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CLASP : a microtubule-based integrator of the hormone-mediated transitions from cell division to elongation… Ruan, Yuan 2015

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  CLASP: A MICROTUBULE-BASED INTEGRATOR OF THE HORMONE-MEDIATED TRANSITIONS FROM CELL DIVISION TO ELONGATION IN ARABIDOPSIS THALIANA   by YUAN RUAN B.Sc., Shandong Normal University, 2005 M.Sc., Shandong Normal University, 2008  A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in The Faculty of Graduate and Postdoctoral Studies (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  October 2015 ©	Yuan Ruan, 2015     iiABSTRACT  Microtubules have long been known to play a vital role in plant growth and development, which is a complex process and needs to be regulated by both environmental and endogenous hormonal signals.  CLASP, an important microtubule-associated protein, has been shown to be involved in both cell division and expansion. The major goal of my thesis was to explore the function of CLASP in new pathways as well as to identify novel factors that are responsible for its function and distribution.  Yeast 2-hybird analysis done independently by two of our collaborators indicated that CLASP interacts strongly with the endocytic membrane-associated protein SNX1. SNX1 had been implicated in the intracellular trafficking of the auxin efflux carrier PIN2. We proved the direct interaction between CLASP and SNX1 by colocalization and BiFC using live cell imaging and we found that clasp-1 mutants have an altered distribution pattern of SNX1, reduced abundance of PIN2 and a series of auxin-related phenotypes such as dwarfism, enhanced lateral branching and aberrant auxin distribution. Drug-induced microtubule disruption caused clasp-1-like defects on PIN2 stability. This study illustrated the role of CLASP and microtubules in polar auxin transport and auxin signalling pathway.  Previous studies revealed a cross-talk between auxin and brassinosteroids. I found that the two major transcription factors in the brassinosteroid signalling pathway directly target the CLASP promoter to repress its transcription. Reduced CLASP expression is correlated with a transverse orientation of cortical microtubules in root meristematic cells, switching them from division to differentiation. Also CLASP and microtubules stabilize the BR receptor BRI1 by preventing its degradation, a similar mechanism as for PIN2. This research highlights the importance of CLASP in BR-modulated meristem cell identity and the potential role of CLASP as a node between auxin and BR pathways. I carried out an immunoprecipitation experiment to identify putative CLASP interactors and obtained TRM19. Further analysis failed to confirm a direct interaction but suggested that TRM19 is a microtubule-associated protein. Its high expression in dividing cells is consistent with prior report that it functions in PPB formation during the cell cycle.  iiiPREFACE  Chapter 1 has been published as: Ruan, Y., and Wasteneys, G.O. (2014) CLASP: a microtubule-based integrator of the hormone-mediated transitions from cell division to elongation. Curr Opin Plant Biol. 22: 149-158 © Copyright Elsevier Ltd., 2014 ( Prof. Geoffrey Wasteneys and Yuan Ruan conceived the idea. Yuan Ruan prepared the figures and wrote the manuscript. Chapter 2 has been published as: Ambrose, C., Ruan, Y., Gardiner, J., Tamblyn L.M., Catching, A., Kirik, V., Marc, J., Overall, R., and Wasteneys, G.O. (2013) CLASP interacts with soring nexin 1 to link microtubules and auxin transport via PIN2 recycling in Arabidopsis thaliana. Dev Cell. 24: 649-659 © Copyright Elsevier Inc., 2013 ( Yeast 2-hybrid screens were conducted separately by our collaborators Dr. John Gardiner at the University of Sydney and Dr. Viktor Kirik at Illinois State University and both identified SNX1. Dr. John Gardiner also predicted and created the 3D structure of the interacting site on SNX1. DR5:GUS analysis was performed by Dr. Viktor Kirik. An undergraduate student, Laura Tamblyn, prepared the RNA samples for RT-PCR. Yuan Ruan carried out BiFC, GFP-SNX1134-219 reporter experiments, analyzed SNX1-GFP in WT and clasp-1, and tracked SNX1-RFP vesicle trajectories. Yuan Ruan also wrote the corresponding materials and methods section. Dr. Chris Ambrose did most of the remaining experiments and processed the data. This work was done under the supervision of Prof. Geoffrey Wasteneys. Chapter 3 has been prepared as a manuscript: Ruan, Y., Ambrose, C., and Wasteneys, G.O. (2015) CLASP modulates its own transcription via brassinosteroid signalling to drive stem cell differentiation in plants. Yuan Ruan and Prof. Geoffrey Wasteneys designed the research. Yuan Ruan performed the experiments, processed the data, prepared figures and wrote the manuscript. For Chapter 4, Prof. Geoffrey Wasteneys and Yuan Ruan designed the research. Yuan Ruan performed the experiments and analyzed the data.    ivTABLE OF CONTENTS ABSTRACT ....................................................................................................................... ii PREFACE ......................................................................................................................... iii TABLE OF CONTENTS ................................................................................................. iv LIST OF TABLES ........................................................................................................... vii LIST OF FIGURES ....................................................................................................... viii LIST OF ABBREVIATIONS .......................................................................................... ix ACKNOWLEDGEMENTS ........................................................................................... xii CHAPTER 1: INTRODUCTION .................................................................................... 1 1.1 Meristem maintenance and plant development .................................................... 1 1.2 Hormones, auxin transport and meristem maintenance ..................................... 1 1.3 Auxin-driven microtubule entrainment ................................................................. 4 1.4 Cytoskeleton control of auxin levels through PIN endocytosis ........................... 6 1.5 Microtubule control of cell division and its link to cellular geometry ................ 8 1.6 The transition to cell division ............................................................................... 10 1.7 Conclusions and future directions ........................................................................ 13 1.8 Thesis objectives .................................................................................................... 15 CHAPTER 2: CLASP INTERACTS WITH SORTING NEXIN 1 TO LINK MICROTUBULES AND AUXIN TRANSPORT VIA PIN2 RECYCLING IN ARABIDOPSIS THALIANA ........................................................................................... 16 2.1 Introduction ........................................................................................................... 16 2.2 Results ..................................................................................................................... 17 2.2.1 SNX1 interacts with CLASP ............................................................................ 17 2.2.2 Central region between PHOX and BAR domains of SNX1 confers interaction with CLASP ............................................................................................................... 18 2.2.3 CLASP686-779 interacts with SNX1 .................................................................... 20 2.2.4 CLASP stabilizes SNX1 endosomes ................................................................. 21 2.2.5 SNX1 endosomes associate with CLASP and MTs .......................................... 24  v2.2.6 SNX1 vesicles form dynamic clusters that associate with GFP-CLASP ......... 27 2.2.7 clasp-1 mutants exhibit enhanced PIN2 degradation but retain PIN2 polar distribution ................................................................................................................. 29 2.2.8 MTs inhibit degradation of PIN2 to facilitate recycling ................................... 32 2.2.9 clasp-1 mutants exhibit auxin-related phenotypes ............................................ 33 2.3 Discussion ............................................................................................................... 37 2.4 Methods .................................................................................................................. 39 2.4.1 Plant materials and growth conditions .............................................................. 39 2.4.2 Microscopy and image analysis ........................................................................ 39 2.4.3 Drug treatments ................................................................................................. 40 2.4.4 Yeast Two-hybrid analysis ................................................................................ 40 2.4.5 RNA extraction following hormone treatments ................................................ 41 2.4.6 BiFC analysis .................................................................................................... 42 2.4.7 Immunofluorescence of PINs in roots .............................................................. 43 CHAPTER 3: A MICROTUBULE-ASSOCIATED CLASP-BRI1-BZR1/2 FEEDBACK LOOP MODULATES BRASSINOSTEROID-DEPENDENT ROOT DEVELOPMENT IN ARABIDOPSIS THALIANA ..................................................... 45 3.1 Introduction ........................................................................................................... 45 3.2 Results ..................................................................................................................... 47 3.2.1 BR modulates microtubule orientation and CLASP subcellular distribution ... 47 3.2.2 BR reduces CLASP expression in a BRI1-dependent manner .......................... 49 3.2.3 BZR1/2 directly binds to the CLASP promoter to repress its expression ......... 51 3.2.4 CLASP sustains BRI1 activity at the plasma membrane .................................. 56 3.2.5 CLASP's association with microtubules is critical for preventing BRI1 degradation ................................................................................................................. 57 3.2.6 BRI1-dependent brassinosteroid signalling is reduced in clasp-1 mutants ...... 59 3.3 Discussion ............................................................................................................... 62 3.4 Methods .................................................................................................................. 64 3.4.1 Plant materials and growth conditions .............................................................. 64 3.4.2 Molecular cloning and plant transformation ..................................................... 65 3.4.3 Drug treatments ................................................................................................. 65 3.4.4 Confocal microscopy and image analysis ......................................................... 66 3.4.5 Total RNA extraction and quantitative Real-Time PCR (qRT-PCR) ................ 66 3.4.6 Electrophoretic mobility shift assay (EMSA) ................................................... 66 3.4.7 Tobacco transient assay ..................................................................................... 67 CHAPTER 4: ARABIDOPSIS THALIANA TRM19: A MICROTUBULE-ASSOCIATED PROTEIN IDENTIFIED THROUGH IMMUNOPRECIPITATION WITH CLASP ................................................................................................................. 70  vi4.1 Introduction ........................................................................................................... 70 4.2 Results ..................................................................................................................... 71 4.2.1 TRM19 was identified through an immunoprecipitation experiment ............... 71 4.2.2 TRM19 is a microtubule-associated protein but does not interact directly with CLASP ....................................................................................................................... 72 4.2.3 TRM19 expression pattern ................................................................................ 73 4.2.4 trm19 mutant phenotype ................................................................................... 75 4.2.5 TRM19 stable line analysis ............................................................................... 76 4.2.6 Expression of TRM19 under various stress conditions ..................................... 78 4.3 Discussion ............................................................................................................... 79 4.4 Methods .................................................................................................................. 80 4.4.1 Plant materials and growth conditions .............................................................. 80 4.4.2 Immunoprecipitation (IP) assay ........................................................................ 80 4.4.3 BiFC .................................................................................................................. 81 4.4.4 Yeast 2-Hybrid analysis .................................................................................... 81 4.4.5 Total RNA extraction and RT-PCR ................................................................... 81 4.4.6 Identification of T-DNA insertions ................................................................... 82 4.4.7 Plasmid construction and generation of stable transgenic lines ........................ 82 4.4.8 Drug treatments ................................................................................................. 82 4.4.9 Confocal imaging .............................................................................................. 83 4.4.10 Histochemical GUS staining ........................................................................... 83 CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS ............................... 85 5.1 Conclusions ............................................................................................................ 85 5.1.1 CLASP and the auxin signalling pathway ........................................................ 86 5.1.2 CLASP and brassinosteroid signalling pathway ............................................... 87 5.1.3 TRM19 .............................................................................................................. 88 5.2 Future directions .................................................................................................... 89 5.2.1 Additional functions of CLASP ........................................................................ 89 5.2.2 Biological function of TRM19 .......................................................................... 91 REFERENCES ................................................................................................................ 92       viiLIST OF TABLES Table 2.1: Putative interaction with CLASP by Yeast 2-Hybrid Assay ............................ 18 Table 2.2: Primers used in this study ................................................................................ 44 Table 3.1: Primers used in this study ................................................................................ 68 Table 4.1: Immunoprecipitation results ............................................................................ 72 Table 4.2: Primers used in this study ................................................................................ 84 Table 5.1: Genes that co-express with CLASP .................................................................. 91                                  viiiLIST OF FIGURES Figure 1.1: Cytoskeletal regulation of PIN ......................................................................... 5 Figure 1.2: Non-transverse CMT arrays in post-cytokinetic cells .................................... 12 Figure 1.3: Microtubule and cell wall-based feedback loops in auxin signalling ............ 14 Figure 2.1: CLASP interacts with SNX1 .......................................................................... 19 Figure 2.2: CLASP686-779 interacts with SNX1 ................................................................. 21 Figure 2.3: CLASP and MTs stabilize SNX1 vesicles ...................................................... 23 Figure 2.4: SNX1 endosomes associate with CLASP and MTs ....................................... 26 Figure 2.5: Association of FM4-64 endosomes with MTs ................................................ 26 Figure 2.6: SNX1 forms CLASP-associated vesicular clusters ........................................ 28 Figure 2.7: PIN2 expression domain and levels are reduced in clasp-1 root tips ............. 30 Figure 2.8: PIN2 levels and PIN2-mediated gravitropic responses in clasp-1 mutants ... 31 Figure 2.9: Depolymerization of  MTs with oryzalin induces PIN2-GFP accumulation within lytic vacuoles .................................................................................................. 33 Figure 2.10: The clasp-1 mutant displays auxin-related defects ...................................... 35 Figure 2.11: DR5:GUS in roots and DR5:GFP in clasp-1 cotyledons exhibit abnormalities .............................................................................................................. 36 Figure 3.1: eBL regulates CLASP localization and microtubule organization ................. 48 Figure 3.2: eBL reduces CLASP transcription in a BRI1-dependent manner ................... 50 Figure 3.3: BZR1 and BES1 directly bind to the CLASP promoter ................................. 53 Figure 3.4: Motifs and probes in CLASP promoter .......................................................... 54 Figure 3.5: 35Spro:GFP, 35Spro:GFP-BZR1 and 35Spro:BES1-GFP are ubiquitously expressed in tobacco leaves in the transient assay ..................................................... 54 Figure 3.6: BZR1 and BES1 repress CLASP expression .................................................. 55 Figure 3.7: Meristem of bzr1-1D ...................................................................................... 56 Figure 3.8: CLASP modulates BRI1 stability and transcription ....................................... 58 Figure 3.9: Oryzalin treatment removes microtubules including CLASP-mediated transfacial bundles ..................................................................................................... 59 Figure 3.10: clasp-1 is hyposensitive to exogenous eBL treatments ................................ 61 Figure 3.11: Model illustrating the function of CLASP in BR-controlled meristem development ............................................................................................................... 64 Figure 4.1: Transient expression of TRM19 in wild-type and clasp-1 cotyledons ........... 73 Figure 4.2: TRM19 expression by RT-PCR ...................................................................... 74 Figure 4.3: TRM19 promoter activity indicated by the GUS assay .................................. 75 Figure 4.4: TRM19 expression by RT-PCR in mutants ..................................................... 76 Figure 4.5: TRM19pro:genomic TRM19-GFP was detected in all root layers ................... 77 Figure 4.6: TRM19-GFP showed diffused signal after plasmolysis ................................. 78 Figure 4.7: UBQpro:GFP-TRM19 showed PM localization with highest fluorescence in root transition zone .................................................................................................... 78         ixLIST OF ABBREVIATIONS 2,4-D 35S  ABA 2,4-dichlorophenoxyacetic acid a very strong constitutive promoter found in Cauliflower mosaic virus (CaMV) abscisic acid  ABP1 Auxin binding protein 1 ABRC  Arabidopsis biological resource center  ARF  ADP ribosylation factor Arabidopsis BAK1 BES1 BFA BiFC BL Arabidopsis thaliana BRI1-ASSOCIATED RECEPTOR KINASE 1 BRI1 EMS SUPPRESSOR-1 brefeldin A bimolecular fluorescence complementation brassinolide BR BRL BRAVO BRI1 BSA  brassinosteroid BRI1-like Brassinosteroids at Vascular and Organizing Centre BRASSINOSTEROID INSENSITIVE 1 bovine serum albumin BZR1 BRASSINAOLE RESISTANT 1 cDNA ChIP complimentary DNA chromatin immunoprecipitation CLASP CLIP CME CMT CLIP-ASSOCIATED PROTEIN CYTOPLASMIC LINKER PROTEIN  clathrin-mediated endocytosis cortical microtubule  Col-0  Columbia-0  DMSO  dimethyl sulfoxide DNA eBL deoxyribonucleic acid 2,4-epibrassinolide  xEDTA  ethylenediaminetetraacetic acid  EMS EMSA En ethyl methanesulphonate electrophoretic mobility shift assay Enkheim ER EtOH  endoplasmic reticulum  ethanol GFP  green fluorescent protein  GUS  β-glucuronidase  IAA IP  indole-3-acetic acid immunoprecipitation  LB  lysogeny broth  Ler  Landsberg erecta  MAP  MAPK MBP MDP40 microtubule-associated  protein mitogen-activated protein kinase maltose binding protein MICROTUBULE DESTABILIZING PROTEIN 40 mRNA MS messenger RNA Murashige-Skoog MT microtubule  NAA  NPA 1-Naphthaleneacetic acid 1-N-Naphthylphthalamic acid PCR PHS1 PI PIN PLT1/2  polymerase chain reaction propyzamide-hypersensitive 1 propidium iodide PIN-FORMED PLETHORA1/2 PM  plasma membrane  PPB PP2A QC qRT-PCR preprophase band phosphatase 2A  quiescent centre quantitative Real-Time PCR  xiRAM root apical meristem RNA  ribonucleic acid  RNAi  RNA interference  RNase RT-PCR SAM ribonuclease reverse transcriptase PCR shoot apical meristem SNX1 sorting nexin 1 T-DNA  transfer-DNA  TIC  TMK TON2/FASS TRM TTP TIME FOR COFFEE transmembrane kinase receptor like kinases TONNEAU2/FASS TON1 Recruiting Motif TON1-TRM-PP2A complex UBQ ubiquitin VPS29  vacuolar protein sorting 29 Ws  Wassilewskija YFP  yellow fluorescent protein Y2H  Yeast 2-Hybrid                   xiiACKNOWLEDGEMENTS  My first and deepest gratitude goes to my supervisor, Dr. Geoffrey Wasteneys. Thanks go to him for giving me the opportunity to work in such a wonderful lab and continued patience and encouragement throughout these years of my PhD study. Without him, I wouldn’t be able to finish my dissertation and obtain so many good results.  I would also like to thank my committee members. Thanks go to Dr. George Haughn for helping me understand genetic knowledge while I was performing EMS mutagenesis. Thanks go to Dr. Lacey Samuels for her valuable advice on the intracellular trafficking part. Special thanks to Dr. Chris Ambrose, for not only providing technical training such as microscopy and image analysis but also for helping to develop my academic interests and scientific thinking. He was important company and witness in this incredible journey.  I would like to thank the postdoctoral researchers who trained me in the techniques needed for my projects. Thanks go to Dr. Sylwia Jancowski for helping with real time PCR analysis. Thanks go to Dr. Bettina Lechner for her guidance on protein expression and purification in bacterial cells. I would also like to thank all the graduate students and undergraduates who helped me with my experiments including Dr. Yan Zhang, at that time a visiting student from Chinese Agricultural University, for root responses to BR treatments; work-learn students Rachel Cederberg and Junsu Kwon for medium preparation, seed planting and cloning, and Salar Fazeli, a directed studies student who assisted with experiments that established CLASP as a target of brassinosteroid signalling. Many thanks to my former and current lab members, Dr. Miki Fujita, Dr. Eric Johnson, Yi Zhang, Ryan Eng, Caitlin Donnelly, Dr. Ankit Walia, Amanda Catching, Dr. Katherine Celler, Adam Mulvihill, for all helpful suggestions and discussions on my projects and precious friendships. Thanks for creating a comfortable and healthy and inspiring work environment and bringing about so many unforgettable memories.  I would like to thank the Department of Botany at Universtiy of British Columbia for providing TAship to support me financially and a chance to communicate with students and develop my teaching skills. Thanks especially to my parents for unconditional love, care and patience. Thanks to my friends who share my feelings. They are the motivation during this long journey of my PhD life. 1  CHAPTER 1: INTRODUCTION 1.1 Meristem maintenance and plant development All plant tissues originate from meristems located at shoot and root apices. Both the shoot apical meristem (SAM) and root apical meristem (RAM) are highly dynamic, integrating versatile developmental as well as environmental inputs into overall plant architecture (Yoshida et al., 2011; Fernández-Marcos et al., 2011). The SAM is dome shaped and organized into three layers (L1-L3) and three zones (central, peripheral and rib) (Irish and Sussex, 1992). The central zone resides at the summit of SAM and slow division events occurring here serve to keep a constant number of stem cell initials. In contrast, peripheral cells sense positional cues, translating them into accelerated proliferation and directional growth as they are recruited into rapidly outgrowing lateral primodia, defined by the surrounding growth-arrested organ boundary cells (Ha et al., 2010). In the RAM, the stem cell niche comprises a mitotically inactive quiescent centre surrounded by initial cells that produce different tissues. Daughter cells undergo rapid elongation and differentiation when they are displaced from the root tip, a process that simultaneously builds the protective root cap and drives the root tip deeper into the soil (Rost, 2011). Ultimately, meristem size control, which is coordinated by overlapping transcription factor networks (Sarkar et al., 2007), relies on the precise balance between maintaining a reservoir of undetermined cells and cell differentiation, giving rise to new tissues and/or organs through a wide spectrum of auxin-dependent activities and hormonal cross-talk (Murray et al., 2012; Perilli et al., 2012; Sozzani et al., 2014; Takatsuka and Umeda, 2014; Schuster et al., 2014). In this chapter, I highlight recent findings that open new avenues to understanding the mechanisms that control meristem activity. In particular, I focus on how hormonal signals are both processed and integrated by cytoskeletal machinery.   1.2 Hormones, auxin transport and meristem maintenance Previous studies have focused on auxin and cytokinin, the two key participants in meristem development, providing evidence for an underlying complex hormone network. The auxin to cytokinin ratio is critical for meristematic cell fate determination, as evidenced by the fact that an auxin maximum at the root tip promotes cell division by 2  upregulating cell cycle genes governing transition states, whereas cytokinin stimulates differentiation, thereby antagonizing auxin’s role. Conversely, relatively high auxin activities are required for lateral organogenesis while high levels of cytokinins are linked with maintenance of uncommitted cells in the SAM (Su et al., 2011; Zhang et al., 2013; Del Bianco et al., 2013). Recent studies have elucidated an intriguing involvement of the brassinosteroid (BR) hormone brassinolide (BL) in the morphogenesis of both the SAM and RAM. Low levels of BL or loss of BL perception in the SAM lead to deeper clefts between the main stem and axillary organs in contrast to the organ-fusion phenotypes, which result from loss of primordium boundaries in BL-hypersensitive mutants (Gendron et al., 2012; Bell et al., 2012). These studies establish the importance of BR homeostasis in the expression of boundary identity genes. By analogy, BL affects quiescent centre (QC) activity and promotes cell cycle progression in the RAM. Disturbance to normal BR signalling reduces RAM size either because of lower division rates in the initial cells or premature differentiation of meristematic cells (Hacham et al., 2011; González-García et al., 2011). It was recently known that the R2R3 transcription factor BRAVO (Brassinosteroids at Vascular and Organizing Centre) is expressed within the QC and counteracts BR-stimulated QC division (Vilarrasa-Blasi et al., 2014). This discovery identifies a cell-specific molecular mechanism that explains how the BR signalling machinery can be dampened in a cell-specific manner, which in the case of the cells in the quiescent centre, is critical for maintaining stem cell identity in the surrounding region. Auxin gradients and local auxin maxima and minima, which drive the differential expression of genes for the body plan, are built up and retained mainly by rate-limiting auxin efflux carriers, most notably the PIN family proteins, which generate directional flow of auxin through their asymmetric (polar) distribution on the plasma membrane (Blilou et al., 2005; Petrásek et al., 2006). PIN1 in the SAM is positioned towards cells accumulating higher concentrations of auxin in the L1 layer and in the developing leaf vasculature, accounting for high auxin flow through the incipient organ primordia and drainage through developing vascular strands (Reinhardt et al., 2003; Petrásek and Friml et al., 2009). Recent work explains the defective leaf production in an erecta family gene triple mutant (Uchida et al., 2012; Chen et al., 2013). Attenuation of PIN1 causes 3  homogenous accumulation of auxin in the L1 layer, and prevents its movement into the vasculature. The enlarged SAM in these mutants can be explained by either reduced recruitment of stem cells or the dramatic expansion of cells in the L1 layer (Uchida et al., 2012; Chen et al., 2013) as a consequence of higher than normal auxin concentrations.  In roots, the flow of auxin is critical to the maintenance of RAM identity and organ elongation. This upside-down fountain pattern is coordinated synergistically through the tissue-specific polar distribution of PINs 1, 2, 3, 4 and 7. Apically polarized PINs 1, 3, 4 and 7 in the stele direct auxin towards the root tip. PINs 3 and 7 account for lateral redistribution in the root cap. PIN2 is instrumental for basipetal movement of auxin through the epidermal layer as well as elongating lateral root cap and cortex tissues. Finally, PINs 2, 7 and 3 complete the loop by diverting shoot-bound auxin into the stele to maintain auxin homeostasis in the root tip (Petrásek and Friml, 2009). It is now well established that perturbing polar auxin transport with chemical inhibitors or mutations affecting the acitivity of auxin tranporters leads to auxin imbalances and concomitant meristem abnormalities. Modulating meristem activity in response to developmental programs and environmental cues is carried out by many complex and overlapping molecular mechanisms, the understanding of which remains far from clear. Consistent with meristem acitivity being integrated with the circadian clock, it was recently reported that PLT1/2 and consequently PIN1, 2, 3 and 7 gene expressions are downregulated in the loss of function tic-2 mutant (Hong et al., 2014). This reduces overall auxin abundance in the root, producing a smaller meristem by compromising stem cell competence to divide. Perturbing just one PIN protein within the network can have similar consequences and may be of particular relevance to modulating meristem activity under changing environmental conditions. The recent discovery that the microtubule-associated protein CLASP has a role in controlling PIN2 levels post-translationally, identifies a previously unknown regulatory mechanism for meristem activity (Ambrose et al., 2013). In mutants lacking CLASP gene expression, specific depletion of PIN2 perturbs the auxin reflux mechanism, causing auxin to pool in the meristem, a feature that is associated with precocious differentiation. This, and other related phenomena are discussed in the following section. 4  1.3 Auxin-driven microtubule entrainment It has long been known that the cytoskeleton serves to convert auxin signals into productive work such as unidirectional growth or organ bending in response to light, touch or gravity stimuli (Nick et al., 1992; Holweg et al., 2004; Lindeboom et al., 2013). The findings of Xu et al. (2014) have clarified the underlying molecular mechanism of interpreting auxin as an extracellular messenger. In this study, auxin was shown to trigger the formation of the ABP1 (auxin-binding protein 1)-TMK (transmembrane kinase receptor like kinases) complex near the cell surface, which in turn activates the ROP GTPase (Rho Of Plants guanosine triphosphatase) signalling cascade to bring about fundamental changes in cytoskeleton configuration and cell shape (Figure 1.1a). Recent studies, however, revealed that the abp1 null mutants do not possess any recognizable auxin-associated phenotypes and that the auxin phenotype of the abp1-5 allele is not linked with the ABP1 loci, putting into serious doubt the role of ABP1 and TMKs in auxin signalling (Enders et al., 2015; Gao et al., 2015). An important insight into how ROP activity can generate microtubule alignment was revealed by Lin et al. (2013), who determined that the ROP effector RIC1 interacts directly with and promotes the activity of the microtubule severing protein katanin. Evidently the release of newly nucleated microtubule branches facilitates their self organization through encounter-based entrainment. Computational simulations predicted a fourfold increase in the time to achieve parallel order without katanin-based severing and release of branch microtubule minus ends (Allard et al., 2010). Subsequent experimental studies with katanin mutants have borne out this prediction, showing that either the normal establishment of parallel order (Lin et al., 2013) or realignment following changes in stress patterns (Sampathkumar et al., 2014) in leaf pavement cells or the blue light-induced shift from transverse to longitudinal orientation in hypocotyls (Lindeboom et al., 2013) are delayed in katanin mutants. Consistent also with the idea that branch-form nucleation generates discordant microtubules for recruitment to new orientations, a recent study indicates that the loss of TON2/FASS-mediated branch form nucleation reduces the rate at which microtubules can reorient in response to hormone treatments (Atkinson et al., 2014). 5                    Figure 1.1: Cytoskeletal regulation of PIN (a) Leaf pavement cell. ROP6/RIC1 stablizes microtubules at the neck region restraining outgrowth while ROP2/RIC4 stabilizes actin filaments at the lobe tip promoting local growth. Stabilized F-actin inhibits PIN1 internalization.  (b) Root cell. ROP6/RIC1 stabilizes actin filament and inhibits PIN2 endocytosis. CLASP facilitates PIN2 recycling to the plasma membrane by direct interaction with SNX1, preventing it from BLOC-1 mediated vacuolar degradation.   6  1.4 Cytoskeletal control of auxin levels through PIN endocytosis More and more studies are demonstrating that the auxin-cytoskeleton relationship can be bidirectional, incorporating complex feedback mechanisms that amplify signals or transport mechanisms. For example, auxin-mediated realignment of actin filaments can further consolidate auxin movement (Nick et al., 2009). In the SAM, the close correspondence between microtubule orientation and PIN1 polarity suggests that microtubule-directed cellular expansion is integrated with auxin-induced organ outgrowth (Heisler et al., 2010). And finally, cellulose-dependent cell wall integrity has been shown to be an important factor in controlling the polar distribution of PIN proteins (Feraru et al., 2011). Thus, PIN-dependent auxin transport, through its control of microtubule entrainment and consequent mechano-chemical wall properties and cell shape, can reinforce its own polarity. Whereas cytoskeletal involvement in auxin responses as well as endocytosis-mediated control of auxin transport has been documented (Paciorek et al., 2005; Maisch and Nick, 2007; Nick, 2010), whether and how the cytoskeleton regulates auxin and auxin polar transport has remained obscure or controversial. Unlike the polar secretion mode described for epithelial cells (Mostov et al., 2003), Arabidopsis PIN proteins are targeted symmetrically to both sides of the cell plate during cytokinesis under the guidance of the microtubule-based phragmoplast and subsequently, PIN polarity is achieved post-mitotically through retrieval from selective cell faces by clathrin-mediated endocytosis (CME) (Geldner et al., 2001; Kitakura et al., 2011). In yeast and animal cells, the actin cytoskeleton is implicated in CME (Yarar et al., 2005; Collins et al., 2011). High resolution microscopy has revealed, for example, that actin patches help the endocytic process by constricting and elongating the neck of clathrin-coated vesicles in cultured mouse cells (Collins et al., 2011). In plant cells, two prominent studies have elucidated the rudimentary role of actin in PIN endocytosis (Nagawa et al., 2012; Lin et al., 2012). These articles show that stabilized actin filaments resulting from the activity of ROP GTPases and their effectors impede clathrin-mediated PIN protein internalization, with ROP2 and RIC4 controlling PIN1 retention in pavement cell lobe regions (Nagawa et al., 2012) and ROP6 and RIC1 controlling PIN2 in root cells (Lin et al., 2012) (Figure 1.1a, b). Integrating the knowledge that auxin can stimulate ROP GTPase activity, the auxin-ROP/RIC-PIN-auxin loop can be described as a self-7  regulating positive feedback system (Xu et al., 2014). Although it was previously shown that RIC1 binds to and promotes microtubule ordering in leaf pavement cells (Fu et al., 2005), according to Lin et al. (2012), PIN2 retention on the plasma membrane is microtubule-independent.  Following internalization, PIN proteins are continuously shuttled between the plasma membrane and endosomal compartments. An intact actin cytoskeleton is necessary for both apical and basal targeting, despite the fact that delivery towards the apical side is more sensitive to actin depolymerization (Kleine-Vehn et al., 2008b). Vesicles carrying PIN proteins have been identified in the phragmoplast, resulting in an unbiased secretion to the expanding cell plate of dividing cells (Geldner et al., 2001). In interphase cells, short-term oryzalin treatment to destabilize microtubules was shown to have little or no effect on PIN polarity, suggesting that microtubules play no role in PIN localization (Boutté et al., 2006). Prolonged microtubule perturbation provoked polarity alteration preferentially for basal localization (Kleine-Vehn et al., 2008b) but this could be due to profound changes to the growth axis and the complete blocking of cytokinesis. In 2013, the description of a function for microtubules, via the microtubule-associated protein CLASP, in the modulation of PIN2 protein levels (Ambrose et al., 2013; chapter 2) broadened our understanding of microtubule involvement in the fine tuning of polar auxin transport (Figure 1.1b). Maintaining PIN homeostasis is achieved both by endocytosis and proteolytic activity. The putative BLOC-1 complex in Arabidopsis mediates the transport of PIN2 for degradation in the vacuole through a direct interaction with SNX1 endosomes (Jaillais et al., 2006; Kleine-Vehn et al., 2008c; Cui et al., 2010). CLASP associates with SNX1 to increase endosome stability along microtubules, thereby facilitating the recycling of PIN2 to the plasma membrane and consequently impeding its transit to the vacuole (Ambrose et al., 2013). In mutants lacking CLASP gene transcription, PIN2 is drastically reduced at the plasma membrane (whereas PIN1 quantity and polarity are unaffected), causing an expanded region of auxin accumulation in the root tip, consistent with impaired PIN2-dependent shootward reflux. Expanded auxin maxima in the RAM are correlated with meristem collapse (Ambrose et al., 2007; Kirik et al., 2007; Ambrose et al., 2013). BLOC-1 complex-related protein degradation is evolutionarily conserved in eukaryotes (Raposo et al., 2007; Hermann et al., 2012).  8  The CLASP-SNX-BLOC-1 mechanism's role in auxin transport appears to be specific to PIN2 and does not influence PIN2 polarity establishment or maintenance. Maintaining the polarity of PIN proteins is also critical for auxin flux patterns. In this regard, the fluid nature of the plasma membrane and the integrity of the plant cell wall are both important. Notably, a deficiency in membrane sterol biosynthesis seems to obliterate PIN polarity (Willemsen et al., 2003). High resolution imaging and computational simulation analysis suggest that non-mobile domains at the plasma membrane minimize PIN lateral diffusion. PIN proteins that occasionally escape from these regions can be retrieved by clathrin-mediated endocytosis (Men et al., 2008; Kleine-Vehn et al., 2011). One obvious mechanism for establishing plasma membrane heterogeneity is the direct targeting of ER-derived and/or Golgi-derived plasma membrane components but the lack of asymmetric distribution of cytoplasmic organelles suggests that other mechanisms are at play (Boutté et al., 2006). Indeed, it appears that the cell wall and, to some extent, the cytoskeleton can constrain lateral diffusion of PIN proteins, suggesting that the cytoskeleton may facilitate connections between the plasma membrane and the cell wall in order  to create spatially discrete domains (Feraru et al., 2011; Martinière et al., 2011; Martinière et al., 2012). COBRA, a potential integrator of microtubules and the cellulosic cell wall (Roudier et al., 2005), warrants further study.  1.5 Microtubule control of cell division and its link to cellular geometry The microtubule-based machinery that coordinates mitosis and cytokinesis is an essential feature of auxin-induced cell division and meristem maintenance. The breakdown and rebuilding of these arrays through the cell cycle are controlled by the activity of microtubule nucleating complexes, and various associated proteins (MAPs) that can promote polymer addition, catastrophe, severing and bundle formation. Recent reviews have already offered excellent information with respect to cytoskeleton and associated protein-dependent PPB formation and phragmoplast guidance, mitotic spindle assembly and the mitogen-activated protein kinase (MAPK) cascade during cytokinesis (Sasabe and Machida, 2012; Rasmussen et al., 2013; Masoud et al., 2013). Hence, we focus here on emerging ideas about cell geometry-based microtubule organization and the potential role that it plays in meristem maintenance. 9  Four distinct microtubule arrays are typically associated with dividing cells. They include the interphase cortical array (CMT) that reinforces the tissue axis between divisions, the preprophase band (PPB) that forms in G2 to determine the future division plane, the mitotic spindle that separates chromosomes and finally the phragmoplast, which completes cytokinesis by building the cell plate (Wasteneys, 2002). The interphase cortical microtubule array situated on the periclinal and radial faces is stereotypically described as being transverse to the cell's axis of elongation. While this is certainly common post-mitotically in the elongation zones of organs, it has been noted for some time now that the arrangement of interphase microtubules in cells within the division zone does not conform to this rule (Sugimoto et al., 2000). Recent studies, which we outline below, have identified three mechanisms that explain how these non-transverse patterns are established and hint at the functional relevance in maintaining cells in an undifferentiated state. Computational simulations of array establishment utilizing microtubule dynamics measurements have shown that cellular geometry can predict the orientation of CMTs (Ambrose et al., 2011). In recently divided cells, the newly formed sharp edges are a formidable barrier to microtubules that encounter them, causing microtubules to undergo catastrophe. Consequently, microtubules become aligned parallel to the sharpest edges of cells. The microtubule-associated protein CLASP, which is enriched at the newly formed sharp cell edges, appears to overcome this default orientation pattern. In addition, CLASP is found along microtubules, and is involved in the anchoring of microtubules to the cortex. Paradoxically, more frequent microtubule detachment in clasp-1 mutants increases the likelihood that microtubule encounters will lead to bundle formation, contributing to a hyperparallel organization (Ambrose and Wasteneys, 2008). Thus, in mutants lacking CLASP altogether, microtubules assume transverse orientation very soon after division (Figure 1.2a). Importantly, this is strongly correlated with precocious exit of cells into terminal elongation, resulting in smaller meristems and shorter roots. In effect, the self-organizing capacity of microtubules can be predicted by the dynamic instability of microtubule polymers and the shape of the polyhedron in which they are assembling. By overcoming the catastrophe-inducing effects of the sharp cell edge, CLASP promotes the formation of robust transfacial bundles that account at least in part 10  for the longitudinal microtubules that are seen at the outer periclinal face of root epidermal cells (Figure 1.2b). At least two other mechanisms involving microtubule nucleation contribute to non-transverse microtubule orientation patterns. Components of the γ-tubulin ring complex are found at the newly formed transverse edges, and serve to generate microtubules that polymerize away from the edge in the longitudinal direction (Ambrose and Wasteneys, 2011) (Figure 1.2c). Edge nucleation might aid in CLASP-dependent transfacial bundle initiation and/or maintenance. Consistent with this, GCPs disappear from edges as cells proceed into elongation and this is coincident with transfacial bundle disappearance from the transverse edges (Ambrose and Wasteneys, 2011). Radial microtubule arrays that form at the nuclear surface in early G1 are also recognized as a distinct array, generally thought to precede and help to populate the cortex with microtubules and nucleating complexes. A recent study indicates that the G1 nucleus can also influence the orientation of cortical arrays (Ambrose and Wasteneys, 2014). As shown for epidermal cells of the root and leaf promoridia, immediately after division, perinuclear microtubule nucleating complexes spawn microtubules that, upon reaching the cell cortex, splay outward to form a bipolar array that is generally longitudinal with respect to the cell long axis (Figure 1.2d). Bipolar CMT arrays have been described in other recent studies (Sambade et al., 2012; Vineyard et al., 2013; Pietra et al., 2013) though any relationship between the nucleus and their formation has not been determined. The bipolar CMT array is a transient feature of recently formed cells, and is lost as nucleating activity at the nuclear surface diminishes. Intriguingly, the nucleus becomes active once again in populating the cell with a bipolar array in root hair-forming cells, and this generates the longitudinally aligned cortical microtubules that persist throughout elongation (Ambrose and Wasteneys, 2014). This suggests that the activity of microtubule nucleating complexes can alternate between the nucleus and the cortex in a development-dependent manner.   1.6 The transition to cell division To date, relatively little research has been conducted to assess the significance of the CMT pattern in the CMT-PPB conversion yet this event establishes the cellular geometry 11  that, as discussed above, influences post-mitotic CMT alignment. Recently, Spinner et al. (2013) identified a TTP (TON1-TRM-PP2A with FASS/TON2 as its regulatory subunit) protein complex as a positive effector for the CMT-PPB transition. Previously characterized ton1 and fass mutants failed to form PPBs, yielding severely mis-positioned cell division and a loss of root organization (Camilleri et al., 2002; Azimzadeh et al., 2008). The recently discovered TRMs (TON1 Recruiting Motif), a superfamily comprising 34 members (Drevensek et al., 2012), appear to function in directing TON1 and PP2A onto microtubules where the PP2A complex presumably exerts its phosphatase activity (Spinner et al., 2013). The critical role played by the TTP complex in PPB formation has been confirmed by mutational analysis of PP2A along with other subunits. These mutants exhibit impaired PPBs and cell plate misorientation. This work demonstrates that the TTP complexes found in acentrosomal plant cells are similar in composition to components of animal centrosomes. Importantly, it is still unclear how the TTP complex controls PPB formation but answers will come with the identification of PP2A substrates. Additionally, the large number of TRM members suggests functional diversification and tissue or organ specificity, which will require careful classification. The transmembrane protein SABRE has been identified as a factor in PPB, spindle and phragmoplast orientation as well as in cortical microtubule alignment, and evidence suggests that it acts upstream of CLASP (Pietra et al., 2013). CLASP has previously been shown to associate with mitotic and cytokinetic arrays and its depletion generates a series of disorders such as poorly developed PPBs, diamond-shaped spindles and shorter phragmoplasts (Ambrose et al., 2007). Despite these defects, clasp-1 mutants are still able to complete mitosis and cytokinesis with regular chromosome segregation and cell plate positioning, in contrast to the severe mitotic defects detected in clasp mutants of yeast and animal cells (Inoue et al., 2004; Maiato et al., 2005). How SABRE affects microtubule structures through CLASP remains obscure, regardless of the synergistic phenotype of sab clasp-1 double mutant and the disrupted CLASP deposition in sab mutants during PPB formation. In meristematic cells of the RAM, SABRE is non-uniformly distributed on apical and basal plasma membranes, forming concentrated patches. It would be interesting to investigate the spatial relationship between these patches and the CLASP-enriched areas that give rise to transfacial microtubule bundles. 12  Strikingly, the distorted cell plates observed in sab mutants are similar to those in mutants of TTP components, necessitating a sequential arrangement of TTP, SABRE, and CLASP guided division events.                             Figure 1.2: Non-transverse CMT arrays in post-cytokinetic cells (a) clasp-1 mutants have transverse CMT (red lines) organization coincident with premature entry to elongation phase. The grey colour indicates the newly formed cell plate between adjacent cells.  (b) CLASP enables CMT growth around sharp edges resulting in the formation of transfacial bundles. The blue colour represents CLASP and the adjacent red lines indicate transfacial bundles. (c) γ-Tubulin ring complex-directed sharp edge nucleation genereates CMTs in the longitudinal direction. The organe dots are GCP2/3 proteins, and microtubules growing away from them are shown in red colour. “+” is representative of microtubule plus ends. 13  (d) Endoplasmic microtubules emanating from the nuclear surface-localized GCP2/3 splay outward to produce a bipolar CMT array when they reach the cell cortex. The brown colour designates nuclei, dotted red lines are endoplasmic microtubules and red lines are bipolar CMTs.                         1.7 Conclusions and future directions Maintaining a population of stem cells, and the tissue initial cells that they give rise to, is a fundamental activity in all multicellular organisms. In plants, the number of cells within division zones is a critical determinant of organ size and growth rate. The key to modulating meristem populations appears to be the rate at which cells exit the division pathway and enter terminal growth and/or differentiation. The transition zones that define the border between dividing and elongating cells are now well known to be under the control of hormones, in particular auxin, cytokinin and brassinosteroids, as well as mechanical inputs that are derived both from the expanding organ itself and the environment that it encounters (Sampathkumar et al., 2014). How some of these inputs converge to align microtubules and feedback upon themselves is shown in Figure 1.3.  Although there are undoubtedly many players involved in developmental phase transitions, the microtubule-associated protein CLASP has generated much interest because it is important for both the integration of hormone signals and for fostering the formation of specific microtubule arrays that promote the continuation of cell division. In this chapter we highlighted CLASP's ability to promote the formation of robust transfacial microtubule bundles, the construction of which is strongly associated with keeping cells mitotically active and inhibiting exit into the differentiation pathway. Overlaid with this is the intriguing and apparently plant-specific role that CLASP has in connecting SNX1 endosomes to microtubules, a process that maintains high levels and activity of the auxin efflux carrier PIN2, which is critical for cycling auxin in the root apical meristem. Thus, since auxin flux is a major determinant of meristem identity and size, CLASP has a second, very specific yet critical function in controlling auxin levels. CLASP’s remarkable dual function in meristem activity corresponds closely with its expression, which is strongest in young, actively dividing tissues and tapers off as cells enter terminal differentiation (Ambrose et al., 2007; Kirik et al., 2007). How the expression of CLASP and/or CLASP protein stability is regulated is thus a critical next 14  step for understanding meristem maintenance. Our own ongoing experiments are exploring a possible connection to the brassinosteroid signalling pathway. Intriguingly, the brassinosteroid receptor BRI1, like PIN2, has been identified in proteomic analysis of SNX1 endosomes (Jaillais et al., 2008). In addition, our preliminary investigations indicate that clasp-1 mutants are hyposensitive to BL, just as they are to auxin. It is tantalizing to speculate that CLASP, which despite its similarity to homologues in other eukaryotic lineages (Al-Bassam and Chang, 2011), has evolved a unique function in plants to modulate the cross-talk between the hormones that mediate transitions from division to differentiation.   Figure 1.3: Microtubule and cell wall-based feedback loops in auxin signalling The black pathway indicates the activation of ROP/RIC and thus katanin-mediated microtubule orientation during auxin-induced expansion. CLASP-SNX1 and microtubule-mediated cell wall mechanical and chemical properties consolidate the function of auxin by promoting PIN stability as shown by the red colour. Microtubules reorient in response to mechanical stress (orange), which converge with auxin signalling at the ROP/RIC point. Meanwhile, mechanical stress (both intrinsic and extrinsic) modulates PIN polarity, influencing auxin distribution and action. Cellular geometry is determined initially by cell division planes, which are predicted by the formation of preprophase bands (PPB). This process (green pathway) involves recruitment of TON1 to microtubules by TRM as well as the activity of SABRE, which may act upstream of CLASP.   15  1.8 Thesis objectives The effect of CLASP on microtubule alignment and plant growth has been well studied (Ambrose et al., 2007; Kirik et al., 2007; Ambrose et al., 2011). The primary aim of this thesis is to reveal new pathways in which CLASP might be involved, and to elucidate the long-sought connection between microtubules and hormones, of which a lot still remains elusive. SNX1, identified in two independent yeast two-hybrid screens for candidate CLASP interactors, was shown previously to mediate PIN2 intracellular trafficking (Jaillais et al., 2006). By studying the interaction of CLASP and SNX1, we hope to gain better insights into how microtubules control auxin polar transport through modulating PIN2 abundance. While studying CLASP and auxin, papers published in the Plant Cell (Wang et al., 2012) and Dev Cell (Lanza et al., 2012) attracted our attention. These papers were the first two papers to reveal the role of the cytoskeleton in brassinosteroid-induced elongation. Prior to these studies, BRI1 was found in the same SNX1-endosomes as PIN2 (Jaillais et al., 2008), suggesting they share a common recycling pathway under the regulation of CLASP. These interesting findings impelled me to investigate the link between CLASP and brassinosteroids, leading to my second thesis objective: to understand the function of CLASP in the brassinosteroid signalling pathway. TRM19 was obtained from an immunoprecipitation experiment aiming to identify CLASP interactors. TRM19 has been reported to form a complex with TON1 and PP2A and is required for PPB formation (Spinner et al., 2013). My third objective is to explore the interaction of CLASP and TRM19, and to determine whether they function together within the same complex.             16  CHAPTER 2: CLASP INTERACTS WITH SORTING NEXIN 1 TO LINK MICROTUBULES AND AUXIN TRANSPORT VIA PIN2 RECYCING IN ARABIDOPSIS THALIANA 2.1 Introduction  In plants, microtubules (MTs) and microfilaments are important mediators of growth and development. In response to myriad morphological, developmental, and environmental cues, these cytoskeletal structures undergo dynamic reorganizations that facilitate normal cellular function, identity and differentiation. In many developmental and environmental contexts, plant hormones are important regulators of cytoskeletal organization and function.  The growth hormone auxin modulates essential processes, including pattern formation, meristem identity, and growth responses. Auxin is synthesized primarily in the shoot apex and subsequently moves toward the root, forming concentration gradients. The distribution of auxin in tissues is mediated by plasma membrane-bound carrier proteins, which move auxin into and out of cells. In accordance with the directional transport of auxin, several of these carriers are enriched at cell ends in polarized tissues, thus facilitating directional flux of auxin (Gälweiler et al., 1998).  Numerous studies show that both microfilaments and MTs respond to auxin and appear to play a role in the trafficking of auxin carrier proteins, although the mechanisms involved are not understood (Wasteneys and Collings, 2007). For example, the actin cytoskeleton is not required for maintaining the pre-established polar localization of the auxin efflux carriers PIN-FORMED 1 (PIN1) and PIN2 (Boutté et al., 2006; Geldner et al., 2001; Rahman et al., 2007) but does play a role in repolarization of PINs. This includes repolarization after brefeldin A (BFA)-induced depolarization (Geldner et al., 2001) and gravity-induced redistribution of PIN3 to lateral cell membranes (Friml et al., 2002). Whether or not MTs directly contribute to auxin carrier protein localizations is far less clear. Short-term treatments with oryzalin to remove MTs show little to no effect on PIN1 or PIN2 polarity (Boutté et al., 2006; Geldner et al., 2001), whereas prolonged MT 17  disruption has been reported to induce polarity reversals of PINs (Boutté et al., 2006; Kleine-Vehn et al., 2008b). However, these changes in PIN polarities could be an indirect effect of tissue depolarization, which results from the aberrations in cell division and expansion (Boutté et al., 2006; Wasteneys and Collings, 2007). Similarly, prolonged MT disruption influences cell wall properties, which have been shown to play a role in maintaining PIN polarities (Feraru et al., 2011). Thus, the debate centers on whether effects of MT drugs on PIN polarities are direct (i.e., involving MT-MT interactions) or indirect (i.e., involving downstream signals following MT depolymerization).  We sought to explore the connection between MTs and auxin transport through genetic strategies. Of many MT-affected mutants studied to date, clasp-1 mutants, which lack expression of the CLASP MT-associated protein, appear to be unique. They display a range of phenotypes characteristic of plants with auxin-related defects, including abundant lateral roots, reduced apical dominance, and a reduction in root apical meristem size (Ambrose et al., 2007; Kirik et al., 2007). Moreover, CLASP can help MTs grow around sharp cell edges (Ambrose et al., 2011).  In our current study, we describe a MT-dependent mechanism that controls the abundance of PIN2 in cells. We show that by directly interacting with the retromer component sorting nexin 1 (SNX1), CLASP promotes endocytic recycling of PIN2 and restricts its degradation.   2.2 Results  2.2.1 SNX1 interacts with CLASP In order to find potential CLASP-interacting proteins, we carried out yeast two-hybrid screens using CLASP as bait (Table 2.1). We identified SNX1 as a putative CLASP-interacting protein. SNX1 is a key component of the retromer, which is a conserved multiprotein complex involved in endosomal recycling of membrane proteins (Seaman, 2005). As shown in Figures 2.1A and 2.1B, SNX1 is 402 amino acids in length and has an N-terminal PHOX domain, which confers binding to phosphoinositide-enriched membranes, and a C-terminal BAR domain incorporating a coiled coil, which is responsible for dimerizing SNX1 with SNX2a and SNX2b (Seaman, 2005). In addition to a near full-length SNX1 (amino acids 12-402), two fragments of SNX1 also were also 18  found: the C-terminal half (amino acids 146-402) as well as a small central fragment (amino acids 134-219), which spans the regions between the PHOX and BAR domain, and partially extends into the BAR domain (Figure 2.1B). To confirm CLASP-SNX1 interaction, we performed bimolecular fluorescence complementation (BiFC) using transient expression in Arabidopsis leaf epidermal cells. Control NYFP/CYFP combinations did not produce fluorescence (Figure 2.1C), whereas a positive control using CYFP-PHS1 and NYFP-MPK18 (Walia et al., 2009) confirmed that the transient expression assay system worked reliably (Figure 2.1D). Cotransfection with full-length SNX1-CYFP and CLASP-NYFP produced YFP fluorescence along cortical MTs (Figure 2.1E). The nonuniform, punctate labeling along MTs was characteristic of GFP-CLASP in expanded cells (Ambrose et al., 2007; Kirik et al., 2007).   Table 2.1: Putative interactors with CLASP by Yeast 2-hybrid assay. Global PBS demonstrates the likelihood of interaction. Sorting Nexin has the maximum likelihood (related to Figure 2.1) Protein function Global PBS Accession Sorting nexin A At5g06140 Tetratricopeptide repeat protein B At5g63200 Polyketide cyclase B At5g08720 DNA polymerase C At2g02480 Zinc figure protein D At1g08290 Elongation factor D At5g13650 CoA-methyltransferase D At4g34050 Beta-glucosidase D At1g52400 tRNA reductase D At1g58290  2.2.2 Central region between PHOX and BAR domains of SNX1 confers interaction with CLASP We observed a similar punctate fluorescence pattern along MTs upon cotransfection with CLASP-NYFP and the small central fragment SNX1134-219 (Figures 2.1F and 2.1G), suggesting that the region between the PHOX and BAR domain is the CLASP-interacting domain. We verified this by expressing a GFP-SNX1134-219 reporter construct in wild-type (WT) plants and clasp-1 mutants. GFP-SNX1134-219 decorated MTs and formed large dots in WT, whereas in clasp-1, the GFP-SNX1134-219 fluorescence was cytosolic, with neither the MT nor the dot localization (Figure 2.1H). The dots that were labeled in WT plants 19  did not appear to be vesicles as judged by their static positions and varied sizes/intensities. Taken together, our Y2H, BiFC, and GFP-SNX1134-219 reporter experiments demonstrate that CLASP interacts with SNX1 and that this directs SNX1 to MTs. The more intensive labelling pattern visualized in Figure 2.1 F,G compared to H is possibly due to enhanced CLASP levels resulting from the transgene expression.    Figure 2.1: CLASP interacts with SNX1 (A and B) Structural model of SNX1 generated by Zhang I-Tasser server (A) and schematic (B). PHOX domain, the lipid-binding domain, is the long α helix highlighted 20  red. The interaction site with CLASP is from amino acids 134-219 based on Y2H assay and is labeled green. (C-G) BiFC assay of the interaction between CLASP and SNX1 in Arabidopsis cotyledon epidermal cells. CLASP and SNX1 were fused at their N or C termini to the N or C termini of split YFP (NYFP or CYFP). (C) Control NYFP and CYFP, (D) positive control CYFP-PHS1 and NYFP-MPK18, (E) full-length CYFP-SNX1 and NYFP-CLASP, (F) CLASP-NYFP and CYFP-SNX1134-219, and (G) CYFP-CLASP and SNX1134-219-NYFP. The YFP signals in (F) and (G) is distributed along cortical MTs, resembling the punctate localization pattern of CLASP.  (H) GFP-SNX1134-219 is associated with MTs in WT but not in mutants lacking CLASP (clasp-1). Scale bars, 10 μm.    2.2.3 CLASP686-779 interacts with SNX1 We carried out Y2H assays to identify the precise region of CLASP that interacts with SNX1. Sequence alignment with human CLASP1 revealed conserved as well as variable regions. Based on this sequence information, a series of truncated CLASP constructs were made and used as baits (Figure 2.2A) and SNX1134-219 as prey. The capacity of the yeast AH109 strain to grow on medium without histidine was used as the marker for physical interaction. As a result, we found a restoration of histidine auxotrophy when full-length CLASP was co-transformed with SNX1134-219 (Figure 2.2), further confirming their interaction. Ultimately, a 93-residue sequence (686-779) in the highly variable region of AtCLASP, which shares only 11.1% identity and 19.2% similarity with the region between the crTOG2 and crTOG3 domains of human CLASP1, was identified as the interacting site. As shown in Figure 2.2, in the human CLASP1, this region contains a SxIP (Ser-x-Ile-Pro) motif, which is not present in AtCLASP (Patel et al., 2012) (Figure 2.2). 21    .Figure 2.2: CLASP686-779 interacts with SNX1 (A) Schematic diagram of human CLASP1 and Arabidopsis CLASP domain organization and the truncated versions of CLASP generated for Y2H. TOG domains are grey, SxIP domain is black and the CLIP-interacting domain of HsCLASP is striped. Conserved regions with HsCLASP2, AtCLASP and DmMAST are shown by the arrows on top (Ambrose et al., 2007).  (B) Y2H results. Full-length and truncated CLASP were cloned into pGBKT7 as baits. SNX1134-219 was cloned into pGADT7 as prey. Yeast cells were co-transformed with bait and prey constructs. The combination of baits with the empty prey vector pGADT7 served as the negative controls. Transformed yeast cells were plated onto SD/-Leu/-Trp medium (-L-T) and SD/-Leu/-Trp/-His medium supplemented with 5 mM 3-amino-1,2,4,-triazole (-L-T-H + 3AT).   2.2.4 CLASP stabilizes SNX1 endosomes To address the functional relationship between MT-associated CLASP and retromer-associated SNX1, we analyzed the behavior of SNX1-associated vesicles in clasp-1 null mutants (Ambrose et al., 2007) using a SNX1-GFP reporter previously used to describe SNX1 endosomes (Jaillais et al., 2006). The distribution of SNX1-GFP showed abnormalities at both the tissue and cell levels in clasp-1. SNX1-GFP fluorescence intensity levels were drastically reduced in clasp-1 root tips (Figure 2.3A). In contrast to 22  the normal punctate SNX1-GFP vesicles seen in WT, SNX1-GFP fluorescence in clasp-1 was diffuse, and vesicles were less numerous and varied in shape and size (Figures 2.3B and C). This resulted in an increased ratio of cytosolic-to-vesicular fluorescence. Quantification showed an approximate 75% reduction in vesicle number (Figure 2.3D) and an approximate 40% reduction in the endosomal signal-to-cytosolic background ratio (Figure 2.3E). SNX1-GFP vesicles in clasp-1 also displayed decreased lifetimes as assessed by their residence times in 4D (XYZT) confocal time series (Figure 2.3F). In some cases, clasp-1 root cells contained enlarged bodies of SNX1-GFP, often surrounding the nuclei. These, however, were difficult to distinguish from the autofluorescence seen when using higher-sensitivity imaging.  Depolymerization of MTs with oryzalin showed clasp-1-like effects on SNX1 endosomes. Short-term treatments with relatively high concentrations (10-30 min, 100 μM oryzalin), designed to avoid potential side effects on vesicle properties, induced a marked reduction in endosome number and SNX1-RFP fluorescence and increased cytosolic background fluorescence relative to mock-treated controls (Figure 2.3G). Hirano et al. (2015) also reported the dissociation of SNX1-mRFP from endosome membrane into cytosol. Taken together, these data indicate a role for CLASP and MTs in maintaining SNX1 vesicle morphology and stability.        23       Figure 2.3: CLASP and MTs stabilize SNX1 vesicles (A-C) Root tip cells of WT and clasp-1 plants expressing pSNX1::SNX1-GFP. Visualization of SNX1-GFP in clasp-1 root tips requires increasing brightness (postacquisition) for morphology comparisons. (A) Sample region of root tips pseudocolored for comparison (original and increased brightness images shown for clasp-1). (B) SNX1-GFP in WT and clasp-1 mutant root tips. SNX1-GFP in clasp-1 root tips is increased in brightness for visibility. In clasp-1, diffuse SNX1-GFP signal is higher relative to vesicular signal than for WT, and vesicles are less numerous. Gray, SNX1-GFP; blue, propidium iodide. Arrowheads point to typical robust vesicle in WT, and typical weak vesicle in clasp-1. (C) Root elongation zone of WT versus clasp-1 overexpressing SNX1-GFP. Fluorescence is diffuse in clasp-1, and vesicles are less numerous. Arrowheads point to vesicle in WT, and diffuse region in clasp-1. (D-F) clasp-1 mutants exhibit reduced vesicle (ves.) number compared to WT (p ≤ 0.0001, t test; n = 8 WT roots [80 cells total] and 10 clasp-1 roots [100 cells total]) (D) 24  reduced signal-to-noise ratio (p ≤ 0.0001, t test; n = 4 WT roots [38 cells total] and 4 clasp-1 roots [35 cells total]) (E) and shorter lifetimes (p = 0.0013, t test; n = 3 WT roots [52 vesicles tracked] and 3 clasp-1 roots [65 vesicles tracked]) (F) Data are taken from root tip epidermal division zone cells. All data are mean ± SEM.  (G) Abnormal SNX1-RFP vesicles in WT roots treated with oryzalin (100 μM, 20 min). Control is 0.1% DMSO for 30 min. Scale bars, 5 μm.  2.2.5 SNX1 endosomes associate with CLASP and MTs Given the requirement of CLASP and MTs for SNX1 vesicle stability, we predicted that SNX1 vesicles would associate with CLASP and MTs. To test this, we stably co-expressed full-length SNX1-RFP along with either GFP-CLASP or the MT reporter GFP-TUB6. SNX1-RFP and GFP-CLASP expression was driven by their native promoters (Ambrose et al., 2011; Jaillais et al., 2006), whereas the GFP-TUB6 expression was driven by the constitutive promoter from cauliflower mosaic virus 35s (Nakamura et al., 2004). Using 4D time-lapse imaging, we found a close and dynamic association between the SNX1-RFP-labeled vesicles and GFP-TUB6-labeled MTs, which are pseudocolored green and red, respectively, in Figure 2.4A for better contrast. Vesicles typically remained in contact with MTs and hopped between them. Tubulations along MTs were also common (indicated by lines in Figure 2.4A). Using the endosomal dye FM4-64, we observed an endosomal association with MTs similar to that observed with SNX1 vesicles (Figure 2.5).  The association between MTs, CLASP and SNX1 vesicles was particularly evident in cells with high levels of CLASP protein accumulation on prominent MT bundles. Specifically, SNX1-RFP colocalized with GFP-CLASP along the coiled MT bundles that form in leaf epidermal cells of the guard cell lineage (Figure 2.4B) (Ambrose et al., 2011; Ambrose et al., 2007; Kirik et al., 2007) as well as at the prominent transfacial MT bundles at recently formed cell edges (Ambrose et al., 2011) (Figure 2.4C). Similarly, taxol treatment generated prominent cortical MT bundles along which GFP-CLASP and SNX1-RFP colocalized (Figure 2.4D). As with GFP-TUB6, the association between SNX1-RFP vesicles and GFP-CLASP was dynamic. In contrast to the complete labeling of MTs by GFP-TUB6, GFP-CLASP 25  typically decorates MTs nonuniformly (Ambrose et al., 2007; Kirik et al., 2007). Analysis of time series revealed that SNX1-RFP vesicles were found overlapping or adjacent to GFP-CLASP domains, where they stayed static or moved around within the vicinity of the associated GFP-CLASP. Similarly, when GFP-CLASP regions moved or changed shape, the accompanying SNX1-RFP vesicle(s) moved with the GFP-CLASP (Figure 2.4E). To quantify the SNX1-RFP vesicle GFP-CLASP association, we used several criteria to define association: (1) vesicles must exhibit matching movement or morphological changes, and (2) static/overlapping vesicles must remain for 60 consecutive seconds or greater. Using these criteria for root division zone cells, we found that 89.5% ± 1.9% of SNX1-RFP vesicles associated with GFP-CLASP (Figure 2.4F; n = 25 cells, 262 vesicles). These results indicate that CLASP’s role in stabilizing SNX1 vesicles is via a direct, dynamic association between MT-associated CLASP and SNX1 vesicles.      26  Figure 2.4: SNX1 endosomes associate with CLASP and MTs (A) Time series of GFP-TUB6-labeled MTs (pseudocolored red for better contrast) and SNX-RFP vesicles (pseudocolored green for contrast). Arrowheads indicate stably associated vesicles. White/yellow lines depict vesicle tubulations along MTs. See also Figure 2.4. (B) Cotyledon cells of the guard cell lineage showing filamentous SNX1-RFP at the GFP-CLASP cortical coils. Arrowheads indicate overlap along filamentous structures. (C) Recently divided cotyledon guard cells showing filamentous SNX1-RFP at the GFP-CLASP stubs at the newly formed cross-wall edge. Arrowheads indicate overlap along filamentous structures.  (D) Taxol treatment (10 μM, 2 hr) shows punctate/filamentous SNX1-RFP colocalized with cortical GFP-CLASP structures in hypocotyls. Arrowheads show examples of colocalized areas.  (E) Time series showing matched dynamic morphological/positional changes between GFP-CLASP and SNX1-RFP. Overlap is near complete at the punctate structure, from which a thin filament (arrowhead) emanates. The filament rotates counterclockwise over time. Other associated punctae are marked with arrowheads. Panel at right is the full 90 s compressed along x axis to emphasize comovement.  (F) Quantification of SNX1-RFP vesicle/GFP-CLASP association. Associated (Assoc.) vesicles were defined as described in the Results (n = 5 roots, 25 cells, and 262 vesicles). p < 0.01, Student’s t test. Scale bars, 5 μm (A-D) and 2.5 μm (E). Time series intervals are 5 s in (D) and 2.5 s in (E).                  Figure 2.5: Association of FM4-64 endosomes with MTs Single time point of hypocotyls cell expressing GFP-TUB6 (green; right panel) and FM4-64 (red; left panel). Merged image shown in middle panel. Arrowheads indicate MT-associated vesicles. Bar, 5 μm.  27  2.2.6 SNX1 vesicles form dynamic clusters that associate with GFP-CLASP We observed that most SNX1 vesicles were not solitary but instead formed tubulo-vesicular clusters that associated with GFP-CLASP- and GFP-TUB6-labeled MTs (Figure 2.6). These vesicular clusters exhibited highly dynamic morphology and movement, with frequent splitting, fusion, tubulation, and formation of interconnecting protrusions. Their appearance and behavior are consistent with that of sorting endosomes in animal cells, wherein dynamic tubulations and budding facilitate recycling of membrane cargoes (Maxfield and McGraw, 2004).  We observed that the vesicular clusters typically changed shape and moved around in relation to GFP-CLASP in a manner reminiscent of groups of balloons on strings tethered to a single anchor point. This dynamic tethering is best illustrated in the time series shown in Figure 2.6D, which shows vesicle cluster/CLASP association. CLASP-associated vesicle clusters can also be shown by projecting all time points into a single image, which shows red (SNX1-RFP) clusters of varying size overlapping with or peripheral to GFP-CLASP domains (Figure 2.6A). To quantify the spatiotemporal relationship between SNX1 vesicles and GFP-CLASP, we tracked the movements of RFP-SNX1 vesicles and mapped their paths onto the corresponding GFP-CLASP channel (Figure 2.6B). We found that 91.1% ± 1.5% of SNX1 vesicles resided within vesicle clusters as opposed to being free (Figure 2.6C; n = 27 cells, 198 vesicles). The montage in Figure 2.6D shows the association between GFP-CLASP and SNX1-RFP vesicles over time. These data suggest that the SNX1 vesicle cluster/CLASP association functions to stabilize SNX1 vesicles, thereby preventing both their accumulation in large aggregates and their dissociation from MTs.           28                            Figure 2.6: SNX1 forms CLASP-associated vesicular clusters (A) Time series projections of GFP-CLASP (green) and SNX1-RFP (red). Clusters appear as large lumpy clusters adjacent to the stable GFP-CLASP domains. Three examples are shown. Total time for each projection is 120 s. Arrowheads correspond by color to the colored traces shown in (B). (B) The same time series projection of GFP-CLASP as in (A) but with SNX1 vesicle life histories mapped. Each colored track represents a different vesicle. (C) Percentage of SNX1 vesicles residing within vesicle clusters. Bars are SE (n = 27 cells, 197 vesicles).  (D) Time series of SNX1-RFP vesicle clusters and GFP-CLASP in root division zone cells. Top row is time projection of the entire 120 s series. Arrowheads indicate individual vesicles within a cluster. Vesicle paths are marked with colored lines in right column. Arrow color in left column matches colored paths in right column.  Scale bars, 5 μm. 29  2.2.7 clasp-1 mutants exhibit enhanced PIN2 degradation but retain PIN2 polar distribution Based on the observed accumulation of SNX1 in aggregates in clasp-1 mutants and the observation of dynamic CLASP-associated SNX1 vesicles, we hypothesized that CLASP-mediated stabilization of SNX1 vesicles assists in the recycling of membrane cargo proteins. To test this, we used the auxin transporter PIN2, which is a known SNX1 endosome cargo protein (Jaillais et al., 2006; Kleine-Vehn et al., 2008c). Using pPIN2:GFP-PIN2 and anti-PIN2 immunostaining, we found that the size of the root tip PIN2 expression domain was strongly reduced, and fluorescence intensity levels were much weaker in clasp-1 root tips ( Figures 2.7A, 2.7B, 2.8A, and 2.8B). However, PIN2 retained its polarized distribution in clasp-1 root tips (Figures 2.7C and 2.7D). The reduced PIN2 expression domain size prompted us to determine if this reduction resulted from reduced PIN2 gene expression. To this end, we performed RT-PCR using cDNAs prepared from root tips of WT and clasp-1 mutants. No apparent differences in transcript levels were found (Figure 2.7E). Protein degradation is a well-characterized mechanism of regulating PIN2 protein levels under a variety of conditions (Kleine-Vehn et al., 2008c). Consistent with this, we frequently observed accumulation of PIN2-GFP in lytic vacuoles in clasp-1 root tips (Figure 2.7F). Because PIN2 is required for root gravitropism, we asked whether clasp-1 mutants show gravity defects. We found no impairment of downward bending in horizontally positioned clasp-1 roots, suggesting that the reduced PIN2 levels are adequate to drive gravitropism (Figure 2.8C). Double clasp-1 pin2 mutants resembled clasp-1 mutants but, as with pin2, were agravitropic (Figure 2.8C). Contrary to a previous report by Jaillais et al. (2006), snx1-1 mutants exhibited normal root gravitropism. Given that snx1-1 mutants show a mild reduction in PIN2 levels (Kleine-Vehn et al., 2008c), these observations suggest that root gravitropism can occur as long as some PIN2 is present and properly polarly localized. To explore the specificity of CLASP’s role in PIN protein targeting, we analyzed the distribution of PIN1 in clasp-1. Using immunofluorescence, we found that PIN1 levels were similar to those observed in WT root tips (Figure 2.7G). 30  Taken together, our findings indicate that the PIN2 polar-targeting machinery remains in clasp-1 but that degradation of PIN2 is enhanced. The data indicate a role for CLASP in PIN2 trafficking, most likely via SNX1/retromer-mediated prevention of degradation.     Figure 2.7: PIN2 expression domain and levels are reduced in clasp-1 root tips (A) Reduced expression domain and fluorescence levels of PIN2-GFP in the clasp-1 mutant background. Boxed areas indicate primary expression domains. Images are pseudocolored for contrast. (B) Reduced PIN2-GFP levels in primary clasp-1 root tips. Maximum projections of 40 confocal slices are shown. (C) PIN2 retains polarized distribution in clasp-1 root tips. Anti-PIN2 immunolabeling of root tip epidermal cells. (D) Fluorescence intensity plot corresponding to dotted lines in (C). Red, clasp-1; blue, WT.  31  (E) PIN2 gene expression is normal in clasp-1 root tips. RT-PCR results from three separate biological replicates (clasp-1 lanes 1-3; WT lanes 4-6). Control is ubiquitin (UBQ1). (F) PIN2-GFP accumulation in enlarged spherical structures in clasp-1 root tips (arrowhead). (G) PIN1 immunostaining in WT and clasp-1 root tips.  Scale bars, 100 μm (A) and 5 μm (B, C, F, and G).                     32  Figure 2.8: PIN2 levels and PIN2-mediated gravitropic responses in clasp-1 mutants (A-B) PIN2 immunostaining in WT and clasp-1 root tips. (A) greyscale images. (B) Pseudocolored images to show intensity variations. (C) Gravity responses. Plants were grown for 5 days and rotated 90 degrees, then imaged after 3 days, as shown in diagram. clasp-1 mutants respond normally, and clasp-1 pin2-1 double mutants are additive. pin2-1 controls show agravitropism. snx1-1 mutants showed normal gravitropism, comparable with WT.  Scale bars in A and B, 100 μm. Immunofluorescence was performed on whole-mounted roots as in Ambrose et al. (2007). Anti-PIN2 and anti-PIN1 antibodies were diluted 1:200 in 3% BSA solution.   2.2.8 MTs inhibit degradation of PIN2 to facilitate recycling The enhanced PIN2 degradation in clasp-1 mutants prompted us to test whether removal of MTs would also induce PIN2 degradation. We treated plants expressing PIN2-GFP with oryzalin to remove MTs using short-term treatments to avoid potential side effects of long-term MT removal. Because CLASP-dependent MT bundles are resistant to oryzalin (Ambrose et al., 2007; Kirik et al., 2007), we used 50 μM oryzalin for 60-90 min to completely remove CLASP-dependent MT bundles. Strikingly, oryzalin-treated plants showed accumulation of PIN2-GFP within enlarged spheroid structures 4 ± 0.2 μm in diameter within 60 min (Figure 2.9A). The size and frequency of the structures match that of lytic vacuoles (Figure 2.9B) (Kleine-Vehn et al., 2008c). Long-term treatments at lower concentrations of oryzalin (10 μM for 16-20 hr) also induced accumulation into bodies, but to a lesser extent (data not shown). Because clasp-1 mutants contain normal MTs except for the lack of CLASP-dependent MT bundles, and low oryzalin does not remove CLASP-dependent bundles, the data suggest a distinct role for these bundles in inhibiting PIN2 degradation.              33                                                  Figure 2.9: Depolymerization of MTs with oryzalin induces PIN2-GFP accumulation within lytic vacuoles (A) Confocal images of PIN2-GFP in WT plants treated with 0.5% DMSO or 50 μM oryzalin for 90 min. Arrowheads indicate polarized PIN2-GFP. Arrow indicates lytic vacuoles. Scale bar, 5 μm. (B) Increased number of lytic vacuoles per cell under oryzalin treatment. Bars indicate SEM (n = 4 roots each, 100 oryzalin-treated cells, and 126 DMSO control cells).  2.2.9 clasp-1 mutants exhibit auxin-related phenotypes Previous reports that clasp-1 mutants are dwarf and display excessive root branching (Ambrose et al., 2007; Kirik et al., 2007) suggest the presence of auxin-related defects. Based on the observed PIN2 depletion in clasp-1, we hypothesized that CLASP controls some aspects of auxin transport. To investigate this possibility, we tested how clasp-1 34  mutant plants respond to exogenously applied auxins and auxin polar transport inhibitors. Over a wide range of concentrations, clasp-1 root growth was less sensitive than that of WT to the growth inhibitory effects of indole-3-acetic acid (IAA) (Figures 2.10A-C), 2,4-D, and NAA. When treated with the auxin transport inhibitor NPA, clasp-1 mutants were more tolerant than WT, as evidenced by clasp-1 plants growing to larger size, forming large root calli and narrow leaves (Figure 2.10D). Similarly, calli occasionally formed on hypocotyls and roots of clasp-1 seedlings under hormone-free conditions, particularly when grown in the dark (Figure 2.10E). These abnormalities are suggestive of anomalies in auxin-response maxima (Chandler, 2008; Kawamura et al., 2010). We tested this directly by using the DR5rev:GFP reporter of auxin activity and found that auxin-response distribution is aberrant in clasp-1 root tips. The DR5:GFP expression domain was shorter than in WT and often extended up into the lateral root cap, in contrast to confinement of GFP expression to the columella of WT root caps (Figure 2.10F). This distribution pattern was also observed with DR5:GUS (Figure 2.11A). The DR5:GFP expression pattern was also altered in the cotyledons of clasp-1 mutants. DR5:GFP fluorescence was higher throughout both faces of the clasp-1 cotyledons and frequently was found to accumulate at tooth-like projections around the cotyledon edge (Figures 2.10G and 2.11B).  Expression of CLASP was not responsive to treatment with auxins (Figure 2.10H). Similarly, the expression of the auxin-responsive gene IAA5 did not change in clasp-1 seedlings, either under normal conditions or following treatment with NAA (Figure 2.10I), suggesting that auxin levels were not globally elevated and that the transcriptional response to auxin is not altered by the clasp-1 mutation. With respect to auxin-dependent tissue patterning and growth responses such as leaf venation, gravitropism, phototropism, and rooting from cuttings, clasp-1 mutants did not differ from WT. Taken together, these findings suggest that CLASP functions in a subset of auxin responses.    35                         Figure 2.10: The clasp-1 mutant displays auxin-related defects (A and B) clasp-1 exhibits reduced sensitivity to IAA-induced root growth inhibition compared to WT. (A) WT and clasp-1 plants grown for 7 days on 10 nM IAA. (B) Primary root lengths as a percentage of control for WT and clasp-1 roots grown for 7 days on 10 nM IAA. (C) Dose-response curve over a range of IAA concentrations.  (D) clasp-1 plants grow to larger sizes on 50 μM NPA. (E) Callus formation on dark-grown clasp-1 seedlings grown on hormone-free medium. Arrowheads indicate calli. 36  (F) Distribution of DR5:GFP in root tips of WT and clasp-1 plants. (See also Figure 2.10A)  (G) Distribution of DR5:GFP in cotyledons of WT and clasp-1 plants. Top images are grayscale; bottom images are pseudocolor to highlight intensity variations. Arrowheads indicate tooth-like projections around the cotyledon edge in clasp-1. (See also Figure 2.10B)  (D) clasp-1 plants grow to larger sizes on 50 μM NPA. (H and I) CLASP gene expression is unaffected by treatment with auxins, and clasp-1 has normal levels and responses of auxin-responsive genes. (H) NAA. (I) IAA. Scale bars, 50 μm. All data are mean ± SEM.     Figure 2.11: DR5:GUS in roots and DR5:GFP in clasp-1 cotyledons exhibit abnormalities (A) Confinement of DR5:GUS to smaller apical zone in clasp-1 root tips. Brackets indicate size of expansion zone. Arrows indicate ectopic expression along lateral root cap in clasp-1.  Scale bar, 10 μm. (B) Enhanced DR5:GFP fluorescence in clasp-1 cotyledons. Images are taken of both sides of each example. Top panel uses grey pixel scale, bottom panel uses rainbow pseudocoloring to enhance contrast.    37  2.3 Discussion Here, we identify a role for MTs and CLASP in controlling SNX1 vesicle morphology and behavior, which links MT function to auxin growth responses. Based on our findings, we propose the following hypothetical model. CLASP and MTs function in PIN2 recycling, but not in PIN2 polarity determination. Recycling to the plasma membrane is achieved by CLASP-dependent recruitment of SNX1/PIN2 vesicles to cortical MTs, thereby increasing vesicle residency time close to the plasma membrane to favour recycling. Importantly, this mechanism provides a means by which changes in the MT cytoskeleton could be used to modulate PIN2 levels. Because MTs change configuration in response to many stimuli (stress, hormones, developmental state), these changes may result in changes in PIN2 abundance and hence polar auxin transport.  Our model is consistent with previous studies on PIN trafficking. SNX1 has been shown to function in maintaining PIN levels, but not polarity (Kleine-Vehn et al., 2008a). Under conditions that perturb SNX1 localization, including wortmannin treatment (Jaillais et al., 2007) and mutation in the retromer component VPS29 (Jaillais et al., 2007), PIN2 remains polarized, whereas some is internalized within intracellular aggregates. ARF-GEFs play a key role in establishing PIN polarity, which results from selective endocytosis following uniform delivery to the plasma membrane. The ARF-GEF inhibitor BFA causes rapid internalization of PINs and loss of polarity (Geldner et al., 2003). Longer treatments result in polarity reversals of PIN1 and PIN2 (Kleine-Vehn et al., 2008a).  The aberrant DR5-based expression patterns in clasp-1 are consistent with the inability to effectively transport auxin and establish normal auxin maxima and gradients. Moreover, the particular auxin maxima abnormalities in clasp-1 are characteristic of plants with aberrant PIN2 distribution. Removal or disruption of PIN2 causes ectopic DR5-reporter expression in lateral root cap cells surrounding the meristem (Jaillais et al., 2006, 2007; Sabatini et al., 1999). The reduction in size and intensity of the PIN2/SNX1 expression domain in root tips is consistent with anomalies in polar auxin transport in clasp-1 mutants.  The identification of a direct link between CLASP and SNX1 explains why SNX1 was identified in a screen for tubulin-binding proteins in Arabidopsis (Chuong et al., 38  2004). Although CLASP homologs in animal and yeast cells are known to function in MT-dependent vesicle trafficking (Chiron et al., 2008; Efimov et al., 2007; Lowery et al., 2010), SNX trafficking appears to involve other factors. In mammalian cells, SNX endosomes require MTs for normal formation and function through direct binding of SNX5 and SNX6 to the dynein/dynactin motor complex on MTs (Hong et al., 2009; Wassmer et al., 2009). Additionally, kinesin motor proteins have been shown to play a role in sorting endosome morphology and budding (Schmidt et al., 2009; Skjeldal et al., 2012). The SNX1-RFP vesicular clusters observed in our analysis closely resemble the recently described secretory vesicle clusters in plant cells (Toyooka et al., 2009). Thus, CLASP might have adopted a unique role in plants for MT-dependent regulation of sorting endosomes. In all eukaryotes, sorting endosomes play a key role in determining the fate of endocytosed cell surface receptors, channels, and other proteins (Johannes and Wunder, 2011). In plants, PINs have been identified as major targets (Jaillais et al., 2006; Johannes and Wunder, 2011). Following endocytosis, two major routes are taken, including targeting to the vacuole/lysosome for degradation (BLOC-1 route) or recycling back to the plasma membrane (SNX route) (Cui et al., 2010). BLOS1, the Arabidopsis BLOC-1 complex, has been shown to interact with SNX1 by yeast two-hybrid and BiFC methods (Cui et al., 2010). RNAi-induced reduction of BLOS1 levels results in increased levels of PIN1 and PIN2, indicating BLOS1-mediated degradation (Cui et al., 2010). Presumably, by reducing the BLOS1-mediated degradation of PINs, the BLOS-1 RNAi treatment increases recycling of PINs to the plasma membrane via SNX1. Taken together, our data provide a possible mechanism by which changes in MT organization influence PIN protein abundance and auxin distribution. Human CLASP1 contains an N-terminal tumor over-expressed gene (TOG1) domain, two cryptic TOG (crTOG2 and 3) domains, a middle Ser-x-Ile-Pro (SxIP) motif and a C-terminal coiled-coil domain (Patel et al., 2012). In vitro biochemical analysis revealed that TOG1 and crTOG2 bind to the MT lattice with high affinity while crTOG3 showed a relatively weaker binding. The SxIP motif confers plus end tracking through binding to EB1. The C-terminal coiled-coil domain is for dimerization as well as associating with other binding partners including CLIP-170 and CENP-E (Hannak and Heald, 2006; Patel 39  et al., 2012). BLAST analysis showed that the homology between HsCLASP and AtCLASP extends throughout the entire length of the protein and is mainly concentrated in the TOG domains and the C-terminal coiled-coil domain (Figure 2.2), suggesting these regions are functionally conserved. The SNX1-interacting region of AtCLASP was mapped to amino acids 686-779. This is a highly variable region among CLASP homologues. Interestingly, this region of HsCLASP contains an SxIP motif that is responsible for EB1 association, and suggested to confer plus end tracking (Hannak and Heald, 2006). These results elucidate the unique interaction between CLASP and SNX1 in plants and call for a phylogeny study to determine when CLASP sequences diverged and the SNX1-interaction, which may be unique to plant lineages, appeared during evolution.      2.4 Methods 2.4.1 Plant materials and growth conditions Arabidopsis thaliana Columbia ecotype plants were grown in continuous light conditions on vertical agar plates containing Hoagland’s medium. Young expanding cotyledon cells were imaged at 3-4 days. Root tips were used for dividing and expanding root cells. For all comparisons between genotypes, fluorescent marker genes were homozygous for both genotypes to ensure accuracy of comparison and measurement.   2.4.2 Microscopy and image analysis Images were acquired on a Zeiss Pascal scanning confocal microscope, or with a PerkinElmer spinning-disk microscope. Images were processed using ImageJ software (, and figures were assembled using CorelDraw software. For analysis of RFP-TUB6 in root tissues, we drove expression under the ubiquitin 1 promoter (Ambrose et al., 2011). DR5:GUS staining was performed as previously described by Sessions et al. (1999). For quantification of SNX1-GFP vesicles in WT and clasp-1 mutants, vesicles were defined as small punctae of <1 μm. The large aggregations in clasp-1 were excluded for quantification of vesicles. Brightness was linearly increased using contrast in Image J.   40  2.4.3 Drug treatments Plants were grown upright on Hoagland’s media as described above. For treatment, seedlings were gently removed from plates and placed in small wells containing 10 or 100 μM oryzalin (made fresh from 10 mM DMSO stock) or 0.1% DMSO. Treatments were typically 10-50 min and are indicated within the figure legends. Using GFP-CLASP to visualize MTs, even after 30 min in 10 μM oryzalin, small GFP-CLASP MT stubs remained. These stubs are the remnants of the superstable transfacial bundles recently described (Ambrose et al., 2011) and are undetectable with fluorescent tubulins due to intense cytosolic background in oryzalin-treated cells. Therefore, complete removal of these stubs under short-term conditions required 100 μM oryzalin for at least 20 min (during which time cells remained viable as assessed by cytoplasmic streaming). For microscopy, roots were cut off at the hypocotyl junction and placed within coverslip chambers (Nunc) and covered with a thin layer of solidified 1%-1.5% bacto agar to hold them in place. Under these imaging conditions, roots stayed healthy for over 8 hr (as assessed by DIC or fluorophor localization patterns).  2.4.4 Yeast Two-Hybrid analysis Two yeast two-hybrid screens were conducted. We first used the Gal4BD-fused CLASP proteins as bait proteins to screen the Arabidopsis cDNA library (derived from flowers, siliques, and green seeds). The screening procedure was performed by mating the bait proteins with pretransformed frozen prey-library yeast cells (Soellick and Uhrig, 2001). The second yeast two-hybrid screen was performed by Hybrigenics (Paris) ( The coding sequence for the full-length Arabidopsis thaliana CLASP (GenBank accession number 42570285) was PCR amplified and cloned into pB27 as a C-terminal fusion to LexA (N-LexA-CLASP-C) and into pB6 as a C-terminal fusion to Gal4 DNA-binding domain (N-Gal4-CLASP-C). The constructs were checked by sequencing and used as a bait to screen a random-primed Arabidopsis thaliana seedlings cDNA library constructed into pP6. pB27, pB66, and pP6 derive from the original pBTM116 (Vojtek and Hollenberg, 1995), pAS2ΔΔ (Fromont-Racine et al., 1997), and pGADGH (Bartel et al., 1993) plasmids, respectively. 41  For the LexA bait construct, 67 million clones (7-fold the complexity of the library) were screened using a mating approach with Y187 (matα) and L40 Δ Gal4 (mata) yeast strains as previously described by Fromont-Racine et al. (1997). For the Gal4 construct, 74 million clones (8-fold the complexity of the library) were screened using the same mating approach with Y187 (matα) and CG1945 (mata) yeast strains. A total of 46 His+ colonies were selected on a medium lacking tryptophan, leucine, and histidine. The prey fragments of the positive clones were amplified by PCR and sequenced at their 5’and 3’ junctions. The resulting sequences were used to identify the corresponding interacting proteins in the GenBank database (NCBI) using a fully automated procedure. A confidence score (predicted biological score) was attributed to each interaction as previously described by Formstecher et al. (2005). In the screen performed by Hybrigenics, SNX1 was the only protein ascribed a confidence score of A, suggesting a high likelihood of a genuine interaction. In the other screen, SNX1 was also identified as the strongest interactor. To map the SNX1-interating domain on CLASP, cDNAs of full-length CLASP and truncations were fused to the GAL4-DNA binding domain in the bait vector pGBKT7 (Clontech) and SNX1134-219 was expressed as the GAL4-DNA activation domain fusion protein in the prey vector pGADT7 (Clontech). pGBKT7 and pGADT7 contain trypotophan (Trp) and leucine (Leu) selection marker, respectively. The yeast strain AH109 co-transformed with the resulting bait and prey constructs were plated onto SD/-Leu/-Trp medium and SD/-Leu/-Trp/-His medium with 5 mM 3-amino-1,2,4,-triazole and grew at 30 °C for 3 d. Growth on SD/-Leu/-Trp medium demonstrated successful co-transformation. Positive interaction activated histidine synthesis which is indicated by growth on SD/-Leu/-Trp/-His medium. 3-amino-1,2,4,-triazole was added to suppress the leaky biosynthesis of histidine. Primers used for cloning are listed in Table 2.2   2.4.5 RNA extraction following hormone treatments WT Arabidopsis thaliana seeds were grown on Hoagland’s medium in 1.2% agar. They were grown for 8 days at 21°C and subjected to 24 hr light. Prior to RNA extraction, seedlings were transferred to 1.2% agar plates containing Hoagland’s media and an IAA concentration of 10 μM for 10 hr. A control group was established that contained seeds 42  transferred to plates containing Hoagland’s medium only (no IAA) for an equivalent time period. Following treatment, root tips were excised and immediately flash frozen in liquid nitrogen. NAA treatments consisted of 4 hr in a 0.1% DMSO solution containing 100 μM NAA or no NAA. Total RNA was extracted using the RNeasy Plant Mini Kit (QIAGEN) and tested for quality and quantity with the NanoDrop 2000 (Thermo Scientific, Wilmington, DE, USA). Each RNA concentration was normalized with ddH20, and cDNA was then synthesized using Superscript III Reverse Transcriptase (Invitrogen). PCR was used to amplify the cDNA transcripts using Taq DNA Polymerase (QIAGEN) and PIN2 primers. cDNA amplification was carried out for 25 cycles at an annealing temperature of 59°C. Primers used are listed in Table 2.2.  2.4.6 BiFC analysis Full-length SNX1 and CLASP cDNA (with and without stop codon) were PCR amplified and cloned into pDONR221 entry plasmid through GATEWAY LR reactions (Invitrogen), following the manufacturer’s protocol. The PCR products were sequenced and recombined through BP reaction into BiFC destination plasmids pUBC-YN/YC and pUBN-YN/YC (a gift from C. Grefen from Plant Science Group, Glasgow, UK), which encoded the two parts of enhanced YFP-YN (residues 1-154) and YC (residues 155-238), respectively, generating fusion proteins with YN or YC at N or C termini (YN-CLASP, YC-CLASP, CLASP-YN, CLASP-YC, YN-SNX1, YC-SNX1, SNX1-YN, and SNX1-YC). The amino acid residues 134-219 of SNX1 were chosen as the partial sequence used for the BiFC assay, based on the prediction of interaction from Y2H. The cloning procedures were the same as above, and primer sequences are shown in the Supplemental Experimental Procedures section. The various binary plasmids were then transformed into A. tumefaciens (GV3101) using the Freeze-Thaw method. Primers for generation of BiFC constructs are listed in Table 2.2. Our BiFC assay was performed as described by Li et al. (2009). Plant cotyledons were examined using a PerkinElmer spinning-disk confocal microscope. YFP signals were detected using a 540 nm emission filter. Typical scan times were 5 s; slice thickness was 0.5 μm. 43  2.4.7 Immunofluorescence of PINs in roots Immunofluorescence was performed on whole-mounted roots as in Ambrose et al. (2007). Anti-PIN2 and anti-PIN1 antibodies were diluted 1:200 in 3% BSA solution.                             44  Table 2.2: Primers used in this study Primer Sequence (5’ to 3’) PIN2RT-F CGTCCATCCTGATATTCTCAGCAC (Cui et al., 2010) PIN2RT-R CATACACCTAAGCCTGACCTGGAA (Cui et al., 2010) UBIQUITINRT-F GCGCGACTGTTTAAAGAATACAAAGAG (Chen et al., 2008) UBIQUITINRT-R TCACCAGATCTTAGAAGATTCCCTGAGT (Chen et al., 2008) CLASPRT-F GACCCTGTCATTCAAAGG CLASPRT-R AGCAACAGCATGGAAAGG IAA5RT-F AACCGGCGAAAAAGAGTC IAA5RT-R GGAACATTTCCCAAGGAAC ACTIN8RT-F ATTAAGGTCGTGGCA (Walia et al., 2009) ACTIN8RT-R TCCGAGTTTGAAGAGGCTAC (Walia et al., 2009) SNX1-attB1  GGGGACAAGTTTGTACAAAAAAGCAGGCTTCATGGAGAGCACGGAGCAGCC SNX1-attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTTTAGACAGAATAAGAAGCTTCAAGT SNX1-attB2-noSTOP GGGGACCACTTTGTACAAGAAAGCTGGGTCGACAGAATAAGAAGCTTCAAGT CLASP-attB1 GGGGACAAGTTTGTACAAAAAAGCAGGCTAAATGGAGGAAGCTTTAGAAAT CLASP-attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTTCAGGTGTCTGCGTCGATAGGGGC CLASP-attB2-noSTOP GGGGACCACTTTGTACAAGAAAGCTGGGTTGGTGTCTGCGTCGATAGGGGC SNX1134-219-attB1 GGGGACAAGTTTGTACAAAAAAGCAGGCTTCATGACCTTTTTGCAAGCAGA SNX1134-219-attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTTTACTCTCTATGCCTCTTGACTAGG SNX1134-219-attB2-noSTOP GGGGACCACTTTGTACAAGAAAGCTGGGTCCTCTCTATGCCTCTTGACTAGG CLASP-F  TCCCCCGGGTATGGAGGAAGCTTTAGAAATG CLASP-R  CGCGGATCCGGTGTCTGCGTCGATAGG CLASP1-501-R  CGCGGATCCATTTATTAGCCTTTGAATGACAG CLASP283-1439-F TCCCCCGGGTTTAATACGAGAATTTGAGAA CLASP502-1229-F TCCCCCGGGTGAAGAAGATGGTGGAAT CLASP502-1229-R CGCGGATCCTCCAGATTTCTTGCTA CLASP502-779-R CGCGGATCCAATTTGCAGTTGGCTATC CLASP686-779-F TCCCCCGGGTTCCGAGTCTCATCCTACTTTTTC     45  CHAPTER 3: A MICROTUBULE-ASSOCIATED CLASP-BRI1-BZR1/2 FEEDBACK LOOP MODULATES BRASSINOSTEROID-DEPENDENT ROOT DEVELOPMENT IN ARABIDOPSIS THALIANA 3.1 Introduction  Plant growth and development involve coordinated activities of cell division, expansion and differentiation. Brassinosteroids (BRs) and the cytoskeleton are both essential for plant growth and development. The BR signalling pathway is initiated from the perception of BR by BRI1, a plasma membrane receptor kinase, and transduced through a series of phoshorylation/dephosphorylation events that ultimately activate BZR1 and BZR2/BES1, the two key transcription factors controlling target gene expression (Clouse, 2011). Earlier studies revealed that BR-deficient or signalling mutants exhibit defective hypocotyl elongation or reduced length of mature root cells (Azpiroz et al., 1998; Clouse et al., 1996; Fujioka et al., 1997; Szekeres et al., 1996), while the gain-of-function mutants with persistently active signalling output result in an opposite effect (González-García et al., 2011; Wang et al., 2002; Yin et al., 2002), attributable to the stimulative role of BR in cell expansion.  As a structural framework for sensing and integrating hormonal signals into directional growth, the cytoskeleton has recently been reported to be involved in BR-induced cell elongation (Lanza et al., 2012; Wang et al., 2012). The gene coding for MICROTUBULE DESTABILIZING PROTEIN40 (MDP40) was originally identified as a putative BZR1 direct target gene from chromatin immunoprecipitation microarray (ChIP-chip) assays (Sun et al., 2010) and further investigations demonstrated that it destabilizes microtubules for their reorientation in response to BR-mediated hypocotyl growth (Wang et al., 2012). Moreover, BR reconfigures actin filaments in elongating root cells (Lanza et al., 2012), providing further evidence that BR functions through the cytoskeleton.  Besides their influence over cell elongation, BRs have been implicated in cell proliferation and thus meristem morphology. The meristem is maintained at a delicate 46  equilibrium between self-renewal and differentiation into new tissues/organs. In the shoot apical meristem, outgrowth of lateral primordia occurs at the periphery of central initials with the growth-arrested boundary between them. BR remains at a low level in the boundary cells, allowing the expression of boundary identity genes. Boundaries are more easily recognized due to insufficient BR biosynthesis or perception, while enhanced BZR1 activity yields organ fusion phenotypes as a result of reduced expression of boundary identity genes and the consequential loss of boundaries (Bell et al., 2012; Gendron et al., 2012). With regard to the root apical meristem, which comprises the quiescent centre with rare cell division surrounded by proliferative initial cells, balanced BR signalling is also needed to achieve an optimal meristem size. Disruption of BR homeostasis reduces meristem size either by promoting the premature elongation of meristematic cells or by slowing down cell production (González-García et al., 2011; Hacham et al., 2011).   Despite the robust correlation between BR and cell division, whether BR’s modulation of cell proliferation relies on the cytoskeleton is still unknown. We sought to understand the connection between microtubules and BR-regulated meristem size using mutants lacking the microtubule-associated protein CLASP. clasp-1 mutants display  smaller meristems resulting from precocious exit from mitosis to elongation (Ambrose et al., 2007), resembling those seen in the constitutively active BR mutants.  BRI1 localizes to the plasma membrane and undergoes constant endocytic trafficking (Russinova et al., 2004; Geldner et al., 2007). BRI1 was found to be transported via the SNX1 retromer complex (Jaillais et al., 2008), which has recently been shown to be stabilized along microtubules through direct association with CLASP (Ambrose et al., 2013). These observations suggest that CLASP and microtubules are involved in the BR signalling pathway.  In the current study, I have identified a microtubule-dependent feedback loop for BR-controlled meristem development. I show that CLASP enhances BR signalling by stabilizing BRI1 at the plasma membrane but that in turn, BR signalling down regulates CLASP expression, leading to changes to microtubule organization that support cell elongation and differentiation.   47  3.2 Results  3.2.1 BR modulates microtubule orientation and CLASP subcellular distribution  To assess BR’s effect on CLASP’s subcellular distribution and microtubule organization patterns, we used wild-type Col-0 lines expressing GFP-CLASP under its endogenous promoter (Ambrose et al., 2011) and GFP-MBD (Marc et al., 1998) driven by a ubiquitin promoter.  In mock treatments, GFP-CLASP formed prominent patches along the transverse edges of epidermal cells within the meristem (Figure 3.1A). After treatment with 10 nM 2,4-epibrassinolide (eBL, a synthetic BR) for 24 h, GFP-CLASP was absent from the transverse edges although some patches were seen along the longitudinal edges (Figure 3.1A). In mock treatments, microtubules were organized into bundles spanning the outer periclinal and transverse anticlinal cell faces, which focussed at the transverse edges in close association with CLASP (Figure 3.1B). The “transfacial ” microtubule bundles bundles were lost after treatment with eBL and replaced by transverse microtubule arrays, which normally develop when epidermal cells enter the elongation zone (Figure 3.1B).  In contrast, GFP-CLASP distribution and microtubule organization remained unaltered when we treated the BR-insensitive mutant, bri1-5 (Noguchi et al., 1999) with eBL (Figure 3.1C, 3.1D). This indicates that activation of the brassinosteroid signal transduction pathway via the BRI1 receptor is required for CLASP’s redistribution and the altered microtubule organization. Notably, the hyperparallel transverse order of cortical microtubules in clasp-1 mutants (Figure 3.1E), which are completely devoid of CLASP expression, closely resembled the microtubule patterns in eBL-treated wild-type cells (Figure 3.1B). Treatment with eBL had no effect on clasp-1 microtubule orientation. .  48   Figure 3.1: eBL regulates CLASP localization and microtubule organization (A and C) eBL induces the redistribution of CLASPpro:GFP-CLASP from transverse (arrowhead) to longitudinal  (arrowhead) in wild type (A) but not in bri1-5 (C). 6-d-old seedlings were transferred to mock or eBL plates for 24 h before confocal imaging. Fluorescence intensity plot corresponding to the dashed white lines, respectively. Red, mock; blue, eBL. (B, D and E) Microtubule alignment in wild type (B), bri1-5 (D) and clasp-1 (E) roots grown on medium with or without eBL for 6 days. Scale bars = 10 µm.  49  3.2.2 BR reduces CLASP expression in a BRI1-dependent manner Based on previous observations that CLASP levels are high in the root division zone and fall off as cells enter the elongation zone (Ambrose et al., 2007), we hypothesized that the eBL-induced reorganization of cortical microtubules is associated with a reduction in the amount of CLASP protein.  To investigate this, we measured the effects of eBL treatment on fluorescence output from the CLASPpro:GFP-CLASP translational reporter in the clasp-1 mutant background, which directly relates fluorescence to CLASP protein levels. Treatment with 10 nM eBL for 24h reduced GFP relative fluorescence by approximately 50% (Figure 3.2A and 3.2D). GFP-CLASP in the bri1-5 mutant background, however, showed no appreciable or significant decline in fluorescence after the same eBL treatment (Figure 3.2B and 3.2D), suggesting that the signal transduction pathway triggered by BRI1 is required for eBL’s reduction of CLASP protein levels. To determine if the CLASP promoter is required for the eBL-induced reduction in CLASP protein levels, we used a 35Spro:YFP-CLASP line reporter line in which YFP-CLASP expression was under the control of the constitutive, BR-unresponsive, 35S promoter (Kirik et al., 2007). In this case, we detected no change in fluorescence or YFP-CLASP distribution patterns upon treatment with 10 nM eBL for 24h (Figure 3.2C and 3.2D), demonstrating that BR represses CLASP gene expression but that it is unlikely to affect the stability of the CLASP protein. To further test whether the CLASP promoter is a target of the BR signalling pathway, we applied eBL to a CLASPpro:GFP reporter line in which GFP fluorescence is an indication of CLASP promoter activity. Results were almost identical to the CLASPpro:GFP-CLASP experiments with relative fluorescence decreasing by 50% in the wild-type background and no effect measured in the bri1-5 background (Figure 3.2E, 3.2F and 3.2G). Thus, BR effects appear to be at the level of transcriptional regulation and not protein stability. Finally, we measured CLASP transcription in response to eBL by quantitative Real Time (qRT)-PCR analysis, using total RNA derived from the 2 mm root tip (Hacham et al., 2011).  According to these results, 10 nM eBL reduced CLASP transcript levels by 30% (Figure 3.2H).   50   Figure 3.2: eBL reduces CLASP transcription in a BRI1-dependent manner (A and B) CLASPpro:GFP-CLASP expression in wild type (A) and bri1-5 (B) root meristems treated with eBL for 24 h.  (C) Response of 35Spro:YFP-CLASP to eBL. (D) Quantification of fluorescence in (A-C), normalizing to mock treatments (n=3, 15 seedlings analyzed for each treatment and genotype).  (E and F) Confocal images of CLASPpro:GFP in wild type (Wassilewskija-2) (E) and bri1-5 (F) root meristems 6DAG, after exposure to eBL (10 nM, 24 h) or mock.   (G) Relative GFP fluorescence in wild type and bri1-5 after eBL application normalized to mock (n=3, 15 seedlings analyzed for each treatment and genotype).  51  (H) CLASP expression is reduced in wild type seedlings grown in the presence of 10 nM eBL for 6 h. ACTIN8 was the reference gene. n=3.  Fluorescence in (A), (B), (C), (E) and (F) is colored to indicate different intensity (see color scales). Error bars denote SD. Asterisks indicate t test results with *p<0.05 and ***P<0.001. Scale bars = 50 µm in (E) and (F) and 20 µm in (A), (B) and (C).   3.2.3 BZR1/2 directly binds to the CLASP promoter to repress its expression BZR1 and BES1/BZR2 are the two major transcription factors in the BR signalling pathway and function both synergistically and independently to regulate BR-responsive gene expression (Wang et al., 2002; Yin et al., 2002). ChIP-chip assays have uncovered many potential BR-regulated and BZR1 and BES1 target genes (Sun et al., 2010; Yu et al., 2011), including the microtubule destabilizing protein 40-encoding gene MDP40, which has been well characterized in hypocotyl growth (Wang et al., 2012). Our sequence analysis of the 2 kb CLASP promoter revealed several putative target motifs for BZR1 and BES1, including one BRRE element  (He et al., 2005) -1500 to -1495 bp upstream of the translation start codon and several E-boxes (Yin et al., 2005) (Figure 3.3A and Figure 3.4). We carried out electrophoretic mobility shift assays (EMSAs) to verify which binding motifs are recognized by these transcription factors. Three promoter fragments were biotin labeled and used as probes (Figure 3.3A and Figure 3.4). Maltose binding protein (MBP) -fused BZR1 and BES1 were expressed in Escherichia coli cells and affinity purified. The result showed that MBP-BZR1 and BES1, but not MBP alone, were able to bind to the P3 (-1531 to -1482) but not the P1 (-346 to -309) or P2 region (-1330 to -1278) as evidenced by the shifted band. Increasing amounts of unlabelled P3 gradually decreased the binding (Figure 3.3B and 3.3D). Furthermore, when we mutated the P3 probe (mP3) by replacing the BRRE motif (CGTGTG) with AAAAAA, which simultaneously damaged the adjacent E-box due to the CGTG overlap, no retarded band was detected and mP3 failed to compete out the binding activity (Figure 3.3C and 3.3E). In summary, our results demonstrated that both BZR1 and BES1 target the CGTG sequence of the CLASP promoter. To evaluate the impact of BZR1 or BES1 binding on CLASP promoter activity, we conducted transient expression assays in Nicotiana benthamiana leaves (Yang et al., 2000). Confocal imaging verified that 35Spro:GFP , 35Spro:GFP-BZR1 and 35Spro:BES1-GFP were expressed ubiquitously in this transient system (Figure 3.5). The reporter, 52  which combines the CLASP promoter DNA sequence with the GUS gene (CLASPpro:GUS), exhibited a substantial expression when co-infiltrated with the 35Spro:GFP negative control (Figure 3.6A). In contrast, cotransfection of CLASPpro:GUS with either 35Spro:GFP-BZR1 or 35Spro:BES1-GFP produced remarkably lower GUS activity, showing that BZR1 and BES1 both inhibit CLASPpro:GUS expression. To further test whether physical binding is required for the repression, we expressed CLASPmpro:GUS harboring the same mutation that abolished BZR1/BES1 binding in the EMSA (Figures 3.3B and 3.3D) and reassessed GUS activity. Overexpression of BZR1 or BES1 displayed no inhibitory action on the mutated reporter (Figure 3.6A). Taken together, these data provide evidence that BZR1/BES1 binding is required for the downregulation of CLASP expression by BR in planta.  Given that BZR1 and BES1 serve as negative regulators for CLASP expression, we reasoned that we would see a decreased CLASP expression and a reduction in meristem size in the gain-of-function alleles of BZR1 and BES1, bzr1-1D and bes1-D, which, through their increased dephosphorylation, cause constitutive activity of BR signalling (Tang et al., 2011). Indeed, as shown in Figure 3.6B, we observed a decline in CLASP mRNA levels as assessed by qRT-PCR in bzr1-1D. In the CLASPpro:GFP-CLASP line, the intensity of GFP signals was also much weaker in bzr1-1D mutant than in wild type (Figure 3.6C and 3.6D). Furthermore, a shorter root apical meristem comprising fewer cells was visualized in bzr1-1D roots (Figure 3.6E, 3.6F and Figure 3.7), which is consistent with previous studies with gain-of-function bes1-D mutants (Yin et al., 2002; González-García et al., 2011). This demonstrated that when endogenously activated, the BR signalling pathway imposes a negative effect on CLASP expression in a similar manner to exogenously applied eBL. 53   Figure 3.3: BZR1 and BES1 directly bind to the CLASP promoter (A) Schematic diagram of the CLASP promoter showing E-box (red open dot), BRRE motif (black dot) and the probes (P1-P3) used in EMSA.  (B and C) EMSA shows BZR1 binds to the P3 region (B) and mutated P3 abolishes the binding (C).  (D and E) EMSA shows BES1 binds to the P3 region (D) and mutated P3 abolishes the binding (E).  54   Figure 3.4: Motifs and probes in CLASP promoter Sequence analysis of the 2 kb CLASP promoter. The translational start codon (ATG) is highlighted with a red box. The BRRE motif (CGTGTG) and E-box (CANNTG) are underlined in red and black, respectively. P1, P2 and P3 are coloured in orange, red and blue.  Figure 3.5 35Spro:GFP, 35Spro:GFP-BZR1 and 35Spro:BES1-GFP are ubiquitously expressed in tobacco leaves in the transient assay (A) 35Spro:GFP. (B) 35Spro:GFP-BZR1. (C) 35Spro:BES1-GFP. Scale bars = 10 μm. 55    Figure 3.6: BZR1 and BES1 repress CLASP expression (A) Transient transcriptional assay in N. benthamiana leaves. The panel indicates the combinations of constructs infiltrated. For quantitative analysis of GUS activity corresponding to each combination, 2,3 are normalized to 1, and 5,6 are normalized to 4 in the left bar graph. 4 is normalized to 1 in the right bar graph. Error bars indicate SD (n=3, 15 leaf areas assessed for each combination).   (B) CLASP expression is reduced in bzr1-1D root meristem. Error bars denote SD in qRT-PCR, n=3.  (C) CLASPpro:GFP-CLASP expression in wild-type and bzr1-1D root meristem at 6 DAG.  (D) Quantification of GFP intensity in (C). Error bars indicate SD (n=3, 20 seedlings assessed for each genotype).  56  (E and F) Quantification of root meristem length (E) and meristem cell number (F) for 7-d-old wild type and bzr1-1D. Error bars indicate SD (n=3, 15 seedlings per genotype).  Asterisks demonstrate t test results with *p<0.05 and ***p<0.001. Scale bars = 20 μm.    Figure 3.7: Meristem of bzr1-1D Root meristem morphology of wild type and bzr1-1D stained with propidium iodide. Meristem regions are marked with white lines. Scale bars = 50 μm.  3.2.4 CLASP sustains BRI1 activity at the plasma membrane Based on the BR receptor BRI1 being identified as a cargo in SNX1 endosomes (Jaillais et al., 2008), along with the recent finding that CLASP tethers SNX1 vesicles to microtubules to assist in recycling of the auxin efflux carrier PIN2 to the plasma membrane (Ambrose et al., 2013), we hypothesized that CLASP also facilitates BRI1 recycling. To test this prediction, we crossed the clasp-1 mutant with a reporter line that expresses BRI1-GFP under the control of the native BRI1 promoter (Geldner et al., 2007). We consistently noted accumulation of BRI1-GFP in intracellular compartments in clasp-1, which was rarely observed in wild-type plants. We determined that these intracellular compartments were lytic vacuoles by prolonged incubation of seedlings in the endocytic tracer FM4-64 (Udea et al., 2001; Kleine-Vehn et al., 2008). FM4-64 labelled the tonoplast surrounding the intracellular BRI1-GFP signal (Figure 3.8A). To determine whether the vacuolar accumulation of BRI1-GFP coincides with its loss from the plasma membrane, we calculated the ratio of plasma membrane-to-intracellular GFP fluorescence intensity. Fluorescence intensity was measured after incubating seedlings in the dark for 4h, which reduces acidity within the vacuolar lumen and hence 57  the loss of GFP fluorescence that occurs under light conditions (Tamura et al., 2003). As shown in Figure 3.8B, the dark treatment enabled BRI1-GFP to be detected in intracellular compartments in the wild type, in accordance with prior descriptions of the BRI1-GFP behaviour under dark conditions (Kleine-Vehn et al., 2008).  In clasp-1 mutants, the BRI1-GFP plasma membrane signal was greatly reduced, with the ratio of plasma membrane to intracellular fluorescence intensity approximately 0.4 in comparison to a wild-type ratio of approximately 1.2, (Figure 3.8C).  To further understand CLASP’s function in BRI1 recycling, we treated the clasp-1 and wild-type BRI1-GFP reporter lines with Brefeldin A (BFA), which has previously been shown to disturb BRI1 recycling and provoke its relocation into aggregated endosomes (Geldner et al., 2007). Internalization of BRI1-GFP occurred normally in clasp-1 mutants, demonstrating that the endocytosis of BRI1-GFP was not affected per se (Figure 3.8D). A profound difference, however, was observed when BRI1-GFP re-established its plasma membrane localization following removal of BFA. Contrary to the wild type in which only 10% of the cells still contained BFA bodies after 2 h of BFA washout, in clasp-1 mutants, prominent BRI1-positive BFA intracellular compartments were still present in about 50% of the cells (Figure 3.8E and 3.8F).  These results are consistent with the impaired BR response seen in clasp-1 mutants, and thus strongly support our hypothesis that CLASP promotes the recruitment of BRI1 to the plasma membrane, protecting it from vacuolar degradation.  In addition to analysis of CLASP’s effects on BRI1 protein levels and distribution, we employed qRT-PCR to ascertain whether CLASP has any influence on BRI1 transcription. We found BRI1 expression in clasp-1 mutants to be approximately 70% that of wild-type levels (Figure 3.8G). This is in contrast to PIN2, which showed no changes in expression in the absence of CLASP (Ambrose et al., 2013), suggesting that CLASP has a positive effect on BRI1 transcript.    3.2.5 CLASP’s association with microtubules is critical for preventing BRI1 degradation To determine if CLASP’s role in maintaining BRI1 levels is dependent on its association with microtubules, we removed microtubules including CLASP-dependent transfacial bundles using short exposure to high concentrations of the microtubule 58  depolymerization drug oryzalin (Ambrose et al., 2007; Ambrose et al., 2013). The oryzalin treatment, which eliminated all microtubules (Figure 3.9), greatly depleted BRI1-GFP abundance (Figure 3.8H and 3.8I), indicating that microtubules play a prominent role in stabilizing the membrane localization of BRI1. Partial depolymerization of microtubules with low concentrations of oryzalin did not reduce BRI1-GFP signal intensity.  Figure 3.8: CLASP modulates BRI1 transcription and stability (A) BRI1-GFP (green) in a clasp-1 root meristem stained with FM4-64 (red) for 2 h. Arrowheads show BRI1-GFP in lytic vacuoles.  59  (B) BRI1-GFP in wild-type and clasp-1 root meristems after 4 h dark incubation. Arrowheads show accumulation of BRI1-GFP in lytic vacuoles.  (C) Quantification of relative BRI1-GFP intensities at the plasma membrane of wild type and clasp-1 root meristematic cells versus intracellular fluorescence intensities in (B) (error bars represent SD, n=3, 10 seedlings analyzed for each genotype, 10 cells per seedling).  (D and E) BRI1-GFP in wild type and clasp-1 root meristematic cells treated with BFA for 1 h (D) and washout for 2 h (E). Arrowheads show aggregation of BRI1-GFP in BFA bodies.  (F) Percentage of epidermal cells with BRI1-GFP positive BFA bodies in wild-type and clasp-1 meristems before and after BFA washout (error bars represent SD, n=3, 20 roots and 10 cells per root analyzed).  (G) Reduced BRI1 transcription in clasp-1 root meristems. Error bars represent SD (n=3). (H) Confocal images of BRI1-GFP abundance in wild type root meristems treated with 100 μM oryzalin or mock for 15 min. Arrowheads show BRI1-GFP in lytic vacuoles. (I) Relative fluorescence of plasma membrane versus intracellular BRI1-GFP in (H) (error bars indicate SD, n=3, 15 seedlings analyzed for each treatment).  Asterisks demonstrate t test results with *p<0.05 and ***p<0.001. Scale bars = 5 μm in (A) and 10 μm in (B), (D), (E), (H).    Figure 3.9: Oryzalin treatment removes microtubules including CLASP-mediated transfacial bundles, shown by GFP-MBD reporter. Scale bars = 10 μm.  3.2.6 BRI1-dependent brassinosteroid signalling is reduced in clasp-1 mutants Since BRI1 maintenance at the plasma membrane is thought to correlate with its receptor activity and activation of the BR signal transduction pathway (Irani et al., 2012), we hypothesized that BR-dependent transcriptional responses would be reduced in clasp-1 mutants. To test this, we first performed quantitative Real-Time PCR (qRT-PCR) on wild-type and clasp-1 plants to compare in the transcriptional activity of DWF4, a BR 60  biosynthetic gene that is known to be down regulated significantly by eBL (Wang et al., 2002; He et al., 2005). The expression of DWF4 in mock-treated clasp-1 mutants was almost twice that of wild-type plants. In addition, DWF4 expression was reduced by only 30% in eBL-treated clasp-1 in comparison to a 60% decrease in wild type (Figure 3.10A).  Thus, BRI1 activity is diminished as a consequence of its reduced retention at the plasma membrane in clasp-1. Consistent with the diminished BR signalling in clasp-1 mutants, we found that the extent to which eBL inhibits root meristem size and cell number was also reduced. Nine day-old wild-type seedlings exposed to 10 nM eBL had a 75% reduction in meristem cell number and an over 50% reduction in meristem length (Figure 3.10B-D). This effect was greatly attenuated in clasp-1 mutants, with 10 nM eBL reducing meristem cell number and length by approximately 30% and 20%, respectively. The above findings indicate that CLASP enhances the ability of high levels of BR to reduce meristem cell number but that it is not required to modulate BR’s effects on cell elongation.  To further explore this relationship, we compared growth responses of wild-type and clasp-1 roots and hypocotyls to a range of eBL concentrations. The rationale for this experiment is that cell division, which is of profound importance to root growth (Beemster and Baskin, 1998), does not contribute to hypocotyl expansion. We found that eBL’s inhibition of relative root length at concentrations at or above 10 nM was diminished in clasp-1 (Figure 3.10E and 3.10F), whereas eBL stimulated hypocotyl elongation to the same extent in clasp-1 and wild type at all concentrations tested (Figure 3.10G). Taken together these results indicate that CLASP modulates BR signalling responses related to cell division. 61    Figure 3.10: clasp-1 is hyposensitive to exogenous eBL treatments. (A) DWF4 expression is increased in clasp-1, and the reduction effect of eBL is alleviated in clasp-1. Plants were grown for 6 days before transfer to 10 nM eBL or mock medium for 6 h. Data were normalized to ACTIN8. Error bars show SD, n=3.  (B) Root meristems of wild type and clasp-1 grown on medium with or without 10 nM eBL for 9 days. Meristem regions are marked with white lines. Scale bars = 50 µm.  (C and D) Quantification of meristem cell number (C) and meristem length (D) in (B) (n=3, 15 seedlings per genotype for each treatment).  (E) Hypocotyl and root phenotypes of wild type and clasp-1 grown for 6 days on medium with or without 50 nM eBL.  62  (F and G) Relative root (F) and hypocotyl (G) length of wild type and clasp-1 mutants on various concentrations of eBL. (n=3, 30 seedlings for each genotype at each concentration). Error bars represent SD.         3.3 Discussion The results of our current study uncover a regulatory feedback system that drives the transition from proliferative cell division in the root apical meristem to terminal differentiation. Previous work established that CLASP’s high expression and abundance in the apical meristem is linked to its function in the formation of prominent transfacial bundles of microtubules that perturb the default transverse parallel order of interphase microtubules (Ambrose et al., 2011). In addition, CLASP’s tethering of SNX1 vesicles to cortical microtubules maintains high levels of PIN2 (Ambrose et al., 2013), the auxin efflux carrier responsible for moving auxin shootward from the root apical meristem (Baluška et al., 2010). Together, these processes foster continual cell division. We have shown here that CLASP enhances brassinosteroid signalling by enabling the recycling of the brassinosteroid receptor BRI1 (and possibly by boosting BRI1 transcript levels). Intriguingly, as a direct target of both BR-activated transcription factors, BZR1 and BES1, CLASP’s expression is down-regulated by the BR pathway. This, in turn, favours the establishment of transversely oriented cortical microtubule arrays that predominate as cells enter the elongation zone (Sugimoto et al., 2000).  BR homeostasis can be achieved through BR biosynthesis (Wang et al., 2002; Tanaka et al., 2005) and metabolism (Neff et al., 1999; Turk et al., 2005; Yuan et al., 2007; Thornton et al., 2010; Zhu et al., 2013), both of which are under tight feedback control by the BRI1 signalling pathway. The significance of the continual cycling of BRI1 between endosomes and the plasma membrane (Russinova et al., 2004; Geldner et al., 2007) for BR signalling has been hotly debated. Elevated BR signalling has been observed when the equilibrium was shifted toward the endosomal pool where BRI1 and its co-receptor BAK1 form functional heterodimers instead of their homodimerization at the plasma membrane (Russinova et al., 2004; Geldner et al., 2007). In contrast, interfering with endocytosis to retain BRI1 at the plasma membrane has been correlated with increased BRI1 signalling (Irani et al., 2012). RAVL1-mediated activation of BRI1 transcription also helps to maintain BR homeostasis (Je et al., 2010). Our present model (Figure 3.11) 63  illustrates a combinatorial role for CLASP in BRI1 transcription and recycling. Similar to its effect on PIN2, CLASP, in association with microtubules, facilitates BRI1 recycling to the plasma membrane. Impairment of BR signalling in clasp-1 mutants, as measured by BR target gene transcript levels, is consistent with the idea that the increased pool of BRI1 at the plasma membrane favours BR signalling. Thus, CLASP’s promotion of BR signalling together with BR’s down-regulation of CLASP expression, constitutes a negative feedback loop that facilitates root differentiation and modulates root apical meristem maintenance when considered in a spatiotemporal context.  Our current findings will need to be considered in the context of tissue- and zone-specific regulation of BR responses in the root apical meristem. In contrast to BRI1’s ubiquitous expression, its homologs BRI1-Like 1 and 3 (BRL1 and 3) are expressed in the central tissues that give rise to the root vasculature (Caño-Delgado et al., 2004; Fàbregas et al., 2013). Although BR signalling in the root epidermis was found to be sufficient to stimulate meristem expansion (Hacham et al., 2011), a recent spatiotemporal translatome analysis revealed that epidermal BR signalling promotes growth while stele signalling antagonizes this effect (Vragović et al., 2015). It was also found that BR-induced and repressed genes prevail in the epidermal cells of the basal meristem and the stele of the apical meristem, respectively (Vragović et al., 2015). Interestingly, CLASP, as a BR-repressed gene, is highly concentrated in the epidermis of the root apical meristem. Our research findings thus highlight the need for a more detailed mapping of transcriptional activity to unravel this complicated regulatory network and to better understand the coherent organ growth.  Auxin-BR cross-talk is implicated in various processes including hypocotyl elongation (Nakamura et al., 2006), lateral root development (Bao et al., 2004) and gravitropic responses (Li et al., 2005). The molecular link between the two hormones has, however, remained a mystery. Lanza et al. (2012) suggested that the actin cytoskeleton may integrate auxin and BR signalling. Adding to the knowledge that CLASP stabilizes PIN2 on the plasma membrane to modulate polar auxin polar transport (Ambrose et al., 2013), our current results demonstrate that CLASP also modulates BR responses. Together, these findings indicate that the microtubule-associated protein CLASP is a critical node between the auxin and BR pathways. 64                      Figure 3.11: Model illustrating the function of CLASP in BR-controlled meristem development   CLASP’s abundance in dividing cells maintains high levels of BRI1 at the plasma membrane by preventing its trafficking to the lytic vacuole. This enhances the BR signalling pathway, which activates the transcription factors BZR1 and BES1. Both of these transcription factors directly bind to the CLASP promoter and down regulate CLASP expression. Initially, this feedback loop buffers the BR signalling pathway because reduced CLASP levels will enhance BRI1 degradation. Eventually, after several rounds of cell division, the reduction in CLASP protein levels leads, through the loss of transfacial microtubule bundle formation, to the establishment of transverse microtubule arrays in early interphase cells, which favours the entry of cells into the elongation zone. Consistent with this model, the complete loss of CLASP expression in the clasp-1 mutant is associated with precocious exit of cells from the meristem.  3.4 Methods 3.4.1 Plant materials and growth conditions Arabidopsis thaliana seeds were geminated on ½ MS (Sigma-Aldrich) plates (1% sucrose, 0.8% agar, pH 5.8) with EtOH or indicated concentrations of eBL (Sigma-Aldrich) and grown vertically for 6 days under continuous light at 22 °C. Plates were scanned and hypocotyl and root length measured using Image J software ( For short-term treatments, 6-d-old seedlings were transferred to ½ MS plate supplemented with EtOH or 10 nM eBL for 6 h (RNA extraction for qRT-PCR) or 24 h (confocal imaging).  65  Cortical microtubules were monitored with GFP-fused microtubule binding domain of MAP4 (Marc et al., 1998), and their angular distribution was quantified as reported (Yao et al., 2008). All plants used for this study are in Columbia-0 ecotype except for bri1-5 (Noguchi et al., 1999) and bes1-D (Yin et al., 2002), which are in Wassilewskija-2 and Enkheim-2 backgrounds, respectively. Some of the other lines were previously described: bzr1-1D (Wang et al., 2002), clasp-1 (Ambrose et al., 2007), CLASPpro:GFP-CLASP (Ambrose et al., 2011), 35Spro:YFP-CLASP (Kirik et al., 2007) and BRI1pro:BRI1-GFP (Geldner et al., 2007). Reporter lines in mutant backgrounds were generated by cross and selection.  3.4.2 Molecular cloning and plant transformation For the microtubule marker pUBQ1:GFP-MBD, smrsGFP followed by a 10× alanine linker (Ambrose et al., 2011) and MAP4-MBD (Mathur and Chua, 2000) were subcloned into pBluescript as a SalI-SpeI and a SpeI-SacI fragment, respectively. Then the entire GFP-MBD sequence was cut out from pBluescript and ligated into a modified pCAMBIA1300 vector containing the Ubiqutin 1 promoter in the HindIII-SalI site (Ambrose et al., 2011). This construct was transferred into Agrobacterium tumefaciens GV3101 using freeze-thaw method. To generate stable transgenic lines, Arabidopsis was transformed by floral dip (Davis et al., 2009). Seeds were screen on ½ MS plates with 50 μg/ml hygromycin. Homozygous plants identified from the T3 generation were used for further analysis. Primers used to make this construct are indicated in Table 3.1.  3.4.3 Drug treatments Oryzalin (Sigma-Aldrich), FM4-64 (Molecular Probes), BFA (Molecular probes) were dissolved in DMSO, H2O and DMSO to make 10 mM, 5 mM and 10 mM stock solutions. Drug treatments were carried out by submerging 6-d-old seedlings into ½ MS liquid containing 50 μM oryzalin, 4 μM FM4-64 and 50 μM BFA. Solvent was added to mock treatments to ensure equivalent content. For BFA washout experiments, seedlings treated with 50 μM BFA for 1 h was briefly rinsed and washed in ½ MS liquid medium for an additional 2 h.  66  3.4.4 Confocal microscopy and image analysis 6-d-old roots were cut and put in the view chamber for microscopy (Ambrose et al., 2013). Root meristem was stained in 10 μg/ml propidium iodide (PI) for 2 min, rinsed with dH2O before visualization. Images were captured using a PerkinElmer spinning-disk microscope (PE) or a Zeiss Pascal scanning microscope. GFP, YFP and PI was excited by the 488, 514 and 561 nm laser of PE and detected through a 525/36, 540/30, and 595/50 nm emission filter, respectively. For Zeiss Pascal, GFP was detected with a 505-600 nm filter. Slice thickness was 0.5 μm. Images were processed with Image J.  Fluorescence intensity was measured using Image J following the instructions ( Microtubule alignment was measured using Angle Tool in Image J.    3.4.5 Total RNA extraction and quantitative Real-Time PCR (qRT-PCR) Total RNA was isolated from the 2 mm root tips (González-García et al., 2011) of 6-d-old seedlings with TRIZOL reagent (Invitrogen, Life Technologies) and then reverse-transcribed using SuperScript® II Reverse Transcriptase (Invitrogen, Life Technologies) to obtain cDNA. qRT-PCR was performed with iQ™ SYBR® Green Supermix (Bio-Rad) in a Bio-Rad iQ5 thermal cycler. ACTIN8 was used as the reference gene. The gene-specific primers used are in Table 3.1.  3.4.6 Electrophoretic mobility shift assay (EMSA) Maltose binding protein (MBP) fused BZR1 and BES1 were expressed in E.coli BL21 cells and purified with amylose resin (NEB). Oligonucleotide probes and mutated P3 probes were synthesized and annealed and labeled with Biotin 3’ End DNA Labeling kit (Pierce). EMSA was carried out using the LightShift Chemiluminescent EMSA kit (Pierce), following the manufacturer’s instructions. Proteins and probes were incubated in 1× binding buffer supplemented with 2.5% glycerol, 5 mM MgCl2, 50 ng/μl Poly (dI·dC) and 0.05% NP-40 in a total volume of 20 μl. The reaction mixtures were separated in a 6.5% native polyacrylamide gel and transferred onto a HybondTM-N+ nylon membrane (Amersham) after incubation at room temperature for 20 min. Biotin-labelled DNA was detected chemiluminescently based on the instructions. 20 fmol of labelled DNA, 4 pmol 67  and 20 pmol of unlabelled competitor DNA were used. Sequences for probes and mutated probes are listed in Table 3.1.   3.4.7 Tobacco transient assay A 2-kb genomic DNA fragment upstream of the ATG start codon of CLASP gene was PCR amplified and cloned into HindIII-SalI sites of pBluescript. As for the CLASP promoter with mutations, site-directed mutagenesis was performed to replace the CGTGTG sequence in P3 region with AAAAAA using the QuikChange II XL Site-Directed Mutagenesis Kit (Agilent Technologies). The two versions of CLASP promoter were then digested from pBluescript and cloned into pBI101.1 binary vector to generate the CLASPpro:GUS and CLASPmpro:GUS reporters. Primers used for mutagenesis are in Table 3.1. BZR1 and BES1 full-length cDNA were PCRed using attB primers and cloned into the gateway compatible entry vector pDONR221 (Invitrogen, Life Sciences) and then shuttled into the binary destination vectors pGWB5 and pGWB6 (Nakagawa et al., 2007) to create the effectors 35Spro:GFP-BZR1/BES1 and 35Spro:BZR1/BES1-GFP. Primers are listed in Table 3.1. Agrobacterium tumefaciens GV3101 containing effector (35Spro:GFP or 35Spro:GFP-BZR1 or 35Spro:BES1-GFP), reporter (CLASPpro:GUS or CLASPmpro:GUS) and p19 silencing suppressor (Lakatos et al., 2004) were co-infiltrated to Nicotiana benthamiana leaves. Co-infiltration and the subsequent GUS activity quantification were described by Liu et al., 2014. This experiment was repeated three times with similar results.                68  Table 3.1: Primers used in this study Primer Sequence (5’ to 3’) DWF4RT-F CACGAGCAACGATATTGAAGTTC (Yan et al., 2009) DWF4RT-R CCTAAGCTCTTCAACGGCTTTAG (Yan et al., 2009) CLASPRT-F ATTTCTGAAATGCTAAAGAG CLASPRT-R CAATAATGGGACAATAACGC (Jancowski et al., 2014) ACTIN8RT-F GAGACATCGTTTCCATGACG (Jancowski et al., 2014) ACTIN8RT-R TTTCAAACCTGCTCCTCCTT BRI1RT-F TGCGATGGATACGCATTTAA BRI1RT-R TCGGACTGACCCTTAGATG smrsGFP-F ACGCGTCGACATGAGTAAAGGAGAAGAAC smrsGFP-R CGGACTAGTTGCTGCTGCTGCTGCTGC MAP4-MBD-F CGGACTAGTATGTCCCGGCAAGAAGAAG MAP4-MBD-R CAAGAGCTCAGATCCCGGGCCCACCTCC P1  TGAAGAAGATAAACGAGAGCATGTGGTTGGCTGGCGTC  GACGCCAGCCAACCACATGCTCTCGTTTATCTTCTTCA P2 CAAAATATATTAAGCATTTGATTAAACTCCGATCAGCTGACATATTAACATGA  TCATGTTAATATGTCAGCTGATCGGAGTTTAATCAAATGCTTAATATATTTTG P3 CGCCGGCGTGACAAGTGACAACAATTGGCCACGTGTGTGTGATTTATTAT  ATAATAAATCACACACACGTGGCCAATTGTTGTCACTTGTCACGCCGGCG mutated P3 CGCCGGCGTGACAAGTGACAACAATTGGCCAaaaaaaTGTGATTTATTAT  CTTAAGATAATAAATCACAttttttTGGCCAATTGTTGTCACTTG CLASPpro-F CCCAAGCTTCACATAAACAAAAATCACTAATAG CLASPpro-R  ACGCGTCGACTTTTTACCAAACCACCGAC mutated CLASPpro-F  CAAGTGACAACAATTGGCCAaaaaaaTGTGATTTATTATCTTAAG mutated CLASPpro-R CTTAAGATAATAAATCACAttttttTGGCCAATTGTTGTCACTTG BZR1-attB1  GGGGACAAGTTTGTACAAAAAAGCAGGCTTCATGACTTCGGATGGAGCTA BZR1-attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTTCAACCACGAGCCTTCCC BZR1-attB2-noSTOP GGGGACCACTTTGTACAAGAAAGCTGGGTCACCACGAGCCTTCCCAT BES1-attB1 GGGGACAAGTTTGTACAAAAAAGCAGGCTTCATGACGTCTGACGGAGCA  69  Table 3.1: Primers used in this study (continued) Primer Sequence (5’ to 3’) BES1-attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTTCAACTATGAGCTTTACCATTTC BES1-attB2-noSTOP GGGGACCACTTTGTACAAGAAAGCTGGGTCACTATGAGCTTTACCATTTCCA                                        70  CHAPTER 4: ARABIDOPSIS THALIANA TRM19: A MICROTUBULE-ASSOCIATED PROTEIN IDENTIFIED THROUGH IMMUNOPRECIPITATION WITH CLASP 4.1 Introduction  As sessile organisms, plants cannot hunt for food and run away from danger. Instead, they respond to the environment by controlling the direction of cell division and elongation. Microtubules are essential for normal plant morphogenesis and architecture by affecting the axes of cell division and elongation.  The onset of cell division is marked by the appearance of the preprophase band (PPB), a transient microtubule structure delineating the plane of future division (Mineyuki, 1999; Dixit and Cyr, 2002).  The PPB is formed in late G2-phase, persists throughout prophase and then is replaced by the bipolar mitotic spindle, on which chromosomes separate during metaphase and anaphase (Dixit and Cyr, 2002). As the cell cycle progresses into cytokinesis, the mitotic spindle is replaced by the phragmoplast, which expands centrifugally with the newly formed cell plate towards the parental wall. Fusion of the new cell plate is at the site occupied previously by PPB (Smith, 2001; Jürgens, 2005; Van Damme et al., 2007; Van Damme and Geelen, 2008). Although the microtubule structures during the entire cell cycle have been well defined, questions remain with respect to how these structures are regulated spatially and temporally, especially how the PPB is formed and predicts the future division site. Mutants of numerous genes exhibit PPB defects, designating their roles in PPB formation. Cells of the mutant mor1-1 (microtubule organization 1-1) fail to form or have aberrant PPBs at the restrictive temperature of 30°C (Kawamura et al., 2006). CLASP is found on PPBs and a majority of the null mutant cells display poorly developed PPBs (Ambrose et al., 2007). The protein SABRE has recently been shown to position CLASP-guided PPB microtubules, mediating PPB and thus cell division orientation (Pietra et al., 2013). Disordered cell files resulting from defective PPBs consisting of loosely and asymmetrically aligned microtubules have also been observed in root cells of bot1, fra2 and lue1, which are all loss of function mutant alleles of the gene encoding the 71  microtubule severing ATPase katanin. These data suggest the importance of katanin in PPB formation (Bichet et al., 2001; Panteris et al., 2011).  Besides the microtubule-associated protein mutants mentioned above, ton1 (tonneau 1) and ton2/fass are the only two so far that have complete abolishment of PPBs, leading to random cell patterning and a severe dwarf phenotype (Traas et al., 1995; Camilleri et al., 2002). The TON1 gene encodes a protein that shares similarity with human centrosomal proteins FOP and OFD1 and associates with CEN1, a homolog of centrin, another eukaryotic centrosomal component (Azimzadeh et al., 2008), while FASS/TON2 encodes the regulatory subunit of PP2A (protein phosphatase 2A) (Camilleri et al., 2002). The TON1 Recruiting Motif protein (TRM) superfamily proteins, comprising 34 members, have been found to interact with TON1 through their C-terminal TON1-interacting motif, and to target TON1 to microtubules via  a central microtubule-binding domain (Drevensek et al., 2012). Interestingly, TRMs have motifs in common with the human centrosomal microtubule-binding protein CAP350, which is responsible for FOP recruitment (Yan et al., 2006). This indicates that a mechanism connected to centrosome function has been conserved during the evolution of plant lineages, in spite of the loss of centrosomes. A recent study by Spinner et al. (2013) revealed the function of a TTP (TON1-TRM-PP2A) complex in the transition from interphase cortical MTs to PPBs. Physical interaction exists between any two components in the complex, which is localized to PPBs via TRM. Similar to ton1 and fass/ton2 mutants, pp2a mutants display impaired PPBs and misoriented cell plates.  This chapter focuses on TRM19, which was identified from an immunoprecipitation experiment as a putative interactor with CLASP. Since both CLASP and TRMs are implicated in PPB formation, we tried to confirm a direct physical interaction and investigate a functional connection between CLASP and TRM19.   4.2 Results 4.2.1 TRM19 was identified through an immunoprecipitation experiment We conducted an immunoprecipitation (IP) experiment to identify CLASP-interacting proteins. Anti-GFP magnetic beads were used to pull down GFP-CLASP from cell 72  lysates of plants expressing GFP-CLASP under the control of the CLASP endogenous promoter. Wild-type plants were used as the negative control to eliminate false positives. The IP experiment followed by mass spectrometry identified two proteins that specifically associated with the CLASP (At2g20190) precipitate, AT1G04820 (TUA4) and AT3G53540 (TRM19) (Table 4.1). TUA4 is an alpha tubulin isoform that is expressed in Arabidopsis roots, leaves and flowers (Abe et al., 2004). As a microtubule-associated protein, CLASP binds to tubulin subunits to regulate microtubule dynamics (Ambrose et al., 2007; Kirik et al., 2007; Ambrose and Wasteneys, 2008). Thus, I focused primarily on TRM19, since its role in microtubule dynamics and organization is unknown.   Table 4.1: Immunoprecipitation results, Atcg and Atmg indicate chloroplast gene and mitochondrial gene, respectively. WT pCLASP:GFP-CLASP/clasp-1 At3g09260 At2g20190 (CLASP)  At1g54010 At3g09260 At3g16460 At1g54010 At3g16420 Atcg00490 At5g26260 At1g04820  At3g53540  Atmg00520  4.2.2 TRM19 is a microtubule-associated protein but does not interact directly with CLASP Among the TRM superfamily, some members localize to microtubules while others show cytoplasmic distribution in transient assays (Drevensek et al., 2012), possibly related to their functional diversity. TRM19 is annotated as a plasma membrane protein and a literature search revealed its presence in cell cortex fractions (Benschop et al., 2007).  In expanding pavement cells of cotyledons I visualized a uniform, punctate fluorescence pattern along microtubules upon transient expression with either N- or C-terminal GFP-fused TRM19 protein, thereby confirming that TRM19 can associate with microtubules.  TRM19 has a distribution pattern resembling that of GFP-CLASP but it could be observed in both wild-type and clasp-1 seedlings (Figure 4.1). This finding 73  indicates that, in contrast to SNX1 (chapter 2 and Ambrose et al., 2013), its microtubule targeting capability is independent of CLASP. Hence, TRM19 was most likely identified from the IP experiment through its binding to microtubules via TUA4, rather than associating directly with CLASP. To distinguish between these two possibilities, I performed BiFC analysis using full length TRM19 and CLASP proteins. The reconstitution of YFP fluorescence was not perceived for any of the eight combinations, while a YFP signal was noted for the positive control pair CLASP-NYFP and SNX1-CYFP, thus ensuring the reliability of the BiFC procedure. Meanwhile, a yeast two hybrid assay also failed to detect TRM19-CLASP interaction. These data do not support the direct interaction between TRM19 and CLASP in vivo.                    Figure 4.1: Transient expression of TRM19 in wild-type (WT) and clasp-1 cotyledons (A) UBQpro:GFP-TRM19 in WT. (B) UBQpro:TRM19-GFP in WT. (C) UBQpro:GFP-TRM19 in clasp-1. (D) UBQpro:TRM19-GFP in clasp-1. Scale bars = 10 μm.   4.2.3 TRM19 expression pattern Although the above analysis shows that TRM19 is not associated with CLASP, little is known about TRM19’s biological function. The TRM19 gene is located on chromosome 3 and contains 5 exons and 4 introns with a total size of 3866 bp. It encodes 74  a protein of 924 amino acids with a calculated molecular mass of 103.27 kDa. RT-PCR revealed a broad pattern of expression in all tissues examined (Figure 4.2). In order to further characterize its spatial and temporal expression profile, a reporter construct was generated by fusing the TRM19 5’UTR promoter region with the coding region of the β-glucuronidase (GUS) gene. Histochemical staining was applied to homozygous transgenic lines carrying this construct. GUS staining observed at various developmental stages completely matched the RT-PCR results but provided a better indication of tissue and organ specific expression. According to the GUS activity assays, TRM19 expression is concentrated in vascular tissues (Figure 4.3B, F) and zones undergoing cell division, including the shoot apical meristem and leaf primordia (Figure 4.3A), the root apical meristem (Figure 4.3E, F) and lateral root primordia (Figure 4.3D, E). This is in agreement with previous finding that TRM1 is involved in cell division control (Spinner et al., 2013). More interestingly, as the seedlings matured, expression shifted to abscission zones such as the inflorescence branching points (Figure 4.3H), and flower and silique receptacles (Figure 4.3I, J), while GUS activity in the leaf vasculature was abolished (Figure 4.3G).                       Figure 4.2: TRM19 expression by RT-PCR S, stems; IF, inflorescences; CL, cauline leaves; RL, rosette leaves; FB, floreal buds and OF, open flowers.  75    Figure 4.3: TRM19 promoter activity indicated by the GUS assay (A) Shoot apical meristem; (B) Cotyledon; (C) Shoot-root junction; (D) Lateral root primordium; (E) Lateral root; (F) Root tip; (G) Rosette leaf; (H) Stem; (I) Silique and (J) Flowers. (A-F) are 7-d-old seedlings and (G-J) are 1-month-old plants. Scale bars = 100 μm in (A-F), 4 mm in (G) and (I) and 2 mm in (H) and (J).  4.2.4 trm mutant phenotype To characterize the biological function of TRM19, I obtained two T-DNA alleles (SALK_049735 and SALK_013820, hereinafter named trm19-1 and trm19-2) from the Salk Institute, both with insertions in exon 5 (Figure 4.4A). RT-PCR showed loss of and reduced level of transcripts in trm19-1 and trm19-2, respectively, and partial transcripts upstream of the T-DNA insertion sites were identified in both lines (Figure 4.4B). No 76  discernible phenotypes were recognized in the growth of trm19 mutants compared with wild type under normal conditions. Moreover, trm19 mutants displayed normal sensitivity to microtubule-disrupting drugs. They exhibited root elongation reduction and left-handed twisting to the same extent as wild type when treated with different concentrations of taxol, oryzalin and propyzamide. The lack of detectable phenotype is possibly due to the redundancy among TRM members or could result from the existence of partial transcripts.                                Figure 4.4: TRM19 expression by RT-PCR in mutants (A) Structure of the TRM19 gene and locations of T-DNA insertions. Black boxes represent exons and black lines denote introns. (B) RT-PCR showing the lack of and reduced level of TRM19 transcripts in trm19-1 and trm19-2 mutants, respectively. Primer pair L1-R1 and L2-R2 in (A) were used to identify full-length and partial transcript, respectively.  4.2.5 TRM19 stable line analysis  Stable Arabidopsis lines expressing the genomic fragment or cDNA of TRM19 with fluorescent tags (GFP or mRFP) coupled either N- or C-terminally under the control of the TRM19 native promoter were established and analyzed. In contrast to the transient expression assays shown in Figure 4.1, none of the stable lines obtained showed a microtubule labelling pattern. TRM19 localized to the plasma membrane (PM) in all root 77  layers and showed the highest fluorescence at the very tip (Figure 4.5). The lack of microtubule binding when driving expression with the endogenous promoter suggests that the microtubule association observed in the transient expression assay could be caused by higher levels of gene expression and accumulation of TRM19 protein, which could lead to false localization information.  To determine if TRM19 is plasma membrane associated, root cells were plasmolyzed with 0.8 M mannitol for 20 min. Instead of co-localizing with the retracted plasma membrane as seen for GFP-PIP2, GFP-TRM19 gave a diffuse cytosolic signal, indicating that TRM19 is located in the cell cortex (Figure 4.6). Interestingly, under plasmolysis conditions, dots with high fluorescence intensity were seen in some cells, which resembled the microtubule distribution patterns observed in the transient expression experiments. We therefore hypothesized that TRM19 associates with microtubules under specific conditions but that under normal circumstances, its level of expression is too low to confer microtubule association.  In order to generate stable lines with higher levels of TRM19 expression, we expressed GFP-TRM19 and TRM19-GFP under a ubiquitin promoter (UBQpro:GFP-TRM19 or UBQpro:TRM19-GFP). Despite the presence of the ubiquitin promoter, the TRM19 fluorescent reporter signals were surprisingly low (Figure 4.7), with similar intensity to the levels observe in the TRM19pro:TRM19-GFP lines. In contrast, fluorescence was also detected in the root transition zone. These observations suggest that TRM19 is under some post-transcriptional regulation, which maintains protein at low levels, and that microtubule association may be triggered by intracellular and extracellular stimuli.      78  Figure 4.5: TRM19pro:genomic TRM19-GFP was detected in all root layers  (A) Root epidermal cells (B) Cortical cells (C) Mid focal plane of the root Scale bars= 10 μm                                                                            Figure 4.6: TRM19-GFP showed diffused signal after plasmolysis  (A) GFP-PIP2 (B) TRM19-GFP Scale bars= 10 μm                                                       Figure 4.7: UBQpro:GFP-TRM19 showed PM localization with highest fluorescence in root transition zone Scale bars= 10 μm  4.2.6 Expression of TRM19 under various stress conditions  Microarray data from Genevestigator ( and the Arabidopsis database ( provided hints to understand how TRM19 functions. According to these data, a variety of stress-related environmental cues such as drought, osmotic stress and high salinity as well as hormones including ABA can 79  trigger the elevation of TRM19 expression. To verify whether TRM19 is involved in responses to these stresses and ABA, I treated trm19 mutants with ABA, salt and mannitol but did not observe any difference in either seed germination rates or post-embryonic development compared to wild type. I also subjected the TRM19pro:TRM19-GFP lines to these same treatments but detected no change in protein levels, as indicated by fluorescence intensity. Interestingly, the UBQpro:TRM19-GFP did show increased fluorescence with ABA treatment and subsequent tests determined that this promoter is activated by ABA.   4.3 Discussion TRM19 belongs to the TON1 Recruiting Motif protein superfamily composed of 34 members, which share 6 conserved motifs. All TRMs contain the C-terminal M2 motif, which binds to TON1. The high sequence diversity apart from conserved motifs within the TRM family and the differential transcriptional patterns shown by the Genevestigator tool suggest functional diversification (Drevensek et al., 2012). To date, only a few studies have been done to establish TRMs’ cellular localization and biological functions. Some TRMs such as TRM1, 2, 8 and 25 are microtubule-associated proteins whereas others such as TRM20 and 26 are cytoplasmically dispersed (Drevensek et al., 2012). TRM1 and 2 were found to be involved in cell elongation. Their dominant mutants show longer leaf blades, leaf petioles, floral organs, cotyledons and seeds (Lee et al., 2006). TRM14 and 19 were detected in cortex fractions (Benschop et al., 2007) while TRM7 displayed a plasma membrane association (Nühse et al., 2003). TRM29 was identified as a nuclear protein interacting with the transcription factor ALCATRAZ which regulates pod shattering in Arabidopsis (Wang et al., 2008).  We identified TRM19 as a putative CLASP interactor from an immunoprecipitation experiment. However, BiFC and Y2H assays failed to validate any direct interaction. Since TUA4 was also obtained, it is reasonable to assume that TRM19 binds to TUA4, which got pulled down via a direct association with CLASP. A transient expression assay revealed that TRM19 decorates microtubules in both wild type and clasp-1 mutant cotyledon cells, suggesting that TRM19 is a microtubule- associated protein and that its 80  targeting capacity is independent of CLASP. This still did not rule out the possibility that other techniques could detect the physical interaction between TRM19 and CLASP. Microtubule targeting was not observed in stable Arabidopsis transgenic lines using either the endogenous promoter or a ubiquitin promoter that drives constitutive expression. Instead, TRM19 was confined to the cell cortex with a diffuse distribution. In some cells we saw structures that resemble the dots seen along microtubules observed in transient overexpression assays, making us reason that low-level TRM19 expression prevents detection. More interestingly, a very dim TRM19-GFP signal was seen even with ubiquitin promoter, suggesting a strict post-trascriptional regulation to maintain a low level under normal conditions. trm19 mutants did not show morphological defects under normal growth conditions, treatments with microtubule-stabilizing and -destabilizing drugs and abiotic stresses imposed by salt, mannitol and ABA. This result was not surprising in terms of the mild phenotypes shown by trm1 and trm2 single mutants, suggesting functional redundancy and compensation among TRM members in such a large family. The TRM family is subdivided into 8 groups plus a few isolated ones based on sequence alignments and TRM19 is in subgroup 5 with TRMs 17, 20, 21, 19 and 26 (Drevensek et al., 2012). Generating and analyzing the phenotypes of multiple (e.g., double, triple, quadruple) mutants would be more informative to unravel TRM19 and its subgroup members’ biological functions.     4.4 Methods 4.4.1 Plant materials and growth conditions  Arabidopsis seeds were surfaced sterilized for 2 mins with 70% ethanol followed by rinsing with sterile water and then plated on Hoagland’s medium or half strength Murashige-Skoog (Sigma-Aldrich) medium. After stratification at 4°C for 48h, plates were transferred vertically to a growth chamber at 22°C with a long-day (16-hour light/8-hour night) regime. Two-week-old seedlings were potted into soil and further grown under the same conditions.   81  4.4.2 Immunoprecipitation (IP) assay 1g of 4-day-old CLASPpro:GFP-CLASP as well as wild-type seedlings were harvested and rapidly frozen in liquid nitrogen. After grinding into fine powder, immunoprecipitation was carried out with µMACS GFP-Tagged Protein Isolation Kit (Miltenyl Biotec) according to the manufacturer’s protocol. Subsequent mass spectrometry was performed at the Centre for High-throughput Biology (CHiBi) at the University of British Columbia.  4.4.3 BiFC TRM19 and CLASP cDNA (with and without stop codons) were PCR-amplified and cloned into pDONR221 entry plasmid through GATEWAY BP reactions (Invitrogen). The PCR products were sequenced and recombined through LR reactions into BiFC destination plasmids. The binary plasmids were transformed into Agrobacterium tumefaciens strain GV3101 using the Freeze-Thaw method. The BiFC assay was carried out as in Chapter 2. Primers used are list in Table 4.2.  4.4.4 Yeast 2-Hybrid analysis pGBKT7 and pGADT7 vectors (BD Biosciences) were used for creation of proteins fused in frame with either the GAL4 DNA binding domain or GAL4 DNA activation domain. CLASP cDNA was PCR-amplified and cloned into pGBKT7 bait vector between the Xma I and Bam HI sites. TRM19 was PCR-amplified as an Xma I fragment and inserted into pGADT7 to generate the prey construct. Primers used are listed in Table 4.2. Saccharomyces cerevisiae strain AH109 (Clontech Laboratories, Inc.) was transformed with bait and prey constructs and plated on minimal medium lacking leucine and tryptophan. After incubating at 30°C for 3 days, individual colonies were selected and spotted onto minimal medium lacking tryptophan, leucine and histidine plus 5 mM 3-amino-1,2,4-triazole (Sigma-Aldrich).     82  4.4.5 Total RNA extraction and RT-PCR Total RNA was isolated with Trizol reagent (Invitrogen). First-strand cDNA was synthesized using SuperScript II reverse transcriptase (Invitrogen) from 1μg of RNA. Subsequent PCRs were conducted using the primers listed in Table 4.2.  4.4.6 Identification of T-DNA insertions T-DNA insertion lines SALK_049735 (trm19-1) and SALK_013820 (trm19-2) were acquired from the Arabidopsis Biological Resource Centre ( Mutant genomic DNA was extracted from cotyledons utilizing the sucrose prep method (Berendzen et al., 2005). Gene-specific primers and T-DNA-specific primers were used to confirm the position of T-DNA insertions. Genotyping primers are listed in Table 4.2.  4.4.7 Plasmid construction and generation of stable transgenic lines TRM19 cDNA was amplified and cloned into pUBC/pUBN-GFP destination vectors through the Gateway strategy. Primers were the same as for the BiFC assay. For the promoter-GUS fusion construct, a fragment covering ~3 kb upstream of the start codon of the TRM19 gene was amplified from Col-0 genomic DNA using primers containing Hind III and Sal I sites and inserted into corresponding sites of PBI101.1 vector harbouring a GUS reporter gene.  To avoid the over-represented phenotype possibly caused by ubiquitin promoter, a 3 kb TRM19 promoter (same as the GUS reporter construct), TRM19 coding region with Sal I and Spe I sites, and GFP coding sequence with Spe I and Pst I sites were subcloned into pCambia 1390 binary vector. Primers are shown in Table 4.2. Arabidopsis plants were transformed by the floral dip method (Clough and Bent, 1998) using Agrobacterium tumefaciens strain GV3101 carrying the constructs above. Positive transgenic plants (T1) were screened based on their antibiotic resistance markers (spectinomycin for UBQpro:TRM19-GFP and UBQpro:GFP-TRM19, kanamycin for TRM19pro:GUS and hygromycin for TRM19pro:TRM19-GFP and TRM19pro:GFP-TRM19), and independent T3 homozygous progenies were used for experiments.   83  4.4.8 Drug treatments 10 mM Taxol, oryzalin and propyzamide (all from Sigma-Aldrich) stock solutions were prepared in DMSO. Stock solutions were added to half MS medium to achieve the indicated final concentrations. DMSO was added so that the final solvent content was the same for drug and control treatments. All plates were observed and pictures recorded after 7 days. Root length and twisting angle were measured using Image J software (    4.4.9 Confocal imaging Seven-day-old seedlings were used for GFP fluorescence analysis. Roots were cut at the hypocotyl junction and placed into view chambers (Nunc) with a layer of solidified 1% bacto agar on top to maintain their position. Images were captured with a Perkin-Elmer spinning-disk microscope. GFP signal was generated by a 488 nm laser and detected through a 502/35 emission filter. Optical slice thickness was 1 μm. Images were processed with Image J software.  4.4.10 Histochemical GUS staining GUS activity was checked as stated by Malamy and Benfey, 1997. Tissues were immersed into a staining solution (1 mg/ml X-Gluc (5-bromo-4-chloro-3-indolyl-b-D-glucuronidase), 20% methanol, 10 mM Na2EDTA, 5 mM K4Fe(CN)6/K3Fe(CN)6, 90 mM sodium phosphate buffer, pH 7.0) and incubated overnight at 37°C. Samples were cleared afterwards by ethanol series treatment. Four developmental stages of TRM19pro:GUS plants were inspected: 5-d-old seedlings, 7-d-old seedlings, 10-d-old seedlings and mature flowering plants.         84  Table 4.2 Primers used in this study Primer Sequence (5’ to 3’) TRM19RT-F ATTTGTCAACTGTCACCTCGGT TRM19RT-R CTAGAAGATTGCACCTACCACA TRM19 partial RT-F GCTTCAACACTGCCCACT TRM19 partial RT-R CAAGAGCATCATTAAACTCTTTAG ACTIN8RT-F ATTAAGGTCGTGGCA (Walia et al., 2009) ACTIN8RT-R TCCGAGTTTGAAGAGGCTAC (Walia et al., 2009) SALK_049735 LP GCAAATCCGGAAACCTAAAAG SALK_049735 RP TGAATCATTCTTCGGTTCCTG SALK_013820 LP ACAACATTCAGTGGCCTGAAC SALK_013820 RP GCTTTCTTTCTAACCGGGTTG LB  ATTTTGCCGATTTCGGAAC CLASPpro-F (Hind III) CCCAAGCTTAAATTTGTTCATATACGAGT CLASPpro-R (Sal I) ACGCGTCGACCTTTCACTCACTGTTCATCT TRM19-F (Sal I) ACGCGTCGAC ATGAACAGATTTCGACTCAG TRM19-R (Spe I) TGCACTAGTGAAGATTGCACCTACCACAA LGFP fwd (Spe I) TGCACTAGTGCAGCAGCAGCAGCAGCA LGFP rev (Pst I) AAAACTGCAGTCATTTGTATAGTTCATCCA CLASP-F (Xma I) TCCCCCGGGTATGGAGGAAGCTTTAGAAATG CLASP-R (Bam HI) CGCGGATCCGGTGTCTGCGTCGATAGG TRM19-F (Xma I) TCCCCCGGGTATGAACAGATTTCGACTCAG TRM19-R (Xma I) TCCCCCGGGGAAGATTGCACCTACCACAA TRM19-attB1  GGGGACAAGTTTGTACAAAAAAGCAGGCTTCATGAACAGATTTCGACTCAG TRM19-attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTCTAGAAGATTGCACCTACCACAA TRM19-attB2-noSTOP GGGGACCACTTTGTACAAGAAAGCTGGGTTGAAGATTGCACCTACCACAA                85  CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS 5.1 Conclusions The major goal of my PhD project was to decipher CLASP’s function and regulation. CLASPs, so-named because they are CLIP-Associated Proteins, belong to the ORBIT/MAST/CLASP family of MAPs, and were initially shown in animal and fungal cells to play a pivotal role in microtubule functions during interphase and mitosis (Maiato et al., 2005; Lansbergen et al., 2006). Due to their higher affinity for the microtubule plus end, where they are thought to modulate plus end dynamics, as well as bind to other cellular structures, such as the plasma membrane and kinetochores, they are termed plus-end-tracking proteins or +TIPs (Akhmanova and Hoogenraad, 2005; Galjart, 2005). It has been reported that +TIPs include a great number of structurally diverse MAPs (e.g. CLASP, EB1, CLIP-170) that form complexes to regulate microtubule activities (Schuyler and Pellman, 2001; Coquelle et al., 2002; Carvalho et al., 2003; Goodson et al., 2003; Howard and Hyman, 2003; Lansbergen et al., 2006; Akhmanova and Hoogenraad, 2005; Mimori-Kiyosue et al., 2005). Depletion or reduction of CLASP imposes a dramatic effect on interphase and mitotic microtubules, thus leading to defective cell motility and disturbed mitosis (Lee et al., 2004; Drabek et al., 2006). The Arabidopsis genome contains a single CLASP gene. Research has shown that AtCLASP is involved in both cell morphogenesis and cell expansion. By labelling it with GFP or YFP, it has been observed CLASP localizes to all microtubule arrays and shows plus end enrichment (Ambrose et al., 2007; Kirik et al., 2007). Interestingly, Arabidopsis plants lacking CLASP gene expression (clasp-1 mutants) display various phenotypes that suggest auxin functions are altered. First, the clasp-1 dwarf morphology results in part from fewer cells being maintained in the meristematic zones, a process that is dependent on auxin. Second, clasp-1 mutants have more lateral roots than wild-type plants, which mimics the effects of high auxin concentrations in the differentiation zone. Finally, clasp-1 mutants show reduced sensitivity to auxin concentrations that inhibit root elongation. Furthermore, a GFP reporter gene driven by the DR5 auxin-responsive promoter also displays an altered distribution pattern of auxin in clasp-1 mutants. Together these observations suggest that CLASP probably plays a role in some aspects of polar auxin transport or in an auxin-dependent signalling pathway.  86  5.1.1 CLASP and the auxin signalling pathway  PIN proteins constantly cycle between endosomal compartments and the plasma membrane (Geldner et al., 2003; Jalliais et al., 2006). The trafficking is, however, quite complicated as reflected by the distinct routes taken by different PIN proteins. For example, PIN1 uses GNOM endosomes whereas PIN2 uses SNX1-defined endosomal compartments for their cycling (Geldner et al., 2003; Jalliais et al., 2006). SNX1 is a plant ortholog of human SNX1 and is a component of the retromer complex, which controls cargo transport from endosomes to the Trans-Golgi Network (TGN) and is highly conserved among eukaryotes (Vanoosthuyse et al., 2003; Seaman, 2005). It contains a core subunit (Vps26-Vps29-Vps35), which provides cargo selection, a PHOX domain for interaction with PtdIns3P (phosphatidylinositol 3-phosphate) on the endosomal membrane, and a BAR domain for dimerization to curve the membrane (Bonifacino and Rojas, 2006).  In Arabidopsis, SNX1 localizes to the prevacuolar compartment (PVC) or multivesicular bodies (MVB) mediating PIN2 recycling as evidenced by the reduced amount of PIN2 in snx1-1 null mutant (Jalliais et al., 2006; Kleine-Vehn et al., 2008c). Cui et al. (2010) found that BLOS1, a subunit of BLOC-1 complex, which in humans and mice participates in the transport from endosomes to lysosomes, physically interacts with SNX1, directing PIN2 to the lytic vacuoles for degradation. An increased level of PIN2 was observed in BLOS1 RNAi lines due to compromised degradative trafficking. This again designates the striking complexity and evolutionary conservation of intracellular trafficking. Two independent Yeast 2-hybrid screens indicated that CLASP is a strong interactor with SNX1 (Chapter 2). We validated the direct association via a variety of methods including BiFC, Y2H and colocalization between GFP-CLASP and SNX1-RFP with live cell imaging. After 3-D structural prediction, I confirmed using BiFC that amino acids 134-219 form the interacting site on SNX1. I then used Y2H assays to identify CLASP686-779 as the SNX1-binding site. When the CLASP gene is mutated or microtubules are disturbed by oryzalin, SNX1 endosomes become misshapen, reduced in density and have shorter lifespans. Accordingly, its cargo PIN2 takes the alternative route to target to the vacuole instead of the PM.  In view of these results, we proposed that CLASP and microtubules constitute essential parts in the large complex that modulates PIN2 trafficking (Ambrose et al., 87  2013). The balance between recycling and degradation is of crucial importance to PIN2 levels for auxin transport, signalling and action. Moreover, our findings demonstrated a reciprocal relationship between microtubules and auxin, suggesting that environmental signals could control auxin polar transport and auxin gradients through CLASP and microtubules to optimize growth.  5.1.2 CLASP and brassinosteroid signalling pathway  The investigation described in Chapter 2 illustrates that CLASP is involved in the auxin signalling pathway. In Chapter 3, I uncovered the relationship between CLASP and another class of hormones - brassinosteroids. Despite the knowledge that the cytoskeleton and brassinosteroids are critical regulators of cell division and expansion (Rasmussen et al., 2013; Smith et al., 2005; Bashline et al., 2014; Singh et al., 2015; Zhiponova et al., 2013; Clouse et al., 2011), the connection between the two has remained unclear. Brassinosteroids have been suggested to play a role in root meristematic cell proliferation and differentiation. Deficiency or excess of brassinosteroids affects root meristem negatively due to distinct mechanisms (González-García et al., 2011; Hacham et al., 2011).   The premature differentiation induced by enhanced brassinosteroid signalling reminded us of the phenotype of the clasp-1 root.  It was therefore reasonable to hypothesize that brassinosteroids control root meristem size through CLASP. Real time PCR and analysis of reporter lines led to two conclusions: (1) brassinosteroid down regulates CLASP transcription and switches its localization from transverse to longitudinal and (2) decreased CLASP expression and altered distribution, especially its loss from the newly formed transverse cell edges, is correlated with a transverse cortical microtubule array that promotes cell elongation, generating a smaller meristem with fewer cells. Further exploration revealed that BZR1 and BES1/BZR2 transcription factors directly target the “CGTG” sequence on the CLASP promoter and repress its expression.  Meanwhile, CLASP and microtubules act as positive regulators for BRI1 recycling, just as they do for PIN2.  Increased vacuolar targeting of BRI1 was observed in clasp-1 mutants or wild-type plants treated with oryzalin to eliminate all microtubules. I also found that CLASP participates in BRI1 transcriptional regulation, using an as yet 88  uncharacterized mechanism. These results elucidate a feedback suppression of CLASP for fine-tuning brassinosteroid signalling during meristem development. Also, it further supports the idea that CLASP and the brassinosteroid signalling pathway have a bidirectional link and that the CLASP-SNX1 module is possibly a junction between auxin and brassinosteroid pathways.  5.1.3 TRM19  Chapter 4 describes my identification and analysis of the proteins TRM19. TRM19 was identified in an immunoprecipitation experiment as a putative CLASP interactor. However, various approaches including BiFC, Y2H and the analysis of TRM19-GFP reporter in wild type and clasp-1 mutants failed to validate their physical interaction. Nevertheless, we still cannot rule out the possibility that other approaches might allow for a positive interaction considering the fact that interactions among some TRM members were identified by tandem affinity purification but not BiFC (personal communication, Martine Pastuglia). And since both TRM19 and CLASP function in PPB formation, it remains possible they function in the same complex. I demonstrated that TRM19-GFP decorates microtubules along their entire length in transient assays but not in stable Arabidopsis transgenic lines with TRM19-GFP driven by either the TRM19 native promoter or the constitutive UBQ10 promoter. In stable lines, TRM19 displayed a dim signal in the cell cortex with some dotted structures reminiscent of the transient assay. My data suggest that TRM19 is a microtubule-associated protein, adding to the previously characterized localizations of TRM members. Intriguingly, even the UBQ10 promoter, which should generate moderate levels of expression (Norris et al., 1993), generated quite faint fluoresence, indicating that the expression of TRM19 is under post-transcriptional modulation to maintain its normally low levels. However, the relatively low expression is the main limiting factor for studying its dynamics and influence over microtubules. I attempted to overcome this limitation by performing in vitro experiments to confirm the binding of TRM19 to microtubules and to evaluate how it regulates microtubule dynamics. Unfortunately, I was unable to express TRM19 in bacterial cells. 89  TRM19 is expressed where high mitotic activity takes place, such as shoot apical meristems, young true leaves, root apical meristems, lateral root primordia, inflorescence branching points and flower and silique receptacles. This is consistent with the previous finding that the TTP (TON1-TRM-PP2A) complex is involved in PPB formation during cell division. Microarray data predicted a stress-related function of TRM19. Yet, trm19 single mutants did not produce discernible changes in phenotype under normal or stress conditions including salt, ABA and mannitol or when treated with microtubule-targeted drugs. Multiple loss-of-function mutants therefore will need to be generated and further investigated in order to understand the biological function of TRM19.  5.2 Future directions The data in my thesis unravel the involvement of CLASP in the auxin and BR signalling pathways. I also carried out and have planned additional projects to understand how CLASP is controlled and to identify additional functions of CLASP. These are described below.  5.2.1 Additional functions of CLASP CLASP binding partners have been identified in yeast, Drosophila and mammalian cells including other MAPs such as EB1, XMAP215/TOG and CLIP at microtubule plus ends (Akhmanova et al., 2001; Watanabe et al., 2009; Lowery et al., 2010), CENP-E at kinetochores (Maffini et al., 2009), LL5β at the cell cortex (Lansbergen et al., 2006) and kinases such as GSK3β and Abl (Kumar et al., 2009, Engel et al., 2014). The binding and phosphorylation of CLASP2 by GSK3β (human) or Abl (Drosophila) modulates its association with MTs and its other binding partners (Watanabe et al., 2009; Engel et al., 2014).  I conducted an extensive forward genetics screen to identify putative CLASP interactors in Arabidopsis. I applied ethyl methane sulfonate (EMS) mutagenesis to CLASPpro:GFP-CLASP/clasp-1 seeds. In this reporter line, the transgene fully compensates for the loss of endogenous CLASP, resulting in plants that are phenotypically identical to wild type. I screened for mutants using a two-step procedure. Seedlings from M2 (the second generation) seeds were screened for dwarf and branchy 90  roots, a phenotype characteristic of CLASP loss of function. Putative mutants were transplanted to soil to generate M3 (the third generation) seeds. Those M3 seedlings that continued to show dwarf and branchy phenotypes were screened for altered GFP-CLASP intensity and/or distribution. This secondary screening strategy was designed to ensure the specificity of the mutations for subcellular distribution and expression levels of CLASP. Four categories of possible mutants were expected: (1) mutations in the GFP-CLASP transgene that disrupt its functionality; (2) mutations in genes that regulate CLASP expression (such as transcription factors); (3) mutations in genes that regulate CLASP protein activity (such as kinases and phosphatases); and (4) mutations in genes encoding proteins that function in the subcellular targeting of CLASP.   Unfortunately, after screening 100,000 seedlings, I did not obtain a single mutant with a stable phenotype. Mutant lines keep segregating wild-type progeny during the M4 and M5 generations. Furthermore, confocal microscopy showed no alteration in the GFP-CLASP localization in the dwarf seedlings that segregated out. The failure of this screen to identify single gene mutations is possibly due to the likely lethality of losing function in a gene that controls, among other things, CLASP distribution or the inability of those point mutations in the transgene to disrupt its function. Reflecting on the success of the mor1-1 mutant screen that obtained temperature-sensitive mutants (Whittington et al., 2001), it may be possible to modify the screen to isolate mutants whose roots only undergo defective development in response to shifting the temperature from 21 to 31 degrees.  In addition to forward genetics, I have also considered using a reverse genetic approach to find potential CLASP interactors. A number of co-expression databases (the Expression Angler, ATTED-II, etc) allow us to infer CLASP function in a reverse genetic system. Genes that exhibit co-expression with CLASP fall into 4 categories as shown in Table 5.1. Many genes among them are well-defined and their mutants as well as GFP/RFP reporter lines are available already. Thus to monitor the double mutant phenotype and changes of GFP patterns, seeds of these lines (both mutants and reporter lines) could be requested and crossed into clasp-1 and GFP-CLASP (and vice versa, GFP-CLASP into the mutant lines). This may help uncover novel pathways in which CLASP might participate. 91  Table 5.1: Genes that co-express with CLASP ENDOMEMBRANE CYTOSKELETON CELL WALL HORMONEAdaptin FASS/TON2 Glycosyltransferase IAA9 COP1-interacting protein-related ACT3/7 KOR1 IAA12 Sec10 SPK1  RGA1 PDR2 Spiral like-3   LRR kinase STICHEL   Importin    PIP kinase    Phosphatidate cytidyltransferase     5.2.2 Biological function of TRM19  In Chapter 4, I observed the microtubule labeling pattern of TRM19 using a transient expression assay in Arabidopsis cotyledons. It would be interesting to examine the effect of TRM19 on microtubule dynamics using in vitro experiments. The expression of TRM19 protein in bacterial cells has, however, been technically challenging. For future work, we could try yeast or insect cell expression systems. Using purified TRM19, it would be possible to perform microtubule co-sedimentation experiments to confirm microtubule binding, tubulin turbidity assays to monitor tubulin assembly, and TIRF (total internal reflection fluorescence) microscopy to visualize the behavior of TRM19 as single particles on microtubules.  Since the trm19 single loss-of-function mutant did not have any noticeable defects, quite possible due to redundancy, multiple mutants among the TRM19 subgroup could be created by crossing and screening T-DNA knock-out lines or by using RNA interference. 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