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Characterizing the role of proteins in modulating microtubule dynamics and cell morphogenesis in Arabidopsis… Eng, Ryan Christopher 2015

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		CHARACTERIZING THE ROLE OF PROTEINS IN MODULATING MICROTUBULE DYNAMICS AND CELL MORPHOGENESIS IN ARABIDOPSIS THALIANA by  Ryan Christopher Eng  B.Sc., The University of British Columbia, 2009  A THESIS SUBMITTED IN PARTIAL FULFILLMENT 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)  December 2015  © Ryan Christopher Eng, 2015 	 ii ABSTRACT  Microtubules are dynamic polymers that are important for the growth and development of plant cells.  Because of their vital role, the dynamics and organization of microtubules need to be tightly modulated by other proteins.  The major focus on this dissertation is to elucidate and characterize the role of specific proteins (ARK1, NEK6, MOR1) that are responsible for orchestrating the dynamics and organization of microtubules in the model plant, Arabidopsis thaliana. The motor protein, ARK1, was previously shown to play an important role in root hair morphogenesis but had an unknown role in modulating microtubule dynamics.  Moreover, evidence showed that ARK1 physically interacts with the NEK6 kinase, although for an undetermined reason. Based on my data, I determined that ARK1 functions as a plus-end tracking protein that has a specific role in promoting the depolymerization of microtubules.  I also discovered that ARK1 has a secondary microtubule-binding domain in addition to the motor domain, which is the canonical microtubule-binding domain. I noted, however, that this secondary microtubule-binding domain is not essential for ARK1’s ability to induce microtubule depolymerization.  While NEK6 and ARK1 both modulate microtubule dynamics, I determined that neither protein requires each other for function or localization, suggesting that they operate independently from each other to control microtubule dynamics and cell elongation.  In addition, I provided evidence that shows that ARK1 has a putative yet unknown role in controlling NEK6 gene expression.  Finally, the microtubule-associated protein MOR1 was confirmed to be a plus-end tracking protein through live-cell imaging of MOR1 fused to a fluorescent reporter (MOR1-3xYpet). In revealing that MOR1 binds to both growing and shrinking microtubule plus ends, my data corroborated previous studies showing reduced microtubule growth and shrinkage in mor1 mutants, and confirm MOR1’s role as a microtubule polymerase.  Comparing MOR1-3xYpet live-cell imaging with previous experiments using fixed samples revealed that the chemical fixation process affects the plus-ends of microtubules, stressing the importance of using non-fixed samples for the most accurate results.  My dissertation thus expands our knowledge about how microtubule dynamics and organization is controlled in plant cells by three distinct proteins.   	 iii PREFACE  Chapter 1 uses figures reproduced with permission from the publishers and are indicated within the chapter.  A portion of Chapter 2 has already been published as: Eng and Wasteneys (2014). The Microtubule Plus-End Tracking Protein ARMADILLO-REPEAT KINESIN1 Promotes Microtubule Catastrophe in Arabidopsis. The Plant Cell. 26: 3372-3386. © Copyright American Society of Plant Biologists, 2014 (www.plantcell.org).  Text, tables, and figures are reproduced with permission from the authors.  Reproduced figures are indicated within the thesis.  The remainder of the unpublished work in this chapter will be incorporated in a manuscript with data from Chapter 3.  Ryan Eng and Dr. Geoffrey Wasteneys designed the experiments and prepared the manuscript.  Ryan Eng performed the experiments and analyzed the data.  Experiments in Chapter 3 use transgenic seed lines and information via personal communications from Dr. Hiroyasu Motose (Okayama University, Japan).  Ryan Eng and Dr. Geoffrey Wasteneys designed the experiments.  Ryan Eng performed the experiments, analyzed the data, and created the figures.  Samuel Livingston, an undergraduate laboratory assistant, helped out with the RNA extraction and the RT-PCR experiments.  Data from Chapter 3 will be submitted for a peer-reviewed publication with Dr. Motose and Dr. Wasteneys as co- and corresponding author, respectively.  Chapter 4 uses a fusion-protein MOR1 construct created by Dr. Jose Alonso (North Carolina State University, USA).  Dr. Geoffrey Wasteneys and Ryan Eng designed the experiments.  Ryan Eng performed the experiments, analyzed the data, and created the figures.  Data from Chapter 4 will be submitted for a peer-reviewed publication with Dr. Alonso and Dr. Wasteneys as senior and corresponding author, respectively.      	 iv TABLE OF CONTENTS  Abstract .......................................................................................................................................... ii	Preface ........................................................................................................................................... iii	Table of Contents ......................................................................................................................... iv	List of Tables ................................................................................................................................ ix	List of Figures ................................................................................................................................ x	List of Abbreviations ................................................................................................................. xiv	Acknowledgements ................................................................................................................... xvii	Dedication ................................................................................................................................... xix	Chapter 1: Introduction ............................................................................................................... 1	1.1	 Microtubule Biochemistry .............................................................................................. 1	1.2	 Microtubule Organization in Plants ................................................................................ 4	1.2.1	 Microtubules in dividing cells .................................................................................... 4	1.2.2	 Microtubules in diffusely expanding interphase cells ................................................ 5 1.2.3	 Microtubules in the tip growing cells of root hairs ..................................................... 5	1.3	 Control of Microtubule Dynamics and Organization: Microtubule-associated Proteins, Kinesins, and Kinases .................................................................................................. 7	1.3.1	 Microtubule-associated proteins (MAPs) ................................................................... 7	1.3.2	 Kinesins ....................................................................................................................... 8 1.3.2	 Kinases ...................................................................................................................... 11 1.4	 Research Objectives and Significance of Findings ....................................................... 13	Chapter 2: ARK1 is a plus-end tracking protein that promotes microtubule catastrophe ................................................................................................................................... 18		 v 2.1.	 Background Information ............................................................................................... 18	2.2.	 Materials and Methods .................................................................................................. 19 2.2.1.	 Plant material and culture ......................................................................................... 19	2.2.2.	 ARK1 construct design and cloning strategies .......................................................... 20 2.2.3.	 Generation of transgenic plant materials .................................................................. 21 2.2.4.	 Live-cell imaging ...................................................................................................... 21 2.2.5.	 Image and data analysis ............................................................................................ 22 2.3.	 Results ........................................................................................................................... 23	2.3.1.	 Microtubule catastrophe frequencies and growth velocities are reduced in ark1-1 mutants .................................................................................................................................. 23	2.3.2.	 Low concentrations of oryzalin rescues the ark1-1 microtubule and root hair morphology phenotype ......................................................................................................... 24 2.3.3.	 ARK1 expression occurs in other cell types in addition to root hairs ....................... 26 2.3.4.	 ARK1 accumulates on plus ends of growing microtubules ...................................... 26	2.3.5.	 ARK1 overexpression increases microtubule catastrophe frequency and the amount of time spent in shrinkage phase .............................................................................. 27 2.3.6.	 The N-terminal microtubule binding domain and the C-terminal Armadillo-repeat domain both play a role in ARK1 microtubule localization ...................................... 28 2.3.7.	 ark1-1 mutant phenotypes are restricted to root hairs despite broad ARK1 expression pattern ................................................................................................................. 29 2.4.	 Discussion ..................................................................................................................... 30	2.4.1.	 ARK1 is a plus-end tracking kinesin that promotes microtubule catastrophe .......... 30	2.4.2.	 ARK1 has two domains that enable microtubule localization .................................. 31 	 vi 2.4.3.	 ARK1 function is redundant in cell types besides root hairs .................................... 32 2.4.4.	 ARK1 is an important factor for microtubule turnover and re-modelling ................ 32 Chapter 3: Elucidating the relationship between NEK6 and ARK1 in microtubule dynamics and cell expansion ...................................................................................................... 59	3.1	 Background Information ............................................................................................... 59	3.2	 Materials and Methods .................................................................................................. 62 3.2.1	 Generation of plant material and culture .................................................................. 62	3.2.2	 ARK1 construct design and cloning strategy ............................................................ 63 3.2.3	 Live-cell imaging ...................................................................................................... 63 3.2.4	 RT-PCR ..................................................................................................................... 64 3.3	 Results ........................................................................................................................... 64	3.3.1	 NEK6-GFP and ARK1-TagRFP co-localize to microtubules .................................. 64	3.3.2	 The TagRFP alters ARK1 microtubule localization but not function ...................... 65 3.3.3	 The number of nek6-1 epidermal protrusions are not significantly different from nek6-1 ark1-1 double mutants .............................................................................................. 65 3.3.4	 NEK6-GFP localization does not change in the ark1-1 background ........................ 66	3.3.5	 ARK1-GFP distribution and function does not change in the nek6 mutant ............. 66 3.3.6	 Exogenous Gibberellic Acid and ACC treatments do not affect the NEK6 and ARK1 interaction .................................................................................................................. 67 3.3.7	 Over-expression of the ARM domain inhibits NEK6-GFP expression .................... 68 3.4	 Discussion ..................................................................................................................... 69	3.4.1	 ARK1 does not require NEK6 for localization, function, phosphorylation, and regulation of activity ............................................................................................................. 69		 vii 3.4.2	 NEK6 does not require ARK1 for microtubule localization or function .................. 70 3.4.3	 ARK1 may play a regulatory role in NEK6 expression ............................................ 71 3.4.4	 Is there a physical NEK6-ARK1 interaction in vivo? ............................................... 72 3.4.5	 Conclusions: NEK6 and ARK1 modulate microtubule dynamics and cell expansion through separate pathways ................................................................................... 73 Chapter 4: MOR1 is a plus-end tracking protein that acts as a microtubule polymerase  . 86	4.1	 Background Information ............................................................................................... 86	4.2	 Materials and Methods .................................................................................................. 89 4.2.1	 Generation of plant material and culture .................................................................. 89	4.2.2	 Chemical fixation of seedlings .................................................................................. 90 4.2.3	 Microscopy ............................................................................................................... 90 4.2.4	 Image processing and data analysis .......................................................................... 90 4.3	 Results ........................................................................................................................... 91	4.3.1	 MOR1pro:MOR1-3xYpet rescues the homozygous lethality of the mor1-23 ............. 91 4.3.2	 MOR1 is expressed in all cell types .......................................................................... 91	4.3.3	 MOR1-3xYpet localizes to mitotic and cytokinetic microtubule arrays .................. 92	4.3.4	 MOR1-3xYpet has an asymmetrical plus-end distribution and is plus-end tracking protein ..................................................................................................................... 93	4.3.5	 MOR1-3xYpet localizes to depolymerizing plus ends of severed microtubules at cross-over sites ...................................................................................................................... 94	4.3.6	 MOR1 is able to bind to the microtubule sidewall in addition to its preferential plus end localization ............................................................................................................. 94		 viii 4.3.7	 Chemical fixation affects the plus ends of microtubules and prevents +TIPs from binding .................................................................................................................................. 96	4.4	 Discussion ..................................................................................................................... 97	4.4.1	 MOR1 is a +TIP that functions in promoting microtubule plus-end dynamics…. ... 97	4.4.2	 How does MOR1 interact with microtubules and plus-end track? ........................... 98 4.4.3	 Chemical fixation inhibits localization of plus-end tracking proteins ...................... 99	Chapter 5: Conclusions ............................................................................................................ 123 5.1	 Majors Findings of the Dissertation ............................................................................ 123	5.1.1	 ARK1 maintains proper root hair tip growth by promoting microtubule catastrophe .......................................................................................................................... 123	5.1.2	 ARK1 and NEK6 regulate microtubule dynamics and cell expansion through independent pathways ......................................................................................................... 125	5.1.3	 MOR1 is a plus-end tracking protein ...................................................................... 126	5.2	 Future Directions ........................................................................................................ 127 5.2.1	 Elucidating the mechanism of ARK1 action .......................................................... 127	5.2.2	 Confirming a physical interaction between NEK6 and ARK1 ............................... 128	5.2.3	 Is the NEK6-ARK1 interaction only manifested under specific stress  conditions? .......................................................................................................................... 128	5.2.4	 Determining the function of ARK1 phosphorylation ............................................. 130 5.2.5	 Do the +TIPs interact to regulate plus-end microtubule dynamics? ....................... 131 Works Cited ............................................................................................................................... 133   	 ix LIST OF TABLES	Table 2.1.  List of primer sequences used for cloning the various ARK1 constructs .................... 35	Table 2.2.  Plus-end growth velocities and catastrophe frequencies of wild-type and ark1-1 microtubules in untreated and 100 nM oryzalin-treated root hairs ............................................... 36   	 x LIST OF FIGURES Figure 1.1.  The growth and shrinkage of dynamic microtubules ................................................ 15 Figure 1.2.  Treadmilling microtubules ........................................................................................ 16 Figure 1.3.  Microtubule organization throughout the different stages of a plant cell ................. 17	Figure 2.1.  Root hairs of ark1-1 are wavy and branched ............................................................. 37		Figure 2.2.  Endoplasmic microtubules in ark1-1 are more abundant than in wild-type root hairs ............................................................................................................................................... 38	Figure 2.3.  Cortical microtubule plus-end velocities and catastrophe frequencies are reduced in ark1-1 root hairs but can be rescued by oryzalin treatment ...................................................... 39	Figure 2.4.  EB1b-GFP moves more slowly in ark1-1 root hairs than in wild-type root hairs .... 41	Figure 2.5.  Treatment with oryzalin causes no obvious differences in microtubule organization between wild-type and ark1-1 root hairs ................................................................. 42	Figure 2.6.  Oryzalin partially rescues the ark1-1 root hair phenotype ........................................ 43	Figure 2.7.  ARK1-GFP rescues the ark1-1 root hair phenotype ................................................. 45	Figure 2.8.  ARK1-GFP localizes to microtubules in elongating and fully-grown root hairs ...... 46	Figure 2.9.  ARK1-GFP is expressed in non-root hair cells and labels different microtubule structures ....................................................................................................................................... 47	Figure 2.10.  ARK1-GFP accumulates on growing microtubule plus ends .................................. 48	Figure 2.11.  ARK1-GFP is not found on microtubule minus ends or depolymerizing plus ends ............................................................................................................................................... 49	Figure 2.12.  ARK1-RFP over-expression leads to increased microtubule catastrophe frequencies .................................................................................................................................... 50	Figure 2.13.  The ARM domain aids in ARK1 localization to microtubules ............................... 52		 xi Figure 2.14.  ARKpro:ARK1ΔARM-GFP is sufficient enough to rescue the ark1-1 root hair phenotype and localizes to microtubules in a similar pattern as full length ARK1 ...................... 53	Figure 2.15.  ARK1pro:ARM-GFP shows plus-end tracking abilities on microtubules but is not able to recue the ark1-1 root hair phenotype .......................................................................... 54	Figure 2.16.  The plus-end velocities and catastrophe frequencies in cells expressing ARK1-GFP and ARK1ΔARM-GFP are not significantly different from each other. .............................. 56 Figure 2.17.  Cell and tissue patterns are not affected in ark1-1 root tips .................................... 57 Figure 2.18.  ark1-1 microtubule phenotype is restricted to root hair-forming cells .................... 58 Figure 3.1.  ARK1-TagRFP and NEK6-GFP have overlapping but non-identical distribution on microtubules ............................................................................................................................. 74		Figure 3.2.  ARK1-TagRFP labelling of the entire microtubule length persists in the ark1-1; 35Spro:EB1b-GFP line  .................................................................................................................. 76 Figure 3.3.  There is no significant difference in the number of epidermal protrusions in nek6-1 and nek6-1 ark1-1 mutants in either light- and dark-grown seedlings .............................. 77	Figure 3.4.  NEK6-GFP microtubule distribution on microtubules is not altered in the ark1-1 mutant ........................................................................................................................................... 79	Figure 3.5.  ARK1-GFP remains associated with microtubule plus ends in the nek6-1 mutant  .......................................................................................................................................... 82	Figure 3.6.  Localization of ARK1-TagRFP and NEK6-GFP to microtubules is unchanged when exposed to GA or ACC ....................................................................................................... 82	 Figure 3.7.  Localization of NEK6-GFP on microtubules does not change when plants are exposed to exogenous GA or ACC ............................................................................................... 83		 xii Figure 3.8.  Microtubule Localization of ARK1-GFP is unchanged when exposed to GA or ACC .............................................................................................................................................. 84	Figure 3.9.  Overexpression of ARM-RFP leads to complete loss of NEK6-GFP expression in ibo1-1 ark1-1; NEK6pro:NEK6-GFP UBQ10pro:ARM-RFP plants ............................................... 85 Figure 4.1.  The MOR1 gene structure and placement of mutations of various mor1 alleles ..... 101 Figure 4.2.  The MOR1pro:MOR1-3xYpet construct rescues the mor1-23 homozygous lethality and supports normal vegetative and reproductive development of A. thaliana plants . 102 Figure 4.3.  The MOR1-3xYpet is expressed in all cell types of Arabidopsis seedlings ............. 104 Figure 4.4.  MOR1-3xYpet localizes to PPB microtubules ........................................................ 106 Figure 4.5.  MOR1-3xYpet localizes to microtubules from the preprophase stage to late anaphase stage ............................................................................................................................. 107 Figure 4.6.  MOR1-3xYpet localizes to the phragmoplast at microtubule plus ends and in microtubule-free domains ........................................................................................................... 109 Figure 4.7.  MOR1-3xYpet is asymmetrically distributed on the plus ends of microtubules and is a plus-end tracking protein ............................................................................................... 111 Figure 4.8.  MOR1-3xYpet localizes to depolymerizing microtubule plus ends ....................... 113 Figure 4.9.  MOR1-3xYpet persists on microtubule plus ends undergoing catastrophe and then rescue .................................................................................................................................. 114 Figure 4.10.  MOR1-3xYpet is not found on microtubule minus ends ...................................... 116 Figure 4.11.  MOR1-3xYpet localizes to newly created plus ends from microtubule severing events at cross-over sites ............................................................................................................. 117 Figure 4.12.  MOR1-3xYpet has preferential binding to microtubule plus ends but still has affinity for the microtubule sidewall ........................................................................................... 119 	 xiii Figure 4.13.  Increased levels of MOR1 protein do not affect development of seedlings .......... 120 Figure 4.14.  Chemical fixation of hypocotyls alters the plus ends of microtubules .................. 121  	 xiv LIST OF ABBREVIATIONS 35Spro  Cauliflower mosaic virus promoter α  Alpha β  Beta Δ  Delta γ  Gamma γTuRC  Gamma Tubulin-ring Complex µL  Microliter µM  Micromolar µM  Micrometer +TIP  Plus-end tracking protein ACC  1-Aminocyclopropane-1-carboxylic acid Arg  Arginine ARK1/2/3 ARMADILLIO-REPEAT KINESIN 1/2/3 ARM  Armadillo-repeat Domain ADP  Adenosine diphosphate ATP  Adenosine triphosphate BAC  Bacterial artificial chromosome Cc  Critical concentration °C  Degrees Celsius cDNA  Complementary deoxyribonucleic acid CMT  Cortical microtubule Co-IP  Co-immunoprecipitation D-box  Destruction Box DNA  Deoxyribonucleic acid EB1b  END-BINDING 1b EGTA  Ethylene glycol tetraacetic acid EMT  Endoplasmic microtubule GA  Gibberellic Acid GAP  GTPase Activating Protein 	 xv GC  Guard cell GDP  Guanosine-5’-diphosphate GFP  Green Fluorescent Protein Glu  Glutamic Acid GTP  Guanosine-5’-triphosphate h  Hour kB  Kilo-base pairs MAP  Microtubule-associated protein MAPK  MITOGEN-ACTIVATED PROTEIN KINASE MBD  Microtubule-binding domain min  Minute MOR1  MICROTUBULE ORGANIZATION 1 mRFP  Monomeric Red Fluorescent Protein MS  Mitotic spindle mRNA  Messenger RNA NEK  NEVER-IN-MITOSIS A KINASE NA  Numerical aperture nm  Nanometer nM  Nanomolar NS  Not significant NTC  No template control OX  Over-expression P  Phragmoplast PC  Pavement cell PCR  Polymerase chain reaction PHS1  PROPYZAMINDE HYPERSENSITIVE 1 PIPES  Piperazine-N,N′-bis(2-ethanesulfonic acid) PPB  Pre-prophase band RFP  Red fluorescent protein RNA  Ribonucleic acid RNAi  RNA interference 	 xvi RT-PCR Reverse transcription polymerase chain reaction s / sec  Second Ser  Serine SD  Standard deviation SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis TAC  Transformation artificial chromosome TagRFP Tag Red fluorescent protein Thr  Threonine T-DNA Transfer-Deoxyribonucleic acid  UBQ10pro UBIQUITIN10 promoter YFP  Yellow Fluorescent Protein	 xvii ACKNOWLEDGEMENTS  I would first like to thank Dr. Geoffrey Wasteneys for being an inspiring, supportive, and creative supervisor.  Geoff has been mentoring me since my undergraduate career where he first piqued my interest in cell biology through a course he was teaching.  I was fortunate enough to join his lab as an undergraduate researcher and finally as a graduate student.  Geoff has taught me to become a critical thinker and a hypothesis-driven researcher, two qualities that I am lucky to have acquired through my time in his lab.  The beginning of my research career was started thanks to Geoff and I thank him for being an outstanding mentor.  I am also very grateful to my committee members: Dr. Eric Cytrynbaum, Dr. George Haughn, and Dr. Lacey Samuels.  They are experts within their own fields and have provided valuable ideas and advice towards my thesis work.  They have always provided constructive feedback and have helped me to stay focused on my big research questions.  A great deal of gratitude goes to my funding sources: an NSERC Postgraduate Scholarship, a UBC Four-Year Fellowship, a Craig Adams Sandercock Memorial Scholarship, Teaching Assistant appointments, and a WoW NSERC CREATE training grant. I am truly grateful for the Botany and Biology Department for all their amazing and unwavering support.  I am especially thankful to the Biology Department for providing the best training for teaching.  Thank you to the dedicated members of the teaching faculty who have taught me so much with regards to the pedagogy of teaching and learning.  I owe most of my lab training to the post-doctoral fellows in my lab: Dr. Chris Ambrose, Dr. Miki Fujita, Dr. Bettina Lechner, and Dr. Sylwia Jankowski. They have taught me all the skills I have utilized in my research and none of the data could have come to fruition without them.  I received training and support for most of the microscopy work from the UBC Bioimaging Facility and thank Kevin Hodgson for all his training and help with using the confocal microscopes.  I would also like to thank members of the Wasteneys Lab (past and present): Amanda Catching, Dr. Katherine Celler, Laryssa Halat, Dr. Eric Johnson, Laura Tamblyn, Samuel Livingston, Adam Mulvihill, Dr. Yuan Ruan, Dr. Ankit Walia, and Yi Zhang.  They have provided me with new ideas on experiments to do and are always available to help out with trouble-shooting. A special thanks to members of the Samuels Lab (Dr. Allan Debono, Miranda Meents, Dr. Mathias Schuetz, Yoshi Watanabe), as they have 	 xviii always been great companions to the Wasteneys Lab and provided me with extra support (and sometimes extra petri dishes).  To the all the members of the Working on Walls (WoW) group and Dr. Brian Ellis: I am honoured and privileged to have been a part of the WoW group.  I am truly grateful that we got to experience so much together, including our amazing trip to Sweden. To my figure skating coach and very first mentor, Tina Leininger: thank for teaching me about hard work ethic: on and off the ice.   I would like to thank my friends for all their unwavering love and encouragement.  I wouldn’t have made it through grad school without you.  To my friends from grad school and now life (Anika Benske, Charles Copeland, Jackie Dee, Kaeli Johnson, Dr. Gabriel Levesque-Tremblay, Dr. Heather McFarlane, Dr. Teagen Quilichini, Dr. Rebecca Smith): you are the only ones who understand the pain that is grad school.  Thank you for keeping grad school fun despite the many lows in research.  To my D-15 friends, far and wide (Adam Belovari, Katelyn Campbell, Natasha Errington, Kara Fernandes, Janelle Lajtar, Kristen Lazuka, Matthew MacMurdo, Lindsey Massoglia, Tara Meyers, Courtney Mew, Ashley Wong): our memories together are unforgettable and keep me smiling and laughing for days, especially when I am missing you all.  To Dr. Michelle Chan, Sayaka Endo, Deanna Guilfoyle, Bryant Jones, Matthew Onas, and Jackie Wu: thank you for shaping me into the person to who I am today. I can always count on you for support, advice, and a good eating session. Thanks to you, I know I can do anything. To all my extended family, aunts, and uncles: thank you for your support and constant encouragement.  For my sister and brothers, Krystle, Matthew, and Craig: thank you for pretending to think that what I do is cool and for not thinking of me as the weird middle child. Finally, this thesis would not have been made possible without my Mom and Dad. They have always supported me in anything I wanted to pursue. I will always appreciate their support with regards to my athletics, education, and life. I could not have achieved any of my goals without you.   	 xix         Dedicated to my Mom and Dad, Rita and Harry  	 1 Chapter 1: Introduction  Microtubules are filamentous protein assemblies that are involved in essential cellular functions of eukaryotic organisms.  The processes of mitosis, cytokinesis, and vesicular/organelle transport, for example, depend on microtubules.   In plants, microtubules contribute to the cell’s ability to grow and develop in order to create and maintain complex cell shapes.  Because plants lack centralized microtubule organizing centers, such as centrosomes, the control of microtubule dynamics and organization is highly important and dependent on other important proteins and cellular mechanisms (Ehrhardt and Shaw, 2006a).  Changes in the microtubule dynamics and organization can disrupt a plant’s ability to properly grow and develop.  Disruption of microtubules, either through genetics or chemical approaches, has been well studied in the model organism, Arabidopsis thaliana.  Unless otherwise noted, most of the background information in the following sections of this chapter refers to information derived from experiments utilizing A. thaliana.  In this chapter, I will discuss the biochemistry of microtubules followed by the various microtubule arrays that are found in A. thaliana.  Furthermore, I will review how microtubule dynamics and organization are modulated through microtubule-associated proteins, kinesins, and kinases.  Finally, I will conclude with the research objectives and summary of the major findings of this dissertation. 1.1. Microtubule Biochemistry Microtubules are dynamic hollow filaments made up of repeating α- and β-tubulin subunits collectively known as tubulin heterodimers that are non-covalently bound together (Weisenberg et al., 1968).  The α- and β-tubulin subunits are longitudinally incorporated into the microtubule polymer in the same orientation with the β-tubulin as the leading subunit (Amos and Klug, 1974; Mitchison, 1993).  The β-subunit thus interacts with the α-subunit of the next incoming tubulin heterodimer resulting in a polymer with alternating α- and β-tubulin subunits (Figure 1.1).  Moreover, the heterodimer subunits assemble and interact with other tubulin subunits that are being incorporated in parallel, resulting in further lateral interactions among the heterodimer subunits.  The inherent polarity of the tubulin heterodimers gives itself the ability to self-assemble in a specific orientation to form the cylindrical protofilaments, creating a polar polymer.  Microtubules have commonly an outer 	 2 diameter of 25 nm with 13 tubulin subunits creating the circumference of the cylinder (Amos and Klug, 1974; Desai and Mitchison, 1997; Ledbetter and Porter, 1963, 1964). The stochastic addition or removal of the tubulin subunits from the microtubule ends causes the microtubule to either grow or shrink, respectively (Figure 1.1).  The transition from a growth to shrinkage event is known as a catastrophe (depolymerization) while the inverse is known as a rescue (polymerization).  In addition, the microtubule ends can sometimes be found in pause in which there is no net addition or removal of tubulin (Desai and Mitchison, 1997; Mitchison and Kirschner, 1984; Shelden and Wadsworth, 1993; Walker et al., 1988).  The growth and shrinkage events are dependent on β-tubulin’s catalytic ability to hydrolyze Guanosine-5’-triphosphate (GTP).  Both α- and β-tubulin possess a binding pocket for GTP.  While GTP within the α-subunit is not hydrolysed and remains part of the heterodimer (Spiegelman et al., 1977), the GTP in β-tubulin can be hydrolyzed to Guanosine-5’-diphosphate (GDP) once incorporated in the microtubule (David-Pfeuty et al., 1977; MacNeal and Purich, 1978).  GTP-bound β-tubulin allows the tubulin heterodimer to be easily added to the microtubule forming a stable, straight polymer due to the GTP-bound tubulin’s structural conformation that promotes strong lateral interactions with its neighboring subunits (Caplow et al., 1994; Hyman et al., 1992). This addition of GTP-bound tubulin forms a structure known as a GTP cap that helps stabilize and promote the growth of microtubules (Mitchison and Kirschner, 1984).  Conversely, hydrolysis of GTP resulting in GDP-bound tubulin weakens tubulin’s lateral interactions with its neighbouring subunits, resulting in instability within the polymer and subsequent dissociation of the tubulin from the microtubule (Hyman et al., 1995; Müller-Reichert et al., 1998).  The weakened lateral interactions with GDP-bound tubulin within the microtubule adopt a curved confirmation.  This confirmation leads to heterodimers “peeling off” from the protofilament resulting in catastrophe (Mandelkow et al., 1991).  Once free, the GDP bound within β-tubulin is exchanged for GTP allowing the heterodimer to be incorporated into the microtubule once again (David-Pfeuty et al., 1977; MacNeal and Purich, 1978). Microtubules have both a characteristic minus and plus end with α- and β-tubulin being exposed at each end, respectively (Figure 1.1).  Plus ends are more dynamic because heterodimers can be quickly and easily added or removed from the microtubule.  The minus end, exposed with α-tubulin, is less dynamic than the plus ends of microtubules due to the 	 3 structural restraints of tubulin being added to microtubule minus end (Allen and Borisy, 1974; Desai and Mitchison, 1997; Mitchison, 1993).  The minus end is sometimes capped with another tubulin isoform called a γ-tubulin that makes up a complex called the γ-tubulin ring complex (γTuRC) along with a series of other proteins called γ-tubulin complex proteins (GCPs) (GCP2-GCP5, NEDD1) (Kong et al., 2010; Liu et al., 1993; Murata et al., 2005; Nakamura et al., 2010a; Zeng et al., 2009).  γTuRCs also make the minus end less dynamic because they cap the minus end and prevent polymerization and depolymerization (Zheng et al., 1995).  Moreover, γTuRCs are essential components for creating new microtubule polymers by assisting in microtubule nucleation.  While plant, yeast, and animals cells all possess γTuRCs, the distribution of these complexes is different in acentrosomal plant cells (Ehrhardt and Shaw, 2006a; Hamada, 2007; Wiese and Zheng, 2006).  Animal cells have the γTuRCs being embedded in the centrosomes creating the microtubule-organizing center (Wiese and Zheng, 2006), while the γTuRCs in plants are distributed throughout the cytoplasm where the γTuRCs are known to localize on pre-existing microtubules as well as the cell cortex and nuclear envelope (Erhardt et al., 2002; Hamada, 2007; Kong et al., 2010; Liu et al., 1993; Murata et al., 2005).  Thus, the majority of new microtubules are created via nucleation and branching from γTuRCs (Teixidó-Travesa et al., 2012) that are localized to extant microtubules, cell cortex, and the nuclear envelope.  A more recent study revealed that microtubule nucleation can actually occur without the canonical nucleation process of using γTuRCs (Lindeboom et al., 2013a). This difference in dynamicity between minus and plus ends also derives from the fact that each end has a different characteristic value known as the critical concentration (Cc).  The Cc is defined as the concentration of free tubulin (GTP or GDP) within a cellular system where the on-rate of tubulin to the microtubule is equal to the off-rate.  The concentration of free tubulin relative to the Cc determines the rate at which the microtubule will grow or shrink. In other words, the microtubule growth or shrinkage rate is directly proportional to the concentration of free tubulin.  If the free GTP-bound tubulin concentration is higher than the Cc, then the microtubule will more readily polymerize and form the GTP cap (Figure 1.1).  Once most of the free GTP-bound tubulin is incorporated within the microtubule, the concentration of free GTP-bound tubulin is lowered, thus slowing down the growth rate.  When the free tubulin concentration falls below the Cc, then the microtubule will shift more 	 4 readily to a depolymerization phase.  This occurs because the rate of GTP-hydrolysis within the tubulin subunit (and subsequent removal of GDP-bound tubulin from the polymer) is faster than the on-rate of GTP-bound tubulin, thus causing microtubule depolymerization.  With respect to the Cc of both the plus and minus ends, plus ends have lower Cc values, which is why the microtubule plus ends are able to more quickly grow and shrink relative to the minus ends.  In other words, minus ends require a much higher threshold of free tubulin concentration before the microtubule minus end can grow or shrink and thus remains more static relative to the plus ends (Desai and Mitchison, 1997; Mitchison and Kirschner, 1984).  A process called treadmilling occurs when the tubulin off rate at the minus end is equivalent to the tubulin on rate at the plus end (Figure 1.2).  This property maintains a constant microtubule length and gives the illusion of a microtubule polymer freely moving within the cell and allowing for microtubule migration (Rodionov and Borisy, 1997; Shaw et al., 2003; Walker et al., 1988)  1.2. Microtubule Organization in Plants 1.2.1. Microtubules in dividing cells Microtubules are required for mitosis and subsequent cytokinesis.  Prior to mitosis, microtubules in interphase are located at the cell cortex.  In the late G2 phase of the plant cell cycle, microtubules form a structure known as the pre-prophase band (PPB).  Microtubules in the PPB form a band at the cortex along the plasma membrane, usually mid-distant to the cell ends (Figure 1.3A).  Moreover, microtubule arrays extend from the nucleus to the microtubule bands at the cell cortex (Wasteneys, 2002).  The PPB is a microtubule array that is thought to predict the placement of the nascent cell plate (Rasmussen et al., 2013).  The most widely known role of microtubules in mitosis is the formation of the mitotic spindle, which is responsible for lining up the chromosomes and separating the chromatids to the two daughter cells.  The mitotic spindle appears after the disassembly of the PPB when microtubules re-assemble to form the spindle at the beginning of mitosis (Figure 1.3B) (Wasteneys, 2002).  Finally, microtubules are a major component of a structure called the phragmoplast that mark the site of and create the new cell plate during cytokinesis (Figure 1.3C, D).  In this structure, microtubules form anti-parallel bundles in the area that will be the new cell plate with the plus ends pointing towards the mid-zone (Euteneuer and 	 5 McIntosh, 1980).  Golgi-derived vesicles containing cell wall and plasma membrane components are transported along microtubule tracks (via kinesin motor proteins) to the site of the nascent cell plate where the new cell wall is being deposited (Jürgens, 2005; Lee et al., 2001; Reddy, 2001; Samuels et al., 1995).  As the new cell plate is synthesized and expands to eventually connect with the parental cell wall, the microtubule bundles in the center disassemble and new microtubules assemble at the periphery (Jürgens, 2005; Rasmussen et al., 2013; Staehelin and Hepler, 1996). 1.2.2. Microtubules in diffusely expanding interphase cells After cytokinesis, early interphase cells have two populations of microtubules that either radiate out from the nucleus to the plasma membrane or are arranged at the cortex of the cell (Figure1.3E).  Diffuse expansion then occurs in interphase cells in which growth is evenly distributed along the longitudinal axis of the cell.  This type of unidirectional growth is typically seen in the root and hypocotyl epidermal cells.  Microtubules in these cells are orientated in parallel arrays perpendicular to the longitudinal growth axis at the cortex along the plasma membrane of the cells, known as the cortical microtubule array (Figure 1.3F).  Diffuse expansion is typically ceased when the perpendicular microtubule arrangement reorients into a longitudinal arrangement (Sugimoto et al., 2000).  Any other microtubule arrangement, such as a disordered microtubule organization, leads to abnormal plant cell growth like isotropic expansion.  This is evident with the application of microtubule-disrupting drugs (such as oryzalin or taxol) (Baskin et al., 1994) or microtubule-related mutants where microtubules are disrupted resulting in loss of anisotropic expansion and leading to radially swollen cells (Whittington et al., 2001). 1.2.3. Microtubules in tip growing cells of root hairs Tip growth occurs in cells where growth is highly localized to a specific area in the cell.   With this process in plants, the plasma membrane and cell wall components are secreted at tip of the cells.  This type of growth is evident in the root hairs and pollen tubes of plants.  Under normal conditions, root hairs are tubular cells that protrude perpendicularly from the root and aid in water and nutrient acquisition as well as anchoring the plant in the soil. Microtubules contribute to proper root hair morphogenesis by establishing and maintaining root hair polarity. The drugs oryzalin and taxol, which cause microtubule 	 6 destabilization and stabilization, respectively, can lead to wavy and branched root hairs as a result of loss of proper tip growth, stressing the importance of the role of microtubules in this process (Bibikova et al., 1999). Studies and visualizations of microtubule organization in root hairs have been previously done (Sieberer and Timmers, 2009; Sieberer et al., 2005) but actual studies of microtubule dynamics have been less extensive.  Electron and confocal imaging of microtubules in root hairs have revealed the presence and characterization of two different types of microtubule populations within the root hairs throughout the initiation and elongation processes: the cortical microtubules (CMTs) and endoplasmic microtubules (EMTs).  Initially reported in legumes (Sieberer et al., 2002), the relative abundance of the two microtubule populations appears to vary during tip growth.  Live-cell imaging of a green fluorescent protein fused to a microtubule-binding domain (GFP-MBD) protein in A. thaliana plants showed that the CMTs are present throughout all developmental stages of the root hair while EMTs were only present in elongating hairs (Bruaene et al., 2004).  During elongation, the abundance of CMTs is relatively low at the apex of the root hair tip and non-existent immediately behind the tip because of the dense actin meshwork and vesicles that prevent microtubules from reaching the tip.  CMTs are abundant along the shank of the root hair, localized along the cortex of the root hair (against the plasma membrane). EMTs are usually found as bundles within the root hairs, extending from the nucleus to the tip of the root hair.  EMTs are most abundant in the apex, possibly indicating a more important role in elongation of the root tip compared to the CMTs (Bruaene et al., 2004; Lloyd et al., 1987) .  Since EMTs do not appear in fully elongated root hairs, it is therefore essential to specifically observe elongating root hairs in order to simultaneously view both populations of microtubules.  Fully elongated root hairs lack EMTs and have CMTs that are sparse and less dense than microtubules in elongating root hairs.  Moreover, CMTs are able to reach the tip of the mature root hair whereas CMT are absent in the tip of elongating root hair (Bruaene et al., 2004).   	 7 1.3. Control of Microtubule Dynamics and Organization: Microtubule-associated Proteins, Kinesins, and Kinases In plants, the control of microtubule dynamics and organization are dependent on microtubule-associated proteins (MAPs), kinesins, and kinases.  Because plant cells are acentrosomal, microtubules need to self-assemble and self organize in order to create the various aforementioned microtubule arrays. 1.3.1. Microtubule-associated proteins (MAPs)  In A. thaliana, there is a large variety of MAPs that work together and are responsible for the nucleation, bundling, cross-linking, and stabilizing/destabilizing of microtubules in order to create the various microtubule arrays (Hamada, 2007).  The aforementioned γTuRCs, for example, are MAPs important for microtubule nucleation and branching.  The large family of MAP65 proteins consists of nine MAP65 genes in A. thaliana that code for proteins that localize to interphase, PPBs, mitotic spindles, and the phragmoplast microtubules.  These proteins are expressed at different cell cycle stages and are shown to bundle and crosslink parallel and anti-parallel microtubules in order to organize the microtubule arrays. The MICROTUBULE ORGANIZATION 1 (MOR1) protein found in A. thaliana is a part of the highly conserved XMAP215/Dis1 family of proteins described for Xenopus laevis (XMAP215) (Gard and Kirschner, 1987), Caenorhabditis elegans (ZYG-9) (Matthews et al., 1998), Drosophila (Msps) (Cullen et al., 1999), Saccharomyces cerevisiae (Stu2) (Wang and Huffaker, 1997), Saccharomyces pombe (Dis1, Alp4) (Garcia et al., 2001; Nabeshima et al., 1995, 1), Nicotiana tabacum cv. BY-2 (MAP200/TMBP200) (Hamada et al., 2004), and Dictyostelium (CP224) (Graf et al., 2000).  MOR1 has been shown to promote microtubule dynamicity by aiding in the polymerization and depolymerization of microtubules in A. thaliana (Kawamura and Wasteneys, 2008).  This is evident in plants with the temperature-sensitive mutant allele, mor1-1, showing fragmented microtubule polymers with reduced dynamics at the restrictive temperature (28°C) while showing no obvious differences from wild-type plants at the permissive temperature (Kawamura and Wasteneys, 2008; Whittington et al., 2001).  In addition to fragmented cortical microtubules, mor1-1 plants, at the restrictive temperature, frequently lack PPBs and possess aberrant microtubules within the spindle and phragmoplast structures (Eleftheriou et al., 2005; Kawamura et al., 2006).  Despite showing abnormal phragmoplast microtubules and misplaced or incomplete cell 	 8 plate formation, cells in mor1-1 were able to complete cell division, albeit in a delayed manner (Kawamura et al., 2006). At a larger scale, mor1-1 plants grown at the restrictive temperature lose their ability for proper cell expansion, leading to seedlings with swollen root tips, twisted aerial and root tissue, and crooked/branched root hairs.  Moreover, mor1-1 plants germinated at the restrictive temperature were extremely dwarfed and unable to produce flowers before reaching lethality (Whittington et al., 2001).  Another MOR1 mutant allele known as gemini pollen1 (gem1) showed cytokinetic defects in pollen development (Twell et al., 2002), indicating MOR1’s importance across different cell types and cell cycle stages. Although MOR1’s function has been well-characterized, in vivo live cell imaging of full-length MOR1 has yet to be accomplished and thus its association with dynamic microtubules has not been visualized.  In vivo live-cell imaging using a fluorescently tagged-MOR1 could elucidate the protein’s exact localization on microtubules.  Previously, immunofluorescence of MOR1 in A. thaliana suspension cultures, protoplast cultures, and intact tissue revealed MOR1 localizing to various microtubule arrays including the cortical array, PPB, mitotic spindle, and phragmoplast (Kawamura et al., 2006; Twell et al., 2002).  These studies using fixed samples and antibodies that recognize different domains of MOR1 showed inconsistent microtubule-labelling patterns and thus its localization on microtubules remains unclear.  Based on previous work using MOR1’s homologous proteins, MOR1 is thought to function as a microtubule plus-end tracking protein known as a +TIP.  +TIPs are MAPs that regulate microtubule dynamics and accumulate on the plus ends of the microtubule.  In vitro studies using the MOR1 homologues, XMAP215 (Brouhard et al., 2008) and Stu2 (Breugel et al., 2003), revealed preferential microtubule plus-end binding  and thus it is predicted that MOR1 may function in a similar manner. 1.3.2. Kinesins  Kinesins are molecular motor proteins that interact with microtubules in order to transport cargo or regulate the dynamics and structure of the microtubule arrays.  Kinesins are able to accomplish these cellular tasks because they are mechanochemical enzymes that utilize the energy released from ATP hydrolysis to generate force and perform various types of work related to microtubule functions (Brady, 1985; Vale et al., 1985).  Most conventional kinesins use this energy to transport cargo within a cell by moving along microtubules tracks 	 9 in a step-wise fashion (Svoboda et al., 1993).  Other kinesins use the energy to promote microtubule depolymerization, which is important for modulating microtubule dynamics (Desai et al., 1999; Walczak et al., 1996). Kinesins are characterized by having a highly conserved motor domain “head” that is able to bind both ATP and the microtubule (Vale and Fletterick, 1997).  This protein motor domain contains a catalytic core that is able to hydrolyze one ATP molecule per kinesin step and exchange the recently hydrolyzed-ADP for a new ATP (Schnitzer and Block, 1997).  The binding of ATP and subsequent hydrolysis in the motor domains leads to conformational changes that allows the cargo-carrying kinesins to “walk” along the microtubule or for the catastrophe-inducing kinesins to break apart the microtubule protofilaments (Vale and Fletterick, 1997).  The kinesin head is typically followed by an internal coiled-coil “stalk”, which allows for homodimerization, heterodimerization, and tetramerization of at least two kinesin peptides.  Kinesins that lack a coiled-coil domain generally function as monomers.  Finally, kinesins have a highly phylogenetically-divergent “tail” domain that is able to bind multiple types of cargo including organelles and other cytoskeletal components (Reddy and Day, 2011; Vale and Fletterick, 1997; Zhu and Dixit, 2012).  There are 14 described families of kinesins across all taxa.  Generally, the location and sequence homology of the motor domain within the protein determines a kinesin’s phylogenetic placement.  The genome of A. thaliana is predicted to encode 61 putative kinesin polypeptides, all of which have varying structures and functions within the plant species.  In land plants, some of the kinesin families have been lost or reduced in the number of members while others have an expanded number of members (Reddy and Day, 2011; Zhu and Dixit, 2012). In A. thaliana and other land plants, the number of kinesins known to be involved in transporting organelles and vesicles is reduced relative to animal systems, possibly because actin-based myosin motor proteins are mainly involved in long distance movement of the organelles/vesicles in plants.  These conventional kinesins in plants are predicted to dictate local positioning and pausing of organelles as well as short-distance trafficking of organelles.  One of the kinesins involved in organelle movement is Kinesin-13A (part of the Kinesin-13 family).  Kinesin-13A is predicted to disperse Golgi-stacks throughout the cell as well as aiding in Golgi-vesicle budding (Lu et al., 2005; Wei et al., 2009).  Kinesin-13A has also been found to have another role related to the regulation of 	 10 microtubules (see below).  Another kinesin, FRA1 (which is part of the Kinesin-4 family), is shown to use ATP hydrolysis to drive movement along cortical microtubules (Zhu and Dixit, 2011).  FRA1’s cargo is predicted to be Golgi-derived vesicles containing pectin destined for the cell wall because fra1-5 mutants were observed to have reduced pectin content within the cell wall (Zhu et al., 2015).  Other characterized kinesins in land plants are responsible for creating and organizing the microtubule arrays.  A member of the Kinesin-5 family, AtKRP125c, is predicted to organize interphase CMT arrays, although the exact function remains unknown (Bannigan et al., 2007).  More complex microtubule structures, such as the mitotic spindle, are also formed by AtKRP125c and two Kinesin-14 members: ATK1 and ATK5. AtKRP125c is thought to bundle anti-parallel inter-polar microtubules in the mid-plane the spindle (Bannigan et al., 2007).  ATK1 and ATK5 have also been shown to bundle anti-parallel microtubules at the spindle mid-zone and to use their motor activity to generate the inward forces that allow the spindle to become straightened and to assume the correct length. In addition, ATK1 and ATK5 are responsible for lining up and focusing the minus ends of microtubules to form the spindle poles (Ambrose and Cyr, 2007; Ambrose et al., 2005; Marcus et al., 2002, 2003).  Since plant cells lack conventional microtubule-organizing centers, the formation of the spindle poles via kinesins is essential for its progression through mitosis.  The number of kinesins in plants known to act as catastrophe factors promoting microtubule depolymerization is small.  To date, Kinesin-13A is the only kinesin shown to be a catastrophe factor. It acts specifically to promote microtubule depolymerization in the secondary cell wall pits of developing xylem cells.  The depolymerizing function of Kinesin-13A is similar to that of its homologues in other organisms (Desai et al., 1999; Oda and Fukuda, 2013). The ARMADILLO-REPEAT KINESIN 1 (ARK1) in A. thaliana is hypothesized to act as a microtubule depolymerase.  This prediction is based on the ark1 mutant root hairs, which display a loss of proper tip growth and an increase in endoplasmic microtubule bundles (Sakai et al., 2008).  Although the motor domain is located at the N-terminal end of the protein, ARK1 cannot be placed into any of the 14 eukaryotic kinesin families (Reddy and Day, 2011; Sakai et al., 2008; Zhu and Dixit, 2012) so the exact function of ARK1 	 11 cannot be easily predicted based on homology.  ARK1 has two other related proteins expressed in A. thaliana, ARK2 and ARK3, which also cannot be placed within the 14 kinesin families. Structurally, all three ARKs are predicted to act as homodimers based on the presence of the coiled-coil domains in all three proteins.  Moreover, the C-termini of the ARKs contain a repeating-domain called Armadillo-repeats that have an unknown function (which will be discussed further in section 4.1).  While ARK1 contains three Armadillo-repeats in-tandem, both ARK2 and ARK3 have four Armadillo-repeats.  ARK2 is predicted to regulate microtubule dynamics in root epidermal cells based on the ark2 mutants exhibiting a root twisting phenotype typically associated with altered microtubule dynamics (Sakai et al., 2008).  Recently, the role of ARK3 has been further characterized.  ARK3 has been found to be cell cycle-dependent (Malcos and Cyr, 2011) and involved in the asymmetrical division of meristemoid cells for proper stomatal development (Lau et al., 2014).  ARK3 driven by its native promoter and fused to YFP showed PPB localization.  Moreover, ARK3 microRNA lines resulted in meristemoid-like cells showing reduced physical asymmetry necessary for stomatal development (Lau et al., 2014).  This suggests that ARK3 is involved in forming the PPB for proper asymmetrical cell division of the meristemoid cells.  However, the exact functions of the ARK kinesins in regulating microtubule dynamics are currently unknown.   1.3.3. Kinases  Kinases are enzymes that are able to phosphorylate (i.e. add phosphate groups to) specific amino acid residues of proteins.  Phosphorylation of these macromolecules can either stimulate or inhibit activity related to macromolecule function.  For example, phosphorylating a protein can turn on its enzymatic activity.  Phosphate groups are removed by phosphatases that reverse the actions related to the activated phosphorylated macromolecules.  The cyclic process of phosphorylating and dephosphorylating the macromolecules acts as a continuous molecular switch that allows the macromolecules to easily shift from one state (e.g. active) to another (e.g. inactive) (Hunter, 1995, 19931993; Ubersax and Ferrell Jr, 2007).  With respect to the cytoskeleton, kinases are able to alter microtubule dynamics by phosphorylating MAPs and the components of the microtubules polymers (α-, β-, γ-tubulin) (MacRae, 1997; Vogel et al., 2001; Westermann and Weber, 2003). 	 12  Kinases are known to phosphorylate MAPs in order to regulate their function.  The A. thaliana MAP65-1 functions in a cell cycle-specific manner and is shown to bundle and form cross-bridges between microtubules during interphase, anaphase, and telophase.  In order for proper mitotic progression, MAP65-1 microtubule-binding is inhibited during prophase and metaphase due to hyper-phosphorylation by a cyclin-dependent kinase (CDK) and the mitogen-activated protein kinase (MAPK) 4 and 6 (MPK4, MPK6) (Beck et al., 2010, 2011; Mao et al., 2005; Smertenko et al., 2006).  CDKs are a family of kinases that are cell cycle-dependent while MAPKs belong to a highly distinct subgroup of protein kinases involved in a diverse and complex kinase signaling transduction network.  It was concluded that normal metaphase spindle organization and its progression to anaphase are dependent on the inactivation/phosphorylation of MAP65-1 (Smertenko et al., 2006).  In vitro microtubule assays using the Nicotania benthamiana MAP65-1a (homologue of A. thaliana’s MAP65-2 and MAP65-3), led to reduced microtubule bundling activity after phosphorylation by a NRK (a MAPK tobacco homologue).  In vivo results revealed reduced microtubule bundling in phragmoplasts as a result of phosphorylated (inactive) NtMAP65-1a.  Reduced microtubule bundling is thought to lead to an increase in microtubule turnover in order for phragmoplast expansion to occur (Sasabe et al., 2006).  Kinases are also known to phosphorylate the tubulin subunits of microtubules in order to alter microtubule dynamics.  For example, during hyperosmotic stress in Oryza satvia and A. thaliana, α-tubulin was shown to be phosphorylated, resulting in microtubule depolymerization (Ban et al., 2013).  Although the study noted that the Thr349 residue was phosphorylated, the exact kinase responsible for the phosphorylation was not found. Similarly, a MAPK phosphatase called PROPYZAMIDE-HYPERSENSITIVE 1 (PHS1) was found to have kinase activity that is only made apparent upon osmotic stress and was shown to phosphorylate the same Thr349 residue of α-tubulin and cause microtubule depolymerization (Fujita et al., 2013).  Two other kinases in A. thaliana have been shown to phosphorylate tubulin in order to regulate microtubules: Casein Kinase 1-like 6 (CKL6) and Never In Mitosis A (NIMA)-related Kinase 6 (NEK6).  CKL6 is part of an evolutionarily conserved family of Ser/Thr protein kinases that is able to associate with microtubules in vivo.  Moreover, CKL6 was shown to phosphorylate both soluble β-tubulin as well as whole microtubule polymers in 	 13 vitro.   Finally, based on overexpression analysis and kinase-inactive mutants showing disorganized microtubules, CKL6’s phosphorylation activity is predicted to regulate microtubule dynamics (Ben-Nissan et al., 2008). NEK6 is one of seven NEKs found in A. thaliana with a known role in regulating epidermal cell expansion.  Like CKL6, NEKs belong to a highly conserved eukaryotic family of Ser/Thr kinases (Sakai et al., 2008; Vigneault et al., 2007).  NEK6 shows in vivo microtubule localization as well as in vitro β-tubulin phosphorylation activity (Motose et al., 2008, 2011).  nek6 mutants have ectopic epidermal protrusions in addition to aberrant microtubule organization and dynamics, suggesting that this kinase phosphorylates β-tubulin to maintain well-ordered microtubules in expanding epidermal cells.  Interestingly, NEK6 was found to interact with the ARK kinesins (Sakai et al., 2008), although the evidence relied on binding in a heterologous system and thus, the exact function for the interaction remains unresolved. It will be discussed further in Chapter 3.  Moreover, NEK6 has been found to homo-dimerize and hetero-dimerize with two other family members, NEK4 and NEK5 (Motose et al., 2011).  1.4. Research Objectives and Significance of Findings  The broad goal of my PhD thesis is to determine the function of microtubule-related proteins in microtubule dynamics, microtubule organization, and cell growth in the model organism, A. thaliana.  More specifically, my thesis addresses the following research objectives: 1. To determine the role and function of the uncharacterized kinesin, ARK1, in microtubule dynamics during root hair tip growth (Chapter 2). 2. To determine if ARK1 and NEK6 depend on each other for function and microtubule association during their activities in modulating microtubule dynamics and cell expansion. (Chapter 3). 3. To determine MOR1’s live cell in vivo distribution on microtubule and how this association occurs during microtubule assembly and disassembly (Chapter 4). Chapter 2 addresses Research Objective 1 in order to elucidate the function of ARK1.  ARK1 is predicted to be a kinesin that participates in microtubule dynamics based on 	 14 aberrant microtubule organization and root hair morphology (Jones et al., 2006; Sakai et al., 2008).  This study used molecular-genetic, chemical, and microscopical approaches to answer my research objective.  In this chapter, I concluded that ARK1 promotes microtubule depolymerization. This is significant because to date only one other kinesin has been found to function in this manner in plants.  Moreover, I further characterized the different domains of the ARK1 protein in order to make predictions on its in vivo mechanism in regulating microtubules.  I discovered that the ARM domain of ARK1 is a second microtubule-binding domain in addition to the canonical microtubule-binding region (the motor domain). Chapter 3 addresses Research Objective 2.  The major goal of this research is to determine the relationship between ARK1 and NEK6.  Previous research has predicted a physical interaction between ARK1 and NEK6 (Sakai et al., 2008) but the exact relationship between these two proteins was unknown.  In this chapter, I determined that NEK6 and ARK1 function independently to help modulate microtubule dynamics and organization.  Specifically, I concluded that ARK1 does not rely on NEK6 for its microtubule localization or for its ability to induce microtubule depolymerization and control root hair morphology.  Similarly, NEK6 does not require ARK1 for microtubule localization of function in epidermal cell elongation.   Chapter 4 addresses Research Objective 3.  Previous studies on MOR1 have only visualized MOR1 through MOR1 immunolabelling of chemically fixed samples.  These studies, however, revealed conflicting and inconclusive results on MOR1’s localization to microtubules.  In this chapter, live-cell imaging of MOR1 was achieved for the first time by fusing the protein to a fluorescent reporter and visualizing it with spinning-disc confocal microscopy.  My results confirmed MOR1’s expression pattern and also revealed a more accurate microtubule localization pattern relative to the previous studies.  I was able to characterize MOR1 and found that MOR1 is specifically localized to growing and shrinking to microtubule plus ends but absent on minus ends.   	 15  Figure 1.1.  The growth and shrinkage of dynamic microtubules. (A) Schematic diagram showing GTP-bound tubulin subunits (left) and GDP-bound tubulin subunits (right).  The black arrow represents the hydrolysis of GTP within the β-tubulin subunit with Pi (phosphate) being removed from GTP.  The α-tubulin and β-tubulin are labelled. (B) Schematic diagram showing the polymerization (“rescue”) of microtubules by the addition of GTP-bound tubulin (left) to the polymer and the depolymerization (“catastrophe”) of microtubules by the peeling away of GDP-bound tubulin (right) from the polymer.  The GTP cap (labelled) is made up of GTP-bound tubulin in polymerizing microtubules.  Over time, the hydrolysis of the GTP in the β-tubulin occurs and leads to microtubule depolymerization.  The plus and minus ends are labelled. Figure 1.1 reproduced and adapted from Figure 1 in © (Al-Bassam and Chang, 2011).  Regulation of microtubule dynamics by TOG-domain proteins XMAP215/Dis1 and CLASP?  Trends in Cell Biology, 21(10), 604-614, page 605. By permission from the publisher.Minus-end Plus-end β-subunit α-subunit GTP cap 	 16  Figure 1.2.  Treadmilling microtubules. Diagram showing the mechanism of a treadmilling microtubule.  The blue arrow represents the microtubule.  The microtubule is polymerized by the addition of tubulin subunits at the plus end.  Simultaneously, the microtubule at the minus end is being depolymerized with the tubulin subunits dissociating from the microtubule at the same rate tubulin is being added at the plus end.  This gives the appearance of a microtubule moving within a cell. Time progresses from left to right.   Time  Polymerizing Plus end  Depolymerizing Minus end 	 17  Figure 1.3.  Microtubule organization throughout the different stages of a plant cell. The arrangement of microtubules (in green) varies throughout the different stages of plant cell development. Nuclei and chromosomes are represented in blue. The top and bottom images are of cells at the same stages but represent the cells at different angles. (A) Microtubules form the pre-prophase band.  Microtubules radiate from the nucleus to the cortex. Microtubules are also found as a band around the nucleus and mark the site for the future nascent cell plate. (B) Microtubules form the metaphase spindle that is responsible for segregating the chromosomes into the daughter cells. (C) Early stage of the cytokinesis phragmoplast.  Microtubules are responsible for proper positioning and formation of the new cell plate separating the two new daughter cells. (D) Later stage of the cytokinesis phragmoplast.  The phragmoplast radiates towards the cell wall. (E) End of cytokinesis and the beginning of interphase. Microtubules radiate out form the nucleus and are arranged at the plasma membrane. (F) Interphase.  Microtubules are arranged along the plasma membrane at the cortex of the cell perpendicular to the longitudinal growth axis. Figure 1.3 reproduced from Figure 1 in © (Wasteneys, 2002).  Microtubule organization in the green kingdom: chaos or self-order?  Journal of Cell Science, (114), 1345-1354, page 1347. By permission from the publisher.   	 18 Chapter 2: ARK1 is a plus-end tracking protein that promotes microtubule catastrophe 2.1. Background Information The ARMADILLO-REPEAT KINESIN 1 (ARK1), originally named MORPHOGENESIS OF ROOT HAIR 2 (MRH2) (Jones et al., 2006), is a kinesin thought to control microtubule dynamics (Sakai et al., 2008).  Because ARK1 is an uncharacterized kinesin belonging to an ungrouped plant-specific family of kinesins, its function cannot be predicted based on phylogenetic placement (Reddy and Day, 2011; Zhu and Dixit, 2012).  Most recently, the moss Physcomitrella patens was found to have an ARK-like kinesin but with an undetermined function (Miki et al., 2014).  However, with previous evidence showing that ark1 mutants have root hairs with wavy/branched morphologies (Figure 2.1) and increased endoplasmic microtubule abundance (Figure 2.2), it has been suggested that ARK1 specifically promotes microtubule depolymerization in root hairs (Sakai et al., 2008; Yoo et al., 2008).  A single ARK1 polypeptide has an N-terminal catalytic motor domain that interacts with microtubules through the binding and hydrolysis of ATP (Sakai et al., 2008).  Much like most conventional kinesins, ARK1 is predicted to function as a homodimer, as evident with an internal coil-coiled domain.  The C-terminus of ARK1 is composed of three tandem Armadillo (ARM) repeat domains with unknown function that gives rise to the kinesin’s name (Sakai et al., 2008).  ARM repeats are made up of a sequence of 42 amino acids that forms three α-helices and, when in tandem, forms a super-helix that allows for its interaction with other proteins.  In ARM-containing protein, the number of ARM repeats can vary and have been shown to function in essential cellular processes related to signaling, the cytoskeleton, and protein-protein interactions in a variety of eukaryotes (Tewari et al., 2010).  In A. thaliana, there are 108 proteins with ARM repeats (Mudgil et al., 2004), but a large majority of them remain uncharacterized.  41 of the 108 predicted ARM-proteins are thought to be members of the U-Box E3 Ubiquitin Ligase family of proteins (Mudgil et al., 2004).  Other studies of characterized ARM-proteins have shown they have roles in F-actin regulation in pollen tube tip growth (Gebert et al., 2008), regulation of ABA response and salt inhibition of seed germination (Bergler and Hoth, 2011), and formation of lateral roots 	 19 (Coates et al., 2006).  However, the ARM-repeats’ role in ARK1 function is still unclear.  Interestingly enough, previous studies using in vitro analyses have shown that the ARM domain of ARK1 is able to bind to actin (Yang et al., 2007) as well as the NIMA-related kinase, NEK6 (see Chapter 3; (Sakai et al., 2008)).   ARK1 is also thought to play a pivotal role in the tightly coordinated tip growth signalling pathway as a potential linker among the cytoskeletal, endomembrane, and GTPases, and GTPase Activating Protein (GAPs) components in order to sustain proper root hair tip growth.  For example, ark1 in the mutant backgrounds of AGD1 (a gene encoding for a GAP) as well as ROP2 and RABA4b (both small GTPases involved in tip growth regulation) showed an enhanced loss of root hair morphology, suggesting an overlap in the tip growth-signaling pathway for the proteins (Yoo and Blancaflor, 2013; Yoo et al., 2008) The major research objective of the work described in this chapter is to determine the exact function of ARK1 in regulating microtubule dynamics in A. thaliana.  Because of the increase in endoplasmic microtubules found in elongating root hairs (Sakai et al., 2008), I hypothesized that ARK1 plays a role in inducing microtubule depolymerization.  Although previous studies have confirmed ARK1’s ability to bind microtubules through fluorescent protein tagging and in vitro microtubule binding assays (Yang et al., 2007), its function in modulating microtubule dynamics and/or organization has not been confirmed.  In this chapter, I further characterize the root hairs and microtubules found in ark1 mutants.  Moreover, I utilized loss and gain of function experiments to show evidence that ARK1 is a microtubule catastrophe factor.  Imaging of ARK1-GFP revealed that this kinesin has plus-end tracking properties and is specifically localized to growing plus ends.  Furthermore, I provide evidence that suggests putative functional roles for each protein domain of ARK1. 2.2. Materials and Methods 2.2.1. Plant material and culture A. thaliana wild-type (Col-0 ecotype) and ark1-1 plants with the 35Spro:GFP-MBD transgene were used as described (Motose et al., 2008). The 35Spro:EB1b-GFP reporter line (Mathur et al., 2003) and the 35Spro:mCherry-MAP4MBD construct (Gutierrez et al., 2009) were kindly provided by Jaideep Mathur (University of Guelph, Canada) and David Erhardt (Carnegie Institute of Science, Stanford, USA) respectively.  	 20 All seeds were sterilized in 70% ethanol, rinsed three times with ddH2O, and plated onto Petri dishes with Hoagland’s media (1.2% Bacto-agar (BD Diagnostics), no sucrose).  Plates with seeds were stored in the dark at 4°C for 2-3 days and transferred to a 21°C growth cabinet (24 h light) where they were grown vertically until imaging.  For the drug studies, seeds were initially vertically grown on Hoagland’s media supplemented with DMSO (Fisher-Scientific) and then transferred to Hoagland’s media containing various concentrations of taxol (Sigma-Aldrich) or oryzalin (Sigma-Aldrich) for two days prior to imaging.  2.2.2. ARK1 construct design and cloning strategies Gateway Cloning Technology (Invitrogen) was used for the ARK1 (At3g54870) genomic and coding sequences.  To generate the ARK1pro:ARK1-GFP construct, the ARK1 genomic sequence (between 888 base pairs upstream of the ATG/start codon and the TGA/stop codon) was amplified from the F28P10 BAC (from the Arabidopsis Biological Resource Center, Ohio State University) using the full length ARK1 genomic sequence primer set (see Table 2.1 for sequences). A second PCR with the attB-adapter primers was performed according to the manufacturer’s protocol (see Invitrogen for attB-adapter sequences).  For the amplification of the ARK1 coding sequences, different primers were used to amplify the cDNA templates for the ARK1, ARK1ΔARM, ARK1ΔMotor, and ARM constructs (see Supplementary Table 1 for primer list).  A second PCR with the attB-adapter primers was performed according to manufacturer’s protocol (Invitrogen).  The mRNA extraction and cDNA synthesis protocol were performed as described (Galway et al., 2011).  Following the BP reaction with the various attB-PCR products and the pDONR221 vector (Invitrogen), a LR reaction was performed with the pMDC107 vector (Curtis and Grossniklaus, 2003) for the ARK1 genomic sequence and the pUBC-RFP DEST vector (Grefen et al., 2010) for the various ARK1 coding sequences.  The constructs were sequenced prior to transformation. For the ARK1ΔARM-GFP and ARM-GFP constructs driven by the ARK1 promoter, Multisite Gateway Cloning technology was used (Invitrogen).  First, the putative ARK1 promoter was amplified from the F28P10 BAC template using primers designed for Multisite Gateway Cloning with same PCR protocol as before (see Table 2.1 for sequences).  The ARK1 promoter PCR product with the attB1/4 sites was then recombined into the 	 21 pDONR221 P4-P1r using the BP Clonase II (Invitrogen).  A multisite Gateway LR reaction was then performed using the pENTR-ARK1pro, pENTR-ARK1ΔARM/pENTR-ARM (generated from above for the UBQ10 constructs), and the binary vector, R4pGWB504 (with a C-terminal sGFP tag) (Nakagawa et al., 2008) using the LR Clonase II plus enzyme (Invitrogen).  The constructs were sequenced prior to transformation. 2.2.3.  Generation of transgenic plant materials For stable transgenic lines, the ARK1 promoter constructs were first transformed into Agrobacterium tumefaciens (GV3101 strain) and then transformed into an Arabidopsis thaliana ark1-1 35Spro:mCherry-MAP4MBD line using the floral dip method (Clough and Bent, 1998). T3 lines homozygous for the ARK1pro:GFP transgenes were segregated and used for further experiments. Transient expression in cotyledons of the four different UBQpro-driven ARK1 fragment-RFP constructs was performed in ark1-1; 35Spro:GFP-MBD plants using the FAST technique (Li et al., 2009).  ark1-1 (SALK_035063) plants were crossed into the 35Spro:EB1b-GFP reporter line  (Mathur et al., 2003) and homozygous F3 ark1-1 and wild-type plants were segregated for imaging.	2.2.4 Live-cell imaging For observing root hair morphology, whole seedlings were mounted on slides with coverslips and bright-field images of root hairs were then collected with a 20x (air) objective lens on a Leica DMR Light Microscope with a Q-CAM digital camera (Leica). Live imaging of the various microtubule reporter proteins, the ARK1pro-driven ARK1-GFP, and the UBQ10pro-driven ARK1-RFP was done using a Perkin Elmer Ultraview VoX Spinning Disc Confocal system (Perkin-Elmer) mounted on a Leica DMI6000 B inverted microscope and equipped with a Hamamatsu 9100-02 electron multiplier CCD camera (Hamamatsu). An argon 488 nm laser line with a complementary GFP (525/36) emission band-pass filter or a 561 nm laser with a complementary RFP (595/50) emission band-pass filter was used.  Images were acquired with a 63x (water) objective lens (NA = 1.2) every 8 seconds for 3-5 minutes with 0.3 – 0.5 µm optical z-slices. For imaging of microtubule dynamics during drug treatments, seedlings grown on Hoagland’s medium with 100 nM oryzalin were mounted in 100 nM oryzalin.  The imaging temperature of the samples was maintained at 21°C using a Bionomic Controller BC-110 with a HEC-400 Heat 	 22 Exchanger, a Bionomic Controller BC-100 (20-20 Technology Inc.) temperature-controlled stage and an objective heater (Bioptechs). For variable-angle Total Internal Reflection Fluorescence (TIRF) microscopy, 7-day dark grown hypocotyls were imaged on a Zeiss Axiovert Z1 microscope with a Zeiss TIRF III slider, a diode-pumped solid state laser (wavelengths: 488 nm, 561 nm), and a Rolera EM-C2 EM-CCD camera.  Images were visualized with a Zeiss 63x (oil) objective lens (NA = 1.46) and acquired every 8 seconds for 3 minutes. Propidium iodide staining of the cell wall in the root tips was done by incubating whole wild-type and ark1-1 seedlings in 10 µg/mL propidium iodide (Calbiochem) for one minute, rinsing them with water, and mounting them on slides and coverslips.  Laser confocal images of the stained root tips were done on a Zeiss AxioImager M1 microscope with a Zeiss PASCAL Excite two-channel LSM 780 system (Carl Zeiss). A Helium-Neon 543 nm laser line and a 560 nm emission Long-pass filter were used.  Images were acquired in 2 µm optical z-slices with a 63x (oil) objective lens. 2.2.5. Image and data analysis All images were processed and analyzed using ImageJ (http://rsbweb.nih.gov/ij/).  For root hair length analysis, line selections were superimposed on root hairs and then measured.  For quantification of endoplasmic and cortical microtubules, confocal images of wild-type and ark1-1; 35Spro:GFP-MBD root hairs were analyzed in Image J and quantified according to Sakai et al. (2008).  Measuring the microtubule velocities of the EB1b-GFP, GFP-MBD, ARK1-GFP, and ARK1ΔARM-GFP constructs were done using the Manual Tracking plug-in (http://rsbweb.nih.gov/ij/plugins/track/track.html).  EB1b-GFP particles were considered endoplasmic when images from the medial confocal plane of the root hair were analyzed in the endoplasmic region of the root hair.  Cortical EB1b-GFP particles were measured and analyzed from cortical plane of the root hairs or along the cortex/plasma membrane in the medial confocal plane of the root hair.  For determining the catastrophe frequency of EB1b-GFP, the inverse of the duration of time spent tracking one EB1b-GFP comet was taken.  This is based on the assumption that the disappearance of the EB1b-GFP signal meant the microtubule was undergoing catastrophe since EB1b-GFP has highest affinity for the growing plus end of microtubules.  The same method for determining 	 23 catastrophe frequency was used with the GFP-MBD marker in the ARK1 overexpression analysis.  For determining the amount of time spent in each phase, the amount of time spent in one phase (i.e., growth, shrinkage, or pause) was divided by the total measured time of a microtubule lifespan (using the GFP-MBD marker).  The mean, standard deviation, F-tests, and T-tests were calculated using Excel (Microsoft). I also sought to measure potential changes in shrinkage velocities and rescue frequencies by imaging the microtubule markers 35Spro:GFP-MBD and UBQ1pro:mRFP-TUB6 in root hairs.  However, I was unable to accurately observe and quantify plus-end dynamics of single microtubules with these markers in root hairs because of increased microtubule bundling, dense microtubule populations, or high background fluorescence.   For quantifying the ratio of fluorescence intensity to mean fluorescence intensity, the equations below were used.  The procedure involved taking 3 µm line scans on microtubule plus ends from ARK1-GFP images and then measuring the fluorescence intensities/grey values (𝑥!) every 0.108 µm along the line-scan using the Plot Profile function in ImageJ.  The fluorescence intensities were then averaged (𝑥) by the total number of points (𝑁) measured in the line scan (Equation 1).  Each fluorescence intensity (𝑥!) value was then divided by the mean fluorescence intensity (𝑥) to get the desired ratio (𝑅!) at each point along the 3 µm line-scan (Equation 2).  All ratios are reported as mean values.  The same measurements were done with the mCherry-MAP4 MBD images as a control. (1) 𝑥 = 𝑥! + 𝑥! + 𝑥!…+  𝑥!𝑁   (2) 𝑅! = 𝑥!𝑥  2.3. Results 2.3.1 Microtubule catastrophe frequencies and growth velocities are reduced in ark1-1 mutants Based on the fact that endoplasmic microtubules are more abundant in the T-DNA insertional mutant ark1-1 (SALK_035063) (Sakai et al., 2008) and later confirmed through my own analysis (Figure 2.2), I hypothesized that microtubule dynamics would be altered. In order to test this hypothesis, I quantified microtubule plus-end growth rates as well as the 	 24 frequency at which microtubules undergo catastrophe by tracking the microtubule plus-end tracking protein END BINDING 1b (EB1b). This reporter was tagged with green fluorescent protein (GFP) and expressed using the 35S Cauliflower Mosaic Virus promoter (35Spro:EB1b-GFP) and thus I was able to visualize microtubule plus ends.  EB1b-GFP movement was measured in wild-type and ark1-1 elongating root hairs by spinning-disc laser confocal microscopy.  Both endoplasmic and cortical microtubule mean growth velocities were significantly reduced in ark1-1 (CMT: 3.5±1.1 µm/minute; EMT: 5.7±1.3 µm/minute) relative to wild type (CMT: 6.5±1.7 µm/minute; EMT: 7.1±1.9 µm/minute; two-sample unequal variance t-test: CMT: P<10-77; EMT: P<10-05) (Figure 2.3A, 2.3C, 2.4; Table 2.2).  The frequency of cortical microtubule catastrophe was significantly reduced by 31% in ark1-1 (0.020±0.011 events/second) relative to wild-type root hairs (0.029±0.015 events/second; two-sample unequal variance t-test: P<10-09) (Figure 2.3B; Table 2-2).  In contrast, I found no significant difference in catastrophe frequency between wild-type (0.048±0.025 events/second) and ark1-1 endoplasmic microtubules (0.055±0.034 events/second) (Figure 2.3B; Table 2-2) despite a greater abundance of endoplasmic microtubules in ark1-1 root hairs (Figure 2.2).  2.3.2 Low concentrations of oryzalin rescues the ark1-1 microtubule and root hair morphology phenotype The decreased frequency of cortical microtubule catastrophe in ark1-1 mutants is consistent with the increased microtubule polymer mass and the occurrence of microtubule bundling. The slower rate of microtubule polymerization, however, is more difficult to explain because decreased polymerization rates should theoretically reduce the polymer mass.  Given that microtubule polymerization rates are proportional to free tubulin concentrations, I considered the possibility that the reduced polymerization rates in ark1-1 are the consequence of a reduced free tubulin concentration, which is likely to result from the reduced catastrophe frequency.  I hypothesized that increasing the availability of free tubulin should restore normal microtubule growth rates in ark1-1. To achieve this, I applied low concentrations of the microtubule destabilizing drug oryzalin and measured microtubule growth and catastrophe rates using the EB1b-GFP reporter. 	 25 Exposure to 100 nM oryzalin increased the ark1-1 plus-end growth rates (as measured by EB1b-GFP) such that they were equivalent to the oryzalin-free wild-type growth rate in both cortical and endoplasmic microtubule populations (Figure 2.3A; Table 2.2).  Microtubules in wild-type root hairs, in contrast, showed reduced plus-end growth rates when exposed to 100 nM oryzalin (Figure 2.3A; Table 2.2), which is consistent with previously published data (Nakamura et al., 2004).  The cortical microtubule catastrophe frequency in 100 nM oryzalin-treated ark1-1 root hairs was increased relative to that of untreated wild-type and ark1-1 cells. The dynamics of endoplasmic microtubules in oryzalin-treated ark1-1 root hairs and untreated wild-type root hairs were not significantly different (Figure 2.3B; Table 2.2).   Despite a clear difference in plus-end growth velocities of oryzalin-treated wild-type and ark1-1 microtubules as assessed with EB1b-GFP, both genotypes exposed to the same oryzalin concentrations had fragmented microtubules that looked indistinguishable from each other when using the 35Spro:GFP-MBD (MBD = microtubule-binding domain of MAP4) marker (Figure 2.5).  This observation possibly resulted from increased resistance to microtubule destabilizing drugs and bundling typically associated with 35Spro:GFP-MBD marker (Lechner et al., 2012; Marc et al., 1998).  Based on the restoration of wild-type microtubule growth rates in the ark1-1 mutant by the oryzalin treatments, I hypothesized that these treatments would ameliorate the ark1-1 root hair morphological defects.  It was previously shown that wavy and branched root hair phenotypes were indicative of a less severe and more severe loss of root hair polarity, respectively, and that the severity of polarity loss was positively correlated with increasing oryzalin and taxol concentrations (Bibikova et al., 1999).  Exposure to 100 nM, 1 µM, and 5 µM oryzalin partially rescued the ark1-1 root hair phenotype by significantly increasing the root hair length relative to untreated ark1-1 root hairs (Figure 2.6A, 2.6B) (α = 0.01).  Moreover, the frequency of the most severe root hair phenotype (branched) appeared to decrease with the less severe phenotype (wavy and bulbous) increasing in frequency at 1 and 5 µM oryzalin concentrations (Figure 2.6C). In the wild type, these same oryzalin concentrations decreased root-hair length and led to wavy and branched root hairs that resembled ark1-1 phenotypically (Figure 2.6A - 2.6C).  10 µM oryzalin was unable to 	 26 partially rescue the ark1-1 phenotype, suggesting a saturation effect of oryzalin on microtubules (Figure 2.6B - 2.6C).  In contrast to the oryzalin treatments, taxol, which promotes microtubule polymerization, further impaired root hair elongation in both wild-type and ark1-1 root hairs (Figure 2.6A, 2.6B) and increased root hair waving/branching (Figure 2-6A). 2.3.3. ARK1 expression occurs in other cell types in addition to root hairs In order to see where ARK1 is localized on the microtubule, I made an ARK1pro:ARK1-GFP construct using the entire ARK1 genomic sequence including the putative 5’-end promoter region (888 base pairs upstream of the ARK1 start codon).  ark1-1 plants expressing 35Spro:mCherry-MAP4MBD were transformed with the ARK1pro:ARK-GFP construct.  ARK1pro:ARK1-GFP expression rescued the ark1-1 root hair phenotype (Figure 2.7), indicating that the GFP-tagged ARK1 protein was fully functional. ARK1-GFP labelled microtubules and was expressed in both elongating and fully-grown root hairs (Figure 2.8) although characterization of ARK1-GFP on microtubules was difficult to analyze in root hairs due to high cytoplasmic fluorescence of ARK1-GFP.  I was, however, able to observe ARK1-GFP expression in other cell types, including root atrichoblasts as well as epidermal cells of the hypocotyl, petiole, cotyledons, and root tip, which is consistent with previous ARK1 gene expression analysis (Figure 2.9) (Sakai et al., 2008; Yang et al., 2007).  ARK1-GFP labelled other microtubule structures including the pre-prophase band (PPB), the mitotic spindle (MS), and the phragmoplast (P) (Figure 2-9). 2.3.4.  ARK1 accumulates on plus ends of growing microtubules In order to best characterize ARK1 on microtubules, I used cotyledon epidermal cells in order to visualize ARK1-GFP under spinning disc confocal microscopy.  In these cell types, I observed ARK1-GFP plus-end tracking on growing microtubules (Figure 2.10A). Time-lapse imaging and kymographic analysis revealed that ARK1-GFP accumulates at and tracks along growing microtubule plus ends (Figure 2.10B).  ARK1-GFP was also observed along the microtubule sidewalls, although the fluorescence was not as strong (Figure 2.10A – 2.10C).  Quantifying the ratio of local fluorescence intensity to the mean fluorescence intensity obtained from ARK1-GFP line-scans confirmed these observations.  As a control, the local to mean fluorescence intensity ratio for the mCherry-MBD fluorescence was 	 27 approximately 1:1 at each point along the length of microtubules, indicative of even distribution of the fluorescent protein (Figure 2.10C).  In contrast, ARK1-GFP showed the highest intensity ratio closest to the microtubule plus end, with the ratio sharply declining at points farther away from the tip (Figure 2.10C). I did not detect ARK1-GFP on the minus ends of treadmilling microtubules (2.11A) or on depolymerizing microtubule plus ends (2.11B), indicating that ARK1 is predominantly found at the plus-end of growing microtubules. 2.3.5. ARK1 overexpression increases microtubule catastrophe frequency and the amount of time spent in shrinkage phase Since I saw decreased microtubule catastrophe frequencies in ark1-1, I hypothesized that ARK1 overexpression would increase the incidence of microtubule catastrophe.  In order to test this, I made an ARK1-RFP translational reporter construct driven by a UBIQUITIN10 promoter (Figure 2.12A) and transiently expressed it in ark1-1 Arabidopsis cotyledons from lines previously stably transformed to express the GFP-MBD microtubule marker under the 35S promoter. This experimental system, using the FAST technique involving Agrobacterium-mediated transformation of germinating seedlings (Li et al., 2009), made it possible to compare microtubule dynamics in cells overexpressing ARK1-RFP with non-transformed cells within the same cotyledon, which act as internal negative controls.  In transformed cells, ARK1-RFP appeared to label the entire length of microtubules and remained bound to both growing and shrinking microtubules (Figure 2.12B), indicative of the high levels of ARK1-RFP expression under the UBQ10 promoter.  The cells expressing ARK1-RFP had a significantly increased catastrophe frequency (0.034 ± 0.032 events/second) relative to cells not expressing ARK1-RFP (0.019 ± 0.011 events/second; two-sample unequal variance t-test: P<0.02807) (Figure 2.12C).  ARK1-RFP microtubules spent a greater amount of time in a shrinking phase (ARK1-RFP: 21.2%; negative control: 14.2%) (Figure 2.12D) yet there was no significant difference in the microtubule shrinkage velocity between cells expressing ARK1-RFP (7.4±4.1µm/minute) and cells not expressing the construct (7.3±4.1 µm/minute) (Figure 2.12-E).  In contrast, the growth velocity of ARK1-RFP overexpressing cells (3.2±1.6 µm/minute) was reduced significantly by approx. 20% relative to cells not expressing ARK1-RFP (3.6±1.8 µm/minute; two-sample equal variance t-test: P<2.6x10-5) (Figure 2.12E).   	 28 Stable transgenic lines expressing the UBQ10pro:ARK1-RFP construct could not be recovered. Some transgenic lines were identified according to antibiotic resistance but, possibly due to post-transcriptional silencing, these lines neither displayed RFP fluorescence nor did they rescue the ark1-1 root hair phenotype.  This suggests that overexpression of the ARK1 kinesin is detrimental to the cell and results in embryo lethality. 2.3.6. The N-terminal microtubule binding domain and the C-terminal Armadillo-repeat domain both play a role in ARK1 microtubule localization To determine the function of the different ARK1 domains, I first performed domain deletion analysis by engineering three RFP-fusion constructs driven by the UBIQUITIN10 promoter: a construct missing the C-terminal Armadillo-repeat domain (ARK1ΔARM-RFP), a construct missing the N-terminal motor domain (ARK1ΔMOTOR-RFP), and a construct with just the Armadillo-repeat domain (ARM-RFP) (Figure 2-13A).  ARK1ΔARM-RFP showed strong microtubule labeling, confirming that the motor domain can bind microtubules (Figure 2.13B).  Surprisingly, the ARK1ΔMOTOR-RFP (which includes the coiled-coil domain) (Figure 2.13B) also labeled microtubules, suggesting that the coiled-coil and/or the C-terminal ARM domain can bind microtubules independently of the motor domain.  The ARM-RFP construct also labelled microtubules (Figure 2.13B), indicating that the ARM repeats comprise a second microtubule-binding domain and that this domain is able to associate with microtubules even in monomeric form (as a result of deleting the coiled-coil domain). Because these constructs were overexpressed and could not be stably transformed into plants, I created two constructs, ARK1ΔARM-GFP and ARM-GFP driven by the ARK1 promoter, to determine how endogenous expression levels of the truncated kinesin affects its function and its ability to associate with microtubules.  These constructs were transformed into the ark1-1; 35Spro:mCherry-MAP4MBD transgenic line.  Surprisingly, the ARK1pro: ARK1ΔARM-GFP construct was able to rescue to the ark1-1 root hair phenotype (Figure 2.14B) and displayed a similar microtubule labeling pattern as the full length ARK1-GFP (compare distribution pattern in Figure 12.14A to Figure 12.14B).  Interestingly, the ARM-GFP had bright GFP punctate labelling at the plus ends of microtubules, reminiscent of the plus-end marker, EB1b-GFP (Figure 12.15A-C).  Like EB1b-GFP, the ARM-GFP labelled only growing plus ends of microtubules (Figure 12.15A).  While ARK1-GFP and 	 29 ARK1ΔARM-GFP were constitutively expressed in various cells types, ARM-GFP had patchy expression (Figure 12.15A).  Moreover, GFP appeared to accumulate in highly-connected vesicular tubules that appear to be the endoplasmic reticulum (ER) (Figure 12.15D).  These observations suggest the instability of this fusion protein leading to either protein degradation or improper translation of the ARM-GFP construct, which may also lead to ARM-GFP’s patchy expression pattern.  Even though ARM-GFP was able to plus-end track like +TIPs, the ARK1pro: ARM-GFP construct was not sufficient to rescue the ark1-1 root hair phenotype (Figure 12.15E). Even though the ARK1pro:ARK1ΔARM-GFP construct rescued the ark1-1 root-hair phenotype and appeared to localize to microtubules in a similar way as the full-length ARK-GFP construct, I wanted to confirm if there were any subtle changes in the ARK1ΔARM-GFP movement.  Using variable-angle total internal reflection fluorescence (TIRF) microscopy, I was able to clearly track ARK1ΔARM-GFP and ARK1-GFP particles and measure microtubule growth velocities and catastrophe frequencies in etiolated hypocotyls.  Velocities of both ARK1 constructs on microtubules (ARK1-GFP: 5.6±0.9 µm/minute; ARK1ΔARM-GFP: 5.9±0.8 µm/minute) and catastrophe frequencies of microtubules in plants expressing both constructs (ARK1-GFP: 0.019±0.009 events/second; ARK1ΔARM-GFP: 0.016±0.008 events/second) were not significantly different from each other, further suggesting that the ARM domain is not critical for ARK1 function. 2.3.7. ark1-1 mutant phenotypes are restricted to root hairs despite broad ARK1 expression pattern The expression of ARK1-GFP in cell types other than root hairs under its endogenous promoter prompted me to explore ARK1 localization on other microtubule populations and to identify potential mutant phenotypes in non-root hair cell types.  In root tips of ark1-1; ARK1pro:ARK1-GFP plants, ARK1-GFP labelled PPBs, mitotic spindles, and phragmoplasts (Figure 2.9).  Despite ARK1’s association with microtubules in mitotic and dividing cells, I did not detect any phenotypes related to cell size or cell plate positioning in ark1-1 mutants after visualization of root tip cells using propidium iodide-labelled cell walls (Figure 2.17).  Moreover, cotyledon pavement cells, atrichoblasts, and hypocotyl epidermal cells in ark1-1 mutants showed no morphological differences from wild-type equivalents as previously shown (Sakai et al., 2008; Yang et al., 2007).  No significant differences in growth velocities 	 30 and catastrophe frequencies were noticed using the EB1b-GFP marker in wild-type and ark1-1 cotyledons (Figure 12.18A – 12.18B).  Similarly, measuring microtubule growth and shrinkage velocities with the GFP-MBD microtubule marker showed no significant differences in wild-type and ark1-1 elongating hypocotyls (Figure 12.18C).  2.4. Discussion 2.4.1. ARK1 is a plus-end tracking kinesin that promotes microtubule catastrophe ARK1 has not been grouped into any of the 14 conventional kinesin families, which means that its function could not easily be predicted based on sequence homology.  Using translational reporters, overexpression and loss-of function analysis, I have determined that ARK1 is a microtubule plus-end tracking catastrophe factor.  The exact mechanism by which it promotes catastrophe remains to be determined. Specifically, it is unknown if the motor/ATP catalytic domain is used for kinesin movement along the microtubule and/or for physically removing tubulin heterodimers from the protofilament (Akhmanova and Steinmetz, 2008).  In order to provide more mechanistic and molecular data, in vitro experiments need to be done using purified ARK1 and tubulin; however, ARK1 synthesis through heterologous gene expression systems (Escherichia coli and S. cerevisiae) and in planta protein extraction-purification methods proved unsuccessful in obtaining purified ARK1. Since my data shows ARK1 is a plus-end tracking depolymerase, we can predict ARK1’s mechanism for promoting microtubule depolymerization by looking at similar kinesins with similar function in other organisms.  For example, ARK1’s activity is potentially similar to that of the S. cerevisiae kinesin, Kip3p.  A member of the Kinesin-8 family, Kip3p was previously shown to be a plus-end tracking protein as well as a microtubule catastrophe factor (Gupta et al., 2006).  Much like ark1-1 mutants, kip3p mutants have increased polymer mass (ie. longer microtubules) (Cottingham and Hoyt, 1997) and reduced catastrophe frequency (Gupta et al., 2006).  Movement of Kip3p along microtubules is ATP-dependent (Gupta et al., 2006; Varga et al., 2006) and highly processive, which allows the kinesin to maintain plus-end association when depolymerization occurs.  ATP hydrolysis in the motor domain was also shown to physically remove tubulin from the plus-end of the microtubule protofilament (Gupta et al., 2006; 	 31 Varga et al., 2006).  Processivity is a kinesin property that allows the kinesin to move a specific number of steps along microtubule before dissociating from the microtubule.  High processivity means a kinesin would be able to travel long distances before falling off the microtubule. 2.4.2. ARK1 has two domains that enable microtubule localization Given that the ARK1ΔARM-GFP and ARM-GFP constructs (driven by the endogenous promoter) were able to remain localized to microtubule (Figure 2.14, 2.15) suggests that ARK1 has two putative microtubule-binding sites.  While it is no surprise that the ARK1ΔARM-GFP construct remains bound to the microtubule (since the motor domain is present), the truncated ARM-GFP construct staying localized to the microtubule plus ends was unexpected (Figure 2.15).  The ARM-GFP construct that is specifically localized to microtubule plus ends could potentially contribute to ARK1’s ability to plus-end track at the growing plus ends.  However, given that the ARK1ΔARM-GFP seems to plus-end track much like full length ARK1-GFP in vivo (Figure 2.14), the ARM may not be involved in aiding in ARK1’s plus-end tracking abilities.  Whether the ARM domain of ARK1 leads to direct or indirect binding to microtubules remains to be determined.  As previously mentioned, the ARM domain is predicted to interact with the NEK6 (Sakai et al., 2008).   The NEK6 kinase has been shown to localize to microtubules (Motose et al., 2008, 2011) and was predicted to aid in ARK1’s microtubule localization via the ARM domain.  However, ARK1’s localization appears to be independent of NEK6 (see Chapter 3). The idea that a kinesin has two independent microtubule-binding domains is not entirely new.  The A. thaliana ATK5, for example, possesses two microtubule-binding domains: the C-terminal motor domain and an additional N-terminal domain that enables microtubule plus-end tracking (Ambrose et al., 2005).  Members of the kinesin-8 family (S. cerevisiae Kip3p and human Kif18A), also contain secondary microtubule-binding domains that allow these kinesins to remain bound to microtubules, resulting in high processivity and plus-end accumulations (Mayr et al., 2011; Stumpff et al., 2011; Su et al., 2011; Weaver et al., 2011). The second microtubule-binding domain of Kif18A also functions independently of ATP (Mayr et al., 2011; Weaver et al., 2011) much like the Armadillo-repeats, which do not have an ATP catalytic site (Tewari et al., 2010).  Based on my evidence, I propose that the ARM domain functions in increasing ARK1 processivity. Motor protein processivity, 	 32 however, can only be confirmed through in vitro assays with purified ARK1 and microtubules/tubulin. 2.4.3. ARK1 function is redundant in cell types besides root hairs  Based on its widespread expression (Figure 2.9), ARK1 appears to function in cell types other than root hair-forming ones, with its expression pattern overlapping with that of its paralogues, ARK2 and ARK3.  This may explain why the ark1-1 phenotype is only seen in root hairs.  ARK2 is thought to modulate microtubule dynamics in root epidermal cells, evident by its T-DNA insertion mutants having a root twisting phenotype (Sakai et al., 2008) typically associated with aberrant microtubule organization and dynamics (Nakajima et al., 2004; Nakamura et al., 2004; Sedbrook et al., 2004).   ARK3/AtKinUa accumulates at PPBs and is cell cycle-regulated during embryogenesis and stomatal development (Lau et al., 2014; Malcos and Cyr, 2011).  Although ARK1-GFP localized to PPBs as well as mitotic spindles and phragmoplasts, it is unlikely to be cell cycle-regulated.  ARK3 was previously found to contain a D-Box motif, a regulatory sequence found in cyclins and cell cycle-regulated proteins, which is absent in ARK1 (Malcos and Cyr, 2011).  Because artificial microRNA against ARK3 led to meristemoid cells with reduced asymmetrical cell divisions, ARK3 is predicted to regulate asymmetrical cell division by regulating the placement of the PPB (Lau et al., 2014).  I did not see any ark1-1 phenotype related to disrupted cell cycle (ie. abnormal cell plate placement, changes in cell shape/size) (Figure 2.17), suggesting the presence of ARK3 is sufficient and necessary for cells to complete cell division while ARK1 is not. 2.4.4 ARK1 is an important factor for microtubule turnover and re-modelling By promoting periodic microtubule catastrophe, ARK1 can facilitate microtubule turnover, an important feature in array remodeling. In addition, our findings suggest that ARK1’s microtubule destabilizing activity ultimately serves to maintain sufficient levels of free tubulin, which is required to sustain rapid microtubule polymerization. Interestingly, the data obtained in the current and some previous studies suggest that rapid microtubule polymerization, rather than polymer level, is the single parameter of microtubule dynamics most critical for maintenance of polar tip growth in root hairs. Previously it was shown that drugs and mutations that either over-stabilize or destabilize microtubules can have the same 	 33 consequence on cell shape (reviewed in (Wasteneys and Ambrose, 2009).  Taxol and oryzalin treatments, which increase and decrease microtubule polymer mass respectively, both cause crooked and branched root hairs to develop (Bibikova et al., 1999). Crooked and branched root hairs are also found in ark1-1 and mor1-1 mutants, which have increased and decreased microtubule polymer mass respectively (Sakai et al., 2008; Whittington et al., 2001). Despite these extremes in polymer mass, the common feature is that microtubule growth rates are consistently reduced in mor1-1 mutants at restrictive temperature (Whittington et al., 2001), in ark1-1 (this study) and in wild-type cells treated with oryzalin (this study and (Nakamura et al., 2004)). Importantly, the application of low concentrations of oryzalin to ark1-1 both increased microtubule polymerization rates and reduced root hair branching. With this evidence, I conclude that microtubule polymerization rates must exceed a threshold value (approximately 6 µm per minute or 100 nm per second at 21ºC) in order to maintain uniform tip growth. This supports the concept that microtubule dynamics need to be within a very narrow “Goldilocks Zone” for maintenance of microtubule array organization and cellular development (Wasteneys and Ambrose, 2009). My results reveal that ark1-1 root hairs accumulate bundles of endoplasmic microtubules despite the fact that these microtubules have reduced growth velocities and catastrophe frequencies not significantly different from endoplasmic microtubules in wild-type root hairs.  This apparently paradoxical result can be explained by the fact that the endoplasmic microtubules originate in the cortex such that their accumulation may be independent of their dynamics.  By resisting catastrophe, the cortical microtubules in the apex of ark1-1 mutant root hairs eventually become detached from the cortex and, via the reverse-fountain cytoplasmic streaming (Sieberer and Emons, 2000), are swept back into the endoplasm where they accumulate in large bundles.	It has also been postulated that the continuously changing positions and orientation of the endoplasmic microtubules are correlated with the highly dynamic cytoplasmic streaming found in the endoplasm of root hairs (Bruaene et al., 2004).  This may have caused me to overestimate endoplasmic microtubule catastrophe frequencies despite my attempts to ensure I was imaging the entire endoplasmic region with frequent acquisition time points for an extended time period. From my mutant, drug, and overexpression analysis, it is clear that the correct modulation of microtubule dynamics encompasses a broad range of cellular factors, 	 34 including free tubulin concentration, and the expression and activation of different MAPs, including ARK1.  Although it was expected that ARK1-RFP OX would result in an increase in catastrophe frequency (Figure 2.12C), I was surprised to observe a reduced microtubule growth rate (Figure 2.12E).  It is possible that over-accumulation of ARK1-RFP, particularly at the microtubule plus-end, inhibits the binding of other microtubule-associated proteins or affects its own function.   In A. thaliana, several MAPs have been reported to interact with microtubule plus ends to control dynamics (e.g. EB1 (Chan et al., 2003); SKU6/SPIRAL1 (Nakajima et al., 2004; Sedbrook et al., 2004), ATK5 (Ambrose et al., 2005), CLASP1 (Ambrose et al., 2007)), the activity of which might be altered by ARK1-RFP OX.  In vitro analysis has previously shown that increasing concentrations of Kip3p and other MAPs can decrease Kip3p’s velocity along microtubules as a result of macromolecular crowding (Leduc et al., 2012).  Given that the plus-end microtubule-associated proteins interact at the nanoscale level, an appropriate ratio of these proteins is required for precise control of microtubule dynamics.  Based on my study, complete removal of ARK1 from microtubule plus ends or its overexpression is detrimental to proper microtubule dynamics. In this study, I was able to conclude that ARK1 has a role in promoting microtubule depolymerization, which is only the second characterized kinesin to do so within the plant kingdom.  ARK1’s role is important for maintaining microtubule polymer mass and dynamics in order for proper root hair morphogenesis to occur.   	 35  Table 2.1.  List of primer sequences used for cloning the various ARK1 constructs. cds = coding sequence.   Forward Primer Reverse Primer PCR Product 5’-AAA AAA GCA GGC TGG ACG ATG ATC AAG AGA TG-3’ 5’-AGA AAG CTG GGT TAT ATA TCT TTA CAC ATA AGT TGT ACA GTG-3’ Full length ARK1 genomic 5’-AAA AAA GCA GGC TTG AAG AGA TAG AAC CAT GAG TTC GTC AAA TTC CTC C-3’ 5’-AGA AAG CTG GGT AGC TTG AGA AGT AAG GGT TTG TTT TG-3’ Full length ARK1 cds 5’-AAA AAA GCA GGC TTG AAG AGA TAG AAC CAT GAG TTC GTC AAA TTC CTC C-3’ 5’-AGA AAG CTG GGT ACT GAC CAG ATA ACG AC-3’ ARK1ΔARM cds 5’-AAA AAA GCA GGC TTG GAT GGA ATT TGA TTA TGA TTA TGA GAG TTT G-3’ 5’-AGA AAG CTG GGT AGC TTG AGA AGT AAG GGT TTG TTT TG-3’ ARK1ΔMotor cds 5’-AAA AAA GCA GGC TTG AAG AGA TAG AAC CAT GAG AGC TAC CAT GGC-3’ 5’-AGA AAG CTG GGT AGC TTG AGA AGT AAG GGT TTG TTT TG-3’ ARM cds 5’-GAA AAG TTG GGT GAA CAG TGG AAG AAC ACG G-3’ 5’-TTT TTT GTA CAA ACT TGC TAG CTG GAG TCT TCA CGA AG-3’ ARK1 promoter 	 36  Table 2.2. Plus-end growth velocities and catastrophe frequencies of wild-type and ark1-1 microtubules in untreated and 100 nM oryzalin-treated root hairs. CMT = cortical microtubules; EMT = endoplasmic microtubules. Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   	 37   Figure 2.1.  Root hairs of ark1-1 are wavy and branched.  Bright-field images of both wild-type and ark1-1 root hairs.  Scale bars represent 50 µm.   Wild type ark1-1 	 38 Figure 2.2.  Endoplasmic microtubules in ark1-1 are more abundant than in wild-type root hairs.  (A) Endoplasmic bundles (indicated by arrows) are more abundant in ark1-1 elongating root hairs than in wild-type elongating root hairs.  Spinning disc confocal images of the mid-plane of elongating root hair expressing the microtubule marker, 35Spro:GFP-MBD. Scale bars represent 5 µm.  (B) Cross-section of the ark1-1 root hair indicated by the dashed-line in (A).  The white circle divides the cortical and endoplasmic region used for analysis in (C). Scale bar represents 5 µm.  (C) Graph showing an increase in relative fluorescence of endoplasmic microtubule bundles to cortical microtubules in ark1-1 and wild-type root hairs.  The ratio of mean fluorescent intensity of microtubules between the endoplasmic to cortical region were measured (indicated in (B)).  Endoplasmic bundles are highest at the tip and decrease as you move further away.  19 wild-type and 13 ark1-1 root hairs were measured.  Images were analyzed according to (Sakai et al., 2008).  Data and bars represent mean and standard deviation, respectively.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org). 	 39 Figure 2.3.  Cortical microtubule plus-end velocities and catastrophe frequencies are reduced in ark1-1 root hairs but can be rescued by oryzalin treatment.  (A) Plus-end velocities of EB1b-GFP of cortical (CMT) and endoplasmic (EMT) microtubules were measured in wild-type and ark1-1 elongating root hairs upon imaging with spinning-disc confocal microscopy. The mean ark1-1 microtubule plus-end velocities (CMT: 3.5±1.1µm/min, n =111; EMT: 5.7±1.3 µm/min, n = 85) are significantly reduced relative to wild-type microtubules (CMT: 6.5±1.7 µm/min, n = 209; EMT: 7.1±1.9, n = 149). Exposure to 100 nm oryzalin rescued the microtubule plus-end velocities of ark1-1 root hairs (CMT: 6.1±1.6µm/min, n =47; EMT: 6.7±2.0 µm/min, n = 106) so that there was no significant difference from oryzalin-free wild-type root hairs (CMT: 6.5±1.7 µm/min, n =209; EMT: 7.1±1.9 µm/min, n = 149).  Oryzalin treatment significantly reduced wild-type velocities (CMT: 5.6±1.6 µm/min, n =134; EMT: 5.9±1.7 µm/min, n = 76). A minimum of five roots (for untreated) and three roots (for oryzalin-treated) (five root hairs per root) from each genetic background were imaged.  (B) Catastrophe frequencies of cortical and endoplasmic microtubules were measured in the same elongating root hairs as in (A).  Cortical microtubule catastrophe frequencies in ark1-1 (0.020±0.011 events/s) are significantly reduced relative to wild-type microtubules (0.029±0.015 events/s). There is no significant difference for endoplasmic microtubules (WT: 0.048±0.025 events/s; ark1-1: 0.055±0.034 events/s). Exposure to 100 nm oryzalin increased the catastrophe frequency of cortical microtubules in ark1-1 root hairs (0.041±0.03 events/s) relative to untreated wild-type root hairs (0.029±0.015 events/s).  The same microtubules in (A) were used to calculate the catastrophe frequency.  0"2"4"6"8"10"A Mean Velocity of EB1b:GFP (µm minute-1 ) Cortical Microtubules Endoplasmic Microtubules **NS * *NS -20 0 20 40 60 80 100 % Change in Plus-end Velocity wild-type ark1-1 Cortical Microtubules Endoplasmic Microtubules B C 0"0.02"0.04"0.06"0.08"0.1"Catastrophe Frequency (s-1) Cortical Microtubules Endoplasmic Microtubules **NS NS NS NS wild-type wild-type + oryzalin ark1-1 ark1-1 + oryzalin 	 40 Figure 2.3.  Cortical microtubule plus-end velocities and catastrophe frequencies are reduced in ark1-1 root hairs but can be rescued by oryzalin treatment.  (C) Exposure to 100 nM oryzalin reduced microtubule plus-end velocities in wild-type root hairs but increased velocities in ark1-1 root hairs.  Data and bars are represented as mean ± SD, respectively. Asterisk shows a significant reduction while NS = no significant difference in velocity relative to untreated wild-type root hairs using a two-sample t-test with unequal variance (α = 0.01).  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   	 41   Figure 2.4.  EB1b-GFP moves more slowly in ark1-1 root hairs than in wild-type root hairs. Montage of spinning-disc laser confocal time-lapse micrographs of EB1b-GFP in wild-type (top) and ark1-1 (bottom) root hairs showing both cortical and endoplasmic microtubule plus ends.  The coloured lines represent the EB1b-GFP trajectories with wild-type showing faster EB1b-GFP movement than in ark1-1. Images were acquired every 8 seconds and represent the medial plane of the root hair.  Scale bars represent 10 µm.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org). 	 42    Figure 2.5.  Treatment with oryzalin causes no obvious differences in microtubule organization between wild-type and ark1-1 root hairs.  Spinning-disc laser confocal micrographs of root hairs expressing the 35Spro:GFP-MBD microtubule marker in wild-type and ark1-1.  No differences were noticeable between the microtubules of wild-type and ark1-1 root hairs treated with 100 nM oryzalin.  Images are merged Z-slices of the entire root hair stack.  Scale bars represent 10 µm.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   ark1-1; 35Spro:GFP-MBD  Oryzalin  Control WT; 35Spro:GFP-MBD ark1-1; 35Spro:GFP-MBD WT; 35Spro:GFP-MBD 	 43  Figure 2.6.  Oryzalin partially rescues the ark1-1 root hair morphology.  (A) Bright-field light images of wild-type and ark1-1 root hairs treated with either 5 µM of taxol or oryzalin showing that the ark1-1 phenotype is partially rescued with oryzalin but not taxol.  Scale bars represent 50 µm.  0"50"100"150"200"250"300"Oryzalin Control Wild type ark1-1 Taxol Root Hair Length (µm) Oryzalin Concentration * * 1 µM 5 µM 10 µM 0 50 100 150 200 250 300 Control 1 uM  5 uM 10 uM Root Hair Length (µm) Wild type ark1-1 Taxol Concentration µM µM µM Wild type ark1-1 A B Frequency of Phenotypes ark1-1 + Oryzalin Wild type + Oryzalin C  100 nM Control*0%"10%"20%"30%"40%"50%"60%"70%"80%"90%"100%"Straight Wavy/Bulbous Branched 0%"10%"20%"30%"40%"50%"60%"70%"80%"90%"100%"Straight Wavy/Bulbous Branched 5 µM  1 µM  100 nM  Control 10 µM  	 44 Figure 2.6.  Oryzalin partially rescues the ark1-1 root hair morphology.  (B) Mean length of wild-type and ark1-1 root hairs treated with various concentrations of taxol and oryzalin.  ark1-1 exposed to 100 nM, 1 µM, and 5 µM oryzalin partially rescues the root hair length phenotype.  Data and bars represent mean and SD, respectively. Asterisks represent a significant increase in mean length relative to untreated ark1-1 root hairs (α = 0.01).  A minimum of 25 root hairs were measured for each genotype and treatment.  (C) Frequency distribution histograms comparing root hair morphology types in wild type and ark1-1 grown on varying oryzalin concentrations.  1 and 5 µM oryzalin partially mitigates the ark1-1 root hair defect by shifting the distribution of the most severe branched phenotype to a milder wavy/bulbous phenotype.  As a control, wild-type root hairs show an increase loss of root hair polarity upon exposure to increasing oryzalin concentrations.  A minimum of ten roots and 150 root hairs were measured for each treatment.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   	 45  Figure 2.7.  ARK-GFP rescues the ark1-1 root hair phenotype  (A) Root hair lengths of ark1-1; ARK1pro:ARK1-GFP and wild type are not significantly different. Data and bars represent mean and standard deviation, respectively.  A minimum of 110 root hairs were measured.  (B) Differential Interference Contrast images of wild-type and ark1-1; ARK1pro:ARK1-GFP root hairs. Scale bars represent 200 µm.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).    0 100 200 300 400 500 Wild-type ark1-1; ARK1pro:ARK1-GFP Root Hair Length (µm) Wild-type ark1-1; ARK1pro:ARK1-GFP A B 	 46   Figure 2.8.  ARK1-GFP localizes to microtubules in elongating and fully-grown root hairs.  (A) Laser confocal micrographs of ark1-1; ARK1pro:ARK1-GFP elongating root hairs.  ARK1-GFP (indicated in red) localizes to microtubules (mCherry-MAP4MBD indicated in cyan).  The cortex and the midplane of the root hairs are both shown.  Arrowheads point to ARK1-GFP localization on microtubules.  The right of the figure shows the magnified representation of the white-boxed area in the merged image.  Scale bar represents 20 µm.  (B) Laser confocal micrographs of ark1-1; ARK1pro:ARK1-GFP fully grown root hairs.  ARK1-GFP (indicated in red) is still expressed in fully-grown root hairs and localizes to microtubules (indicated in cyan).  The cortex and the midplane of the root hairs are both shown.  Arrowheads point to ARK1-GFP localization on microtubules.  The right of the figure shows the magnified representation of the white-boxed area in the merged image. Scale bar represents 20 µm.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   Cortex Midplane ARK1-GFP mcherry-MAP4MBD mcherry-MAP4MBD Merge Merge ARK1-GFP A ARK1-GFP ARK1-GFP Cortex Midplane mcherry-MAP4MBD mcherry-MAP4MBD Merge Merge B >">">">">">">">"	 47  Figure 2.9.  ARK1-GFP is expressed in non-root hair cells and labels different microtubule structures. ARK1-GFP is expressed in epidermal cells of the root, hypocotyl, cotyledon, petiole, and root tip.  ARK1-GFP was co-expressed in ark1-1 35Spro:mCherry-MAP4MBD plants. GC = guard cell; PC = pavement cell; MS= mitotic spindle; PPB = pre-prophase band; P = phragmoplast. Scale bars represent 10 µm. Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   Root Epidermal Hypocotyl Epidermal Cotyledon Epidermal GC PC Petiole Epidermal ARK1-GFP mCherry- MAP4MBD Merge Root Tip Epidermal MS P PPB 	 48  Figure 2.10.  ARK1-GFP accumulates on growing microtubule plus ends.  (A) A time-lapse montage showing ARK1-GFP accumulating on microtubule plus ends (mCherry-MAP4MBD) in cotyledon epidermal cells.  Arrowheads indicate the growing microtubule plus-end.  Scale bar represents 5 µm. Time for each frame is indicated.  (B) A kymograph of the yellow line scan in the last panel (A) showing ARK1-GFP moving along growing microtubules.  (C) Graph showing that ARK1-GFP accumulates at the plus-end of microtubules but still remains bound to the sidewall of microtubules.  Lines represent the ratio between fluorescence intensity of one point to the mean fluorescence intensity of one line-scan.  The ratio was highest within 1 µm of the microtubule plus-end but decreased further from the plus-end. As a control, the same ratio was calculated for microtubules with mCherry-MAP4MBD. 46 line-scans were used to measure ARK1-GFP and 26 line-scans were used to measure mCherry-MAP4MBD.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).    ARK1-GFP mCherry-MAP4MBD Merge >" >" >">" >" >">"0 s 8 s 24 s 16 s 32 s 40 s 48 s 0.5 1 1.5 0 1 2 3 Ratio of  Fluorescence Intensity: Mean Fluorescence Intensity Distance from MT Plus-End (µm) mCherry-MAP4MBD ARK1-GFP time ARK1-GFP mCherry-MAP4MBD Merge B A C >">">">">">" >">">">">">">">"	 49  Figure 2.11.  ARK1-GFP is not found on microtubule minus ends or depolymerizing plus ends.  (A) A time-lapse montage showing that ARK1-GFP localizes specifically to plus ends but not minus ends. Arrowheads indicate a growing microtubule plus-end. Arrows label the microtubule minus end that is depolymerizing. Scale bars represent 5 µm.  (B) A time-lapse montage showing that ARK1-GFP does not localize to shrinking microtubule plus ends.  The microtubule plus-end grows (indicated by the arrowhead) and eventually depolymerizes (indicated by arrow).  ARK1-GFP disappears upon microtubule depolymerization. Scale bars represent 5 µm.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).  ARK1-GFP mCherry-MAP4MBD Merge >">">">">">">">">">">">">"ARK1-GFP mCherry- MAP4MBD Merge >">">">">">">">"A 0 s 8 s 16 s 24 s 32 s 40 s 0 s 16 s 8 s 24 s 32 s 48 s 40 s 56 s 64 s 72 s 80 s 88 s 96 s 104 s B 	 50  Figure 2.12.  ARK1-RFP overexpression leads to increased microtubule catastrophe frequencies.  (A) Diagram of UBIQUITIN10 promoter translational fusion construct used for transient expression in Arabidopsis thaliana cotyledons.  ARM = Armadillo-repeats  (B) Confocal laser micrographs of the UBQ10pro:ARK1-RFP OX construct in ark1-1; 35Spro:GFP-MBD plants. Negative control (UBQ10pro:RFP) shows cytoplasmic RFP. Scale bar represents 10 µm.  (C) Cells transiently overexpressing ARK1-RFP have a significant increase in microtubule catastrophe frequency (0.034±0.032 events/s) relative to cells not expressing ARK1-RFP (0.019±0.011 events/s). For ARK1-RFP OX and the negative control, 26 and 42 microtubules were measured over a time period of 1700 seconds and 2500 seconds, respectively.  (D) Distribution of time spent in each phase (growth, shrinkage, and pause) for cells overexpressing and not overexpressing ARK1-RFP in ark1-1 35Spro:GFP-MBD plants.  Microtubules in cells overexpressing ARK1-RFP spent a greater amount of time shrinking as a result of increased catastrophe frequency shown in (C). For ARK1-RFP OX and the negative control, 86 and 82 microtubules were visualized over a time period of 8000 seconds and 5800 seconds, respectively.   Motor Coiled-coil ARM-ARM-ARM N- -RFP UBQ10pro:ARK1-RFP ARK1-RFP GFP-MBD Merge AB0 2 4 6 8 10 12 14 Growth Shrinkage 0% 10% 20% 30% 40% 50% 60% 70% 80% 90% Growth Shrinkage Pause Velocity (µm minute-1 ) Time Spent per Phase 0 0.01 0.02 0.03 0.04 0.05 0.06 0.07 0.08 Negative Control ARK1:RFP OX Catastrophe Frequency (events second-1) **Negative Control ARK1:RFP OX Negative Control ARK1:RFP OX C D E-RFP (neg control) 	 51 Figure 2-12.  ARK1-RFP overexpression leads to increased microtubule catastrophe frequencies.  (E) The mean microtubule growth and shrinkage velocities in cells overexpressing and not overexpressing ARK1-RFP.  There is a significant reduction in growth velocities in ARK1-RFP overexpression cells (3.2±0.1.6 µm/min) relative to the negative control (3.6±1.8 µm/min).  There is no significant difference in shrinkage velocities between the two treatments (ARK1-RFP= 7.4±4.1 µm/min; negative control= 7.3±4.2 µm/min). For each treatment, a minimum of 700 growth events and 150 shrinkage events were measured.  Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).    	 52  Figure 2.13.  The ARM domain aids in ARK1 localization to microtubule. (A) Diagram of all the truncated UBIQUITIN10 promoter translational fusion constructs used for transient expression in Arabidopsis thaliana cotyledons. (B) Confocal laser micrographs of the various ARK1 OX constructs transiently expressed in ark1-1 35Spro:GFP-MBD plants.  All truncated ARK1 proteins remain localized to microtubules even with just the ARM domain.  Scale bars represent 10 µm. Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).    Coiled-coil ARM-ARM-ARM N- -RFP Motor Coiled-coil N- -RFP ARM-ARM-ARM N- -RFP UBQ10pro:ARK1ΔARM-RFP UBQ10pro:ARK1ΔMotor-RFP UBQ10pro:ARM-RFP ARK1ΔARM-RFP ARK1ΔMotor-RFP GFP-MBD GFP-MBD Merge Merge ARM-RFP GFP-MBD Merge AB	 53  Figure 2.14.  ARKpro:ARK1ΔARM-GFP is sufficient to rescue the ark1-1 root-hair phenotype and localizes to microtubules in a similar pattern as full length ARK1.   (A) Bright-field image of the ark1-1 root hair phenotype being rescued by the ARK1pro:ARK1ΔARM-GFP construct. Root hairs are straight with normal polarity (similar to wild-type root hairs).  Scale bar represents 100 µm.  (B) Variable-angle TIRF micrographs of ARK1-GFP (in red) localizing to microtubule plus ends in etiolated hypocotyls.  Microtubules are labelled with mCherry-MAP4MBD (in cyan).  ARK1-GFP and microtubule micrographs are merged in the right figure. Arrowheads point to ARK1-GFP particles. Scale bars represent 10 µm. (C) Variable-angle TIRF micrographs of ARK1ΔARM-GFP (in red) showing similar microtubule plus-end localization as the full length ARK1-GFP construct in etiolated hypocotyls. Arrowheads point to ARK1-GFP particles that localize to plus ends.  Scale bars represent 10 µm.    A ark1-1; ARK1Δ ARM-GFP ARK1-GFP mCherry-MAP4MBD Merge ARK1Δ ARM-GFP mCherry-MAP4MBD Merge B C > > > > > > > > > > > > > > > > > > Wild type 	 54  Figure 2.15.  ARK1pro:ARM-GFP shows plus-end tracking abilities on microtubules but is not able to rescue the ark1-1 root hair phenotype. (A) Spinning disc confocal micrographs of ARM-GFP (in red) as bright, punctae localizing to growing microtubule plus ends (seen in the merged image) in light-grown hypocotyls.  ARM-GFP expression is patchy and generally expressed at low levels in cells.  The asterisk represents a neighbouring cell that is not expressing ARM-GFP but is expressing the mCherry-MAP4MBD microtubule marker (in cyan).  Arrowheads point to ARM-GFP particles.  Scale bar represents 10 µm. (B) Spinning disc confocal micrographs of ARM-GFP as bright, punctae similar to the EB1b-GFP marker in light-grown hypocotyls.  Scale bar represents 10 µm.  A B C D E Merge ARM-GFP *"*"*"ARM-GFP ark1-1; ARM-GFP ARM-GFP ARM-GFP mCherry-MAP4MBD > > > > > > > > > > > > > > > 	 55 Figure 2.15.  ARK1pro:ARM-GFP shows plus-end tracking abilities on microtubules but is not able to rescue the ark1-1 root hair phenotype. (C) Spinning disc confocal micrographs showing ARM-GFP trajectories on growing microtubule plus ends in light-grown hypocotyls.  The image represents a merged stack of time-lapse images acquired every 8 seconds for 3 minutes.  Scale bar represents 10 µm.  (D) Spinning disc confocal micrographs showing ARM-GFP as bright, punctae (indicated by arrowheads) as well as fluorescence accumulating in tubules, possibly part of the endoplasmic reticulum (indicated by arrows).  Scale bar represents 10 µm. (E) Bright-field image of the ark1-1 root hair phenotype remaining unaffected by expression of the ARK1pro:ARM-GFP construct. Root hairs remained crooked, bulbous, and branched.  Scale bar represents 100 µm	 56  Figure 2.16.  The plus-end velocities and catastrophe frequencies in cells expressing ARK1-GFP and ARK1ΔARM-GFP are not significantly different from each other. (A) The growth velocities of microtubules in etiolated hypocotyls expressing ARK1-GFP (5.6±0.9 µm/minute; n = 199 particles measured) or ARK1ΔARM-GFP (5.9±0.8 µm/minute; n = 70 particles measured) were not significantly different from each other.  Constructs are in the ark1-1 35Spro:mCherry-MAP4MBD transgenic lines. (B) The catastrophe frequencies of microtubules in plants expressing ARK1-GFP (0.019±0.009 events/second; n = 199 particles measured) and ARK1ΔARM-GFP (0.016±0.008 events/second; n = 70 particles) were not significantly different.  The same particles in (A) were used for quantifying catastrophe frequencies. Data and bars are represented as mean ± SD, respectively. Two-sample t-tests with unequal variances were measured using α = 0.01. 0 1 2 3 4 5 6 7 0 0.005 0.01 0.015 0.02 0.025 0.03 0.035 Catastrophe Frequency (events/s) Velocity (µm/min) ARK1-GFP ARK1-GFP ARK1Δ ARM-GFP ARK1Δ ARM-GFP 	 57  Figure 2.17.  Cell and tissue patterns are not affected in ark1-1 root tips. Confocal images of propidium iodide-stained root tips of the lateral root cap (left) and epidermal cells (right).  The size of cells and cell plate placement are not altered in ark1-1 (bottom) relative to wild type (top).  Scale bars represent 30 µm. Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).   	 58  Figure 2.18.  ark1-1 microtubule phenotype is restricted to root hair-forming cells (A) Microtubule plus-end growth velocities are not significantly different in the pavement cells of wild-type (3.7±0.6 µm/min, n = 409) and ark1-1 (3.7 ±0.6 µm/min, n = 153) cotyledons. Four wild-type and ark1-1 plants expressing the 35Spro:EB1b-GFP marker each were imaged.  (B) Microtubule catastrophe frequencies in ark1-1 (0.020±0.013 events/s) and wild-type cotyledon pavement cells (0.020±0.011 events/s) are not significantly different.  Catastrophe frequencies of microtubules were measured in the same plants and cells as in (A). (C) Microtubule growth and shrinkage velocities in wild-type (growth: 5.0±2.8 µm/min, n = 371; shrinkage: 15.3±9.2 µm/min, n = 37) and ark1-1 (growth: 5.3±2.8 µm/min, n = 286; shrinkage: 13.6±6.3 µm/min, n = 34) elongating hypocotyls are not significantly different from each other. Two (wild-type) and seven (ark1-1) plants expressing the 35Spro:GFP-MBD marker were imaged.  Data and bars are represented as mean ± SD, respectively.  Two-sample t-tests with unequal variances were measured using α = 0.01. Reproduced with permission from Eng and Wasteneys (2014); Copyright American Society of Plant Biologists (www.plantcell.org).  Mean EB1b-GFP Plus-end Velocities in Cotyledons (µm minute-1 ) Wild-type ark1-1 0"0.005"0.01"0.015"0.02"0.025"0.03"0.035"Catastrophe Frequency in Cotyledons (events s-1 ) Wild-type ark1-1 0"0.5"1"1.5"2"2.5"3"3.5"4"4.5"5"A B 0"5"10"15"20"25"Growth" Shrinkage"Wild-type ark1-1 Mean Velocities in Elongating Hypocotyls(µm minute-1 ) C 	 59 Chapter 3: Elucidating the relationship between NEK6 and ARK1 in microtubule dynamics and cell expansion 3.1. Background Information  The Never In Mitosis A (NIMA)-related Kinases (NEKs) belong to a highly conserved group of Serine/Threonine kinases found in a variety of eukaryotic organisms and function in processes related to the cell cycle and the regulation of microtubule dynamics.  The first NEK was discovered in Aspergillus nidulans (a filamentous fungi) and was noted to regulate the transition from the G2 to the M-phase of the cell cycle.  NEK mutants were found to be never in mitosis, thus contributing to the kinase’s name.  A. nidulans’ NEK was thought to be important for phosphorylating/activating downstream targets that are responsible for cells entering mitosis, nuclear envelope break down, and spindle organization (Osmani et al., 1991, 1988).  More evidence has shown that A. nidulans’ NEK phosphorylates histone H3 in order for chromatin condensation to occur (De Souza et al., 2000).  While it appears that NEKs in fungi and mammals are specifically involved in regulating mitotic events (e.g. mitotic initiation, centrosome duplication and separation, spindle formation, cytokinesis) (O’Connell et al., 2003), the NEKs in plants appear to be primarily involved in regulating cell growth and microtubule dynamics.  The exact functions of plant NEKs, however, are less clear relative to the studies of fungal and animal systems.  The unicellular ciliated green alga, Chlamydomonas reinhardtii, has two characterized NEKs (Fa2p and Cnk2p) that are responsible for regulating the length of the organism’s flagellum, a microtubule-containing structure.  Fa2p has been specifically shown to induce microtubule severing within the flagella as well as acting as a cell-cycle regulator (Finst et al., 2000; Mahjoub et al., 2002, 2004).  The second Nek found in Chlamydomonas, Cnk2p, has been shown to regulate the flagellar length and the size of the unicellular organism (Bradley and Quarmby, 2005).  A larger number of NEKs are also represented in higher plants such as in A. thaliana (seven NEKs), Oryza sativa (six NEKs), and Populus trichocarpa (nine NEKs).  In these organisms, NEKs are predicted to regulate plant development, such as organ development and vascularization (Cloutier et al., 2005; Vigneault et al., 2007). 	 60 The NEK6 kinase is one the most studied NEKs in A. thaliana and is one of the seven NEKs found in the Arabidopsis genome.  NEK6 is known to function as a dimer: as a homodimer with itself or as heterodimers with the closely related NEK4 and NEK5.  Knocking out the function of any one of the three kinases leads to loss of cell elongation resulting in ectopic epidermal cell outgrowths (Motose et al., 2011). The family of NEK proteins consists of a highly conserved N-terminal Ser/Thr kinase domain followed by a PEST sequence that is predicted to control NEK regulation and activity (since PEST motifs are generally involved in protein degradation) (Rogers et al., 1986). The C-terminal end of the peptide contains a plant NEK C-terminal (PNC) motif that is the putative site for hetero or homo-dimerization of NEK6 with NEK4 and NEK5 (Motose et al., 2011). Because various characterized nek6 mutant alleles showed ectopic epidermal protrusions in hypocotyl and petiole cells (of non-stomatal lineages), NEK6 is thought to play in a role in cell elongation, specifically in suppressing ectopic outgrowths of epidermal cells.  Outside of hypocotyls and petioles, the nek6-2 allele was also shown to have a larger proportion of two-branched trichomes (versus the wild-type trichomes that typically have three or more branches) (Motose et al., 2011).   However, another nek6 mutant allele, ibo1-2, showed an increase in trichome branching (Motose et al., 2012), suggesting nek6 allele-specific phenotypes.  nek6 mutants were also shown to have disorganized cell files in the root epidermis, with atrichoblast cells ectopically growing root hairs.  Moreover, nek6 roots were shown to display a leftward skew when growing vertically on agar (Sakai et al., 2008).  Another allele, nek6-1/ibo1-4, showed irregular cell files in the roots and hypocotyls as well as aberrant cell plates (Motose et al., 2012).  Since NEK6 is expressed in all cell types, it is hypothesized that NEK6 plays a role in microtubule dynamics in varying cell types and throughout different stages of the cell cycle.   Several observations and experimental evidence support the idea that NEK6 helps modulate microtubule dynamics.  First, the aforementioned nek6 morphological phenotypes are consistent with microtubule-related defects.  NEK6 has also been shown to bind to microtubules, both growing and shrinking as well as both minus and plus ends (Motose et al., 2011).  In addition, nek6 mutants expressing microtubule marker lines revealed less dynamic (i.e. more stable) and disorganized microtubules in the protruded hypocotyl epidermal cells.  Specifically, nek6 microtubules spent a greater than normal amount of time in the pause state 	 61 and deviated from the transverse and oblique microtubule patterns typical for diffusely elongating hypocotyl epidermal cells (Motose et al., 2008, 2011).  Furthermore, microtubules in nek6 mutants were frequently seen in whorled arrays and bundles (Motose et al., 2012).  NEK6 could function in regulating microtubule dynamics by phosphorylating tubulin as evidenced by the demonstration of NEK6’s ability to phosphorylate the β-tubulin subunits, TUB4 and TUB6, in vitro (Motose et al., 2011).  Another member of the NEK family in A. thaliana, NEK2, is also predicted to function in regulating microtubule dynamics and plant growth.  NEK2 RNAi plants showed aberrant microtubule organization as well as delayed plant development and reduced plant size (Agueci et al., 2012). Besides its effects on microtubule dynamics, NEK6 might act as regulator for cell cycle progression (much like in fungi and animals).  In fungi and animals, NEK proteins regulate the cell cycle by activation and localization of Cyclin B, a protein that is involved in cell cycle progression (O’Regan et al., 2007). Similarly, one study overexpressing the A. thaliana NEK6 reported increased expression levels of the Cyclin B genes, CYCB1;1 and CYCA3;1.  Conversely, the expression levels of the same two Cyclin B genes were significantly reduced in the nek6 mutants.  Plants overexpressing NEK6 were also found to have increases in the number of lateral roots, rosette (leaf and petiole) size, and silique length as a result of increased mitotic events driven by increased levels of CYCB1;1 and CYCA3;1 (Zhang et al., 2011). NEK6 is predicted to interact with ARK1 by binding to the latter protein’s ARM-repeat domain.  This interaction was discovered through a yeast two-hybrid screen of potential ARK1 interactors (Sakai et al., 2008).  The ARK1-NEK6 physical interaction was further confirmed through an in vitro pull-down using ARK1 and NEK6 protein fragments created through heterologous gene expression (Sakai et al., 2008).  Another in vitro assay showed NEK6’s ability to phosphorylate ARK1 (H. Motose, personal communication).  Both proteins’ in vivo physical and phosphorylation interaction has yet to be confirmed.  However, previous studies characterizing kinesins in A. thaliana have already shown that kinesin activity can be regulated through phosphorylation.  The kinesins, KCA1 and KCA2, are known to have multiple functions such as light-dependent chloroplast movement (Suetsugu et al., 2010) and dictating the position of the cell plate formation in dividing cells (Vanstraelen et al., 2006).  In terms of regulation through phosphorylation, KCA1/2 was 	 62 found to interact in vivo with a cyclin-dependent protein kinase, CDKA;1, where the kinase phosphorylates the kinesins’ tail-domain, enabling the proper dimerization or conformation of the kinesins for functional activation (Vanstraelen et al., 2004). The major biological question in this chapter is to elucidate the relationship between NEK6 and ARK1.  Specifically, it is unknown if either of these proteins depend on each other for microtubule localization and/or function.  In this chapter, I determined that ARK1 does not require NEK6 for localization and function.  Similarly, NEK6 does not require ARK1 for its localization to microtubules or its function. These findings led to my conclusion that NEK6’s regulation of microtubule dynamics in elongating epidermal cells of the hypocotyl and petiole is independent of its interaction with ARK1.  Finally, I demonstrate that NEK6 expression is potentially regulated by ARK1.  3.2. Materials and Methods 3.2.1.  Generation of plant material and culture A. thaliana ibo1-1; NEK6pro:NEK6-GFP lines were provided by Dr. Hiroyasu Motose (The University of Okayama, Japan) and crossed into the ark1-1 line (see Chapter 2) and F3 segregants were used for further experiments.  Similarly, the ark1-1; ARK1pro:ARK1-GFP lines used in Chapter 2 were crossed into the nek6-1/ibo1-4 (SALK_152782) and F3 segregants were used for further experiments.  For stable transgenic lines, the constructs of ARK1pro:ARK1-TagRFP and UBQ10pro:ARM-RFP were first transformed into Agrobacterium tumefaciens (GV3101 strain) and then transformed into either ark1-1 ibo1-1 NEK6pro:NEK6-GFP or ark1-1; 35Spro:EB1b-GFP lines (used in Chapter 2) using the floral dip method (Clough and Bent, 1998). T3 lines homozygous for each TagRFP/GFP construct were segregated and used for further experiments. All seeds were sterilized in 70% ethanol, rinsed three times with ddH2O, and plated onto Petri dishes with Hoagland’s medium (1.2% Bacto-agar (BD Diagnostics), no sucrose).  For hormone treatments, ACC (Sigma), GA (Sigma), or DMSO (Fisher Scientific) was added to the Hoagland’s media to the desired final concentrations.  Plates with seeds were stored in the dark at 4°C for 2-3 days and transferred to a 21°C growth cabinet (24 h light) where they were grown vertically until imaging.  For untreated samples, 5 to 7 day-old seedlings were used for imaging.  For hormone treated samples, seedlings were grown on 	 63 normal Hoagland’s media for 4 days before being transferred to the plates with hormones for two days prior to imaging.   3.2.2.  ARK1 construct design and cloning strategy Gateway Cloning Technology (Invitrogen) was used for the ARK1 (At3g54870) genomic and coding sequences.  To generate the ARK1pro:ARK1-TagRFP construct, the ARK1 genomic sequence (between 888 base pairs upstream of the ATG/start codon and the TGA/stop codon) was amplified from the F28P10 BAC (from the Arabidopsis Biological Resource Center, Ohio State University) using the full length ARK1 genomic sequence primer set (see Table 2.1 for sequences). A second PCR with the attB-adapter primers was performed according to the manufacturer’s protocol (see Invitrogen for attB-adapter sequences). Following the BP reaction with the various attB-PCR products and the pDONR221 vector (Invitrogen), a LR reaction was performed with the pGWB659 vector (Nakamura et al., 2010b).  The construct was sequenced prior to transformation. The same UBQ10pro:ARM-RFP construct used in Chapter 2 was used for stable transgenic lines in this chapter. 3.2.3.  Live-cell imaging Live imaging of the various fluorescent reporter lines was done using a Perkin Elmer Ultraview VoX Spinning Disc Confocal system (Perkin-Elmer) mounted on a Leica DMI6000 B inverted microscope and equipped with a Hamamatsu 9100-02 electron multiplier CCD camera (Hamamatsu). An argon 488 nm laser line with a complementary GFP (525/36) emission band-pass filter or a 561 nm laser with a complementary RFP (595/50) emission band-pass filter was used.  Images were acquired with a 63x (water) objective lens (NA = 1.2) with 0.3 – 0.5 µm optical z-slices. Seedlings were mounted in either water or the hormone solutions during imaging. Imaging of the hypocotyl and petiole epidermal cells utilized a Leica M216FA Stereomicroscope (Leica Microsystems) equipped with a DFC 350 FX R2 camera (Leica Microsystems).  For the light- and dark-growth experiments, 10-day old seedlings were grown in either light or complete dark at 21°C.   	 64 3.2.4.  RT-PCR  20 seven day-old seedlings of Col-0, ark1-1 ibo1-1 NEK6pro:NEK6-GFP, and ark1-1 ibo1-1 NEK6pro:NEK6-GFP UBQ10pro:ARM-RFP were grown on Hoagland’s media (no sucrose) under continuous light conditions.  Whole seedlings were removed from the media and flash frozen in liquid nitrogen. Seedlings were ground with a mortar and pestle and total RNA was extracted as described (Galway et al., 2011).  cDNA was synthesized using 500 ng total RNA, Oligo d(T)18 (Integrated DNA Technologies), and SuperScript® III Reverse Transcriptase (Life Technologies).  A PCR was done using GFP primers (Forward: GAC GTA AAC GGC CAC AAG T; Reverse: GTA GCG GCT GAA GCA CTG) as well as F-actin primers for the control (Forward: TTT CAA ACC TGC TCC TCC TT; Reverse: GAG ACA TCG TTT CCA TGA CG).  30 cycles of annealing and extension steps were done.  Equal volumes of the PCR reaction were run on a 2% agarose gel with EtBr.  Three biological replicates were performed.  3.3. Results 3.3.1. NEK-GFP and ARK1-TagRFP co-localize to microtubules  Previously, NEK6 was shown to have various microtubule localization patterns.  Transient expression of 35Spro:NEK6-GFP in N. benthamiana revealed an uneven GFP distribution with bright GFP spots along the length of the microtubule.  Moreover, NEK6-GFP co-localized with NEDD1-RFP, a component of γ-tubulin (Motose et al., 2008).  γ-tubulin is part of the microtubule nucleating complex that in plant cells has been shown to bind to pre-existing microtubules and act as nucleation sites for new microtubules.  Later studies using stable expression of NEK6pro:NEK6-GFP in A. thaliana characterized NEK6-GFP dynamics, revealing its association with both growing and shrinking microtubules plus ends (Motose et al., 2011).  Based on NEK6’s microtubule localization, I wanted to determine if NEK6 localizes to growing microtubule plus ends where ARK1 was previously shown to accumulate and if this co-localization is constitutive.  More specifically, I wanted to test whether ARK1 was responsible for recruiting NEK6 to the microtubule plus ends.  In order to accomplish this, I created an ARK1-TagRFP construct driven by the ARK1 promoter, transformed the construct into an ibo1-1 ark1-1; NEK6pro:NEK6-GFP transgenic line, and collected live-cell images of both proteins using spinning-disc confocal microscopy.  ibo1-1 	 65 is a nek6 mutant allele with a point mutation (an Arg substitution at Glu177 in the kinase domain) that results in complete loss of kinase function (Motose et al., 2008).  While the ARK1pro:ARK1-TagRFP construct proved functional as evident by the rescued ark1-1 root hair phenotype (Figure 3.1A), its microtubule distribution pattern was different from the ARK1-GFP plus-end localization described in Chapter 2.  Rather than having an asymmetrical distribution on microtubule plus ends as with the ARK1-GFP construct, the ARK1-TagRFP construct labelled the entire length of the microtubule (observed in root and hypocotyl epidermal cells) (Figure 3.1B, C).  Moreover, ARK1-TagRFP remained bound to microtubules in both rescue and catastrophe events, whereas ARK1-GFP labelling disappeared upon microtubule catastrophe.   3.3.2.  The TagRFP alters ARK1 microtubule localization but not function Due to these differences in microtubule labelling between the ARK1-GFP (Chapter 2) and ARK1-TagRFP construct, I hypothesized that there was an abnormal interaction between NEK6 and ARK1 from the GFP and TagRFP C-terminal tags, respectively, which could have resulted in altered ARK1-TagRFP microtubule localization.  In order to test this hypothesis, I transformed the same ARK1pro:ARK1-TagRFP construct into an ark1-1; 35Spro:EB1b-GFP line to see if the ARK1-TagRFP microtubule labeling pattern persisted.  Once again, this construct rescued the ark1-1 phenotype (Figure 3.2A) and labelled the entire length of the microtubule (Figure 3.2B).  This suggested that the aberrant ARK1-TagRFP labeling possibly resulted from the TagRFP itself rather than an abnormal interaction with NEK6-GFP.  This aberrant microtubule localization, however, did not disrupt ARK1 function as the construct rescued the ark1-1 root hair phenotype (Figure 3.2A).  Given that the labelling pattern of ARK1-TagRFP and NEK6-GFP seen in section 3.2.1 indicated overlapping but not identical distributions, these results suggest that NEK6 and ARK1 associate with microtubules independently. 3.3.3.  The number of nek6-1 epidermal protrusions are not significantly different from nek6-1 ark1-1 double mutants  Since NEK6 and ARK1 are predicted to interact, I wanted to confirm if both proteins function within the same pathway through double mutant analysis.  While it is already known that the nek6 and ark1 mutants have non-overlapping phenotypes (Sakai et al., 2008), 	 66 I wanted to determine if ark1 mutants would enhance the nek6 epidermal protrusion phenotype in hypocotyls and petioles by creating nek6-1 ark1-1 double mutants.  The nek6-1 T-DNA insertion mutants previously showed a complete knock out of NEK6 expression (Sakai et al., 2008).  In light-grown hypocotyls, ark1-1 did not enhance or suppress the nek6-1 phenotype (Figure 3.3A-B), as the number of ectopic epidermal protrusions was not significantly different between the ark1-1 nek6-1 double mutants (7.3±5.6 protrusions) and the nek6-1 single mutants (8.0±4.0 protrusions) (Figure 3.3D).  The severity of nek6-1 epidermal protrusions was previously reported to be reduced in dark-grown (etiolated) hypocotyls (Motose et al., 2008) and I noted the same effect.  Similarly, etiolation suppressed the number of ectopic protrusions in nek6-1 ark1-1 to the same extent as in nek6-1 (Figure 3.3C, D). Epidermal protrusions were completely absent in both light- and dark-grown wild-type and ark1-1 plants (Figure 3.3A, C). These results suggest that NEK6 functions independently of ARK1 in the role it plays in elongation of hypocotyl epidermal cells and possibly in the regulation of microtubule dynamics/organization. 3.3.4. NEK6-GFP localization does not change in the ark1-1 background  Based on previous data that showed NEK6 interacting with the ARM domain of ARK1, I hypothesized that NEK6’s microtubule association is dependent on ARK1 and that NEK6-GFP should mis-localize in the ark1-1 background. To visualize NEK6-GFP in the ark1-1 background, I used spinning-disc confocal microscopy. Upon analysis of root hairs, root epidermal, and hypocotyl epidermal cells, NEK6-GFP distribution was no different in the wild-type and ark1-1 backgrounds (Figure 3.4).  In both genotypes, NEK6-GFP had uneven and punctate distribution along microtubules.  NEK6-GFP displayed normal dynamics in ark1-1 in that NEK6-GFP remained associated with shrinking and growing microtubules similar to NEK6-GFP in the nek6 complemented background as previously reported in Motose et al. (2008, 2011).  These observations suggest that NEK6 localization is independent of ARK1. 3.3.5.  ARK1-GFP distribution and function does not change in the nek6 mutant  To test if ARK1 microtubule distribution depends on the presence of NEK6, I localized ARK1-GFP in the nek6-1 mutant co-expressing a mCherry-MBD microtubule reporter.  It was previously shown through RT-PCR that the nek6-1 T-DNA insertion lines 	 67 were true knock-outs with null transcript (Sakai et al., 2008).  In root hairs, ARK1-GFP continued to associate with microtubules in nek6-1 (Figure 3.5A).  Moreover, plants expressing ARK1-GFP in the nek6-1 ark1-1 double mutant maintained straight root hairs, which suggests that ARK1-GFP is still able to complement the ark1-1 phenotype despite the complete absence of NEK6 (Figure 3.5B).  Because nek6-1 mutants have epidermal protrusions in hypocotyls and petioles, I then decided to look at these cells types to see if ARK1-GFP localization is altered in these cells and contributes to the protrusion phenotype.  However, ARK1-GFP appeared to remain associated with microtubule plus ends in the epidermal protrusions (Figure 3.5C).  These results suggest that ARK1 distribution is independent of NEK6 and does not require phosphorylation by NEK6 for function and/or microtubule association. 3.3.6.  Exogenous Gibberellic Acid and ACC treatments do not affect the NEK6 and ARK1 interaction  Because I saw no differences in ARK1-GFP and NEK6-GFP localization in the nek6-1 and ark1-1 mutant backgrounds, respectively, I concluded that ARK1 and NEK6 do not interact with each other under normal conditions.   However, previous results have shown changes in the number of epidermal protrusions upon plant hormone treatments with either Gibberellic Acid (GA) or the ethylene precursor, ACC, in the nek6 mutant background. Under normal conditions, hypocotyl cell elongation is driven by etiolation, which involves enhanced GA signalling.  Moreover, much like the effect of etiolation on nek6 hypocotyls, GA was also shown to inhibit the ibo1/nek6 ectopic protrusions.  ACC, however, was shown to do the opposite of GA and increase the number of protrusions in the nek6 background (Motose et al., 2008). Normally, ethylene promotes radial growth, particularly in inducing ectopic root hair formation and epidermal protrusions (Masucci and Schiefelbein, 1996; Pitts et al., 1998; Tanimoto et al., 1995).  Given that the nek6 mutants showed an increase in ectopic protrusions with ACC, NEK6 is predicted to act antagonistically to ethylene to inhibit outgrowths (Motose et al., 2008).  The relationship with ethylene/ACC and NEK6 is predicted to function as a negative feedback loop.  ACC was shown to induce NEK6 expression while NEK6 overexpression led to reductions in the expression levels of ethylene-related genes and ethylene biosynthesis (Zhang et al., 2011).  Exposure to ACC and 	 68 GA had no effect on wild-type hypocotyl and petioles as epidermal protrusions remained non-existent in these cell types (Motose et al., 2008). Based on these observations, I hypothesized that perhaps the interaction of ARK1 and NEK6 is conditional (i.e. dependent on specific hormonal conditions).  In order to test this hypothesis, I decided to visualize ARK1-TagRFP and NEK6-GFP using confocal microscopy upon exposure to ACC (50 µM) and GA (10 µM).  In ibo1-1 ark1-1 plants complemented by expressing ARK1-TagRFP and NEK6-GFP, treatments with ACC and GA did not alter both proteins’ localization pattern to untreated plants (Figure 3.6).  In untreated (mock), GA, and ACC seedlings, ARK1-TagRFP and NEK6-GFP distribution on microtubules remained the same.  This observation is consistent with the lack of epidermal protrusions in wild-type seedlings exposed either to GA or ACC (Motose et al., 2008), which suggests that neither GA nor ACC alter the NEK6-ARK1 interaction or either protein’s association with microtubules. I then hypothesized that NEK6 or ARK1 function/localization may be altered in either the ark1 or nek6 background, respectively, when exposed to exogenous GA or ACC.  The NEK6-GFP in the ark1-1 background showed no changes with ACC and GA (Figure 3.7). Similarly, the ARK1-GFP in the nek6-1 background revealed no changes in ARK1’s plus-end localization in GA and ACC treatments (Figure 3.8). Taken together, ACC and GA treatments do not reveal hormone-specific interactions and functions between NEK6 and ARK1. 3.3.7.  Over-expression of the ARM domain inhibits NEK6-GFP expression  Because previous yeast two-hybrid and pull-down data revealed a direct interaction between C-terminal domains of NEK6 and the ARM domain of ARK1, I wanted to determine if ARM over-expression would reveal a putative conditional NEK6 interaction (i.e. will ARM over-expression lead to abnormal NEK6-GFP localization?).  In order to test this, I over-expressed an ARM-RFP translational fusion protein driven by the UBIQUITIN 10 promoter and transformed the construct into the ibo1-1 ark1-1; NEK6pro:NEK6-GFP line.  Previous attempts to over-express full-length ARK1-RFP in stable transgenic lines were unsuccessful, likely as a consequence of ARK1’s role in microtubule catastrophe induction (see Chapter 2).  Of 30 independent ibo1-1 ark1-1; NEK6pro:NEK6-GFP lines transformed with the UBQ10pro:ARM-RFP, only two lines exhibited ARM-RFP fluorescence, and in both 	 69 lines, ARM-RFP was distributed along the entire microtubule length (Figure 3.9A). In these same two lines, NEK6-GFP could not be detected, either along microtubules or in the cytoplasm.  While the remaining 28 transgenic lines were also BASTA-resistant (indicative of the presence of the UBQ10pro:ARM-RFP transgene construct inserted into the genome), these lines lacked RFP microtubule labeling but NEK6-GFP microtubule-labelling was present. This suggested ARM-RFP expression specifically caused the loss of NEK6. To determine if the lack of NEK6-GFP microtubule labeling resulted from increased cytosolic NEK6-GFP or reduced NEK6-GFP expression, RT-PCR was done to detect NEK6-GFP expression in transgenic lines expressing ARM-RFP.  In both transgenic lines expressing ARM-RFP, no bands were detected using GFP-specific primers, thus suggesting inhibition of NEK6-GFP transcription or degradation of mRNA (Figure 3.9B).  To exclude the possibility that lack of NEK6-GFP gene expression was a result of the absence of the NEK6pro:NEK6-GFP transgene, the same lines with absent NEK6-GFP transcript were genotyped using GFP-specific primers.  In all lines, the NEK6pro:NEK6-GFP transgene was present.  Unexpectedly, however, the nek6 epidermal protrusions were absent in plants with ARM-RFP overexpression, indicating continued nek6 complementation by NEK6pro:NEK6-GFP despite the absence of NEK6-GFP transcript in the same plants.  Nevertheless, these results suggest that the ARM domain of ARK1 may have some function in the regulation of NEK6 expression, either directly or indirectly.  3.4.  Discussion  3.4.1.  ARK1 does not require NEK6 for localization, function, phosphorylation, and regulation of activity  Based on ARK1-GFP showing unchanged plus-end localization and dynamics in the nek6-1 background relative to the wild-type background, particularly in root hairs, where the role of ARK1 is most critical (Figure 3.5), I can conclude that ARK1’s microtubule localization does not depend on NEK6’s ability to bind microtubules or its kinase activity. In Chapter 2, I showed that the ARK1-RFP and ARM-RFP proteins driven by the UBQ10 promoter labelled the entire length of the microtubule (Figure 2.12B, 2.13B) (Eng and Wasteneys, 2014).  Based on the observation that these RFP constructs labelled microtubule domains that lacked NEK6-GFP (based on NEK6-GFP punctate and uneven microtubule 	 70 distribution) (Figure 3.4), I can conclude ARK1 is able to localize to microtubules without any direct interaction with NEK6.  This conclusion is further supported by the observations from the ARK1-TagRFP and NEK6-GFP co-localization study in which ARK1-TagRFP bound to microtubule domains where NEK6-GFP was absent (Figure 3.1). The findings of my investigation further indicated that ARK1’s function does not rely on NEK6 phosphorylation activity because ARK1-GFP activity, as assessed by rescue of the crooked root hair phenotype, appeared normal in nek6-1 knock out mutant lines.  This in vivo observation, however, is inconsistent with in vitro data showing ARK1’s ability to be phosphorylated by NEK6 (H. Motose, personal communication).  Given that both NEK4 and NEK5 are co-expressed with NEK6, it could be predicted that ARK1 phosphorylation activation remains in nek6-1 because it is able to be phosphorylated by NEK4 and NEK5 despite there being no NEK6.  However, we can rule out this hypothesis because NEK4 and NEK5 have been shown to only be functional when they act as heterodimers with NEK6 (Motose et al., 2011).  In some kinesins, the cargo-binding tail domain is able to regulate kinesin activity through autoinhibition.  For example, without any cargo binding to the tail-domain, the domain is able to adopt a folded conformation to block the motor domain’s ATPase function or the microtubule binding of the kinesin.  However, binding of cargo to the tail-domain releases the autoinhibition to activate kinesin function (Ganguly and Dixit, 2013; Hirokawa et al., 2009).  With respect to ARK1 and NEK6, I can conclude that ARK1 does not require NEK6-binding for regulating its activity.   This is evident by the observation that ARK1 activity and plus-end microtubule localization continues without the presence of NEK6.  In other words, ARK1 is not autoinhibited when NEK6 is absent. Together, the data suggest that ARK1 functions independently from NEK6. 3.4.2.  NEK6 does not require ARK1 for microtubule localization or function  Several observations indicate that NEK6 localization and function do not depend on ARK1. I noted that NEK6-GFP localization to microtubules is not altered when ARK1 is missing (Figure 3.4), which demonstrate that NEK6 localizes to microtubules independently of ARK1.  This conclusion is further supported by the fact that NEK6-GFP localizes to microtubule regions not occupied by ARK1 (seen with the ARK1-GFP reporter), particularly at points along the microtubule sidewalls, as well as depolymerizing microtubule plus ends, 	 71 and microtubule minus ends.  Finally, in the nek6-1 ark1-1 double mutants, the number of epidermal protrusions was not significantly different from the nek6-1 single mutants (Figure 3.3), suggesting that NEK6 function is unaffected by the loss of ARK1. There are two possible explanations for the above observations. First, the findings indicate that there is no ARK1 and NEK6 genetic or physical interaction, at least during elongation of hypocotyl and petiole epidermal cells.  Alternatively, the absence of any changes in NEK6 localization, or number of epidermal protrusions in single and double nek6-1 mutants, could result from ARK1 functional redundancy with its homologue, ARK2, which is also expressed in the hypocotyls and petioles (Sakai et al., 2008). ARK2 might compensate for the lack ARK1, thus leading to unchanged NEK6-GFP microtubule labeling in ark1-1.  In the future, NEK6-GFP should be crossed into an ark1 ark2 double mutant to check for changes in NEK6-GFP localization. 3.4.3. ARK1 may play a regulatory role in NEK6 expression  The unexpected lack of NEK6-GFP expression in UBQ10pro:ARM-RFP plants is an intriguing finding (Figure 3.9).  Although the initial (and most logical) explanation was that the over-expression of ARM-RFP was simply out-competing NEK6 for the microtubule-binding sites, leading to increased cytosolic NEK6, I ruled this out, first by the lack of GFP fluorescence in the ARM-RFP overexpressing plants, and second, by determining that NEK6-GFP transcript was absent.  The lack of NEK6 transcript also rules out the idea that NEK6-GFP is absent due to protein degradation.  It is unknown why the plants absent with NEK6-GFP expression still lacked the nek6 mutant protrusions given that there appears to be no NEK6-GFP expression to complement the nek6 phenotype.  Based on this the lack of NEK6-GFP caused by ARM-RFP over-expression, I can hypothesize that the ARK1 may be responsible for transcriptional regulation of NEK6 through an unknown mechanism. At this point, it remains possible that NEK6-GFP expression is either absent due to suppression of NEK6-GFP transcription or degradation of NEK6-GFP transcript or a combination of both.    Previous studies that have identified involvement of motor proteins in modulating gene expression have indicated a role in the spatial distribution of mRNA (Martin and Ephrussi, 2009) rather than a direct function in gene transcription.  More interestingly, work on the O. sativa kinesin, BRITTLE CULM12 (BC12) was shown to have the ability to bind microtubules as well as to the promoter of a gene related to GA biosynthesis (Li et al., 2011, 	 72 12).  While work on BC12 revealed cytoplasmic and nuclear localization, suggesting a role in transcriptional regulation, ARK1 has only been characterized to be cytoplasmically distributed (Eng and Wasteneys, 2014).  Based on this precedent, future studies on ARK1-NEK6 interactions, should consider elucidating a role for microtubules and ARK1 in regulating NEK6 expression. 3.4.4.  Is there a physical NEK6-ARK1 interaction in vivo?  Based on my data, the functional importance of a NEK6 and ARK1 interaction remains unclear.  As previously mentioned earlier in the discussion, changes in NEK6 and ARK1 distribution patterns along microtubules did not occur in the ark1-1 and nek6-1 mutant backgrounds, respectively.  Thus, it appears that, at least under normal culture conditions, there is not an in vivo physical interaction between NEK6 and ARK1.  I initially attempted to confirm the reported NEK6-ARK1 interaction (Sakai et al., 2008) through co-immunoprecipitation (co-IP) experiments using an anti-GFP to pull down any putative binding partners to ARK1-GFP.  My co-IP experiment did not work since I was unable to obtain sufficient amounts of ARK1-GFP protein to be visualized by western blot.  As previously mentioned, the NEK6-ARK1 interaction was first recovered from a yeast two-hybrid library screen of potential ARK1 interactors but also supported by an in vitro pull-down experiment (Sakai et al., 2008). The yeast two-hybrid screen used, however, only used the C-terminal ARM-repeats of ARK1 as the bait that eventually isolated NEK6 as an interactor. Since there are 108 predicted proteins in the Arabidopsis genome that possess ARM-repeats (Mudgil et al., 2004), NEK6 could have potentially interacted non-specifically with the ARM domain of ARK1.  For example, a yeast two-hybrid screen revealed that NEK6 interacts with an ABA positive-regulator protein called ARIA that also possesses an ARM-repeat domain (Lee et al., 2010). Because ARM-repeats generally confer protein-protein interactions (Tewari et al., 2010), this could have led to a false-positives in the both the yeast two-hybrid and in vitro pull down assay between ARK1 and NEK6 due to the non-specificity binding activity of the of ARM domain.  The non-specificity of the ARM domain has also demonstrated through a previous in vitro experiment which showed that the ARM domain of ARK1 was able to co-sediment with actin filaments (Yang et al., 2007) despite in vivo data showing that the ARM domain is specifically localized to microtubules (Eng and Wasteneys, 2014) .  Thus, finding stronger evidence for the NEK6-ARK1 interactions needs 	 73 to be accomplished.  Further experiments to confirm the physical interaction between ARK1 and NEK6 will be discussed in section 5.2.2. 3.4.5.  Conclusion: NEK6 and ARK1 modulate microtubule dynamics and cell expansion through separate pathways  The genetic analysis of nek6-1 ark1-1 and nek6-1 mutants showed no significant difference in the number of epidermal protrusions in light grown hypocotyls suggesting that ARK1 does not play a role in regulating microtubules and cell expansion, specifically in the hypocotyl.  However, as previously mentioned, the one caveat to this conclusion is the presence of ARK2 redundancy with ARK1 in the hypocotyls, which and may explain why there is no difference in the number of epidermal protrusion in between nek6-1 ark1-1 and nek6-1 mutants.  This conclusion is further supported by the fact the proteins are not dependent on each other for localization and/or function under normal conditions or in the presence of enhanced GA or ethylene signalling.  While the results of Chapter 2 (Eng and Wasteneys, 2014) have confirmed that ARK1 can induce microtubule catastrophe, the exact microtubule regulatory function of NEK6 still remains unclear.  Previous work showed that microtubules in nek6 mutants were more frequently in the pause state (while other microtubule dynamic parameters were no different between wild-type and nek6 plants) and NEK6 was shown to be able to phosphorylate tubulin in vitro (Motose et al., 2008).  This suggests that NEK6 phosphorylates tubulin in order to increase microtubule turnover by a mechanism that is independent of ARK1. The intriguing finding that overexpressing of the ARM domain of ARK1 can eliminate NEK6 transcript, however, leaves open the question as to whether ARK1 and NEK6 interact.   	 74  Figure 3.1.  ARK1-TagRFP and NEK6-GFP have overlapping but non-identical distribution on microtubules. (A) Bright field images of ibo1-1ark1-1; NEK6pro:NEK6-GFP ARK1pro:ARK1-TagRFP (left panel), wild-type (right panel), and ibo1-1ark1-1; NEK6pro:NEK6-GFP (right panel) roots.  ark1-1 root hairs are rescued by the ARK1pro:ARK1-TagRFP construct showing complete functionality of ARK1-TagRFP. Scale bars represent 100 µm. (B) Confocal micrograph image of ARK1-TagRFP (in red), NEK6-GFP (in cyan), and the merged image of the both micrographs.  ARK1-TagRFP labels the entire length of the microtubule while ARK1-GFP has an asymmetric microtubule distribution as previously reported (see Chapter 2).  Images are of root epidermal cells. Scale bars represent 10 µm.  ibo1-1 ark1-1; NEK6pro:NEK6-GFP ARK1pro:ARK1-TagRFP A B ARK1-TagRFP NEK6-GFP Merge C ARK1-TagRFP NEK6-GFP Merge ARK1-GFP ARK1-GFP Root Epidermal Cells Hypocotyl Epidermal Cell Wild type ibo1-1 ark1-1; NEK6pro:NEK6-GFP 	 75 Figure 3.1.  ARK1-TagRFP and NEK6-GFP have overlapping but non-identical distribution on microtubules. (C) Confocal micrograph image of ARK1-TagRFP (in red), NEK6-GFP (in cyan), and the merged image of the both micrographs.  ARK1-TagRFP labels the entire length of the microtubule while ARK1-GFP (right-most panel) has an asymmetric microtubule distribution as previously reported (see Chapter 2).  Images are of hypocotyl epidermal cells. Bright puncta in the NEK6-GFP channel are autofluorescence from chloroplasts.  Scale bars represent 10 µm.   	 76  Figure 3.2.  ARK1-TagRFP labelling of the entire microtubule length persists in the ark1-1; 35Spro:EB1b-GFP line.  (A) Bright field images of ark1-1; 35Spro:EB1b-GFP ARK1pro:ARK1-TagRFP (left panel), wild-type (center panel), and ark1-1; 35Spro:EB1b-GFP (right panel) roots.  ark1-1 root hairs are rescued by the ARK1pro:ARK1-TagRFP showing complete functionality of ARK1-TagRFP (left panel) but is not rescued by 35Spro:EB1b-GFP	(right panel). Scale bar represents 100 µm. (B) Confocal micrograph image of ARK1-TagRFP (red), EB1b-GFP (in cyan), and a merged image of both micrographs.  Images are of a hypocotyl epidermal cell. The microtubule-labelling pattern of ARK1-TagRFP in the ark1-1; 35Spro:EB1b-GFP background is consistent to the labeling pattern in the ark1-1 ibo1-1; NEK6pro:NEK6-GFP background (see Figure 3.1B, C).  Scale bar represents 10 µm.  ark1-1; 35Spro:EB1B-GFP ARK1pro:ARK1-TagRFP ARK1-tagRFP EB1b-GFP Merge A B  ark1-1; 35Spro:EB1B-GFP Wild type 	 77   Figure 3.3.  There is no significant difference in the number of epidermal protrusions in nek6-1 and nek6-1 ark1-1 mutants in either light- or dark-grown seedlings.  (A) Images of hypocotyl and petioles of light-grown seedlings.  Ectopic protrusions (labelled with red arrow heads) are evident in nek6-1 and nek6-1 ark1-1 mutants but not in wild-type (Col-0) and ark1-1 seedlings.  Scale bars represent 1 mm.  (B) Magnified image of ectopic epidermal protrusions in light-grown nek6-1 ark1-1 plants.  The image is magnified from the red box in (A).  Scale bar represents 0.5 mm. Col-0 ark1-1 nek6-1 Dark-grown nek6-1 ark1-1 Number of Epidermal Protrusions per plant 0 2 4 6 8 10 12 14 Col-0 ark1-1 nek6-1 nek6-1 ark1-1 Col-0 ark1-1 nek6-1 nek6-1 ark1-1 Light-grown Dark-grown nek6-1 nek6-1 ark1-1 n = 20 n = 20 n = 80 n = 83 n = 59 n = 40 n = 94 n = 89 B C ark1-1 Col-0 > > > > > > > > Light-grown A D NS NS 	 78 Figure 3.3.  There is no significant difference in the number of epidermal protrusions in nek6-1 and nek6-1 ark1-1 mutants in either light- or dark-grown seedlings.  (C) Images of hypocotyl and petioles of dark-grown (etiolated) seedlings.  Epidermal protrusions are absent in all genotypes.  The black bracket in the Col-0 panel indicates the hypocotyl region. Scare bars represent 1 mm. (D) Bar graph representing the mean number of epidermal protrusions in hypocotyls and petioles of light- and dark-grown seedlings.  The difference in number of protrusions between nek6-1 and nek6-1 ark1-1 genotypes is not significant in both light -and dark-grown seedlings  (measured using a two-sample t-test with unequal variance; α = 0.01).  Ectopic protrusions are absent in dark-grown plants of all genotypes. Data and bars are represented as mean ± SD, respectively. n = number of plants sampled; NS = no significant difference using α = 0.01.   	 79  Figure 3.4.  NEK6-GFP microtubule distribution on microtubules is not altered in the ark1-1 mutant. Confocal micrograph images of NEK6-GFP in different cell types (root hair, hypocotyl epidermal, root epidermal).  The distribution of NEK6-GFP in the ibo1-1 (complemented) background (left panels) is not different from NEK6-GFP in the ibo1-1 ark1-1 background (right panels). Images are merged optical z-slices. Scale bars represent 30 µm ibo1-1; NEK6pro:NEK6-GFP  ibo1-1 ark1-1; NEK6pro:NEK6-GFP Root Hair Hypocotyl Epidermal Root Epidermal 	 80    Figure 3.5.  ARK1-GFP remains associated with microtubule plus ends in the nek6-1 mutant.  A ARK1-GFP Merge mCherry-MAP4MBD Cortical Medial C ARK1-GFP in nek6-1 ark1-1 ARK1-GFP in ark1-1 ARK1-GFP in nek6-1 ark1-1 B ARK1-GFP Merge mCherry-MAP4MBD ARK1-GFP Merge mCherry-MAP4MBD ARK1-GFP mCherry-MAP4MBD Merge 	 81 Figure 3.5.  ARK1-GFP remains associated with microtubule plus ends in the nek6-1 mutant.   (A) Confocal micrograph images of ARK1-GFP in the nek6-1 ark1-1 double mutant.  ARK1-GFP (in red) remains bound to microtubules (mCherry-MAP4MBD marker in cyan) in elongating root hairs.  Images are taken from the cortical (top panel) and medial (bottom panel) sections of an elongating root hair.  Merged panels of both channels are on the right.  Arrowheads indicate areas of ARK1 and microtubule co-localization.  Scale bar represents 5 µm.   (B) Bright-field image of root hairs with ARK1-GFP in the nek6-1 ark1-1 background showing straight root hairs.  Root hairs are still complemented by the ARK1-GFP construct, suggesting continued ARK1-GFP function despite the lack of NEK6.  Scale bar represents 200 µm.    (C) Confocal images of ARK1-GFP in the ark1-1 (complemented) background (top row) and the nek6-1 ark1-1 double mutant (bottom row).  ARK1-GFP (in red) remains associated with microtubule plus ends (mCherry-MAP4MBD marker in cyan) in hypocotyl epidermal protrusions found in the nek6-1 ark1-1 background. Merged panel of both channels is on the right.  The yellow-outlined panels are of enlarged images outlined in yellow in the merged panels.  Scale bar represents 10 µm.   	 82  Figure 3.6.  Localization of ARK1-TagRFP and NEK6-GFP to microtubules is unchanged when exposed to GA or ACC. (A) Confocal images of cells expressing ARK1-TagRFP NEK6-GFP in ibo1-1 ark1-1 double mutant.  Images are of untreated (mock) hypocotyl epidermal cells. (B) Confocal images of the same transgenic lines seen in (A) but exposed to 10 µM GA.  ARK1-TagRFP and NEK6-GFP microtubule distribution remained the same.  Images are of petiole epidermal cells. (C) Confocal iamges of the same transgenic lines seen in (A) but exposed to 50 µM ACC.  ARK1-TagRFP and NEK6-GFP microtubule distribution remained the same.  Images are of petiole epidermal cells. Plants were grown on Hoagland’s media (no sucrose) for four days before being transferred to media supplemented with the GA or ACC for two days.  Scale bars represent 20 µm.  ARK1-TagRFP NEK6-GFP Merge A Mock B + 50 µM ACC   C + 10 µM GA   ARK1-TagRFP ARK1-TagRFP NEK6-GFP NEK6-GFP Merge Merge 	 83  Figure 3.7.  Localization of NEK6-GFP on microtubules does not change when plants are exposed to exogenous GA or ACC. Confocal images of NEK6-GFP in plants grown either without hormones (mock), with 10 µM GA, or with 50 µM ACC.  NEK6-GFP remains localized to microtubules in a punctate and uneven fashion in hormone-treated seedlings and appears no different from the mock treatment.  Images are of hypocotyl epidermal cells.  Plants are of ibo1-1 ark1-1; NEK6pro:NEK6-GFP lines.  Plants were grown on Hoagland’s media (no sucrose) for four days before being transferred to media supplemented with the GA or ACC for two days.Scale bars represent 20 µm.   Mock  + 10 μM GA  + 50 μM ACC 	 84  Figure 3.8.  Microtubule Localization of ARK1-GFP is unchanged when exposed to GA or ACC. (A) Confocal images of cells expressing ARK1-GFP in nek6-1 ark1-1 with a mCherry-MAP4MBD microtubule marker.  As previously seen in Chapter 2, ARK1-GFP is specifically localized to growing microtubule plus ends.  Images are of untreated (mock) hypocotyl epidermal cells. (B) Confocal micrographs of the same transgenic lines seen in (A) but exposed to 10 µM GA.  ARK1-GFP still associates with microtubule plus ends.  Images are of hypocotyl epidermal cells. (C) Confocal micrographs of the same transgenic lines seen in (A) but exposed to 50 µM ACC.  ARK1-GFP still associates with microtubule plus ends.  Images are of hypocotyl epidermal cells. Plants were grown on Hoagland’s media (no sucrose) for four days before being transferred to media supplemented with the GA or ACC for two days.  Scale bars represent 20 µm.A Mock B + 50 µM ACC   C + 10 µM GA   ARK1-GFP mCherry-MAP4MBD Merge ARK1-GFP mCherry-MAP4MBD Merge ARK1-GFP mCherry-MAP4MBD Merge 	 85  Figure 3.9.  Overexpression of ARM-RFP leads to complete loss of NEK6-GFP expression in ibo1-1 ark1-1; NEK6pro:NEK6-GFP UBQ10pro:ARM-RFP plants. (A) Confocal micrograph images of ibo1-1 ark1-1; NEK6pro:NEK6-GFP UBQ10pro:ARM-RFP hypocotyls expressing either ARM-RFP (top panel) or no ARM-RFP (bottom panel).  Hypocotyls overexpressing ARM-RFP lack NEK6-GFP.  In plants that are not expressing UBQ10pro:ARM-RFP, NEK6-GFP expression is still present.  ARM-RFP labels the entire length of the microtubule.  Scale bars represent 50 µm. (B) RT-PCR gel using GFP-specific primers (to detect NEK6-GFP) and F-actin primers (as a control). NEK6-GFP expression is knocked-out when UBQ10pro:ARM-RFP is being expressed.  NTC = no template control. Lanes (left to right): 1.  Col-0 (wild-type)  2.  ibo1-1 ark1-1; NEK6pro:NEK6-GFP 3.  ibo1-1 ark1-1; NEK6pro:NEK6-GFP UBQ10pro:ARM-RFP 4.  NTC   Merge no NEK-GFP Hypocotyls without ARM-RFP expression Hypocotyls with ARM-RFP expression A F-actin NEK6-GFP B Col-0 ibo1-1 ark1-1; NEK6-GFP ibo1-1 ark1-1; NEK6-GFP ARM-RFP NTC ARM-RFP no ARM-RFP NEK-GFP Merge 	 86 Chapter 4: MOR1 is a plus-end tracking protein that acts as a microtubule polymerase 4.1. Background Information  MICROTUBULE ORGANIZATION 1 (MOR1) is an important microtubule-associated protein essential for the growth and development of A. thaliana. The indispensible nature of MOR1 protein is evident from one if its mutant alleles, gem1-1,  having homozygous embryo lethality (Twell et al., 2002).  MOR1 belongs to the highly conserved XMAP/Dis1 family of proteins that have a variety of functions in microtubule dynamics and organization in fungi, plants, and animals.  Proteins in the family all have repeating domains at the N-terminus called TOG (tumour overexpressed gene) domains (named after the human homologue, ch-TOG).  The number of TOG domains, however, varies depending on the species.  MOR1 contains five TOG domains as do the human (ch-TOG), Xenopus laevis (XMAP215), and Drosophila melanogaster (MSPS) homologues, while other species, such as yeast, have homologues with only two TOG domains (Saccharomyces cerevisae: Stu2; Schizosaccharomyces pombe: Dis1, Alp14) (Gard et al., 2004).  Each TOG domain contains six HEAT (Huntington, Elongation Factor 3, Protein Phosphatase 2A, TOR PI-3 Kinase) repeats that are folded into side-by-side α-helices connected by intra-HEAT repeat loops (Al-Bassam et al., 2007; Slep and Vale, 2007; Al-Bassam and Chang, 2011; Lechner et al., 2012).  The role of the TOG domains (and intra-HEAT repeats) varies across the species as well as across the different TOG domains within the same protein.  For example, the first two TOG domains of XMAP215 are involved in binding free tubulin and its subsequent polymerization into the microtubule polymer (Widlund et al., 2011).  Conversely, the first two TOG domains (TOG12) of MOR1 are predicted to bind microtubule polymers rather than tubulin subunits (Lechner et al., 2012).  This prediction is based on the finding that the TOG12 domain contains a conserved microtubule binding motif normally found in regions outside of the first two TOG domains in the MOR1 homologues, XMAP215 and Msps (Lechner et al., 2012; Currie et al., 2011; Widlund et al., 2011).  The importance of the TOG12 domain was made apparent through experiments that revealed that the domain alone was able to promote microtubule polymerization in vitro and rescue the mor1-1 mutant phenotype in vivo (Lechner et al., 2012).  The mor1-1 and mor1-2 mutants, which both have 	 87 point mutations occurring within the TOG12 domain, have drastically altered microtubule dynamics and organization (Whittington et al., 2001; Kawamura et al., 2006; Kawamura and Wasteneys, 2008) and highlight the necessity of the TOG12 domain for MOR1 function.  Based on this data, it is predicted that the presence of just the TOG12 domain is sufficient for function, which is entirely possibly given that the Stu2p and Dis1p proteins are able to function with just two TOG domains.    MOR1 is predicted to act as a microtubule polymerase but might also assist with depolymerization.  This prediction is based on research visualizing microtubules in the temperature-sensitive mutant allele, mor1-1, which has a leucine to phenylalanine amino acid substitution in the first TOG domain (Figure 4.1).  At the restrictive temperature, cortical microtubules of mor1-1 plants were fragmented and shortened instead of forming the typical long microtubules organized into parallel arrays (Whittington et al., 2001).  Furthermore, microtubules within the PPB, spindle, and phragmoplast were either not present or disrupted, resulting in aberrant mitotic activity of mitotic cells (Kawamura et al., 2006).   Live-cell imaging of microtubules in mor1-1 revealed reduced microtubule polymerization and depolymerization rates at the restrictive temperature.  In addition, microtubules were shown to have spent a greater amount of time in the pause phase, stressing the importance of MOR1 in promoting dynamic microtubules (Kawamura and Wasteneys, 2008).  Studies on MOR1 homologs have also shown similar functions in regulating microtubule dynamics.  For example, work on XMAP215 and Stu2 revealed that these proteins also act as microtubule polymerases (Breugel et al., 2003; Brouhard et al., 2008; Podolski et al., 2014). While the general function of MOR1 has been determined, its mechanism of function and microtubule association remains unclear.  With regards to microtubule localization, two different studies using immunofluorescence with antibodies targeted towards different epitopes of MOR1 resulted in conflicting observations.  In one study, antibodies were raised against a C-terminal fragment of the protein (Twell et al., 2002). Immunofluorescence of Arabidopsis suspension and protoplast cultures showed weak punctate distribution along interphase microtubules but increased labelling at the mid-zone of the mitotic spindles and phragmoplasts, suggesting that MOR1 is a plus-end localized protein (Twell et al., 2002).  In a second study, antibodies raised against a synthetic peptide from the N-terminus of MOR1 	 88 showed labelling of the entire microtubule length of multiple arrays throughout the cell cycle (Kawamura et al., 2006). Thus, immunolabelling has been of limited use for understanding MOR1’s distribution in relation to its function in promoting microtubule assembly and disassembly, which are activities confined to the dynamic plus end.  These studies using MOR1 immunofluorescence are at odds with localization studies of the MOR1 homologues from animal and fungal systems. In vitro work of non-fixed samples (XMAP215: (Brouhard et al., 2008); Stu2: (Breugel et al., 2003)) and in vivo live-cell imaging (Alp14: (Garcia et al., 2001; Al-Bassam et al., 2012)) of various MOR1 homologs revealed plus-end binding on polymerizing microtubules, thus classifying these MAPs as +TIPs.  Based on the fact that these proteins have similar function to MOR1, the data suggests that MOR1 may also function as a +TIP.  In order to confirm this, live-cell imaging of MOR1 needed to be accomplished. Due to the considerable size of the MOR1 gene (about 14 kB), many previous attempts to clone the gene and fuse it to a fluorescent protein tag coding sequence for live-cell imaging have been unsuccessful.  Live-cell imaging has been achieved, however, with GFP fused to the C-terminus of the N-terminus of MOR1 comprising the first two TOG domains (TOG12-GFP) and, consistent with the immunofluorescence results of Kawamura et al. (2006), this reporter protein distributed along the entire microtubule sidewall (Lechner et al., 2012). In spite of this N-terminal MOR1 fragment being able to rescue the mor1-1 temperature-sensitive phenotype, it did not bind to microtubules in the presence of full-length wild-type MOR1. Thus the TOG12-GFP reporter is not fully functional and of limited use in understanding MOR1’s localization. To circumvent the problems related to the cloning of MOR1, a bacterial homologous recombination system (recombineering) (Zhou et al., 2011) was used in this study to create a MOR1pro:MOR1-3xYpet construct (Ypet is a variant of the yellow fluorescent protein) that enabled live-cell imaging of MOR1.  Recombineering takes advantage of the bacteriophage recombination machinery and allows for the creation of a full-length gene fused to a fluorescent reporter sequence.  Recombineering technology is advantageous relative to traditional restriction enzyme-based cloning for multiple reasons.  First, it allows for all gene regulatory regions to be included in the construct, which increases the chances of the expression for the gene of interest. Second, since the gene of interest is already in the desired 	 89 expression vector, PCR amplification is no longer required and thus, large sized genes can be easily tagged without the limitations associated with PCR. In this study, the full length MOR1 gene was fused at its 3’-end to a 3xYpet coding sequence.  A detailed explanation of the recombineering protocol can be found in 4.2.1.  The major research objective of this chapter is to characterize the localization of the MOR1 protein by visualizing MOR1-3xYpet through live-cell imaging.  By visualizing MOR1’s localization in corroboration with previous mor1-1 loss of function data (Whittington et al., 2001; Kawamura et al., 2006; Kawamura and Wasteneys, 2008; Lechner et al., 2012), a stronger hypothesis on how MOR1 functions can be made.  In this chapter, I discuss the distribution of MOR1 on microtubules and propose multiple functions for MOR1 in modulating microtubule dynamics.  4.2. Materials and Methods 4.2.1. Generation of transgenic plant material and culture A MOR1pro:MOR1-3xYpet construct was generated by Dr. Jose Alonso (North Carolina State University) through recombineering (Zhou et al., 2011).  Briefly, a transformation-competent artificial chromosome (TAC) vector with the MOR1 gene was identified and then transformed into a recombineering strain of E. coli.  A 3xYpet fluorescent protein (a YFP variant with 3 copies) cassette was synthesized using PCR.  Using specially designed primers, a PCR product coding for the 3xYpet cassette was flanked by sequences homologous to the desired regions of the TAC vector at the site of recombination downstream of the MOR1 gene.  The 3xYpet cassette was then recombined into the TAC clone at the 3’-end of the MOR1 gene.  Finally, the recombineered construct was transferred into Agrobacterium tumefaciens (GV3101 strain), which was then used to transform into an the Arabidopsis heterozygous SALK_032056 T-DNA insertion line (herein referred to as mor1-23) using a technique to enhance transformation efficiency of large TAC constructs (Alonso and Stepanova, 2014). After selection of T1 lines by BASTA-resistance and selfing, T2 seedlings homozygous for the Salk T-DNA insertion and the MOR1pro:MOR1-3xYpet transgene were recovered, selfed and T3 lines were used for subsequent experiments.  The same homozygous T3 lines were transformed with a 35Spro:mRFP-TUB6 construct (provided by Dr. Richard Cyr, Pennsylvania State University) and homozygous T3 lines were used for 	 90 further experiments.  The same mor1-23; MOR1pro:MOR1-3xYpet lines were also crossed into a EB1bpro:EB1b-mCherry reporter line and F3 lines (homozygous for the MOR1pro:MOR1-3xYpet and EB1bpro:Eb1b-mCherry transgenes, azygous for the mor1-23 T-DNA insertion) were used for further experiments.  Seeds for all experiments were sterilized and germinated with the same methodology as described in Chapters 2 and 3. All wild-type plants used are Columbia (Col-0) ecotype. 4.2.3. Chemical fixation of seedlings  6 day-old etiolated seedlings were fixed in PME buffer (25 mM PIPES, 0.5 mM MgSO4, 2.5 mM EGTA, pH 7.2) with 0.5% glutaraldehyde (Electron Microscopy Sciences) and 1.5% paraformaldehyde (Electron Microscopy Sciences) for 30 minutes under vacuum infiltration.  Seedlings were then washed three times with PMET buffer (PME buffer with 0.05% Triton-X 100).   4.2.3. Microscopy Imaging of the live- and fixed-samples was done using a Perkin Elmer Ultraview VoX Spinning Disc Confocal system (Perkin-Elmer) mounted on a Leica DMI6000 B inverted microscope and equipped with a Hamamatsu 9100-02 electron multiplier CCD camera (Hamamatsu). An argon 516 nm laser line with a complementary YFP (540/30) emission band-pass filter or a 561 nm laser with a complementary RFP (595/50) emission band-pass filter was used.  5 to 7 day old seedlings (either light- or dark-grown) were used for imaging.  Time-lapse images were acquired every 5 seconds with a 63x (glycerol) objective lens (NA = 1.3).  Optical z-stacks of 0.2 – 1.0 µm slices were acquired. 4.2.4. Image processing and data analysis  Stacks of z-slices were merged using the Z-project function in ImageJ (http://rsbweb.nih.gov/ij/).  A cross-section of a z-stack was created using the Reslice function.  Images were stitched together using the Stitching plug-in (Preibisch et al., 2009).  To measure the MOR1-3xYpet microtubule distribution, the fluorescence intensity was measured using the RGB Line Profiler plug-in (http://rsb.info.nih.gov/ij/plugins/rgb-profiler.html).  Specifically, 12 line-scans (of 25 pixels in length) were taken of microtubule plus ends and the fluorescence intensity of both Ypet and mRFP were measured.  The average intensity of each point and channel was measured.  The background fluorescence 	 91 was also measured and subtracted from the measured fluorescence of the line-scan. A similar method was used to measure MOR1-3xYpet length on microtubules.  4.3. Results 4.3.1. MOR1pro:MOR1-3xYpet rescues the homozygous lethality of mor1-23 Due to the size of the MOR1 gene, engineering a full-length MOR1-fluorescent fusion protein has been unsuccessful and thus, live-cell imaging of MOR1 has yet to be done.  In order to visualize live-cells expressing MOR1 tagged with a fluorescent reporter, a MOR1pro:MOR1-3xYpet construct (Ypet is a variant of the yellow fluorescent protein) was generated using a bacterial homologous recombination system (recombineering) (Zhou et al., 2011).  In this study, the full length MOR1 gene was fused at its 3’-end to a 3xYpet coding sequence.  Once created, the MOR1pro:MOR1-3xYpet construct was transformed into a mor1 mutant line.  Since the already characterized mor1 mutant alleles (mor1-1, mor1-2) are known to produce mutant MOR1 proteins (Whittington et al., 2001), and have no phenotype under standard culture conditions, it was preferable to use, if possible, a null transcript allele that would provide unequivocal evidence that the MOR1-3xYpet protein is functional. To accomplish this, I used a MOR1 T-DNA insertion line, SALK_032056 (with its T-DNA inserted into MOR1’s last intron) (Figure 4.1), which will herein be referred to as mor1-23. Owing to MOR1’s essential nature, progeny homozygous for the transgene are embryo lethal but heterozygous mor1-23 plants are fully viable and wild type in appearance.  After transforming heterozygous mor1-23 plants with the MOR1-3xYpet construct, I was able to recover one line (out of 42 T2 lines) that was both homozygous for mor1-23 and the MOR1pro:MOR1-23xYpet transgene.  Since MOR1pro:MOR1-3xYpet was able to completely complement the homozygous lethality phenotype of the mor1-23, I was able conclude complete functionality of the fusion protein (Figure 4.2). 4.3.2. MOR1 is expressed in all cell types.  A previous study looking at MOR1 expression patterns was based on RT-PCR experiments using whole tissue samples (e.g. roots, cotyledons, rosette leaves) (Whittington et al., 2001).  This study, however, did not look at MOR1 expression patterns at the microscopic and cell-specific level.  Based on the breadth of mor1-1 phenotypes across cell types at the restrictive temperature, it has been predicted that MOR1 is expressed and 	 92 required for microtubule functions throughout all cell types. Using spinning disc confocal microscopy, I was able to visualize MOR1-3xYpet in all cell types using the mor1-23; MOR1pro:MOR1-3xYpet transgenic line.  As expected, MOR1-3xYpet was widely expressed in all cell-types of the both aerial and root tissue (Figure 4.3).  The MOR1-3xYpet distribution pattern in each cell type is reminiscent of the canonical microtubule orientation known in each cell type (Wasteneys and Ambrose, 2009; Ehrhardt and Shaw, 2006b; Bruaene et al., 2004).  In the root tip, MOR1-3xYpet labelled various types of mitotic microtubule structures, which will be discussed further in section 4.3.3.  Moreover, MOR1’s exact distribution on microtubules will be further explained. 4.3.3. MOR1-3xYpet localizes to mitotic and cytokinetic microtubule arrays  In order to determine MOR1’s specific distribution on microtubules, the mor1-23; MOR1pro:MOR1-3xYpet line was transformed with a 35Spro:mRFP-TUB6 microtubule marker construct. Using spinning-disc confocal microscopy, I was able to visualize MOR1-3xYpet on microtubules in the same cell types mentioned in 4.3.2.  In the root tip where cells are undergoing mitosis and cell division, MOR1-3xYpet localized to PPBs and the nuclear envelope (Figure 4.4, 4.5), mitotic spindles (Figure 4.5), and phragmoplasts (Figure 4.6).  Upon closer inspection, I noted that MOR1-3xYpet localizes to the microtubules throughout prophase, prometaphase, metaphase, and anaphase (Figure 4.5). MOR1-3xYpet localization on mitotic microtubule structures (spindle, phragmoplast) was similar to the MOR1 immunofluorescence labelling seen in Twell et al. (2002). During telophase, phragmoplast microtubules are arranged with their plus ends pointing inwards towards the mid-zone of the dividing cell (Rasmussen et al., 2013) (Figure 4.6A).  In these cells, it was noticed that MOR1-3xYpet had increased fluorescence in the domain of the phragmoplast where microtubule plus ends are occupied (Figure 4.6B) as previously reported with another study using MOR1 immunofluorescence labelling (Twell et al., 2002). In order to quantify MOR1-3xYpet’s plus-end distribution in the phragmoplast, I measured MOR1-3xYpet’s fluorescence intensity along the length of the phragmoplast and found an increase in intensity in domains closer to the microtubule plus ends.  Conversely, in the phragmoplast domains of increased MOR1-3xYpet fluorescence intensity, mRFP-TUB6 intensity was reduced relative to the rest of the phragmoplast microtubules (Figure 4.6B, C).  	 93 These observations suggest that MOR1 accumulates more at the plus ends of microtubules in the phragmoplast. 4.3.4. MOR1-3xYpet has an asymmetrical plus-end distribution and is a plus-end tracking protein To further test MOR1’s plus-end localization on microtubules, I examined etiolated hypocotyls, in which it is easier to visualize microtubules and therefore get a more accurate representation of MOR1’s localization and dynamics related to microtubules.  MOR1-3xYpet was noted to label all growing plus ends (Figure 4.7A, B, D), although not as punctate relative to another plus-end tracking MAP such as EB1b.  Instead, MOR1-3xYpet showed highest fluorescence intensity right at the plus-end followed by diminished intensity immediately behind the plus ends of microtubules.  This distribution results in a comet-like appearance.  100% of polymerizing microtubules had MOR1-3xYpet localized to the plus-end (n = 100 polymerizing microtubule events in 7 cells).  In order to quantitatively determine MOR1’s asymmetric distribution on microtubules, I measured the fluorescence intensity of MOR1-3xYpet along the growing plus ends of microtubules using a line scan (Figure 4.7B, C).  The fluorescence intensity of MOR1-3xYpet is shown to be highest at the microtubule plus end with the intensity decreasing distally, which is consistent with the visual observations.  As a control, the fluorescence intensity of the microtubule (mRFP-TUB6) was also measured, which showed even fluorescence distribution along the line scan.   To further characterize MOR1-3xYpet, I looked at MOR1-3xYpet’s association with microtubules in various microtubule states.  Depolymerizing microtubules were shown to have MOR1-3xYpet on their plus ends 93% of the time (n = 60 depolymerizing microtubule events in 7 cells) (Figure 4.8).  Moreover, MOR1-3xYpet was consistently shown to remain bound to microtubules during both catastrophe and rescue events (Figure 4.9).  Much like polymerizing plus ends, MOR1-3xYpet also had the comet-like appearance on depolymerizing plus ends.  On treadmilling microtubules, which have free (unanchored) minus-ends, MOR1-3xYpet was absent on minus ends but still remained on the plus end (Figure 4.10).  	 94 4.3.5. MOR1-3xYpet localizes to depolymerizing plus ends of severed microtubules at cross-over sites  To test the ability of MOR1 to remain associated with shrinking microtubule plus ends, I examined MOR1-3xYpet distribution following severing events (Figure 4.11A).  Depending on the cell type, when a polymerizing microtubule encounters an obstacle microtubule at a steep angle (>45°), the polymerizing microtubule cross over the obstacle microtubule (Dixit and Cyr, 2004; Wasteneys and Ambrose, 2009; Allard et al., 2010).  In specific cell types or stages of cell development such as etiolated hypocotyls, the polymerizing microtubules tend to undergo severing at the point of cross-over, resulting in depolymerization at the newly generated plus end (Wightman and Turner, 2007; Lindeboom et al., 2013b; Zhang et al., 2013) (Figure 4.11B).  I documented four such post cross-over severing events. In each case, the MOR1-3xYpet accumulated on the newly formed depolymerizing plus end but was absent from the newly created minus end lacked MOR1-3xYpet. This suggests that MOR1 can be rapidly recruited to the shrinking plus end. 4.3.6. MOR1 is able to bind to the microtubule sidewall in addition to its preferential plus-end localization MAPs that are plus-end trackers (known as +TIPs), such as EB1, have dynamic accumulations on plus ends, which lead to the appearance of +TIPs “surfing" on polymerizing microtubule plus-end.  The mechanism for this “surfing” appearance is currently unknown; however, the putative mechanism that has been studied the most is known as treadmilling (not to be confused with previously discussed microtubule treadmilling).  In +TIP treadmilling, the +TIPs are first targetted to growing microtubule plus ends (possibly due to higher affinity towards the plus end) where +TIPs are able to accumulate.  As the microtubule continues to polymerize, +TIPs are stationary relative to the plus-end, remaining bound to the microtubule at the protofilament sidewall domain behind the growing plus end.  This is followed by quick dissociation of the +TIPs from the microtubule sidewall as the microtubule continues to polymerize, leading to the comet-like appearance of +TIPs surfing along the microtubule plus end (Akhmanova and Steinmetz, 2008; Carvalho et al., 2003).  Thus, +TIPs are able to bind to two microtubule domains: the plus end and the sidewalls/lattice of microtubules.  It is currently unknown how +TIPs recognize and bind to specific microtubule domains. 	 95 In order to determine if MOR1 has specific preference for either the microtubule plus end or the microtubule sidewall, I decided to see if differential levels of MOR1 would affect its known plus-end microtubule distribution.  To accomplish this, I increased levels of the MOR1 protein by expressing MOR1pro:MOR1-3xYpet in wild-type plants, thus producing plants that express both endogenous MOR1 and MOR1-3xYpet proteins.  In contrast, plants expressing MOR1-3xYpet in the mor1-23 background would produce lower levels of MOR1 (in the form of MOR1-3xYpet) and lack endogenous MOR1 protein. Using confocal microscopy, I noted that the distribution of MOR1-3xYpet on microtubules appeared significantly different in the wild-type background compared to the mor1-23 background (Figure 4.12A).  MOR1-3xYpet in the wild-type background had increase distribution along the sidewalls of the microtubule relative to MOR1-3xYpet’s asymmetrical plus-end distribution in the mor1-23 background.  Quantification of MOR1-3xYpet’s distribution in the wild-type background showed that the protein is localized on microtubules up to 9.1±2.6 µm from the plus end.  Comparatively, MOR1-3xYpet in the mor1-23 background was more tightly distributed at the plus end with the protein being localized no more than 1.2±0.4 µm from the plus end, which is a significant reduction from the wild-type background (P<10.7x10-61) (Figure 4.12B).  Moreover, while MOR1-3xYpet in the mor1-23 background is shown as bright fluorescent punctate with a comet tail at the plus end, a large proportion of the MOR1-3xYpet fluorescence in the wild-type background appeared more even along the microtubule sidewall and lacked that bright punctae (Figure 4.12A).  Because increased levels of MOR1 may affect normal plant development of plant by potentially altering microtubule dynamics, I decided to compare seedling development in the MOR1-3Ypet-expressing wild-type and mor1-23 lines.  Phenotypically, the morphology of wild-type seedlings expressing MOR1pro:MOR1-3xYpet appeared no different from the morphology of seedlings with the mor1-23 background, suggesting that having higher MOR1 protein levels in wild-type plants had no effect on cell growth and development (Figure 4.13).  Based on the observations of MOR1-3xYpet localization in both genetic backgrounds, I can conclude that MOR1 has preferential binding to microtubule plus end but is still capable of binding older parts of the microtubule sidewalls before dissociating from the microtubule.  	 96 4.3.7. Chemical fixation affects the plus ends of microtubules and prevents +TIPs from binding Through live-cell imaging of MOR1-3xYpet, I determined that MOR1 is plus-end localized.  While one previous immunofluorescence study showed MOR1 distributed in the mid-zone of the phragmoplast and spindle (Twell et al., 2002), consistent with my data seen in Figures 4.5 and 4.6, this study did not specify or conclude that MOR1 is plus-end localized.  Moreover, these samples were of Arabidopsis suspension and protoplast cell cultures rather than whole, living seedlings used in my study. Another immunofluorescence study using fixed seedlings showed MOR1 distribution along the entire microtubule sidewall (Kawamura et al., 2006) rather than the asymmetrical distribution seen with MOR1-3xYpet.  To understand the discrepancy between the antibody labelling along the full length of microtubules and the MOR1-3xYpet plus-end distribution, I hypothesized that the microtubule plus end is depolymerized during the chemical fixation process, causing the loss of MOR1.  To test this hypothesis, I chemically-fixed mor1-23; MOR1pro:MOR1-3xYpet 35Spro:mRFP-TUB6 etiolated seedlings using a 30 min treatment with 0.5% glutaraldehyde and 1.5% formaldehyde, and then visualized microtubules and MOR1-3xYpet in elongating hypocotyls with confocal microscopy (Figure 4.14A, B).  While the microtubules remained intact in the expected transverse orientation associated with elongating hypocotyls, MOR1-3xYpet could no longer be detected on the microtubules (Figure 4.14B).  Instead, the MOR1-3xYpet signal was diffuse, suggesting cytoplasmic distribution. The loss of MOR1-3xYpet from the microtubule plus end could have derived from the fixation method affecting either the properties/structure of the plus end or the MOR1 protein itself.  To test both of these possibilities, I used the same fixation method but with a wild-type line expressing both EB1bpro:EB1b-mCherry and MOR1pro:MOR1-3xYpet.  If chemical fixation does in fact alter the plus-end properties/structure of microtubules, then the EB1b-mCherry distribution should be altered since the protein is a +TIP (Figure 4.14C).  Upon chemical fixation, the EB1b-mCherry signal became cytoplasmic (Figure 4.14D) much like MOR1-3xYpet, indicating that the fixation method does in fact alter the properties and/or structure of the plus ends.  Moreover, since MOR1-3xYpet remained localized to the microtubule sidewall domain after fixation (Figure 4.14D), the fixation method appears to not affect MOR1’s ability to interact with microtubules.  Based on these observations, I can 	 97 conclude that the chemical fixation method affects the plus ends rather than specifically affecting the MOR1 protein.  4.4. Discussion 4.4.1. MOR1 is a +TIP that functions in promoting microtubule plus-end dynamics MOR1 has been previously predicted to promote the polymerization and depolymerization of microtubules (Kawamura and Wasteneys, 2008).  Prior to my research, however, it was unknown if MOR1 functioned as a +TIP like its well-studied homologue, XMAP215 and Stu2 (Brouhard et al., 2008; Podolski et al., 2014).  My data revealing plus-end localization to growing and shrinking microtubules fits the model in which MOR1 is a +TIP that participates in controlling plus-end dynamics (Kawamura and Wasteneys 2008).  Specifically, the model suggests that MOR1 is able to increase microtubule polymerization and depolymerization frequency and rates by binding to both growing and shrinking plus ends.  The lack of MOR1 localization on microtubule minus ends reveals a specific plus-end role in microtubule regulation, similar to other previously characterized +TIPs (Carvalho et al., 2003; Akhmanova and Steinmetz, 2008). My findings also reinforce the idea of functional diversity of proteins in the XMAP215/Dis1 family.  For example, XMAP215 in vitro has been shown to promote the polymerization of microtubules by binding shrinking and growing plus ends (Brouhard et al., 2008) like MOR1.  In the same study, XMAP215 was also shown to promote microtubule depolymerization; however, this only occurred at diluted levels of tubulin concentration and thus, XMAP215 may not necessarily function as a depolymerase in vivo.  Comparatively, S. pombe’s Alp14, is shown to only promote microtubule polymerization and specifically bind to growing plus ends (Al-Bassam et al., 2012).  Moreover, while Stu2 in S. cerevisiae has been shown to have similar functions as XMAP215, Stu2 localizes to both plus and minus ends where it promotes the growth of both ends (Podolski et al., 2014). The data showing MOR1 localizing to newly created plus ends of severed microtubules is consistent with the idea that MOR1 may promote depolymerization after microtubule severing (e.g. by the action of the protein katanin (Lindeboom et al., 2013b; Wightman et al., 2013; Zhang et al., 2013)), which is a novel mechanism within the XMAP/Dis1 family of proteins as well as +TIPs in Arabidopsis.  Despite this novel 	 98 observation, it is currently unknown how MOR1 is recruited to the new plus end or if MOR1 has a role in microtubule severing. 4.4.2. How does MOR1 interact with microtubules and plus-end track?  The differences in MOR1-3xYpet labelling of microtubules between the wild-type and mor1-23 genetic background may have been a consequence of the amount of MOR1 being expressed.  Previous research looking at cells overexpressing +TIPs, for example, revealed increased microtubule sidewall labelling relative to cells expressing endogenous levels of +TIPs (Carvalho et al., 2003).  With respect to my observations, the increase in MOR1-3xYpet microtubule sidewall labelling in the wild-type background could have also resulted from increased MOR1 protein levels.  Specifically, endogenous MOR1 could have stronger affinity for the plus-end than MOR1-3xYpet and thus, MOR1-3xYpet is competitively inhibited from binding to the plus ends.   Subsequently, MOR1-3xYpet must bind further behind the microtubule plus end, leading to the observed increase in microtubule sidewall binding. Conversely, because there is no endogenous MOR1 protein in the mor1-23 background, MOR1-3xYpet is able to freely bind to and accumulate on the plus-end, resulting in the typical +TIP comet-like appearance. MOR1 is able to bind to the plus-end as well as the microtubule sidewall.  It is unknown how MOR1 targets the plus end and the microtubule sidewall.  One hypothesis suggests that +TIPs are able to recognize specific biochemical or structural characteristics of the plus ends.  For example, +TIPs could recognize the GTP cap (Zanic et al., 2009) or specific tubulin residues, the microtubule seam, or protofilament structures that are characteristic for microtubule plus ends (Carvalho et al., 2003; Sandblad et al., 2006; Akhmanova and Steinmetz, 2008).  Given that there are MOR1-3xYpet accumulations on the plus ends of depolymerizing microtubules, the hypothesis that MOR1 recognizes the GTP cap can be ruled out because GTP caps are not present in depolymerizing microtubules.  Whether MOR1 binds to other plus end structural features needs to be determined.  Since MOR1 binds to both polymerizing and depolymerizing microtubules, MOR1 must recognize a feature that is common for depolymerizing and polymerizing plus ends or have distinct mechanisms that allow it to remain bound to plus ends in both types of events.  With +TIPs that are thought to undergo treadmilling, the comet-like tail is thought to derive from quick dissociation of the +TIPs from the microtubule sidewall.  For example, 	 99 studies using EB1 from X. laevis egg extracts concluded that EB1 had strongest affinity for the plus-end and reduced affinity for the sidewall, leading to quicker dissociation from the microtubule sidewall region (and the appearance of a comet-like tail) (Tirnauer et al., 2002).  The change in microtubule affinity has been proposed to be regulated by kinases that phosphorylate +TIPs in order to decrease its affinity for microtubules. (Carvalho et al., 2003; Akhmanova and Steinmetz, 2008).  This is based on previous research showing inhibition of microtubule binding of CLIP-170 (a +TIP) when phosphorylated (Rickard and Kreis, 1991; Diamantopoulos et al., 1999; Perez et al., 1999), although the exact kinase is unknown.  The idea of +TIP dissociation as a consequence of decreased microtubule affinity via phosphorylation could explain the observations seen with MOR1-3xYpet in the mor1-23 background.  With this hypothesis, MOR1-3xYpet behind the plus-end is phosphorylated and is then quickly dissociated from the microtubule sidewall.  This mechanism, however, fails to describe what is observed with MOR1-3xYpet’s increase in microtubule sidewall labelling in the wild-type background.  In this case, the observation suggests that there is reduced MOR1-3xYpet dissociation occurring within the sidewall region.  This decrease in MOR1 dissociation may have resulted from an imbalance between the amount of MOR1 (both endogenous and Ypet fusion proteins) and the amount of kinase(s) that phosphorylate MOR1.  Since there are increased levels of MOR1 protein in the wild-type background, there may not be enough kinase(s) to phosphorylate MOR1 to signal its dissociation from the sidewall.  Because of this imbalance, MOR1-3xYpet cannot be phosphorylated and remains attached to the sidewall, as indicated through my observations.  The increase in microtubule sidewall labelling does not appear to affect microtubule dynamics or cell morphogenesis given that seedlings expressing MOR1pro:MOR1-3xYpet in the mor1-23 and wild-type backgrounds are not morphologically different from each other.  This suggests that proper regulation of microtubule dynamics occurs strictly at the plus ends of microtubules and that +TIPs, such as MOR1, has a negligible effect on microtubule dynamics at the sidewall domain. 4.4.3. Chemical fixation inhibits localization of plus-end tracking proteins Previous published data using chemical fixation of whole seedling samples (Twell et al., 2002; Kawamura et al., 2006) may have led to artefacts since MOR1’s microtubule distribution pattern in these experiments differs from my observations using live-cell 	 100 imaging of MOR1-3xYpet.  My data showing plus-end localization with live-cell imaging should be considered artefact-free since the MOR1pro:MOR1-3xYpet construct was able to completely rescue the mor1-23 homozygous lethality without causing any vegetative or reproductive defects.  It is unknown why MOR1 is shown to localize to the entire microtubule sidewall in the Kawamura et al. (2006) study, especially since the authors provided several experimental data showing the specificity of the MOR1 antibody and accuracy of MOR1’s immunolabelling. The difference in MOR1 localization between live-cells and fixed-samples stresses the potential drawbacks of using chemical fixation and immunolabelling for imaging experiments.  Chemical fixation appears to have altered the plus ends of microtubules so that +TIPs, such as EB1b and MOR1, can no longer bind to the plus ends.  As a result, both of these proteins became cytoplasmically dispersed.  What remains unclear is how chemical fixation alters the plus ends; however, the structure or biochemical properties could potentially be affected since +TIPs use these characteristics to target the plus ends. In this chapter, I was able to use live-cell imaging to visualize MOR1-3xYpet and determine that it is a plus-end tracking protein.  By utilizing previous loss of function data with the mor1-1 mutants (Whittington et al., 2001; Kawamura et al., 2006; Kawamura and Wasteneys, 2008), I was able to conclude that MOR1 functions as microtubule polymerase.  My data also reveals that MOR1 is able to bind to the sidewalls of microtubules in addition to its preferential plus-end binding.  Finally, I demonstrate that chemical fixation can affect the plus-end of microtubules, thus stressing the importance of using live-cell imaging to visualize +TIPs and plus-end dynamics. 	 		 101  Figure 4.1.  The MOR1 gene structure and placement of mutations of various mor1 alleles. The MOR1 gene structure with both exons (vertical bars) and introns (horizontal line).  The different alleles either have point mutations or T-DNA insertions located in the labelled regions.  ATG = protein start codon; TAG = protein stop codon; UTR = untranslated region; C = Cytosine; T = Thymine; G = Guanine; A = Adenine.  Numbers between the nucleotides represent the residue position of the point mutation   14,980 K14,978 K14,976 K14,974 K14,972 K14,970 K14,968 K14,966 KFEI2NM_129116.4NM_001202755.1MOR1NM_129117.4 NP_565811.2UBQ7NM_129118.4 NP_565812.1AT2G35637GenesSNPmor1-23 (SALK_032056) ATG TAG 5’-UTR 3’-UTR mor1-1 (C1626T) mor1-2 (G1689A) gem1-1 (T6369G) 14,980 K14,978 K14,976 K14,974 K14,972 K14,970 K14, 68 14,966 KFEI2NM_129116.4NM_00120275 .1MOR1NM_129117.4 NP_565811.2UBQ7NM_129118.4 NP_565812.1AT2G35637GenesSNP14,980 K14,978 K14,976 K14,974 K14,972 K14,970 K1 ,96  K14,9 6 KFEI2NM_ 291 6.4NM 01202755.1MOR1NM_12 117.4 NP_565811.2UBQ7NM_129118.4 NP_565812.1AT2G35637GenesSNP	 102  Figure 4.2.  The MOR1pro:MOR1-3xYpet construct rescues the mor1-23 homozygous lethality and supports normal vegetative and reproductive development of A. thaliana plants. (A) Seven day-old Col-0 and mor1-23; MOR1pro:MOR1-3xYpet seedlings grown on vertically held Petri dishes.  There appears to be no obvious developmental differences between the lines.  Six plants from each line are shown.  Scale bar represents 1 cm. (B) Nineteen day-old Col-0 and mor1-23; MOR1pro:MOR1-3xYpet plants showing no difference in rosette size.  Two plants from each line are shown.  Scale bar represents 2 cm. A Col-0 mor1-23; MOR1pro:MOR1-3xYpet  B Col-0 mor1-23; MOR1pro:MOR1-3xYpet  CCol-0 mor1-23; MOR1pro:MOR1-3xYpet  DCol-0 mor1-23; MOR1pro:MOR1-3xYpet  	 103 Figure 4.2.  The MOR1pro:MOR1-3xYpet construct rescues the mor1-23 homozygous lethality and supports normal vegetative and reproductive development of A. thaliana plants. (C) Thirty-six day-old Col-0 and mor1-23; MOR1pro:MOR1-3xYpet plants showing no difference in inflorescence stem height. Scale bar represents 5 cm. (D) Fully-grown siliques of Col-0 and mor1-23; MOR1pro:MOR1-3xYpet plants showing no difference in silique size.  Five siliques from each line are shown.  Scale bar represents 1 cm.	 104  Figure 4.3.  MOR1-3xYpet is expressed in all cell types of Arabidopsis seedlings. Spinning-disc confocal images of MOR1pro:MOR1-3xYpet in mor1-23 showing the MOR1 expression pattern. (A) MOR1pro:MOR1-3xYpet in mor1-23 is expressed in all cells of the root tip. The image is a representation of two stitched images that were merged from a stack of z-slices (taken every 0.5 µm).  Scale bar represents 20 µm. (B) MOR1-3xYpet is expressed in the root elongation zone.  The image is a merged stack of z-slices (taken every 0.5 µm).  Scale bar represents 20 µm. (C) A cross-section of the root elongation zone showing MOR1-3xYpet being expressed in epidermal (trichoblasts, atrichoblasts), cortex, and endodermal cells.   The root cross-section was taken from the yellow line in (B).  a = atrichoblast; t = trichoblast; c = cortex cell; e = endodermal cell.  Scale bar represents 20 µm.  Root Tip A B D E F a a c t c a a G PC GC C e 	 105 Figure 4.3.  MOR1-3xYpet is expressed in all cell types of Arabidopsis seedlings. (D) MOR1-3xYpet is expressed in elongating (left) and mature (right) root hairs.  The image is a cortical slice of the root hair.  The image is a merged stack of z-slices (taken every 0.2 µm).  Scale bar represents 10 µm. (E) MOR1-3xYpet is expressed in petiole epidermal cells. The image is a merged stack of z-slices (taken every 0.5 µm).  Scale bar represents 20 µm. (F) MOR1-3xYpet is expressed in etiolated epidermal hypocotyl cells. The image is a merged stack of z-slices (taken every 0.5 µm).  Scale bar represents 20 µm. (G) MOR1-3xYpet is expressed in pavement and guard cells of cotyledons.  GC = guard cells; PC = pavement cells.  The image is a merged stack of z-slices (taken every 0.5 µm).  Scale bar represents 20 µm.   	 106  Figure 4.4.  MOR1-3xYpet localizes to PPB microtubules. Spinning-disc confocal micrographs of MOR1-3xYpet (in cyan), mRFP-TUB6 (in red), and a merged image of both channels.  Arrows label the microtubule bands co-localized with MOR1-3xYpe that surround the nucleus at the mid-zone of the cell.  Arrowheads label microtubules and MOR1-3xYpet that surround the nuclear envelope.  Images are of medial sections from a lateral root cap cell.  Scale bar represents 10 µm.    MOR1-3xYpet mRFP-TUB6 Merged <" <"<" <" <" <"	 107   Figure 4.5.  MOR1-3xYpet localizes to microtubules from the preprophase stage to late anaphase stage. (A) Merged time-lapse spinning-disc confocal micrographs of MOR1-3xYpet (in cyan) and mRFP-TUB6 (in red) being expressed in dividing cells in the root tip.  MOR1-3xYpet localizes to microtubule structures in all stages of mitosis.  Each individual image shows two cells at different stages of mitosis and is either labelled with a white or yellow bracket.  The white bracket labels a cell starting from the pre-prophase to anaphase stage.  Note that MOR1-3xYpet localization occurs near the nuclear envelope during its breakdown followed by localization to the mitotic spindle. The yellow bracket labels a cell that is has progressed further in mitosis from metaphase to late anaphase. Time progresses from left to right.  Images were acquired every 60 seconds.  Scale bar represents 10 µm.   ["["Merged MOR1-3xYpet mRFP-TUB6 A B C 	 108 Figure 4.5.  MOR1-3xYpet localizes to microtubules from the preprophase stage to late anaphase stage. (B) Time-lapse spinning-disc confocal micrographs of MOR1-3xYpet.  The images are of the same cells in (A). (C) Time-lapse spinning disc confocal micrograph of mRFP-TUB6.  The images are of the same cells in (A). 	 109  Figure 4.6.  MOR1-3xYpet localizes to the phragmoplast at microtubule plus ends and in microtubule-free domains. (A) Schematic diagram of a plant cell during late telophase where cytokinesis is occurring.  Note that the microtubule plus ends align at the mid-plane.  MOR1 localizes to the plus ends of the microtubules.  The + and – indicate the plus and minus end of the microtubule, respectively.      ++__Microtubule MOR1 MOR1-3xYpet mRFP-TUB6 Merged 0$20$40$60$80$100$120$0$0.648$1.296$1.944$2.592$3.24$3.888$4.536$5.184$5.832$0$20$40$60$80$100$120$140$160$0$0.648$1.296$1.944$2.592$3.24$3.888$4.536$5.184$5.832$Fluorescent Intensity for White Line Fluorescent Intensity for Yellow Line Distance Along Line Scan (in µm) Distance Along Line Scan (in µm) Pixel Intensity (grey value) Pixel Intensity (grey value) mRFP-TUB6 MOR1-3xYpet ABC1 2 	 110 Figure 4.6.  MOR1-3xYpet localizes to the phragmoplast at microtubule plus ends and in microtubule-free domains. (B) Spinning-disc confocal micrographs of root tip cells expressing MOR1-3xYpet (in cyan), mRFP-TUB6 (in red), and a merged image of both channels.  Images show two different cells at different stages of telophase: a cell with in late telphase (labelled “1”) and a cell in early telophase (labelled “2”).  Note the decrease in mRFP-TUB6 fluorescence intensity at the middle of the phragmoplast (labelled with white arrow), which is the newly formed cell plate that excludes microtubules.  Also, note the increase in MOR1-3xYpet fluorescence intensity at the middle of the phragmoplast (labelled with white arrow).  This suggests an accumulation of MOR1-3xYpet at the plus ends of the microtubules.  Images are of a root tip cell.  Scale bar represents 10 µm. (C) Fluorescence intensity of line scans of the phragmoplasts seen in the merged image panel of (B).  Each graph shows a line scan of either the late phragmoplast (1) with the white line or the early phragmoplast (2) with the yellow line.  Note the decrease in mRFP-TUB6 intensity (black arrow on graph) correlates with the confocal micrograph (white arrow in (B)).  Also, note the increase in MOR1-3xYpet intensity (black arrow on graph) correlates with the confocal micrograph seen in B (white arrow).   	 111  Figure 4.7.  MOR1-3xYpet is asymmetrically distributed on the plus ends of microtubules and is a plus-end tracking protein. (A) A spinning-disc confocal image of an etiolated hypocotyl epidermal cell expressing MOR1-3xYpet (in red) and mRFP-TUB6 (in cyan) in the mor1-23 background.  MOR1-3xYpet has the strongest fluorescence intensity at microtubule plus ends.  Scale bar represents 20 µm. (B) Enlarged images taken from the yellow-boxed area of the cell in (A) showing MOR1-3xYpet and mRFP-TUB6 in separate channels.  A merged image is also shown.  Scale bar represents 5 µm. A B 0 20 40 60 80 100 120 140 1 3 5 7 9 11 13 15 17 19 21 23 Distance from plus-end (in pixels) Pixel Intensity (grey value) MOR1-3xYpet   mRFP-TUB6 C D 0 s 5 s 10 s 15 s 20 s 25 s 30 s 35 s 55 s 50 s 45 s 40 s 60 s 65 s 70 s 75 s MOR1-3xYpet mRFP-TUB6 Merge 	 112 Figure 4.7.  MOR1-3xYpet is asymmetrically distributed on the plus ends of microtubules and is a plus-end tracking protein. (C) Graph showing the fluorescence intensity of line scans taken at the microtubule plus ends.  MOR1-3xYpet is quantitatively shown to have its highest accumulation on the microtubule plus end with lower accumulations at the microtubule side wall behind the plus end.  As a control, the intensity of mRFP-TUB6 was also measured and remained constant.  The intensity of both MOR1-3xYpet and mRFP-TUB6 were measured by a line scan similar to the dotted line in the merged image in (B) (except with the line superimposed on the microtubule). 12 line scans (of 25 pixels in length) of microtubule plus ends were measured and averaged out.  (D) Time-lapse montage of spinning-disc confocal images showing MOR1-3xYpet’s (in red) ability to plus-end track with growing microtubules (in cyan). The yellow circles indicate microtubule plus ends. Images were acquired every 5 seconds.  Scale bar represents 10 µm.   	 113  Figure 4.8.  MOR1-3xYpet localizes to depolymerizing microtubule plus ends. Time-lapse montage of spinning-disc confocal images showing MOR1-3xYpet (in red) localizing to the plus ends of depolymerizing microtubules (in cyan) (in the mor1-23 background).   Merged images of both channels are also shown.  Arrowheads indicate two depolymerizing microtubules.  Images are of etiolated hypocotyls and were acquired every 5 seconds with time progressing from left to right.  Scale bar represents 5 µm. MOR1-3xYpet mRFP-TUB6 Merge > > > > > > > > > > > > > > > > > > > > > > > > 	 114 > > > > >>>> > >> > >>> > >>> > > >> > >> > >> > >>	 115 Figure 4.9.  MOR1-3xYpet persists on microtubule plus ends undergoing catastrophe and then rescue. A time-lapse montage of spinning-disc confocal images taken of MOR1-3xYpet (in red) and mRFP-TUB6 (in cyan) in the mor1-23 mutant background.  MOR1-3xYpet labels microtubules during polymerization (labelled with arrowhead) and remains bound to microtubule plus ends during catastrophe (indicated by the arrow).  Time progresses from left to right.  Images were taken from etiolated hypocotyls every 5 seconds.  Scale bar represents 5 µm.   	 116  Figure 4.10.  MOR1-3xYpet is not found on microtubule minus ends. Time-lapse montage of spinning-disc confocal images of MOR1-3xYpet (in red) and mRFP-TUB6 (in cyan) showing a treadmilling microtubule.  A merged image of both channels is also shown.  The polymerizing plus end is labelled with MOR1-3xYpet (indicated by the arrowhead).  The depolymerizing minus end of the same microtubule has no MOR1-3xYpet signal (indicated by the arrow).  Time progresses from left to right with images being acquired every 5 seconds.  Images are taken of etiolated hypocotyls in the mor1-23 background.  Scale bar represents 5 µm.   MOR1-3xYpet mRFP-TUB6 Merge > > >>>> >> >>>> >> > >>>>> >>>> >> >	 117  Figure 4.11.  MOR1-3xYpet localizes to newly created plus ends from microtubule severing events at cross-over sites.  (A) Time-lapse montage of spinning-disc confocal images of MOR1-3xYpet (in red), mRFP-TUB6 (in cyan), and merged images. MOR1-3xYpet labels the growing microtubule plus end (labelled with white circle).  The polymerizing microtubule eventually crosses over a pre-existing microtubule between t = 25 s – 30 s.  At t = 30, the microtubule is severed (indicated by white arrowhead), and a new plus-end with MOR1-3xYpet is created that undergoes depolymerization (labelled with a yellow circle).  Images are of etiolated hypocotyls acquired every 5 seconds.  Scale bar represents 5 µm.        +"_"+"_">+"_"+"_"+"t1 +"t4 t3 t2 >"20 s 10 s 5 s 0 s 15 s 25 s 30 s 35 s 40 s 45 s MOR1-3xYpet mRFP-TUB6 Merge A B 	 118 Figure 4.11.  MOR1-3xYpet localizes to newly created plus ends from microtubule severing events at cross-over sites.  (B) Schematic diagram of a polymerizing microtubule crossing over a pre-existing microtubule followed by microtubule severing and depolymerization events.  A polymerizing microtubule (in red) crosses over a pre-existing microtubule (in blue)  (from t1 - t2).  Once the red microtubule has crossed over (indicated by arrowhead), the red microtubule is severed, creating a new plus end that undergoes depolymerization.  MOR1 (indicated by yellow circle) binds to the new plus end that is depolymerizing (in t3 – t4). The + and – indicate the plus and minus end, respectively.   	 119  Figure 4.12.  MOR1-3xYpet has preferential binding to microtubule plus ends but still has affinity for the microtubule sidewalls.  (A) Confocal micrographs of MOR1-3xYpet in wild-type and mor1-23 backgrounds. MOR1-3xYpet is more tightly localized to the tip in the mor1-23 background while having a great distribution along the microtubule sidewall in the wild-type background.  Images are of etiolated hypocotyls.  Scale bars represent 10 µm.  (B) Bar graph showing the mean length of MOR1-3xYpet on microtubules in wild-type and mor1-23 backgrounds.  The length of MOR1-3xYpet fluorescence was measured in each background.  MOR1-3xYpet has significantly less distribution along the microtubule in the mor1-23 background (n = 150 microtubules) relative to the wild-type (n = 125 microtubules) background. Data and bars are represented as mean ± SD, respectively. Asterisk shows a significant difference using a two-sample t-test with unequal variance (P<10.7x10-61).  10 cells from four different plants in each genetic background were sampled.   MOR1pro:MOR1-3xYpet in mor1-23 background  MOR1pro:MOR1-3xYpet in Wild-type background  A 0 2 4 6 8 10 12 14 B Wild-type background  mor1-23 background  MOR1-3xYpet length along microtubule  (in µm) * 	 120  Figure 4.13.  Increased levels of MOR1 protein do not affect development of seedlings.  Increased levels of MOR1 in the wild-type (Col-0) line expressing MOR1-3xYpet (right) do not affect seedling development relative to the mor1-23 (left).  Six day-old seedlings were grown vertically on bacto-agar. Scale bar represents 1 cm.   mor1-23; MOR1pro:MOR1-3xYpet 35Spro:mRFP-TUB6 Col-0; MOR1pro:MOR1-3xYpet 	 121  Figure 4.14.  Chemical fixation of hypocotyls alters the plus ends of microtubules  (A) Confocal micrographs of live (unfixed control) seedlings expressing both MOR1-3xYpet (in red), mRFP-TUB6 (in cyan) in the mor1-23 background.  MOR1-3xYpet is plus-end localized in live cells as previously described.  Compare to (C).      A Merge mRFP-TUB6 MOR1-3xYpet MOR1-3xYpet EB1b-mCherry Merge MOR1-3xYpet mRFP-TUB6 Merge C MOR1-3xYpet EB1b-mCherry Merge B D 	 122 Figure 4.14.  Chemical fixation of hypocotyls alters the plus ends of microtubules  (B) Confocal micrographs of live (unfixed control) seedlings expressing both MOR1-3xYpet (in red), EB1b-mCherry (in cyan) in the wild-type background.  MOR1-3xYpet is localized along the microtubule sidewall as previously described and EB1b-mCherry is plus-end localized.  Compare to (D).  (C) Confocal micrographs of chemically fixed seedlings expressing both MOR1-3xYpet (in red), mRFP-TUB6 (in cyan) in the mor1-23 background. MOR1-3xYpet is no longer localized to microtubules and instead becomes cytoplasmic (compare to the unfixed control in (A)).  (D) Confocal micrographs of chemically fixed seedlings expressing both MOR1-3xYpet (in red), EB1b-mCherry (in cyan) in the wild-type background. EB1b-mCherry becomes cytoplasmic as it is no longer plus-end localized due to the chemical fixation.  MOR1-3xYpet appears unaffected by the chemical fixation and remains bound to the microtubule sidewall much like seen in (B).  Scale bars represent 20 µm.   	 123 Chapter 5: Conclusions 5.1. Major Findings of the Dissertation  Microtubules play an essential role in the growth and morphogenesis of plants. Thus, it is important to understand how the dynamics and organization of microtubules are orchestrated by the activities of microtubule-associated proteins and kinases.  Within this dissertation, I describe studies on a variety of factors that control microtubule dynamics in the model system, Arabidopsis thaliana.  Specifically, I looked at the ARK1 kinesin motor protein (Chapter 2), the NEK6 kinase (Chapter 3), and the MOR1 microtubule-associated protein (Chapter 4), which each play different but potentially interrelated roles in microtubule dynamics and cell morphogenesis. 5.1.1.  ARK1 maintains proper root hair tip growth by promoting microtubule catastrophe  In Chapter 2, I characterized the function of the kinesin, ARK1, and determined the importance of its role in microtubule dynamics and tip growth in root hairs.  Prior to my work, ARK1 was known to play a role in root hair morphogenesis based on the fact that ark1 mutants had root hairs with loss of polarity (Jones et al., 2006; Sakai et al., 2008).  Phenotypically, ark1 mutants showed an abnormal abundance of endoplasmic microtubules (Sakai et al., 2008) and thus, it was hypothesized that ARK1 promotes the depolymerization (catastrophe) of microtubules.  In order to test this hypothesis, I first measured the incidence of microtubule catastrophe in root hairs using spinning-disc confocal microscopy and found that catastrophe frequency was significantly reduced in the ark1-1 mutant, thus confirming the hypothesis.  Interestingly, I also found that ark1-1 mutants had reduced plus-end growth velocities relative to wild-type root hairs.  By applying small concentrations of the microtubule-destabilizing drug oryzalin, it was possible to restore normal microtubule growth velocities, which indicates that the reduced microtubule growth velocity in ark1-1 results from reduced free tubulin turnover, consistent with the decrease in catastrophe events. Importantly, this treatment also largely rescued the crooked and branched root hair phenotype of ark1-1, which suggests that microtubule polymerization rate is a critical factor in maintaining tip growth.  	 124  To directly test the hypothesis that ARK1 is a microtubule catastrophe factor, I overexpressed the ARK1 gene. It is important to note here that attempts to stably transform Arabidopsis with ARK1 overexpression constructs failed, supporting the idea that the activity of a catastrophe factor must be finely balanced. Nevertheless, it was possible to use an Agrobacterium-mediated transient expression system to overexpress ARK1 in leaf pavement cells and this clearly demonstrated an increase in microtubule catastrophe events in cells overexpressing ARK1-RFP relative to cells with endogenous ARK1 expression levels.  Taking together the results of the loss of function and gain of function analysis, I was able to conclude that ARK1 does in fact promote the periodic depolymerization of microtubules and that this is required to maintain uniform tip growth in root hairs. In addition to the genetic analysis, I used cloning strategies to construct an ARK1-GFP translational reporter driven by its native promoter.   Visualization of the reporter protein using spinning-disc confocal microscopy revealed that ARK1 is exclusively a plus-end tracking protein (+TIP) that only associates with growing microtubules.  The specific localization of ARK1 to the growing plus end of microtubules is consistent with its activity as a plus-end acting catastrophe factor and suggests a simple mechanism whereby ARK1 accumulates at the plus end to induce catastrophe before dissociating from the microtubule. To gain insight into the function of the C-terminal “cargo”domain of ARK1, I generated reporter constructs that truncated the ARM domain or expressed it without the motor and coiled-coil domains. This strategy enabled me to determine that the ARM domain of ARK1 does not play an essential role in ARK1’s function as a catastrophe factor.  The ARK1 ΔARM-GFP construct driven by the ARK1 promoter was shown to rescue the ark1-1 root hair phenotype and maintain its ability to track microtubule plus ends in the same manner as the full length ARK1-GFP.  The ARM-GFP construct was also localized to the microtubule plus ends, suggesting that it may confer an additional microtubule interaction domain. Its expression, however, did not rescue the ark1-1 root hair phenotype. Taken together, these truncation experiments suggest that the motor domain can induce microtubule catastrophe without the ARM domain and that the ARM domain’s function remains unclear.  Finally, my translational reporter analysis confirmed that that ARK1 is broadly expressed throughout many cell types and cell-cycle stages in A. thaliana and not specific to root hair development as was initially indicated by the study that identified ARK1 on the 	 125 basis of being highly expressed in developing root hairs (Jones et al., 2006). Moreover, ARK1-GFP is also expressed throughout cellular development and associates with the PPB, mitotic spindle, phragmoplast, and interphase cortical microtubules, suggesting that it might also function in turnover of microtubules during array transitions as cells progress through mitosis and cytokinesis.  Whether ARK1 acts as a catastrophe factor in dividing cells or in the elongation zone remains uncertain because the ark1-1 phenotype is confined to root hairs. I hypothesized that the ark1-1’s lack of phenotype beyond root hairs results from functional redundancy with two closely related proteins, ARK2 and ARK3, which have overlapping expression with ARK1, except in developing root hairs (Sakai et al., 2008).  ARK3, for example, is localized to the PPBs in leaf epidermal cells and has a role in asymmetrical cell division during stomatal development (Lau et al., 2014). The ARK1/ARK2/ARK3 kinesin family have, to date, remained uncategorized phylogenetically in relation of the 13 kinesin families described so far (Reddy and Day, 2011; Zhu and Dixit, 2012).  One of the significant aspects of establishing the function of ARK1 is that now, the function of ARK2 and ARK3 can be predicted to have a similar role in promoting microtubule turnover. ARK1 is also one of only a handful of kinesins (inside and outside the plant kingdom) known to function as microtubule catastrophe factors. The apparent restriction of ARKs to plant lineages may help to map the functional diversification of kinesins throughout eukaryotic evolution. 5.1.2.  ARK1 and NEK6 affect microtubule dynamics and cell expansion through independent pathways  In Chapter 3, I was able to provide evidence that there is no NEK6 and ARK1 interaction and that these two proteins function independently in controlling microtubules and cell expansion under standard conditions.  Using non-in vivo techniques, a previous study revealed ARK1’s interaction with the kinase, NEK6 (Sakai et al., 2008).  Despite both proteins showing microtubule-localization in vivo (Chapter 2; Eng and Wasteneys, 2014; Motose et al., 2008, 2011), the function and physical interaction of ARK1-NEK6 remained unclear.  In the work described in Chapter 3, I used confocal microscopy to visualize ARK1 and NEK6 fluorescently tagged proteins in the nek6 and ark1 mutant backgrounds, respectively.  I was able to question if either protein depends on the other for its association with microtubules.  In the mutant backgrounds, the distribution and dynamics of both ARK1 	 126 and NEK6 remained unchanged relative to the wild-type background.  This suggests that NEK6 and ARK1 associate to microtubules independently.  However, as previously mentioned, the functional redundancy of ARK2 and ARK3 might compensate for the loss of ARK1 and could potentially explain why we see no differences with NEK6-GFP in the ark1-1 single mutant background.  This problem will be addressed further in 5.2.2. Because ARK1-GFP function remains unchanged in the nek6-1 background, I concluded that ARK1 does not require phosphorylation by NEK6 for proper function.  I next hypothesized that the NEK6 and ARK1 interaction is hormone-dependent.  This was based on previous data showing changes in the number of epidermal protrusions seen in nek6 mutants when exposed to specific hormones, ACC (an ethylene precursor) and GA.  However, exogenous exposure to ACC and GA revealed no changes in NEK6 and ARK1 microtubule distribution patterns in both wild-type and mutant backgrounds. Mutant analysis between nek6-1 ark1-1 double mutants and nek6-1 single mutants showed no significant difference in the number of hypocotyl epidermal protrusions while protrusions in ark1-1 were non-existent.  These observations suggest that NEK6 plays an independent role in microtubule dynamics and cell expansion in the hypocotyls.  My data showing how overexpression of the ARM domain caused a complete knock out of NEK6-GFP gene expression, however, suggests that ARK1 may play a regulatory role in NEK6 expression. This intriguing finding also provides a possible lead to understanding the function of the ARM cargo domain of the ARK proteins, especially since it evidently is not required for the catastrophe-inducing activity.  5.1.3.  MOR1 is a plus-end tracking protein  Chapter 4 focused on characterizing the MOR1 protein using live-cell imaging of MOR1 fused to a Ypet fluorescent protein tag.  Prior to my research, creating a MOR1 fusion protein was difficult due to cloning limitations.  Fortunately, the recombineering technology successful generated a MOR1 reporter tagged with a fluorescent protein, which finally allowed for live-cell imaging of full length and fully functional MOR1.  Using spinning-disc confocal microscopy, I was able to confirm that MOR1 is a plus-end tracking protein (+TIP) and that it associates with both the growing and shrinking plus ends.  MOR1 also rapidly localized to newly created plus ends, which are the consequence of microtubule severing events at microtubule cross-over sites.  My observations of MOR1 plus-end 	 127 microtubule localization is consistent with the hypothesis that MOR1 promotes the polymerization of microtubules at the plus ends (Kawamura and Wasteneys, 2008).  I also confirmed that MOR1 plays an important role in modulating microtubule dynamics and structures throughout all cells in M-phase and interphase. Live-cell imaging of MOR1 also allowed me to observe the effects of chemical fixation methods and the artefacts that arise from the process.  My data revealed that the chemical fixation method utilized in most microtubule-based immunofluorescence experiments can affect microtubule plus ends, causing the +TIPs to dissociate from microtubules.  Finally, my data demonstrated that MOR1 preferentially binds microtubule plus ends.  Increasing MOR1 protein levels led to an increase in microtubule sidewall binding but not plus-end binding and did not disrupt the microtubule-dependent process of cell morphogenesis.  This finding suggests that MOR1 has a specific role in driving microtubule dynamics at the plus ends but not the sidewall/lattice region.  5.2. Future Directions 5.2.1. Elucidating the mechanism of ARK1 action  Although my data shows that ARK1 promotes microtubule depolymerization in vivo, there are still outstanding questions about the mechanism by which ARK1 achieves this.  It is unknown whether or how ARK1 utilizes its ability to hydrolyze ATP to perform its microtubule catastrophe-inducing function.  Previous studies on the depolymerizing kinesin, Kip3, have shown that ATP hydrolysis is responsible for both kinesin motility and for inducing microtubule catastrophe (Gupta et al., 2006; Su et al., 2011; Varga et al., 2006).  To address the function of ARK1’s ATP-dependent mechanism, in vitro assays should be conducted using high-resolution fluorescence microscopy (e.g. near-Total Internal Reflection Fluorescence (TIRF) microscopy) to track ARK1-fluorescent protein on microtubules polymerized from fluorescently-tagged tubulins, in the presence and absence of ATP.  This experimental set-up would allow me to address two hypotheses. First, to determine if ARK1 uses ATP to induce catastrophe, the frequency of catastrophe could be measured in conditions with and without ATP by visualizing the in vitro system with TIRF microscopy.  If microtubule depolymerization occurs more frequently with the addition of ATP, then it can be concluded that ARK1 uses ATP to induce catastrophe.  Second, to determine if ARK1 	 128 uses ATP for motility along the microtubule lattice and/or for plus-end tracking, we can observe ARK1 movement along microtubule lattice and plus ends (or lack thereof) with and without ATP.  Loss of ARK1 movement on microtubules without ATP would suggest that ARK1 utilizes ATP for motility.  If movement along the microtubule lattice still occurs without ATP, then it can be predicted that this movement is a result of ATP-independent diffusion along the lattice, which has been demonstrated for the microtubule depolymerizing kinesin, MCAK (Helenius et al., 2006). 5.2.2. Confirming a physical interaction between NEK6 and ARK1.  Despite previous research showing a physical interaction between NEK6 and ARK1 (Sakai et al., 2008), I was unable to confirm any interaction in my study.  It remains possible, however, that these proteins do interact and additional experiments could be conducted to specifically determine this. First, a reciprocal yeast two-hybrid assay should be performed using full-length ARK1 as the prey and full length NEK6 as the bait.  Previous experiments only used the ARM domain of ARK1 as the bait (Sakai et al., 2008).  Second, this interaction can be tested in vivo using bimolecular fluorescent complementation (BiFC) in Arabidopsis. Not only will these results elucidate whether the two proteins interact, but it will also show where on the microtubule the proteins interact.  Finally, a more sensitive co-immunoprecipitation experiment should be done to obtain enough ARK1-GFP protein followed by proteomic analysis of potential proteins pulled down with ARK1-GFP.  Proteomic analysis should potentially recover potential ARK1 interactors (such as NEK6).  Finally, the observation that the NEK6-GFP localization to microtubules is not altered in ark1-1 mutants could be explained by ARK1’s possible functional redundancy with ARK2 and ARK3.  To address this, NEK6-GFP should be observed in an ark1 ark2 ark3 triple mutant background to rule out or determine NEK6-GFP’s functional dependency on ARK1, ARK2, and/or ARK3. 5.2.3. Is the NEK6-ARK1 interaction only manifested under specific stress conditions? Although previous unpublished data has shown ARK1’s ability to be phosphorylated by NEK6 (or NEK4 or NEK5) in vitro (Dr. H Motose, personal communication), it is currently unknown if ARK1 is a target of NEK6 in vivo. Lack of evidence for ARK1-NEK6 phosphorylation interaction under normal conditions in vivo still leaves open the possibility 	 129 that the phosphorylation of ARK1 can occur under specific conditions.  Although I failed to notice any changes in ARK1-GFP/TagRFP and NEK6-GFP localization and function upon exposure to exogenous ACC and GA, which exacerbate and mitigate the nek6 mutant phenotype respectively, many other conditions remain to be tested. Previous analysis has shown NEK6 is involved in salt and osmotic stress so these stresses would be obvious areas to explore. When plants were exposed to salt and osmotic stress, NEK6 expression levels increased and plants modified to overexpress NEK6 had increased tolerance to these stresses (Zhang et al., 2011).  Phosphorylation activity in some kinases can be induced under specific stress conditions. For example, PHS1, which has both MAP kinase phosphatase (Walia et al., 2009) and kinase capabilities, is activated during hyperosmotic stress and subsequently phosphorylates α-tubulin to regulate microtubule dynamics (Fujita et al., 2013).  In a similar manner, NEK6 might phosphorylate ARK1 and/or tubulin in order to modulate microtubule function under specific stress conditions.  Previous analysis has shown NEK6’s ability to phosphorylate β-tubulin in vitro (Motose et al., 2011) much like is seen with NEK6 being able to phosphorylate ARK1 (Dr. H Motose, personal communication).  Based on these findings and my own data, I can hypothesize that ARK1 phosphorylation by NEK6 only occurs during stress as a way to modulate ARK1 function. To determine if ARK1 phosphorylation by NEK6 is dependent on salt and osmotic stress conditions, ARK1-GFP proteins should be purified from plants under stress and non-stress conditions followed by analysis of its phosphorylation status using an electrophoretic technique such as an SDS- polyacrylamide gel (SDS-PAGE) with a Phos-tag (Kinoshita, 2006). SDS-PAGE with a Phos-tag is used for elucidating the phosphorylation status of proteins by causing a mobility shift of phosphorylated proteins, which is subsequently visualized as slower migrating bands relative to dephosphorylated proteins on the gel.  Thus, this technique should reveal if ARK1-GFP proteins are in fact phosphorylated.  Finally, to confirm if ARK1 phosphorylation occurs specifically by one of the NEKs and not any other kinase, the same experimental set up should be performed with ARK1-GFP proteins extracted from the nek4 nek5 nek6 triple mutant line.  If ARK1-GFP extracted from the triple mutant is no longer phosphorylated relative to the wild-type background, then it can be concluded that ARK1 is indeed a specific target of one or all of the NEKs. 	 130 While the previous experiments would test if ARK1 is a phosphorylation target of NEK6 (through a transient physical interaction) during specific stress conditions, further work would also need to be done to determine if a more stable ARK1-NEK6 physical interaction occurs under specific stress conditions.  To test if this, visualization of both proteins tagged with a fluorescent reporter should be done under specific stress conditions to see if the ARK1-NEK6 association with each other or with the microtubules becomes altered.  Similarly, the same assays used to determine in vivo physical interactions in 5.2.2 should be done using the same stress conditions as previously mentioned. 5.2.4. Determining the function of ARK1 phosphorylation.  Finally, the purpose of ARK1 phosphorylation should be elucidated (i.e. how does the phosphorylation of ARK1 affect its function?).  Kinesin phosphorylation has been shown to promote the association/dissociation of its cargo as well as to modulate a kinesin’s ability to bind to microtubules (Hirokawa et al., 2009).  In the case of ARK1, which promotes microtubule catastrophe (Eng and Wasteneys, 2014), phosphorylation under stress conditions could enhance or inhibit ARK1’s effects on microtubule dynamics.  Hypothetically, NEK6 phosphorylation of ARK1 could affect the interaction between ARK1 and its association with NEK6 itself, a microtubule plus-end, or another unknown protein.  Using the assumption that ARK1 is capable of being phosphorylated (see previous section, 5.2.3), then the next objective would be to determine if phosphorylation affects the localization and/or function of ARK1.  To address this, putative ARK1 phosphorylation sites should be identified using phospho-proteomic programs that are able to predict putative phosphorylation sites of proteins.  Once putative phosphorylation sites have been identified, site-directed mutagenesis of these sites that make up the ARK1-GFP proteins should be performed.  By altering the putative phosphorylation sites, the phosphorylation of ARK1-GFP at these specific sites should be inhibited.  Upon expressing the mutagenized ARK1-GFP proteins in Arabidopsis, a pull-down of the ARK1-GFP proteins should be performed and its phosphorylation status confirmed using SDS-PAGE with a Phos-tag.  The same mutagenized ARK1-GFP proteins can then be observed using live-cell imaging with confocal microscopy to see if altered ARK1-GFP function and/or localization is a direct consequence of the absence of phosphorylation at the mutagenized sites. 	 131 5.2.5. Do the +TIPs interact to modulate plus-end microtubule dynamics?  Because MOR1 and ARK1 are both plus-end tracking proteins, it could be possible that these two +TIPs work together to control dynamics at the microtubule plus ends.  The idea that +TIPs are functionally dependent on each other to regulate microtubules is not a novel one.  Previous research has shown that XMAP215 works synergistically with EB1 to increase microtubule growth rates and regulate mitotic spindle assembly, albeit through two separate, non-redundant mechanisms (Kronja et al., 2009; Zanic et al., 2013).  Furthermore, XMAP215 localization to microtubule plus ends has been shown to be dependent on EB1 (Kronja et al., 2009).  XMAP215 and the microtubule catastrophe-inducing kinesin, XKCM1, are also known to control microtubule dynamics through antagonistic mechanisms (Tournebize et al., 2000).  Specifically, XMAP215 was found to promote microtubule stabilization while XKCM1 was found to act as a microtubule destabilizer.  Since MOR1 promotes microtubule polymerization (like XMAP215) and ARK1 is a microtubule catastrophe-factor (like XKCM1), it is possible that MOR1 and ARK1 may also function antagonistically to control microtubule plus-end dynamics.   In order to determine if MOR1 and ARK1 work together to regulate the plus-end dynamics (specifically through an antagonistic relationship), an in vitro system should be used to see how each protein functions with the absence of other microtubule-associated proteins that is normally found with in vivo systems.  By removing other MAPs that contribute to microtubule dynamics in vivo, MOR1 and ARK1’s exact function can be more accurately determined.  To accomplish this, TIRF microscopy should be used to visualize microtubule dynamics with fluorescently-tagged tubulins as well as purified MOR1 and ARK1 proteins.  By analyzing how each individual protein affects microtubule dynamics, the exact functions of MOR1 and ARK1 can be determined.  For example, given that MOR1’s predicted function is to promote microtubule polymerization in vivo, then the addition of MOR1 to the in vitro system should increase microtubule polymerization rates.  The subsequent addition of ARK1 to the system should act as an agonist to MOR1 and thus, lead to an increase in frequency of microtubule depolymerization. High-resolution imaging with TIRF microscopy has previously been used to determine if specific +TIPs bind to distinct or the same regions of the microtubule plus-end.  For example, the +TIPs, ch-TOG and EB1, were found to bind to distinct regions of the 	 132 microtubule plus ends in human HeLa cells.  Furthermore, that study concluded that both proteins function independently from each and thus were determined to contribute to microtubule dynamics through different mechanisms (Nakamura et al., 2012).  In order to determine if MOR1 and ARK1 are dependent on each other for function in vivo, near-TIRF microscopy should be used to visualize MOR1-3xYpet and ARK1-RFP to determine the nanoscale distribution of both proteins on microtubule plus ends.  If both proteins have high co-localization on microtubule plus ends, then one prediction would be that the proteins are potentially dependent on each other for function.  Conversely, if both proteins are shown to bind to distinct regions of the microtubule, like with ch-TOG and EB1 (Nakamura et al., 2012), then it could be hypothesized that MOR1 and ARK1 function independently from each another to control microtubule dynamics.  Finally, the MOR1 and ARK1 interaction could be further elucidated by utilizing a mor1-23; MOR1pro:MOR1-3xYpet line crossed into an ark1-1 mutant expressing EB1bpro:EB1b-mCherry.  By using the EB1b-mCherry as a plus-end microtubule marker, any changes in MOR1-3xYpet plus-end localization without the presence of ARK1 could be visualized with TIRF microscopy.  As a control, the same mor1-23; MOR1pro:MOR1-3xYpet EB1bpro:EB1b-mCherry transgenic line (but azygous for ark1-1) should be used to see where MOR1-3xYpet normally localizes to the plus end when ARK1 is present.  Moreover, this experimental control can also reveal if MOR1 and EB1b bind to distinct regions of the microtubule plus end, much their corresponding homologues in humans (Nakamura et al., 2012). If the TIRF data reveals high co-localization between MOR1 and ARK1, then an additional hypothesis would be that both proteins physically interact with each other for function.  To discover a potential physical interaction between ARK1 and MOR1, a yeast two-hybrid assay should be done followed by in vivo experiments using BiFC.  Finally, co-immunoprecipitation should be done by pulling down either the MOR1-3xYpet or ARK1-GFP proteins to see if the proteins interact with each other (or any other novel or known proteins).  Performing co-immunoprecipitation might also identify new proteins that interact with MOR1, ARK1, and NEK4/5/6.  Given that MOR1, ARK1, and the NEKs appear to have separate functions, there may be other uncharacterized proteins with novel and diverse functions involved in the precise control of microtubule dynamics and organization.  	 133 Works Cited Agueci, F., Rutten, T., Demidov, D., and Houben, A. (2012). Arabidopsis AtNek2 Kinase is Essential and Associates with Microtubules. Plant Mol. Biol. Report. 30: 339–348. Akhmanova, A. and Steinmetz, M.O. (2008). Tracking the ends: a dynamic protein network controls the fate of microtubule tips. Nat. Rev. Mol. Cell Biol. 9: 309–322. Allard, J.F., Ambrose, J.C., Wasteneys, G.O., and Cytrynbaum, E.N. (2010). A Mechanochemical Model Explains Interactions between Cortical Microtubules in Plants. Biophys. J. 99: 1082–1090. Allen, C. and Borisy, G.G. (1974). Structural polarity and directional growth of microtubules of Chlamydomonas flagella. J. Mol. Biol. 90: 381–402. Alonso, J.M. and Stepanova, A.N. (2014). Arabidopsis transformation with large bacterial artificial chromosomes. Methods Mol. Biol. Clifton NJ 1062: 271–283. Ambrose, J.C. and Cyr, R. (2007). The Kinesin ATK5 Functions in Early Spindle Assembly in Arabidopsis. Plant Cell Online 19: 226–236. Ambrose, J.C., Li, W., Marcus, A., Ma, H., and Cyr, R. (2005). A Minus-End–directed Kinesin with Plus-End Tracking Protein Activity Is Involved in Spindle Morphogenesis. Mol. Biol. Cell 16: 1584–1592. Ambrose, J.C., Shoji, T., Kotzer, A.M., Pighin, J.A., and Wasteneys, G.O. (2007). The Arabidopsis CLASP Gene Encodes a Microtubule-Associated Protein Involved in Cell Expansion and Division. Plant Cell 19: 2763–2775. Amos, L.A. and Klug, A. (1974). Arrangement of Subunits in Flagellar Microtubules. J. Cell Sci. 14: 523–549. Bannigan, A., Scheible, W.-R., Lukowitz, W., Fagerstrom, C., Wadsworth, P., Somerville, C., and Baskin, T.I. (2007). A conserved role for kinesin-5 in plant mitosis. J. Cell Sci. 120: 2819–2827. Ban, Y., Kobayashi, Y., Hara, T., Hamada, T., Hashimoto, T., Takeda, S., and Hattori, T. (2013). α-Tubulin is Rapidly Phosphorylated in Response to Hyperosmotic Stress in Rice and Arabidopsis. Plant Cell Physiol. 54: 848–858. Baskin, T.I., Wilson, J.E., Cork, A., and Williamson, R.E. (1994). Morphology and Microtubule Organization in Arabidopsis Roots Exposed to Oryzalin or Taxol. Plant Cell Physiol. 35: 935–942. Al-Bassam, J. and Chang, F. (2011). Regulation of microtubule dynamics by TOG-domain proteins XMAP215/Dis1 and CLASP. Trends Cell Biol. 21: 604–614. 	 134 Al-Bassam, J., Kim, H., Flor-Parra, I., Lal, N., Velji, H., and Chang, F. (2012). Fission yeast Alp14 is a dose-dependent plus end–tracking microtubule polymerase. Mol. Biol. Cell 23: 2878–2890. Al-Bassam, J., Larsen, N.A., Hyman, A.A., and Harrison, S.C. (2007). Crystal Structure of a TOG Domain: Conserved Features of XMAP215/Dis1-Family TOG Domains and Implications for Tubulin Binding. Structure 15: 355–362. Beck, M., Komis, G., Müller, J., Menzel, D., and Šamaj, J. (2010). Arabidopsis Homologs of Nucleus- and Phragmoplast-Localized Kinase 2 and 3 and Mitogen-Activated Protein Kinase 4 Are Essential for Microtubule Organization. Plant Cell Online 22: 755–771. Beck, M., Komis, G., Ziemann, A., Menzel, D., and Šamaj, J. (2011). Mitogen-activated protein kinase 4 is involved in the regulation of mitotic and cytokinetic microtubule transitions in Arabidopsis thaliana. New Phytol. 189: 1069–1083. Bergler, J. and Hoth, S. (2011). Plant U-box armadillo repeat proteins AtPUB18 and AtPUB19 are involved in salt inhibition of germination in Arabidopsis: PUB-ARM proteins in stress response. Plant Biol. 13: 725–730. Bibikova, T.N., Blancaflor, E.B., and Gilroy, S. (1999). Microtubules regulate tip growth and orientation in root hairs ofArabidopsis thaliana. Plant J. 17: 657–665. Bradley, B.A. and Quarmby, L.M. (2005). A NIMA-related kinase, Cnk2p, regulates both flagellar length and cell size in Chlamydomonas. J. Cell Sci. 118: 3317–3326. Brady, S.T. (1985). A novel brain ATPase with properties expected for the fast axonal transport motor. Nature 317: 73–75. Breugel, M. van, Drechsel, D., and Hyman, A. (2003). Stu2p, the budding yeast member of the conserved Dis1/XMAP215 family of microtubule-associated proteins is a plus end–binding microtubule destabilizer. J. Cell Biol. 161: 359–369. Brouhard, G.J., Stear, J.H., Noetzel, T.L., Al-Bassam, J., Kinoshita, K., Harrison, S.C., Howard, J., and Hyman, A.A. (2008). XMAP215 Is a Processive Microtubule Polymerase. Cell 132: 79–88. Bruaene, N.V., Joss, G., and Oostveldt, P.V. (2004). Reorganization and in Vivo Dynamics of Microtubules during Arabidopsis Root Hair Development. Plant Physiol. 136: 3905–3919. Caplow, M., Ruhlen, R.L., and Shanks, J. (1994). The free energy for hydrolysis of a microtubule-bound nucleotide triphosphate is near zero: all of the free energy for hydrolysis is stored in the microtubule lattice. J. Cell Biol. 127: 779–788. Carvalho, P., Tirnauer, J.S., and Pellman, D. (2003). Surfing on microtubule ends. Trends Cell Biol. 13: 229–237. 	 135 Chan, J., Calder, G.M., Doonan, J.H., and Lloyd, C.W. (2003). EB1 reveals mobile microtubule nucleation sites in Arabidopsis. Nat. Cell Biol. 5: 967–971. Clough, S.J. and Bent, A.F. (1998). Floral dip: a simplified method forAgrobacterium-mediated transformation ofArabidopsis thaliana. Plant J. 16: 735–743. Cloutier, M., Vigneault, F., Lachance, D., and Séguin, A. (2005). Characterization of a poplar NIMA-related kinase PNek1 and its potential role in meristematic activity. FEBS Lett. 579: 4659–4665. Coates, J.C., Laplaze, L., and Haseloff, J. (2006). Armadillo-related proteins promote lateral root development in Arabidopsis. Proc. Natl. Acad. Sci. 103: 1621–1626. Cottingham, F.R. and Hoyt, M.A. (1997). Mitotic Spindle Positioning in Saccharomyces cerevisiae Is Accomplished by Antagonistically Acting Microtubule Motor Proteins. J. Cell Biol. 138: 1041–1053. Cullen, C.F., Deák, P., Glover, D.M., and Ohkura, H. (1999). mini spindles A Gene Encoding a Conserved Microtubule-Associated Protein Required for the Integrity of the Mitotic Spindle in Drosophila. J. Cell Biol. 146: 1005–1018. Currie, J.D., Stewman, S., Schimizzi, G., Slep, K.C., Ma, A., and Rogers, S.L. (2011). The microtubule lattice and plus-end association of Drosophila Mini spindles is spatially regulated to fine-tune microtubule dynamics. Mol. Biol. Cell 22: 4343–4361. Curtis, M.D. and Grossniklaus, U. (2003). A Gateway Cloning Vector Set for High-Throughput Functional Analysis of Genes in Planta. Plant Physiol. 133: 462–469. David-Pfeuty, T., Erickson, H.P., and Pantaloni, D. (1977). Guanosinetriphosphatase activity of tubulin associated with microtubule assembly. Proc. Natl. Acad. Sci. 74: 5372–5376. Desai, A. and Mitchison, T.J. (1997). Microtubule polymerization dynamics. Annu. Rev. Cell Dev. Biol. 13: 83–117. Desai, A., Verma, S., Mitchison, T.J., and Walczak, C.E. (1999). Kin I Kinesins Are Microtubule-Destabilizing Enzymes. Cell 96: 69–78. Diamantopoulos, G.S., Perez, F., Goodson, H.V., Batelier, G., Melki, R., Kreis, T.E., and Rickard, J.E. (1999). Dynamic Localization of CLIP-170 to Microtubule Plus Ends Is Coupled to Microtubule Assembly. J. Cell Biol. 144: 99–112. Dixit, R. and Cyr, R. (2004). The Cortical Microtubule Array: From Dynamics to Organization. Plant Cell Online 16: 2546–2552. Ehrhardt, D.W. and Shaw, S.L. (2006a). Microtubule Dynamics and Organization in the Plant Cortical Array. Annu. Rev. Plant Biol. 57: 859–875. 	 136 Ehrhardt, D.W. and Shaw, S.L. (2006b). Microtubule Dynamics and Organization in the Plant Cortical Array. Annu. Rev. Plant Biol. 57: 859–875. Eleftheriou, E.P., Baskin, T.I., and Hepler, P.K. (2005). Aberrant Cell Plate Formation in the Arabidopsis thaliana microtubule organization 1 Mutant. Plant Cell Physiol. 46: 671–675. Eng, R.C. and Wasteneys, G.O. (2014). The Microtubule Plus-End Tracking Protein ARMADILLO-REPEAT KINESIN1 Promotes Microtubule Catastrophe in Arabidopsis. Plant Cell Online: tpc.114.126789. Erhardt, M., Stoppin-Mellet, V., Campagne, S., Canaday, J., Mutterer, J., Fabian, T., Sauter, M., Muller, T., Peter, C., Lambert, A.-M., and Schmit, A.-C. (2002). The plant Spc98p homologue colocalizes with γ-tubulin at microtubule nucleation sites and is required for microtubule nucleation. J. Cell Sci. 115: 2423–2431. Euteneuer, U. and McIntosh, J.R. (1980). Polarity of midbody and phragmoplast microtubules. J. Cell Biol. 87: 509–515. Finst, R.J., Kim, P.J., Griffis, E.R., and Quarmby, L.M. (2000). Fa1p is a 171 kDa protein essential for axonemal microtubule severing in Chlamydomonas. J. Cell Sci. 113: 1963–1971. Fujita, S., Pytela, J., Hotta, T., Kato, T., Hamada, T., Akamatsu, R., Ishida, Y., Kutsuna, N., Hasezawa, S., Nomura, Y., Nakagami, H., and Hashimoto, T. (2013). An Atypical Tubulin Kinase Mediates Stress-Induced Microtubule Depolymerization in Arabidopsis. Curr. Biol. 23: 1969–1978. Galway, M.E., Eng, R.C., Schiefelbein, J.W., and Wasteneys, G.O. (2011). Root hair-specific disruption of cellulose and xyloglucan in AtCSLD3 mutants, and factors affecting the post-rupture resumption of mutant root hair growth. Planta 233: 985–999. Ganguly, A. and Dixit, R. (2013). Mechanisms for regulation of plant kinesins. Curr. Opin. Plant Biol. 16: 704–709. Garcia, M.A., Vardy, L., Koonrugsa, N., and Toda, T. (2001). Fission yeast ch-TOG/XMAP215 homologue Alp14 connects mitotic spindles with the kinetochore and is a component of the Mad2-dependent spindle checkpoint. EMBO J. 20: 3389–3401. Gard, D.L., Becker, B.E., and Josh Romney, S. (2004). MAPping the Eukaryotic Tree of Life: Structure, Function, and Evolution of the MAP215⧸Dis1 Family of Microtubule-Associated Proteins. In B.-I.R. of Cytology, ed (Academic Press), pp. 179–272. 	 137 Gard, D.L. and Kirschner, M.W. (1987). A microtubule-associated protein from Xenopus eggs that specifically promotes assembly at the plus-end. J. Cell Biol. 105: 2203–2215. Gebert, M., Dresselhaus, T., and Sprunck, S. (2008). F-Actin Organization and Pollen Tube Tip Growth in Arabidopsis Are Dependent on the Gametophyte-Specific Armadillo Repeat Protein ARO1. Plant Cell 20: 2798–2814. Graf, R., Daunderer, C., and Schliwa, M. (2000). Dictyostelium DdCP224 is a microtubule-associated protein and a permanent centrosomal resident involved in centrosome duplication. J. Cell Sci. 113: 1747–1758. Grefen, C., Donald, N., Hashimoto, K., Kudla, J., Schumacher, K., and Blatt, M.R. (2010). A ubiquitin-10 promoter-based vector set for fluorescent protein tagging facilitates temporal stability and native protein distribution in transient and stable expression studies. Plant J. 64: 355–365. Gupta, M.L., Carvalho, P., Roof, D.M., and Pellman, D. (2006). Plus end-specific depolymerase activity of Kip3, a kinesin-8 protein, explains its role in positioning the yeast mitotic spindle. Nat. Cell Biol. 8: 913–923. Gutierrez, R., Lindeboom, J.J., Paredez, A.R., Emons, A.M.C., and Ehrhardt, D.W. (2009). Arabidopsis cortical microtubules position cellulose synthase delivery to the plasma membrane and interact with cellulose synthase trafficking compartments. Nat. Cell Biol. 11: 797–806. Hamada, T. (2007). Microtubule-associated proteins in higher plants. J. Plant Res. 120: 79–98. Hamada, T., Igarashi, H., Itoh, T.J., Shimmen, T., and Sonobe, S. (2004). Characterization of a 200 kDa Microtubule-associated Protein of Tobacco BY-2 Cells, a Member of the XMAP215/MOR1 Family. Plant Cell Physiol. 45: 1233–1242. Helenius, J., Brouhard, G., Kalaidzidis, Y., Diez, S., and Howard, J. (2006). The depolymerizing kinesin MCAK uses lattice diffusion to rapidly target microtubule ends. Nature 441: 115–119. Hirokawa, N., Noda, Y., Tanaka, Y., and Niwa, S. (2009). Kinesin superfamily motor proteins and intracellular transport. Nat. Rev. Mol. Cell Biol. 10: 682–696. Hunter, T. (1995). Protein kinases and phosphatases: The Yin and Yang of protein phosphorylation and signaling. Cell 80: 225–236. Hyman, A.A., Chrétien, D., Arnal, I., and Wade, R.H. (1995). Structural changes accompanying GTP hydrolysis in microtubules: information from a slowly hydrolyzable analogue guanylyl-(alpha,beta)-methylene-diphosphonate. J. Cell Biol. 128: 117–125. 	 138 Hyman, A.A., Salser, S., Drechsel, D.N., Unwin, N., and Mitchison, T.J. (1992). Role of GTP hydrolysis in microtubule dynamics: information from a slowly hydrolyzable analogue, GMPCPP. Mol. Biol. Cell 3: 1155–1167. Jones, M.A., Raymond, M.J., and Smirnoff, N. (2006). Analysis of the root-hair morphogenesis transcriptome reveals the molecular identity of six genes with roles in root-hair development in Arabidopsis. Plant J. 45: 83–100. Jürgens, G. (2005). Plant cytokinesis: fission by fusion. Trends Cell Biol. 15: 277–283. Kawamura, E., Himmelspach, R., Rashbrooke, M.C., Whittington, A.T., Gale, K.R., Collings, D.A., and Wasteneys, G.O. (2006). MICROTUBULE ORGANIZATION 1 Regulates Structure and Function of Microtubule Arrays during Mitosis and Cytokinesis in the Arabidopsis Root. Plant Physiol. 140: 102–114. Kawamura, E. and Wasteneys, G.O. (2008). MOR1, the Arabidopsis thaliana homologue of Xenopus MAP215, promotes rapid growth and shrinkage, and suppresses the pausing of microtubules in vivo. J. Cell Sci. 121: 4114–4123. Kinoshita, E. (2006). Phosphate‐binding tag, a new tool to visualize phosphorylated proteins. Mol Cell Proteomics 5: 749–757. Kong, Z., Hotta, T., Lee, Y.-R.J., Horio, T., and Liu, B. (2010). The γ -Tubulin Complex Protein GCP4 Is Required for Organizing Functional Microtubule Arrays in Arabidopsis thaliana. Plant Cell 22: 191–204. Kronja, I., Kruljac-Letunic, A., Caudron-Herger, M., Bieling, P., and Karsenti, E. (2009). XMAP215–EB1 Interaction Is Required for Proper Spindle Assembly and Chromosome Segregation in Xenopus Egg Extract. Mol. Biol. Cell 20: 2684–2696. Lau, O.S., Davies, K.A., Chang, J., Adrian, J., Rowe, M.H., Ballenger, C.E., and Bergmann, D.C. (2014). Direct roles of SPEECHLESS in the specification of stomatal self-renewing cells. Science 345: 1605–1609. Lechner, B., Rashbrooke, M.C., Collings, D.A., Eng, R.C., Kawamura, E., Whittington, A.T., and Wasteneys, G.O. (2012). The N-terminal TOG domain of Arabidopsis MOR1 modulates affinity for microtubule polymers. J. Cell Sci. 125: 4812–4821. Ledbetter, M.C. and Porter, K.R. (1963). A “microtubule” in Plant Cell Fine Structure. J. Cell Biol. 19: 239–250. Ledbetter, M.C. and Porter, K.R. (1964). Morphology of Microtubules of Plant Cell. Science 144: 872–874. Leduc, C., Padberg-Gehle, K., Varga, V., Helbing, D., Diez, S., and Howard, J. (2012). Molecular crowding creates traffic jams of kinesin motors on microtubules. Proc. Natl. Acad. Sci. 109: 6100–6105. 	 139 Lee, S., Cho, D.-I., Kang, J., Kim, M.-D., and Kim, S.Y. (2010). AtNEK6 interacts with ARIA and is involved in ABA response during seed germination. Mol. Cells 29: 559–566. Lee, Y.-R.J., Giang, H.M., and Liu, B. (2001). A Novel Plant Kinesin-Related Protein Specifically Associates with the Phragmoplast Organelles. Plant Cell 13: 2427–2439. Li, J. et al. (2011). Mutation of Rice BC12/GDD1, Which Encodes a Kinesin-Like Protein That Binds to a GA Biosynthesis Gene Promoter, Leads to Dwarfism with Impaired Cell Elongation. Plant Cell 23: 628–640. Li, J.-F., Park, E., Arnim, A.G. von, and Nebenführ, A. (2009). The FAST technique: a simplified Agrobacterium-based transformation method for transient gene expression analysis in seedlings of Arabidopsis and other plant species. Plant Methods 5: 6. Lindeboom, J.J., Lioutas, A., Deinum, E.E., Tindemans, S.H., Ehrhardt, D.W., Emons, A.M.C., Vos, J.W., and Mulder, B.M. (2013a). Cortical Microtubule Arrays Are Initiated from a Nonrandom Prepattern Driven by Atypical Microtubule Initiation. Plant Physiol. 161: 1189–1201. Lindeboom, J.J., Nakamura, M., Hibbel, A., Shundyak, K., Gutierrez, R., Ketelaar, T., Emons, A.M.C., Mulder, B.M., Kirik, V., and Ehrhardt, D.W. (2013b). A Mechanism for Reorientation of Cortical Microtubule Arrays Driven by Microtubule Severing. Science 342: 1245533. Liu, B., Marc, J., Joshi, H.C., and Palevitz, B.A. (1993). A gamma-tubulin-related protein associated with the microtubule arrays of higher plants in a cell cycle-dependent manner. J. Cell Sci. 104: 1217–1228. Lloyd, C.W., Pearce, K.J., Rawlins, D.J., Ridge, R.W., and Shaw, P.J. (1987). Endoplasmic microtubules connect the advancing nucleus to the tip of legume root hairs, but F-actin is involved in basipetal migration. Cell Motil. Cytoskeleton 8: 27–36. Lu, L., Lee, Y.-R.J., Pan, R., Maloof, J.N., and Liu, B. (2005). An Internal Motor Kinesin Is Associated with the Golgi Apparatus and Plays a Role in Trichome Morphogenesis in Arabidopsis. Mol. Biol. Cell 16: 811–823. MacNeal, R.K. and Purich, D.L. (1978). Stoichiometry and role of GTP hydrolysis in bovine neurotubule assembly. J. Biol. Chem. 253: 4683–4687. MacRae, T.H. (1997). Tubulin Post-Translational Modifications. Eur. J. Biochem. 244: 265–278. Mahjoub, M.R., Montpetit, B., Zhao, L., Finst, R.J., Goh, B., Kim, A.C., and Quarmby, L.M. (2002). The FA2 gene of Chlamydomonas encodes a NIMA family kinase with roles in cell cycle progression and microtubule severing during deflagellation. J. Cell Sci. 115: 1759–1768. 	 140 Mahjoub, M.R., Rasi, M.Q., and Quarmby, L.M. (2004). A NIMA-related Kinase, Fa2p, Localizes to a Novel Site in the Proximal Cilia of Chlamydomonas and Mouse Kidney Cells. Mol. Biol. Cell 15: 5172–5186. Malcos, J.L. and Cyr, R.J. (2011). An ungrouped plant kinesin accumulates at the preprophase band in a cell cycle-dependent manner. Cytoskeleton 68: 247–258. Mandelkow, E.M., Mandelkow, E., and Milligan, R.A. (1991). Microtubule dynamics and microtubule caps: a time-resolved cryo-electron microscopy study. J. Cell Biol. 114: 977–991. Mao, G., Chan, J., Calder, G., Doonan, J.H., and Lloyd, C.W. (2005). Modulated targeting of GFP-AtMAP65-1 to central spindle microtubules during division. Plant J. 43: 469–478. Marc, J., Granger, C.L., Brincat, J., Fisher, D.D., Kao, T., McCubbin, A.G., and Cyr, R.J. (1998). A GFP–MAP4 Reporter Gene for Visualizing Cortical Microtubule Rearrangements in Living Epidermal Cells. Plant Cell Online 10: 1927–1939. Marcus, A.I., Ambrose, J.C., Blickley, L., Hancock, W.O., and Cyr, R.J. (2002). Arabidopsis thaliana protein, ATK1, is a minus-end directed kinesin that exhibits non-processive movement. Cell Motil. Cytoskeleton 52: 144–150. Marcus, A.I., Li, W., Ma, H., and Cyr, R.J. (2003). A Kinesin Mutant with an Atypical Bipolar Spindle Undergoes Normal Mitosis. Mol. Biol. Cell 14: 1717–1726. Martin, K.C. and Ephrussi, A. (2009). mRNA Localization: Gene Expression in the Spatial Dimension. Cell 136: 719–730. Masucci, J.D. and Schiefelbein, J.W. (1996). Hormones act downstream of TTG and GL2 to promote root hair outgrowth during epidermis development in the Arabidopsis root. Plant Cell 8: 1505–1517. Mathur, J., Mathur, N., Kernebeck, B., Srinivas, B.P., and Hülskamp, M. (2003). A Novel Localization Pattern for an EB1-like Protein Links Microtubule Dynamics to Endomembrane Organization. Curr. Biol. 13: 1991–1997. Matthews, L.R., Carter, P., Thierry-Mieg, D., and Kemphues, K. (1998). ZYG-9, A Caenorhabditis elegans Protein Required for Microtubule Organization and Function, Is a Component of Meiotic and Mitotic Spindle Poles. J. Cell Biol. 141: 1159–1168. Mayr, M.I., Storch, M., Howard, J., and Mayer, T.U. (2011). A Non-Motor Microtubule Binding Site Is Essential for the High Processivity and Mitotic Function of Kinesin-8 Kif18A. PLoS ONE 6: e27471. Miki, T., Naito, H., Nishina, M., and Goshima, G. (2014). Endogenous localizome identifies 43 mitotic kinesins in a plant cell. Proc. Natl. Acad. Sci. 111: E1053–E1061. 	 141 Mitchison, T.J. (1993). Localization of an exchangeable GTP binding site at the plus end of microtubules. Science 261: 1044–1047. Mitchison, T. and Kirschner, M. (1984). Dynamic instability of microtubule growth. Nature 312: 237–242. Motose, H., Hamada, T., Yoshimoto, K., Murata, T., Hasebe, M., Watanabe, Y., Hashimoto, T., Sakai, T., and Takahashi, T. (2011). NIMA-related kinases 6, 4, and 5 interact with each other to regulate microtubule organization during epidermal cell expansion in Arabidopsis thaliana. Plant J. 67: 993–1005. Motose, H., Takatani, S., Ikeda, T., and Takahashi, T. (2012). NIMA-related kinases regulate directional cell growth and organ development through microtubule function in Arabidopsis thaliana. Plant Signal. Behav. 7: 1552–1555. Motose, H., Tominaga, R., Wada, T., Sugiyama, M., and Watanabe, Y. (2008). A NIMA-related protein kinase suppresses ectopic outgrowth of epidermal cells through its kinase activity and the association with microtubules. Plant J. 54: 829–844. Mudgil, Y., Shiu, S.-H., Stone, S.L., Salt, J.N., and Goring, D.R. (2004). A Large Complement of the Predicted Arabidopsis ARM Repeat Proteins Are Members of the U-Box E3 Ubiquitin Ligase Family. Plant Physiol. 134: 59–66. Müller-Reichert, T., Chrétien, D., Severin, F., and Hyman, A.A. (1998). Structural changes at microtubule ends accompanying GTP hydrolysis: Information from a slowly hydrolyzable analogue of GTP, guanylyl (α,β)methylenediphosphonate. Proc. Natl. Acad. Sci. 95: 3661–3666. Murata, T., Sonobe, S., Baskin, T.I., Hyodo, S., Hasezawa, S., Nagata, T., Horio, T., and Hasebe, M. (2005). Microtubule-dependent microtubule nucleation based on recruitment of γ-tubulin in higher plants. Nat. Cell Biol. 7: 961–968. Nabeshima, K., Kurooka, H., Takeuchi, M., Kinoshita, K., Nakaseko, Y., and Yanagida, M. (1995). p93dis1, which is required for sister chromatid separation, is a novel microtubule and spindle pole body-associating protein phosphorylated at the Cdc2 target sites. Genes Dev. 9: 1572–1585. Nakagawa, T., Nakamura, S., Tanaka, K., Kawamukai, M., Suzuki, T., Nakamura, K., Kimura, T., and Ishiguro, S. (2008). Development of R4 Gateway Binary Vectors (R4pGWB) Enabling High-Throughput Promoter Swapping for Plant Research. Biosci. Biotechnol. Biochem. 72: 624–629. Nakajima, K., Furutani, I., Tachimoto, H., Matsubara, H., and Hashimoto, T. (2004). SPIRAL1 Encodes a Plant-Specific Microtubule-Localized Protein Required for Directional Control of Rapidly Expanding Arabidopsis Cells. Plant Cell 16: 1178–1190. 	 142 Nakamura, M., Ehrhardt, D.W., and Hashimoto, T. (2010a). Microtubule and katanin-dependent dynamics of microtubule nucleation complexes in the acentrosomal Arabidopsis cortical array. Nat. Cell Biol. 12: 1064–1070. Nakamura, M., Naoi, K., Shoji, T., and Hashimoto, T. (2004). Low Concentrations of Propyzamide and Oryzalin Alter Microtubule Dynamics in Arabidopsis Epidermal Cells. Plant Cell Physiol. 45: 1330–1334. Nakamura, S., Grigoriev, I., Nogi, T., Hamaji, T., Cassimeris, L., and Mimori-Kiyosue, Y. (2012). Dissecting the Nanoscale Distributions and Functions of Microtubule-End-Binding Proteins EB1 and ch-TOG in Interphase HeLa Cells. PLoS ONE 7: e51442. Nakamura, S., Mano, S., Tanaka, Y., Ohnishi, M., Nakamori, C., Araki, M., Niwa, T., Nishimura, M., Kaminaka, H., Nakagawa, T., Sato, Y., and Ishiguro, S. (2010b). Gateway Binary Vectors with the Bialaphos Resistance Gene, <I>bar</I>, as a Selection Marker for Plant Transformation. Biosci. Biotechnol. Biochem. 74: 1315–1319. Ben-Nissan, G., Cui, W., Kim, D.-J., Yang, Y., Yoo, B.-C., and Lee, J.-Y. (2008). Arabidopsis Casein Kinase 1-Like 6 Contains a Microtubule-Binding Domain and Affects the Organization of Cortical Microtubules. Plant Physiol. 148: 1897–1907. O’Connell, M.J., Krien, M.J.E., and Hunter, T. (2003). Never say never. The NIMA-related protein kinases in mitotic control. Trends Cell Biol. 13: 221–228. Oda, Y. and Fukuda, H. (2013). Rho of Plant GTPase Signaling Regulates the Behavior of Arabidopsis Kinesin-13A to Establish Secondary Cell Wall Patterns. Plant Cell 25: 4439–4450. O’Regan, L., Blot, J., and Fry, A.M. (2007). Mitotic regulation by NIMA-related kinases. Cell Div. 2: 25. Osmani, A.H., O’Donnell, K., Pu, R.T., and Osmani, S.A. (1991). Activation of the nimA protein kinase plays a unique role during mitosis that cannot be bypassed by absence of the bimE checkpoint. EMBO J. 10: 2669–2679. Osmani, S.A., Pu, R.T., and Morris, N.R. (1988). Mitotic induction and maintenance by overexpression of a G2-specific gene that encodes a potential protein kinase. Cell 53: 237–244. Perez, F., Diamantopoulos, G.S., Stalder, R., and Kreis, T.E. (1999). CLIP-170 Highlights Growing Microtubule Ends In Vivo. Cell 96: 517–527. Pitts, R.J., Cernac, A., and Estelle, M. (1998). Auxin and ethylene promote root hair elongation inArabidopsis. Plant J. 16: 553–560. 	 143 Podolski, M., Mahamdeh, M., and Howard, J. (2014). Stu2, the Budding Yeast XMAP215/Dis1 Homolog, Promotes Assembly of Yeast Microtubules by Increasing Growth Rate and Decreasing Catastrophe Frequency. J. Biol. Chem. 289: 28087–28093. Preibisch, S., Saalfeld, S., and Tomancak, P. (2009). Globally optimal stitching of tiled 3D microscopic image acquisitions. Bioinformatics 25: 1463–1465. Rasmussen, C.G., Wright, A.J., and Müller, S. (2013). The role of the cytoskeleton and associated proteins in determination of the plant cell division plane. Plant J. 75: 258–269. Reddy, A.S.N. (2001). Molecular motors and their functions in plants. In B.-I.R. of Cytology, ed (Academic Press), pp. 97–178. Reddy, A.S.N. and Day, I.S. (2011). Microtubule Motor Proteins in the Eukaryotic Green Lineage: Functions and Regulation. In The Plant Cytoskeleton, B. Liu, ed, Advances in Plant Biology. (Springer New York), pp. 119–141. Rickard, J.E. and Kreis, T.E. (1991). Binding of pp170 to microtubules is regulated by phosphorylation. J. Biol. Chem. 266: 17597–17605. Rodionov, V.I. and Borisy, G.G. (1997). Microtubule Treadmilling in Vivo. Science 275: 215–218. Rogers, S., Wells, R., and Rechsteiner, M. (1986). Amino Acid Sequences Common to Rapidly Degraded Proteins: The PEST Hypothesis. Science 234: 364–368. Sakai, T. et al. (2008). Armadillo repeat-containing kinesins and a NIMA-related kinase are required for epidermal-cell morphogenesis in Arabidopsis. Plant J. 53: 157–171. Samuels, A.L., Giddings, T.H., and Staehelin, L.A. (1995). Cytokinesis in tobacco BY-2 and root tip cells: a new model of cell plate formation in higher plants. J. Cell Biol. 130: 1345–1357. Sandblad, L., Busch, K.E., Tittmann, P., Gross, H., Brunner, D., and Hoenger, A. (2006). The Schizosaccharomyces pombe EB1 Homolog Mal3p Binds and Stabilizes the Microtubule Lattice Seam. Cell 127: 1415–1424. Sasabe, M., Soyano, T., Takahashi, Y., Sonobe, S., Igarashi, H., Itoh, T.J., Hidaka, M., and Machida, Y. (2006). Phosphorylation of NtMAP65-1 by a MAP kinase down-regulates its activity of microtubule bundling and stimulates progression of cytokinesis of tobacco cells. Genes Dev. 20: 1004–1014. Schnitzer, M.J. and Block, S.M. (1997). Kinesin hydrolyses one ATP per 8-nm step. Nature 388: 386–390. 	 144 Sedbrook, J.C., Ehrhardt, D.W., Fisher, S.E., Scheible, W.-R., and Somerville, C.R. (2004). The Arabidopsis SKU6/SPIRAL1 Gene Encodes a Plus End–Localized Microtubule-Interacting Protein Involved in Directional Cell Expansion. Plant Cell 16: 1506–1520. Shaw, S.L., Kamyar, R., and Ehrhardt, D.W. (2003). Sustained Microtubule Treadmilling in Arabidopsis Cortical Arrays. Science 300: 1715–1718. Shelden, E. and Wadsworth, P. (1993). Observation and quantification of individual microtubule behavior in vivo: microtubule dynamics are cell-type specific. J. Cell Biol. 120: 935–945. Sieberer, B. and Emons, A.M.C. (2000). Cytoarchitecture and pattern of cytoplasmic streaming in root hairs ofMedicago truncatula during development and deformation by nodulation factors. Protoplasma 214: 118–127. Sieberer, B.J., Ketelaar, T., Esseling, J.J., and Emons, A.M.C. (2005). Microtubules guide root hair tip growth. New Phytol. 167: 711–719. Sieberer, B.J. and Timmers, A.C.J. (2009). Microtubules in Plant Root Hairs and Their Role in Cell Polarity and Tip Growth. In Root Hairs, P.D.A.M.C. Emons and D.T. Ketelaar, eds, Plant Cell Monographs. (Springer Berlin Heidelberg), pp. 233–248. Sieberer, B.J., Timmers, A.C.J., Lhuissier, F.G.P., and Emons, A.M.C. (2002). Endoplasmic Microtubules Configure the Subapical Cytoplasm and Are Required for Fast Growth of Medicago truncatulaRoot Hairs. Plant Physiol. 130: 977–988. Slep, K.C. and Vale, R.D. (2007). Structural Basis of Microtubule Plus End Tracking by XMAP215, CLIP-170, and EB1. Mol. Cell 27: 976–991. Smertenko, A.P., Chang, H.-Y., Sonobe, S., Fenyk, S.I., Weingartner, M., Bögre, L., and Hussey, P.J. (2006). Control of the AtMAP65-1 interaction with microtubules through the cell cycle. J. Cell Sci. 119: 3227–3237. De Souza, C.P.C., Osmani, A.H., Wu, L.-P., Spotts, J.L., and Osmani, S.A. (2000). Mitotic Histone H3 Phosphorylation by the NIMA Kinase in Aspergillus nidulans. Cell 102: 293–302. Spiegelman, B.M., Penningroth, S.M., and Kirschner, M.W. (1977). Turnover of tubulin and the N site GTP in chinese hamster ovary cells. Cell 12: 587–600. Staehelin, L.A. and Hepler, P.K. (1996). Cytokinesis in Higher Plants. Cell 84: 821–824. Stumpff, J., Du, Y., English, C.A., Maliga, Z., Wagenbach, M., Asbury, C.L., Wordeman, L., and Ohi, R. (2011). A Tethering Mechanism Controls the Processivity and Kinetochore-Microtubule Plus-End Enrichment of the Kinesin-8 Kif18A. Mol. Cell 43: 764–775. 	 145 Suetsugu, N., Yamada, N., Kagawa, T., Yonekura, H., Uyeda, T.Q.P., Kadota, A., and Wada, M. (2010). Two kinesin-like proteins mediate actin-based chloroplast movement in Arabidopsis thaliana. Proc. Natl. Acad. Sci. 107: 8860–8865. Sugimoto, K., Williamson, R.E., and Wasteneys, G.O. (2000). New Techniques Enable Comparative Analysis of Microtubule Orientation, Wall Texture, and Growth Rate in Intact Roots of Arabidopsis. Plant Physiol. 124: 1493–1506. Su, X., Qiu, W., Gupta Jr., M.L., Pereira-Leal, J.B., Reck-Peterson, S.L., and Pellman, D. (2011). Mechanisms Underlying the Dual-Mode Regulation of Microtubule Dynamics by Kip3/Kinesin-8. Mol. Cell 43: 751–763. Svoboda, K., Schmidt, C.F., Schnapp, B.J., and Block, S.M. (1993). Direct observation of kinesin stepping by optical trapping interferometry. Nature 365: 721–727. Tanimoto, M., Roberts, K., and Dolan, L. (1995). Ethylene is a positive regulator of root hair development in Arabidopsis thaliana. Plant J. 8: 943–948. Teixidó-Travesa, N., Roig, J., and Lüders, J. (2012). The where, when and how of microtubule nucleation – one ring to rule them all. J. Cell Sci. 125: 4445–4456. Tewari, R., Bailes, E., Bunting, K.A., and Coates, J.C. (2010). Armadillo-repeat protein functions: questions for little creatures. Trends Cell Biol. 20: 470–481. Tirnauer, J.S., Grego, S., Salmon, E.D., and Mitchison, T.J. (2002). EB1–Microtubule Interactions in Xenopus Egg Extracts: Role of EB1 in Microtubule Stabilization and Mechanisms of Targeting to Microtubules. Mol. Biol. Cell 13: 3614–3626. Tournebize, R., Popov, A., Kinoshita, K., Ashford, A.J., Rybina, S., Pozniakovsky, A., Mayer, T.U., Walczak, C.E., Karsenti, E., and Hyman, A.A. (2000). Control of microtubule dynamics by the antagonistic activities of XMAP215 and XKCM1 in Xenopus egg extracts. Nat. Cell Biol. 2: 13–19. Twell, D., Park, S.K., Hawkins, T.J., Schubert, D., Schmidt, R., Smertenko, A., and Hussey, P.J. (2002). MOR1/GEM1 has an essential role in the plant-specific cytokinetic phragmoplast. Nat. Cell Biol. 4: 711–714. Ubersax, J.A. and Ferrell Jr, J.E. (2007). Mechanisms of specificity in protein phosphorylation. Nat. Rev. Mol. Cell Biol. 8: 530–541. Vale, R.D. and Fletterick, R.J. (1997). The Design Plan of Kinesin Motors. Annu. Rev. Cell Dev. Biol. 13: 745–777. Vale, R.D., Reese, T.S., and Sheetz, M.P. (1985). Identification of a Novel Force-Generating Protein, Kinesin, Involved in Microtubule-Based Motility. Cell 42: 39–50. Vanstraelen, M., Acosta, J.A.T., Veylder, L.D., Inzé, D., and Geelen, D. (2004). A Plant-Specific Subclass of C-Terminal Kinesins Contains a Conserved A-Type Cyclin-	 146 Dependent Kinase Site Implicated in Folding and Dimerization. Plant Physiol. 135: 1417–1429. Vanstraelen, M., Van Damme, D., De Rycke, R., Mylle, E., Inzé, D., and Geelen, D. (2006). Cell Cycle-Dependent Targeting of a Kinesin at the Plasma Membrane Demarcates the Division Site in Plant Cells. Curr. Biol. 16: 308–314. Varga, V., Helenius, J., Tanaka, K., Hyman, A.A., Tanaka, T.U., and Howard, J. (2006). Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner. Nat. Cell Biol. 8: 957–962. Vigneault, F., Lachance, D., Cloutier, M., Pelletier, G., Levasseur, C., and Séguin, A. (2007). Members of the plant NIMA-related kinases are involved in organ development and vascularization in poplar, Arabidopsis and rice. Plant J. 51: 575–588. Vogel, J., Drapkin, B., Oomen, J., Beach, D., Bloom, K., and Snyder, M. (2001). Phosphorylation of γ-Tubulin Regulates Microtubule Organization in Budding Yeast. Dev. Cell 1: 621–631. Walczak, C.E., Mitchison, T.J., and Desai, A. (1996). XKCM1: A Xenopus Kinesin-Related Protein That Regulates Microtubule Dynamics during Mitotic Spindle Assembly. Cell 84: 37–47. Walia, A., Lee, J.S., Wasteneys, G., and Ellis, B. (2009). Arabidopsis mitogen-activated protein kinase MPK18 mediates cortical microtubule functions in plant cells. Plant J. 59: 565–575. Walker, R.A., O’Brien, E.T., Pryer, N.K., Soboeiro, M.F., Voter, W.A., Erickson, H.P., and Salmon, E.D. (1988). Dynamic instability of individual microtubules analyzed by video light microscopy: rate constants and transition frequencies. J. Cell Biol. 107: 1437–1448. Wang, P.J. and Huffaker, T.C. (1997). Stu2p: A Microtubule-Binding Protein that Is an Essential Component of the Yeast Spindle Pole Body. J. Cell Biol. 139: 1271–1280. Wasteneys, G.O. (2002). Microtubule organization in the green kingdom: chaos or self-order? J. Cell Sci. 115: 1345–1354. Wasteneys, G.O. and Ambrose, J.C. (2009). Spatial organization of plant cortical microtubules: close encounters of the 2D kind. Trends Cell Biol. 19: 62–71. Weaver, L.N., Ems-McClung, S.C., Stout, J.R., LeBlanc, C., Shaw, S.L., Gardner, M.K., and Walczak, C.E. (2011). Kif18A Uses a Microtubule Binding Site in the Tail for Plus-End Localization and Spindle Length Regulation. Curr. Biol. 21: 1500–1506. 	 147 Wei, L., Zhang, W., Liu, Z., and Li, Y. (2009). AtKinesin-13A is located on Golgi-associated vesicle and involved in vesicle formation/budding in Arabidopsis root-cap peripheral cells. BMC Plant Biol. 9: 138. Weisenberg, R.C., Broisy, G.G., and Taylor, E.W. (1968). Colchicine-binding protein of mammalian brain and its relation to microtubules. Biochemistry (Mosc.) 7: 4466–4479. Westermann, S. and Weber, K. (2003). Post-translational modifications regulate microtubule function. Nat. Rev. Mol. Cell Biol. 4: 938–948. Whittington, A.T., Vugrek, O., Wei, K.J., Hasenbein, N.G., Sugimoto, K., Rashbrooke, M.C., and Wasteneys, G.O. (2001). MOR1 is essential for organizing cortical microtubules in plants. Nature 411: 610–613. Widlund, P.O., Stear, J.H., Pozniakovsky, A., Zanic, M., Reber, S., Brouhard, G.J., Hyman, A.A., and Howard, J. (2011). XMAP215 polymerase activity is built by combining multiple tubulin-binding TOG domains and a basic lattice-binding region. Proc. Natl. Acad. Sci. 108: 2741–2746. Wiese, C. and Zheng, Y. (2006). Microtubule nucleation: γ-tubulin and beyond. J. Cell Sci. 119: 4143–4153. Wightman, R., Chomicki, G., Kumar, M., Carr, P., and Turner, S.R. (2013). SPIRAL2 Determines Plant Microtubule Organization by Modulating Microtubule Severing. Curr. Biol. 23: 1902–1907. Wightman, R. and Turner, S.R. (2007). Severing at sites of microtubule crossover contributes to microtubule alignment in cortical arrays. Plant J. 52: 742–751. Yang, G., Gao, P., Zhang, H., Huang, S., and Zheng, Z.-L. (2007). A Mutation in MRH2 Kinesin Enhances the Root Hair Tip Growth Defect Caused by Constitutively Activated ROP2 Small GTPase in Arabidopsis. PLoS ONE 2: e1074. Yoo, C.-M. and Blancaflor, E.B. (2013). Overlapping and divergent signaling pathways for ARK1 and AGD1 in the control of root hair polarity in Arabidopsis thaliana. Front. Plant Sci. 4. Yoo, C.-M., Wen, J., Motes, C.M., Sparks, J.A., and Blancaflor, E.B. (2008). A Class I ADP-Ribosylation Factor GTPase-Activating Protein Is Critical for Maintaining Directional Root Hair Growth in Arabidopsis. Plant Physiol. 147: 1659–1674. Zanic, M., Stear, J.H., Hyman, A.A., and Howard, J. (2009). EB1 Recognizes the Nucleotide State of Tubulin in the Microtubule Lattice. PLoS ONE 4: e7585. Zanic, M., Widlund, P.O., Hyman, A.A., and Howard, J. (2013). Synergy between XMAP215 and EB1 increases microtubule growth rates to physiological levels. Nat. Cell Biol. 15: 688–693. 	 148 Zeng, C.J.T., Lee, Y.-R.J., and Liu, B. (2009). The WD40 Repeat Protein NEDD1 Functions in Microtubule Organization during Cell Division in Arabidopsis thaliana. Plant Cell 21: 1129–1140. Zhang, B. et al. (2011). NIMA-related kinase NEK6 affects plant growth and stress response in Arabidopsis. Plant J. 68: 830–843. Zhang, Q., Fishel, E., Bertroche, T., and Dixit, R. (2013). Microtubule Severing at Crossover Sites by Katanin Generates Ordered Cortical Microtubule Arrays in Arabidopsis. Curr. Biol. 23: 2191–2195. Zheng, Y., Wong, M.L., Alberts, B., and Mitchison, T. (1995). Nucleation of microtubule assembly by a γ-tubulin-containing ring complex. Nature 378: 578–583. Zhou, R., Benavente, L.M., Stepanova, A.N., and Alonso, J.M. (2011). A recombineering-based gene tagging system for Arabidopsis. Plant J. 66: 712–723. Zhu, C. and Dixit, R. (2012). Functions of the Arabidopsis kinesin superfamily of microtubule-based motor proteins. Protoplasma 249: 887–899. Zhu, C. and Dixit, R. (2011). Single Molecule Analysis of the Arabidopsis FRA1 Kinesin Shows that It Is a Functional Motor Protein with Unusually High Processivity. Mol. Plant 4: 879–885. Zhu, C., Ganguly, A., Baskin, T.I., McClosky, D.D., Anderson, C.T., Foster, C., Meunier, K.A., Okamoto, R., Berg, H., and Dixit, R. (2015). The FRA1 kinesin contributes to cortical microtubule-mediated trafficking of cell wall components. Plant Physiol.: pp.114.251462.  

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