareol is a toxic to plants, ABCG/PDR transporters can export sclareol out of the root. Sclareol has been used to characterize the detoxification phenotype of the Arabidopsis mutant pdr12/abcg40, where wild- type roots can export sclareol and mutant root elongation was inhibited due to lack of transport activity (Campbell et al., 2003). In this study, we tested an inhibitor, FK506 (Okuhara et al., 1987) for its ability to inhibit sclareol export from roots. FK506 has been shown to be a potent inhibitor in yeast of the ABCG subfamily, which is closely related to the plant PDRs.  If FK506  125 inhibits sclareol export from plant roots, then I predicted that in the presence of FK506, plants normally resistant to sclareol would lose their mechanism of detoxification, and consequently growth would be inhibited.  The Arabidopsis seedling root is a good model for the study of a lignification (Dharmawardhana et al., 1992). When meristematic cells differentiate to xylem cells, the procambium cells elongate, build a thick secondary cell wall, and then deposit lignin into the cell wall. The process of xylogenesis is comparable to that in Zinnia tracheary elements and the basic differentiation is comparable to differentiation of lignifying cells in wood formation. Moreover, there are only a few lignifying tracheary elements at the very end of the vascular bundle in the root tip, and thus it is relatively easy to visualize the newly lignified tracheary element using florescence light microscopy, which takes advantage of the lignin autofluorescence. 4.1.1 Plant PM H + -ATPase is not inhibited with ABC transporter inhibitors Preliminary studies in the Samuels lab (Banno and Samuels, unpublished) tested whether inhibitors of ABC transporters would also inhibit the plasma membrane H+-ATPase. Barley roots were germinated in sand, grown without added potassium, and then transferred into hydroponic medium where the pH, due to H+-ATPase activity, could be monitored (Glass et al., 1981).  Upon addition of potassium, the pH of the solution surrounding the roots dropped, which could be expressed as H+ flux per gram of fresh weight of roots.  When the P-type ATPase inhibitor vanadate was added, flux was dramatically decreased as previously reported (Palmgren, 2001).  However, in contrast, the ABC transporter inhibitors Reversin 121 and Fumitremorgin C (Fig. 4.1) did not significantly change proton pump activity.  Similar results were found in preliminary experiments using FK506 (data not shown).  126    Figure 4.1 Average proton fluxes for potassium (K + ) starved barley roots when exogenous potassium was added. Average H+ fluxes were calculated and then normalized by root mass of the treatment. N=4 for the K+ and DMSO treatment, N=3 for both DMSO treatment as well as the K+ and vanadate treatment. N=2 for the K+ and Rev and K+and FTC treatments. Error bars, where present, represent standard deviations. (Brian Banno, Biol 448 report).  127  These data suggested that ABC transporter inhibitors, except vanadate, could be used in plants without destroying normal ion traffic mediated by the plasma membrane proton pump. The effects of ABC transporter inhibitors on lignification were tested in pine secondary xylem tissue using autoradiography (Dr. Kim Rensing, Samuels Lab unpublished data).  Both Reversin 121 and FTC reduced incorporation of 3H-phenylalanine by 60% -70% in developing xylem and ray parenchyma (Fig. 4.2). At high magnification, there did not appear to be a build up of incorporated 3H-phenylalanine in the cytosol of tracheids in pine, implying that reversin and FTC treatment reduced monolignol biosynthesis or phenylalanine uptake rather than inhibiting monolignol transport.  This is in contrast to Arabidopsis ABC transporter mutants such as wbc11 and cer5 where blocking the ABC transporter led to the accumulation of lipidic inclusions in the cytoplasm (Bird et al. 2008, Pighin et al. 2003).        128            Figure 4.2 Quantification of grey levels in light microscopy autoradiographs of pine developing secondary xylem. Dissected cambium and associated tissue samples were fed 3H-phenylalanine and ABC transporter inhibitors for 4 hours.  Two ABC transporter inhibitors were used. Reversin 121 (Rev), an ABCB/MDR subfamily specific inhibitor and FK506, an ABCG/PDR subfamily specific inhibitor, were fed with 3H-phenylalanine. Grey levels reflect intensity of radiolabel incorporation, where 0 is black and 1024 is white, were inverted and scaled to mean white background to give positive mean+SE values. Developing xylem indicates incorporation into axial tracheids while Ray Cytoplasm indicates incorporation into rays crossing from phloem to mature xylem (K. Rensing and L. Samuels, unpublished data).   129 The objective of this study was to use diverse lignifying cell types to test if ABC transporter inhibitors would block lignification. The secondary xylem system of poplar forms abundant monolignols in the lignification of the secondary cell wall (see Chapter 2). The dissected tissues were fed 3H-phenylalanine with inhibitors, and the effects of inhibitors were assessed on the basis of changes in radioactive phenylpropanoid localization at the light microscope level. If ABC transporters were involved in monolignol export, the label would be expected to increase in cytoplasm. Secondly, an Arabidopsis seedling root assay was used to test if lignification in the primary xylem of the root could be halted by application of ABC transporter inhibitors. Arabidopsis seedlings grown on agar plates were transferred to plates with inhibitors. As the roots grew, the vascular cylinder differentiated to keep pace with growth. The lignification process of tracheary elements in the root was a good system to screen the effects of inhibitors because an effective lignin autofluorescence protocol was available (Dharmawardhana et al., 1992). Because this is much faster than autoradiography, a wider variety of inhibitors could be tested. If ABC transporters were involved in the lignification process, lignin deposition in the root should be slowed or stopped. Timing of lignification during development was assayed by the presence of autofluorescence staining as the lignin content in the helical secondary cell wall of tracheary elements increased. This could be quantified as the distance from the tip of the root cap to the first lignified tracheary element.  Although this length includes the root cap and meristem, for convenience, it was named the “procambium length”. The treatment of ABC transporter inhibitors was predicted to decrease lignin export without changing development, leading to an increased procambium length.   130 4.2 Results In order to investigate whether ABC transporters are directly involved in the lignification process during secondary cell wall formation in poplar secondary xylem, two different ABC transporter inhibitors were used for autoradiography experiments with 3H-phenylalanine feeding, based on previous results obtained in the Samuels lab (Fig. 4.2).  For ABCB/MDR subfamily inhibitor, reversin 121 was used and for the ABCG/PDR subfamily inhibitor, FK506 was used.  Dissected cambium and associated developing xylem from poplar were incubated in 3H-phenylalanine solution with/without ABC transporter inhibitors followed by high pressure freezing, freeze substitution and autoradiography, as described in Chapter 2 and Kaneda et al., (2008).  Autoradiography was performed on light microscope sections and dark autoradiographic signals were quantified by measuring grey scale levels.  In the controls, 3H- phenylalanine was distributed heavily in the phloem, developing xylem and ray parenchyma (Fig. 4.3).  With reversin treatment, dark autoradiographic deposits on phloem and developing xylem were 40 to 50% more intense than the controls. However, the intracellular localization of decay events was very difficult to determine, making it difficult to discriminate between the secondary cell wall and cytoplasm at this level of resolution. Meanwhile with FK506, there was no significant changes of 3H-phenylalanine incorporation or decay distribution in any cell type, compared to controls.   131   Figure 4.3 Quantification of grey levels in light microscopy autoradiographs of poplar developing secondary xylem following ABC transporter inhibitor treatment. Dissected cambium and associated tissue samples were fed 3H-phenylalanine and various inhibitors for 4 hours.  Two ABC transporter inhibitors were used: Reversin 121 (Rev), an ABCB/MDR subfamily specific inhibitor and FK506, an ABCG/PDR subfamily specific inhibitor.  Inhibitors were fed together with 3H-phenylalanine, and then samples were cryofixed, sectioned and dipped for autoradiography. Dev. Xy; Axial lignifying cells of the developing xylem, Mat. Xy; axial vessels and fibers of mature xylem. Grey levels, where 0 is black and 1024 is white, were inverted and scaled to mean white background to give positive mean+SE values.  132 Another approach to test inhibitors was the Arabidopsis root assay.  To test if the root assay can detect changes in lignification, roots were treated with the cinnamate-4-hydroxylase inhibitor piperonylic acid (PA) (Schalk et al., 1998).  PA treated seedling roots continued to elongate normally (Fig. 4.5B).  However, the distance from the tip of the root cap to the first lignified tracheary element, the ‘procambium length’ was increased from 1.8 mm to 6 mm (Fig. 4.5A). The inhibition of phenylpropanoid pathway could potentially impair seedling growth because of the defective lignified cell wall in the vascular bundle but, at least in this short term growth assay, root growth was fully normal. Vanadate is know to be a P-type ATPase transporter inhibitor in plants, which inhibits ABC transporters and the plasma membrane H+-ATPases without influencing the function of H+-ATPases in the vacuole (Martinoia et al. 2002).  Vanadate treatment of Arabidopsis roots produced a mean procambium length that was 50% shorter than controls. That data indicated that there is an effect on lignification by inhibiting these transporters. However, root growth rate was severely compromised. Thus, vanadate had a strong impact on whole root development and impaired seedling growth. 1-N-naphthylphthalamic acid (NPA) is known to be auxin efflux inhibitor and it binds directly to several ABCB/MDR-type ABC transporters.  Auxin is directionally transported by ABCB/MDR subfamily proteins, and multiple ABCB/MDR members are NPA sensitive (Geisler and Murphy, 2006).  Auxin is required for normal vascular development (Mattson et al. 1999).  When Arabidopsis seedlings were grown on plates containing NPA and 1% sucrose, root growth was normal (Fig.4.4). With NPA, the length of the no vasculature zone near procambium increased as tracheary development stopped (Fig. 4.4). The procambium length of NPA treated seedling roots was comparable to that of PA treated seedling root (Fig. 4.5A).  In  133 contrast, high auxin treatment inhibited root growth, induced lateral root growth and had no affect on the procambium length (Fig. 4.4). On the other hand, four other subfamily-specific ABC transporter inhibitors, reversin 121, fumitremorgin C, FK506 and cyclosporin A did not show any effect on root growth or tracheary element lignification. After the treatment with inhibitors, seedlings still grew normally and showed lignification in tracheary elements typical of controls, suggesting that either ABC transporter proteins are not involved in direct lignin deposition or these inhibitors did not interrupt the target ABC transporters.  The procambium length in all of these treatments was the same as in the controls (Fig. 4.5).   134   Figure 4.4 The procambium length from the root tip to the first lignified TE in Arabidopsis seedling root is increased by auxin transport inhibition. NPA treatment inhibited tracheary element formation/lignification in Arabidopsis primary root. IAA treatment inhibited root elongation and induced lateral root growth. Mean+SE values   135  Figure 4.5  Procambium length after variety of inhibitor treatments in seedling roots (A) Five different ABC transporter inhibitors did not alter tracheary element lignification. Wild-type of Arabidopsis seedling were grown by 48 hours on medium containing different ABC transporter inhibitors. Treated seedling were fixed and stained in Basic Fuschin to see lignified tracheary elements. These measurements demonstrate that known ABC transporter inhibitors did not affect the lignification process in Arabidopsis primary xylem. Mean+SE values. (B) Growth rates of 7 days old Arabidopsis seedling roots were measured during the treatment period.  Only vanadate inhibited growth strongly, with FK506 giving a slight reduction in growth. Bars represent standard error. Black stars indicate differences were significant (p<0.05).  136 The lack of effect of putative ABC transporter inhibitors on lignification introduced the possibility that these inhibitors might not be effective on the plant ABC transporters.  Previous studies that demonstrated inhibitor effectiveness took advantage of the function of ABCBG/PDR subfamily members, which includes detoxification in higher plants where herbicides and fungicides are exported from roots (Van den Brule et al., 2002).  The ABCG/PDR specific inhibitor, FK506 was used to inhibit sclareol efflux from roots in barley. Barley was used in this assay since this was the system in which the sensitivity to the proton pump was tested using the barley root medium acidification assay of Glass et al. (1981). When sclareol was in the medium, barley seedling growth was identical to controls (Fig. 4.6), which suggested that ABCG/PDR proteins were able to pump out the toxic compound, sclareol, from the root. On the other hand, when the inhibitor FK506 was added, the growth rate of barley seedlings were reduced in a dose-dependent manner (Fig. 4.6).  These results are consistent with the findings of Campbell et al. (2003) who showed that sclareol was only toxic to Arabidopsis seedlings when pdr12 was knocked out.  Thus, it appears that, while it does not inhibit the proton pump, the inhibitor FK506 is effective against ABCG/PDR targets in plants and stops the export of the toxin, sclareol.  Thus the lack of inhibition of lignification by FK506 argues against a role for ABCG/PDR subfamily members as monolignol exporters.     137  Figure 4.6 FK506 inhibited sclareol detoxification in barley root. Whole length of barley seedlings was measured after 10 days of incubation in B5 medium. Sclareol did not effect growth of seedling and 5 uM FK506, but FK506 reduced 50% in 50 m and 60% in 100 m, suggesting that FK506 inhibited sclareol export from root cells.  138  4.3 Discussion Although the preliminary studies investigating lignification in pine indicated a sensitivity of lignin export to ABC transporter inhibitors, closer examination of the system revealed no accumulation of monolignols inside the cells upon transporter inhibition (unpublished data, Samuels lab).  In addition, reverse genetic studies in Arabidopsis failed to demonstrate any reduced lignin phenotypes for single gene knock-out mutants that might support a role for ABC transporters in lignification (Chapter 3). With the broader aim of inactivating suites of ABC transporters, inhibitors were employed to test if lignification would be altered upon treatment. However, as the data in this chapter shows, ABC transporter inhibitors did not block lignification in either the secondary xylem of poplar or the primary xylem of Arabidopsis. In poplar secondary xylem system, increasing radioactive incorporation in xylem parenchyma (or ray parenchyma cells) and developing xylem were observed by treatment with reversin.  Due to the large central vacuole occupying most of the volume in these cells, it was difficult to distinguish cytosolic or extracellular localization of the signal.  This is important because if lignification were blocked, then an intracellular signal would be expected to accumulate and the total signal could be higher than control.  Although cytoplasmic signal could not be definitively separated from cell wall signal, the thickened secondary cell walls did still appear to be lignified.  With FK506, lignification in Arabidopsis root tracheary elements and poplar secondary cell wall were identical to control.  In both cases, ABC transport inhibition does not appear to decrease lignification. It has been reported that multiple phenylpropanoid pathway enzymes such as PAL and 4CL are present in xylem ray parenchyma cells (Takabe et al. 2001) so an increased signal in this cell type after reversin or FK506 treatment would potentially be interesting if it were  139 accompanied by a decreased signal in the developing xylem.  However, this was not the case as incorporation into both axial lignifying cells and the rays was increased.  Tracheary element lignification continues after programmed cell death (PCD) in Zinnia culture system (Demura and Fukuda, 2007), suggesting that neighboring cells such as xylem parenchyma or ray cells can contribute monolignols to continue the lignification in xylary cells after PCD.  Due to lack of response to the ABC transporter inhibitors, data in this study did not verify if the ‘good neighbor’ hypothesis is valid in vivo.  The simplified system of Arabidopsis root tracheary element lignification allowed good access of applied inhibitors to cells due to the lack of a cuticle, and it provided a fast assay for lignification.  The assay was validated by the controls, but ABC transporter inhibitors did not produce any changes in lignin deposition. NPA treated Arabidopsis roots showed an enlarged procambium zone, indicating that NPA treatment interfered with normal tracheary element formation. Because auxin efflux carriers such as PGP19/ABCB19 and PGP1/ABCB1 and influx carrier PGP4/ABCB4 show high sensitivity to NPA (Noh et al., 2001, Murphy et al., 2002, Geisler et al. 2005, Terasaka et al., 2005), it is presumed that polar auxin transport would be disturbed by NPA treatment (Geisler and Murphy, 2006). Both auxin and cytokinin are required for normal expression of VND6 and VND7, which are xylem transcriptional factors known to induce differentiation into protoxylem and metaxylem (Kubo et al. 2005, Demura and Fukuda, 2007). The right auxin distribution established by auxin transporters is needed for xylem differentiation. With NPA treatment of Arabidopsis roots in this study, the larger zone of procambium length might be induced by procambium left behind as undifferentiated cells because of abnormal auxin distribution in plants by NPA, rather than direct inhibition of monolignol transport by ABCB/MDR proteins.  140  In conclusion, my data suggest that ABC transporter inhibitors did not affect lignification in primary and secondary xylem. The possibility of toxicity or secondary effects was minimized in the Arabidopsis roots by measuring growth and observing changes in auxin- related processes such as lateral root formation. The ABC transporter inhibitors, with the exception of reversin, have been shown in other studies to inhibit ABC transporters (Geisler et al., 2005; Lee et al., 2008). In the case of FK506, we could demonstrate that this inhibitor does work on presumed ABCG/PDR subfamily targets, suggesting that these are not responsible for lignification in the root. Thus, either ABC transporters are not involved in monolignol transport, or known ABC transporter inhibitors used did not inhibit the relevant plant ABC transporters. 4.4 Materials and methods 4.4.1 ABC transporter inhibitor treatment in poplar with autoradiography Active growing poplar tree stems (Populus trichocarpa x P. deltoides) were prepared as described in chapter 2 and Kaneda et al., (2008).  The dissected thin sections containing phloem, cambium and xylem were incubated in 3H-phenylalanine solution with ABC transporter inhibitors, reversin 121 (50 m) and FK506 (50 m) for 4 hours, then rinsed twice with 0.2M sucrose solution.  For autoradiography of incorporated 3H-phenylalanine distribution, tissues were cryofixed by HPF (High Pressure Freezing), and embedded in Spurr’s resin, then prepared as described in chapter 2 and Kaneda et al., (2008). 4.4.2 ABC transporter inhibitor treatment in Arabidopsis seedling root.  Plants were grown for 7 days at 21oC on AT agar (0.7%) plates, then transferred to plates containing ABC transporter inhibitors: vanadate (100 m), cyclosporin A (100 m),  141 fumitremorgin C (FTC, 50 m), tacrolimus (also known as FK506 or Fujimycin, 50 m), phenylpropanoid pathway inhibitors; piperonylic acid (1 m), or polar auxin transporter inhibitor; 1-N- naphthylphthalamic acid (NPA). After 48 hours treatment on AT plates, seedlings were fixed with 75% ethanol and stained by 0.001% Basic Fuchsin as lignin stain. Timing of lignification during development was assayed by the presence of autofluorescence as the lignin content in the helical secondary cell wall of tracheary elements increased, and this could be quantified as the distance from the tip of the root cap to the first lignified tracheary element.  Although this length includes the root cap and meristem, for convenience, it was named the “procambium length”.  Results are shown from 10-20 roots per treatment in four replicate experiments.  In addition, to testing toxicity of ABC transporters inhibitors for seedling development, newly grown root length of seedlings were measured after 48 hours of inhibitor treatment. 4.4.3 Barley growth test with inhibitors The barley seeds were surface-sterilized with 15% hypochlorite for 15 min and then rinsed 3 times with sterile distilled water and planted in a magenta box (15 x 15 x15 cm) on 30 ml of Gamborg’s B5 basal medium (Q-BioGene, USA) containing 0.7% (w/v) agar (Fisher).  The culture media contained antifungal diterpene sclareol (100 m) and ABCG/PDR subfamily inhibitor FK506 (5 m, 50 m or 100 m dissolved 0.1% DMSO). The culture boxes were sealed with plastic lid and incubated to grow under constant light at 21oC for 10 days. The heights of grown seedlings were then measured.  142  4.5 Bibliography  Al-Shawi MK, Senior AE. (1993) Characterization of the adenosine triphosphatase activity of Chinese hamster P-glycoprotein. J Biol Chem 268, 4197-4206. Bird D, Beisson F, Brigham A, Shin J, Greer S, Jetter R, Kunst L, Wu X, Yephremov A and Samuels L. (2007) Characterization of Arabidopsis ABCG11/WBC11, an ATP binding cassette (ABC) transporter that is required for cuticular lipid secretion. The Plant Journal 52, 485–498 Campbell EJ, Schenk PM, Kazan K, Penninckx IA, Anderson JP, Maclean DJ, Cammue BP, Ebert PR, Manners JM. (2003) Pathogen-responsive expression of a putative ATP- binding cassette transporter gene conferring resistance to the diterpenoid sclareol is regulated by multiple defense signaling pathways in Arabidopsis. Plant Physiol 133, 1272-1284. Cardenas ME, Cruz MC, Del Poeta M, Chung N, Perfect JR, Heitman J. (1999) Antifungal activities of antineoplastic agents: Saccharomyces cerevisiae as a model system to study drug action. Clin Microbiol Rev 12, 583-611. Dean M, Rzhetsky A, Allikmets R. (2001) The human ATP-binding cassette (ABC) transporter superfamily. Genome Res 11, 1156-1166. Demura T, Fukuda. (2007) Transcriptional regulation in wood formation. Trends in Plant Science 12, 65-70. Dharmawardhana DP, BE Ellis, JE Carlson (1992) Characterization of the vascular system of Arabidopsis.  Can J Bot 70, 2238-2244. Geisler M, JJ Blakeslee, R Bouchard, O Lee, V Vincenzetti, A Bandyopadhyay, B Titapiwatanakun, WA Peer, A Bailly, EL Richards, KFK Ejendal, AP Smith, C Baroux, U Grossniklaus, A Muller, CA Hrycyna, R Dudler, AS Murphy, and E Martinoia. (2005) Cellular efflux of auxin catalyzed by the Arabidopsis MDR/PGP transporter AtPGP1 Plant J 44, 179–194 Geisler M, Murphy AS. (2006) The ABC of auxin transport: the role of p-glycoproteins in plant development. FEBS Lett 580, 1094-1102. Glass AD, Siddiqi MY, Giles KI. (1981) Correlations between Potassium Uptake and Hydrogen Efflux in Barley Varieties : A potential screening method for the isolation of nutrient efficient lines. Plant Physiol 68, 457-459. Gottesman MM, Pastan I. (1993) Biochemistry of multidrug resistance mediated by the multidrug transporter. Annu Rev Biochem 62, 385-427. Jasinski M, Stukkens Y, Degand H, Purnelle B, Marchand-Brynaert J, Boutry M. (2001) A plant plasma membrane ATP binding cassette-type transporter is involved in antifungal terpenoid secretion. Plant Cell 13, 1095-1107. Kaneda M, Rensing KH, Wong JCT, Banno B, Mansfield SD, Samuels AL. (2008) Tracking monolignol during wood development in lodgepole pine. Plant Physiol. 147, 1-11. Klein M, Martinoia E, Hoffmann-Thoma G, Weissenbock G. (2001) The ABC-like vacuolar transporter for rye mesophyll flavone glucuronides is not species-specific. Phytochemistry 56, 153-159.  143 Kubo M, M Udagawa, N Nishikubo, G Horiguchi, M Yamaguchi, T Mimura, T Demura (2005) Transcription switches for protoxylem and metaxylem vessel formation Genes & Devel 19, 1855–1860. Lowes S, Simmons NL. (2002) Multiple pathways for fluoroquinolone secretion by human intestinal epithelial (Caco-2) cells. Br J Pharmacol 135, 1263-1275. Martinoia E, Klein M, Geisler M, Bovet L, Forestier C, Kolukisaoglu U, Muller-Rober B, Schulz B. (2002) Multifunctionality of plant ABC transporters--more than just detoxifiers. Planta 214, 345-355. Mattsson J, Sung ZR, Berluth T. (1999) Responses of plant vascular systems to auxin transport inhibition. Development 126, 2979-2991. Murphy AS, Hoogner KR, Peer WA, Taiz L. (2002) Identification, purification, and molecular cloning of N-1-naphthylphthalmic acid-binding plasma membrane-associated aminopeptidases from Arabidopsis. Plant Physiol 128, 935-950. Noh B, Murphy A, Spalding EP. (2001) Multidrug resistance-like genes of Arabidopsis required for auxin transport and auxin-mediated development. Plant Cell 13, 2441- 2454. Okuhara M, Kohsaka M, Aoki H, et al. (1987) FK-506, a novel immunosuppressant isolated from a Streptomyces. II. Immunosuppressive effect of FK-506 in vitro. J Antibiot (Tokyo) 40, 1256-1265. Ozvegy C, Varadi A, Sarkadi B. (2002) Characterization of drug transport, ATP hydrolysis, and nucleotide trapping by the human ABCG2 multidrug transporter. Modulation of substrate specificity by a point mutation. J Biol Chem 277, 47980-47990. Palmgren MG. (2001) Plant plasma membrane H+-ATPases: Powerhouses for Nutrient Uptake. Annu Rev Plant Physiol Plant Mol Biol 52, 817-845. Pighin J, H Zheng, LJ Balakshin, IP Goodman, TL Western, R Jetter, L Kunst, AL Samuels. (2001) Plant Cuticular Lipid Export Requires an ABC Transporter. Science 306, 702. Robey RW, Medina-Perez WY, Nishiyama K, Lahusen T, Miyake K, Litman T, Senderowicz AM, Ross DD, Bates SE. (2001) Overexpression of the ATP-binding cassette half- transporter, ABCG2 (Mxr/BCrp/ABCP1), in flavopiridol-resistant human breast cancer cells. Clin Cancer Res 7, 145-152. Schalk M., F Cabello-Hurtado, M-A Pierrel, R Atanossova, P. Saindrenan, D Werck- Reichhart (1998) Piperonylic Acid, a Selective, Mechanism-Based Inactivator of the trans-Cinnamate 4-Hydroxylase: A New Tool to Control the Flux of Metabolites in the Phenylpropanoid Pathway. Plant Physiol 118, 209-218. Shitan N, Bazin I, Dan K, Obata K, Kigawa K, Ueda K, Sato F, Forestier C, Yazaki K. (2003) Involvement of CjMDR1, a plant multidrug-resistance-type ATP-binding cassette protein, in alkaloid transport in Coptis japonica. Proc Natl Acad Sci USA 100, 751- 756. Takabe K, Takeuchi M, Sato T, Ito M, Fujita M. (2001) Immunocytochemical localization of enzymes involved in lignification of the cell wall. J Plant Res114, 509-515. Terasaka K, Shitan N, Sato F, Maniwa F, Ueda K, Yazaki K. (2003) Application of vanadate- induced nucleotide trapping to plant cells for detection of ABC proteins. Plant Cell Physiol 44, 198-200. Terasaka K, Blakeslee JJ, Titapiwatanakun B, Peer WA, Bandyopadhyay A, Makam SN, Lee OR, Richards EL, Murphy AS, Sato F, Yazaki K. (2005) PGP4, an ATP binding  144 cassette P-glycoprotein, catalyzes auxin transport in Arabidopsis thaliana roots. Plant Cell 17, 2922-2939. van den Brule S, Muller A, Fleming AJ, Smart CC. (2002) The ABC transporter SpTUR2 confers resistance to the antifungal diterpene sclareol. Plant J 30, 649-662  145                    CHAPTER 5:  CELL SPECIFIC DEPOSITION OF HEMICELLULOSE IN DEVELOPING SECONDARY CELL WALL OF HYBRID POPLAR (POPULUS DELTOIDES X P. TRICHOCARPA) VESSELS AND FIBERS 1               1. A version of a manuscript that will be published M. Kaneda and A.L. Samuels. Cell specific deposition of hemicellulose in developing secondary cell wall of hybrid poplar (Populus deltoides x P. trichocarpa) vessels and fibers  146 CHAPTER 5: CELL SPECIFIC DEPOSITION OF HEMICELLULOSE IN DEVELOPING SECONDARY CELL WALL OF HYBRID POPLAR (POPULUS DELTOIDES X P. TRICHOCARPA) VESSELS AND FIBERS  5.1 Introduction Poplar has ecological and economical value, and its diverse cell types also provide substantial material to study wood formation.  During secondary xylem differentiation, tangential and longitudinal cell division in cambial initial cells produce xylem daughter cells that undergo cell differentiation: cell elongation, secondary cell wall deposition, programmed cell death and lignification (Larsson, 1994).  When developing vessels and fibers reach their mature length and size, cell elongation stops and a thick secondary cell wall is deposited, consisting of three major components.  The most abundant component, cellulose, is assembled at plasma membrane by the cellulose synthase complex where parallel cellulose microfibrils are systematically accumulated inside of the primary cell wall to form layers. The secondary cell wall is normally made of three layers called S1, S2 and S3 layers defined by different angles of cellulose microfibrils relative to the cell length axis (Donaldson, 2001).  The second component is hemicellulose, which helps to aggregate cellulose microfibrils by providing cross-links between cellulose and hemicellulose via hydrogen bonds.  During secondary cell wall formation, a cellulose microfibril- hemicellulose matrix is assembled before lignification and provides the environment for monolignols to polymerize into lignin (Awano et al., 2002).  When monolignols polymerize, polysaccharides bond to lignin polymers, by both covalent and non-covalent interactions to form a lignin-polysaccharide complex (Houtman and Atalla, 1995).  Thus, the hemicellulose-  147 lignin connection is a key to organizing the secondary cell wall (Atala et al. 1993; Donaldson. 2001).  The cell types in poplar wood are more diverse in function and shape than in gymnosperm wood.  Vessel elements, wood fibers and axial parenchyma cells all differentiate from one meristematic tissue, the cambium.  Tracheids and vessel elements conduct water and ions, while fibers provide support for the plant.  Differences in function and development of these lignified cell types might require diverse hemicelluloses in the secondary cell wall, or they might use the same hemicelluloses with differences in timing of deposition.  The predominant hemicelluloses in woody angiosperms secondary cell walls are glucuronoxylans (Ebringerova and Heinze, 2000).  Monoclonal antibodies, LM10 and LM11, antibodies raised against a synthetic pentose xylan conjugated to BSA can recognize glucuronoxylans from beech wood (McCartney et al. 2005, Knox et al. 2005).  These antibodies were used to localize xylan in metaxylem and protoxylem secondary cell walls in wild-type Arabidopsis inflorescence stems (Persson et al. 2007).  The importance of this xylan for secondary cell wall production in Arabidopsis was demonstrated in the irregular xylem8 (irx8) mutant that showed significant reduction of xylan and collapsed xylem with uneven cell wall deposition in stem vascular bundles and interfascicular fibers (Persson et al. 2007).  Xylan distribution in the secondary cell wall of Fagus crenata was studied with field emission SEM and increasing accumulation of xylan was seen as secondary cell wall deposition progressed (Awano et al. 2002). Another hemicellulose, mannan, is known to be abundant in the lignified secondary cell walls of gymnosperms.  Mannan epitopes were localized in thickened secondary cell walls of xylem elements, xylem parenchyma and interfascicular fiber in Arabidopsis inflorescence stem  148 (Handford et al. 2003).  Monoclonal antibodies raised against galactomannan were tested in this study for binding to poplar secondary xylem walls (Pettolino et al. 2001).  The goal of this chapter was to define the cellular structures involved in xylogenesis, specifically hemicellulose secretion in poplar vessels and fibers.  In the course of the autoradiographic studies of developing wood (Kaneda et al. 2008) and published studies of gymnosperm wood prepared with cryofixation  (Inomata et al. 1992; Samuels et al., 2002), it became clear that the quality of organelle structure preservation in cryofixed developing wood was very high.  In contrast, published reports of cell structure in developing poplar fibers and vessels relied on chemical fixation and demonstrated poor preservation (Arend and Fromm, 2003).  To date, hemicellulose synthesis, secretion and distribution in poplar secondary xylem cell types have not been studied. Therefore, I used immuno-label light and electron microscopy on cryofixed poplar wood, to place the distribution of xylan and mannan in different cell types, and within the secretory apparatus of a cell, in a spatial /structural context. 5.2 Results 5.2.1 Secondary xylem and ultrastructure during xylogenesis Understanding cell structure and morphological changes during secondary xylem development is important because the cells undergo a dramatic transformation during their development and they are responsible for the production of all secondary cell wall materials. However, observation of cell structure at the electron microscopic level during xylogenesis is exceptionally difficult because xylem cells are highly sensitive to conventional chemical fixatives. To determine important cell structural information in poplar xylogenesis, cryofixation using high pressure freezing (HPF) and freeze substitution were used. This rapid freezing  149 technique allows high quality preservation of ultrastructure, especially the endomembrane systems.  The ultrastructure of cambium and developing xylem cells were observed using TEM. In cambial cells, mitotic cells undergo cell plate deposition to form daughter cells.  The forming cell plate was observed being laid down between daughter cells, in association with the phragmoplast and multiple Golgi (Fig. 5.1A).  The phragmoplast, rich in microtubules and microfilaments, excludes the bulkier organelles such as mitochondria to its periphery.  As reported for pine (Bailey, 1919, Rensing et al., 2002), the phragmoplast of the secondary vascular system is unlike the hoop-like phragmoplast of dividing cells in primary meristems of higher plant cells.  It is contained in a bolus of cytoplasm that migrates the length of the cambial cell and two opposing boli move from the newly divided nuclei to the distal ends of the cells.  Xylem mother cells, adjacent to the cambium, had a large central vacuole and cortical cytoplasm (Fig.5.1B), in contrast to the newly divided ray cells in which several smaller vacuoles surround the nucleus (Fig. 5.1C). In cells actively depositing secondary cell wall, the cell structure again was dominated by the large central vacuole with the cytoplasm restricted to the cell ends and cortex.  Cortical microtubules were abundant in these cells and aligned with cellulose microfibril orientation (Fig. 5.1D).  150   Figure 5.1 Ultrastructure during xylogenesis in poplar A) A cambial cell undergoing cell division with a long vertical cell plate (black arrows). B) Newly developed cambial cells with very thin cell wall and large central vacuoles with many Golgi. C) Ray parenchyma cells with a large central nucleus and dense cytoplasm. D) and E) Developing fiber cells with many Golgi with multiple vesicles associated with cortical microtubules. Large cisternae  and reduced trans Golgi network (TGN) with tubular cluster shown (arrowheads). F) Mature vessel associated with vitally active ray parenchyma cells. The black lines on cell wall  151 During the differentiation of xylem from cambium into fibers and vessels, the Golgi structure changed.  In the cambium, the Golgi morphology was typical of plant parenchyma cells with 5-6 cisternae and a trans-Golgi network with clusters of vesicles (Fig. 5.2 A,B).  In contrast, the Golgi stacks in developing xylem cells were unusual in that they contained many cisternae, often associated with a large number of small vesicles (Fig. 5.2 C-O). Tube like structures, probably a part of the trans-Golgi network, were seen but the grape-like clusters of Golgi vesicles that were found in pine (Samuels et al. 2002) were not observed. Vesicles could be observed fusing with plasma membrane, forming “slit-like or horseshoe shaped structures” (Fig. 5.1D arrows), which have been reported in cryofixed pine developing tracheids (Samuels et al. 2002) and carrot and sycamore-maple suspension cultured cells (Staehelin and Chapman 1987). These prominent structures of Golgi and vesicle clusters in secondary xylem were correlated with secretion of cell wall as the secondary cell wall increased in thickness as distance from the cambium increased (Fig. 5.2E).   152   Figure 5.2 Golgi morphology during xylogenesis in poplar xylem cells. A) and B) TEM of the Golgi of cambial cells. Multiple Golgi with typical anatomy of cis/trans polarity. C) to O). Golgi from developing xylem cells showing characteristic Golgi with multiple middle stacks and less or indistinct TGN with small vesicles. CW; cell wall. Bar = 500 nm.  153 5.2.2 Xylan and mannan distribution in poplar secondary xylem An overview of poplar secondary xylem development was obtained using light microscopy sections stained with toluidine blue (Fig. 5.3A), which showed the developmental gradient from cambium to mature xylem.  In such transverse sections, it is clear that vessels showed rapid radial expansion during their development, followed by secondary wall deposition. Fibers showed slower, more gradual radial expansion and in longitudinal sections (not shown), they displayed axial elongative growth. Xylan distribution in poplar secondary xylem was tested using the anti-xylan antibody LM10 (McCartney et al. 2005).  LM10 bound to the tissue in a general distribution on fibers, vessels and ray parenchyma cells, demonstrated by secondary antibody Alexa 594 label on tangential xylem samples (Fig. 5.3B-D).  The secondary cell walls, but not the middle lamella, of fibers and vessels (Fig. 5.3B,C) as well as mature radial tracheids (Fig. 5.3D) were evenly labeled.  Controls, which were not incubated with primary antibody, but instead incubated in buffer then secondary antibody, did not have significant fluorescent signals in the red spectrum used to detect the Alexa fluorochrome. Mannan distribution was studied using monoclonal anti-glaucomannan primary antibodies (Pettolino, 2001), detected with secondary antibody-Alexa 594.  Anti-mannan label was strongly enriched in fiber cell walls, especially the S2 layer (Fig. 5.4), suggesting that vessel, fiber and ray have different hemicellulose compositions. In addition, anti-mannan binding to ray tracheids increased late in poplar xylem development (data not shown), which demonstrates that ray hemicellulose deposition is dramatically delayed compared to axial xylem development.  In no-primary antibody controls, sections did not have significant fluorescent signal in the red spectrum used to detect the Alexa fluorochrome.  154  Figure 5.3 Poplar secondary xylem and immunolabeling of LM10 (xylan) localization A) Light micrograph of radial sections of inner bark of hybrid poplar (Populus deltoides x P. trichocarpa) showing all cell types in poplar secondary xylem from cambium (Cam)(Left) to developing xylem (Dev.Xy) and mature xylem (Mat.Xy). V: vessel, F: fiber. Bar is 50 m. B), C) and D) Monoclonal antibody LM10 was used to detect poplar xylan distribution in secondary xylem. Secondary antibody label was linked to Alexa 594. V: vessel, F: fiber. Bar is 50 m  155  Figure 5.4 Immunolabeling of anti-mannan in poplar secondary xylem Poplar secondary xylem showing a strong mannan immunolabeling in fiber secondary cell wall. Relatively lower label in vessel and ray secondary cell wall. Monoclonal rat anti-mannan antibody was used to localize xylan and anti-rat conjugated to Alexa 594 was used as secondary antibody. Bars = 50 m.   156 To study the secretion of hemicellulose during secondary cell wall deposition at the subcellular level, samples were prepared for TEM and the grids incubated with monoclonal ß-1-4-mannan antibody and colloidal gold-conjugated secondary antibody.  Gold labeled the secondary cell walls of fibers but little label was detected on the walls of rays or vessels (Fig. 5.5A,B), supporting the immuno-fluorescence results.  In the cytoplasm of developing fibers, gold particles were detected on the periphery of the Golgi stacks and associated with vesicles (Fig. 5.5C).  157   Figure 5.5 Anti-mannan immunogold labeling in poplar developing xylem. A) Labeling periphery of Golgi (G) or Golgi associated vesicles. B) Profuse labeling in fiber secondary cell wall, especially S2 layer (Arrows) but less labeling in vessel secondary cell wall. C) Abundant labeling in fiber cell wall but less in ray parenchyma secondary cell wall.   158  5.3 Discussion The cryofixation and freeze substitution method for preservation of poplar allowed a remarkable high resolution ultrastructural examination by electron microscopy. In developing secondary xylem of poplar following cryo-fixation, cell structure did not show signs of osmotic disturbance as seen in conventional preparations.  Here the cells showed intact organelles, smooth membranes on the Golgi and tonoplasts, and no infoldings of plasma membrane. In contrast, extensive infoldings of plasma membrane penetrating into cytosol were seen in poplar developing xylem prepared by conventional chemical fixation (Arend and Fromm, 2003).  Unlike the secretion of monolignols, the involvement of Golgi apparatus for the production and secretion of hemicelluloses in secondary cell wall was supported. Novel Golgi morphology during xylogenesis was observed in poplar, in particularly multiple medial cisternae in the Golgi and a large number of small vesicles at the periphery of the Golgi apparatus. Further work is necessary to characterize the phenomenon of multiple medial cisternae in the Golgi and to define the cargo of Golgi mediated vesicles.  Given that these Golgi appear to have an unusual structure, the arrangement of glycosyl transferases producing the hemicelluloses in poplar might be different or the way that the polysaccharide product is packaged in cisternae compared to vesicles could be different. Golgi structure observed was unlike other secretory polysaccharide systems such as pine where TGN were made up of grape- like clusters (Samuels et al. 2002) or seed coat cells where the TGN consists of interconnected vesicular clusters (Young et al. 2008).  In poplar secondary xylem, changes in cell structure were correlated with the production of the secondary cell walls during development.  As the secondary cell wall thickened, the cells contained many Golgi stacks with cisternae and hemicelluloses were  159 deposited.  Xylan epitopes showed a general distribution in developing xylem using monoclonal anti-xylan LM10.  That data agreed with studies of secondary xylem in tobacco, showing that LM10 localized to all cell types in the secondary cell walls in secondary xylem (McCartney et al. 2005). Xylan epitopes were distributed abundantly in secondary cell wall in Arabidopsis inflorescence stems; IRX8 protein is know to strongly influence the production of xylan and secondary cell wall formation itself (Pena et al., 2007). Overall, the localization of xylan in poplar supported that xylan is a major, general hemicellulose in secondary cell walls of diverse cell types.  In contrast, the distribution of mannan epitopes in poplar xylem showed cell type specific localization.  Mannan epitopes were enriched in the fiber secondary cell walls, especially the S2 layer, but at low levels in vessel element cell walls and rays.  In Arabidopsis, mannans are found in secondary cell walls (Handford et al. 2003).  However, the authors mentioned that xylem vessels consistently showed lower labeling of mannan epitopes than fibers and xylem parenchyma cells (Handford et al. 2003). These data are consistent with fiber specific mannan localization in poplar xylem. The basic structure of secondary cell walls in vessel elements and fibers were similar but distinct. Both have abundant cellulose and lignin in thick secondary cell wall, however; lignin composition in the two cell types is different. Vessel cell walls show high guaiacyl (G) lignin content, while fiber cell walls contain more syringyl (S) lignin (Fukushima and Terashima, 1990). The differences of hemicellulose matrix in secondary cell walls might be linked to the differences in lignin content, e.g. hemicellulose could help to organize selective monolignol binding.  Atalla discussed the influence of the hemicellulose–cellulose matrix, which provides a link between lignin and the polysaccharide matrix, with hemicelluloses in particular playing a central role in selecting lignin precursors  160 (reviewed in Atalla, 2005). This histo- and cytochemical study established that mannans and xylans occur in different patterns in poplar wood.  Xylans are generally found in all secondary cell wall domain while mannans appear in discrete domains. It is possible that hemicellulose may play a role in cell wall construction, defining the cell wall properties and allowing cell type specific lignin deposition. 5.4 Materials and methods 5.4.1 Plant materials and growth conditions  Hybrid poplar; Populus deltoides x P. trichocarpa (H11-11) were grown in pots (15x15x25 cm) in the green house after whips were propagated from a large poplar tree (see chapter 2). The active poplar stems were dissected for cryofixation, freeze substitution and Spurr resin embedding for ultrastructural analysis as described in chapter 2 and Kaneda et al. (2008). 5.4.2 Immunofluorescent localization in xylem tissue by LM10 antibody and anti mannan antibody. For immunoflorescent labeling, cryofixed samples were fixed and substituted with 0.25% glutaraldehyde in DMP/acetone, then infiltrated and embedded in LR-white resin.  Teflon coated multiwell slides (EMS cat#63424-06) were coated with poly-L-lysine and left to dry. Sections (300, 500 nm) were placed on a drop of water in each slide well and air dried over night in a 37oC incubator. Slides were placed in a coplin jar filled with 5% non-fat milk blocking buffer with TBST for 20 min. After washing for 10 min with TBST, sections were incubated with 1/100 concentration anti- (1-4)-beta-D-mannan and galacto-(1-4)-beta-D- mannan (catalogue #400-4) monoclonal antibody (Biosupplies Australia Pty Ltd) or anti-xylan LM10 antibody (kind gift of Dr. J. Paul Knox: www.plantprobes.co.uk) for 1 hour, followed by  161 1/100 secondary antibody-Alexa 543 for 1 hour. Fluorescent localization was observed by Leica light microscope (Leica, DRM) using a Texas Red filter.  5.4.3 Immuno-gold labeling for xylem tissue by monoclonal mannan antibody.  LR-White or Spurr resin blocks were cut into 60 to 80 nm sections and mounted on formvar coated nickel grids. Before immuno-gold labeling, Spurr sections were treated in 10% H2O2 solution for 20 min. for etching reaction, and then rinsed with ddH2O. For blocking non specific protein binding, grids were incubated with 5% (w/v) bovine serum albumin (BSA) (Sigma) in TBST for 20 min. Grids were floated on a drop of primary antibody of anti-mannan with antibody solution (1% BSA in TEST) for 1 hour at room temperature. After washing in TBST, grids were transfer to secondary antibodies (1:100 dilution in antibody solution). Secondary antibodies were conjugated to 15 nm colloidal gold (Ted Pella) with goat anti- mouse IgG + IgM for anti-mannan and goat anti-rat for LM10. Following washing in TBST and ddH2O, sections were poststained with 2% (w/v) uranyl acetate for 15 min and Reynold’s lead citrate for 5 min.  162  5.5 Bibliography Arend M., Fromm J. (2003) ultrastructural changes in cambial cell derivatives during xylem differentiation in poplar. Plant Biol 5, 255–264 Atalla, RH. (2005) The role of the hemicelluloses in the nanobiology of wood cell walls : a systems theoretic perspective. Proceedings of the Hemicelluloses Workshop 2005 : 2005 January 10-12, The Wood Technology Research Centre, University  of Canterbury, Christchurch, NZ. Christchurch, NZ : Wood Technology Research Centre, University of Canterbury, 2005: Pages 37-57. Awano T., Takabe K, Fujita M. (2002) Xylan deposition on secondary wall of Fagus crenata fiber. Protoplasma 219, 106-115. Bailey, I.W. (1919) Phenomena of cell division in the cambium of arborescent gymnosperms and their cytological significance. Proc. Natl. Acad. Sci USA. 5, 283-285. Esau K. (1965) Plant anatomy. Wiley, New York Ebringerová, A., and Heinze, T. (2000). Xylan and xylan derivatives—Biopolymers with valuable properties. I. Naturally occurring xylans structures, isolation procedures and properties. Macromol. Rapid Commun. 21, 542–556. Handford MG., Baldwin TC, Goubet F, Prime TA, Miles J, Yu X, Dupree P. (2003) Localisation and characterisation of cell wall mannan polysaccharides in Arabidopsis thaliana. Planta 218, 27-36. Houtman C. and RH Atalla (1995) Cellulose-Lignin Inderactions Plant Physiol 107:977-984 Inomata F, Takabe K, Saiki H. (1992) Cell wall formation of conifer tracheid as revealed by rapid-freeze and substitution method. J. Electron Microsc 41, 369-374. Kaneda M., Rensing KH, Wong JCT, Banno B, Mansfield SD, Samuels AL. (2008) Tracking monolignols during wood development in lodgepole pine. Plant  Physiol. 147, 1-11. Larsson, P.R. (1994) The Vascular Cambium: Development and Structure. Berlin Heidelberg. Springer. McCartney L., Blake AW, Flint J, Bolam DN, Boraston AB, Gilbert HJ, Knox JP. (2006) Differential recognition of plant cell walls by microbial xylan-specific carbohydrate- binding modules. Proc Natl Acad Sci U S A 103, 4765-4770. Moore PJ., Swords KMM, Lynch MA, Staehelin LA. (1991. Spatial organization of the assembly pathways of glycoproteins and complex polysaccharides in the Golgi apparatus of plants. J Cell Biol 112, 589-602. Persson S., Caffall KH, Freshour G, Hilley MT, Bauer S, Poindexter P, Hahn MG, Mohnen D, Somerville C. (2007) The Arabidopsis irregular xylem8 mutant is deficient in glucuronoxylan and homogalacturonan, which are essential for secondary cell wall integrity. Plant Cell 19, 237-255. Pettolino, FA., Hoogenraad, NJ, Ferguson, C, Bacic, A, Johnson, E, Stone, BA (2001). A (14)-beta-mannan-specific monoclonal antibody and its use in the immunocytochemical location of galactomannans. Planta 214, 235-42. Rensing, KH., Samuels, A.L., and Savidge, R.A. 2002. Ultrastructure of vascular cambial cell cytokinesis in pine seedlings preserved by cryofixation and substitution. Protoplasma 220,39-49.  163 Samuels AL., Rensing KH, Douglas CJ, Mansfield SD, Dharmawardhana DP, Ellis BE. (2002) Cellular machinery of wood production: differentiation of secondary xylem in Pinus contorta var. latifolia. Planta 216, 72-82. Staehelin LA and RL Chapman (1987) Secretion and membrane recycling in plant cells: novel intermediary structures visualized in ultrarapidly frozen sycamore and carrot suspension-culture cells Planta 171:43-57 Takabe K., Takeuchi M, Sato T, Ito M, Fujita M. (2001) Immunocytochemical localization of enzymes involved in lignification of the cell wall. J Plant Research 114, 509-515. Taylor NG., Laurie S, Turner SR. (2000) Multiple cellulose synthase catalytic subunits are required for cellulose synthesis in Arabidopsis. Plant Cell 12, 2529-2540. Turner S., Gallois P, Brown D. (2007) Tracheary Element Differentiation. Annu Rev Plant Biol 58, 407-433. Young RE., McFarlane HE, Hahn MG, Western TL, Haughn GW, Samuels AL. (2008) Analysis of the Golgi apparatus in Arabidopsis seed coat cells during polarized secretion of pectin-rich mucilage. Plant Cell 20, 1623-1638. Zhong R., Pena MJ, Zhou GK, Nairn CJ, Wood-Jones A, Richardson EA, Morrison WH, 3rd, Darvill AG, York WS, Ye ZH. (2005) Arabidopsis fragile fiber8, which encodes a putative glucuronyltransferase, is essential for normal secondary wall synthesis. Plant Cell 17, 3390-3408. Zhang GF., Staehelin LA. (1992) Functional compartementation of the Golgi apparatus of plant cells. Plant Physiol. 99, 1070-1083. Zhang GF., Driouich A, Staehelin LA. (1993) Effect of monensin on plant Golgi: re- examination of the monensin-induced changes in cisternal architecture and functional activities of the Golgi apparatus of sycamore suspension-cultured cells. J Cell Sci 104, 819-831.  Zhong R., Pena MJ, Zhou GK, Nairn CJ, Wood-Jones A, Richardson EA, Morrison WH, 3rd, Darvill AG, York WS, Ye ZH (2005) Arabidopsis fragile fiber8, which encodes a putative glucuronyltransferase, is essential for normal secondary wall synthesis. Plant Cell 17, 3390-3408.   164            CHAPTER 6:  CONCLUSION AND FUTURE DIRECTIONS  165 CHAPTER 6: CONCLUSION AND FUTURE DIRECTIONS 6.1 Summary of the results and conclusion This thesis examines the secretion of the secondary cell walls, a lignin-rich thickened cell wall that is important for plant function.  Secondary cell walls are formed in fibers and vessel elements of angiosperms such as poplar or tracheids of gymnosperms such as pine and are essential for plant strength and rigidity. The secondary cell wall is formed by systematic construction of cellulose, hemicellulose and lignin in a strictly regulated fashion during xylogenesis, requiring active secretion and dynamic cytoskeleton activity. However, in order to study this process at the ultrastructural level, good preservation of cell structure in protoplasts secreting secondary cell walls is required.  In the past, it has proven difficult to preserve plant cells during secondary cell wall secretion using conventional experimental approaches. In my Ph.D. thesis studies, I had the goal of getting accurate information on the endomembrane system in developing lignifying cells, in order to study the mechanisms of lignin precursor transport and hemicellulose secretion during wall deposition.  I was able to meet this goal by using cryofixation to preserve the lignifying cells and their contents.  Cryofixation allowed high quality observations of well preserved ultrastructure in poplar lignifying cells. In lignifying cells of poplar, intact organelles, well preserved Golgi, tonoplasts, endomembrane system and smooth plasma membrane were seen by TEM.  Unique Golgi morphology during secondary cell wall deposition was observed in poplar such as multiple medial cisternae in the Golgi and a large number of small vesicles at the periphery of Golgi apparatus. Golgi apparatus are involved in for the production and secretion of hemicelluloses in secondary cell wall. The arrangement of glycosyl transferases producing the  166 hemicelluloses in poplar might be different or the way that the polysaccharide product is packaged in cisternae compared to vesicles could be different.  Providing regulated hemicellulose-cellulose matrix structure for lignin accumulation is fundamental in secondary cell walls. Hemicelluloses distribution and their mechanisms of secretion are important for understanding the environment in which monolignols polymerize. Using cryofixed cells, I was able to confirm the Golgi-mediated secretion of hemicelluloses into the secondary cell wall.  Using immunolabeling, the xylan distribution was demonstrated throughout secondary cell walls in vessel elements, fibers and matured rays. On the other hand, mannan showed fiber-specific localization but not in the ray and vessel walls.  This hemicellulose distribution supported my hypothesis that different cell types have different hemicelluloses, in this case varying amounts of mannan: xylan.  This finding is consistent with the idea that different hemicellulose components may play a role in differential lignin construction during secondary cell wall formation. Since there are areas enriched in S-lignin or G-lignin in wood, beyond the random radial coupling reactions, Atalla hypothesized that hemicelluloses play a central role in selective monolignol linking by providing an interactive environment (Atalla, 2005). For example, it may be speculated that the mannans in the fibers help to organize the deposition of S-lignin in this cell type.  Following hemicellulose deposition in lignified cells, monolignols are secreted from the protoplast to the apoplast.  In chapter 2, the currently accepted model of monolignol transport, that monolignols are secreted from the cell by Golgi-mediated vesicle transport, was re-examined by investigating the subcellular distribution of monolignols in developing xylem.  Derivatives of 3H-phenylalanine were analyzed using HPLC and scintillation counting and, somewhat surprisingly, coniferin was not strongly radioactive in our pine  167 samples although monolignols were. The radiolabeled phenylpropanoids did not accumulate inside developing tracheids, but were quickly deposited in the lignifying secondary cell wall. Within the cell, cytoplasm and Golgi were labeled following treatment with 3H-phenylalanine. Using a combination of inhibitors of phenylpropanoid metabolism or protein synthesis, the Golgi signal was revealed to be protein, rather than monolignol, in nature.  These data are more consistent with a transporter-mediated export of monolignols, rather than a Golgi- mediated export hypothesis. The new approach used in this study was the combination of data from both the biochemical assay and autoradiographic quantification analysis that provided evidence that the compounds of interest were labeled and localized with high fidelity. 3H-compounds in autoradiography were identified as lignin precursors by HPLC-MS and lignin polymer by thioacidolysis analysis. The reason I used 3H-phenylalanine, rather than compounds more down stream of monolignol biosynthesis, as a 3H-lignin precursor was that downstream compounds were inefficiently incorporated compared to phenylalanine (Anterola et al., 2002). Cinnamic acid inhibits PAL activity, which leads to decreased flow into the whole phenylpropanoid pathway (Blount et al. 2000), which would have led to potential problems of reduced lignin production or even less uptake of 3H-lignin precursors during the feeding experiment. On the other hand, adding phenylalanine solution induced p-coumaryl alcohol and coniferyl alcohol production, and PAL, 4CL, CCoOMT and CCR transcript levels (Anterola et al. 2002). The effect of tissue dissection from the trees and incubation in solution for hours could have been a problem in this assay, however; electron microscopy confirmed that dissected tissues were still intact and cell structure was not disrupted in endomembrane systems such as Golgi apparatus, ER, vesicles and plasma membrane. In conclusion, this  168 study demonstrated that monolignol export was not mediated by the Golgi in pine tracheids during lignification. Excluding a Golgi-mediated export mechanism of monolignols, then how are monolignols leaving the cytoplasm of cells during lignification? The first possibility is diffusion, driven by the force of a chemical gradient.  In this case, polymerization could remove monomers from solution, generating the concentration gradient.  Without knowing what the transport substrate really is, it is hard to judge whether diffusion would be operating, e.g. it could make a difference if the export substrate was coniferyl alcohol or the glucoside. Studies of model membranes using liposomes or lipid bilayer disks demonstrated partitioning of model phenolic substrates into model liposomes or lipid disks (Boija and Johansson, 2006; Boija et al., 2007).  The authors did not explain why they did not use the monolignol compounds but instead used related compounds like alpha-methylsyringyl alcohol. It is not clear how active desorption of these compounds from the hydrophobic environment of the membrane into the apoplast would occur in this model. A possible alternative mechanism is membrane transport proteins moving monolignols across the plasma membrane as seen in the movement of molecules such as amino acids and ions. These transporters may assist transport of a specific substrate by non-energy requiring facilitated diffusion or move compounds across a concentration gradient using ATP dependent transports, which use energy from ATP provided from H+-gradient. In chapter 3, a possible role of active transporters in monolignol export was tested. ATP-binding cassette (ABC) transporters represent the largest transporter superfamily in plants so it was necessary to restrict the scope of this study. Previous gene expression studies demonstrated a correlation between expression of phenylpropanoid biosynthetic genes and a  169 subset of genes encoding ABC transporters, especially in the ABCB/Multi-drug resistance (MDR) and ABCG/Pleiotropic drug resistance (PDR) subfamilies (ABCB11, ABCB14, ABCB15, and ABCG33). Homozygous T-DNA insertional single mutant lines (abcb11; abcb14; abcb15, and abcg33) showed no apparent alterations in xylem and interfascicular fiber lignification and morphology in comparison to wild-type.  However, in abcb14 null mutants, vascular morphology was disorganized, with decreased phloem area in the vascular bundle and decreased xylem vessel lumen diameter. In addition, abcb14 mutants also showed decreased polar auxin transport through whole stems from apex to base.  abcb11 and abcb14 showed similar polar auxin transport phenotypes while abcg33 had wild-type polar auxin transport. These results identify these ABCB/MDR-type of ABC transporters as required for polar auxin transport in Arabidopsis, a result consistent with the auxin transporting roles of other ABCB/PGP/MDR transporters such as PGP1 and PGP19 (Geisler and Murphy, 2006). Given the demonstration by Geisler et al., (2005) of direct efflux of auxin by PGP19, it is reasonable to propose that ABCB11, ABCB14 and ABCB15 represent a new set of auxin transporters that are active in the stem of Arabidopsis. Promoter::GUS results of these genes showed that they are vascular cylinder specific but show no expression could be detected in the root tip or elongation zone. The auxin transporters in ABCB subfamily are expressed in immature root or elongation zone in root, therefore this new subset of stem ABCB/PGP/MDR transporters is distinct from the ones that had already been characterized in roots. However, an important stem auxin transporter, ABCB19/PGP19, has also been described and the relative contribution of these proteins is not known. These results suggest an important role of ABCB14 and possibly other B-subfamily ABC transporters in vascular differentiation related auxin transport in the stem.  170 Although we can conclude that these ABC transporters are required for normal polar auxin transport, what about their putative role in monolignol export? Overlapping expression patterns of these proteins, as revealed by promoter::glucuronidase reporters, raises the possibilities of gene redundancy, making it difficult to draw conclusions about additional functions for these transporters such as monolignol export. Could these ABC transporters move both auxins and monolignols? In mammalian cells, overexpressed MDR1 gene gained resistance to multiple cancer drugs (Gottesman et al. 2002). Since ABC transporters, especially ABCB/MDR subfamily proteins, can transport a wide variety of structurally unrelated substrates at the same time, one ABC protein might have multiple substrates. Therefore, if plant ABCB/MDR has same unique features as mammalian ABC transporters, Arabidopsis candidate ABC transporters in ABCB subfamily (ABCB11, ABCB14 and ABCB15) might transport monolignols in addition to auxin. Multiple gene disruption by T- DNA insertion or point mutation might create auxin-related problems so short term inducible reduction of transcript levels by RNAi construct might be a way to test if these ABC transporters are active in monolignol transport.  In chapter 4, in order to disable wide range of ABC transporter proteins, chemical inhibitors were used, which have been tested to inhibit ABC transporters in other biological systems, either plants, mammals or fungus. In poplar secondary xylem, two ABC transporter inhibitors were used to test whether ABC transporters are involved in monolignol transport and, unlike in developing pine wood, there was no effect on the incorporation of 3H- phenylalanine derivatives.  The new assay that I developed to quantify lignification and potential inhibition of lignification in Arabidopsis roots had the benefit of being fast and simple.  Tracheary element development was easily followed along the axis of the root and  171 the expected controls such as PA and NPA gave me confidence that if lignification were altered, I could detect it.  Since ABC inhibitors did not affect lignification in Arabidopsis seedling root tracheary elements, these results suggest that ABC transporters are not involved directly in monolignol export.  However, the possibility exists that the inhibitors used in this study, which can be subfamily specific, were not effective against a subset of ABC transporters responsible for monolignol transport in the root. I tried to avoid this problem by using a broad range of inhibitors including some that have been proven in plant cells like cyclosporin A. For some of these inhibitors, especially Reversin 121, which is a potent ABCB/MDR inhibitor for mammals, their efficacy against plant ABC transporters has not been proven.  Reversin 121 did not inhibit gravitropism in Arabidopsis roots grown on plates (unpublished data), which is an ABCB/MDR mediated process, so it is hard to be confident about that inhibitor.  But for FK506, efficacy could be demonstrated using a sclareol export assay.  This assay had been used previously to characterize as Arabidopsis ABC transporter mutant where wild-type plants were resistant to sclareol poisoning but mutant plants were sensitive.  In my study, FK506 plants were sensitive to sclareol poisoning but controls were not.  Therefore, I have more confidence that ABC transporters of the ABCG/PDR subfamily were effectively inhibited and yet lignification proceeded normally.  If there is one common lignification mechanism acting for all lignified cell types and experimental models were equally valid, then my results suggest that monolignol export is not a Golgi-mediated process and probably not ABC transporter mediated. That suggests other unknown transporters like multidrug and toxin exporter (MATE) transporters, which is a large plant and bacteria specific transporter known to transport flavonoids (Marinova et al. 2007) and glutathione conjugates (Diener et al. 2001), or  172 uncharacterized transporters that use the energy of the plasma membrane proton gradient, still need to be discovered. The question here is whether we can eliminate ABC transporters as a potential mechanism of monolignol transport based on my reverse genetics studies and ABC transporter inhibitor assays. Although knocking out best candidate ABC transporters did not result in reduced lignin phenotype, they were single knockout mutants. In the Arabidopsis root assay, a wide variety of inhibitors were tested and none stopped lignification. But reversin 121 was not confirmed to inhibit plant ABCB/MDR subfamily transporters. Thus, I conclude still ABC transporter should not be fully excluded as potential monolignol exporters. Further if the ABCB/MDR subfamily members can transport both monolignols and auxin, it will be hard to disrupt them without disturbing differentiation events well upstream of lignification. In retrospect, it may have been naïve to think that we could use single gene knockouts to characterize candidate ABC transporters involved in lignification when the correlation data set of Ehlting et al. (2005) contained so many ABC transporters, and no study has shown that these ABC transporters are involved in the lignification process. Although my thesis did not find the exact gene products that transport monolignols, it has made other contributions such as re-considering the Golgi-mediated vesicle transport theory for monolignol transport and proposing new theory; membrane transport of monolignol export, and new discovery of polar auxin transport proteins in Arabidopsis stem.  173  6.2 Bibliography  Anterola, AM., Jeon JH, Davin LB, Lewis NG. (2002) Transcriptional control of monolignol biosynthesis in Pinus taeda: factors affecting monolignol ratios and carbon allocation in phenylpropanoid metabolism. J Biol Chem 277, 18272- 18280. Atalla, RH. (2005) The role of the hemicelluloses in the nanobiology of wood cell walls : a systems theoretic perspective. Proceedings of the Hemicelluloses Workshop 2005 : 2005 January 10-12, The Wood Technology Research Centre, University of Canterbury, Christchurch, NZ. Christchurch, NZ : Wood Technology Research Centre, University of Canterbury, 2005: Pages 37-57 Boija, E., Johansson G. (2006) Interactions between model membranes and lignin-related compounds studied by immobilized liposome chromatography. Biochim Biophys Acta 1758, 620-626. Boija, E., Lundquist A, Edwards K, Johansson G. (2007) Evaluation of bilayer disks as plant cell membrane models in partition studies. Anal Biochem 364, 145-152. Blount, JW., Korth KL, Masoud SA, Rasmussen S, Lamb C, Dixon RA. (2000) Altering expression of cinnamic acid 4-hydroxylase in transgenic plants provides evidence for a feedback loop at the entry point into the phenylpropanoid pathway. Plant Physiol 122, 107-116. Diener, AC, Gaxiola RA, Fink GR. (2001) Arabidopsis ALF5, a multidrug efflux transporter gene family member, confers resistance to toxins. Plant Cell 13, 1625-1638. Ehlting, J, Mattheus N, Aeschliman DS, Li E, Hamberger B, Cullis IF, Zhuang J, Kaneda M, Mansfield SD, Samuels L, Ritland K, Ellis BE, Bohlmann J, Douglas CJ. (2005) Global transcript profiling of primary stems from Arabidopsis thaliana identifies candidate genes for missing links in lignin biosynthesis and transcriptional regulators of fiber differentiation. Plant J 42, 618-640. Geisler, M, JJ Blakeslee, RBouchard, OR Lee, V Vincenzetti, A Bandyopadhyay, B Titapiwatanakun, WA Peer, A Bailly, EL Richards, KFK Ejendal, AP Smith, C Baroux, U Grossniklaus, A Muller, CA Hrycyna, R Dudler, AS Murphy2 and E Martinoia (2005) Cellular efflux of auxin catalyzed by the Arabidopsis MDR/PGP transporter AtPGP1. Plant J  44, 179–194 Geisler, M, Murphy AS. (2006) The ABC of auxin transport: the role of p-glycoproteins in plant development. FEBS Lett 580, 1094-1102. Gottesman, MM, Fojo T, Bates SE. (2002) Multidrug resistance in cancer: role of ATP- dependent transporters. Nat Rev Cancer 2, 48-58. Marinova, K, Pourcel L, Weder B, Schwarz M, Barron D, Routaboul JM, Debeaujon I, Klein M. (2007) The Arabidopsis MATE transporter TT12 acts as a vacuolar flavonoid/H+ -antiporter active in proanthocyanidin-accumulating cells of the seed coat. Plant Cell 19, 2023-2038. Pena MJ, Zhong R, Zhou G.-K, Richardson EA, O’Neill MA, Darvill AG, York WS, Ye Z.-H. (2007) Arabidopsis irregular xylem8 and irregular xylem9: implications for the complexity of glucuronoxylan biosynthesis. Plant Cell 19:549–563.

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            data-media="{[{embed.selectedMedia}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
https://iiif.library.ubc.ca/presentation/dsp.24.1-0066871/manifest

Comment

Related Items