University of Cincinnati

UNIVERSITY OF CINCINNATI

Date: 1-Oct-2010

I, Jason Matthew Puglise , hereby submit this original work as part of the requirements for the degree of: Doctor of Philosophy in Cell & Molecular Biology It is entitled: Roles of the Rac/Cdc42 effector proteins Pak and PIX in cytokinesis,

ciliogenesis, and cyst formation in renal epithelial cells

Student Signature: Jason Matthew Puglise

This work and its defense approved by: Committee Chair: Robert Brackenbury, PhD Robert Brackenbury, PhD

11/1/2010 1,117

Roles of the Rac/Cdc42 effector proteins Pak and PIX in cytokinesis,

ciliogenesis, and cyst formation in renal epithelial cells

A dissertation submitted to the Graduate School of the University of Cincinnati in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

in the Graduate Program of Cancer and Cell Biology of the College of Medicine

Jason M. Puglise M.Sc., Wright State University 2005

Committee Chair: Robert Brackenbury, Ph.D.

ABSTRACT

Puglise, Jason M. Ph.D., Cancer and Cell Biology Program. University of Cincinnati, 2010. Roles of the Rac/Cdc42 effector proteins Pak and PIX in cytokinesis, ciliogenesis, and cyst formation in renal epithelial cells.

The p21-activated kinase 1 (Pak1) is a putative Rac/Cdc42 effector molecule and a multifunctional enzyme implicated in a wide range of cellular and biological activities.

Although well-established as a regulator of cytoskeletal and microtubule dynamics, Pak1 influences centrosome behavior and plays a part in the cell cycle. We examine the role Pak1 and its binding partner Pak1-interacting exchange factor (PIX) play in centrosome dynamics and in

cell cycle events in renal epithelial cells. Utilizing Madin-Darby canine kidney (MDCK) cells as our model system, we provide evidence that Pak1 and PIX function are important for proper centrosome functioning and cell division. We found that inhibition of the normal function of Pak or PIX leads to centrosome and cytokinesis defects. Moreover, our results reveal that Pak1 and

PIX play a crucial role in renal epithelial morphogenesis and ciliogenesis, a function not previously described for Pak. Furthermore, our findings stress the importance of the Pak-PIX interaction in regulating cilia and acetylated tubulin dynamics, including lumen formation.

Taken together, our work suggests that Paks are important for renal biology.

iii

JASON M. PUGLISE

2010

ACKNOWLEDGMENTS

I want to thank various past and present members of the Zegers/ter Beest lab from the

University of Cincinnati and from the University of Chicago for their contributions to my work.

I want to thank my committee members: Dr. John Bissler, Dr. Robert Brackenbury (Committee

Chair), and Dr. Wallace Ip for their time and cooperation. I want to express my gratitude to Dr.

Karl Matlin, who played a role in my admittance and timely departure from the program. I’d like to sincerely thank my advisor, Dr. Mirjam Zegers, for her help and guidance on my dissertation. In addition, I’d like to thank my good friend, Steve Kioko, and my brother, Scott, for their advice and proofreading of my dissertation.

Lastly, I’d like to dedicate this thesis to my family—particularly my father, stepmother, brother, sister, and step-uncle for their unwavering support and encouragement. Without their aid, I could not have overcome the obstacles nor endured the hardships that were imposed upon me. I am truly indebted to them for all their help, and am appreciative for having such a caring and understanding family. Truly, blood is thicker than water.

“Our greatest glory is not in never failing, but in rising up every time we fail.”

(Ralph Waldo Emerson)

TABLE OF CONTENTS

Page

INTRODUCTION...... 1

CHAPTER 1. General background and scope of dissertation

I. Introduction to Rho GTPases and Pak family of kinases...... 2

Background...... 2

Structure of Pak...... 6

Modes of action…………………………………………………………………..7

II. Paks as multifunctional kinases……………………………………………………..8

Regulation of cytoskeleton dynamics...... 8

Role of Paks and small GTPases in cell cycle, cytokinesis,

and spindle regulation……………………………….………………...…..……10

Role of Paks and small GTPases in microtubule dynamics...... 13

III. Centrosomes...... 18

General...... 18

Function………………………………………………………………………...19

Centrosome duplication...... 20

Role of Paks and small GTPases in centrosome dynamics……………………..20

Consequences of centrosome malfunctioning and disease...... 21

IV. Cilia………………………………………………………………………………….23

General………………………………………………………………………….23

Structure………………………………………………………………………...23

Function………………………………………………………………………...24

TABLE OF CONTENTS (continued)

Page

Primary cilia as mechanosensors...... 24

Primary cilia formation is linked to the centriole and cell cycle...... 25

Diseases associated with ciliary dysfunction……………………...……………27

V. Scope of dissertation...... 27

CHAPTER 2. Pak and PIX regulate ciliogenesis and cytokinesis

I. Introduction…………………………………………………………………………...33

II. Results………………………………………………………………………………..35

III. Discussion…………………………………………………………………………..47

CHAPTER 3. Pak and PIX control lumen formation

I. Introduction…………………………………………………………………………...77

II. Results………………………………………………………………………………..82

III. Discussion…………………………………………………………………………..84

CHAPTER 4. Materials and methods

I. Cell culture and cell lines……………………………………………………………..94

II. Western blotting (WB) and immunoprecipitation (IP)………………………………96

III. siRNA………………………………………………….…………...…………….…97

IV. Immunofluorescence (IF) confocal microscopy…………………………………….98

V. BrdU incorporation…………………………………………………………………100

vii

TABLE OF CONTENTS (continued)

Page

CHAPTER 5. General discussion/ future goals & directions

I. General discussion………………………………………………………………..…102

II. Future goals & directions……………………………………………………..…….107

REFERENCES...... 110

viii

LIST OF FIGURES

Page

CHAPTER 1.

Figure 1. Diagram of Rho GTPases (Rho, Rac, and CDC42) in their inactive and active states...... 29

Figure 2. Schematic diagram indicating features of Pak1 structure…………………………...30

Figure 3. General structure of cilia…………………………………………………………….31

Figure 4. The assembly and disassembly of primary cilia are coordinated with the centriole and cell cycle………………………………………………………………………….32

CHAPTER 2.

Figure 1. Overexpression of dominant-negative Cdc42 promotes ciliogenesis……………….55

Figure 2. GIT1 and PAK localize to centrosome……………………………………………...58

Figure 3. Active Pak1 inhibits ciliogenesis over time…………………………………………60

Figure 4. Diminished ciliogenesis in active Pak1 mutant cells partially depends on Pak-PIX interaction……………………….…………………………………………….………62

Figure 5. Loss of βPIX leads to formation of Ac-tub rich cell extensions…………………….64

Figure 6. Ac-tub rich cell extensions induced by βPIX knockdown do not emanate from a basal body and associate with mitotic cells……………………………………………...66

Figure 7. Loss of Pak leads to formation of Ac-tub rich cell extensions……………………...68

Figure 8. Loss of PIX or Pak induces multinucleation and supernumerary centrioles………..70

Figure 9. Expression of kinase-dead Pak1 induces the formation of Ac-tub rich cell extensions which depends on Pak-PIX interaction…………………………………………...... 73

Figure 10. Expression of PIX mutant unable to bind to Pak induces Ac-tub rich cell extensions...... 75

LIST OF FIGURES (continued)

Page

CHAPTER 3.

Figure 1. Expression of active Pak1 induces formation of cyst-like lumens in two-dimensional MDCK culture……………………………………………………………...... 89

Figure 2. Formation of cyst-like lumens stimulated by Pak1-L107F depends on interaction with βPIX...... 91

Figure 3. Expression of Pak1-L107F affects LN synthesis and secretion……………………..92

Figure 4. Accumulation of LN in lumens of 14-day old cultures of Pak1-L107F-expressing cells…………………………………………………………………...93

INTRODUCTION

The Rho GTPases Rho, Rac, and Cdc42 are pleiotropic regulators of many crucial

cellular processes. As molecular switches, they interact and activate multiple downstream

effectors to trigger signaling pathways. Pak kinases are putative effectors of Rac and Cdc42, and

as multifunctional kinases, take part in a variety of biological activities. This dissertation focuses

on the Rho GTPases and Paks and their participation in specific cellular activities. The scientific

literature, as reviewed in Chapter 1, points to their well-established roles in the cytoskeleton and microtubule dynamics, and includes their involvement in the cell cycle, spindle regulation, cytokinesis, and centrosome behavior. While the research mentioned in the literature utilized several cell lines and different model systems, my work primarily focuses on functions of Pak1 specifically in renal epithelial cells. In Chapter 2, my research examines the importance of the

Rac/Cdc42 effector proteins Pak1 and PIX in centrosome behavior and cytokinesis in MDCK cells. Additionally, my results in Chapter 2 illustrate that Pak and PIX play a part in ciliogenesis, a novel function not previously described in the literature. And finally in Chapter 3, my research highlights the significance of Pak and PIX in epithelial morphogenesis. In toto, my work

supports and strengthens the notion that Paks play a part in a plethora of vital cellular functions

in epithelial cells. Ultimately, my findings suggest that Paks are important for renal biology.

CHAPTER I. General background and scope of dissertation

I. Introduction to Rho GTPases and Pak family of kinases

Background

Guanine nucleotide-binding proteins (G proteins) are important signal transducers that fall into two major classes: the heterotrimeric G proteins comprising of α, β, and γ subunits and the monomeric G proteins. The Rho GTPases are monomeric G proteins that hydrolyze guanosine triphosphate (GTP) to guanosine diphosphate (GDP). As molecular switches, they cycle between two conformational states, a GDP-bound and a GTP-bound state (see Figure 1 ) which involves either GTP hydrolysis or GTP-binding in the conserved G domain (54, 95). When

bound to GDP, they are in an inactive confirmation. Conversely when bound to GTP, they are in

the active conformational state. The activity of Rho GTPases can be governed by Guanine

nucleotide Dissociation Inhibitors (GDIs), GTPase Activating Proteins (GAPs), and Guanine

nucleotide Exchange Factors (GEFs). GDIs sequester inactive Rho GTPases and inhibit the

dissociation of GDP from Rho proteins, and thus, prevent GTP activation via GEFs and

interactions with effector molecules. Further, GDIs regulate the cycling of the Rho GTPases

between the cytosol and the plasma membrane; whereupon, Rho GTPases can be activated. In

humans, three Rho GDIs have been identified that are differentially expressed in different

tissues. They are: the ubiquitously expressed RhoGDI, the hematopoietic cell-selective D4GDI, and RhoGDIγ –which is expressed in the brain, lung, and testis (36). GAPs stimulate the intrinsic GTP hydrolase activity of GTPases, thereby promoting the hydrolysis of GTP to GDP, and thus, favor the GDP-bound form of the GTPase. Over 70 RhoGAPs have been identified

from several different model systems (e.g., human, mouse, and fly) (124). Some examples

include: p50RhoGAP, BPGAP1, Bcr, ARAP proteins (ARAP1, ARAP2, and ARAP3), and

srGAP proteins (srGAP1, srGAP2, srGAP3). Several RhoGAPs are specific to only one Rho

GTPase member, whereas others are more promiscuous. Moreover for some GAPs, the Rho

GTPase specificity changes depending whether it’s in vitro or in vivo (124). In contrast, GEFs

catalyze the nucleotide exchange from GDP to GTP, and thus, promote the GTP-bound state.

Two distinct families of GEFs for Rho GTPases are the Dbl-related and the DOCK 180-related

GEFs. Dbl-related GEFs contain a Dbl homology (DH) domain and a Pleckstrin homology

(PH) domain that are tandemly linked. The DH domain is responsible for catalyzing the guanine

exchange activity, while the PH domain is important for the intracellular localization of the GEF

protein (39). Dock 180-related GEFs contain a Dock homology region-1 (DHR-1) domain and a

Dock homology region-2 (DHR-2) which is essential for GEF activity (32). Examples of Dbl-

related and Dock180-related GEFs include: Cool/PIX proteins (Cool-1/βPIX, Cool-2/αPIX), Sos proteins (Sos1, Sos2), Vav proteins (Vav1, Vav2, Vav3), TIAM proteins (TIAM-1, TIAM-2), and the Docks proteins (Docks1-11). GEFs can preferentially activate one specific Rho GTPase or multiple Rho GTPases. For example, Tiam-1 can activate Rac1, Rac2, and Rac3, and Vav1 can activate RhoA, Rac1, RhoG, and Cdc42 (55). On the other hand, p115RhoGEF/Lsc and

FRG only activate specifically one member, RhoA and Cdc42, respectively (39). Dock 180- related proteins are a class of GEFs specific for Rac and Cdc42 (77).

When GTPases are in the GTP-bound state, they interact and activate multiple downstream effectors to trigger complex, interconnected signaling pathways. They bind to effector proteins via GTPase-binding motifs found within the effector molecule. Examples include: the

Cdc42/Rac-interacting binding (CRIB) domain found in effector proteins such as Paks, WASP,

WASP-N, MEKK-4, MRCKα, and MRCKβ; the Rho effector homology (REM) domain found in effector proteins such as PRK1, PRK2, Rhophilin, and Rhotekin; and the ROK-kinectin

homology (RKH) domain found in Kinectin, ROKα, and ROKβ (18). Mechanistically, Rho

GTPases typically activate effector proteins by disrupting intramolecular autoinhibitory

interactions that allow exposure of functional domains within the effector molecule.

Rho GTPases are a family of small GTPases that belong to the Ras superfamily of GTPases.

In mammals, the Rho family consists of approximately twenty members that are fall into eight

subfamilies: subfamily one comprises of Cdc42, TC10, and TCL; subfamily two consists of

Rac1, Rac2, Rac3, and RhoG; subfamily three comprises of RhoA, RhoB, and RhoC; subfamily

four consists of RhoBTB1 and RhoBTB2; subfamily five comprises RIF (a.k.a. RhoF) and

RhoD; subfamily six is made up of RhoH; subfamily seven consists of CHP (a.k.a. RhoV) and

WRCH1 (a.k.a. RhoU); and finally, subfamily eight is made up of Rnd1, Rnd2, and Rnd3 (a.k.a.

RhoE) (55). Although some members are considered atypical Rho GTPases which reside mainly

in the GTP-loaded conformation (e.g., members of the RhoBTB and Rnd subfamilies), the

majority of the members are characterized as classical Rho GTPases, which cycle between the

inactive GDP-bound and active GTP-bound forms (as previously described above) (55).

Probably the most recognized and extensively studied members of the Rho family are Rho, Rac,

and Cdc42. These classical Rho GTPases are 21 kDa proteins that are well-established

regulators of the cytoskeleton, and participate in a variety of other biological activities (e.g., cell

polarity, gene transcription, and epithelial morphogenesis (43)).

Paks (p21-activated kinases) are a family of enzymes that are putative effectors of Rac and

Cdc42. Paks 1, 2, and 3 (rat αPak , γPak , and βPak respectively) were initially identified by

screening for binding partners of Rho GTPases in rat brain cytosol (83). Initial reports indicated

that these newly discovered proteins bind to Rac and Cdc42, but not Rho. The GTP-bound

active forms of Rac and Cdc42 directly interacted with these kinases, resulting in increased Pak

phosphotransferase activity, whereas the GDP-bound inactive forms did not stimulate Pak activity (83). Later studies further established that Paks were genuine Rac and Cdc42 targets (8,

67, 68, 86). Additional studies indicate that besides Rac and Cdc42, other lesser studied Rho

GTPases such as CHP, TC10, and Wrch-1 proteins also stimulate Pak activity, but not Rho family members RhoA-G or other members of the Ras superfamily (19, 156).

Paks are highly conserved 62-68 kDa serine/threonine kinases found in a plethora of

eukaryotes. Members of the Pak family have been found in yeasts (e.g., Saccharomyces

cerevisiae, Schzosaccharomyces pombe), protozoans (e.g, Acanthamoeba castellani, Entamoeba histolytica, Dictylostelium discoideum) worms (e.g., Caenorhabditis elegans), insects (e.g.,

Drospholia melanogaster), amphibians (e.g., Xenopus laevis), and mammals (e.g., Mus

musculus, Homo sapiens) (57, 122). The fact that one or more Paks can be found in simple and

higher eukaryotes illustrates their ancient origin and emphasizes the importance of these signal

transducers in several vital cellular processes.

To date, six isoforms of Pak in mammals have been discovered and classified into two

structurally and functionally distinct subfamilies, Group I/A and Group II/B Paks. Group I/A

Paks comprise Pak1, Pak2, and Pak3. As the first GTPase-regulated kinases to be identified, they are the conventional Paks (19). The non-conventional Group II/B Paks are Pak4, Pak5, and

Pak6. Emerging evidence suggests that each group is regulated differently, with members from both groups having unique and diverse functions. Moreover, the six isoforms are selectively and/or differentially expressed in various tissues (5). For example, all the Paks are expressed in the brain. Paks 1 and 5 are highly expressed in the brain, and Pak3 is predominantly expressed

5 in this tissue. Additionally, Pak1 is also highly expressed in the spleen and muscle. Paks 2 and

4 are widely expressed in most tissues, with high expression of Pak4 found in prostate, testis, and colon. Expression of Pak6 can be found in the testis, prostate, kidney, and placenta.

Structure of Pak

Overall, Group I/A and Group II/B Paks are similar in sequence and structure. Structurally, all the Pak isofoms have a regulatory N-terminal domain with proline rich regions and a catalytic C- terminal domain (see Figure 2). Both subfamilies contain a PDB (p21-binding domain)/GTPase-binding domain (CRIB) that resides within the N-regulatory domain that confers binding to active Rac and Cdc42.

Group I/A Paks have an AID (auto-inhibitory switch domain) that overlaps the PBD (5,

57). The AID mediates the dimerization of Pak molecules, maintaining them in an inactive conformation as trans-inhibited homodimers via inhibiting the catalytic domain of the other Pak molecule in the dimer (100). The PBD domain of Pak2 and Pak3 is 88% and 90% identical to the PBD domain of Pak1, respectively (5). Additionally, Paks 1-3 have several proline rich regions with canonical PxxP SH3 (Src homology 3)-binding domains, two of which bind to the adapters Nck and to Grb2, as well as a noncanonical PxP SH3-binding site for PIX (Pak- interacting exchange factor). The kinase domain of Pak2 and Pak3 is 93% and 95% identical to the kinase domain of Pak1, respectively (5).

Group II/B Paks lack a defined AID as well as a PIX binding site (5, 57). The PBD domain of Pak5 and Pak6 is 72% and 60% identical to the PBD domain of Pak4, respectively

(5). Whereas GroupI/A Paks are strongly activated upon binding to GTPases, Group II/B Paks are not appreciably activated upon binding. Such binding is thought to be more important for

localization of Group II/B Paks rather than their activation (5). The kinase domain of Pak5 and

Pak6 is approximately 54% identical to the kinase domain of Pak4 (5).

Modes of activation

Amongst the six isoforms in both groups, Pak1 is the best characterized isoform in the literature.

The X-ray crystal structure of Pak1 in the autoinhibitory conformation has been resolved (75).

In an inactive state, Pak1 exists in solution and in cells as a homodimer in which the N-terminal

regulatory domain of one Pak1 molecule binds and inhibits the C-terminal catalytic domain of

the other. Disruption of this trans-inhibited dimer results in conformation changes in Pak1 that

can induce an active-state conformation. As previously mentioned, GTPase-binding stimulates

the kinase activity of Pak1. Their binding results in the rearrangement of the AID and unfolding

of the rest of the Pak molecule. This induces dimer dissociation by removing the kinase inhibitor

(KI) region (aa 136-150) within the AID of one Pak molecule from the catalytic site of the other

Pak molecule (99). The restructuring of the kinase domain into a catalytically competent state

permits Pak as a monomer to be phosphorylated, either by autophosphorylation or by

transphosphorylation. Particularly crucial is the phosphorylation at Thr 423 in the activation

loop of the Pak1 catalytic domain, which is needed for its full catalytic function. For example,

activation can be potentially stimulated by kinases (e.g., PDK1, Akt) and by lipids at the

membrane since Rac and Cdc42, due to their prenylated C-terminus, also localize there. Other means of stimulating Pak kinase activity that are not necessarily Cdc42/Rac-dependent include protease-mediated activation, and interaction with SH3-containing proteins such as the adaptor proteins Nck and Grb2, and by PIX (Pak-interacting exchange factor).

II. Paks as multifunctional kinases

Paks have been implicated in wide range of cellular and biological activities including apoptosis

and survival signaling, hormone signaling, gene transcription, cell cycle progression, contact

inhibition and epithelial wound healing, neurogenesis, angiogenesis, and cancer metastasis (19,

41, 51, 65, 123, 125, 149, 151). As multifunctional kinases, it is implausible to give an all- encompassing review on the functions of Paks. Consequently, this dissertation will focus primarily on their role in the cytoskeleton, cell cycle progression, spindle regulation, microtubule dynamics, and centrosome behavior.

Regulation of Cytoskeleton Dynamics

Paks are probably best recognized for their role in cell motility, cytoskeletal reorganization, and their participation in focal adhesions. Focal adhesions are large protein complexes that link the cell through its cytoskeleton to the extracellular matrix (ECM). They function as

mechanosensors through integrin-mediated anchorage to the ECM. As dynamic structures, they

are constantly changing their morphology and composition in response to cell spreading and

migration. One of the first clues that suggested Paks were involved with the actin cytoskeleton,

cell motility, and focal adhesions came from experiments with mammalian fibroblasts. In this

study, microinjection of active Pak1 into quiescent Swiss 3T3 cells induced formation of

lamellopodia, filopodia, and membrane ruffling (115). Additionally, the expression of specific

Pak1 mutants in these fibroblasts induced the formation of vinculin-containing focal adhesions

(115). In another study, the expression of the Pak kinase inhibitor domain KID (Pak83-149 ; a.k.a.

AID) which inhibits Pak activity led to an accumulation of Pak in focal adhesions in fibroblasts.

Similarly, the co-expression of glutathione S-transferase (GST)-tagged KID and Flag-αPak

(Pak1) in HeLa cells led to an accumulation of αPak at vinculin-rich focal complexes. On the other hand, activation of Pak required autophosphorylation and caused focal complex disassembly (157). Numerous other studies have confirmed that Paks are involved with actin remodeling, cell motility, and modulate focal adhesions (82, 116, 151).

The mechanism by which Pak elicits these effects on the actin cytoskeleton and focal adhesions is partially dependent on its kinase activity, which can be stimulated by Rac and

Cdc42 (116). These two GTPases interact with Pak to modulate cell morphology and motility via actin dynamics. They induce the formation of actin-rich surface protrusions commonly referred to as membrane ruffles, lamellopodia, and filopodia. In addition, they temporally and spatially regulate Pak activity (100). Numerous studies have indicated, however, that Pak can elicit these effects on the cytoskeleton and focal complexes by means of independent Rac/Cdc42 and kinase-independent activation mechanisms (34, 47, 79, 93, 115, 119). In many of these cases, Pak interacts with SH3 domain-containing proteins, which bind to the proline-rich regions within the N-terminus of Pak. Examples include the adaptor proteins Nck and Grb2, as well as

PIX (157).

Many Pak functions depend on being recruited by PIX. PIX proteins are members of the

DBL (diffuse B-cell lymphoma) family of Rho GEFs. They exhibit GEF activity towards Cdc42 and Rac. There are two isoforms, αPIX (or Cool-2) and βPIX (or Cool-1), although the latter has many splice variants (110). PIX facilitates the formation of large oligomeric complexes within the cell and associates with GIT1 (G protein-coupled receptor kinase-interacting target), an Arf

GAP that localizes to focal adhesions by directly interacting with the focal adhesion protein

9 paxillin (110). In general, this PIX-GIT complex operates as a cassette in control of cell shape and polarity, which can target Paks to various intracellular sites, resulting in Pak activation (46).

Role of Paks and small GTPases in cell cycle, cytokinesis, and spindle regulation

As previously mentioned, the small Rho GTPases are pleiotropic regulators, and participate in many cellular activities. These activities, while diverse, include their role in microtubule dynamics, the cell cycle, and cytokinesis. Similarly, Paks influences these biological processes.

A brief discussion of their role in these activities is warranted since these topics are pertinent to my dissertation work.

In eukaryotes, the cell cycle is the series of events within a cell leading to its division and duplication. The cell cycle consists of five distinct phases (G0, G1, S, G2, and M) which are divided in two periods: interphase and mitosis. In interphase (G1, S, G2), the cell grows, replicates its DNA, and collects nutrients needed for mitosis. Alternatively in interphase, the cell may be quiescent (G0)—essentially nonproliferative. In mitosis (M), the division of the mother cell takes place, and splits by means of cytokinsis that result in two "daughter cells." These series of events are controlled by specific regulatory proteins, namely cyclins and cyclin- dependant kinases (CDK). In addition, the Cip/Kip and INK4a/ARF family of cyclin-dependent proteins inhibit cell cycle progression.

Studies involving modulating Rho, Rac, and Cdc42 activity have elucidated their role in

G1/S progression. In human myelomonocyctic cells, Rho was inhibited with botulinum C3 exoenzyme, which in situ ADP-ribosylates Rho. In this system, inhibition of Rho resulted in reduced proliferation and inhibition of cell division (1). Similar studies in fibroblasts showed that botulinum C3 exoenzyme inhibited RhoA and caused cell cycle arrest at the G1 phase (94).

Moreover in fibroblasts, microinjection of dominant-negative mutants of Cdc42 and Rac blocked

serum-induced DNA synthesis. Conversely, fibroblasts microinjected with the active forms of

these GTPases promoted G1 progression and DNA synthesis. While these studies point to their role in G1/S, the mechanism by which Rho GTPases accomplish this is unclear, although it

involves anchorage- or adhesion-dependent signals for proliferation and the Ras/ERK pathway

(64, 143). It is thought that Rho GTPases control the timing and levels of cyclin D1, a key

regulator of G1/S phase progression and is the first cyclin produced in the cell cycle in response

to extracellular signals. They regulate cyclin D1 expression transcriptionally or translationally by modulating ERK activity. Moreover, they regulate levels of Cdk2 inhibitors, p21cip1 and p27kip1

by promoting their degradation, or by inhibiting their transcription and/or translation (64).

Rho GTPases also regulate other stages of the cell cycle and were shown to control G2/M progression, specific stages in mitosis, and play a role in cytokinesis. HeLa cells treated with

Clostridium difficile Toxin B, a mono-glucosyltransferase that glucosylates and thereby inactivates Rho GTPases, interfered with G2/M progression. There was a delay in Ser10-histone

H3 phosphorylation as well as delayed activation of cyclin B/Cdk1 complex and Aurora A at the

centrosome (3). As for mitosis, they have important roles during prophase, prometaphase, and in

metaphase. In prophase, astral microtubules along with actin-myosin filaments, which are

regulated by Rho and its effector ROCK, are required for correct centrosome positioning. In

prometaphase, Cdc42 and mDia3 regulate microtubule attachment to kinetochores (89), which

are protein structures on chromosomes where the spindle fibers attach during division. Depletion

of mDia3 or inhibition of Cdc42 results in improper attachment of chromosomes to the spindle

microtubules and leads to mitotic arrest (145). Asymmetric division is regulated by Cdc42, the

Par proteins (PAR1-6), and atypical protein kinase C (aPKC). Such division generates daughter

cells of different sizes and specification, which is central for Caenorhabditis elegans

development and oocyte maturation, and neuroblast division in Drosophilia (43). When cells

undergo cytokinesis, Rho and its known effectors ROCK and Citron localize to the cleavage

furrow, an indentation that begins the cleavage process (64). The correct assembly and the

positioning of the contractile ring, which is made up actin and myosin II filaments, depend on

Rho activity. The contractile ring is involved in cell abscission to generate two separate

daughter cells. Rho-dependant protein kinases ROCK and Citron promote myosin activation by phosphorylating myosin light chain (MLC) which drives contraction of the actin filament ring

(64). Inhibition of the Rho and Cdc42 in mammalian cells and in Xenopus embryos disrupts contractile ring assembly (141).

Emerging evidence indicates that Rho GTPases can use Paks as downstream effectors to regulate the cell cycle. Specifically, roles of Paks in proliferation, cyclin D expression, mitotic progression, spindle formation, and cytokinesis have been proposed. Overexpression of active

Pak1 (T423E) in murine mammary glands induced hyperplasia (132). Furthermore, the mammary glands of T423 Pak1 transgenic mice had increased cyclin D1 staining in contrast to wild-type mice, suggesting a correlation between Pak1 and cyclin D1 expression (9). Upon further analysis utilizing several cell lines (e.g., breast cancer cells, HC11 cells, HeLa cells)

Balasenthil et al. (9) demonstrated that Pak1 regulates cyclin D1 promoter activity, cyclin D expression, and cyclin D transcription in a NFκB-dependent manner. In mitotic fibroblasts,

Cdc2-mediated threonine phosphorylation of Pak1 occurred at the centrosomes. Although phosphorylation at this site did not alter its kinase activity, phosphorylation of Pak1 at T212 had profound effects on mitotic organization of MTs and postmitotic cell spreading (10, 125). In breast cancer cells, activated Pak1 localized to centrosomes and centromeric regions of

chromosomes and induced multi-polar spindles after long exposure (76, 129). Moreover,

activated Pak phosphorylated histone H3 on Ser 10, a phosphorylation event tightly correlated

with chromosomal condensation and segregation and considered a marker for mitotic

progression (76). In addition, Pak1 contributes to G2/M progression by regulating polo-like kinase1 (Plk1) and Aurora A (AurA), which are mitotic kinases that play a central role in centrosome maturation and spindle formation. Active Pak1 stimulated Plk1 and AurA activity by phosphorylating them at key sites (Ser 49 for Plk1; Thr288 and Ser342 for AurA) necessary for their activation in mitosis (84, 155). Inhibition of Pak function resulted in spindles defects, and a delay in centrosome maturation and G2/M progression (84, 155). Additionally, Chung and

Firtel (30) identified and characterized a putative PAK-family gene, PAKa in Dictyostelium discodium. In Dictyostelium discodium cells at the aggregation stage, the ectopic expression of

HA tagged-PAKa localized to the cleavage furrow. On the other hand, cells which lacked Paka had defects in cytokinesis. These paka null cells were multinucleated, had defects in F-actin organization and chemotactic movement, and demonstrated a delay in development (30). And finally in Saccharomyces cerevisiae, the Pak homologue Cla4p, whose activity peaked in mitosis and was stimulated by GTP-bound Cdc42p, was also implicated in cytokinesis (14).

Role of Paks and small GTPases in microtubule dynamics

Microtubules (MTs) are hollow cylindrical filaments made up of polymerized α- and β-tubulin.

As structural components within the cell, they are involved in many cellular processes including mitosis, cytokinesis, and cell motility and are the principle structural component of cilia and flagella. As dynamic structures, MTs are in a constant state of flux, either shrinking or growing in response to external or internal cues. Catastrophe events involve a switch from MT growth to

13 shrinkage, whereas a switch from shrinkage to growth is called a rescue. MTs are organized and nucleated by microtubule-organizing centers (MOTCs). MTs are polarized and are typically bound to the MOTC at their minus-ends (-) while their dynamic plus-ends (+) are towards the cell periphery. The plus ends (+) of MTs alternate between growth and shrinkage (referred to as dynamic instability) while exploring intracellular spaces. During this search process, they are eventually captured and stabilized at target destinations within the cell. At these sites, MTs interact with numerous proteins such as microtubule-associated proteins (MAPs) and plus-end binding proteins (+ TIPs) that not only stabilize MTs but facilitate interactions with other proteins at various cellular locales such as the cell cortex. This search-and-capture of MTs is important for defining cell shape, polarity, and the orientation of intracellular organelles. For example in many cell types, MTs appear to control the polarity of migrating cells. In motile fibroblasts, the centrosome along with stable MTs preferentially reorient towards the direction of migration at the leading edge (140).

Recent work indicates that Rho GTPases influence MTs through their effectors by interacting or modifying the activities of +TIPs and MAPs. For example, the constitutive expression of active Rac or Cdc42 promotes CLIP-170 binding to the Rac/Cdc42 effector

IQGAP, which enhances MT plus-end capture (48, 134). Also, mDia binds to the plus-end binding-protein EB1 and the adenomatous polyposis coli (APC) tumor suppressor, implicating a role for Rho in MT capture (64). Furthermore, the association of APC with MTs at the plus- ends, which is important for establishing cell polarity of migrating astrocytes, is regulated by

Cdc42 via the Par6/PKCζ effector complex (42). Rac and Cdc42 regulate MT castastrophe via

Op18/stathmin, a protein which promotes depolymerization of MTs. Phosphorylation of

Op18/stathmin at any one of four distinct serine sites results in its inactivation, thereby

14 promoting MT polymerization. In HEp-2 cells, Op18/stathmin phosphorylation at Ser 16 was mediated by Rac and Cdc42. Constitutively activated Rac and Cdc42 promoted Ser 16 phosphorylation whereas inhibition of Rac and Cdc42 activity abolished or diminished phosphorylation at this specific site (35). As a common downstream target of Rac and Cdc42,

Pak was implicated in phosphorylating Op18/Stathmin.

While the roles of Rho GTPases in regulation of the actin cytoskeleton are widely known and studied, accumulating evidence points to their influence on the MT network as well. In one study, inhibition of either Rac1 or Cdc42 led to a random distribution of MTs throughout the lamella in contrast to control cells whose MTs were organized in a typical fibroblast-type radial array (52). Moreover, the inactivation of Rac and Cdc42 stabilized MT plus-ends and minus- ends, whereas constitutively activated Rac1 and Cdc42 restored the dynamic instability of MT plus-ends and minus-ends to control levels. In another study, RhoA was implicated in stabilizing microtubules. Detyrosinated tubulin or "Glu-tubulin" is post-translationally modified tubulin which accumulates in stable MTs. Lysophospatidic acid (LPA), a component of serum, promotes the rapid assembly of Glu MTs (i.e., stabilized MTs) and also activates Rho—inducing the formation of actin stress fibers and focal adhesions (31, 107). It was shown that RhoA is necessary for induction of Glu MTs by LPA. Microinjection of C3 toxin (which blocks RhoA activity) into LPA-treated fibroblasts diminished their capacity to generate Glu MTs. On the other hand, the microinjection of active RhoA into serum-starved fibroblasts promoted the formation of Glu MTs. Palazzo et al. (2001) identified mDia as the likely Rho effector responsible for the Rho-regulated assembly and organization of Glu MTs (96). In PtK1 cells, a marsupial kidney epithelial cell line, expression of constitutively active Rac1 induced lamellipodia and enhanced MT turnover and plus-end growth. In contrast, expression of

dominant negative Rac1 lacked lamellipodia, exhibited little net change in MT plus-end growth

over time and no retrograde flow of MTs (138).

Conversely, MTs can regulate the activity of the Rho GTPases. MT growth can direct

sites of actin poymerization and promote the formation of lamellipodia. MTs can be

depolymerized with the drug nocodazole, and regrown after washing out this drug. In fibroblasts

after nocodazole washout, MT growth activated Rac1 and induced the formation of lamellipodia

(135). In contrast, MT disassembly can activate Rho, resulting in myosin contractility and stress

fiber formation (40). The molecular mechanisms that link MT dynamics to Rho GTPases were

unclear until the identification of several guanine-nucleotide-exchange factors (GEFs) that not

only modulate Rho, Rac, and Cdc42 activity, but co-localize and bind to MTs. Examples include p190RhoGEF, GEF-H1, TrioGEF1, and Asef (140). For example in HeLa cells, GEF-H1

modulated cell shape and actin organization through the activation of Rho, which was MT- dependent (72). The expression of MT-associated GEF-H1 resulted in no discernible effects on overall cell morphology or the actin cytoskeleton, and GEF activity towards Rho was decreased.

On the other hand, expression of GEF-H1 mutants deficient in MT-binding caused drastic changes in cell shape, leading to an increase in the number and intensity of actin stress fibers.

Under these conditions, GEF-H1 activity was high. Furthermore, the treatment of HeLa cells with nocodazole caused alterations in cell morphology and actin organization that were similar to the changes induced by expression of the highly active MT-unbound GEF-H1 mutants. These nocodazole-induce morphological changes were inhibited by expression of Rhotekin RBD, which inhibits Rho activity, and GEF-H1(DHmut ), which acts as a dominant negative and inhibits

GEF-H1 function. Based on these observations and other supporting data, a model was proposed

on how MTs regulate GEF-H1 activity. When GEF-H1 is bound to MTs, its nucleotide

exchange activity is inactive. However, when GEF-H1 disassociates from MTs due to physiological or drug-induce MT depolymerization, its activity increases and promotes Rho activation. Consequently, activated Rho elicits changes in cell morphology and actin organization. This model suggests that MTs negatively regulate GEF-H1 activity.

Recent data indicate the involvement of Paks in MT dynamics. Paks 1 and 4 Pak directly interact with GEF-H1, a GEF whose activity is regulated by MT dynamics. Pak4 binds to GEF-

H1 through its GEF interaction domain (GID), phosphorylates GEF-H1 at key regulatory sites

(Ser 67 and 810), and mediates morphological changes in fibroblasts via GEF-H1 (24). Pak1 binds to and phosphorylates GEF-H1 at Ser 885, which promotes binding of 14-3-3 to GEF-H1; whereupon, a fraction of 14-3-3 translocates to MTs upon binding to phosphorylated GEF-H1

(152). Pak has also been found at centrosomes and spindle MTs of many cells undergoing mitosis (76, 129). In fibroblasts at metaphase, exposure of a phosphorylated peptide mimicking phosphorylated Pak1 at T212 resulted in an increase in the length of astral MTs (10).

Microinjection of the Pak1 fragment PBD/AID(H83L), which inhibits the kinase activity of Paks but cannot sequester activated Rac1, into active Rac1-expressing PtK1 cells or PtK1 cells alone reduced MT net growth and retrograde flow while increasing MT catastrophe frequencies (138).

Pak also influences the activities of proteins involved in depolymerization and polymerization of

MTs. Pak controls the activity of Op18/Stathmin, which destabilizes MTs. As previously mentioned above, phosphorylation of Op18/Stathmin is mediated by Rac and Cdc42, and phosphorylation of Op18/Stathmin disrupts its function. Whereas constitutively activated Rac and Cdc42 promoted phosphorylation of Op18/Stathmin at Ser 16, inhibition of Pak in EGF- induced HEp-2 cells expressing active Cdc42 and Rac1 prevented phosphorylation of

Op18/Stathmin at this site. This suggested that Pak activity was required for Rac1- and Cdc42-

mediated phosphorylation of Op18/Statmin at Ser 16 (35). A later study further confirmed that

Rac-mediated phosphorylation of Op18/Stathmin is Pak1-dependent and that Pak1 directly

phosphorylated Op18/Stathmin at Ser 16 in vitro, thereby inhibiting its catastrophe promoting

activity (139). Additionally, the tubulin cofactor TCoB, which aids in the assembly of tubulin heterodimers, is a physiological substrate of Pak1. Pak1 can directly phosphorylate TCoB on

Ser 65 and 128, and thus, activate it and promote its tubulin polymerization activity (130).

III. Centrosomes

General

The centrosome is considered the primary microtubule organizing center (MOTC) of most eukaryotic cells that regulates the spatial organization and nucleation of MTs. Near the nucleus, the centrosome consists of two orthogonally arranged centrioles, which are barrel-like structures made up of nine triplet MTs in a pinwheel arrangement. The older of the two centrioles is called the mother centriole whereas the daughter centriole is assembled during the previous cell cycle.

These centrioles are surrounded by the pericentriolar material (PCM), an amorphous mass containing γ-tubulin ring complexes (γTuSC), and other proteins responsible for nucleation (e.g.,

γ-tubulin, pericentrin), anchoring (e.g., ninein, centriolin, dynactin), and release (e.g., katanin) of

MTs from the centrosome (7, 16). Moreover, the centrosome houses a large and diverse number of other centrosomal proteins with multifunctional capabilities. At least 500 proteins have been identified that associate with centrosomes by mass spectroscopy, whereas a conservative estimate on the number of centrosome proteins is about 100 with about 60 being present in the interphase centrosome (2, 111). Examples of centrosome proteins are Aurora and Polo-like

kinases, which regulate centrosomal behavior and mitotic progression and includes proteins such

as Cep55, CP100, and BBS6 which are purported to regulate cytokinesis (6).

Function

As multifunctional organelles, the centrosome participates in a variety of cellular activities.

Centrosomes are involved in the organization and subcellular localization of proteasomes,

neuronal migration, and axonal growth (7). Centrosomes influence specific steps of cell

migration in neurons, particularly nucleokinesis, which is nuclear translocation into the leading

edge of migrating cells. Centrosomal-associated proteins such as lissenchephaly 1 protein

(L1S1), doublecortin (DCX), nudE nuclear-distribution gene E homologue 1 (NDE1), nudE

nuclear-distribution gene E homologue-like 1 (NDEL1), and 14-3-3ε influence MT dynamics

that promote nucleokinesis. Other proteins enriched at the centrosome such as spastin and

NA14, which regulate MTs and influence cytoskeletal dynamics, are important for axonal

elongation and targeting.

During interphase, MTs radiate from the centrosome that function as a scaffold to direct

organelle and vesicle trafficking in epithelial cells. Moreover, the mother centriole converts into

the basal body, a structure that is required for cilia formation in non-proliferating cells (see

section IV). As cells prepare for cell division, centrosome maturation takes place, a process in

which tubulin levels rise and microtubule-associated proteins are recruited for mitotic spindle

formation. The maturation of the centrosome is facilitated by kinases such as Polo and Aurora A

(21, 38, 87). During mitosis, centrosomes serve as mitotic spindle poles that mediate the proper

assembly of the bi-polar spindle, thus promoting proper alignment and equal segregation of chromosomes. They may also function in regulating cytokinesis (104).

Centrosome Duplication

Centrosomes undergo duplication once before cell division. Centrosome/centriole duplication

consists of several steps that are tightly coordinated with the cell cycle progression. Generally,

the centrosome/centriole cycle begins at the time of the G1/S transition and ends before the onset

of mitosis. This process entails centriole disengagement, duplication and elongation, and maturation and separation (16). In centriole disengagement, which occurs during mitotic exit- early G1 phase, the tight link between the two centrioles is lost. During late G1-S, a new procentriole (a precursor centriole) grows in close orthogonal association with a pre-existing

centriole and elongates. These procentrioles reach full length and mature, then finally separate,

which happens at G2-M. This whole process ensures that when cells finally undergo division,

the resulting daughter cells inherit one centrosome (consisting of two centrioles) per cell cycle

event.

Role of Paks and small GTPases in centrosome dynamics

The correct positioning of the centrosome during the cell cycle is crucial to its functions. Rho

GTPases regulate centrosome positioning during interphase and mitosis. Cdc42 along with the

GEF Tuba and the Golgi-associated GM130 form a complex that regulates centrosome

organization (69). Inhibition of GM130, Tuba, and Cdc42 induced centrosome defects in U20S

and HeLa cells. These abnormal centrosomes during interphase were unable to organize MTs

and contained multiple (>2) centrin2-positive foci on top of the nucleus in U20S cells.

Furthermore in HeLa cells, the inactivation of Cdc42 promoted disorganized spindle MTs which

resulted in multipolar spindles. In prometaphase, activated Rho and its effectors ROCK and

mDia regulate the attachment of aster MTs to the cell cortex which is crucial for centrosome

separation and spindle orientation (90). Moreover, the Rho GTPases influence centrosome

behavior via Aurora A activation. The inactivation of Rho GTPases via Clostridium difficile

Toxin B in HeLa cells resulted in the delayed activation of the centrosome protein Aurora A,

which is crucial for centrosome maturation and separation and the activation of the cyclinB/Cdk1

complex (3).

Previous work has illustrated the importance of Pak at the centrosome. It has been shown

that overexpression of active Pak1 in MCF-7 breast cancer cells led to an increase in centrosome

numbers as well as multipolar spindles (129), but the underlying mechanism was not explored in

this study. Later studies showed that Pak1 localized at the centrosomes of mitotic cells, where it

is recruited to the GIT-PIX complex (125, 155). At the centrosomes of HeLa cells, Pak2, which

is the dominant isoform in this cell line, can be both be activated by Rho GTPases (3) or in a Rho

GTPase independent manner (155). One of these studies also showed that active Pak

phosphorylated and activated Aurora A at the centrosome, whereas inhibition of Pak delayed

Aurora A accumulation in centrosomes (155). Together, these data suggest that Pak1 is an

important upstream regulator of centrosomes, and may act through Aurora A.

Consequences of centrosome malfunctioning and disease

Perturbation of centrosome function can cause dire consequences to epithelial cells.

Centrosome dysfunction can involve abnormal MT nucleation, aberrant centrosome duplication, and a failure of centrosomes to separate during mitosis. The consequences of centrosome dysfunction can give rise to centrosome amplification, spindle abnormalities (e.g., mono-polar, multipolar), and cytokinesis defects in cells. Furthermore, these incidents may induce

multinucleation as well as the abnormal segregation of chromosomes, and thus, promote

genomic instability. Karyotypic alterations due to genomic instability may lead to aneuploidy.

Errors in centrosome function that induce aneuploidy may contribute to or drive cancer

formation. Aneuploidy is the most common characteristic of human solid tumors (70).

Centrosome alterations occur frequently in many human tumor types. Cancer cell centrosomes

may have the following characteristics: an increase in centrosome number and volume,

accumulation of excess pericentrolar material, supernumerary centrioles, inappropriate

phosphorylation of centrosome proteins, and increased MT nucleation activity (71). For example,

centrosome amplification can be found in several solid human tumors in the brain, breast, lung,

colon, prostate, pancreas, bile duct, head, and neck. Centrosome abnormalities are also present in hematologic cancers such as Hodgkin's lympohomas and acute leukemias (71). Centrosome

dysfunction has also been implicated in several human ailments including neurodegenerative

diseases, autoimmune disorders, and viral or bacterial infections (71).

Finally, as will be discussed below, centrosome function is intrically linked to the function of primary cilia, and defects in either organelle are linked to the pathogenesis of cystic diseases of the kidneys. Protein products of genes that are mutated in cystic diseases of the kidney in humans, mice, and zebrafish, which are collectively referred to as cystoproteins, are expressed in cilia, basal bodies, or centrosomes (56). Examples of diseases in which cystoproteins localize to centrosomes or basal bodies include nephronophthisis and Bardet-Biedl syndrome (56).

IV. Cilia

General

Cilia are antenna-like organelles commonly found in many eukaryotic cells. They are evolutionary conserved, MT-based structures found in a board spectrum of organisms. In

vertebrates, these organelles are virtually ubiquitous and can found on the apical surface of

epithelial cells. Cilia can be motile or nonmotile, with cells possessing either one cilium

(primary cilium) or multiple cilia. Whether cells are monociliated or multiciliated depends on

the cell type. For example in humans, cells from the kidney, liver, pancreas, bone/cartilage, and

the eye are monciliated; in contrast, brain ependymal cells, and epithelial cells lining the

fallopian tubes and from the lungs are multiciliated (45). The magnitude of different cilia types

on various tissues from various species indicates that such organelles have numerous functions.

Indeed, it has been shown that cilia act as mechanosensors, environmental sensors, and play a

critical role in development (15).

Structure

Structurally, cilia can be divided into four subcompartments that include an axoneme, basal body, ciliary membrane, and ciliary tip (see Figure 3). The axoneme, the cilium core, is made up of microtubule doublets consisting of A- and B-tubules. It is sheathed in a ciliary membrane, which is continuous with the plasma membrane. In motile cilia, these nine peripheral microtubule doublets surround two central microtubule singlets. Accordingly, motile cilia are characterized as possessing a 9+2 configuration. The adjacent doublets are connected by nexin

links and held in place by radial spokes. Additionally, inner and outer dynein arms are attached

to the A-tubules and mediate cilium bending by means of interacting with tubule B. In contrast, nonmotile or primary cilia lack the central microtubule singlets, dynein arms, and radial spokes that enable ciliary movement and have a 9+0 microtubular arrangement. Despite such differences, the ciliary axoneme emanates from the basal body, a triplet microtubular structure.

Made up of A-, B-, and C-tubules, the basal body is a barrel-shaped structure that anchors the cilium to the apical surface of epithelial cells.

Function

Long ignored as being vestigial structures, primary cilia recently have been proposed to have a multiple of functions. They play roles in environmental sensing, which includes roles in photoreception and olfaction. Further, they play different roles during mammalian development,

by being involved in sonic hedgehog, Wnt, and PDGF signaling pathways. As these wide-

ranging roles are beyond the scope of this thesis, they will not be further discussed in this thesis.

Primary cilia as mechanosensors

In renal tubule cells, primary cilia act as sensors for fluid flow by monitoring the composition

and flow rate of urine in the nephron. The bending of these cilia in response to luminal fluid

flow induces an intracellular calcium signal which triggers signal transduction pathways that are

crucial in maintaining kidney tissue homeostasis (105, 126, 154). Intracellular calcium fluxes

are regulated by polycystin1 (PC1) and polycystin2 (PC2) (45, 126) which are transmembrane

proteins that are localized in primary cilia of renal epithelial cells. Together, they interact as a

mechanosensory complex that translates the bending of the cilia axoneme into signals involved

24 in regulating growth and renal tubular cell differentiation. Functional disruption of PC1 and PC2 abolishes flow sensing, deregulates calcium signaling, and promotes unregulated cell proliferation and expansion which results in the formation of cysts. Precisely how the disruption of calcium signaling leads to cyst formation is unclear. PC1 and PC2 are mutated and their functions are impaired in autosomal dominant polycystic kidney disease (ADPKD), an inherited disorder characterized by the progressive development of multiple fluid-filled cysts (126, 136).

These cysts are detrimental and can cause a decline in kidney function, and can lead to renal failure.

Primary cilia formation is linked to the centriole and cell cycle (see Figure 4)

The assembly of primary cilia relies on the mother centriole for its formation. The basal body is derived from the mother centriole and is responsible for axoneme nucleation of cilia. The assembly and disassembly of primary cilia are therefore coupled to the centriole and cell cycle

(97). Consistent with that, proteins involved in the regulation of the centrosome can also affect ciliogenesis. For instance, the centriolar proteins CP110 and Cep97 also act upon cilia.

Depletion of the centriolar protein CP110 by means of RNAi resulted in centrosome duplication defects and centrosome separation, whereas expression of a CP110 mutant that cannot be phosphorylated by cyclin-dependent kinases (CDKs) induced polyploidy (29). Additionally, this protein interacts with the centriolar protein Cep97. Knockdown of Cep97 in human U2OS osteosarcoma cells resulted in the formation of monopolar and multipolar spindles and binucleated cells implicating cytokinesis defects. Moreover, these two proteins form a complex that negatively regulates cilia formation (102, 118).

How the mother centriole transforms into the basal body is not completely understood.

Nevertheless, its correct positioning at the apical surface of epithelial cells is important for the

proper assembly of cilia. The non-canonical Wnt pathway (a.k.a. planar cell polarity pathway)

may be involved in basal body docking in cells with primary cilia since this pathway has been

implicated in basal body docking in multiciliated cells (131). The PCP pathway regulates convergent extension in vertebrates and cell polarity via Rho and Rac activation which results in reorganization of the actin cytoskeleton. In the mucociliary epithelium of Xenopus embryos,

planar cell polarity (PCP) signaling was important for the proper docking and polarization of

basal bodies to the plasma membrane; Dishevelled MO morphants had stunted cilia and

exhibited defects in the actin cytoskeleton (98). Once the basal body is properly docked at the apical surface, the assembly of the cilium can commence. This entails the elongation of the cilium axoneme and requires intraflagellar transport (IFT ), a cellular process essential for

axonemal growth and the proper maintenance of cilia, since protein synthesis does not occur

inside cilia. The construction and maintenance of cilia depend on a steady source of

components which are transported by IFT particles across the axoneme towards the ciliary tip

and back to the ciliary base by use of kinesin and dynein motors. The assembly of primary cilia

happens during interphase when cells are in a quiescent and/or differentiated state. When cells

near or approach mitosis, primary cilia are reabsorbed, a process mediated by Aurora A.

Activated Aurora A promotes primary cilia disassembly via activation/phosphorylation of

histone deacetylase 6 (HDAC6) which deacetylates the tubulin in the axoneme (106). As the

primary cilium resorbs, the centrioles replicate at S phase. With the disassembly of the cilium

and basal body, the centrioles are liberated to form centrosomes. During the G2/M phase,

centrosome maturation and separation takes place, and the two centrosomes form the bipolar

spindle. Once cells exit mitosis and enter interphase or growth arrest, primary cilia reassemble.

Diseases associated with ciliary dysfunction

Cilia dysfunction and defects associated with mutations in ciliary or basal body proteins have

been implicated in several human diseases termed ciliopathies. Examples include Bardet-Biedl

syndrome, Alstrom syndrome, Joubert syndrome, nephronophthisis, polycystic kidney disease,

polycystic liver disease, retinal degeneration, and Senior-Loken syndrome. These disorders are defined by unique clinical criteria, but many have overlapping phenotypes such as retinal degeneration, polydactyly, situs inversus, mental retardation, encephalocele, and cysts in the kidney liver, and pancreas (50).

V. Scope of dissertation

The scientific literature summarized in Chapter 1 indicates that Rho GTPases, Pak, and PIX regulate centrosome dynamics, spindles, and aspects of cell division in a variety of different cell types such as fibroblasts, and includes tumorigenic cell types such as breast cancer and HeLa cells. Given their influence on cell cycle events and centrosome behavior in these cell types, we think it is likely that Pak and PIX influences these aspects similarly in renal epithelial cells.

Given our lab’s focus on epithelial biology and interest in Pak function, we wanted to ascertain whether Pak1 as well as its binding partner PIX play a part in centrosome dynamics and cell division in Madin-Darby canine kidney cells. MDCK cells have been utilized as a model for the renal distal tubule and are commonly used to study epithelial architecture, apical-basal polarity, and epithelial morphogenesis. Further, based on a study on the centriolar proteins CP110 and

Cep97 which regulate centrosomes and ciliogenesis (118) and a report that link Pak1 and PIX to

the activation of Aurora A (33, 155), a centrosome kinase which has been implicated in

regulating cilia retraction (106), we think that Pak and PIX may not only control centrosome dynamics, but may also regulate cilia formation. Consequently (in Chapter 2), we set out to explore the potential roles of Pak and PIX in centrosome dynamics, aspects of cell division, and cilia formation in renal epithelial cells—and to possibly determine mechanistically how these proteins regulate them.

Previous studies in MDCK cells have shown that Rac1 activation is crucial for proper tubulocyst and cyst formation in 2-D and 3-D cultures respectively (91, 146). Given that Rac1 as

well as Cdc42 (85) regulate lumen formation in MDCK cells, we postulate that the putative

Rac1/Cdc42 effector Pak1is likely involved in lumen formation as well, and thus, is important

for epithelial morphogenesis. In Chapter 3, we explore the potential role of Pak as well as PIX in

lumen formation in MDCK cells.

In conclusion, we anticipate that our combined findings on Pak (from Chapters 2 and 3)

will emphasize their importance in renal biology. Additionally, since polycystic kidney disease

is associated with centrosome and cilia dysfunction, including the formation of cysts which are

detrimental to normal kidney functioning, we think our findings on Pak function may point to a

potential role in the pathogenesis of polycystic kidney disease.

Guanine exchange factors (GEFs)

Signals GEF

Rac-GDP Rac-GTP Effectors inactive active

GAP

GTPase activating proteins (GAPs)

Figure 1. Diagram of Rho GTPases (Rho, Rac, and CDC42) in their inactive and active states. Rho GTPases cycle between two conformational states, a guanosine diphosphate (GDP)- bound and a guanosine triphosphate (GTP)-bound state. When bound to GDP, they are in an inactive state. When bound to GTP, they are in the active state and interact with multiple downstream effectors to trigger complex, interconnected signaling pathways. The activity of Rho

GTPases can be governed by Guanine nucleotide Dissociation Inhibitors (GDIs), GTPase

Activating Proteins (GAPs), and Guanine nucleotide Exchange Factors (GEFs). GDIs (not shown in figure) sequester inactive Rho GTPases and inhibit the dissociation of GDP from Rho proteins, and thus, prevent GTP activation via GEFs and interactions with effector molecules.

Further, GDIs regulate the cycling of the Rho GTPases between the cytosol and the plasma membrane; whereupon, Rho GTPases can be activated. GAPs promote hydrolysis of GTP back to GDP, whereas GEFs facilitate the nucleotide exchange from GDP to GTP.

Figure 2. Schematic diagram indicating features of Pak1 structure. The structure of Pak1 consists of two basic domains, a kinase domain displayed in blue and a regulatory domain.

Within the regulatory domain, there are: the p21-binding domain (PDB) shown in grey where the active, GTP-bound Rho GTPases Rac and Cdc42 bind, an autoinhibitory domain (AID) shown in red that overlaps with the PBD, a proline-rich PIX binding domain displayed in yellow, and five canonical PXXP SH3-binding motifs shown in green.

Ciliary tip Axoneme

Ciliary membrane 9+0 (nonmotile)

9+2 (motile)

Cell membrane Basal Body

9x3

Figure 3. General structure of cilia. The structure of the cilium includes an axoneme, a basal

body, a ciliary membrane, and a ciliary tip. The axoneme, the cilium core, is made up of

microtubule doublets consisting of A- and B-tubules. It is sheathed in a ciliary membrane, which

is continuous with the plasma membrane. In motile cilia, these nine peripheral microtubule

doublets surround two central microtubule singlets and are characterized as possessing a 9+2

configuration. Nonmotile cilia have a 9+0 microtubular arrangement. The ciliary axoneme emanates from the basal body, a microtubular structure which is made up of A-, B-, and C- tubules in a 9x3 configuration. The basal body anchors the cilium to the apical surface of epithelial cells.

G0 G1

M S G2

Nucleus Centrosome Mother centriole Daughter centriole Primary cilium

Figure 4. The assembly and disassembly of primary cilia are coordinated with the centriole

and cell cycle. The assembly of primary cilia happens during G1 or G0 when cells are in a

quiescent and/or differentiated state. The assembly of primary cilia relies on the mother

centriole for its formation. The basal body is derived from the mother centriole. When cells

near or approach mitosis, primary cilia are reabsorbed and their basal bodies are disassembled.

Once the centrioles are liberated, centrioles replicate and DNA synthesis happens at S phase.

During the G2/M phase, centrosome maturation and separation takes place, and the two

centrosomes form the bipolar spindle. Once daughter cells exit mitosis and enter G1 or G0,

primary cilia reassemble.

CHAPTER II. Pak and PIX regulate ciliogenesis and cytokinesis

I. INTRODUCTION

The centrosome, consisting of two orthogonally arranged centrioles surrounded by the pericentriolar material, is the primary MOTC which regulates the spatial organization and nucleation of MTs. As mitotic spindle poles, centrosomes mediate the proper assembly of the bipolar spindle and promote the proper alignment and equal segregation of chromosomes. During interphase, the mother centriole converts into the basal body which organizes the assembly of the ciliary axoneme which is required for cilia formation.

Centrosome biogenesis and ciliogenesis are tightly coordinated with the cell cycle progression. Centriole duplication starts at the time of the G1/S transition and ends before the onset of mitosis. Miscoordination of the centrosome cycle with the cell cycle can lead to centrosome amplification, spindle abnormalities, and cytokinesis defects which can give rise to multinucleated cells.

Studies on the Rho GTPase Cdc42 have emphasized its importance in centrosome dynamics. Cdc42 along with the GEF Tuba and the Golgi-associated GM130 form a complex that regulates centrosome organization (69). Inhibition of GM130, Tuba, and Cdc42 induced centrosome defects in U20S and HeLa cells. These abnormal centrosomes during interphase were unable to organize MTs and contained multiple (>2) centrin2-positive foci on top of the nucleus in U20S cells. In HeLa cells, the inactivation of Cdc42 promoted disorganized spindle

MTs which resulted in multipolar spindles. Additionally, the inactivation of Cdc42 as well as

Rac1 and Rho via Clostridium difficile Toxin B in HeLa cells resulted in the delayed activation

33 of the centrosome protein Aurora A, which is crucial for centrosome maturation and separation and the activation of the cyclinB/Cdk1 complex (3).

Moreover, the putative Cdc42 effector Pak and Cdc42 GEF PIX influence centrosome behavior. Overexpression of active Pak1 in MCF-7 breast cancer cells led to an increase in centrosome numbers as well as multipolar spindles (129). Additionally, Paks 1 and 2 regulate spindle orientation in HeLa cells. Pak2 regulates spindles in a β-PIX and Cdc42/Rac1 dependent manner (88). Furthermore, Pak has been found at centrosomes of mitotic cells (125) and influences centrosome behavior by interacting with Aurora A (33, 155). Pak1 is recruited to the centrosome via GIT-PIX complex and is activated. Active Pak1 phosphorylates and activates

Aurora A. Inhibition of Pak and depletion of PIX delayed Aurora A accumulation in centrosomes.

Given that Cdc42, Pak, and PIX influence centrosome behavior in a variety of different cell lines, we were interested whether Pak and PIX regulate centrosomes specifically in kidney epithelial cells. We extended our interests to ciliogenesis, postulating that that these two proteins not only influence centrosome dynamics, but also regulate cilia formation. For example, the centriolar proteins CP110 and Cep97 regulate centrosomes and form a complex that negatively regulates ciliogenesis. Loss of CP110 or Cep97 results in centrosome abnormalities as well as aberrant outgrowth of cilia (118). Furthermore, Pak interacts with Aurora A, a centrosome kinase which promotes primary cilia shortening (106). We used Madin-Darby canine kidney cells (MDCK) as our model system to explore the role of Pak and PIX in centrosome behavior and ciliogenesis. In this study, we provide evidence that Pak and PIX influence centrosome behavior and cytokinesis dynamics. Additionally, we demonstrate a novel ciliary function of

Cdc42 and its effector Pak in regulating primary cilia of MDCK cells. And lastly, we

demonstrate that the Pak-PIX interaction is important for controlling the formation of cilia and

acetylated microtubules.

II. RESULTS

Overexpression of DN Cdc42 promotes ciliogenesis.

The small GTPase Cdc42 regulates aspects of the cell cycle (e.g., G1/S-phase progression), MT dynamics (e.g., cortical control of MT stability and polarization, and MT attachment to kinetochores) and centrosome organization (53, 69, 89, 94). Given its role in influencing these

aspects we were interested whether the inactivation of Cdc42 affects ciliogenesis. We first

stained cilia in control MDCK cells, using acetylated tubulin (Ac-tub) as a marker. As cilia nucleate from the mother centriole, we also stained for gamma-tubulin, which is a centrosome marker. In MDCK cells line, two centrioles are not always tightly associated and can be spaced apart up to 13 µm (23), thus allowing us to easily distinguish the two individual centrioles that comprise the centrosome. Staining with Ac-tub revealed short cilia as well as elongated dots

(Figure 1A', 1A"). Since these Ac-tub-positive dots terminated in one of the centrioles (Figure

1A, 1A", presumably the mother centriole), these dots, which we will call here "punctate cilia", indeed represent small cilia. To investigate possible effects on monocilia, we used MDCK cells that express a myc-tagged dominant-negative Cdc42 mutant, Cdc42-N17, under the control of the Tet-Off system (44, 94). In this system, expression is suppressed in the presence of 20 ng/ml doxycycline (dox). Proteins are expressed in the absence of doxycycline. We plated control cells (+dox) and cells expressing Cdc42-N17 (-dox) at confluent densities on Transwell filters. Seven days after plating, we fixed the cells and immunostained cells for myc to detect

Cdc42-N17-expressing cells and for Ac-tub. In both the control cells and cells expressing

Cdc42-N17, cilia were often short and punctate. We found that the percentage of cells that contained a primary cilium appeared higher in cells that express Cdc42-N17 (detected in red in

Figures 1C') as compared to control cells (Figure 1B). Quantification of our data confirmed this and showed a significant increase in cells with a primary cilium upon expression of Cdc42-

N17 (Figure 1D).

Cdc42 was previously been shown to be essential for progression through the G1 phase of the cell cycle as expression of Cdc42-N17 causes cell cycle arrest in fibroblasts (94). As cilia are assembled when cells become quiescent, the increased percentage of cells with a primary cilium may therefore be due to inhibition of the cell cycle by Cdc42-N17. To test this, we quantified proliferation levels in our cells with a BrdU incorporation assay. We previously showed that confluent MDCK cells establish contact inhibition of proliferation 3-6 days after plating at confluent densities. As a result of that, proliferation rates are low in monolayers that have been grown for more than 6 days (78). Consistent with these findings, we found that proliferation was low in control cells when grown for 7 days on a Transwell filter. When we expressed Cdc42-N17, we did not observe any difference in proliferation (Figure 1E). Together, these data indicate that a signaling pathway downstream of Cdc42 exists that inhibits ciliogenesis via a mechanism that is not the indirect effect of cell cycle arrest.

Pak and GIT1 localize to centrosome in MDCK cells

One of the best characterized downstream effector molecules of Cdc42 is p21-activated kinase 1

(Pak1) (83). Many of Pak1's functions depend on its ability to form a complex with βPIX. βPIX was originally identified as a guanine exchange factor (GEF) for Cdc42, but may also have

distinct roles in localizing Pak1 to specific intracellular locations (46). Both Pak1 and βPIX can directly bind to Cdc42 (142). In fibroblasts, βPIX constitutively localizes to the centrosome by interacting with GIT1, and recruits and activates Pak1 at the centrosome during mitosis (155).

We hypothesized that Cdc42 controls ciliogenesis by a mechanism that involves Pak1- and

βPIX-dependent regulation of the centrosome.

We first wanted to ascertain whether GIT1, βPIX, and Pak localize to centrosomes of kidney epithelial cells as well. We found that the expression of GFP-tagged GIT1 in MDCK cells localized to the centrosome by evidence of γ-tubulin staining, a centrosome marker which denotes the centrioles (Figure 2A). Moreover through the use of an antibody that detects phosphorylated, active Paks 1/2/3 (P-Pak), we found that the active P-Pak localizes to the centrosomes of mitotic MDCK cells, but in not cells in interphase (Figure 2B). Indeed, in

Figure 2B3, staining with the nuclear stain DAPI shows one cell on the left of the figure with condensed chromosomes, indicating that this cell is in prophase, while the cell on the right is in interphase. Note that P-Pak (Figure 2B, B3) only co-localizes with the centrosome (Figure 2B,

B2, and Figure 2C) in the cell in prophase, but not with the cell in interphase. Similar results were found in mIMCD-3 cells, a kidney epithelial cell lined derived from the murine inner medullary collecting duct cells (Michael Hunter, data not shown). P-Pak was found at the spindle poles of MDCK cells at various stages of mitosis (Figure 2D-E). Interestingly, during late stages of mitosis, P-Pak also localized at the midbody of cells undergoing cytokinesis

(Figure 2E, F). We were unable to demonstrate that βPIX localizes to centrosomes utilizing various commercially available antibodies that recognize endogenous βPIX or by means of a cell line that overexpresses myc-tagged human βPIX. This may be due to inaccessibility of the epitope, as previous studies also showed that βPIX at centrosomes could only be demonstrated

with one particular non-commercial antiserum (155). Nevertheless, our laboratory (not shown)

and others have found that most cellular PIX is constitutively associated with GIT1 in many

cells, including MDCK cells (46), and thus, we consider it likely that both GIT1 and βPIX localize to centrosomes.

Active Pak1 inhibits ciliogenesis over time

To investigate the role of Pak1 in ciliogenesis, we utilized MDCK cells that inducibly express myc-tagged Pak1-L107F. This Pak1 mutant is constitutively active because it has a point mutation in its autoinhibitory domain (AID) (22). Initial experiments suggested that active Pak1- expressing cells (detected in red) were inhibited in their ability to form cilia (Figure 3A, B). We quantitated this change by utilizing two clones from this active Pak1-expressing cell line with different expression levels (clone 7 and clone 120). Western blot analysis of lysates of cells grown in the absence or presence of dox showed that expression levels of Pak1-L107F in clone

120 were higher as compared to clone 7, but that neither clone expressed the Pak1 mutant in the presence of dox (Figure 3C). To analyze ciliogenesis in these two clones, control cells (+dox) and cells expressing active Pak1 (-dox) were plated at confluent densities on Transwells. Cells were fixed at designated time points, which spanned a total of seven days, and stained for Ac-tub and either the myc tag or an anti-Pak1 antibody that only recognized the human mutant protein.

Consistent with our previous findings that MDCK cells become quiescent and contact inhibited around 3-6 days after plating at confluent densities (78), we found that the percentage of cells with a primary cilium increased over time (Figure 3D, blue bars). Ciliogenesis was decreased in cells expressing active Pak1 as compared to control cells at all time points (Figure 3D, red bars). Additionally, we found that the higher expression levels of Pak1-L107F in clone 120

corresponded with a greater reduction of cilia as compared with clone 7, suggesting that the

decrease in cilia was dependent on expression levels of the Pak1 mutant.

Instead of forming a single monolayer as seen in control cells, we noticed that 4-7 day

old monolayers of MDCK cells expressing active Pakl formed multilayers of cells (not shown).

As this suggested an increase of proliferation we assayed proliferation levels at Day7 using

Clone 120—since this reflected the greatest difference in ciliation between active Pak1-

expressing cells and control cells. We found a significant difference in proliferation levels

between the mutant and control cells of clone 120 seven days after plating (Figure 3E) which

could partially explain the disparity in cilia formation. However, proliferation in 2 day-old cell

cultures was not different in control and induced cells, whereas in seven days we already found a

significant reduction of cilia in Pak1-L107F-expressing cells (Figure 3E). Together, our data

indicate that the decrease in cilia in Pak1-L107F cells is at least partially independent of the cell cycle.

Diminished ciliogenesis found in active Pak1 mutant cells partially depends on Pak-PIX interaction

Since Pak1 was reported to be recruited to centrosomes by interacting with βPIX, we next tested if the decrease in ciliogenesis by Pak1-L107 was dependent on its ability to interact with βPIX.

To test this, we generated a new cell line that inducibly expresses active Pak1 that cannot bind to

βPIX (Pak1 L107F, R193G, P194A; hereafter called Pak1-L107FΔPIX). Immunoprecipitation experiments confirmed that Pak1-L107F, but not Pak1-L107FΔPIX binds to βPIX (Figure 4A).

We discovered that cells with myc-tagged Pak1-L107FΔPIX do possess less cilia when compared to control cells seven days after plating. However, expression of Pak1-L107FΔPIX

had a lesser effect on cilia inhibition when compared to active Pak1-expressing cells (Figure

4B). Additionally, cells expressing Pak1-L107FΔPIX occasionally multilayered as seen in active

Pak1 cells. When we assayed these cells for proliferation using a BrdU assay, we occasionally found increased proliferation, but statistical analysis showed that these changes were not significant (Figure 4C). Together, our results suggest that inhibition of cilia is at least partially dependent on the ability of Pak1 to interact with βPIX.

Inhibition of Pak and PIX function leads to formation of Ac-tub-rich cell extensions.

Upon expression of Pak1-L107FΔPIX, we found that the inhibition of ciliogenesis is at least partially dependent on the ability of Pak1 to interact with βPIX. Next, we decided to knock down PIX in our active Pak1 cells as an alternative approach to see whether the loss of βPIX expression in our active Pak1 cells yields similar results when compared to cells expressing an active Pak1 mutant that is unable to bind to βPIX (a.k.a. Pak1-L107FΔPIX). We used a siRNA oligonucleotide to transiently knockdown canine βPIX in Pak1-L107F-expressing cells. As a control, we used a scrambled siRNA oligonucleotide. After transfection of siRNA oligonucleotides, nucleofected cells were plated confluently on Transwell filters. Two days after plating, cells were fixed and stained for the myc tag, Ac-tub, and DAPI. Unexpectantly and surprisingly, we observed a dramatic phenotype in active Pak1 cells with βPIX knockdown

(Figure 5B). These cells had unusually long extensions that stained positive for Ac-tub which were robustly seen throughout the Transwell filter in contrast to active Pak1 cells transfected with control siRNA (Figure 5A). Western blot analysis showed that we were able to efficiently knock down expression of βPIX (Figure 5C). Based off this result, we decided to investigate the role of PIX in more detail. Consequently, we used siRNA oligonucleotides to transiently

knock down canine βPIX expression in normal MDCK cells. Previously, we have shown that

knockdown of canine βPIX expression using two different siRNA oligonucleotides increased

proliferation in confluent MDCK cells. This effect could be rescued by ectopic expression of

human βPIX, which is resistant to siRNA-mediated knockdown, thus showing that the effects of the siRNA against βPIX were specific (78). To investigate the role of βPIX, we stained cells in

which βPIX was knocked down with a siRNA oligonucleotide against canine βPIX for the ciliary

marker Ac-tub (Figure 5E). As a control, we used a scrambled siRNA oligonucleotide (Figure

5D). Western blot analysis showed that we were able to efficiently knock down expression of

βPIX up to three days (Figure 5F). For this experiment, we used another parental cell MDCK

cells line (designated MDCK F7), as compared to the control we used the Tet-off system

(MDCK T23, which harbors the Tet-repressible transactivator (12)). As seen in Figure 5D, the

MDCK F7 control cell line grows longer cilia as compared to the Tet-inducible cell lines, but

otherwise behaves identical to the MDCK T23 parental line.

Examination of βPIX knockdown cells stained for Ac-tub revealed a dramatic phenotype

as well, in which many cells showed extremely long extensions that stained positive for Ac-tub

(Figure 5E). To test if these Ac-tub rich extensions represented cilia, we stained for proteins

that were previously reported to accumulate in the primary cilia in MDCK cells. These proteins

included atypical PKCzeta (44) and von Hippel-Lindau protein (112). Unfortunately, we were

unable to stain cilia using these antibodies, even in control cells. We therefore co-stained cells

for Ac-tub and gamma-tubulin to investigate if the long structures originated in one of the

centrioles. Results of these experiments showed that unlike Ac-tub staining in control cells,

which showed cilia that all originated at a centriole (Figure 6A), the Ac-tub rich long extensions

in βPIX knockdown cells generally did not originate in a centriole (Figure 6B), although we still

did observe smaller cilia that emanate from a centriole (not shown). Moreover, the Ac-tub rich

extensions were sometimes seen to associate with mitotic cells (Figure 6C). Unlike in control

cells, the centrosomes in confluent βPIX-knockdown cells often did not localize exclusively at the apical aspect of the cell. As a consequence, imaging of these centrosomes as accompanied by increased background signals, which appeared to be non-specific, but which sometimes

overlapped with Ac-tub staining. To avoid this, we also stained centrosomes in subconfluent

cells, in which centrosomes localize at one focal plane because these cells are quite flat. Thus,

even though subconfluent cells do not form cilia, they do allow for better analysis of the

centrosomes. In subconfluent cells, we found strong co-localization of Ac-tub with both

centrioles upon knockdown of βPIX (Figure 6F). This was in contrast to control cells, where we

did not observe any co-localization of Ac-tub with either centriole (Figure 6E).

As Pak function often relies on βPIX, we also tested how knockdown of Pak affected Ac- tub staining. MDCK cells express both Pak1 and Pak2, and both isoforms are able to bind βPIX.

We previously identified siRNA oligonucleotides that effectively knocked down either Pak1 or

Pak2 (78). Using these oligonucleotides, we found that Pak2 in particular, induced Ac-tub rich

extensions (Figure 7C), which appeared identical to those observed in the βPIX knockdown

cells. Cells nucleofected with a scrambled siRNA accumulated Ac-tub in small cilia (Figure B).

Upon knockdown of Pak1, the Ac-tub rich structures did also form, but they tended to be

narrower and smaller as compared to the structures in the Pak2 or βPIX knockdown cells.

Further, the aberrant Ac-tub-rich extensions we observed in Pak2 or βPIX knockdown cells were

most abundant 1-2 days after plating, whereas the structures in Pak1 knock-down cells mostly

appeared in cells that had been cultured for 3 days (Figure 7B).

Cytokinesis defects upon loss of Pak or PIX

Apart from being at cilia, Ac-tub is also found at the midbody during cytokinesis. Since we

found that Pak is activated at centrosomes as well as at the midbody, we hypothesized that Pak

and PIX may play a role in regulating the behavior of centrosomes and the central spindle during

mitosis. Cytokinesis defects will lead to bi- or multinucleated cells and supernumerary

centrosomes. To test if transient knockdown of Pak or βPIX resulted in cytokinesis defects, we

stained cells with the cell-cell adhesion marker E-cadherin to demarcate individual cells and with

gamma-tubulin to stain centrosomes. Nuclei were stained with the DNA-stain DAPI. We

introduced oligonucleotides against canine βPIX, Pak1, and Pak2 into MDCK cells by

electroporation. Cells were fixed and stained 1, 2, and 3 days after plating. To quantify

multinucleated cells we scored cells at five random microscopic fields. All individual cells that

were delineated by E-cadherin and contained more than one nucleus were scored as

multinucleated. Although electroporation by itself induced some artifactual multinucleation in

control cells, which diminished over time, knockdown of canine βPIX, Pak1 and Pak2 gave rise

to a significant increase in multinucleated when compared to cells nucleofected with control

siRNA (Figure 8A). Interestingly, even though we previously observed that Pak2 induced Ac-

tub rich extensions faster and to a larger extent as compared to Pak1 (see Figure 7), knockdown of either isoform was equally efficient in inducing multinucleation. Knockdown of both isoforms did not increase the effect. The multinucleation as induced by knockdown of βPIX could be rescued by overexpressing human βPIX (Figure 8B). For this, we knocked down canine βPIX in MDCK cells that inducibly express human βPIX under the control of the Tet-Off

System. Using this system, we were able to restore (human) βPIX levels to levels similar to those observed in control cells (Figure 8C). We were unable to perform similar rescue

experiments for Pak1 and Pak2, because the expression levels of human Pak1 in the Tet-Off

system were too heterogeneous. Further, consistent with another reported study, our laboratory

has been unable to generate cells that overexpress human Pak2 for reasons that are not

completely understood (M. ter Beest and M. Zegers, unpublished). However, knockdown of

Pak1 and/or Pak2 with two different oligonucleotides per isotype yielded identical results,

suggesting that these effects of Pak1 or Pak2 knockdown were also specific.

When we analyzed centrosomes, we found that the majority of the multinucleated cells in

the Pak- and βPIX knockdown cells also had supernumerary centrioles (Figure 8D).

Occasionally, we also observed cells with one large nucleus and supernumerary centrioles, which may suggest that these cells did not progress beyond metaphase. Together, our results suggest that loss of Pak1 or 2 or βPIX result in defects in cytokinesis. Our results further suggest that the Ac-tub rich extensions represent highly elongated central spindles between cells that have been unable to complete cytokinesis.

Pak-PIX complex is required for normal cell division

Previous work from our laboratory and that of others have suggested that the kinase-dead Pak1 mutant Pak1-K299R inhibits different endogenous group I Pak isoforms and is a potent inhibitor of the turnover of protein complexes that contain Pak1 and βPIX (62, 78). Overexpression of

this mutant can lead to inhibition of disassembly of Pak-PIX containing protein complexes and

may result in inappropriate signaling by the Pak1 N-terminal domain, while inhibiting Pak

functions that rely on its kinase function. We used MDCKs that inducibly express

haemagglutinin (HA)-tagged Pak1-K299R. Cells that express HA-tagged Pak1-K299R

multilayer and are more proliferative than controls cells (151). We plated control cells (+dox)

and HA-expressing Pak1-K299R cells at confluent densities on Transwell filters. We then fixed

the cells and then stained for Ac-tub, DAPI, and the HA tag at various time points leading up to

185 hours after plating. We found that cells expressing HA-tagged Pak1-K299R again formed

Ac-tub rich cell extensions, whereas control cells had punctate cilia (compare control cells in

Figure 9A to Pak1-K299R-expressing cells in Figures 9B, C). We examined several clones

from this Pak1-K299R-expressing cell line and found that these clones exhibited the same

phenotype (data not shown). As was the case for cells in which Pak or βPIX was knocked

down, most of the Ac-tub rich extension did not originate from a centriole, but rather connected

two or more cells (see Figure 9B). At earlier time points, many of the Ac-tub rich structures

were observed, appearing as early as 18 hours after plating. However, the formation of these

structures diminished over time, and roughly seven days after plating, the phenotype had mostly

disappeared (data not shown). As many of the Ac-tub rich extensions originated in cells with

condensed nuclei (see Figure 9B), it is likely that the formation of these structures causes

apoptosis, thus explaining the decrease of the Ac-tub rich extensions over time.

Our laboratory previously showed that the dominant-negative function of the Pak1-

K299R mutant relies on its ability to be recruited to specific intracellular sites via its interaction

with βPIX. As we showed that loss of function of either βPIX or Pak1 or Pak2 induces Ac-tub

rich structures, we tested whether the Pak1-K299R-induced phenotype depended on its

interaction with βPIX. For this, we used cells that inducibly express myc-tagged Pak1-K299R with mutations in its PIX binding domain (151) under control of the Tet-Off system. This triple mutant is Pak1 kinase-dead and cannot bind to PIX (Pak1-K299R, R193G, P194A; hereafter called Pak1-K299RΔPIX). We cultured both Pak1-K299R and Pak1-K299RΔPIX in the presence (control, Figures 9D, F) or absence (Figures 9E, G) of dox. Cells were grown for

three days, and then fixed and stained for Ac-tub and either the HA tag or human Pak1. Cells

expressing Pak1-K299RΔPIX possess punctate cilia similar to control cells (Figure 9G) and do

no longer exhibit the abnormal Ac-tub rich extensions found in Pak1-K299R-expressing cells.

Immunoprecipitation experiments confirmed that the triple mutant cannot bind to endogenous

βPIX and that kinase-dead Pak1 alone can bind to βPIX (Figure 9H).

Since the results with Pak1-K299RΔPIX-expressing cell lines suggested to us that PIX-

dependent recruitment of dominant-negative Pak1 was required for the morphological effects of this Pak1 mutant, we next wanted to examine whether PIX-dependent recruitment of endogenous

Pak was required for normal cell behavior. For this, we inducibly overexpressed a myc-tagged

βPIX mutant (βPIX-W43K) that is unable to bind endogeneous Pak1 due to alterations in its

SH3 domain. As controls, we used MDCK cells overexpressing wt (wild type; human)-βPIX, as described in Figure 8C. Immunoprecipitation experiments confirmed that the βPIX-W43K mutant cannot bind to Pak1 and that wt-βPIX can bind to endogenous Pak1 (Figure 10A).

Induced (-dox, Figures 10C, E) and non-induced (+dox, Figures 10B, D) cells were plated at confluent densities, fixed, and stained for Ac-tub, DAPI, and the myc tag. We found cells expressing wt-βPIX show punctate cilia comparable to the control cells after three days of plating (Figure 10C). Preliminary quantification of ciliated cells over time showed similar percentages between control and wt-βPIX-expressing cells (data not shown). Conversely, cells expressing mutant βPIX-W43K have Ac-tub rich extensions, that are very similar as those seen in cells that express Pak1-K299R, or cells in which Pak and βPIX was knocked down (Figure

10E). Thus, the absence of interaction of endogenous Pak1 with βPIX caused the formation of

Ac-tub rich extensions.

III. DISCUSSION

Uncoupling of ciliogenesis from cell cycle

We show that the Rho GTPase Cdc42 and its putative downstream effector Pak 1 regulate ciliogenesis in MDCK cells, a novel function for these proteins which has not been previously described in the literature. Whereas the inhibition of Cdc42 activity promotes the formation of cilia, the constitutive activation of Pak1 inhibits cilia assembly over time. Furthermore, we show that diminished ciliogenesis found in active Pak1 mutant cells partially depends on its interaction with Cdc42 GEF βPIX. In all cases, our data suggest that the expression of these mutants partially uncouples cilia formation from the cell cycle since our proliferation data suggested no real difference between the control and the mutant.

Involvement of Aurora A in Pak1-mediated regulation of cilia formation

Cdc42 and PIX are not regulating cilia formation directly. Cdc42 and PIX are likely regulating ciliogenesis indirectly by interacting with a downstream effector molecule which controls cilia formation. We suggest that Cdc42 and PIX operate through Pak1 to elicit changes on cilia.

How the putative Cdc42 effector Pak1 mechanistically mediates it effects on cilia is unknown.

Given published reports that Pak1 binds, phosphorylates, and activates Aurora A (155) and that activated Aurora A promotes primary cilia shortening (106), it is possible that active Pak inhibits ciliogenesis through the mitotic kinase Aurora A. If active Pak1 activates Aurora A, which is downstream of Pak1 and promotes cilia shortening, then inhibiting Aurora A would consequently inhibit cilia retraction. To begin to explore these notions, we first looked at whether endogenous or the active Pak 1 mutant can bind directly to Aurora A. Co-

immunopreciptation experiments failed to show a direct interaction between endogenous Pak1 or

the active Pak1 mutant, and Aurora A in Pak1-L107F MDCK cells (data not shown). We also

assayed for Aurora A activity utilizing a K-LISA Aurora Kinase Activity Kit (Calbiochem), theorizing that active Pak1 cells would have higher levels of Aurora A activity. Unfortunately, these experiments yielded highly inconsistent results from multiple experiments, and we were

therefore unable to determine whether Pak1 activated Aurora A in MDCK cells (data not

shown). To determine the effect of Aurora A inhibition on Pak1-induced changes in cilia in

MDCK cells, we utilized two approaches: siRNA targeted against Aurora A, and pharmacological Aurora A inhibitors. We used three different RNA oligonucleotides to transiently knock down Aurora A in Pak1-L107F-expressing cells. Even though we were

successful with siRNA-mediated knockdown for Pak1, Pak2 and βPIX, we were unable to knock

down Aurora A. A possible explanation is that cells that have reduced levels of Aurora A are not

viable. Additionally, the use of Aurora A inhibitors, and particularly VX-680, yielded strange

Ac-tub rich cell structures that did not resemble cilia. Together, our unsuccessful attempts to measure Aurora A activity or to successfully knock down Aurora A precluded us from determining whether Aurora A acted downstream of active Pak in terms of inhibiting cilia formation.

Interestingly at higher doses, the Aurora A inhibitor VX-680 slowed down proliferation, and induced spindle defects and multinucleated cells; however at these levels, the inhibitor is expected to be non-specific and likely inhibits Aurora A, B, and C. Nevertheless, as will be discussed below, similar phenotypes were observed upon loss of function of Pak1, Pak2 or βPIX.

At this point it is unclear if the Pak-PIX complex and Aurora act upon a common pathway, or that they independently affect mitosis.

Taking all these experiments into account, our results were inconclusive, and thus, we were unable to confirm whether Aurora A is involved in Pak1-mediated regulation of ciliogenesis in MDCK cells. Clearly, more work needs to done to delineate the pathway. Future work may require transfecting Aurora A into MDCK cells or switching to an alternative kidney epithelial cell line. One of the issues of our co-immunoprecipitation experiments was that the antibodies we have available are much more reactive against human, and to some extent, mouse

Pak and Aurora A as compared to the canine counterparts in MDCK. Thus, performing our experiments in a human or mouse kidney epithelial cell line may yield more robust results.

Speculation on possible Pak1 substrates responsible for defects in cytokinesis and supernumerary centrioles

Previous reports have implicated PIX and Pak in regulating centrosome duplication and spindles in nonpolarized, tumorigenic cells such as HeLa cells (75, 139). Given these reports, we hypothesized that PIX and Pak may influence centrosome dynamics and aspects of cell division in polarized, kidney epithelial cells. First, we wanted to ascertain whether these molecules localize to the centrosome in kidney epithelial cells, similarly to what was found in fibroblasts.

In fibroblasts, βPIX constitutively localizes to the centrosome by interacting with GIT1, and recruits and activates Pak1 at the centrosome during mitosis (155). We provide evidence that

GIT1 as well as active Pak1 localizes to the centrosome. Although we were unable to demonstrate βPIX localization at the centrosome, results from our βPIX knockdown experiments implicate a role in centrosome dynamics. The inhibition of either βPIX or Paks 1 and 2 induces multinucleation and supernumerary centrioles. These observations, including the formation of

Ac-tub rich extensions, suggest defects in cytokinesis. Mechanistically how Pak and PIX

49 participate in cytokinesis is not clear. We hypothesize that PIX by means of a GIT-PIX complex recruits Pak to the centrosome; whereupon, Pak is activated and participates in cytokinesis in mitotic cells, and thus, inhibiting Pak function at the centrosome results in defects in cytokinesis.

Several studies have implicated a role for Pak in cytokinesis (30, 76, 109). Pak 1 was found at the midbody of mitotic breast cancer cells and fibroblasts (76). Consistent with these reports, we found active Pak at the midbody of mitotic MDCK cells. The precise role that Paks play in cytokinesis is uncertain. We speculate that possible downstream targets of Pak that likely play a role are the Aurora and Polo-like family of kinases which act during anaphase to promote cytokinesis. In view of that, relevant Pak1 substrates are the centrosome kinases Polo-like kinase-1 (Plk1) and Aurora A (84, 155). Although both kinases are known to regulate centrosome maturation, chromosome segregation, and spindle formation, Plk1 has mainly been implicated in regulating cytokinesis. A role for Aurora A in cytokinesis is not well-established in the literature, rather the evidence points towards Aurora B. Aurora B is a chromosomal passenger protein, which together with INCENP, Survivin, and the protein Borealin—are part of a chromosomal passenger complex (CPC) which is purportedly required at several steps during cytokinesis in animal cells (128). However in some instances, Aurora A can phosphorylate

Aurora B substrates such as INCEP and Survivin in vitro (25). Moreover, the function of Aurora

C seems to overlap with that of Aurora B during cytokinesis (144). Nonetheless, Aurora A and

Aurora B are fundamentally distinct. Furthermore, Pak 1 and 2 have only been reported to interact with Aurora A (155), not these other isoforms.

On the other hand, Plk1 is a good candidate since reports indicate that Plk1 regulates cleavage furrow by controlling RhoA activity in mammalian cells and budding yeasts (11). In

MDCK cells, we noticed the incorrect positioning of active Rho at the cleavage furrow upon

50 expression of Pak1-K299R (Liu, F and Zegers, M, data not shown). Interestingly, Rho activity also affects HDAC6 activity, which has been implicated in cilia retraction by deacetylating the tubulin in the axoneme (106). The inhibition of Rho was correlated with an increase in MT stabilization and acetylation in osteoclasts (37). The presence of Ac-tub rich cell extensions found in our cells upon loss of PIX and Pak function may possibly be due to the inhibition of

Rho which is regulating HDAC6 activity. When cells are undergoing cytokinesis, Rho activation at the cleavage furrow is required for the correct assembly and the positioning of the contractile ring, which is involved in cell abscission. Rho-dependant protein kinases ROCK and

Citron promote myosin activation by phosphorylating myosin light chain (MLC) which drives contraction of the actin filament ring. Regulators of Rho activity during cytokinesis include the

Rho GEF Ect2 and GAP MgcRacGAP (11). Plk1 activity is required for recruitment of RhoA

GEF ECT2 to the central spindle. The inhibition of Plk1 prevents RhoA accumulation at the cortex and abolishes cleavage furrow formation. Rho-dependant protein kinases ROCK1,

ROCK2, Citron, and MYPT1 can bind to the Polo-box domain (PBD) of Plk1 in a phospho- dependent manner, suggesting that these proteins interact (80). Moreover, Plk1 along with RhoA synergistically activate ROCK2 (80).

Further, the mechanism by which Pak and PIX affect centrosome duplication is unclear.

There are two possibilities on why the loss of PIX and Paks may induce supernumerary centrioles. One possibility is that Pak and PIX may be involved in the centriole duplication process, and thus, the loss of these molecules induces aberrant duplication. However, recent studies have uncovered an immense number of proteins that regulate centrosome/centriole duplication. For example, studies involving RNA interference (RNAi) screens in

Caenorhabditis elegans have identified a group of proteins which includes ZYG-1, SPD-2, SAS-

5, SAS-6, and SAS-4 that form part of a centriole-assembly protein module that regulate the centriole cycle (16). These proteins (with the exception of SAS-5 for which no counterpart has been found for) have been identified in other model systems, stressing the importance of these proteins in centriole duplication. However, Pak and PIX have not been implicated in regulating these proteins. Secondly, the supernumerary centrioles we see upon loss of Paks and PIX may be due simply to cytokinesis failure, consequently, leading to multinucleated cells. Two likely

Pak1 substrates, Aurora A and Plk1, are probably involved with this phenotype observed in our cells. Aurora A and Plk1 are critical for centrosome maturation, and predominantly, Plk1 has been shown to be important for cytokinesis in mammals as previously mentioned above.

Ac-tub rich cell extensions are regulated by PAK-PIX complex

We show that the inhibition of Pak function leads to formation of Ac-tub-rich cell extensions.

Lengthwise, many of these Ac-tub-rich cell extensions are long, typically longer than 5 and up to

100 microns in length. Several had a branching or fragmented phenotype, including others displayed an intriguing "beads on a string" appearance. Although these structures stained for acetylated tubulin, a ciliary marker, we could not demonstrate that these cell extensions emanated from a basal body, or modified centriole via gamma-tubulin staining. The attempt to isolate these Ac-tub structures via the "peel-off method" for cilia isolation (61) and probe for proteins purportedly localized in primary cilia of MDCK cells failed. Taking these observations into account and given that transient knockdown of both molecules induces multinucleation and supernumerary centrioles in a small percentage of cells, our results suggest that these Ac-tub rich extensions are likely due to defects in cytokinesis. Interestingly, whereas the transient knockdown of Pak2 in cells resulted in these Ac-tub extensions being robustly expressed, the

abundance of these structures was much less in Pak1 knockdown cells—even though both

displayed similar levels of multinucleation. An explanation for this discrepancy is unclear;

however, it suggests that Pak2 has a larger role in the formation of these Ac-tub extensions than

Pakl. We used the kinase-dead Pak1 mutant Pak1-K299R as an alternative approach to inhibit

Pak function. This mutant is thought to act as a dominant-negative for both Pak1 and Pak2 in

MDCK cells (78). Utilizing MDCK expressing Pak1-K299R, we observed a robust presence of these Ac-tub rich extensions early on (18 hrs after plating) upon expression of this mutant.

Many of these structures associated with apoptotic cells, as they originated from cells with condensed nuclei. We also noticed that these Ac-tub rich extensions disappear over time as proliferation levels drop. We think the disappearance of these Ac-tub rich extensions is connected to the lack of multinucleation upon Pak1-K299R expression, which is contrary to what is observed with loss of Paks which induces multinucleation. We speculate that a lack of multinucleation upon Pak1-K299R overexpression is because cells unable to complete cytokinesis may die. Hence, apoptotic cells with these structures are present at early time intervals (when proliferation levels are high) and as these cells disappear so does these structures

(when proliferation decreases).

Our data clearly demonstrates that PIX plays a role in the formation of these Ac-tub rich extensions as the loss of this molecule in normal MDCK cells and in cells that express active

Pak1 induces the formation of these structures. Interestingly, our preliminary results suggest that the robustness of these Ac-tub rich extensions is enhanced upon loss of βPIX expression in active Pak1-expressing cells than in normal MDCK cells with βPIX knockdown, suggesting that the expression of active Pak1 combined with βPIX knockdown synergizes the effect.

Intriguingly, the Ac-tub rich extensions as induced by βPIX knockdown were not rescued by

overexpressing human wt-βPIX, even though multinucleation levels were rescued and

comparable to control levels. And lastly, we show that the interaction between PIX and Pak is

important. Cells expressing Pak1-K299RΔPIX possess punctate cilia similar to control cells

(Figure 6C) and abolish the abnormal Ac-tub rich extensions found in Pak1 kinase-dead cells.

Furthermore, we demonstrate that MDCK cells that express a PIX mutant that can't bind to endogenous Pak also form these Ac-tub extensions. Together, our results indicate that inhibition of formation of an endogenous Pak-PIX complex will give rise to formation of Ac-tub rich extensions. Although we cannot rule out the possibility that some of these Ac-tub rich extensions may represent aberrant cilia, our observations suggest that these structures are likely the results of defects in cytokinesis. Examination of the ultra-structure of these Ac-tub rich cell extensions via electron microscopy could provide further insights.

Conclusion

In conclusion, our work emphasizes the importance of PIX and Pak in ciliogenesis, centrosome dynamics, and cytokinesis. How these molecules regulate these aspects is uncertain; however, future work involving Plk1, Aurora A, and RhoA could provide some clues.

Figure 1. Overexpression of dominant-negative Cdc42 promotes ciliogenesis. (A-A")

Control cells, grown in the presence of dox, were plated at confluent densities on coverslips.

Three days after plating, cells were briefly extracted with CSK buffer prior to fixation in

PFA/MeOH, and stained for γ-tubulin (green), acetylated tub (Ac-tub; red), and DAPI (blue).

Notice that primary cilia (indicated by arrow in A' and A" and detected in red in A") emanate

from a centriole, most likely representing the basal body) in A". Arrowheads in A and A"

indicate centrioles. Scale bar is 10 µm. (B-C'). Control cells (B), grown in the presence of dox,

or cells that were induced to express dominant-negative Cdc42 (Cdc42-N17) by growing them in the absence of dox (C) were plated at confluent densities Transwell filters (B-C'). Cells were grown for 7 days and fixed and stained for the myc tag (red), Ac-tub (green), and DAPI (blue).

Images represent projections from confocal stacks of MDCK cells that inducibly express Cdc42-

N17. Notice that cultures of MDCK cells expressing Cdc42-N17 (C, detected in red) have more ciliated cells. Cilia are indicated by arrows in B and C. Scale bar is 10 µm. (D) Quantification of percentage of cells with primary cilia seven days after plating. n=3. Error bars represent standard deviation (SD). *P < 0.01. (E) Proliferation data of DN Cdc42 cell line. Cell proliferation was determined by BrdU incorporation 7 days after plating cells confluently on

Transwell filters. Error bars represent SD. n=4. Expression of Cdc42-N17 does not significantly change proliferation (P= 0.76).

Figure 2. GIT1 and PAK localize to centrosome. (A) A single confocal image of MDCK

cells expressing GFP-tagged GIT1. Cells were fixed with PFA and stained for γ-tubulin and

DAPI. Note that GIT1 colocalizes with γ-tubulin (arrowheads), a centrosome marker which denotes the centrioles. (B-F) Phosphorylated active Pak (P-Pak) localizes to centrosome of mitotic MDCK cells. MDCK cells were plated subconfluently on coverslips. Three days after plating, cells were fixed with PFA/MeOH and later stained for phosphorylated active Pak (green;

P-Pak), either Ac-Tub (red) or γ-tubulin (red), and DAPI (blue). Imaging was performed via confocal microscopy. (B-B3) Note that P-Pak only co-localizes with centrosomes in the cell in prophase (on the left), but not with the cell in interphase (on the right). (B) Merged images of P-

Pak, γ-tubulin, and DAPI staining. (B1) Black and white image of P-Pak staining. (B2) Black and white image of γ-tubulin staining. (B3) Black and white image of DAPI staining. (C-F)

Black and white images on the right of merged images represent P-Pak, Ac-tub, and DAPI

staining (top to bottom) within dotted areas in the merged images. C) MDCK cell in prophase

(in dotted area). P-Pak is at the centrosome. (D) MDCK cell in metaphase. P-Pak at the spindle poles. (E-F) MDCK cell undergoing cytokinesis. P-Pak is at the midbody. (E) Early cytokinesis. (F) Late cytokinesis. Scale bars in A-F are 10 µm.

Figure 3. Active Pak1 inhibits ciliogenesis over time. (A and B) Projections of MDCK cells that inducibly express Pak1-L107F. Control cells (+dox) and cells expressing active Pak1 (-dox) were plated at confluent densities on Transwell filters. Seven days after plating, cells were fixed and stained for the myc tag (red), acetylated tubulin (Ac-tub; green), and DAPI (blue). Notice that cultures of MDCK cells expressing active Pak1 (detected in red) have less ciliated cells.

Scale bar is 10 µm. (C) Control cells (+dox) and cells expressing active Pak1 (-dox) from

Clones 7 and 120 were plated at confluent densities on plastic. Cells were lysed after three days and analyzed via Western blotting. Cell lysates were stained for ectopically-expressed (human)

Pak1, myc tag, and GAPDH as a loading control. (D) Cilia Time Courses. Control cells (blue

bars) and cells expressing active Pak1 (red bars) from Clones 7 and 120 were plated on

Transwell filters at confluent densities. At the designated times (Days 1, 2, 3, 4, and 7) the cells

were fixed and then later stained for either the myc tag (Clone 7) or human Pak1 (Clone 120),

Ac-tub (marker for cilia), and DAPI. Imaging was performed via confocal microscopy, and the

number of ciliated and non-ciliated cells was counted. The average percentage of ciliated cells at

given times was quantitated. n=3. Error bars represent SD. *P < 0.05. (E) Proliferation data of

Pak1-L107F cell line (Clone 120). Cell proliferation was determined by BrdU incorporation 2 and 7 days after plating cells confluently on Transwell filters. For Day 2: n=4, P= 0.426. For

Day 7: n=4, *P < 0.02. Error bars represent SD.

Figure 4. Diminished ciliogenesis in active Pak1 mutant cells partially depends on Pak-PIX interaction. (A) Western blot of endogenous βPIX coimmunoprecipitating with Pak1-L107F but not Pak1-L107FΔPIX. Ectopically-expressed active Pak1 mutants (-dox) were immunoprecipitated with anti-c-Myc (mAb 9E10) ascites and associated β-PIX was detected by mouse monoclonal anti-β-PIX. Immunoprecipitations using anti-mouse IgG were performed in parallel as a negative control. Total lysate (TL). (B) Quantification of cells expressing either

Pak1-L107F or Pak1-L107FΔPIX with primary cilia calculated as a percentage of the control.

Control ciliated cells (blue bars) and ciliated cells expressing active Pak1 mutants (red bars).

For comparison, the percentage of ciliated cells in both controls was normalized to 100% and corresponds to 43% and 12% for the Pak1-L107F and Pak1-L107FΔPIX-expressing cells, respectively. n= 5 for Pak1-L107FΔPIX and n=3 for Pak1-L107F. *P < 0.05. Error bars represent SD. (C) Proliferation data of Pak1-L107F (Clone 120) and Pak1-L107FΔPIX cell line.

Cell proliferation was determined by BrdU incorporation 7 days after plating cells confluently on

Transwell filters. For Pak1-L107F: n=4, *P < 0.02. Expression of Pak1-L107FΔPIX does not significantly change proliferation (n=5, P= 0.13). Error bars represent SD.

Figure 5. Loss of βPIX leads to formation of Ac-tub rich cell extensions. (A-C). Cells expressing Pak1-L107F were grown in the absence of dox to allow for mutant Pak1 expression and nucleofected with siRNA oligonucleotide against βPIX (B) or a scrambled oligonucleotide

(A, control). After nucleofection, cells were grown for two days on Transwell filters. Cells were then fixed and stained for Ac-tub and DAPI to stain nuclei (A-B). Images show projections of confocal images. C shows protein levels of myc to indicate mutant Pak1 expression and βPIX to show knockdown of βPIX. (D-F) shows untransfected MDCK F7 cells nucleofected with a scrambled siRNA control (D, F) or a siRNA oligonucleotide against βPIX (E, F). Cells were plated and stained as described in A-C. F shows Western blot of cell lysates. β-tubulin was used as a loading control. Note that all cells with βPIX knockdown have Ac-tub rich cell extensions whereas control cells have cilia. Scale bar is 10 µm.

Figure 6. Ac-tub rich cell extensions induced by βPIX knockdown do not emanate from a

basal body and associate with mitotic cells. (A-C) Projections of cells transfected with control siRNA or βPIX siRNA. Scale bar is 10 µm. Nucleofected cells were plated confluently on

Transwell filters. Three days after plating, cells were briefly extracted with CSK buffer prior to

PFA/MeOH fixation, and stained for γ-tubulin (green), Ac-tub (red), and DAPI (blue). Note that cilia emanate from a centriole in control cells (A) whereas Ac-tub rich extensions induced by

βPIX knockdown do not (B). (C) An Ac-tub rich extension associating with a mitotic MDCK cell. (E-F") Single confocal images of cells transfected with control siRNA or βPIX siRNA.

Nucleofected cells were plated subconfluently on Transwell filters. Cells were briefly extracted with CSK buffer prior to fixation in PFA/MeOH and stained for γ-tubulin (green), Ac-tub (red), and DAPI (blue). Note the co-localization of Ac-tub with both centrioles upon knockdown of

βPIX in subconfluent MDCK cells (F-F"), in contrast to control cells where no co-localization was observed (E-E"). Scale bar is 10 µm.

Figure 7. Loss of Pak leads to formation of Ac-tub rich cell extensions. (A) Western blot of cell lysates nucleofected with oligonucleotides against Pak1 or Pak2, showing that MDCK cells express both isoforms, which can be knocked down effectively by siRNA. Con (Control), cells transfected with a scrambled oligonucleotide. β-tubulin was used as a loading control. (B-D')

Projections of control cells (B-B'), cells with Pak2 knockdown (C-C'), and cells with Pak1 knockdown (D-D'). Three days after plating, cells were briefly extracted with CSK buffer prior to fixation in PFA/MeOH, and stained for γ-tubulin (green), acetylated tub (Ac-tub; red), and

DAPI (blue). Notice that loss of Pak1 and Pak2 leads to formation of Ac-tub rich cell extensions. In contrast, control cells have punctate cilia. Scale bar is 10 µm.

Figure 8. Loss of PIX or Pak induces multinucleation and supernumerary centrioles. (A)

Percentage of multinucleation in MDCK cells with knockdown of βPIX, Pak1, and Pak2. Two

different oligonucleotides against Pak1 or Pak2 were used to demonstrate specificity. Percentage

of multinucleation cells one day after plating (blue bars) or two days after plating (red bars).

n=3. *P < 0.05. Error bars represent SD. (B) Percentage of multinucleation in MDCK cells that

inducibly express human βPIX with knockdown of endogenous canine βPIX. Percentages of

multinucleated cells one day after plating (blue bars) or two days after plating (red bars). Notice

that expression of human βPIX (-dox) in cells in which endogenous canine βPIX was knocked

down, multinucleation levels were rescued and comparable to control levels. n=3. *P < 0.05.

Error bars represent SD. (C) Western blots of lysates stained for polyclonal anti- βPIX antibody

that recognizes both endogenous canine and human βPIX. The top band in induced cells (-dox)

represents human βPIX, which can be detected as a slightly slower migrating band due to its myc

tag. (D) Projections of MDCK cells nucleofected with control siRNA (left image) or βPIX

siRNA (right image). Three days after plating, cells were briefly extracted with CSK buffer

prior to fixation in PFA/MeOH, and stained for E-cadherin (red), γ-tubulin (green), and DAPI

(blue). Notice that βPIX knockdown induces multinucleated cells with supernumerary centrioles

(>2 centrioles per cell). Cells with arrowheads show multinucleated cells and cells with asterisks show supernumerary centrioles. Scale bar is 10 µm.

Figure 9. Expression of kinase-dead Pak1 induces the formation of Ac-tub rich cell

extensions which depends on Pak-PIX interaction. (A-C') Projections of cells that inducibly

express Pak1-K299R. Three days after plating, cells were fixed in PFA/MeOH prior to CSK

extraction, and stained for γ-tubulin (green), Ac-tub (red), and DAPI (blue). Notice that control

cells (+dox; A-A') have cilia that emanate from a modified centriole whereas cells that express

Pak1-K299R (-dox; B-C') have Ac-tub rich cell extensions that do not originate from a centriole.

Scale bar is 10 µm. (D-G') Projections of cells that inducibly express Pak1-K299R or Pak1-

K299RΔPIX. Control cells (+dox) and cells expressing kinase-dead Pak1 mutants (-dox) were

plated at confluent densities on Transwell filters. Three days after plating, cells were fixed and

stained for either the HA tag (red) or human Pak1 (red), Ac-tub (green), and DAPI (blue).

Notice MDCK cells expressing active Pak1-K299R (E, E') have Ac-tub rich cell extensions,

indicated with arrowheads, whereas cells expressing Pak1-K299RΔPIX appear to have punctate

cilia (G, G'). Scale bar is 10 µm. (H) Western blot of endogenous βPIX

coimmunoprecipitating with Pak1-K299R but not Pak1-K299RΔPIX. Ectopically-expressed

Pak1-K299R (-dox) was immunoprecipitated with anti-HA (mAb; Clone 12CA5) ascites and

Pak1-K299RΔPIX (-dox) was immunoprecipitated with anti-c-Myc (mAb; Clone 9E10) ascites, and associated βPIX was detected by mouse monoclonal anti-β-PIX. Immunoprecipitations using anti-mouse IgG were performed in parallel as a negative control. Total lysate (TL).

Figure 10. Expression of a PIX mutant unable to bind to Pak induces Ac-tub rich cell

extensions. (A) Western blot of endogenous Pak1 coimmunoprecipitating with wt-βPIX but not

βPIX-W43K. Ectopically-expressed wt-βPIX (-dox) and βPIX-W43K (-dox) were immunoprecipitated with anti-c-Myc (mAb; Clone 9E10) ascites and associated Pak1 was detected by rabbit polyclonal anti-Pak1. Immunoprecipitations using anti-mouse IgG were

performed in parallel as a negative control. Total lysate (TL). (B-E') Projections of cells that inducibly express wt-βPIX or βPIX-W43K. Control cells (+dox) and cells expressing wt-βPIX or βPIX-W43K were plated at confluent densities on Transwell filters. Three days after plating, cells were fixed and stained for the myc tag (red), Ac-tub (green), and DAPI (blue). Notice

MDCK cells expressing βPIX-W43K (E, E') have Ac-tub rich cell extensions, indicated with arrowheads, whereas cells expressing wt-βPIX appear to have punctate cilia. Scale bar is 10 µm.

CHAPTER III. Pak and PIX control lumen formation

I. INTRODUCTION

Epithelial morphogenesis refers to the developmental processes by which epithelial cells contribute to organogenesis and body shape. Individual cells collectively coordinate their behaviors with regard to proliferation, migration, adhesion, polarization, differentiation, and death to form coherent sheets of cells. These polarized sheets of cells called epithelia, which can be one-cell thick or several cells thick, cover and protect the outside of the organism as well as form the lining of internal cavities, ducts, and organs. Moreover, epithelia function in a variety of other physiological processes including: absorption, diffusion, sensory reception, excretion, and secretion.

Numerous mammalian epithelial organs consist of typically two basic building blocks - cysts and tubules, which are lumen-enclosing monolayers of polarized epithelia (73, 92). Cysts are spherical monolayers that enclose a central lumen, whereas tubules are lumen-enclosing monolayers that are cylindrical and elongated, rather than spherical. Cells that form cysts and tubules display apical-basolateral polarity, and thus, orientate their apical surfaces towards the lumen, while their lateral membrane domains are facing neighboring cells, and their basal surfaces are towards the extracellular matrix (ECM). Lumen formation is therefore a crucial aspect of organogenesis and is important for the structure and function of epithelial organs.

Renal epithelial cells form the nephron, the functional unit of the mammalian kidney.

The nephron is a tubular structure that is responsible for filtering and excreting wastes from the blood and for producing urine. The renal epithelial cells from the nephron have primary cilia that protrude from their apical surface into the lumens of the ducts. Renal primary cilia are

77 thought to act as mechanosensors for fluid flow and are considered indispensable in maintaining kidney tissue homeostasis. In polycystic kidney disease (PKD), multiple fluid-filled cysts develop from the tubular epithelium, which hamper the healthy functioning of the kidney. PKD, which can be congenital or acquired, are a major cause of end-stage renal failure. In autosomal dominant PKD (ADPKD), an inherited form of PKD, multiple fluid-filled cysts arise from every renal tubule segments as cystic outpouchings which progressively enlarge and eventually become disconnected from the rest of the renal tubule (136). Recent studies have linked defects in the assembly and function of primary cilia, including the expression of ciliary cystoproteins, with the pathological formation of renal cysts (56, 101). Although the mechanisms that underlie cyst formation are unclear, cysts probably arise from a combination of factors including: an imbalance between cell proliferation and death, polarization defects, and defects in cell-cell and cell-matrix interactions (136).

MDCK cells as a model system

The Madin-Darby canine kidney (MDCK) cell line is an attractive in vitro model system to study epithelial and branching morphogenesis, epithelial architecture, and apical-basal polarity.

Derived from the kidney tissue of an adult female cocker spaniel, they are highly differentiated renal epithelial cells that form polarized monolayers in 2D-culture systems. MDCK cells grown on permeable supports have a basolateral domain that faces the filter and neighboring cells, and an apical surface that faces the medium. Monolayers of MDCK cells give rise to lumen- containing structures named tubulocysts when overlaid with type I collagen. In this system, application of collagen I gel overlay to the apical domain of a polarized monolayer of MDCK cells induces dramatic changes in the organization of the cells and the orientation of their

78 polarity. Specifically, the apical domain is reoriented away from the new ECM-cell contact site, and multilayered, polarized lumen-containing structures called tubulocysts are formed (114,

158). The formation of tubulocysts provides an easily tractable system in which formation of lumens can be investigated. In 3D-culture systems, lumen formation can be studied by suspending single MDCK cells in a collagen matrix. These cells form polarized cysts consisting of a single epithelial monolayer enclosing a central lumen (92). MDCK cyst morphology resembles epithelial organization in vivo. In cysts, the apical surfaces of cyst cells are towards the central lumen, while their basolateral domains face neighboring cells and the ECM.

Exposing MDCK cells to hepatocyte growth factor (HGF) induces the formation of branching tubules (147).

Role of Rho GTPases and Pak in tubulocyst and cyst formation

Three-dimensional cultures and collagen gel overlay experiments in MDCK cells have illustrated the importance of Cdc42, Rac1, and Rho in tubulocyst and cyst formation. Martin-Belmonte et al. demonstrated that depletion of Cdc42 interferes with proper lumen formation. Knockdown of

Cdc42 via siRNA in cysts caused malformation of the central lumen, resulting in mature cysts with intracellular and intercellular lumens (85). Previous work has shown that Rac1 activation is crucial for proper tubulocyst and cyst formation in MDCK cells. MDCK cells expressing dominant-negative Rac1 (Rac1-N17) overlaid with collagen fail to form tubulocysts (146).

Cysts that express Rac1-N17 have inverted apical polarity, as evidenced by the apical markers podocalyxin/gp135, which incorrectly localized to the cyst periphery (91). Additionally, these cysts lack lumens and have an abnormal organization of the basement membrane component laminin (LN) on their surfaces. It was subsequently shown that inhibiting the function of β1-

integrins via AIIB2 induced similar defects in cystogenesis, and that expression of an active form

of Rac1 was sufficient to rescue cystogenesis and lumen formation under these conditions (146).

Together, these results suggest that Rac1 is involved in cystogenesis, but the downstream

signaling is still not understood.

As there is commonly crosstalk between the Rho GTPases, in many systems, Rac and

Rho are known to antagonize each other (113). As the loss of β1-integrin or Rac1 function led to

cysts with inverted polarity, in a subsequent study, it was shown that the inversion of polarity in

MDCK cysts due to inhibition of β1-integrin signaling led to a decrease in Rac1 activity, but

resulted in elevated RhoA activity. Additionally, treating cysts with cytotoxic necrotizing factor

(CNFy), which induced RhoA activity, resulted in cysts with inverted polarity that mimicked

AIIB2-treated cysts. In contrast, it was observed that AIIB2-treated cysts with knockdown of

RhoA had normal polarity, and thus, indicated that the inverted polarity induced by AIIB2 treatment was rescued. Moreover, inhibiting or depleting RhoA kinase (ROCK), a RhoA effector, by means of a pharmacological inhibitor or by shRNA in AIIB2-treated cysts phenocopied RhoA-depleted cysts. Further, the study found that inhibiting myosin II activity via blebbistatin in AIIB2-treated cysts rescued the AIIB2 phenotype (i.e., inverted polarity), and that myosin II likely acts downstream from ROCK. And finally, it was noted that Rac1-N17-cysts treated with pharmacological agents that inhibit ROCK and myosin II activity revert cysts to normal polarity. Taking all these findings into account, Yu et al. (2008) concluded that the

RhoA-ROCK I-myosin II pathway controlled the inversion of polarity seen in cysts expressing dominant-negative Rac1 or in cysts where β1-integrin signaling was blocked by AIIB2 antibody

(148).

In previous work, we implicated Pak1 in crosstalk between cell-matrix and cell-cell

adhesion, by showing that Pak1 controls cell-cell contact mediated inhibition of migration and proliferation during MDCK scrape wound healing (78, 151). We demonstrated that Pak1 localizes to focal contacts in subconfluent cells, but is found at cell-cell contacts upon establishment of cell-cell contacts. Inhibition of endogenous Pak1 causes an accumulation of

Pak1 at focal contacts, inhibits recruitment to lateral membranes and blocks the ability of cells to undergo contact inhibition of proliferation. Based on these data we hypothesize that Pak1 is part of the mechanism by which cells sense the extracellular environment in kidney epithelial cells, and may control cellular behavior in a cell adhesion-dependent manner. Further, we showed in chapter 2 of this dissertation that the loss of Pak function leads to mitotic defects, as evidenced by the presence of multinucleated cells and supernumerary centrioles. As these findings point to a potential role in the pathogenesis of PKD, we hypothesize that Pak may play a part in cyst formation in PKD.

Given that Cdc42 and Rac1 affect lumen formation in MDCK cells and Pak is a

Rac/Cdc42 downstream effector, we investigated the role of Pak1 in lumen formation. Already, our lab has demonstrated that Pak1 plays a role in cyst formation in 3-D culture systems through the use of a kinase-dead Pak1 mutant (Michael Hunter’s work; see the Discussion section for more details). Here we show that active Pak1 plays a role in lumen formation in MDCK cells in

2-D culture systems. Additionally, we demonstrate that Pak1’s capacity to stimulate the formation of lumens depends on its interaction with PIX. Taken together, our results suggest that the Pak-PIX interaction is important in regulating cyst formation in renal epithelial cells.

II. RESULTS

Active Pak1 cells form spontaneous cyst-like lumens without collagen overlay

Given that Rac1 regulates lumen formation in 2-D and 3-D cultures of MDCK cells, we investigated the role of the putative Rac1 effector Pak1 in lumen formation. To examine a potential role for Pak1 in lumen formation, we used MDCK cells that inducibly express constitutively active Pak1 (Pak1-L107F) under the control of the Tet-off system. Six days after plating cells confluently on Transwell filters, we discovered that MDCK cells that express active

Pak1 display a dramatic phenotype. This phenotype was dissimilar to the typical polarized monolayer phenotype found in the control cells (+dox) (Figure 1A-A”). Active Pak1 cells (- dox) multilayer, with cells growing in clusters and on top of each other (Figure 1B-B”).

Furthermore, when we stained for podocalyxin/gp135, an apical marker, we noticed that these cells form spontaneous lumens without collagen overlay in 2D culture systems (Figure 1B-B”).

These lumens resemble the cyst-like lumens found in normal cells overlaid with collagen.

Interestingly, staining with the apical marker podocalyxin/gp135 revealed that the epithelial layer that covers the lumens has two apical surfaces: one that faces the medium, and the other toward the interior of the lumen (Figure 1B). Long-term culture of the cells for up to fourteen days resulted in a reorientation of the apical surface at the lumen interior to a single apical surface facing the medium (Figure 1C-C”). Together, our results show that Pak1 has the capability to trigger the formation of cyst-like lumens without the need of the ECM collagen I.

Active Pak1 requires binding to PIX to stimulate the formation of spontaneous cyst-like lumens

Since our previous work emphasized the importance of the Pak-PIX interaction (in Chapter 2) in renal epithelial cells, we looked at whether Pak’s ability to form spontaneous cyst-like lumens depends on its interaction with PIX. To investigate this notion, we used MDCK cells that inducibly express active Pak1 which is unable to bind to βPIX (i.e., Pak1-L107FΔPIX). We plated control cells (+dox) and cells that express Pak1-L107FΔPIX (-dox) confluently on

Transwell filters. After seven days, we fixed the cells and stained for podocalyxin/gp135 to visualize potential cyst-like lumens. As expected, the control cells formed a polarized monolayer and gp135 staining indicated no cyst-like lumens (Figure 2A, 2A’). Upon expression of Pak1-L107FΔPIX, we noticed that these cells multilayer similarly to active Pak1 (i.e., Pak1-

L107F) cells do (Figure 2B, 2B’). However, we observed that cells that express Pak1-

L107FΔPIX failed to form spontaneous cyst-like lumens after seven days (Figure 2B, 2B’).

These data suggest that Pak1 regulates spontaneous lumen formation by a PIX-dependent mechanism.

Expression of Pak1-L107F affects LN synthesis and secretion

Since Rac regulates the proper basement membrane (BM) assembly of laminin (LN) on MDCK cyst surfaces which is in turn crucial for lumen formation including cyst polarity (91, 146), we postulated that the putative Rac effector Pak1 plays a role in lumen formation through the BM assembly of LN. To explore this idea, we first looked at LN expression in 2-D cell culture. We plated control cells (+dox) and cells that express active Pak1 (-dox) on Transwell filters for

Western blotting and immunofluorescence microscopy. Protein lysates were collected and

stained for LN utilizing an antibody raised against LN-111. This antibody recognizes the β1-

and γ1-LN chains and most likely stains LN-511 in MDCK cells (78, 131). We normalized protein lysates to β-tubulin for analysis for one day up to six days after plating, We found that cells that express active Pak1 have lower LN levels than controls cells for all days (Figure 3C), although there was no discernible pattern between the days. Moreover upon microscopic examination, we noticed a difference in LN distribution between control cells and active Pak1 cells. LN expression was diffuse and even in control cells (Figure 3A), whereas in cells that express active Pak1—the LN expression appeared more localized and concentrated (Figure 3B).

Interestingly, we found that 14 day-old cultures of active Pak1-expressing cells had an

accumulation of LN in the lumens (Figure 4B, 4B’) in contrast to 6 day-old cultures of active

Pak1-expressing cells where only little LN accumulated into the lumens (Figure 4A, 4A’).

Altogether, these data suggests that Pak1 plays a part in lumen formation through the BM

assembly of LN.

III. DISCUSSION

Pak and LN deposition

Previous work on cysts demonstrated that proper LN assembly into the BM requires Rac1

activity (91, 146). As a putative effector of Rac1, we examined Pak1’s involvement in LN

assembly. We originally proposed that active Pak1 promotes LN assembly into the BM which in

turn is crucial for lumen formation including cyst polarity. Our data with our active Pak1 cells

suggest that there is a contribution of a Pak-LN pathway since we found a difference in LN

expression. This difference in LN may be connected with the spontaneous cyst-like lumen

formation we observed in 2-D cultures. However, we found that LN levels were lower from

active Pak1-expressing cells, suggesting that active Pak1 probably does not promote the proper

BM assembly of LN at cyst surfaces. On the other hand, where our results suggest that there is a contribution of a Pak-LN pathway in lumen formation, other reports from our lab involving work with DN-Pak1 cysts are discordant with this notion. No alterations in LN deposition were found in MDCK cysts upon expression of DN-Pak1 (Michael Hunter; (62)). Additionally, whereas there was an alteration in cyst polarity upon inhibition of Rac1, there was no change in cyst polarity in DN-Pak1 cysts. In contrast, in active Pak1 cells we found an inversion of polarity such that the apical surface of the cells which make up the spontaneous cyst-like lumens was misorientated two weeks after plating them on Transwell filters. Additionally, LN accumulated in the lumens in 14 day-old cultures of active Pak1-expressing cells, whereas in six days we only found small spots of LN in the cyst-like lumens. The LN accumulation coincided with a change in cell polarization, in that we see apical markers line the cyst interior after 6 days, but these markers disappear and localize only at the medium-apical cell interphase after 14 days. We propose that this shift in the localization of apical markers is due to the accumulation of LN into the cyst lumens as it was previously shown that the deposition of LN on surfaces causes the orientation of the apical domain towards the opposite surface (91).

Based on the data presented here and findings from our lab on kinase-dead Pak1 expression in cysts, we propose that active Pak1 and the kinase-dead Pak1 (a.k.a. DN-Pak1 or

Pak1-K299R) may act through different pathways to elicit changes involving lumen formation and cyst morphology. Whereas the expression of kinase-dead Pak1 mutant in cysts resulted in morphological changes (e.g., spontaneous branching and irregular cyst shape) probably attributed to its scaffolding function by recruiting key signaling proteins to subcellular locations

such as focal contacts (DN-Pak1 sequesters βPIX into large focal contacts and dysregulates focal adhesion turnover in many cell types), the pathway involving active Pak1 depends on its kinase activity. For example, our work suggests that the catalytic activity of Pak1 is required to alter

LN expression and deposition since expression of DN-Pak1 in cysts resulted in no discernible

changes in LN synthesis and secretion. Active Pak1 may be interacting with proteins that

regulate LN production transcriptionally or post-transcriptionally, whereas the kinase-dead Pak1

does not. Further, the changes in polarity upon expression of active Pak1 in our cyst-like lumens

suggest that active Pak1 behaves differently than DN-Pak1. DN-Pak1 cysts do not display a

change in apical-basolateral polarity, and thus, active Pak1 may interact with different proteins

through a different pathway to elicit these changes.

Lumen formation and cyst morphology are regulated by Pak-PIX complex

We show that Pak1 is involved in lumen formation and cyst morphology in MDCK cells in 2-D

as well as in 3-D culture systems. The active form of Pak1 can induce spontaneous cyst-like

lumens in the absence of collagen overlay. The formation of these spontaneous cyst-like lumens

depends on Pak’s interaction with PIX since cells that express an active Pak1 mutant that cannot

bind to PIX do not form lumens. Also, the kinase activity of Pak1 seems important for the

formation of spontaneous lumens in 2-D culture systems as the kinase-dead form of Pak1 (Pak1-

K299R; a.k.a. DN-Pak1) multilayer but do not form these spontaneous cyst-like lumens. Our lab

has found that active Pak1 cells do form normal cysts in 3D culture systems (data not shown).

On the other hand, our lab discovered that kinase-dead Pak1 cysts were aberrantly shaped and

not spherical (Michael Hunter; (62)). These cysts have a reduced ability to form clear lumens

and possess a spontaneous branching phenotype. Moreover, DN-Pak1 requires binding to PIX to

generate branches and increased contractility in 3D culture. Cells that express a DN-Pak1

mutant that can’t bind PIX do not form branching cysts, and their increase in contractility in

collagen is lost.

To date, we have not examined the effects on cysts upon expression of Pak1-L107FΔPIX,

but given the importance of this Pak-PIX complex in lumen formation and cyst morphology in 2-

D and 3-D culture systems, we think such expression is likely to have some observed effects on

cysts. Nevertheless, our data combined with other findings from our lab suggest that Pak’s

interaction with PIX is crucial for lumen formation and branching morphogenesis.

In conclusion, our results emphasize the importance of Pak and PIX in lumen formation

and cyst morphology in renal epithelial cells. Precisely how Pak and PIX regulate renal

epithelial morphogenesis is uncertain. From studies on DN-Pak1 cysts, we know that Pak1 regulates branching morphology, which depends on PIX interaction and β1-integrin signaling

(62). These studies also suggested that the Pak-PIX complex regulates myosin II activity since the expression of DN-Pak1 induced increased contractility in 3-D culture by a PIX-dependent mechanism. The exact way in which this complex affects myosin activity is unclear since our lab did not find the activation of Rho in DN-Pak1 cells in 3-D culture. Altogether, these results suggested that Pak and PIX may influence cyst formation by regulating the dynamics of the cytoskeleton and focal contacts.

Another possibility, although speculative at this point, is that the multilayering and cyst- like lumens we see upon expression of our active Pak1 mutant in MDCK cells is due to a pathway that is regulated by cAMP. In an old report, Ullrich et al. (1993) found that MDCK cells multilayer and form lumens in response to elevated cAMP levels (127). Similar to what we observe, the cAMP-induced lumens were enclosed by cells that apparently have two apical

surfaces while lacking a basal surface. cAMP also enlarged lumens in MDCK cysts (81) and

promoted the formation of intracellular and intercellular apical lumens in primary hepatocytes

and the hepatoma cell line, HepG2 (20, 150). Finally, cAMP levels are elevated in cystic tissue from various PKD models, and lowering cAMP levels with vasopressin-2 receptor antagonist

inhibits the development and progression of PKD in models for autosomal recessive polycystic

kidney disease (133). Some of the cAMP-induced effects on lumen formation are likely due to activation of transporters that eventually lead to increased chloride and fluid secretion into lumens (121). Additionally however, cAMP has clear effects on polarized trafficking and cytoskeletal rearrangements and it is possible that cAMP and Pak share the signaling pathway that drives the formation of the intercellular lumens. cAMP can activate Rac and Cdc42, and can thus potentially activate Pak1 (59). The cAMP-dependent protein kinase (112) was shown to regulate the activity and translocation of βPIX (27, 28). Further, PKA phosphorylates and inhibits Pak1 in a matrix-dependent manner in endothelial cells and fibroblasts (49, 60). At present, it remains to be seen whether cAMP and Pak act together to generate a lumen phenotype, but as is clear from the literature, there are several different possibilities for such a scenario.

Figure 1. Expression of active Pak1 induces formation of cyst-like lumens in two- dimensional MDCK culture. (A-C") MDCK cells that inducibly express the active Pak1 mutant Pak1-L107F under control of the Tet-off system were grown in the presence of dox

(control), or in the absence of dox for 3 days prior to plating to induce expression of the Pak1-

L107F mutant. Cells were then plated at confluent densities on Transwell filters and grown in the presence (+dox, A-A") or absence (-dox, B-B") for 6 or 14 days (C-C"). Cells were then fixed and stained for gp135/podocalyxin, β-catenin, and ZO-1, using mouse, rabbit, and rat antibodies, respectively. Nuclei were stained with DAPI. Panels A, A', B, B', C, and C' represent merged X-Y (horizontal) and X-Z (vertical) images of gp135 (green), β-catenin (red) and nuclei (blue). Arrowheads in A, B and C indicate the location of the X-Y planes in A', B' and C'. Panels A", B", and C" represent merged X-Z images of ZO-1 (green), β-catenin (red) and nuclei (blue). The area indicated with the dotted line in B" indicates the location of the apical lumen, as delineated by the gp135/podocalyxin staining in B. Note that cells facing the lumen in B, B' have apical domains that face both the apical medium and the enclosed lumen. In contrast, cells facing lumens in C, C' have lost the apical surface surrounding the enclosed lumen. Scale bars are 10 µm.

Figure 2. Formation of cyst-like lumens stimulated by Pak1-L107F depends on interaction

with βPIX. (A-B') MDCK cells that inducibly express the Pak1-L107FΔPIX mutant under the

control of the Tet-off system were grown in the presence of dox (control), or in the absence of

dox for 3 days prior to plating to induce expression. Cells were then plated at confluent densities

on Transwell filters and grown in the presence of dox (+dox, A-A') or absence (-dox, B-B') for six days. Cells were then fixed and stained for gp135/podocalyxin, β-catenin, and ZO-1, using mouse, rabbit, and rat antibodies respectively. Nuclei were stained with DAPI. Panels A and B represent merged X-Z images of gp135, β-catenin, and nuclei. A' and B' represent merged X-Z images of ZO-1, β-catenin, and nuclei. Scale bars are 10 µm.

Figure 3. Expression of Pak1-L107F affects LN synthesis and secretion. (A, B) Images

represent LN staining in one-day old cultures of control (A) and Pak1-L107F-expressing cells

(B). Bar is 10 µm. (C), Western blots showing that LN synthesis is diminished upon Pak-L107F expression in one- and six-day old cultures. Myc staining indicates expression of myc-tagged

Pak1-L107F in the absence of doxycycline (dox).

Figure 4. Accumulation of LN in lumens of 14-day-old cultures of Pak1-L107F-expressing cells. (A-B') MDCK cells that inducibly express the active Pak1 mutant Pak1-L107F under control of the Tet-off system, were grown for 6 or 14 days as described in the legends for Figure

1. Cells were fixed and stained for LN (green), p120 (red) using rabbit and mouse antibodies respectively. Nuclei (blue) were stained with DAPI. Panels A, A' and B, B' represent X-Z images and X-Y images respectively. Arrowheads in A and B indicate the location of the X-Y planes in A' and B'.

CHAPTER IV. Materials and methods

I. Cell culture and cell lines

Maintenance of cells

MDCK cells that inducibly express myc tagged- DN Cdc42, HA tagged- Pak1-K299R

(9.49.5.10), myc tagged- Pak1-R193G, P194A, K299R (hereafter referred to as Pak1-

K299RΔPIX), myc tagged- Pak1-L107F (Clones 7 and 120), myc tagged- wt-βPIX, myc tagged-

βPIX-W43K, and myc tagged- Pak1-L107F R193G, P194A (hereafter referred to as Pak1-

L107FΔPIX) by use of the Tet-off system were grown in modified Eagle's medium (MEM) with

L-glutamine, 10% fetal calf serum (FCS), and 1% penicillin-streptomycin (P/S) in a humidified atmosphere at 37°C in 5% CO2. Cells were maintained in growth medium with 20 ng/ml

doxycycline to suppress transgene expression. To induce transgene expression, doxycycline was

removed by several washings with PBS followed by addition of growth medium (without

doxycycline).

Generation of Pak1-L107FΔPIX cell line (Clone 135)

To generate the DNA segment encoding Pak1-L107FΔPIX, the Pak1 DNA fragment harboring

the L107F mutation in the AID was excised from the pAMtet8 vector and ligated to the Pak1

DNA fragment harboring R193G and P194A mutations in the PIX binding domain from the

pCMV6 vector. Then the DNA segment encoding Pak1-L107FΔPIX was subcloned into the

pAMtet8 vector. The DNA sequence of Pak1-L107FΔPIX was verified via the University of

Chicago Cancer Research Center-DNA Sequencing Facility (UCCRC-DSF). To generate

MDCK cells which inducibly express Pak1-L107FΔPIX, the pAMtet8 vector containing the

DNA insert encoding Pak1-L107FΔPIX was transfected into cells using the calcium phosphate co-precipitation method. In brief, 20 µg of DNA and 31 µL of 2.0 M CaCl2 were added to 0.5 ml

Hepes buffered saline in a 1.5 mL eppendorf tube. The tube was "flicked" for 20 seconds and left at room temperature for 30 minutes. MDCK cells were trypsinized off a 10 cm dish and resuspended in 1ml growth medium containing 10% FCS plus 1% P/S (i.e., normal growth medium). The cell suspension (containing 1 x 106 cells) was transferred to a 10 cm dish, and

500 µL of DNA-Ca coprecipitate was slowly added to cell suspension while agitating. The cell suspension on the plate was incubated at room temperature for 20 minutes. Then 3.5 mL of growth medium supplemented with 10% FCS, 1% P/S and 200 µM chloroquine was added to dish. The plate was agitated to spread the cells, and cells on the dish were incubated at 37° C for approximately 6 hours to permit cell attachment. The medium was removed from the plate, and cells were washed once with normal growth medium. Next, 2 mL of 15 % glycerol/HBS was added gently to the dish, and cells were incubated exactly 1 minute at 37° C. Glycerol/HBS was removed from the dish, and cells were gently washed twice with normal growth medium.

Finally, 10 mL of fresh growth medium was added to the plate, and cells were grown for three days before undergoing selection. Clones expressing Pak1-L107FΔPIX were selected using 6

µg/mL blasticidin S.

Generation of Pak1-L107F cell line (Clone 120)

To generate MDCK cells which inducibly express Pak1-L107F, the pAMtet8 vector containing the DNA insert encoding Pak1-L107F was transfected into cells using the calcium phosphate co-

precipitation method (see above for details). Clones expressing Pak1-L107F were selected using

6 µg/mL blasticidin S.

II. Western blotting (WB) and immunoprecipitation (IP)

WB analysis

Cells were lysed in lysis buffer containing1% SDS and boiled for 5 minutes at 100°C and stored

at -20°C. To prepare for SDS-PAGE, protein determination of cell lysates was done by

bicinchoninic acid (BCA) assay (Pierce). Normalized cell lysates were mixed with 4x Laemmli

buffer and boiled for 5 minutes at 100°C. Equal volumes of lysates were loaded onto 10% SDS-

polyacrylamide gels and transferred onto polyvinylidene fluoride (PVDF) membranes

(Millipore). Blots were imaged utilizing an Odyssey detector (LI-COR, Lincoln, NE) or

visualized by enhanced chemiluminescence (ECL, GE Healthcare). Secondary antibodies

utilized for Odyssey detection are Alexa Fluor IR Dye 800-conjugated or Alexa Fluor 680-

conjugated donkey anti-mouse and donkey anti-rabbit IgGs (Invitrogen). Secondary antibodies

for ECL were HRP-conjugated donkey ant-rabbit IgG (Jackson Immunoresearch Laboratories).

IPs

Cells grown on 10 cm dishes were washed twice with cold PBS, lysed in cold IP lysis buffer

(125 mM NaCl, 20 mM Hepes ph 7.4, 1% Igepal) containing protease (cocktail 1:1000, AESBF

1:1000, and aprotinin 1:1000) and phosphatase inhibitors (1mM Na3VO4, 1mM NaF), and

scraped with a plastic cell lifter (Costar® cell lifter). Cell lysates were precleared for 30 minutes

at 4°C by adding 40µl of 25% CL-4B bead slurry per sample. Beads were spun down and

supernatants were transferred to new eppendorf tubes. Protein determination of supernatants was

done by bicinchoninic acid (BCA) assay (Pierce) and cell lysates were normalized. Primary

antibodies were added to normalized cell lysates, and samples in eppendorf tubes were rotated

for 1 hour at 4° C. In addition to primary antibodies, anti- mouse IgG (H+L) and anti-rabbit IgG antibodies (Pierce Biotechnology) were also added to normalized cell lysates as a negative control. Afterwards, 40µl of 12.5% washed protein A beads was added to cell lysates. Samples were incubated and rotated overnight at 4° C. The next day, samples were washed three times with cold IP lysis buffer, the beads were spun down, and the supernatant was removed with a

Hamilton syringe. Dry beads were stored at -20°C. To prepare for SDS-PAGE, 30µl of 4X

Laemmli buffer was added to beads and boiled for 5 minutes at 100°C. Equal volumes of the supernatants were then loaded onto 10% SDS-polyacrylamide gels.

Primary Antibodies utilized for WB and IPs

The mouse monoclonal anti-c-Myc (Clone 9E10) and anti-HA (Clone 12CA5) antibodies were acquired from Keith Mostov’s lab. Rabbit anti-Pak1 antibody was from Cell Signaling. The mouse anti-β-PIX antibody was obtained from BD Transduction Laboratories. The mouse anti-

GAPDH antibody was from Biodesign.

III. siRNA

Stealth siRNA (Invitrogen) against canine β-PIX (β-PIX KD2: target sequence: taaactttgctctcactaccagttg) was utilized to knock down β-PIX expression. A scrambled Stealth oligonucleotide was used as a control. All Pak siRNAs were Stealth siRNA (Invitrogen) and

comprised the following targeting sequences: Pak1 KD1: ggaugcggcuacaucuccuauuuca; Pak1

KD-2: cagccgaagaaagagcugauuauua: Pak2 KD1: gccagaacagugggcucgauuauua; Pak2 KD2:

cagaggugguuacacggaaagcuua. Scrambled Stealth oligonucleotides were used as controls for Pak

knockdown experiments. For transient transfection, 4x106 cells were electroporated with 100 pmoles of siRNA, using an Amaxa nucleofector (program T23, buffer T).

IV. Immunofluorescence (IF) confocal microscopy

PFA fixation of TWs

Cells were plated at confluent densities on 12 mm 0.4-µm pore polycarbonate Transwell filters

(Corning-Costar). At designated times, cells on Transwells (TWs) were washed twice with PBS, and then fixed with 4% paraformaldehyde (PFA)/PBS for 15 minutes. Afterwards, 4% PFA/PBS was removed, and PBS was added to TWs in 12-well multiplates. Plates were stored at 4° C for later staining. In preparation for staining, cells on TWs were washed twice with PBS, quenched in 50 mM NH4Cl/ PBS for 15 minutes, and washed three times with PBS. Then cells were blocked and permeabilized in 5% normal donkey serum in 0.1% Triton-X100/PBS for 30

minutes. Incubation of primary antibodies at various dilutions (1:100 to 1:500) in the same

buffer was done overnight at 4° C. The following day, cells were washed three times for 5

minutes with 0.1% Triton-X100/PBS. Next, secondary antibodies were added at 1:200 dilution

in 5% normal donkey serum in 0.1% Triton-X100/PBS for 30 minutes. Secondary antibodies

utilized for indirect immunofluorescence staining were Alexa Fluor™ 488/555 anti-mouse IgG

(H+L), Alexa Fluor™ 488/555 anti-rabbit IgG (H+L), and Alexa Fluor™ 555 anti-rat IgG

(Invitrogen). Cells were then washed three more times with 0.1% Triton-X100/PBS for 5

minutes, and finally once with PBS. Sections of the polycarbonate Transwell filter were cut out

and mounted on glass slides in Fluorosave (Calbiochem) with 4'-6-diamidino-2-phenylindole

(DAPI; 1:500) to stain nuclei. Confocal image stacks (Z-stacks) of cells were taken on a Zeiss

510 LSM confocal microsocope with an Axiovert 200M microscope and a C-Apochromat

63x/1.2W Corr lens.

PFA/Methanol fixation of coverslips

Cells were plated subconfluently on coverslips in a 6-well multiplate. Three days after plating, cells were washed twice with cold PBS and fixed with cold 4% PFA/PBS for 2 minutes.

Afterwards, cells on coverslips were washed again with cold PBS and fixed for 10 minutes with

cold methanol in the freezer. Then, cells were washed thrice briefly with 0.1% Triton-

X100/PBS. Cells were blocked and permeabilized in 5% normal donkey serum in 0.1% Triton-

X100/PBS for 30 minutes. Incubation of primary antibodies at various dilutions (1:100 to 1:500)

in the same buffer was done overnight at 4° C. The following day, cells were washed three times

for 5 minutes with 0.1% Triton-X100/PBS. Next, secondary antibodies were added at 1:200

dilution in 5% normal donkey serum in 0.1% Triton-X100/PBS for 30 minutes. Secondary

antibodies utilized for indirect immunofluorescence staining were Alexa Fluor™ 555 anti-mouse

IgG (H+L) and Alexa Fluor™ 488 anti-rabbit IgG (H+L) (Invitrogen). Cells were then washed three more times with 0.1% Triton-X100/PBS for 5 minutes, and finally once with PBS. Cells on coverslips were mounted on glass slides in Fluorosave (Calbiochem) with DAPI (1:500) to stain nuclei.

Primary Antibodies utilized for IF microscopy

The mouse monoclonal anti-acetylated tubulin and anti- γ-tubulin were obtained from Sigma-

Aldrich. The rabbit anti-phospho-Pak1/2/3 antibody was obtained from Invitrogen. Rat anti- myc, rabbit anti-HA, and rabbit anti-Pak1 C-19 antibodies were acquired from Santa Cruz

Biotechnology.

V. BrdU incorporation

Addition of BrdU and PFA Fixation

Cells were plated at confluent densities on TWs in 12-well multiplates. Seven days after plating, the growth medium was removed from TWs and replaced with medium containing 50 µM 5- bromo-2-deoxyuridine (BrdU) for 3 hours at 37° C. Cells on Transwells (TWs) were washed thrice in PBS with Ca2+ and Mg2+, and then fixed with 4% paraformaldehyde/ PBS for 15

minutes. Afterwards, 4% PFA/PBS was removed, and PBS was added to TWs in 12-well

multiplates. Plates were stored at 4° C for later staining.

HCl Treatment for BrdU staining

In preparation for BrdU staining, cells were washed once with PBS, quenched in 50 mM NH4Cl/

PBS for 10 minutes, and washed thrice with PBS. To denature DNA, cells on TWs were treated

with 2N HCl for 60 minutes. HCl was neutralized with three washes of 10 mM Tris-HCl (pH

8.0). Then cells were blocked and permeabilized in 5% normal donkey serum in 0.1% Triton-

X100/PBS for 30 minutes. Cells were incubated with mouse anti-BrdU (Calbiochem, 1:100) in

5% normal donkey serum in 0.1% Triton-X100/PBS overnight at 4° C. The following day, cells

100

were washed three times for 5 minutes with 0.1% Triton-X100/PBS. Next, cells were incubated

with the secondary antibody Alexa Fluor™ 488 anti-mouse IgG (H+L) at 1:200 dilution in 5% normal donkey serum in 0.1% Triton-X100/PBS for 30 minutes. Cells were then washed three more times with 0.1% Triton-X100/PBS for 5 minutes, and finally once with PBS. Sections of the polycarbonate Transwell filter were cut out and mounted on glass slides in Fluorosave

(Calbiochem) with DAPI (1:500) to stain nuclei.

DNAse Treatment for Co-staining (BrdU and Myc tag)

Cells were washed once with PBS, quenched in 50 mM NH4Cl/ PBS for 10 minutes, and

washed thrice with PBS. Then cells were blocked and permeabilized in 5% normal donkey

serum in 0.1% Triton-X100/PBS for 30 minutes. To denature DNA and for co-staining, cells

were incubated with the primary antibodies mouse anti-Myc tag (clone 9E10; 1:500) and rat anti-

BrdU (Calbiochem, 1:100) in DNAse I solution (50 µg/mL DNAse I, 2.5 mM MgCl2, in block

buffer) overnight at 4° C. The following day, cells were washed three times for 5 minutes with

0.1% Triton-X100/PBS. Next, cells were incubated with the secondary antibodies Alexa

Fluor™ 555 anti-mouse IgG (H+L) and Alexa Fluor™ 488 anti-rat IgG at 1:200 dilution in 5% normal donkey serum in 0.1% Triton-X100/PBS for 30 minutes. Cells were then washed three more times with 0.1% Triton-X100/PBS for 5 minutes, and finally once with PBS. Sections of the polycarbonate Transwell filter were cut out and mounted on glass slides in Fluorosave

(Calbiochem) with DAPI (1:500) to stain nuclei.

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CHAPTER V. General Discussion/ Future Goals & Directions

I. General Discussion

In this dissertation work, we provide evidence that Pak and Pix regulate ciliogenesis, a novel Pak

and PIX function not previously described in the literature. The activation of Pak1 inhibits cilia

formation, and thus, reduces the number of ciliated cells. Furthermore, Pak and PIX can interact

with each other to regulate cilia formation. In addition, we demonstrate that Pak and PIX

influence centrosome behavior and regulate cytokinesis since the loss of these molecules induces

supernumerary centrioles and multinucleation. And finally, we show that Pak and PIX regulate

spontaneous lumen formation in kidney epithelial cells.

In normal tissue, epithelial cells generally arrange into coherent sheets of cells. These epithelial sheets form a barrier between the outside world and body interior, and form the tubular and glandular architecture of many organs. To maintain organ homeostasis, epithelia undergo constant renewal and repair in order to maintain epithelial architecture. This requires that epithelial cells balance important aspects such as cell proliferation, death, and motility in order to sustain tissue integrity and functionality. The disruption of epithelial architecture due to an imbalance of levels could quite possibly lead to organ system malfunctioning or failure.

In a nutshell, our work indicates that PIX and Pak regulate important aspects in renal epithelial cells. Aberrant signaling from these proteins yields mitotic defects, a loss of primary cilia, and the formation of lumens instead of a simple monolayer. For all our experiments, we used MDCK cells, which have been utilized as a model for the renal distal tubule and are typically used to study epithelial architecture, apical-basal polarity, and epithelial

102

morphogenesis. Given this notion and since the kidneys require normal cell division, proper

shape/form, and primary cilia as mechanosensors for normal maintenance and functioning of its

tissue, our work suggests that Pak and PIX are important for epithelial biology, and perhaps

kidney homeostasis.

Although our findings suggest Pak and PIX are important for renal biology, our work raises many questions. First, if Pak and PIX are negatively regulating ciliogenesis as suggested from our work, what is the mechanism of inhibition? The inhibition of cilia formation may be attributed to proliferation effects. From our findings, we observed that upon expression of active

Pak, cells were more proliferative, and thus, the absence of primary cilia in active Pak1- expressing cells may be due to proliferation effects since primary cilia are absent in proliferating cells. Our results suggest, however, that the expression of active Pak1 partially uncouples cilia formation from the cell cycle, and thus, the inhibition of cilia formation due to proliferation effects can only partially explain how Pak is negatively regulating primary cilia formation.

Other means by which Pak could inhibit cilia formation may involve its regulation of MTs— either by promoting depolymerization or polymerization of MTs via Op18/Stathmin or TCoB, respectively. Aside from Pak1 affecting MT dynamics, Pak may inhibit cilia formation by activating the mitotic kinase Aurora A, which has been implicated in regulating cilia retraction.

Taking all these factors into consideration, it is evident that more research needs to be done to understand precisely how Pak and PIX are functioning to inhibit ciliogenesis—whether it’s due to the indirect effects of proliferation, or due to independent effects involving MT dynamics and mitotic kinases, or a combination of these factors. Obviously, further investigation is needed.

Possible lines of investigation are discussed later in "Future Goals & Directions".

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Another probably related issue our research raises is the nature of the Ac-tub rich cell extensions that form when Pak-PIX complexes are inhibited either by expression of DN-Pak1 or

βPIX-W43K, or by siRNA-mediated downregulation of Pak1, Pak2, or βPIX. What do these structures represent? Clearly these long extensions are microtubule-based, but are they aberrant cilia? They do stain positive for acetylated tubulin, which is a ciliary marker. Although we cannot rule out the possibility that these long extensions—or at least that some of them—are cilia, our data suggest otherwise since the Ac-tub rich cell extensions we observed did not appear to emanate from a basal body. Our data suggest that they are likely spindle remnants due to abnormal cell division. Secondly, if these structures are mitotic spindle remnants, how does the inhibition of the Pak-PIX complex bring about the presence of these structures? Mechanistically, are these structures regulated similarly as the Ac-MTs found in cilia? For example, does it involve Pak regulation of MT dynamics via Op18/Stathmin or TCoB? Or does it entail Pak interacting with some other protein such as NAT10 (N-acteyltransferase 10), which regulates cytokinesis and the stabilization of MTs via acetylation (117)? We can only speculate at this point, but it is plausible that TCoB and Op18/Stathmin, two Pak1 substrates implicated in MT dynamics, may be involved—since both proteins influence mitotic spindles and centrosome behavior. Deregulation of TCoB and Op18/Stathmin activity results in spindle abnormalities and centrosome amplification (63, 130). On the other hand, currently, there is no connection between Pak and NAT10, although both proteins localize to the midbodies of mitotic cells (76,

117). Nevertheless, the activation of Pak1 induces the loss of Ac- MTs in cilia, whereas the loss of function of Pak or PIX gives rise to the increase of these structures. Clearly, further investigation of these structures is needed. For a discussion on the possible approaches we could take to investigate these structures, see "Future Goals & Directions".

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Thirdly, what is the physiological impact of our results? In other words, under which

(patho) physiological conditions would such structures appear? It is difficult to assess the

physiological significance of the Ac-tub cell extensions without knowing precisely what they

represent and how they are regulated. Nevertheless, our data suggest that they are likely the

results of defects in cytokinesis since the loss of Pak and PIX function induces multinucleation

and supernumerary centrioles. However, the other phenotypes we observe—multinucleation and

centrosome amplification—are associated with cancer. Multinucleation due to improper

cytokinesis may promote spindle abnormalities and genomic instability, and consequently,

induce aneuploidy which is common in human tumors (120). Also, centrosome amplification is

detected in several human tumor types, for example, such as in the brain, breast, and colon (71).

Related to this, altered Pak expression has been found in cancers of the brain, breast, and

colon. For example, Balasenthil et al. examined Pak1 expression in 60 breast cancers specimens

via immunohistochemical analysis (9). Overexpression of Pak1 was found in 34 out of 60

tumors (55%). Of the Pak1-positive tumors, cytoplasmic Pak1 staining was observed in 19

tumors (56%), nuclear staining of Pak1 was seen in 5 tumors (15%), and 10 samples had both nuclear and cytoplasmic Pak1 staining (29%). In another breast cancer study which attempted to correlate Pak1 expression and subcellular localization with tamoxifen resistance, Holm et al. examined Pak1 expression in breast tumors, and among the 403 specimens analyzed, they observed that 19% of the specimens had high cytoplasmic Pak1 expression, whereas 13% had nuclear Pak1 staining (58). Moreover in a report on Pak1 expression in normal and neoplastic

colorectal tissues, Carter et al. observed that Pak1 expression was significantly overexpressed in

cancerous colorectal tissues (26). Pak1 expression was significantly higher in adenomas,

invasive carcinomas, and lymph node metastases than normal colon tissue. Further, they

105 correlated Pak1 expression with colorectal carcinoma (CRC) progression, and found that Pak1 expression is significantly increased with disease progression in CRC. And lastly, Aoki et al. examined and observed that tissue samples of normal brain had no cytoplasmic and no/or very weak nuclear phosphorylated Pak1 staining, whereas phosphorylated Pak1 staining in tumor samples was more robustly seen in the cytoplasm and nucleus of glioblastoma cells (4).

Additionally, they found that the median overall survival was significantly shorter in patients with glioblastoma tumors with moderate/high levels of cytoplasmic phosphorylated Pak1 than in patients with no cytoplasmic phosphorylated Pak1. Given these studies mentioned above as well as our data, our research may be pertinent to cancer.

Our research may also be relevant for understanding the pathogenesis of PKD.

Centrosome amplification can be found in two major forms of PKD, autosomal dominant PKD

(ADPKD) and autosomal recessive PKD (ARPKD). For example in a recent study, Zhang et al.

(2010) found that fibrocystin/polyductin, the product of the PKHD1 gene which has been implicated in ARPKD in humans, localizes to centrosomes, mitotic spindles, and the midbodies of dividing renal epithelial cells. Moreover, inhibition of fibrocystin/polyductin function in two renal cell lines, MDCK and mIMCD3 cells, led to centrosome amplification, chromosome lagging, and multipolar spindle formation (153). Consistent with their in vitro findings, they also detected centrosome amplification in the kidneys of ARPKD patients (153). In another report, Battini et al. (2008) noticed that functionally inhibiting polycystin1, the product of the

PKD1 gene which is defective in persons afflicted with ADPKD, in IMCD3 cells led to multipolar spindles and centrosome amplification. Also, the inhibition of polycystin1 in primary human renal cells resulted in genomic instability and centrosome amplification (13).

106

Furthermore, it’s plausible that polycystin1 may indirectly affect the activity and function

of Pak1. Polycystin1 is a glycoprotein which functions as a G protein-coupled receptor, acts as a

integral membrane protein involved in cell-matrix interactions, modulates intracellular calcium fluxes via interaction with polycystin2 , and serves as an activator of transcription via regulated intramembrane proteolysis (66, 126, 137). Polycystin1 participates in several signaling pathways

such as Wnt, Jak-STAT, and G protein-coupled signaling pathways. Although subcellular localization of polycystin1 is variable and depends on the developmental stage and cell polarization of the tubular epithelium, polycystin1 has been detected in tight junctions, adherens junctions, desmosomes, focal adhesions, apical vesicles, and primary cilia (126). Besides

polycystin1 and Pak1 localizing to similar subcellular locations such as the centrosome, the

midbody, and focal adhesions, both polycystin1 and Pak1 are involved in the Jak-STAT

pathway. Polycystin1 induces STAT1 and STAT3 activation, and physically interacts and

activates Jak2 (with help from polycystin2) (17). Activated Rac stimulates Jak2 activation and

STAT dependent transcription (103). Jak2 phosphorylates Pak1 on tyrosine residues 153, 201,

and 285 and regulates Pak1 activity and function (108). Phosphorylation at these sites on wild type Pak1 decreases induced apoptosis and increases cell motility (108). Taking all these studies into account with our results, our findings may very well be pertinent to PKD.

II. Future goals & directions

Future goals will be to elucidate the mechanisms by which Pak and Pix influence centrosome behavior, cilia and lumen formation, and finally cytokinesis in kidney epithelial cells. Although it is unclear how these proteins regulate these activities mechanistically, elucidating their precise

107

role may provide valuable biological knowledge. Such insight into basic biological processes

may be fundamental in understanding several human diseases related to ciliary and centrosome

dysfunction such as PKD and other ciliopathies. Moreover, since centrosome dysfunction is

associated with tumorigenesis and altered Pak expression is seen in human tumors (74), our

results may provide basic insight into cancer as well. Ultimately, the knowledge obtained may

be used eventually to develop treatments for persons afflicted with these diseases.

Future directions will be to determine whether centrosome kinases Aurora A and/or Plk1 act downstream from Pak and PIX to influence centrosome behavior and regulate ciliogenesis and cytokinesis. Since we were unable to confirm whether Aurora A is involved in Pak1- mediated regulation of ciliogenesis in MDCK cells, future work will involve either transfecting

Aurora A into MDCK cells or switching to an alternative kidney epithelial cell line.

Additionally, future work will also entail further analysis and characterization on these Ac-tub- rich cell extensions found in our cell lines by means of electron microscopy and the use of ciliary markers. Although our data suggest that these structures are due to defects in cytokinesis, we cannot rule out the possibility that some of these are aberrant cilia, and thus, elucidating what these structures precisely represent and how they originate may provide clues into cilia and cytokinesis dynamics. In examining the role of Pak and PIX in cytokinesis, we could utilize the

Pak cell lines we have available, and determine whether Plk1 is involved by employing inhibitor studies, immunoprecipitation experiments, and kinase activity assays. Results from possible experiments may provide answers on whether the induction of supernumerary centrioles and multinucleation in MDCK cells upon the loss of Pak and PIX is due to inhibition of Plk1 activity.

Future work would also include the examination of Rho activity since cytokinesis is dependent on Plk1 activity, which promotes the recruitment of GEFs required for the activation of Rho at

108 the cleavage furrow, and since recent results from our laboratory indicate that expression of

Pak1-K299R induces an incorrect positioning of active Rho at the cleavage furrow in MDCK cells.

Additionally, future work could entail examining the roles of Pak and PIX in lumen formation and epithelial morphogenesis. We have several cell lines at our disposal (e.g., Pak and

PIX inducible knockdown, βPIX-W43K, and wt-βPIX cell lines) which could further elucidate their roles in lumen formation. Utilizing such cell lines, we could examine whether they form tubulocysts (upon collagen overlay) in 2-D culture or cysts in 3-D culture. Additionally, we could further look at the importance of the Pak-PIX interaction in cyst formation (in 3-D) upon expression of Pak1-L107FΔPIX. Results from our lab have already emphasized to some degree the importance of this Pak-PIX interaction, and thus, it is likely that future studies with this cell line, including the ones suggested above, will yield insightful results.

And lastly in possible future studies, we could collect tissue samples and make use of animal models to explore the potential involvement of Pak in PKD. We could use mouse models of polycystic kidney disease and examine Pak1 expression in the kidneys. Alternatively, we could obtain polycystic tissue samples and examine Pak1 expression and/or activation to see if a correlation exists. In addition, we could utilize transgenic mice which conditionally express mutant forms of Pak in the kidneys and examine the effects on ciliogenesis, cytokinesis, and renal epithelial development. These approaches could potentially shed some light on Pak’s role in PKD.

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