KALLIKREIN-RELATED PEPTIDASE 4 ACTIVATION

OF PROTEASE-ACTIVATED RECEPTOR FAMILY

MEMBERS AND ASSOCIATION

WITH PROSTATE CANCER

ANDREW RAMSAY

BACHELOR OF APPLIED SCIENCE (HONOURS)

THE INSTITUTE OF HEALTH AND BIOMEDICAL INNOVATION,

SCHOOL OF LIFE SCIENCES, QUEENSLAND UNIVERSITY OF TECHNOLOGY.

A THESIS SUBMITTED FOR THE DEGREE OF DOCTOR OF PHILOSOPHY OF

THE QUEENSLAND UNIVERSITY OF TECHNOLOGY

2008

KEYWORDS

Prostate cancer, Prostate cancer progression, Bone metastasis, Protease Activated Receptor (PAR), G Protein Coupled Receptor (GPCR), Kallikrein-related peptidase (KLK), Serine protease, Signal transduction, Internalisation

i ABSTRACT

Two areas of particular importance in prostate cancer progression are primary tumour development and metastasis. These processes involve a number of physiological events, the mediators of which are still being discovered and characterised. Serine proteases have been shown to play a major role in cancer invasion and metastasis. The recently discovered phenomenon of their activation of a receptor family known as the protease activated receptors (PARs) has extended their physiological role to that of signaling molecule. Several serine proteases are expressed by malignant prostate cancer cells, including members of the kallikrein- related peptidase (KLK) serine protease family, and increasingly these are being shown to be associated with prostate cancer progression.

KLK4 is highly expressed in the prostate and expression levels increase during prostate cancer progression. Critically, recent studies have implicated KLK4 in processes associated with cancer. For example, the ectopic over-expression of KLK4 in prostate cancer cell lines results in an increased ability of these cells to form colonies, proliferate and migrate. In addition, it has been demonstrated that KLK4 is a potential mediator of cellular interactions between prostate cancer cells and osteoblasts (bone forming cells). The ability of KLK4 to influence cellular behaviour is believed to be through the selective cleavage of specific substrates. Identification of relevant in vivo substrates of KLK4 is critical to understanding the patho- physiological roles of this enzyme.

Significantly, recent reports have demonstrated that several members of the KLK family are able to activate PARs. The PARs are relatively new members of the seven transmembrane domain containing G protein coupled receptor (GPCR) family. PARs are activated through proteolytic cleavage of their N-terminus by serine proteases, the resulting nascent N-terminal binds intramolecularly to initiate receptor activation. PARs are involved in a number of patho-physiological processes, including vascular repair and inflammation, and a growing body of evidence suggests roles in cancer. While expression of PAR family members has been documented in several types of cancers, including prostate, the role of these GPCRs in prostate cancer development

ii and progression is yet to be examined. Interestingly, several studies have suggested potential roles in cellular invasion through the induction of cytoskeletal reorganisation and expression of basement membrane-degrading enzymes.

Accordingly, this program of research focussed on the activation of the PARs by the prostate cancer associated enzyme KLK4, cellular processing of activated PARs and the expression pattern of receptor and agonist in prostate cancer. For these studies KLK4 was purified from the conditioned media of stably transfected Sf9 insect cells expressing a construct containing the complete human KLK4 coding sequence in frame with a V5 epitope and poly-histidine encoding sequences. The first aspect of this study was the further characterisation of this recombinant zymogen form of KLK4. The recombinant KLK4 zymogen was demonstrated to be activatable by the metalloendopeptidase thermolysin and amino terminal sequencing indicated that thermolysin activated KLK4 had the predicted N-terminus of mature active KLK4 (31IINED). Critically, removal of the pro-region successfully generated a catalytically active enzyme, with comparable activity to a previously published recombinant KLK4 produced from S2 insect cells.

The second aspect of this study was the activation of the PARs by KLK4 and the initiation of signal transduction. This study demonstrated that KLK4 can activate PAR-1 and PAR-2 to mobilise intracellular Ca2+, but failed to activate PAR-4. Further, KLK4 activated PAR-1 and PAR-2 over distinct concentration ranges, with KLK4 activation and mobilisation of Ca2+ demonstrating higher efficacy through PAR-2. Thus, the remainder of this study focussed on PAR-2. KLK4 was demonstrated to directly cleave a synthetic peptide that mimicked the PAR-2 N- terminal activation sequence. Further, KLK4 mediated Ca2+ mobilisation through PAR-2 was accompanied by the initiation of the extra-cellular regulated kinase (ERK) cascade. The specificity of intracellular signaling mediated through PAR-2 by KLK4 activation was demonstrated by siRNA mediated protein depletion, with a reduction in PAR-2 protein levels correlating to a reduction in KLK4 mediated Ca2+mobilisation and ERK phosphorylation.

The third aspect of this study examined cellular processing of KLK4 activated PAR- 2 in a prostate cancer cell line. PAR-2 was demonstrated to be expressed by five

iii prostate derived cell lines including the prostate cancer cell line PC-3. It was also demonstrated by flow cytometry and confocal microscopy analyses that activation of PC-3 cell surface PAR-2 by KLK4 leads to internalisation of this receptor in a time dependent manner.

Critically, in vivo relevance of the interaction between KLK4 and PAR-2 was established by the observation of the co-expression of receptor and agonist in primary prostate cancer and prostate cancer bone lesion samples by immunohistochemical analysis.

Based on the results of this study a number of exciting future studies have been proposed, including, delineating differences in KLK4 cellular signaling via PAR-1 and PAR-2 and the role of PAR-1 and PAR-2 activation by KLK4 in prostate cancer cells and bone cells in prostate cancer progression.

iv TABLE OF CONTENTS

Keywords………………………………………………………………………… i Abstract…………………………………………………………………………. ii Table of contents……………………………………………………………….. v List of figures………………………………………………………………….... x List of tables…………………………………………………………………….. xii List of abbreviations…………………………………………………………… xiii List of publications…………………………………………………………….. xv Statement of originality………………………………………….………...…... xvi Acknowledgements…………………………………………………………….. xvii

CHAPTER 1: INTRODUCTION……………………………………………... 1 1.1 Prostate cancer…………………………………………………………. 2 1.2 Proteases……………………………………………………………...... 3 1.2.1 Serine proteases…………………………………………………………. 3 1.3 Kallikrein-related peptidases………………………………………..... 7

1.3.1 Kallikrein-related peptidases in the prostate…………………………..... 8

1.3.2 Kallikrein-related peptidase 4………………………………………...... 11 1.4 Protease Activated Receptors………………………………………..... 12 1.4.1 Protease Activated Receptor-1………………………………………….. 13 1.4.2 Patho-physiological roles of Prostate Activated Receptor-1……………. 18 1.4.3 Protease Activated Receptor-2………………………………………...... 19

1.4.4 Patho-physiological roles of Prostate Activated Receptor-2……………. 20 1.4.5 Protease Activated Receptor-3………………………………………...... 21 1.4.6 Physiological roles of Prostate Activated Receptor-3………………...... 22 1.4.7 Protease Activated Receptor-4………………………………………….. 22 1.4.8 Patho-physiological roles of Prostate Activated Receptor-4………...... 23 1.5 Protease Activated Receptors in cancer…………………………...... 23 1.5.1 Protease Activated Receptor expression in prostate cancer…………...... 25 1.5.2 Consequences of Protease Activated Receptor activation in prostate cancer cell lines………………………………………………………..... 27

v 1.6 PAR cleavage by prostatic trypsin-like kallikrein-related peptidases……………………………………………………………..... 29 1.7 Study aims……………………………………………………………… 31

CHAPTER 2: MATERIALS AND METHODS...…………………………… 33 2.1 Materials……………………………………………………………...... 34 2.1.1 Reagents………………………………………………………………… 38 2.1.2 Buffers…………………………………………………………………... 35 2.1.3 Enzymes and kits………………………………………………………... 35

2.1.4 Oligonucleotides………………………………………………………… 36 2.1.5 Vectors………………………………………………………………….. 36 2.1.6 Bacterial growth media………………………………………………..... 36 2.1.7 Bacterial culture plates………………………………………………….. 38 2.2 Methods……………………………………………………………….... 39 2.2.1 Cell culture…………………………………………………………….... 39 2.2.2 Reverse transcription polymerase chain reaction……………………….. 39 2.2.2.1 RNA extraction…………………………………………………………. 39 2.2.2.2 Reverse transcription………………………………………………….... 40 2.2.2.3 Polymerase Chain Reaction…………………………………………….. 40 2.2.2.4 Agarose gel electrophoresis…………………………………………….. 40 2.2.3 DNA cloning……………………………………………………………. 41 2.2.3.1 Coding region generation and purification……………………………… 41 2.2.3.2 Ligation and transformation……………………………………………. 41 2.2.3.3 Screening transformed clones…………………………………………... 42 2.2.3.4 Plasmid DNA isolation and purification………………………………... 42 2.2.3.5 Restriction digestion…………………………………………………..... 42 2.2.3.6 DNA sequencing……………………………………………………...... 42 2.2.4 Protein analysis………………………………………………………..... 43 2.2.4.1 Whole cell protein extraction…………………………………………… 43 2.2.4.2 Protein quantification…………………………………………………… 43 2.2.4.3 Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis………… 43 2.2.4.4 Protein staining………………………………………………………..... 44 2.2.4.5 Western blotting analysis……………………………………………...... 44 2.2.5 Expression Constructs and transfections……………………………...... 45 vi 2.2.6 Expression and purification of recombinant pro-KLK4-V5-His……….. 45 2.2.7 Recombinant pro-KLK4-V5-His activation…………………………..... 46 2.2.8 Kinetic measurements of activated KLK4-V5-His……………………... 46 2.2.9 Ca2+ flux assays…………………………………………………………. 47 2.2.10 siRNA knockdown of PAR-2 expression……………………………..... 47 2.2.11 Cell surface biotinylation……………………………………………...... 48 2.2.12 Flow cytometry………………………………………………………...... 48 2.2.13 Confocal microscopy………………………………………………...... 49 2.2.14 Immunohistochemistry……………………………………………...... 50

CHAPTER 3: GENERATION AND CHARACTERISATION OF

RECOMBINANT HUMAN KALLIKREIN-RELATED PEPTIDASE 4………... 51 3.1 Introduction...... 52 3.2 Materials and Methods………………………………………………... 53 3.3 Results…………………………………………………………………... 54 3.3.1 Production of recombinant KLK4 from Sf9 insect cells………………... 54 3.3.2 N-glycosylation of KLK4…………………………………………...... 56 3.3.3 Activation of KLK4 by thermolysin…………………………………….. 58 3.3.4 Characterisation of thermolysin activated KLK4……………………….. 60 3.3.5 Defining the importance of the catalytic serine residue in KLK4 activity…………………………………………………………………... 63 3.3.6 Inhibition of KLK4 activity……………………………………………... 66 3.4 Discussion…………………………………………………………...... 69

CHAPTER 4: ANALYSIS OF PROTEASE ACTIVATED RECEPTOR

ACTIVATION BY KALLIKREIN-RELATED PEPTIDASE 4………………… 75 4.1 Introduction………………………………………………………...... 75 4.2 Materials and Methods……………………………………………….. 77 4.3 Results……………………………………………………………...... 78 4.3.1 Cytosolic calcium release in response to PAR activation by KLK4...... 78 4.3.1.1 Determination of PAR-1 activation by KLK4…………………………. 79 4.3.1.2 Determination of PAR-2 activation by KLK4………………………...... 79 4.3.1.3 Determination of PAR-4 activation by KLK4………………………...... 80

vii 4.3.1.4 Characterisation of KLK4 mediated calcium mobilisation through PAR-1 and PAR-2……………………………………………... 82

4.3.2 Cleavage of the PAR-2 N-terminus by KLK4………………………….. 84 4.3.3 ERK phosphorylation in response to PAR-2 activation by KLK4……... 86 4.3.4 Depletion of PAR-2 by siRNA and consequences for signal transduction……………………………………………………...... 90 4.3.5 Discussion……………………………………………………...... 95

CHAPTER 5: EXPRESSION AND LOCALISATION OF PROTEASE

ACTIVATED RECEPTOR-2 IN PROSTATE CANCER TISSUE AND CELL

LINES AND PROCESSING FOLLOWING KALLIKREIN-RELATED

PEPTIDASE 4 ACTIVATION…………………………………………………. 101 5.1 Introduction…………………………………………………………… 102 5.2 Materials and Methods………………………………………………. 103 5.3 Results………………………………………………………………..... 104 5.3.1 Expression of PAR-2 in normal, neoplastic and metastatic prostate tissues…………………………………………………...... 104 5.3.2 Transcription and expression of PAR-2 in normal and neoplastic prostate epithelial cell lines………………………………………...... 106 5.3.3 Localisation of endogenous PAR-2 in a prostate cancer cell lines……... 108 5.3.4 KLK4 activation of endogenous PAR-2 in a prostate cancer cell line..... 112 5.3.5 KLK4 mediated internalisation of endogenous PAR-2 by flow cytometry……………………………………………………………..... 112 5.3.6 KLK4 mediated internalisation of exogenous PAR-2 in a prostate cancer cell line………………………………………………………...... 114 5.4 Discussion……………………………………………………………… 117

CHAPTER 6: GENERAL DISCUSSION AND FUTURE DIRECTIONS….…. 123 6.1 Key observations……………………………………………………… 124 6.2 Implications of PAR-1 and PAR-2 activation for prostate cancer tumourigenesis……………………………………………………...... 124 6.2.2 Potential contributions of KLK4 activation of PARs to prostate cancer bone metastasis………………………...... 126 viii 6.3 Mechanistic considerations of KLK4 activation of PAR-1 and PAR-2…………………………………………………………...... 127 6.4 Potential in vivo regulation mechanisms…………………………..... 128 6.4.1 Activation of the KLK4 zymogen…………………………………...... 129 6.4.2 Regulation of KLK4 activity………………………………………...... 130 6.5 PARs as therapeutic targets in prostate cancer…………………...... 131 6.6 Future directions……………………………………………………… 133 6.7 Summary…………………………………………………………….... 135

Bibliography………………………………………………………………….. 137

ix LIST OF FIGURES

Figure 1.1 The general activation mechanism for S1 serine proteases………... 5 Figure 1.2 Serine protease catalytic mechanisms……………………………... 6 Figure 1.3 Serine protease mediated PAR activation and disarming…………. 14 Figure 1.4 PAR family structural features……………..……………………… 16 Figure 1.5 PAR-1 G protein coupling……………………………………...... 17 Figure 2.1 Vectors utilised in this PhD program of study……………………. 38 Figure 3.1 Expression and purification of recombinant pro-KLK4-V5-His from Sf9 insect cells……………………………………………….. 56 Figure 3.2 N-glycosylation of insect cell and human cell expressed pro-KLK4-V5-His………………………………………………… 57 Figure 3.3 Removal of pro-KLK4-V5-His pro-region by thermolysin……….. 59 Figure 3.4 Active site titration of KLK4-V5-His……………………………... 61 Figure 3.5 pH spectrum of KLK4-V5-His activity…………………………… 62 Figure 3.6 Velocities of KLK4-V5-His and trypsin hydrolysis of the F-V-R-pNA tri-peptide………………………………………….... 63 Figure 3.7 Comparison of insect cell expressed recombinant mutant pro-KLK4-V5-His and wild-type pro-KLK4-V5-His…………….. 65 Figure 3.8 Comparison of wild-type and mutant KLK4-V5-His velocities….. 66 Figure 3.9 Aprotinin complex formation with zymogen and mature form KLK4-V5-His…………………………………………………….. 67 Figure 3.10 Inhibition of KLK4-V5-His activity by aprotinin………………... 68 Figure 3.11 Crystal structure of KLK4………………………………………... 72 Figure 4.1 PAR-1 mediated calcium mobilisation………………………….... 80 Figure 4.2 PAR-2 mediated calcium mobilisation…………………………… 81 Figure 4.3 PAR-4 mediated calcium mobilisation…………………………… 82 Figure 4.4 Functional PAR-1,-2 or -4 remaining after KLK4 treatment…….. 83 Figure 4.5 KLK4 dose responses of PAR-1 and PAR-2 activation and effect on calcium mobilisation…………………………………………... 85 Figure 4.6 Characterisation of KLK4 cleavage of a peptide spanning the PAR-2 activation site…………………………………………….. 87

x Figure 4.7 Time dependent phosphorylation of ERK1/2 in response to PAR-2 activation……………………...... 88 Figure 4.8 KLK4 dose response of PAR-2 activation and effect on phosphorylation of ERK1/2………………………………………. 89 Figure 4.9 siRNA depletion of PAR-2 protein in PAR-2-LMF cells………... 91 Figure 4.10 Reduction of calcium mobilisation by PAR-2 siRNA mediated protein depletion………………………………………………….. 92 Figure 4.11 Reduction of ERK1/2 phosphorylation by PAR-2 siRNA mediated protein depletion……………………………………….. 94 Figure 5.1 Immunohistochemistry of PAR-2 and KLK4 expression in primary prostate cancers………………………………………….. 105 Figure 5.2 Immunohistochemical analyses of PAR-2 and KLK4 expression in prostate cancer bone metastasis………………………………... 107 Figure 5.3 The transcription of PAR-2 in prostate epithelial derived cell lines………………………………………………………………. 107 Figure 5.4 The expression of PAR-2 in prostate epithelial derived cell lines… 108 Figure 5.5 Surface biotinylation of endogenous PAR-2 in PC-3 cells………. 109 Figure 5.6 Confocal microscopy analysis of endogenous PAR-2 in PC-3 cells……………………………………………………………….. 110 Figure 5.7 Examination of PAR-2 lipid raft localisation…………………….. 111 Figure 5.8 KLK4 initiated changes in intracellular Ca2+ ion concentration in prostate cancer derived PC-3 cell lines…………………………... 113 Figure 5.9 Quantitation of endogenous PAR-2 internalisation in PC-3 cells... 114 Figure 5.10 KLK4 treatment of PC-3 cells transiently transfected with a PAR-2-GFP expression construct………………………………... 115 Figure 5.11 Quantitation of PAR-2 GFP internalisation in PC-3 cells………… 116

xi LIST OF TABLES

Table 1.1 Summary of studies examining proteolytic substrate specificity and prostate expression of kallikrein-related peptidases………….... 9 Table 1.2 PAR agonist peptides………………………………………………. 14 Table 1.3 Summary of studies examining protease activated receptor expression in normal and cancerous prostate……………………… 26 Table 1.4 Summary of studies examining PAR activation by tryptic KLKs…. 30 Table 2.1 Oligonucleotide primer sets used in PCR experiments…………….. 37 Table 2.2 siRNA oligonucleotide templates used in the synthesis of siRNAs by pSilencer3.1-H1 puro siRNA Construction Kit…………………. 37

xii LIST OF ABBREVIATIONS

μg microgram μL microliter μM micromole per litre AP agonist peptide BCA bicinchoninic acid assay bp base pairs BPH benign prostatic hyperplasia BSA bovine serum albumin cDNA complementary DNA DEPC diethylpryrocarbonate DMSO dimethylsulphoxide DNA deoxyribonucleic acid dNTP deoxynucleotide triphosphate DAPI 4, 6-diamidino-2-phenylindoledihydrochloride hydrate DTT dithiothreitol EDTA ethylene diamine tetra acetate EMSP-1 enamel matrix serine protease-1 ECM extra-cellular matrix ERK extra-cellular regulated kinase g gram GFP green fluorescent protein GPCR G protein coupled receptor h hour IPTG isopropyl-β-D-thiogalactoside kb kilobase kDa kilo Dalton KLK kallikrein-related peptidase LB Luria Bertani LMF lung murine fibroblasts mM millimole per litre M mole per litre

xiii mL millilitre mRNA messenger RNA MUGB 4-methylumbelliferyl pguanidinobenzoate NaCl sodium chloride NaOH sodium hydroxide ng nanogram nmol nanomole nM nanomole per litre OD optical density PAP prostatic acid phosphatase PAR protease activated receptor pNA paranitroaniline PBS phosphate-buffered saline PCR polymerase chain reaction PIN prostatic intraepithelial neoplasia PSA prostate specific antigen RNA ribonucleic acid rpm revolutions per minute RT room temperature RT-PCR reverse transcription polymerase chain reaction SDS sodium dodecyl sulfate SDS-PAGE sodium dodecyl sulfate-polyacrylamide gel electrophoresis SEM standard error of the mean SSC standard saline citrate TAE Tris-acetate, EDTA TBS Tris-buffered saline TTSP type II transmembrane serine proteases U units UV ultraviolet

xiv LIST OF PUBLICATIONS

PUBLICATIONS ARISING DIRECTLY FROM THIS PhD PROGRAM OF STUDY

AUTHORS (in order of authorship): Ramsay AJ, Dong Y, Hunt ML, Linn M, Samaratunga H, Clements JA and Hooper JD TITLE: Kallikrein-related peptidase (KLK) 4 initiates intracellular signalling via protease-activated receptors (PARs). KLK4 and PAR-2 are co-expressed during prostate cancer progression. JOURNAL: The Journal of Biological Chemistry DATE OF PUBLICATION: Accepted 29 February 2008 IF 5.808

AUTHORS (in order of authorship): Ramsay AJ, Reid JC, Adams MN, Samaratunga H, Dong Y, Clements JA and Hooper JD TITLE: Prostatic trypsin-like kallikrein-related peptidases (KLKs) and other prostate expressed tryptic proteases as regulators of signalling via protease activated receptors (PARs) JOURNAL: Biological Chemistry DATE OF PUBLICATION: Accepted 15 February 2008 IF 2.752

ADDITIONAL PUBLICATIONS DURING THIS PhD PROGRAM OF STUDY

AUTHORS (in order of authorship): He Y, Ramsay AJ, Hunt ML, Whitbread AK, Myers SA, Hooper JD TITLE: N-glycosylation analysis of the human Tweety family of putative chloride ion channels supports a penta-spanning membrane arrangement. Impact of N- glycosylation on cellular processing of Tweety homologue 2 (TTYH2). JOURNAL: The Biochemical Journal DATE OF PUBLICATION: Accepted 11 February 2008 IF 4.100

AUTHORS (in order of authorship): Ramsay AJ, Reid JC, Velasco G, Quigley JP and Hooper JD TITLE: The type II transmembrane serine protease Matriptase-2 – identification, structural features, enzymology, expression, pattern and potential roles. JOURNAL: Frontiers in Bioscience DATE OF PUBLICATION: 1 January 2008 IF 3.600

xv STATEMENT OF ORIGINALITY

The work in this thesis has not been previously submitted for a degree or a diploma at any other higher education institute. To the best of my knowledge and belief, the thesis contains no material previously published or written by another person except where due reference is made.

Signed:………………………………….. Andrew Ramsay, B.App. Sci. (Hons)

___/___/___

xvi ACKNOWLEDGEMENTS

This work was supported by a Queensland Cancer Fund PhD scholarship and a Clive and Vera Ramaciotti Foundation grant awarded to Dr. John Hooper.

I would firstly like to thank my supervisors, Dr. John Hooper and Prof. Judith Clements, whose guidance, advice, constructive criticisms and encouragement have been a driving force throughout the duration of my PhD. John in particular has invested countless hours in guiding me through the many aspects required for a successful career in science.

I have been fortunate enough to have been surrounded by many good people during my time at QUT, many of whom have become friends. To my good friends Brett Hollier, Stephen Myers and Shea Carter, thanks for all the good times we have shared. To Mitch Lawerence, Johnny Lai and Brett Willams, thanks for the laughs. I am also grateful to members of the Hooper group and hormone dependent cancer program who have provided invaluable insights and advice to me over the years.

To Michelle O’Connor, who gave me her patience, understanding, encouragement and support during my PhD, this significant achievement would not have been possible without you. Finally, I can never repay my parents and family for their constant love and support throughout my life, I hope this makes you all proud.

xvii

CHAPTER 1

INTRODUCTION

1.1 Prostate cancer

In men from western countries, prostate cancer is estimated to account for 33% of all newly diagnosed cancers for the year 2006 (Jemal et al., 2006). Of men diagnosed with prostate cancer, 9% are expected to die from the disease (Jemal et al., 2006). Risk factors for the development of prostate cancer include genetic, environmental, dietary, social and hormonal factors, with the incidence of prostate cancer strongly increasing with age (Haas and Sakr, 1997). Prostate cancer manifests itself in indolent or aggressive forms of the disease (Harnden et al., 2008). In its initial stages, when confined to the prostatic capsule, prostate carcinoma is essentially curable by surgical intervention and/or radiation therapy (Wilt and Thompson, 2006). Critically, if not detected early, or in more aggressive forms of the disease, prostate cancer can advance to stages characterised by local invasion of the seminal vesicles, followed by metastasis primarily to the bone, ultimately resulting in death (Li et al., 2008).

The majority of prostate cancer research has focussed on aspects of the clinical progression pathway that are pertinent issues for patient outcome. Central to these aspects has been the identification of prognostic markers that distinguish aggressive forms of prostate cancer from the majority of indolent cancers. In contrast, less attention has been focused on the mechanisms underlying prostate cancer initiation, and on defining the parameters of cancer progression in molecular terms. As such, the molecular mechanisms underpinning the development and progression of prostate cancer remain largely undefined.

Key components in both the development and progression of cancer are proteases. Tumour-associated proteolytic activities were first observed at the beginning of the 20th century when tumour cells were found to have enhanced plasma clot-degrading activities in tissue culture. The significance of this observation was the implication that proteolytic activities of tumour cells could be a molecular mechanism for cancer progression and metastasis. In 1946 Fisher proposed that tumour-associated proteases were potentially responsible for degradation of the extra-cellular matrix (ECM) and the subsequent invasion of surrounding tissues (Fisher, 1946). This hypothesis is now strongly supported, with the contribution of extracellular proteolysis to tumour invasion and metastasis through ECM degradation now having

2 been recognised for decades (Duffy et al., 2008). Importantly, it is currently acknowledged that proteases of all classes contribute to all stages of tumour progression, not just the later stages of disease as was originally proposed by Fisher (Egeblad and Werb, 2002; Borgono and Diamandis, 2004; Mohamed and Sloane, 2006).

1.2 Proteases

Proteolytic enzymes (proteases) are effectors of numerous biological events either as non-specific catalysts of protein degradation or highly selective mediators involved in tightly regulated physiological events (López-Otín and Bond, 2008). The current classes of proteases are recognised on the basis of their catalytic mechanisms and include serine, cysteine, aspartic, glutamic, asparagines, glycine, metallo- and threonine proteases (Neurath, 1984; López-Otín and Bond, 2008) . With the advent of whole genome sequencing this classification system has expanded by necessity to encompass the diversity of proteases now recognised (López-Otín and Matrisian, 2007). Barrett and Rawlings have devised a classification system that divides proteases into clans based on catalytic mechanism and families on the basis of common ancestry. Each clan is identified with two letters, the first representing the catalytic type of the families included in the clan (with the letter 'P' being used for a clan containing families of more than one of the catalytic types). At present, over 66000 protease protein sequences have been classified into 50 clans and 184 families (Rawlings et al., 2006). The largest of the protease classes are the serine proteases, with over 26000 members grouped into 13 clans and 40 families (Rawlings et al., 2006).

1.2.1 Serine proteases

Serine proteases are proteolytic enzymes that exploit serine for their catalytic activity and display ubiquitous organism expression, being found in viruses, bacteria and eukaryotes (Rawlings and Barrett, 1994). Over 20 families (denoted S1 - S66) of serine protease have been identified, these being grouped into clans on the basis of structural similarity and other functional evidence (Rawlings et al., 2006).

3 Serine proteases belonging to clan PA are the largest and most well described group of enzymes to date (Page and Di Cera, 2008). The cleavage site specificity of these enzymes is indicated by the residue six amino acids before the catalytic serine. In the active conformation of the protease, this residue is located at the bottom of a pocket in which the side-chain of the scissile bond of the substrate is inserted. An aspartate or, rarely, a glutamate at this site indicates that the serine protease will preferentially cleave after substrate arginine or lysine residues (trypsin-like or tryptic substrate specificity); a small hydrophobic residue in this position specifies preferential cleavage after large hydrophobic amino acids (chymotrypsin-like substrate specificity); and larger, usually non-polar residues at this site specify preferential proteolysis following small hydrophobic amino acids (elastase-like substrate specificity) (Perona and Craik, 1995; Perona and Craik, 1997).

The majority of serine proteases in clan PA belong to the S1 family, which encompasses two distinct sub-families, S1A and S1B. S1A and S1B are phylogenetically distinct groups of enzymes that share common structural architecture. The S1B proteases are ubiquitously expressed in all cellular life and are responsible for intracellular turnover, while S1A proteases mediate a variety of extracellular processes and display a limited distribution in plants, prokaryotes and the archea (Page and Di Cera, 2008).

Generally the catalytic activity of S1 serine proteases requires an initial conformational change to expose the S1 substrate binding pocket of the enzyme. This is catalysed by formation of a salt bridge between the isoleucine of the nascent N- terminus, exposed following pro-region removal from the zymogen, and the invariant aspartate located immediately before the catalytic serine (Page and Di Cera, 2008) (Figure 1.1). Further, removal of the pro-region from zymogen serine proteases leads to exposure of the pre-formed active site portion that is sterically blocked by pro-segment residues (Khan and James, 1998). Successful substrate pocket formation and active site exposure allows substrate binding and proteolytic cleavage.

4

Figure 1.1 The general activation mechanism for S1 serine proteases. The conformational change generally required for serine protease catalytic activity is initiated by cleavage to remove the pro-region from the zymogen enzyme, allowing the formation of a salt bridge (marked by dashed line) between α-amino group of the released isoleucine and the β-carboxylate group of the aspartate (represented by white filled circles) located immediately before the catalytic serine. The catalytic residues histidine, aspartate and serine are marked and represented by grey filled circles in the linear arrangement of the clan PA serine proteases.

Proteolytic cleavage proceeds through the carbonyl carbon of the substrate P1 amino acid (definition derived from (Schechter and Berger, 1967)) being attacked by the oxygen (nucleophilic attack) of the catalytic serine to form a tetrahedral intermediate, which undergoes water molecule-catalysed deacylation to release the enzyme and the amino terminal portion of the substrate (Page and Di Cera, 2008) (Figure 1.2).

Nucleophilicity of the catalytic serine is typically dependent on a catalytic triad consisting of aspartate, histidine and serine residues (Blow et al., 1969) (Figure 1.2). The geometric orientations of the catalytic residues are similar between families, despite different protein folds (Rawlings and Barrett, 1994). The linear arrangements of the catalytic residues commonly reflect clan relationships. For example the catalytic triad in the chymotrypsin clan (PA) is ordered histidine-aspartate-serine, but is ordered aspartate-histidine-serine in the subtilisin clan (SB) and serine-aspartate- histidine in the carboxypeptidase clan (SC) (Rawlings and Barrett, 1993; Rawlings and Barrett, 1994).

5

Figure 1.2 Serine protease catalytic mechanisms. An overview of the serine protease proteolytic mechanism mediated by the catalytic triad residues histidine57, aspartate102 and serine195 (numbering based on chymotrypsin). Starting with the left side and descending, the first stage of hydrolysis involves insertion of the recognised portion of the peptide substrate into the enzyme active site. This leads to the formation of a covalent acyl-enzyme tetrahedral intermediate, in which the carboxyl of the substrate is esterified to the hydroxyl group of serine195. The nucleophilicity of the serine –OH is markedly enhanced by histidine57, which accepts a proton from serine as serine attacks the carbonyl carbon atom of the substrate. The negatively charged aspartate102 functions to stabilise the resulting positively charged histidine through electrostatic interactions. The negative charge on the tetrahedral transition state is also stabilised by hydrogen bonding to two main-chain NH groups in the oxyanion hole. The second stage (right hand side of panel), deacylation, is in essence a reverse of the acylation, with H2O substituting for the amine component. (Taken from http://www.mansfield.ohiostate.edu/~sabedon/biochem511.htm [2/2/08])

The S1A serine proteases, which encompass as many as 140 (Rawlings et al., 2006) of the total 569 human degradome complement (López-Otín and Matrisian, 2007), modulate a variety of cellular processes by selective cleavage of specific substrates to influence cell behaviour (Turk, 2006). Well known physiological examples

6 include the proteases of the blood coagulation (eg. thrombin; (Davie et al., 1991)), digestive (eg. trypsin; (Yamashina, 1956)) and wound healing (eg. plasmin; (Castellino and Ploplis, 2005)) cascades. It is also clear that dys-regulated activity of S1A serine protease facilitates progression of various diseases including cancer (Dano et al., 2005; Turk, 2006; Szabo and Bugge, 2007). Landmark studies in the 1970s lead to the identification of plasminogen activators as proteolytic enzymes associated with oncogenic transformation (Ossowski et al., 1973; Unkeless et al., 1973; Quigley et al., 1974). The current knowledge of the plasminogen activation (PA) system illustrates the importance of S1A serine proteases for prostate cancer progression. The serine protease plasmin, formed by the cleavage of plasminogen by urokinase plasminogen activator (uPA), plays a critical role in the proteolytic processes associated with progression of malignant tumours (Sidenius and Blasi, 2003). Plasmin can either directly degrade the basement membrane and ECM or activate other zymogen proteases such as procollagenase, promoting neoplastic spread (Sidenius and Blasi, 2003). A family of S1A serine proteases with documented roles in prostate cancer development and progression are the KLKs.

1.3 Kallikrein-related peptidases

The largest contiguous collection of S1A serine protease encoding genes is located within the KLK locus at chromosome 19q13.3-13.4 (Gan et al., 2000; Harvey et al., 2000; Yousef et al., 2000). The 15 encoded KLK serine proteases are involved in the post translational processing of polypeptide precursors to either their active or inactive forms (Clements et al., 2004).

The KLK gene family was originally thought to encode only three proteins; tissue kallikrein (KLK1), glandular kallikrein (KLK2) and prostate specific antigen (PSA/KLK3) (Clements et al., 2001). However, it is now recognised that the KLK family consists of 15 members (Gan et al., 2000; Harvey et al., 2000; Yousef et al., 2000). The human KLK1-15 genes range in size from 4-10 Kb and display a conserved genomic organisation, with each member containing five coding exons. All KLKs are synthesised as preproenzymes, which range in size from 244 to 305 amino acids (Clements et al., 2004). At the N-terminus of each KLK is a

7 hydrophobic region that ranges from 15 to 73 amino acids in length. This feature instructs secretion with the signal peptide being subsequently cleaved to give an inactive precursor. The proenzyme contains an amino-terminal activation sequence, which is cleaved to yield the mature active enzyme (Borgono and Diamandis, 2004; Clements et al., 2004). Additionally, the region of the exons responsible for encoding the catalytic triad (histidine (H), aspartate (D) and serine (S)) is conserved across all fifteen genes in humans (Clements et al., 2001).

Generally, either an aspartate183 (trypsin-like) or serine183 (chymotrypsin-like) residue is located at the bottom of the first substrate-binding pocket and determines substrate cleavage specificity, with a preference for basic (arginine/lysine) or hydrophobic amino acids respectively. Substrate specificity analysis of the KLKs revealed KLK 1, 2, 4, 5, 6, 8 and 10-15 are predicted or have been shown to possess trypsin-like activity (Clements et al., 2001) (Table 1.1). The KLKs are expressed by numerous tissues within the body, including the skin, breast, ovary and prostate. However, there are significant differences in the levels of expression of each KLK in these tissues (Clements et al., 2004).

1.3.1 Kallikrein-related peptidases in the prostate

A common site of expression for the KLKs is the prostate, with this tissue a predominant site of mRNA expression for KLK2, KLK3, KLK4, KLK11, KLK14 and KLK15 (Clements et al., 2004; Shaw and Diamandis, 2007). In addition, mRNAs of KLK6, KLK9, KLK12 and KLK13 are highly expressed in this tissue, while low or very low prostatic mRNA expression has been reported for each of the remaining KLKs (Clements et al., 2004; Shaw and Diamandis, 2007). Importantly, as summarised in Table 1.1, all family members, except KLK8, have been detected at the protein level in normal or cancerous prostate.

8 Table 1.1 Summary of studies examining proteolytic substrate specificity and prostate expression of kallikrein-related peptidases 1

Protein Expression in Normal and Cancerous Prostate

Specificity 2 3 KLK P1 preference Localization Cancer

1TArg, Met, Phe > Lys SF; N ND

(Borgono et al., 2007b; Bourgeois et al., 1997; Chagas et al., 1995; (Fink et al., 1985; Geiger and Clausnitzer, 1981; Shaw and Diamandis, 2007) Chen et al., 2000; Sueiras-Diaz et al., 1994)

2TArg SF; N; B; P; C up

(Darson et al., 1999; Darson et al., 1997; Deperthes et al., 1995; Herrala et al., (Bourgeois et al., 1997; Cloutier et al., 2002; Lovgren et al., 1999a; 2001; Lovgren et al., 1999b; Magklara et al., 2000; Shaw and Diamandis, 2007; Mikolajczyk et al., 1998) Tremblay et al., 1997; Veveris-Lowe et al., 2005; Young et al., 1996)

3 C Leu, Tyr, Phe, Gln, His, Asn, Leu, Ser SF; N; B; P; C down

(Akiyama et al., 1987; Christensson et al., 1990; Coombs et al., (Darson et al., 1999; Darson et al., 1997; Herrala et al., 2001; Ishida et al., 2003; 1998; Denmeade et al., 1997; Malm et al., 2000; Takayama et al., Magklara et al., 2000; Shaw and Diamandis, 2007; Simone et al., 2000; Tremblay 1997) et al., 1997; Veveris-Lowe et al., 2005)

4TArg > Lys SF; N; B; P; C up

(Borgono et al., 2007a; Debela et al., 2006a; Debela et al., 2006b; (Day et al., 2002; Dong et al., 2004; Klokk et al., 2007; Obiezu et al., 2005; Matsumura et al., 2005; Obiezu et al., 2006; Takayama et al., Obiezu et al., 2002; Shaw and Diamandis, 2007; Veveris-Lowe et al., 2005) 2001b)

5TArg > Lys, Phe, Tyr, Gly, Asp SF; N ND

(Borgono et al., 2007a; Borgono et al., 2007b; Brattsand et al., 2005; Debela et al., 2006b; Michael et al., 2005; Oikonomopoulou (Shaw and Diamandis, 2007) et al., 2006)

6TArg > Lys, Leu, Phe, Tyr, Leu, Gly, Ser, Thr, Pro SF; N; B; C down

(Bernett et al., 2002; Borgono et al., 2007b; Debela et al., 2006b; Little et al., 1997; Magklara et al., 2003; Oikonomopoulou et al., (Diamandis et al., 2000; Petraki et al., 2003a; Petraki et al., 2001) 2006; Okui et al., 2001)

7 C Try > Phe, Leu SF; N ND

(Egelrud, 1993; Skytt et al., 1995) (Kishi et al., 2004; Shaw and Diamandis, 2007)

8TArg, Lys Not detected ND

(Kishi et al., 2006; Rajapakse et al., 2005; Stefansson et al., 2008)

9 C ND SF; N ND

(Shaw and Diamandis, 2007)

10 T Arg > Lys SF; N; B; C down

(Luo et al., 2001b; Petraki et al., 2003a; Petraki et al., 2002; Shaw and (Debela et al., 2006b) Diamandis, 2007)

4 11 T Arg SF; N; B up

(Luo et al., 2006; Mitsui et al., 2000) (Diamandis et al., 2002; Nakamura et al., 2003b; Shaw and Diamandis, 2007)

12 T Lys > Arg SF; B; C ND

(Memari et al., 2007b) (Memari et al., 2007a; Shaw and Diamandis, 2007)

13 T Arg > Lys SF; N; B; C down

(Borgono et al., 2007a; Kapadia et al., 2004; Sotiropoulou et al., (Kapadia et al., 2003; Petraki et al., 2003a; Petraki et al., 2003b; Shaw and 2003) Diamandis, 2007)

4 14 T Arg > Lys, Tyr, Phe,Tyr, Gly, Leu SF; N up

(Borgono et al., 2007a; Borgono et al., 2007b; Borgono et al., 2007c; Brattsand et al., 2005; Felber et al., 2005; Oikonomopoulou (Borgono et al., 2007c; Shaw and Diamandis, 2007) et al., 2006; Stefansson et al., 2008)

15 T Arg > Lys SF; N ND

(Takayama et al., 2001a) (Shaw and Diamandis, 2007; Shaw et al., 2007)

Notes: 1. T, tryptic; C, chymotryptic. 2. SF, seminal fluid; N, normal prostate tissue; B, benign prostate disease; P, prostatic intraepithelial neoplasia; C, cancer. 3. Up or down-regulated in prostate cancer. ND, not done; up/down, increase/decrease in detectable levels compared with benign or normal. 4. Determined from serum.

9 At present, KLK2 and KLK3 are the most well described of the prostate expressed KLKs with respect to roles in normal physiology and cancer. PSA is secreted by the prostate into the seminal fluid where it is involved in semen liquification, which is required for sperm motility (Kumar et al., 1997). However, PSA is known more for its diagnostic importance in prostate cancer (Filella et al., 1997). In the prostate, PSA is thought to promote cellular proliferation and growth by activating growth factors. For example, PSA is reported to cleave the insulin growth factor binding proteins 3 and 4 (IGFBP-3 and IGFBP-4) and thus increase levels of free IGF (Okabe et al., 1999; Rehault et al., 2001).

The proliferative effects of PSA are also mediated by activation of pro-transforming growth factor beta-2 (proTGF-β2) to TGF-β2 (Killian et al., 1993; Dallas et al., 2005). PSA is also thought to be involved in prostate cancer metastasis to bone by cleaving parathyroid hormone related protein (PTHrP), a protein that is important in regulating osteoclast destruction of bone (Russell et al., 1998). Furthermore, studies investigating secreted PSA from the prostate cancer cell line LNCaP, suggest that PSA plays a role in ECM degradation, a key step in the metastatic cascade, by degrading fibronectin and laminin (Webber et al., 1995). More recently, clinical evaluation of KLK2 showed promising results as a complementary marker to PSA to improve sensitivity and specificity of PSA detection (Haese et al., 2003). Similar to PSA, KLK2 functions to degrade both seminogelins I and II in vivo, however, KLK3 cleaves these substrates at a higher efficiency and at different sites compared with KLK2 (Deperthes et al., 1996). In common with KLK2, KLK4 displays largely prostate restricted expression. Significantly, KLK4 protein levels are elevated in malignant prostate compared with normal tissue (Dong et al., 2005; Klokk et al., 2007), while prostate cancer patient sera contains antibodies that bind recombinant KLK4 (Day et al., 2002). In vivo roles for KLK4 are yet to be elucidated, although limited studies have implicated KLK4 in seminal clot liquidification (Takayama et al., 2001b) and ECM remodeling (Obiezu et al., 2006). Due to limited knowledge of physiological roles and elevated expression in prostate cancer, KLK4 is the primary focus of this study.

10 1.3.2 Kallikrein-related peptidase 4

KLK4 was originally cloned using a subtractive hybridisation approach from human prostate tissue (Nelson et al., 1999) and by positional candidate gene approaches (Stephenson et al., 1999; Yousef et al., 1999). The KLK4 gene encodes a preproprotein of 254 amino acid residues that is converted by a signal peptidase to a 228 residue zymogen. The zymogen is converted to the 224 amino acid mature enzyme by an unknown activating enzyme. KLK4 contains the highly conserved canonical serine protease catalytic triad, histidine57, aspartate102 and serine195, in the active site that acts as a charge relay system for proteolytic hydrolysis (Rawlings and Barrett, 1994). Due to the presence of an aspartate residue at the bottom of the first substrate pocket (aspartate189) (Nelson et al., 1999), KLK4 has a trypsin-like specificity, with preference for cleavage following basic amino acids (Takayama et al., 2001a; Obiezu et al., 2006; Debela et al., 2006b). KLK4 is predominately a secreted protein and shares 72% amino acid homology to the previously characterised porcine orthologue enamel matrix serine protease 1 (EMSP-1) that was identified in the early stages of tooth development (Simmer et al., 1998 615). EMSP- 1 is expressed during the early maturation stages of mammalian tooth formation, functioning in enamel layer mineralisation by degrading the ECM surrounding enamel crystallites (Hu et al., 2000). Consistent with the role of EMSP-1 in ECM turnover, KLK4 has been reported to degrade the ECM components type I and type IV collagen in vitro (Obiezu et al., 2006).

In addition to ECM components, several macromolecular substrates of potential patho-physiological relevance have been identified for KLK4 from in vitro studies including another KLK family member, pro-PSA (Takayama et al., 2001a) and pro- uPA and the uPA receptor (Beaufort et al., 2006), as well as fibrinogen (Obiezu et al., 2006). These in vitro KLK4 substrates have reported roles in prostate cancer development and metastasis (Russell et al., 1998; Okabe et al., 1999; Rehault et al., 2001; Sidenius and Blasi, 2003), although conclusive data linking KLK4 to alterations in prostate cancer cellular behaviour are lacking. Using cell based assays, work done in our laboratory has shown that stable over-expression of KLK4 in prostate cancer PC-3 cells resulted in an increased ability of these cells to migrate, accompanied by a transition from an epithelial morphology to a fibroblastic shape

11 and, consistently, a significant decrease in E-cadherin protein levels and an increase in vimentin expression (Veveris-Lowe et al., 2005). In addition, using an inducible expression system, Klokk and colleagues demonstrated that over-expression of an amino-terminally truncated isoform of KLK4, results in significantly increased colony formation, migration and proliferation of the PC-3 and DU145 prostate cancer cell lines (Klokk et al., 2007). Most recently using co-culture systems, it has been shown that KLK4 is a potential mediator of cellular interactions between prostate cancer cells and osteoblasts (bone forming cells) in bone metastases (Gao et al., 2007).

Expanding the known substrates of KLK4 is critical to delineating in vivo physiological roles which may be altered to facilitate carcinogenesis. Over the last 15 years it has been increasingly recognised that many of the cellular effects elicited by trypsin-like serine proteases are mediated by cleavage of cell surface proteins known as PARs. Further, recent studies by Oikonomopoulou and colleagues have identified members of the KLK family with trypsin-like specificity as activators of the PARs (Oikonomopoulou et al., 2006).

1.4 Protease Activated Receptors

GPCRs represent one of the largest and most diverse groups of proteins encoded by the genome (Bjarnadottir et al., 2006). GPCRs serve to regulate intracellular signaling pathways by primarily interacting with the heterotrimeric G proteins, α, β and γ (Oldham and Hamm, 2008). Receptor activation, caused by agonist binding, causes conformational changes within the receptor, which in turn catalyses nucleotide exchange between heterotrimeric G proteins. Guanine nucleotide exchange is a key event in many cellular processes such as metabolic enzyme function, modulation of gene expression and cytoskeletal reorganisation. Nearly 800 different human genes encode receptors for various ligands; including hormones, neurotransmitters, sensory stimuli (Bjarnadottir et al., 2006) and more recently, trypsin-like serine proteases.

Members of the GPCR sub-family comprising PAR-1 to PAR-4, in contrast to other GPCRs which are activated by docking of soluble ligands, are irreversibly activated

12 by the action of proteases (Coughlin, 2000; Macfarlane et al., 2001). Indeed, PAR activation is almost exclusively mediated by trypsin-fold serine proteases with substrate specificity for cleavage following arginine or lysine residues. As shown in Figure 1.3 cleavage occurs within the activation amino-terminal exodomain of the receptor generating a new amino-terminal that serves as a tethered ligand which binds intramolecularly, causing allosteric changes within the PAR, followed by receptor coupling to heterotrimeric G-proteins and signal transduction (Coughlin, 2000). It is well established that for PAR-1, -2 and -4, short synthetic peptides designed on the tethered ligand sequence can serve as selective receptor agonists (agonist peptides; AP) for the respective PAR (Table 1.2), and as such, have served as critical reagents in PAR activation studies (Hollenberg and Compton, 2002). Importantly, it is also possible for proteases to cleave downstream of the activation site leading to disarming of the receptor (Oikonomopoulou et al., 2006) (Figure 1.3, disarming).

Summarised in the following sections are the currently known biochemical properties and patho-physiological roles of the PAR family members.

1.4.1 Protease Activated Receptor-1

To date PAR-1 is the most widely studied and well characterised of the PAR receptors. PAR-1 was discovered in 1991 by Coughlin and colleagues using a dilution cloning approach (Vu et al., 1991). Sequencing of the functional clone in the study revealed a 3.5 kb insert, containing an open reading frame which encoded a 425 amino acid protein. The protein was deduced by hydropathy analysis, to be a member of the seven transmembrane domain receptor superfamily. PAR-1 is activated proteolytically by the serine protease, thrombin and exhibits an extended N-terminus containing both a putative thrombin cleavage site (LDPR41/S42) and a hirudin (thrombin inhibitor)-like domain (D52KYEPF), a binding site for the anion exosite of thrombin, essential for high affinity binding and the potent effects of thrombin (Vu et al., 1991) (Figure 1.4).

13

Figure 1.3 Serine protease mediated PAR activation and disarming. This schematic shows the structure of the protease activated receptor family members. Note the seven transmembrane regions and extended N-terminus containing the tethered ligand sequence. N-terminal cleavage by a serine protease can result in either receptor activation or disarming. Receptor activation involves cleavage at an arginine or lysine to expose the tethered ligand which in turn binds intramolecularly to induce an allosteric change in the receptor. Receptor disarming involves cleavage distal to the activation site causing removal of the tethered ligand and inhibition of receptor activation.

Table 1.2 PAR agonist peptides.

Receptor activation Protease-Activated Receptor Human tethered ligand sequence Receptor agonist peptide PAR-1 SFFLRN SFFLRN activates both PAR-1 and PAR-2 TFLLRN selectively activates PAR-1 - identified in structure function studies

PAR-2 SLIGKV SLIGKV selectively activates PAR-2

PAR-3 TFRGAP No tethered ligand sequence from any species activates PAR-3 PAR-4 GYPGQV GYPGQV selectively activates PAR-4 - low potency agonist AYPGKF selectively activates PAR-4 - identified in structure function studies as high potency agonist

14 The importance of the N-terminal region in the binding and subsequent activation of thrombin has been explored in several studies. A mutant PAR-1 receptor with the LDPR/S site replaced by an enterokinase site was responsive to enterokinase, demonstrating that there are no additional mechanisms for receptor activation (Vu et al., 1991). In support of this finding, a mutant PAR-1 receptor lacking the N-terminal thrombin cleavage site was both inactive and unresponsive to thrombin stimulation (Chen et al., 1994). Although not essential for receptor activation, the hirudin-like domain is of particular importance to the potency of thrombin’s activation of PAR-1. A study utilising γ-thrombin (which lacks the anion exosite) to stimulate PAR-1 activation, demonstrated that γ-thrombin was a 100 fold less potent than thrombin in receptor activation (Bouton et al., 1995). It is important to note, the unique thrombin binding domain of PAR-1 does not restrict activation by other proteases. PAR-1 can be activated by Factor Xa (Riewald et al., 2001), plasmin (Pendurthi et al., 2002) or activated protein C (APC) (Riewald et al., 2002). The ability of PAR-1 to be activated intramolecularly by its tethered agonist was demonstrated in studies showing receptor activation by the AP mimicking the new N-terminal sequence (SFLLRN) created after proteolytic cleavage (Vu et al., 1991).

In the case of PAR-1 its AP, SFLLRNPNDKYEFP, was able to activate both mutant (LDPR/S site replaced by an enterokinase) and wild-type receptors (Vu et al., 1991). The ability of the PAR-1 AP to activate a mutant receptor, lacking the N-terminal extracellular domain, suggests a site or number of sites through which receptor activation can proceed. Experiments utilising antibodies generated against different segments of PAR-1 demonstrated that both the N-terminal exodomain and the second extracellular loop (ECL) (Figure 1.4) determine binding of the SFLLRN AP (Bahou et al., 1994). Furthermore, studies have demonstrated that the second ECL determines the specificity of this interaction (Lerner et al., 1996).

In common with other helipthical receptors, PAR-1 couples to G-proteins following receptor activation. The intracellular signaling event that first demonstrated G protein/PAR-1 coupling was stimulation of phospholipase C-catalysed hydrolysis of polyphosphoinositides by the Gq/11 class of G proteins (Hung et al., 1992). Coupling between PAR-1 and Gq/11 results in the formation of inositol 1, 4, 5-trisphosphate 2+ (InsP3), mobilisation of intracellular Ca and the production of diacylglycerol,

15 which in turn activates protein kinase C (Clapham, 2007) (Figure 1.5). More recent studies have determined that PAR-1 also induces cellular signals via the G proteins

Gi2 (Hung et al., 1992), G12 and G13 (Offermanns et al., 1994) (Figure 1.5).

Figure 1.4 PAR family structural features. The regions highlighted (boxed) include; the N-terminal showing the protease cleavage sites and the hirudin like binding domain, the conserved extracellular loop 2 which contains the region for receptor/tethered agonist interactions, and the C-terminal tail that is involved in signal transduction and desensitisation. (Taken from (Macfarlane et al., 2001)).

Coupling between Gi and PAR-1 is implicated in the activation of the ERK cascade (van Corven et al., 1993). ERK1/2 are also named the “switch kinases”, due to their key role in integrating various extracellular signals into a co-ordinated cellular response (Payne et al., 1991), which is largely the initiation of mitogenesis (Pages et al., 1993). Currently the paradigm for activation of ERK1/2, of which PAR-1 incorporates many components, is that which has been established for tyrosine kinase linked receptors. Initiation of this pathway involves phosphotyrosine residues within the intracellular domain of an activated receptor interacting with the adapter protein SHC that subsequently recruits GRB-mSos for increased guanine-nucleotide exchange by p21ras, leading to plasma membrane translocation of Raf-1 isoforms for

16 the activation of RAS and the downstream activation of ERK kinase, the direct activator of ERK1/2 (Figure 1.5) (Malarkey et al., 1995).

Collectively the initiation of intracellular signaling through PAR-1/G protein coupling leads to alterations in a number of cellular functions, including; growth, morphology, motility and soluble factor release.

Figure 1.5 PAR-1 G protein coupling. PAR-1 G protein coupling to G12/13, Gi/2 and Gq/11 proteins. PAR-1 coupling to Gq/11 and Gi/2 initiates the well characterised intracellular signaling pathways leading to cytosolic Ca2+ release (grey filled) and activation of the ERK1/2 (black filled) respectively. Dashed lines represent formation of breakdown products. PLC, phospholipase C; PIP2, phosphatidylinositol 4,5-biphosphate; IP3, inositol 1,4,5-trisphosphate; DAG, diacylglycerol; PKC, protein kinase C; MEK, extracellular signal-regulated kinase kinase; ERK, extracellular signal-regulated kinase.

17 1.4.2 Patho-physiological roles of Prostate Activated Receptor-1

The cloning of thrombin receptors PAR-1, PAR-3 and PAR-4 (the latter 2 receptors are comprehensively covered in subsequent sections) ended the search for the cell receptor(s) responsible for mediating the various cellular responses thrombin initiates (Vu et al., 1991; Ishihara et al., 1997). Consequently, it is now widely accepted that many of the early studies of thrombin mediated cellular effects are due to activation of the thrombin receptors.

There are presently a number of studies indicating that thrombin plays a critical role in vascular injury responses associated with thrombosis, arthrosclerosis and balloon angioplasty (Stouffer et al., 1996). Platelet adhesion, aggregation and soluble factor release are critical early events in initiating the vascular response to injury (Eidt et al., 1989). Thrombin mediates platelet shape change and their release of the inflammatory mediators, serotonin (5-HT), adenosine triphosphate, thromboxane A2 and other granule contents (Harmon and Jamieson, 1986). Thrombin also stimulates the plasma membrane localisation of integrin aIIb/β3, catalysing the binding of fibrinogen and von Willebrand factor, resulting in platelet aggregation (McGregor et al., 1989). Additionally, thrombin stimulation leads to the translocation of P-selectin and CD40 ligand to the plasma membrane causing the binding of platelet to endothelial cells (Henn et al., 1998). Activation of endothelial cells is another important response to thrombin activation of the thrombin receptors. Thrombin released from platelets stimulates the release of von Willebrand factor, cell surface redistribution of P-selectin, and increased expression of tissue factor and adhesion molecules, ICAM-1, VCAM-1, and E-selectin on endothelial cells (Hattori et al., 1989). The net effect of these processes is to provide rapid adherence of neutrophils, monocytes and lymphocytes in sites of vessel damage.

Platelet derived thrombin also acts as a potent mitogen for cells of mesenchymal origin. Thrombin has been shown to stimulate increased DNA synthesis in endothelial, fibroblast and vascular smooth muscles cell (Chen and Buchanan, 1975). Thrombin also has an indirect effect on cell growth by stimulating the synthesis and release of the mitogenic growth factors, platelet derived growth factor and endothelin-1 (Kohno et al., 1992).

18

The activation of the thrombin receptors also plays a role in the wound healing process. The breakdown of the basement membrane along with endothelial cell migration and proliferation are important events in catalysing angiogenesis, a process which is essential for normal tissue healing (Singer and Clark, 1999). Consistently, thrombin mediates the increased synthesis and activation of basement membrane degrading enzymes, such as MMP-1, MMP-2 and MMP-3 and progelatinase A (Nguyen et al., 1999). Further, thrombin stimulation of angiogenesis has been confirmed in vitro with endothelial tube formation in matrigel (Haralabopoulos et al., 1997), as well as in vivo with induction of angiogenesis in the chick chorioallantoic membrane system (Tsopanoglou et al., 1993).

Since the discovery of PAR-1, three additional receptors, PAR-2, PAR-3 and PAR-4, have been cloned and a number of their patho-physiological roles delineated.

1.4.3 Protease Activated Receptor-2

PAR-2 was first identified from a mouse genomic library using hybridisation of probes corresponding to regions of the bovine substance K receptor (Nystedt et al., 1994). The complete mouse PAR-2 cDNA contains an 1197 nucleotide open reading frame (Nystedt et al., 1995). The gene encoding human PAR-2 was isolated from a human genomic library, hybridising with a probe based on the 3’ exon of the known mouse PAR-2 gene (Nystedt et al., 1995). Significant differences between human PAR-1 and PAR-2 exist in the N-terminus. The 397 amino acid, PAR-2 is shorter than PAR-1 due to its lack of a hirudin-like binding domain (Figure 1.4). PAR-2 is unique amongst the PARs, as it is the only receptor described to date which is not activated by thrombin. The activation region of PAR-2 displays no distinct protease specificity, which has lead to the concept of a 'generic PAR' that serves as the signal transducer for a variety of proteases like the TTSPs (Hooper et al., 2001) such as mast cell tryptase (Molino et al., 1997b), membrane-type serine protease-1 (MTSP- 1)/matriptase (Takeuchi et al., 2000) and TMPRSS2 (Wilson et al., 2005), as well as other proteases such as sperm protease acrosin (Smith et al., 2000), bacterial proteases (Lourbakos et al., 1998) and coagulation factors (F)VIIa (Camerer et al., 2000) and FXa (Riewald and Ruf, 2001).

19

Serine protease cleavage of PAR-2 at Arg36/Ser37 generates the N-terminal sequence, SLIGKV (Figure 1.4). In Chinese hamster ovary cells transfected with human PAR- 2, both the AP of PAR-1 and PAR-2, SFLLRNP and SLIGKV respectively, were able to mediate PAR-2 activation (Nystedt et al., 1995), demonstrating cross reactivity between the agonists. This was resolved in experiments using a combinatorial peptide design approach and monitoring PAR-1 and PAR-2 activation in human embryonic kidney 293 (HEK293) cells. This approach successfully identified an AP with the sequence TFLLR which selectively activated PAR-1 with high potency in HEK 293 cells without activating or desensitising PAR2 (Hollenberg et al., 1997) (Table 1.2). In addition, this study demonstrated that the PAR-2 AP SLIGKV activates PAR-2 with high potency without cross-activating PAR-1 (Hollenberg et al., 1997) (Table 1.2).

2+ Similar to PAR-1, PAR-2 couples to Gq/11 to initiate Ca mobilisation. Furthermore, pertussis toxin sensitive Ca2+ mobilisation has been demonstrated in PAR-2 transfected Xenopus oocytes in response to trypsin (Nystedt et al., 1994), indicating the potential involvement of Gi dependent transduction mechanism in a manner similar to PAR-1 (Schultheiss et al., 1997). Although the mechanisms remain uncharacterised, activation of PAR-2 by trypsin has been shown to initiate ERK activation (Belham et al., 1996). At present, no direct studies have examined coupling to other G proteins such as G12 and G13.

1.4.4 Patho-physiological roles of Prostate Activated Receptor-2

The expression of PAR-2 in a number of highly vascularised human and animal tissues suggests a role in the regulation of vascular tone (Nystedt et al., 1994) (Nystedt et al., 1995; Bohm et al., 1996; D'Andrea et al., 1998). Trypsin and PAR-2 AP treatment induced endothelium relaxation in rat aorta (al-Ani et al., 1995), porcine coronary (Hwa et al., 1996) and basilar arteries (Sobey and Cocks, 1998). PAR-2 mediated endothelium relaxation is due to the activation of nitric oxide synthase by Ca2+ mobilisation induced by PAR-2 (Sobey et al., 1999). Strong expression of PAR-2 has been detected in cells of the gastrointestinal tract, including enterocytes in both basolateral and apical membranes (D'Andrea et al., 1998). PAR-2

20 activation by its AP at the apical site stimulated intracellular Ca2+ and the secretion of prostaglandin (PGE)2 and PGF1α (Kong et al., 1997). Expression of PAR-2 has been noted in other digestive organs, most significantly the pancreas, where its expression and activation has been linked to release of amylase (Bohm et al., 1996). Additionally, a role in exocrine secretion in salivary, parotid and sublingual glands has proposed a common function for PAR-2 in secretion for tissues of the intestinal tract (Kawabata et al., 2000).

Expression of PAR-2 at moderate to high levels has been noted within layers of epidermal keratinocytes (Steinhoff et al., 1999). Interestingly, expression was found to differ between layers of the epidermis, with expression of PAR-2 higher in the more differentiated granular layer compared to that of the proliferative basal layer. It remains yet to be elucidated what the physiological consequences are for these differences, although it has been suggested that PAR-2 may function to control keratinocyte terminal differentiation. Consistently, activation of PAR-2 in epidermal keratinocytes inhibits cellular proliferation and differentiation (Derian et al., 1997).

1.4.5 Protease Activated Receptor-3

PAR-3 was first described in mouse platelets, which were deficient in PAR-1, but were fully responsive to thrombin (Ishihara et al., 1997). As a part of this study, a candidate receptor was cloned from rat platelet mRNA, using a series of degenerate primers corresponding to conserved regions of PAR-1 and PAR-2. Subsequently, human PAR-3 was identified using primers based on the identified rat PAR-3 clone (Ishihara et al., 1997). The 374 amino acid human PAR-3 receptor shares 27% homology with human PAR-1 and 28% with human PAR-2. The consensus sequence for both a serine protease cleavage site (Lys38/Thr39) and a hirudin-like domain were identified in the N-terminal of the receptor (Figure 1.4). Confirming the identified consensus serine protease site, the study by Ishihara et al showed PAR-3 receptor activation in receptor transfected COS-7 cell line, was dependent upon the presence of the Lys38/Thr39 site (Ishihara et al., 1997).

Interestingly, AP that mimic the putative tethered ligand sequence have been demonstrated to be ineffective in activating PAR-3 (Ishihara et al., 1997) (Table

21 1.2), but rather activate PAR-1 and PAR-2 in PAR-expressing cells. It has been suggested by Ishihara and colleagues that amino-terminal cleavage of PAR-3 is sufficient to induce an active structural state, independent of amino-terminal binding to extracellular loop 2 (Ishihara et al., 1997). Due to lack of a specific activator, molecular and physiological effects due to PAR-3 activation remain poorly defined.

1.4.6 Patho-physiological roles of Prostate Activated Receptor-3

Interestingly, in mouse platelets PAR-3 expression was found to be necessary for full activation of platelet aggregation and secretion by thrombin, as PAR-3 deficient mouse platelets expressing only PAR-4 were delayed and less sensitive to thrombin stimulation (Kahn et al., 1998). As murine PAR-3 does not signal directly, it may behave as a co-receptor that increases the efficiency of PAR-4 cleavage by thrombin (Nakanishi-Matsui et al., 2000). As PAR-3 cannot be specifically activated, it will be omitted from examination in this study.

1.4.7 Protease Activated Receptor-4

PAR-4 was identified by two laboratories employing bioinformatic approaches (Kahn et al., 1998; Xu et al., 1998). The PAR-4 cDNA was isolated from a lymphoma daudi cell line library, using a 600bp probe identified from EST sequence searches (Xu et al., 1998). Human PAR-4 is 385 amino acids in length and like other members of the family, PAR-4 possesses a putative serine protease cleavage site (Arg47/Gly48) in the N-terminal sequence (Figure 1.4). Interestingly like PAR-2, PAR-4 lacks a hirudin-like binding domain (Figure 1.4). Due to the lack of a hirudin-like binding domain, PAR-4 is relatively insensitive to thrombin activation, with an EC50 for the serine protease at approximately 50 fold higher than the amount required for PAR-1 activation (Xu et al., 1998). Along with thrombin, trypsin is an activator of PAR-4, with potency equal to that of thrombin.

The tethered ligand peptide for PAR4, GYPGKF, lacks potency and is of limited utility (Faruqi et al., 2000). In a combinatorial peptide design approach study, similar to that used to identify the specific PAR-1 AP TFLLR, Faruqi and colleagues demonstrated that the AP with the sequence AYPGKF was approximately 10-fold

22 more potent than GYPGKF and displayed selective activation for PAR-4 (Faruqi et al., 2000) (Table 1.2). Whereas PAR-1 couples to Gq/11 and Gi/2, PAR4 couples to

Gq/11 to induce calcium mobilisation, but not Gi/2 (Faruqi et al., 2000).

1.4.8 Patho-physiological roles of Prostate Activated Receptor-4

Of the thrombin-activated receptors, PAR-1 and PAR-4 account for thrombin signaling in human platelets (Trejo et al., 1996). Both receptors are expressed by platelets, and activation of either PAR-1 or PAR-4 with their respective AP is sufficient to trigger platelet ATP secretion and aggregation (Faruqi et al., 2000). PAR-1 blocking antibodies inhibit platelet secretion and aggregation at low but not high concentrations of thrombin, while PAR-4 blocking antibodies alone have little effect on platelet activation. Importantly, inhibition of both PAR-1 and PAR-4 with blocking antibodies markedly attenuates platelet activation even at high concentrations of thrombin (Kahn et al., 1999). Thus, it has been suggested that PAR-4 functions as a low affinity thrombin receptor, helping to sustain platelet aggregation in response to thrombin during physiological events of elevated thrombin release when PAR-1 is rapidly inactivated (Kahn et al., 1999).

PAR-4 has also been reported to play a role in nociceptive (physical pain) threshold. Intraplantar injection of a PAR-4 AP significantly increased nociceptive threshold in response to thermal and mechanical noxious stimuli. Further, co-injection of the PAR-4 AP with the excipient carrageenan, significantly reduced the induced inflammatory hyperalgesia (increased sensitivity to pain) and allodynia (painful response to a characterisitcally non-painful stimulus), but had no effect on inflammatory parameters such as oedema and granulocyte infiltration (Asfaha et al., 2007).

1.5 Protease Activated Receptors in cancer

A number of studies in recent years have focussed on the expression and possible role of PARs in several cancers. The expression of the PARs has been noted in cells originating from pancreatic tumours (Rudroff et al., 1998), colon adenocarinoma (Wojtukiewicz et al., 1995), lung adenocarcinoma (Jin et al., 2003), breast carcinoma

23 (Even-Ram et al., 1998), melanoma cell lines (Even-Ram et al., 2001) and gastric carcinoma (Caruso et al., 2006). Importantly, an increasing number of reports have demonstrated that the expression of PARs increases as a result of cancer progression. For example, Even-Ram and colleagues demonstrated that the level of PAR-1 mRNA expression in human breast carcinoma biopsy specimens is proportional to the degree of invasiveness of the tumour (Even-Ram et al., 1998). Similarly, the metastatic potential of human melanoma cells correlates with over-expression of PAR-1 (Nierodzik et al., 1998). Further, immunohistochemical analysis demonstrated that PAR-2 expression is up-regulated during ovarian cancer progression (Jahan et al., 2007). Interestingly, the expression of PAR-1 and PAR-2 is also increased in stromal fibroblasts of malignant tissues, but remains unchanged in fibroblasts of benign and normal origin (D'Andrea et al., 2001). Significantly, in vitro studies using proliferating, human dermal fibroblasts, and scrape-wounded human dermal fibroblasts demonstrated the presence of PAR-1 and PAR-2 not detected in quiescent fibroblasts (D'Andrea et al., 2001).

In addition to increased expression, studies have demonstrated PAR-1 involvement in cellular processes necessary for cancer progression. For example, introduction of antisense PAR-1 cDNA into the aggressively metastatic breast carcinoma cell line MDA-435 caused the reduction of in vitro invasiveness to a level comparable to non- metastatic cell lines (Even-Ram et al., 1998). In addition, targeted over-expression of human PAR-1 in mouse mammary glands leads to precocious hyperplasia, characterised by a dense network of ductal side branching and accelerated proliferation (Yin et al., 2006). Further, Yin and colleagues demonstrated that mouse mammary gland over expression of PAR-1 activates the Wnt and β-catenin pathway, both critical components for tumour progression in numerous cancers (Yin et al., 2006). Stromal derived MMP-1, which plays a central role in remodeling the tumour- stromal microenvironment, was also recently determined to activate PAR-1 to cause breast cancer cell migration and invasion (Boire et al., 2005).

Increasing in vitro and animal model data have emerged implicating PAR-1 involvement in human melanoma metastasis. A study by Nierodzik et al found that pulmonary metastasis of B16F10 murine melanoma cells, which express only PAR- 1, increased in vivo experimental pulmonary murine metastasis model, following

24 exposure to the PAR-1 AP (Nierodzik et al., 1998). Furthermore, B16F10 cells transfected with PAR-1 displayed enhanced adhesion to the extracellular protein fibronectin following treatment with thrombin (Nierodzik et al., 1998). Interestingly, thrombin mediates increased in vitro invasion of melanoma cells through the synergistic activation of PAR-1 and PAR-2 through direct and indirect activation mechanisms respectively (Shi et al., 2004). Similarly, co-operative activation of PAR-1 and PAR-4 by thrombin, by direct cleavage mechanisms, leads to increased hepatoma carcinoma cell migration (Kaufmann et al., 2007).

Critical to this PhD program of study, a number of recent reports examined the expression of the PARs in prostate cancer and delineated consequences of their activation.

1.5.1 Protease Activated Receptor expression in prostate cancer

The mRNA and protein expression patterns of PARs in prostate, analysed by a number of approaches, are summarised in Table 1.3. Tissue microarray analysis of a large number of patient samples indicated elevated PAR-1 expression in benign prostatic hyperplasia compared to normal prostate stroma (Tantivejkul et al., 2005). Increased PAR-1 expression was also observed in high grade prostatic intraepithelial neoplasial (PIN) and prostate adenocarcinoma in comparison to normal prostate stroma, BPH and atrophic prostate tissues (Tantivejkul et al., 2005). Kaushal and colleagues have also recently reported correlation of increased PAR-1 mRNA and protein levels with prostate cancer stage (Kaushal et al., 2006). These workers, employing quantitative PCR analysis of early versus late stage prostate cancer tissue, showed an increase in PAR-1 mRNA in the late stage cancer samples. Increasing PAR-1 mRNA with prostate cancer stage was similarly reflected in PAR-1 protein expression, as Western blot analysis of PAR-1 protein levels showed a 2.75 fold increase in late versus early stage disease. This observation was consistent with their immunohistochemical analyses that showed low PAR-1 expression in 56% (15 of 27) of BPH samples, low to medium expression levels in 44% (14 of 32) and strong staining in 22% (7 of 32) of early stage prostate cancer, while strong staining was also observed in all 22 advanced prostate cancer tissues. Interestingly, approximately

25 50% of the strong staining seen in early and advanced tumours was in areas of blood vessels and a weak correlation was found between levels of PAR-1 and the pro- angiogenic factor vascular endothelial growth factor (VEGF)-2 (Kaushal et al., 2006).

Table 1.3 Summary of studies examining protease activated receptor expression in normal and cancerous prostate.

PAR Expression

PAR Prostate Tissue Type1 Level Localisation2 Reference

1 mRNA Matched N versus PCa (n=6) Increased in cancer (5 of 6) n/a Black et al., 2007 E (n=9) versus Ad (n=7) PCa Increased in advanced 2.39 fold n/a Kaushal et al., 2006 PCa pre- and post- Decreased in cancer after n/a Salah et al., 2007 androgen ablation (n=9) ablation (semi-quantitative) protein E (n=9) versus Ad (n=6) PCa Increased in advanced 2.75 fold n/a (Western blot analysis) Kaushal et al., 2006 BPH (n=27) Absent/low (12/15 of 27) VE E PCa (n=32) Absent/low-medium (12/14 of 32) VE and epithelial origin Ad PCa (n=22) Strong (22 of 22) VE and epithelial origin Normal prostate stroma (n=128) Stain intensity 1.1 epithelium Tantivejkul et al., 2005 BPH (n=303) Stain intensity 1.4 epithelial origin PIN (n=82) Stain intensity 2.7 epithelial origin PCa (n=82) Stain intensity 2.5 epithelial origin Matched N versus C (n=40) Increased in cancer (45%) epithelial origin Black et al., 2007 2 mRNA Matched N versus PCa (n=6) Increased in cancer (4 of 6) n/a Black et al., 2007 protein Matched N versus C (n=40) Increased in cancer (42%) epithelial origin Black et al., 2007 3 protein Matched N versus C (n=40) Absent n/a Black et al., 2007 4 mRNA Matched N versus PCa (n=6) Increased in cancer (3 of 6) n/a Black et al., 2007 protein Matched N versus C (n=40) Increased in cancer (68%) epithelial origin Black et al., 2007 Notes: 1. N, normal; E, early; Ad, advanced; BPH, benign prostatic hyperplasia; PIN, prostatic intraepithelial neoplasia; PCa, prostate cancer. 2. n/a, not applicable; VE, vascular endothelium.

Similar to PAR-1, other members of this receptor family are also elevated in prostate cancer. Black and co-workers demonstrated that six matched normal and cancerous patient samples showed increased mRNA levels of PAR-1 (5 of 6), PAR-2 (4 of 6) and PAR-4 (3 of 6), while PAR-3 levels were consistently low (Black et al., 2007). In addition, immunohistochemical analysis of 40 patient tissues showed little PAR-1 (7.5%) and PAR-2 (13%) in normal prostate glands, with PAR-4 staining apparent in approximately 55% of samples and PAR-3 expression completely absent.

PAR expression in prostate cancer samples was elevated in comparison to normal prostate glands for PAR-1 (45%), PAR-2 (42%) and PAR-4 (68%), with elevated

26 expression most notable in cancerous epithelial cells. Additionally, higher Gleason grade cancers (G7-10) had increased expression of PAR-1 in periglandular stromal cells compared to lower grade tumours (G5-9), signifying a shift from epithelial to periglandular expression in prostate cancer progression (Black et al., 2007). It is apparent that PARs are expressed in both normal and cancerous prostate. The presence of these receptors and the known roles of PARs as sentinels of tryptic serine protease activity, indicates the potential that these cell surface receptors will be important transducers of signals initiated by serine proteases expressed during prostate cancer initiation and progression.

1.5.2 Consequences of Protease Activated Receptor activation in prostate cancer cell lines

Activation of PARs in prostate cancer cell lines initiates a variety of cellular processes that are associated with cancer progression including proliferation, secretion of tumour promoting factors, changes in morphology and increased migration (Arora et al., 2007). As indicated below most of these studies have employed APs to induce signaling via PARs. In addition, several studies, recognising the role of thrombin as the endogenous activator of PAR-1 and PAR-4 in the vasculature (Vu et al., 1991; Xu et al., 1998), the correlation between hyperactivation of the coagulation system and tumour progression (Rickles et al., 1992) and the access of thrombin to cancer cells during cancer progression (Trikha and Nakada, 2002), have also employed this protease in studies of the consequences of PAR activation in prostate derived cells. Thrombin as well as trypsin stimulated LNCaP cell migration towards fetal calf serum in a concentration dependent manner. The observed protease induced migration was mimicked by PAR-1 and PAR-2 AP, but not by PAR-4 AP, signifying the importance of PAR-1 and PAR-2 in prostate cancer cell migration (Greenberg et al., 2003). Additionally, a low concentration of thrombin stimulated PC-3 and DU145 cell migration towards fibronectin (Shi et al., 2004). Interestingly, higher concentrations of thrombin resulted in unchanged cell migration of PC-3 cells on fibronectin and significantly decreased cell migration on plastic, collagen I and IV and laminin (Loberg et al., 2007) while synergistic treatment with PAR-1 and PAR-2 APs enhanced migration similar to thrombin in these cell lines. Interestingly, PAR-1

27 AP alone stimulated no migration, while PAR-2 AP resulted in only marginal migration. Thrombin in this study was implicated as an indirect activator of PAR-2, with de-sensitisation experiments pre-treating M24met cells with PAR-2 agonist abolishing the effect of thrombin induced migration. Furthermore, experiments employing a PAR-2 cleavage blocking antibody demonstrated that thrombin mediated effects were cleavage independent (Shi et al., 2004). One explanation is that PAR-1 and PAR-2 are forming heterodimers at the cell surface with thrombin mediated changes via PAR-1 requiring the interaction of these receptors (Shi et al., 2004).

The study by Greenberg and co-workers also indicated the ability of thrombin and trypsin to induce changes in the morphology of LNCaP cells. Proteolytic treatment induced cytoskeletal reorganisation after 8 hours, and, after 48 hours, the projection of filapodia containing actin filaments and the development of punctate focal adhesions connected with stress fibres consisting of thick actin bundles (Greenberg et al., 2003). Consistently, PAR-1 and PAR-2 APs were able to initiate activation of the cytoskeleton reorganisation mediated through GTPases RhoA, Rac1 and Cdc42 (Hall, 2005) in a manner similar to thrombin and trypsin in LNCaP cells (Greenberg et al., 2003). Loberg and colleages have also shown that thrombin treatment of PC-3 cells causes retraction of the actin cytoskeleton and numerous microspike extensions from the cell body (Loberg et al., 2007). This treatment was accompanied by rapid activation of RhoA and Cdc42 with no change in the level of Rac-1 activation (Loberg et al., 2007). Thrombin has also been reported to promote adhesion of prostate cancer cells to ECM proteins. For example, Liu and colleagues have shown that thrombin increases DU145 adhesion to fibronectin and laminin via a mechanism involving PAR-1 (Liu et al., 2004). In contrast to DU145 cells, treatment of PC-3 cells with a similar concentration of thrombin and PAR-1 AP decreased cell adhesion to collagen I and IV and laminin and resulted in no change in adhesion to fibronectin (Loberg et al., 2007).

In addition to cell migration and adhesion, examples of the co-opting of other normal cellular processes essential for cancer progression, such as tissue remodelling, cell growth, angiogenesis and mechanisms regulating apoptosis, have recently been shown to be mediated via PARs. For example, Wilson and colleagues

28 have demonstrated that activation of PAR-2, but not PAR-1, in LNCaP cells induced increased activity of the tissue remodelling enzyme MMP-2. In addition, these workers demonstrated that activation of both PAR-1 and PAR-2 induced increased MMP-2 activity in PC-3 and DU145 cells, while PAR-2 activation stimulated the activity of another matrix remodelling protease, MMP-9, in PC-3 cells (Wilson et al., 2004).

The importance of PAR-1 for growth of prostate cancer derived cells in mice has also recently been reported. Yin and co-workers demonstrated that rat Dunning prostate carcinoma cells expressing human PAR-1 displayed a 3.7 fold increase in tumour mass above controls when grown subcutaneously in rats (Yin et al., 2003). In addition, low concentrations of thrombin, have been reported to modulate proliferation of DU145, LNCaP and PNT1A cells (Liu et al., 2003). In another study, high concentrations of thrombin, acting through PAR-1, impaired the proliferation of DU145 cells by a mechanism proposed to involve the induction of apoptosis (Zain et al., 2000). In contrast, pre-treatment of DU145 and PC-3 cells with thrombin or PAR1 AP provided a protective effect against docetaxel-induced apoptosis by mediating increased expression of the anti-apoptotic protein Bcl-xL (Tantivejkul et al., 2005). Also of interest, thrombin treatment of DU145 cells has been shown to induce, via PAR-1 activation, elevated secretion of two cytokines that are implicated in tumour cell growth, interleukin (IL)-8 and VEGF (Liu et al., 2006). This study was consistent with a previous report showing that thrombin stimulation mediated increased VEGF mRNA levels in PC-3 cells (Huang et al., 2001).

It is apparent that PARs have the capacity to transduce signals in response to multiple tumour-generated proteases. Prostate cancer tumours are replete with tryptic serine proteases with the potential to activate tumour cell expressed PARs. One such group of prostate cancer expressed serine proteases are the tryptic members of the KLK family.

1.6 PAR cleavage by prostatic trypsin-like kallikrein-related peptidases

Studies examining the ability of tryptic KLKs to regulate signaling via PARs are summarised in Table 1.4. In a significant recent report Oikonomopoulou and

29 colleagues examined the ability of three family members (KLK5, 6 and 14) to regulate signaling via PAR-1, PAR-2 and PAR-4 (Oikonomopoulou et al., 2006). Mass spectral analysis of peptides spanning the activation site of each receptor indicated that a major cleavage site for each KLK is at the known receptor activation site. In addition, both KLK5 and KLK14 also cleaved up-stream of the PAR-2 activation site. Interestingly, only KLK14 was able to cleave at a site downstream of the activation site of the peptide of one of the receptors, PAR-1 (which would induce disarming of the receptor in intact cells; Figure 1.3).

Table 1.4 Summary of studies examining PAR activation by tryptic KLKs

Serine Protease Structure1 PAR Cleavage2 Reference

KLK1 secreted Unable to cleave PAR-2 activation peptide sequence Molino et al., 1997a

Unable to activate PAR-1 or PAR-2 in intact cells Molino et al., 1997b

Activates PAR-4 in rodent paw oedema model Houle et al., 2005

KLK5 secreted Cleaves at activation site of PAR-1, -2 and -4 peptides Oikonomopoulou et al., 2006

Cleaves cell surface hPAR-2 and rPAR-2 Oikonomopoulou et al., 2006; Stefansson et al., 2008

Induces Ca2+ mobilisation via hPAR-2 and rPAR-2 Oikonomopoulou et al., 2006; Stefansson et al., 2008

KLK6 secreted Cleaves at activation site of PAR-1, -2 and -4 peptides Oikonomopoulou et al., 2006

Cleaves cell surface rPAR-2

Induces Ca2+ mobilisation via rPAR-2

KLK7 secreted Unable to induce Ca2+ mobilisation via hPAR-2 Stefansson et al., 2008

KLK8 secreted Unable to induce Ca2+ mobilisation via hPAR-2 Stefansson et al., 2008

KLK14 secreted Cleaves at activation site of PAR-1, -2 and -4 peptides Oikonomopoulou et al., 2006

Cleaves downstream of activation site of PAR-1 peptide Oikonomopoulou et al., 2006

Cleaves cell surface hPAR-2 and rPAR-2 Oikonomopoulou et al., 2006; Stefansson et al., 2008

Induces Ca2+ mobilisation via h/rPAR-2 and h/rPAR-4 Oikonomopoulou et al., 2006; Stefansson et al., 2008

Disarms hPAR-1 in intact cells Oikonomopoulou et al., 2006

Notes:

1. TTSP, type II transmembrane serine protease 2. h, human; r, rat; m, mouse.

Consistent data were obtained from microscopy analyses of cells stably expressing rat PAR-2. Interestingly, these workers also showed that KLK5 is less efficient than KLK6 and KLK14 at cleaving at the PAR-2 activation site. Furthermore, experiments analysing Ca2+ mobilisation yielded important insights on the ability of KLKs to induce differential signaling in intact cells via PARs. These workers have proposed that PAR-2 is activated by low concentrations of KLK6, whereas higher enzyme concentration results in receptor inactivation. Equally as interesting is the proposal that, in cells expressing both PAR-1 and PAR-2, the former receptor is disarmed by KLK14 whereas the latter is activated by this enzyme. Adding another

30 layer of complexity, KLK14 was also shown to signal via PAR-4, whereas KLK5 and KLK6 were unable to induce signaling through this receptor at concentrations that initiated signaling via PAR-2 (Oikonomopoulou et al., 2006). The observations of Oikonomopoulou and co-workers indicate that individual KLKs can function as both activator and disarmer of individual PARs as well as have the ability to cleave multiple PARs. These findings highlight the potential complexity of regulation of PAR signaling; particularly in situations of dys-regulated trypsin-like serine protease activity and expression of multiple PARs such as occur in prostate cancer.

The data of Oikonomopoulou and colleagues have been partially confirmed by others. Stefansson and co-workers confirmed the ability of KLK5 and KLK14 to activate PAR-2 by examining calcium mobilisation in cells stably expressing the human receptor (Stefansson et al., 2008). This group also examined the ability of the related enzymes KLK7 (chymotryptic) and KLK8 (tryptic) to signal via PAR-2. Not surprisingly, KLK7 could not signal via PAR-2. Interestingly however, KLK8 was also unable to signal through this receptor (Stefansson et al., 2008). In contrast, whereas Oikonomopoulou and colleagues have reported that KLK6 cleaves peptides spanning the activation sites of PAR-1, PAR-2 and PAR-4, Angelo et al demonstrated that while KLK6 is able to efficiently cleave a PAR-2 activation site spanning peptide, this enzyme is unable to cleave PAR-1 or PAR-4 activation site spanning peptides or a PAR-3 peptide (Angelo et al., 2006).

Early studies by Molino and co-workers indicated that KLK1 is unable to cleave PAR-1 or PAR-2 in endothelial cells (Molino et al., 1997a) or to cleave a peptide spanning the PAR-2 activation site (Molino et al., 1997b). However, more recently Houle and colleagues used a rodent paw oedema model to demonstrate that this protease plays a role in inflammation by direct or indirect activation of PAR-4 (Houle et al., 2005).

1.7 Study aims

In summary, a number of members of the KLK family, including KLK4, are associated with prostate cancer. It is also clear that activation of PARs initiates

31 cellular changes (such as proliferation, migration and growth factor release) which facilitate cancer progression.

Accordingly, the hypothesis of this PhD program of study is that KLK4 will activate members of the PAR family and this activation will be functionally important in prostate cancer.

Therefore the specific aims of this project were:

1. To generate recombinant KLK4, develop a protocol to activate the zymogen and characterise the kinetics of the activated enzyme.

2. To examine the ability of KLK4 to activate PAR-1, PAR-2 and PAR-4 and the consequences of their activation in terms of Ca2+ mobilisation and initiation of the ERK cascade in a mouse PAR-1 knockout cell line reconstituted with human PAR-1, PAR-2 or PAR-4.

3. To examine expression and localisation of PARs in prostate derived cell lines and prostate cancer tissue and to examine the cellular processing of KLK4 activated PARs.

32

CHAPTER 2

MATERIALS AND METHODS

2.1 Materials

2.1.1 Reagents

General reagents were purchased from the following vendors: PAR-2 activating peptide (AP; SLIGKV-NH2), PAR-1 AP (TFLLR-NH2), PAR-4 AP (AYPGKF-NH2) as the carboxyl amide, a peptide spanning the PAR-2 serine protease activation site (ortho-aminobenzoic acid (Abz)- serine(S)-lysine(K)-glycine(G)-arginine(R)-serine( S)-↓-leucine(L)-isoleucine(I)-glycine(G)-lysine(K)(N-[2,4dintrophenyl]ethyleneedia mine (Dnp))-Asp-OH; “↓” indicates the cleaved peptide bond), and a tri-peptide substrate for kinetic studies benzoyl(Bz)-phenylalanine(F)-valine(V)-arginine(R)- paranitroaniline(pNA) were from Auspep (Parkville, Australia); 4′,6-diamidino-2- phenylindole hydrochloride (DAPI), FURA-2 acetoxymethyl ester and TRIzol reagent from Invitrogen (Mount Waverley, Australia); trypsin from Worthington Biochemical (Lakewood, NJ); thermolysin from Calbiochem (San Diego, CA); poly- L-lysine, bovine serum albumin (BSA), 3,3'-diaminobenzidine (DAB), imadazole, sodium orthovanadate, NaF, protease inhibitor cocktail (4-(2- aminoethyl)benzenesulfonyl fluoride (AEBSF), E-64, bestatin, leupeptin, aprotinin, and sodium EDTA), ampicillin, kanamycin, 4-methylumbelliferone, 4- methylumbelliferyl p-guanidinobenzoate (MUGB), bovine α-thrombin, and the thermolysin inhibitor phosphoramidon from Sigma (Sydney, Australia); Ni-NTA superflow resin from Qiagen (Doncaster, Australia); Odyssey blocking buffer from LiCOR (Millennium Science, Surrey Hills, Australia); and EZ-link NHSSS-Biotin and ImmunoPure immobilized streptavidin from Pierce (Quantum Scientific, Murarrie, Australia).

Cell culture reagents were purchased from the following vendors: Dulbecco’s modified Eagle’s medium, SF9002 serum free media, fetal calf serum, penicillin, streptomycin, hygromycin B, versene, lipofectamine and cellfectin from Invitrogen; and blasticidin and puromycin from InvivoGen (San Diego, CA)

Antibodies were purchased from the following vendors: phospho-specific monoclonal antibody to ERK1/2 and rabbit anti-ERK1/2 antibody, Cell Signalling (Beverly, MA); monoclonal anti-PAR-1 (ATAP2), anti-PAR-2 (SAM11) and anti-

34 GFP antibodies, Santa Cruz Biotechnology (Santa Cruz, CA); and rabbit anti- GAPDH antibody, Abcam (Sapphire Bioscience Pty Ltd, Redfern, Australia).

2.1.2 Buffers

Where pH values are stated, the baseline pH was quantitated by an Orion 91-02 general purpose combination pH electrode (GENEQ Inc, Montreal, Canada), before the addition of either 1M HCl or 2M NaOH to obtain the desired pH.

Phosphate-buffered saline (PBS): 137 mM NaCl, 10 mM sodium phosphate buffer pH 7.4, 2.7 mM KCl. TAE: 40 mM Tris-acetate, 1 mM EDTA (pH 8.0). Agarose gel loading dye: 0.09% (w/v) bromophenol blue, 0.09% (w/v) xylene cyanol FF, 60% (v/v) glycerol, 60 mM EDTA, 1M Tris (pH 8.0) Tris-buffered Saline (TBS): 100 mM Tris, 137 mM NaCl (pH 7.6). Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) loading dye: 100 mM Tris pH 6.8, 4% (w/v) SDS, 0.2% (w/v) bromophenol blue, 20% (v/v) glycerol, 200 mM β-mercaptoethanol. 5X SDS-PAGE running buffer: 125 mM Tris, 960 mM glycine, 0.5% (w/v) SDS (pH

8.3) diluted to 1X in ddH2O before use. Protein transfer buffer: 20 mM Tris, 192 mM glycine, 20% (v/v) methanol (pH 7.4). Coomassie blue gel staining buffer: 10% (v/v) acetic acid, 40% (v/v) methanol, 0.1%

(w/v) Coomassie Brilliant Blue R (Sigma) in ddH2O, filtered before use.

De-stain buffer: 10% (v/v) acetic acid, 45% (v/v) methanol in ddH2O. Protein total extraction buffer: Triton X-100 (1% v/v), 150 mM NaCl, 50 mM Tris- HCl (pH 7.4) Nickel (Ni) column wash buffer: 50 mM Tris, 300 mM NaCl (pH 8.0)

Extracellular medium: 121 mM NaCl, 5.4 mM KCl, 0.8 mM MgCl2·6H2O, 1.8 mM

CaCl2, 5.5 mM glucose, 25 mM HEPES (pH 7.4) containing 0.2% (w/v) BSA Bacterial lysis buffer: 0.2 M NaOH, 1% (w/v) SDS

2.1.3 Enzymes and kits

T4 DNA ligase (Promega, Sydney, Australia); SuperScript III RNase H-cDNA

35 synthesis kit (Invitrogen); Platnium Taq DNA polymerase (Invitrogen); PfuUltra DNA polymerases (Stratagene, La Jolla, CA); Restriction enzymes (Fermentas, Hanover, MD); QIAprep Spin Miniprep kit (Qiagen); High Pure PCR Cleanup kit (Roche); RNeasy Mini kit for RNA extraction (Qiagen); Deoxyribonuclease 1 (DNAse I) (Roche); ABI BigDye Terminator Cycle Sequencing kit (Applied Biosystems, Foster City, CA); EnVision+ peroxidise polymer detection system (Dako, Botany, Australia); Silver Stain Plus Kit (Bio-Rad laboratories, Gladesville, Australia); Bicinchoninic acid (BCA) assay kit (Pierce, Murarrie, Australia)

2.1.4 Oligonucleotides

Oligonucleotides used in polymerase chain reaction (PCR), DNA sequencing and in vitro transcription of siRNA duplexes were synthesised by Sigma. Oligonucleotides and experimental conditions utilised are listed in Table 2.1 (PCR and DNA sequencing) and Table 2.2 (siRNA).

2.1.5 Vectors

The following vectors were utilised in experiments performed in this PhD program of study, pGEM-T easy vector (Promega) (Figure 2.1A) for the sub-cloning of amplified PCR products, pEGFP-N1 vector (Clontech, Mountain View, CA) (Figure 2.1B) to express full-length PAR-2 with the addition of green fluorescent protein (GFP) at the carboxyl terminal, pIB-V5-His vector (Invitrogen) to express the recombinant protein pro-KLK4-V5-His in Spodoptera frugiperda Sf9 cells (Figure 2.1C) and pSilencer 3.1-H1 puro (Ambion, Austin, TX) for in vitro transcription of siRNA duplexes (Figure 2.1D).

2.1.6 Bacterial growth media

Bacterial culture was conducted in Luria-Bertani (LB) broth (1% (w/v) Bacto- Tryptone, 0.5% (w/v) Bacto-yeast extract and 0.5% (w/v) NaCl, pH 7.0). The post- transformation growth of bacteria performed for up to 1 h in SOC medium (2% (w/v) Bacto-Tryptone, 0.5% (w/v) yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM

MgCl2, 10 mM MgSO4 and 20 mM glucose) before plating onto culture plates.

36

Table 2.1 Oligonucleotide primer sets used in PCR experiments.

Gene Name Sequence Genbank Amplicon Annealing Ascension # Position Temperature PAR-21 F 5’-AGAAGCCTTATTGGTAAGGTT-3’ NM_005242 260-841 59°C R 5’-AACATCATGACAGGTGGTGAT-3’ PAR-22 F 5'-CTCGAGCTTCCAGGAGGATG NM_005242 144-1348 60°C CGG -3' (Xho1 site underlined) R 5'-GAATTCGATAGGAGGTCTTA ACAGTGGTTGAAC -3' (EcoR1 site underlined) Human β-actin F 5’CAAGATCATTGCTCCTCCTG-3’ NM_001101.2 1057-1367 54°C R 5’-TGTCACCTTCACCGTTCCA-3’ Mouse β-actin F 5’-CGTGGGCCGCCCTAGGCACCA-3’ NM_007393.2 181-423 63°C R 5’-TTGGCCTTAGGGTTCAGGGGGG-3’ Vector name Sequence Vector Vector Annealing Company position Temperature pGEMT-easy F 5'-TAATACGACTCACTATAGGG-3' Promega 2999-3 52°C R 5'-ATTTAGGTGACACTATAGAA-3' 139-158 pEGFP-N1 F 5'-CGCAAATGGGCGGTAGGCGTG-3' Clontech 518-539 60°C R 5'-CTCGACCAGGATGGGCACCAC-3' 745-724 pSilencer 3.1-H1 F 5'-GTTTTCCCAGTCACGAC-3' Ambion 359-375 60°C puro R 5'-GAGTTAGCTCACTCATTAGGC-3' 682-662 pIB-V5-His 5´-CGCAACGATCTGGTAAACAC-3´ Invitrogen 511-530 59°C 5´-GACAATACAAACTAAGATTTAGTCAG-3´ 762-737 1Primer set used for routine PCR mRNA transcript detection 2 Primer set used for generation of full-length PAR-2 for DNA cloning

Table 2.2 siRNA oligonucleotide templates used in the synthesis of siRNAs by pSilencer3.1-H1 puro siRNA Construction Kit.

Target Oligonucleotide Sequence Genbank Target Gene Name Designation Accension # Position PAR-2 PAR-2 siRNA sense 5'-GATCCAGGAAGAAGCCTTATTGGTTTCAA NM_005242 254-274 construct #1 GAGAACCAATAAGGCTTCTTCCTTTTTTTGGAAA-3' anti-sense 5'-AGCTTTTCCAAAAAAAGGAAGAAGCCT TATTGGTTCTCTTGAAACCAATAAGGCTTCTTCCTG-3' PAR-2 PAR-2 siRNA sense 5'-GATCCAGTAGACTTGGTGTGAAGATTCAA NM_005242 1635-1655 construct #2 GAGATCTTCACACCAAGTCTACTTTTTTTGGAAA-3 anti-sense 5'-AGCTTTTCCAAAAAAAGTAGACTTGGTG TGAAGATCTCTTGAATCTTCACACCAAGTCTACTG-3' PAR-2 PAR-2 siRNA sense 5'-GATCCGTAGTCGTGAATCTTGTTCATTCAA NM_005242 2190-2210 construct #3 GAGATGAACAAGATTCACGACTATTTTTTGGAAA-3' anti-sense 5'-AGCTTTTCCAAAAAATAGTCGTGAATCT TGTTCATCTCTTGAATGAACAAGATTCACGACTACG-3'

37

Figure 2.1 Vectors utilised in this PhD program of study. Vector maps of the (A) pGEMT-easy (taken from http://promega.com/vectors/t_vectors.htm [2/2/08]), (B) pEGFP-N1 (taken from http://www.clontech.com/images/pt/dis_vectors/PT3027- 5.pdf [2/2/08]), (C) pIB-V5-His (taken from http://tools. itrogen.com/content/sfs/vectors/pibv5his_map.pdf [2/2/08]) and (D) pSilencer 3.1- H1 puro vectors (taken from http://www.ambion.com/techlib/misc/vectors/3.1_ puro.html [2/2/08]) showing the multiple cloning sites and main functional elements.

2.1.7 Bacterial culture plates

Bacterial cultures were plated onto LB-agar plates with the appropriate antibiotic for selection of the plasmid of interest. Ampicillin and kanamycin were used at a concentration of 100 µg/mL and 50 µg/mL, respectively. Blue-white colony screening was performed for pGEM-T vector on LB-agar plates supplemented with 50 µl X-Gal (20 µg/mL) and 30 µl isopropyl-β-D-thiogalactoside (IPTG, 100 mM).

38 2.2 Methods

2.2.1 Cell culture

The non-tumorigenic and tumorigenic prostate epithelial derived cell lines, RWPE-1 and RWPE-2 respectively, and the prostate cancer derived cell lines LNCaP, PC-3, DU145 were from American Type Culture Collection (Manassas, VA). Lung murine fibroblasts (LMF) from PAR-1 null mice (Darrow et al., 1996) stably expressing human PAR-1, PAR-2 or PAR-4 (Andrade-Gordon et al., 1999) were from Johnson & Johnson Pharmaceutical Research and Development (Spring House, PA). These cells are designated PAR-1-LMF, PAR-2-LMF and PAR-4-LMF, respectively. Cells were grown in Dulbecco’s modified Eagle’s medium containing 10% (v/v) fetal calf serum, 100 units/mL penicillin and 100 μg/mL streptomycin and propagated in 95% air/5% CO2, at 37°C. Cultures of PAR-1-LMF, PAR-2-LMF and PAR-4-LMF murine fibroblasts were supplemented with 200 μg/mL hygromycin B. Insect Sf9 cells were grown in SF9002 serum free media containing 100 units/mL penicillin and 100 μg/mL streptomycin at 27°C.

2.2.2 Reverse transcription polymerase chain reaction

2.2.2.1 RNA extraction

Total RNA was extracted from harvested cells that were grown in six-well culture flasks using the TRIzol reagent according to the instructions of the manufacturer, except the final RNA precipitation step was carried out at -20°C instead of room temperature (RT). The precipitated RNA pellet was resuspended in 20 μL of diethylpyrocarbonate (DEPC) treated water. The resuspended RNA was treated with 1 U of DNase I for 15 mins at 37°C and then purified using an RNeasy Mini kit in accordance with the instructions of the manufacturer. RNA concentration was determined by spectrophotometric analysis (GeneQuant, Pharmacia Biotech,

Victoria, Australia) where 1 optical density (OD) at A260nm is equal to approximately

40 μg/mL of RNA. The 260/280 ratios were also assessed to ensure the sample was free from protein and DNA contamination (A ratio of 2.0 at λ=260 nm to λ=280 nm

39 indicates a preparation essentially free from protein and DNA contamination). All RNA preparations were stored at -80°C.

2.2.2.2 Reverse transcription

First-strand cDNA synthesis was carried out using the commercial Moloney murine leukemia virus reverse transcriptase (MMLV RT) SuperScript III. Typically, 2 µg of total RNA was combined with 10 nmol deoxyribonucleotides (dNTPs; Roche) and

250 ng oligo dT15 (Promega), and heated at 70°C for 5 min, followed by a chilling period longer than 1 min on wet ice. 5X first-strand buffer (4 µL), 2 µL 0.1 M dithiothreitol (DTT), and 200 U SuperScript III was mixed and the reaction was then incubated at 50°C SuperScript III for 1 h. The reverse transcription was heat- inactivated at 70°C for 10 min.

2.2.2.3 Polymerase Chain Reaction

PCRs were carried out in a PTC-200 Peltier thermal cycler DNA Engine (Bresatec, South Australia). A typical PCR reaction mixture contained 200 nM of primers (Table 2.1), 0.2 mM of dNTPs, and 1.25 U of Taq polymerase in the reaction buffer supplied by the companies and 10-100 ng of DNA template. Cycling conditions varied with the primers, template and requirement of enzymes, including a 5-min initial denaturation at 94-96oC, 20-35 cycles of amplification period consists of a denaturation at 94-95°C for 15-30 sec, an annealing1 at 55-65°C for 30 sec, and an extension2 at 65-72°C for 1-2 min per kb of the amplicon size, and a final extension at 65-72°C for 7 min. 1 Annealing temperatures are primer specific. Specific annealing temperatures are indicated in Table 2.1. 2 The extension temperature for PfuUltra DNA polymerase was 68°C and 72°C for Platinum Taq.

2.2.2.4 Agarose gel electrophoresis

DNA was separated on 0.8-2.0% (w/v) agarose gels, depending on the size of DNA

40 fragments. Ethidium bromide (0.5 μg/mL) was added into the gel solution prior to casting. Agarose loading dye (section 2.1.2) was spiked into samples and the gels were run in TAE buffer (section 2.1.2) at 100 V for 40-60 min depending on fragment size, and finally visualized in darkness using a ultra-violet (UV) transilluminator and photographed.

2.2.3 DNA cloning

2.2.3.1 Coding region generation and purification

PCR was carried out as described in section 2.2.2.3 with PfuUltra DNA polymerase. PCR amplicons were visualised by agarose gel electrophoresis and ethidium bromide staining under UV illumination. The band of interest (DNA insert) was excised from the agarose gel using a sterile scalpel blade. The DNA insert was purified by using a High Pure PCR Product Purification Kit according to the instructions of manufacturer.

2.2.3.2 Ligation and transformation

Where necessary, vectors and inserts were cut with appropriate restriction enzymes (Table 2.1), gel purified and examined by agarose gel electrophoresis. The inserts annealed from two chemically synthesised oligonucleotides (siRNA experiments) were treated with T4 polynucleotide kinase (T4 PNK) in a reaction containing 20 pmol DNA duplex, 1x T4 PNK buffer, 1.5 mM dATP, and 10 U of T4 PNK. A standard ligation was typically made up by DNA fragments, 1x T4 DNA ligation buffer and 1-5 U of T4 DNA ligase, with an overnight incubation at 4oC. Ligation reactions were used directly for the transformation of heat-shock competent cells.

Heat-shock competent cells in glycerol were thawed on wet ice, mixed with DNA, and incubated on wet ice for 30 min. Then the cells were heat-shocked using Falcon 2059 tubes in a 42°C water bath for 45 sec, chilled on wet ice for 2 min before a 1 h recovery in SOC medium 37°C. Recovery culture (10-200 µL) was plated on an LB agar plate containing the appropriate antibiotic for the transformed vector.

41 2.2.3.3 Screening transformed clones

Blue-white colony screening was performed for those vectors with the cloning site within a lacZ gene, such as pGEMT-easy vector. For other vectors, a modified alkaline lysis protocol was used. Bacterial lysis buffer (200 µl; section 2.1.2) was added directly to 200 µL of bacterial culture. After being neutralized by addition of 200 µL 5 M potassium acetate (pH 5.0), the mixture was centrifuged at 16,000 x g for 3 min. The plasmid DNA was precipitated by addition of an equal volume of isopropanol, followed by centrifuging at 16,000 x g for 2 min. The pellet was aspirated and reconstituted in 20 µL water. The colonies with plasmids of the expected size when visualised by agarose gel electrophoresis and ethidium bromide staining under UV illumination were selected for further plasmid purification by miniprep (section 2.2.3.4). DNA sequencing was then performed to check if the colony had the expected insert (section 2.2.3.6).

2.2.3.4 Plasmid DNA isolation and purification

High quality plasmid DNA for gene cloning and transfection was extracted and purified using the QIAprep Spin Miniprep kit following the instructions of the manufacturer. The quality of plasmids was assessed by agarose gel electrophoresis, and their concentrations were determined by spectrophotometric analysis at λ=260 nm.

2.2.3.5 Restriction digestion

Purified DNA was digested with restriction endonuclease at a ratio of 2-5 U per microgram of DNA for 1 h at 37°C before heat inactivation at 65°C for 20 minutes.

2.2.3.6 DNA sequencing

DNA sequencing reactions were carried out using an ABI BigDye Terminator Cycle Sequencing kit, with Australian Genomic Research Facility (AGRF) resolving the DNA products. A 20 μL sequencing reaction consisting of 2 μL BigDye terminator mix, 4 pmol primer specific for the vector (Table 2.1) and 400-600 ng of plasmid

42 DNA. Following a 5 min denaturation period at 94oC, a cycling condition of 96°C for 10 sec, 50°C for 10 s, and 60°C for 4 min was repeated for 35 cycles. Products were precipitated by addition of 75% (v/v) isopropanol before submission to AGRF.

2.2.4 Protein analysis

2.2.4.1 Whole cell protein extraction

Monolayer-cultured cells were lysed in a protein total extraction buffer (section 2.1.2) in the presence of a protease inhibitor cocktail (diluted 1:20 in the extraction buffer). After incubation on a rolling shaker for 1 h at 4oC, the samples were centrifuged at 14,000 rpm for 40 min to collect the supernatant. For ERK1/2 phosphorylation, cells were grown to 50% confluence, serum-deprived for 24 hours before treatment then collection of whole cell lysates as above except the lysis buffer also contained 1 mM sodium orthovanadate and 50 mM NaF.

2.2.4.2 Protein quantification

Protein concentrations were determined using a BCA assay kit according to the instructions of the manufacturer. A BSA protein standard ranging from 0.1-1 mg/mL was generated against which unknown protein concentrations were measured. Then 200 μL of working reagent (supplied in the BCA kit) was added to each well. Twenty-five μL of each BSA standard, blank control (diluent) and protein samples under test were added to each well, mixed by gentle rocking and then incubated for 30 min at 37oC. Once cooled to RT, absorbances of the samples were measured at 560 nm in a microplate reader.

2.2.4.3 Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis

SDS-PAGE using 10% or 12% resolving gels and 4% stacking gels was used to separate protein samples by molecular size. Approximately 20 μg of total protein was boiled for 5 min in loading buffer (section 2.1.2) and electrophoresed in running buffer (section 2.1.2) on a Protean II Minigel Apparatus (BioRad). The gel was run

43 at 100 V until the loading dye was approximately 1 cm from the bottom of the resolving gel. A pre-stained molecular weight protein marker (low range, BioRad) was used to determine the molecular sizes of the resulting bands.

2.2.4.4 Protein staining

SDS-PAGE gels were stained with either Silver Stain or Coomassie blue where indicated. Silver staining was performed using the Silver Stain Plus Kit according to instructions provided by the manufacturer. Coomassie blue staining was performed by incubating the polyacrylamide gel overnight at RT with gentle rocking in Coomassie blue gel staining buffer (section 2.1.2). The gels were then incubated with

10% (v/v) acetic acid, 45% (v/v) methanol in ddH2O (section 2.1.2) at RT with gentle rocking until protein bands were distinct from background staining.

2.2.4.5 Western blotting analysis

Proteins from whole cell lysates were resolved by SDS-PAGE and transferred to a nitrocellulose membrane in transfer buffer (section 2.1.2) at 100V for 1 h at 4°C. The membrane was blocked overnight at 4°C with Odyssey blocking buffer containing 0.1% (v/v) Tween 20 and then incubated overnight at 4°C with an anti-PAR2 (1:1000), anti-GAPDH (1:1000), anti-ERK (1:1,000), or anti-phospho-ERK (1:2,000) antibody. The membrane was then washed in TBS (section 2.1.2) containing 0.1% (v/v) Tween20 before incubation with species-appropriate fluorescently conjugated secondary antibodies for 1 h at room temperature, followed by a second series of washes in TBS containing 0.1% (v/v) Tween20 and 0.01% (v/v) SDS. Membranes were analysed using an Odyssey Infrared Imaging System (LiCOR) and where relevant signal intensity determined using LiCOR imaging software and exported to Microsoft Excel for graphical representation as mean +/- SEM. Significance was examined using a Student’s t-test with a P value of <0.05 considered significant.

44 2.2.5 Expression Constructs and transfections

Using the methods outlined in section 2.2.1, the previously published full-length KLK4 (NM_004917) coding sequence was excised using the restriction enzymes Xho1 and BamH1 from the previously published pDNA3.1:K4 construct (Harvey et al., 2003) then ligated into the insect expression Xho1/BamH1 digested plasmid pIB- V5-His in frame with the V5-His tags and stop codon (Figure 2.1). This construct generates the complete KLK4 amino acid sequence followed by V5 (GKPIPNPLLGLDST) and 6 x histidine tags. Insect Sf9 cells were transfected with this KLK4-pIB/V5-His construct using Cellfectin in accordance with the instructions of the manufacturer.

A mammalian expression construct encoding PAR-2 with GFP at the C-terminus was generated in the pEGFP-N1 vector. PCR employing Pfu DNA polymerase was used to amplify the PAR-2 coding region. Following amplification and purification, the PAR-2 PCR product was ligated into the pGEMT-easy shuttle vector. The pGEMT- easy vector containing the full-length PAR-2 cDNA was propagated, purified and restriction digested with Xho1/EcoR1 and ligated into Xho1/EcoR1 digested pEGFP- N1. Mammalian cells were transiently transfected using Lipofectamine in accordance with the instructions of the manufacturer. The sequences of both constructs were verified using DNA sequencing.

2.2.6 Expression and purification of recombinant pro-KLK4-V5-His

Following transfection with the pIB:KLK4-V5-His construct, stable Sf9 cells were selected with 50 μg/mL Blasticidin for a 2 week period. pro-KLK4-V5-His was purified from conditioned media from these cells using Ni-NTA superflow resin by following the instructions of the manufacturer. This involved washing the column 3 times with Ni column washer buffer (section 2.1.2), 3 times with Ni column wash buffer containing 30 mM imadazole, before pro-KLK4-V5-His was eluted from the column with column wash buffer containing 250 mM imadazole in 1 mL fractions. pro-KLK4-V5-His containing fractions, identified by analysis of a Coomassie stained polyacrylamide gel, were pooled then concentrated by nitrogen gas mediated

45 evaporation. Following dialysis against PBS (section 2.1.2) at 4°C overnight pro- KLK4-V5-His was aliquoted and stored at -80°C.

2.2.7 Recombinant pro-KLK4-V5-His activation

Recombinant pro-KLK4-V5-His was incubated with the metalloendopeptidase thermolysin in PBS at pH 7.4 for 1 hour at 37°C (KLK4: thermolysin; 80:1). The amount of active enzyme produced was quantified by active site titration using the pseudo-suicide inhibitor methylumbelliferyl ester of p-guanidinobenzoate (Jameson et al., 1973). pro-KLK4-V5-His and thermolysin treated pro-KLK4-V5-His were examined on a Coomassie stained polyacrylamide gel, protein bands were excised and the amino terminal of the excised treated KLK4-V5-His sequenced by Edman degradation at the Australian Proteome Analysis Facility (North Ryde, Australia). Thermolysin activity was stopped by the addition of phosphoramidon (10 μM).

2.2.8 Kinetic measurements of activated KLK4-V5-His

Determination of kinetic parameters was performed using 40 nM of active KLK4 against increasing concentrations (0 μM, 32.5 μM, 75 μ M, 150 μM, 300 μM, 600 μM) of the tri-peptide substrate benzoyl(Bz)-phenylalanine(F)-valine(V)- arginine(R)-paranitroaniline(pNA)(referred to in subsequent chapters as F-V-R- pNA). Assays were performed in 50 mM Tris–HCl (pH 7.4), 20 mM CaCl2, 0.01% (v/v) Tween20 at 28°C by following pNA release photometrically at 405 nm.

Experiments to determine the kinetics of cleavage of a fluorescence quenched peptide o-aminobenzoic acid-(Abz)-serine(S)-lysine(K)-glycine(G)-arginine(R)- serine(S)-leucine(L)-isoleucine(I)-glycine(G)-lysine(K)-2,4- Dinitrophenol(Dnp) (referred to in subsequent chapters as Abz-SKGRSLIGK-Dnp) with sequence spanning the PAR-2 serine protease activation site, was performed over the concentration range of 0 μM, 0.5 μM, 2.5 μM, 5 μM, 10 μM, 25 μM, 50 μM and 100

μM of substrate at 37°C in 50 mM Tris–HCl (pH 7.4), 20 mM CaCl2, 0.01% (v/v) Tween20 with continuous monitoring of fluorescence at 440 nm following excitation at 330 nm. Emissions were monitored over 15 minutes using a Polarstar Optima plate reader (BMG Labtech, Melbourne, Australia). Enzyme activity was determined from

46 a standard curve generated from the fluorescence obtained following complete cleavage by trypsin of known amounts of peptide using an absorption coefficient of 104 M−1 cm−1.

To obtain kcat and Km values, initial velocity data at each substrate concentration were fitted to the Michaelis Menten equation by non-linear regression analysis using GraphPad Prism 4.0 (GraphPad Software, San Diego, CA). Experiments were performed in triplicate on 3 independent occasions with results displayed as means +/- standard error of the mean (SEM).

2.2.9 Ca2+ flux assays

Cells grown to 80% confluence were washed with PBS, detached non-enzymatically with versene, resuspended (4×106 cells/mL) in extracellular medium (section 2.1.2), then loaded with the fluorescence indicator FURA-2 acetoxymethyl ester (1.0 μM) at room temperature for 60 minutes. Cells were then pelleted followed by resuspension in EM (without BSA) at a concentration of 2×106 cells/mL for fluorescence measurements. The ratio of fluorescence at 510 nm after excitation at 340 nm and 380 nm was monitored using a Polarstar Optima fluorescent plate reader. Single agonist treatments were active KLK4 (300 nM), trypsin (10 nM), thrombin (10 nM), PAR-1 AP and PAR-2 AP (100 μM) and PAR-4 AP (500 μM). Displayed data are representative of experiments performed in triplicate and repeated on 3 independent occasions. Concentration response experiments were performed over the KLK4 concentration range 0.1 to 1000 nM in triplicate and performed on 3 independent occasions with results displayed as means +/- SEM.

2.2.10 siRNA knockdown of PAR-2 expression

The mammalian siRNA expression vector pSilencer 3.1-H1 puro was used to reduce expression of PAR-2. Candidate PAR-2 siRNA target sequences were designed as previously described (Pei and Tuschl, 2006) then aligned against the human genome database using the BLAST algorithm to eliminate those with significant homology to other genes. The sequences selected (Table 2.2) were inserted into the pSilencer 3.1-

47 H1 puro vector as outlined in section 2.2.3. Fibroblasts from Par-1 null mice stably expressing PAR-2 (PAR-2-LMF) were transfected with the PAR-2 pSilencer 3.1-H1 puro constructs or the supplied pSilencer 3.1-H1 puro negative control using Lipofectamine. After 48 hours 2 μg/mL of puromycin was added to the medium and stably transfected cells selected following a 2 week selection period. PAR-2 protein levels and agonist induced induction of ERK phosphorylation were examined by Western blot analysis (section 2.2.4).

2.2.11 Cell surface biotinylation

PC-3 cells were washed 3 times with cold PBS then plasma membrane proteins biotinylated by incubation with 1.22 mg/mL EZ-link NHS-SS Biotin at 4°C for 1 hour with gentle agitation. The cells were then washed in PBS prior addition of protein total extraction buffer. After centrifugation, bead immobilised streptavidin was added into the supernatant and incubated for 15 min on ice to capture biotinylated proteins. The streptavidin beads were pelleted by centrifugation and the supernatant recovered for analysis of cytoplasmic (non-biotinylated) proteins. The beads were then washed thoroughly and associated cell surface (biotinylated) proteins eluted into Laemmli sample buffer (60 mM Tris-HCl, pH 6.8, 10% (v/v) glycerol, 2% (w/v) SDS, 1% (v/v) 2-mercaptoethanol and 0.002% (w/v) bromophenol blue). Cytoplasmic and cell surface fractions were examined for PAR-2 by Western blot analysis.

2.2.12 Flow cytometry

Cells grown in serum free media overnight were detached non-enzymatically with versene, washed in PBS then resuspended at 2×106 cells/mL. Cells were subjected to non-permeabilising fixation in 1% (w/v) formaldehyde on ice for 5 min, washed with PBS with 0.2% (w/v) BSA. Following incubation with an anti-PAR-2 antibody (SAM11; 2 μg/106 cells) on ice for 60 min, cells were washed then incubated with fluorescently tagged anti-mouse secondary antibody. Cell surface PAR-2 was assessed using an FC500 flow cytometer (Beckman Coulter, Gladesville, Australia).

48 2.2.13 Confocal microscopy

Cells plated on sterile poly-L-lysine coated glass coverslips were allowed to adhere overnight and then fixed with 4% (w/v) formaldehyde for 30 min at room RT and permeabilised with 0.1% Triton X-100 in PBS for 10 min. Cells were blocked for 30 mins in 0.2% BSA in PBS and incubated with the primary antibody for 1 h at 37°C. The cells were then incubated with the species appropriate fluorescently conjugated secondary antibody, counterstained with DAPI diluted in PBS and mounted on slides.

For the detection of PAR-2 GFP, the PAR-2-pEGFP-N1 or only the pEGFP-N1 vectors were transfected into cells adhered to poly-L-lysine coated glass coverslips. For agonist treatments cells were incubated with either KLK4 (100 nM) or PAR-2 AP (100 μM) for 10 minutes at 37°C before fixation with 4% (v/v) formaldehyde. Nuclei were stained by incubating cells for 5 minutes at room temperature with DAPI (1:1500) in PBS. Coverslips were mounted on slides and cells imaged with a Leica SP5 confocal microscope (Leica Microsystems, Sydney, Australia). Images were processed using Adobe Photoshop CS3 and displayed using CorelDraw.

The amount of PAR-2 on the cell surface was quantified by examining the fluorescence of randomly selected untreated and KLK4 treated cells (n = 15) using ImageJ software (National Institutes of Health, Bethesda, MA) adapting the approach of Scherrer and co-workers for quantifying GPCR cell surface expression (Scherrer et al., 2006). Briefly, three cellular regions of interest were defined - the whole cell, the intracellular region and the nucleus. The signal obtained for the whole cell and the intracellular region was corrected for background signal by subtracting nuclear fluorescence. Cell surface fluorescence was then obtained by subtracting the corrected value for the intracellular region from the corrected value for the whole cell. These values were then divided by the number of pixels contained within each region to give fluorescence density values for the cell surface (Di surface) and cytoplasm (Di cytoplasm). The ratio of Di surface to Di cytoplasm was determined to normalize data across the counted cell population. Results are displayed graphically as mean +/- SEM and significance was examined using a Student’s t-test with a P value of <0.05 considered significant.

49

2.2.14 Immunohistochemistry

Archival formalin-fixed paraffin-embedded blocks from primary prostate cancers from men with more advanced disease (n = 6; Gleason scores 3+4 to 4+5 (medium to poorly differentiated)) and prostate cancer bone metastases (n = 2) were obtained from Sullivan Nicolaides Pathology (Taringa, Australia) and the Royal Prince Alfred Hospital (Sydney, Australia), respectively, following institutional ethics approval. Immunohistochemistry was performed as previously described (Hooper et al., 1999). Briefly, serial sections (4 μm), were deparaffinised then rehydrated and after antigen retrieval in urea (5% w/v) in 0.1M Tris buffer (pH 9.5) incubated in H2O2 (3% v/v) to quench endogenous peroxidise. Sections were then blocked in normal goat serum (10% w/v) and incubated overnight at 4°C with either an anti-PAR-2 monoclonal (SAM11; 1:1000 dilution) or an anti-KLK4 rabbit polyclonal (1:250 dilution) antibody raised against the first 12 residues of the amino terminal of the active enzyme (Harvey et al., 2003). As negative controls mouse and rabbit immunoglobulins replaced the SAM11 and anti-KLK4 primary antibodies respectively. The EnVision+ peroxidise polymer detection system was used with DAB as the chromogen. The sections were counterstained with Mayer’s haematoxylin, visualized by microscopy (Leitz, Laborlux S, Germany) and using a Nikon OXM1200 digital camera. Images were processed using Adobe Photoshop CS3 and displayed using CorelDraw.

50

CHAPTER 3

GENERATION AND CHARACTERISATION OF RECOMBINANT HUMAN

KALLIKREIN-RELATED PEPTIDASE 4

51 3.1 Introduction

As stated in Chapter 1 the hypothesis of this project is that the trypsin-like serine protease KLK4, may be an activator of PARs in the prostate. To perform the in vitro assays required to test this hypothesis, recombinant KLK4 was needed.

As insect expressed recombinant proteins have many of the post-translation modifications of mammalian proteins (Possee, 1997), a feature bacterial and yeast expression systems lack (Sahdev et al., 2008), an insect cell expression system was chosen for expression of KLK4. In recent years, vector systems that utilise baculovirus promoters to access insect host cell transcriptional machinery have been developed, eliminating the need for viral infection for recombinant protein production in insect cells (Possee, 1997). The pIB/V5-His vector utilised in this study, drives protein expression through the baculiovirus immediate-early promoter OpIE2, derived from the baculovirus Orgyia pseudotsugata. Although the natural host for this virus is the Douglas fir tussock moth, the promoters allow efficient protein expression in a variety of cell lines, including Sf9 cells, derived from Spodoptera frugiperda ovarian cells (Hegedus et al., 1998), which were utilised in this study. Before in vitro PAR activation studies could be performed, protocols needed to be developed to activate pro-KLK4, to perform active site titration of the active protease and to investigate other kinetic parameters.

Active site titration is a method to establish the fraction of enzyme sites that are active. Due to the presence of an aspartate residue at the bottom of the first substrate binding pocket (aspartate189) (Nelson et al., 1999), KLK4 has trypsin-like specificity, which is the preference for cleavage following basic amino acids (Takayama et al., 2001a; Obiezu et al., 2006; Debela et al., 2006b). A common approach for titrating the active sites of trypsin-like serine proteases (such as trypsin, plasmin and plasma kallikrein) is to employ the substrate nitrophenyl ester p- guanidinobenzoate (NPGB) (Chase and Shaw, 1969). The mechanism for this reaction involves the rapid acylation of the enzyme with a stoichiometric release of nitrophenol followed by very slow deacylation of the resultant acyl enzyme, allowing spectroscopic measurement of released nitrophenol (Chase and Shaw, 1969). The primary limitation of this method, a lack of sensitivity, is overcome by using the

52 fluorogenic analog methylumbelliferyl ester of p-guanidinobenzoate (Jameson et al., 1973). While the ester is non-fluorescent, the released product 4- methylumbelliferone emits fluorescence and can be quantified against a standard curve of known concentrations of this reaction product.

During our generation of KLK4, Matsumura and colleagues reported production of a recombinant form of this enzyme. These researchers generated KLK4 from S2 insect cells in which the endogenous activation site had been replaced by an enteropeptidase cleavage site (DDDDK). The peptide substrate specificity of the active enzyme, defined by screening a tetrapeptide positional scanning synthetic combinatorial library, was P1-arg; P2-gln/leu/val; P3-gln/ser/val and P4-ile/val (Matsumura et al., 2005). Screening proteolytic activity towards pNA tri-peptides revealed that a peptide with the sequence P1-arg, P2-val and P3-phe displayed the greatest KLK4 hydrolysis rate (Matsumura et al., 2005). Accordingly, to permit direct comparison to previously published data, this substrate was used to determine the kinetic parameters of our recombinant KLK4.

Described in this chapter are experiments used to generate and purify full-length V5/His tagged recombinant KLK4 from the Sf9 insect cell line. The biochemical properties of recombinant KLK4 were characterised by assessing N-glycosylation content, zymogen activation, active site titration of the mature enzyme, peptide hydrolysis rates and enzyme inhibition.

3.2 Materials and Methods

These are comprehensively described in Chapter 2. The reagents used specifically in experiments performed in this chapter are described below.

The previously published full-length KLK4 (NM_004917) coding sequence was excised using the restriction enzymes Xho1 and BamH1 from the previously published pDNA3.1:K4 construct (Harvey et al., 2003) then ligated into the insect expression Xho1/BamH1 digested plasmid pIB-V5-His in frame with the V5-His tags and stop codon (Figure 2.1). The integrity of the construct was confirmed by dye terminator sequencing. (Sub-cloning was performed by Mr Nigel Bennet).

53

To generate an insect cell expression construct encoding the serine to alanine mutant pro-KLK4-V5-His, site directed mutagenesis using a Stratagene mutagenesis kit was performed according to the instructions of the manufacturer on the pDNA3.1:KLK4- V5-His construct. Mutant KLK4 was excised and inserted into the insect expression plasmid pIB-V5-His as described above. The integrity of the construct was confirmed by dye terminator sequencing. (Mutagenesis was performed by Mr Nigel Bennet).

For production of recombinant wild-type and mutant pro-KLK4-V5-His, Sf9 insect cells were transfected with wild-type and mutant pIB:KLK4-V5-His constructs. Stable Sf9 cells were selected with 50 μg/mL blasticidin following a 2 week selection period. Recombinant pro-KLK4-V5-His proteins were purified from conditioned media from these cells using Ni-NTA superflow resin by following the instructions of the manufacturer. (Insect cell work and protein purification was performed by Ms Melanie Hunt).

3.3 Results

3.3.1 Production of recombinant KLK4 from Sf9 insect cells (the expression constructs used here were generated by Mr Nigel Bennett. Ms Melanie Hunt performed the work with insect cells to generate and purify recombinant KLK4).

To perform the in vitro assays required to test the hypothesis of KLK4-mediated activation of PAR family members, recombinant KLK4 was produced in Sf9 insect cells. The full-length KLK4 expression construct encodes a 274 amino acid protein (NP_004908) which includes pre- and pro- regions (Figure 3.1A). KLK4 also contains histidine57, aspartate102 and serine195 residues in the highly conserved consensus motif which is essential for the catalytic activity of serine proteases and 6 disulfide bridges formed between cysteine residues, which are also highly conserved among members of this protease family (Figure 3.1A). Further, the recombinant pro-KLK4-V5-His also contains the V5 epitope for detection by Western blot analysis and Histidine x 6 (His) tag at the C-terminus for protein purification (Figure 3.1A).

54

One litre of media from polyclonal Sf9 insect cells that had been selected with 50 μg/mL blasticidin for stable incorporation of the pIB:KLK4-V5-His plasmid was applied to a nickel resin column and the histidine tagged recombinant pro-KLK4-V5- His allowed to bind to the nickel groups. Following a series of low stringency salt washes, bound KLK4 was eluted in 1 mL fractions from the resin with buffer containing increasing concentrations of imadazole (elutions 1-5; Figure 3.1B). The elutions, starting material (SM) and flowthrough (FT) were resolved by SDS-PAGE on a 10% gel under reducing conditions. Commassie staining the gel showed the presence of pro-KLK4-V5-His in each fraction migrating at ~35 kDa. KLK4 contains a consensus N-glycosylation site at 169NVS (N-X-S motif; where X can be any amino acid residue). Post-translational modification at this site may account for the difference in size of recombinant pro-KLK4-V5-His at 35 kDa compared to the predicted 29.3 kDa (KLK4 =27.0, V5 and His= 2.3 kDa) (Figure 3.1B).

55

Figure 3.1 Expression and purification of recombinant pro-KLK4-V5-His from Sf9 insect cells. A. Schematic representation of the full-length 274 amino acid (aa) pro-KLK4-V5-His protein produced from stable incorporation of the pIB:KLK4-V5- His vector into Sf9 insect cells. Structural features marked on the schematic include; 26 amino acid pre-region, 6 amino acid pro-region, the catalytic triad residues; histidine57, aspartate102 and serine195, disulphide bridges (designated by dashed line), N-linked glycosylation of asparagine169, V5 epitope for detection by Western blot analysis and Histidine x 6 (His) tag for protein purification. B. Commassie stained 10% gel run under reducing conditions of conditioned media from Sf9 cells stably expressing pro-KLK4-V5-His (starting material; SM), with unbound proteins allowed to flow through (FT). Successive imadazole elutions removed the bound pro-KLK4-V5-His from the column as a ~35 kDa protein.

3.3.2 N-glycosylation of KLK4

The molecular weight of the peptide backbone of the full-length untagged and V5- His tagged purified KLK4 (pro-KLK4-V5-His) is predicted to be 27 kDa and 29.3 kDa respectively. However, purified pro-KLK4-V5-His migrated at ~35 kDa on a Commassie stained 10% gel run under reducing conditions. As indicated in section 3.3.1 KLK4 contains a consensus N-glycosylation site at 169NVS (N-X-S motif;

56 where x can be any amino acid residue). To determine if N-glycosylation contributes to the additional molecular weight of pro-KLK4-V5-His, deglycosylation with the enzyme N-glycosidase F was employed.

N-glycosidase F (PNGase F) removes N-linked glycosylation by cleaving amide bonds between the N-acetylglucosamine moiety and the asparagine amino acid of high mannose, hybrid, and complex oligosaccharides (Maley et al., 1989). Purified recombinant pro-KLK4-V5-His was incubated with this enzyme and Western blot analysis using an anti-V5 antibody following SDS-PAGE on a 10% gel run under reducing conditions indicated that PNGase F removed ~2 kDa of N-linked glycosylation, reducing the molecular weight of pro-KLK4-V5-His to ~33 kDa (Figure 3.2). For comparative purposes, untreated and PNGase F treated lysates and media from HeLa cells transfected with a mammalian pro-KLK4-V5-His expression construct were analysed. Western blot analysis with an anti-V5 antibody indicated that pro-KLK4-V5-His from human cells migrated at a similar molecular weight to insect cell produced pro-KLK4-V5-His and that this protein is modified to approximately the same extent by N-glycosylation in human and insect cells. (Figure 3.2)

Figure 3.2 N-glycosylation of insect cell and human cell expressed pro-KLK4- V5-His. Western blot analysis with an anti-V5 antibody following SDS-PAGE on a 10% gel run under reducing conditions, indicated that deglycosylation by N- glycosidase F (PNGase F) of recombinant pro-KLK4-V5-His purified from Sf9 insect media and cell lysate and conditioned media from HeLa cells transiently transfected with pcDNA3.1:KLK4-V5-His, removed ~2 kDa of sugar residues from both the insect and human expressed recombinant pro-KLK4-V5-His proteins.

57 3.3.3 Activation of KLK4 by thermolysin

For kinetic characterisation and PAR activation experiments, conversion of recombinant zymogen KLK4 (pro-KLK4-V5-His) to its active mature form (KLK4- V5-His) was required. Unlike the zymogen forms of other KLK family members which have a basic residue (arginine or lysine) at the activation site, the pro-region of KLK4 ends in a glutamine, with the start of the mature peptide beginning with an isoleucine. The metalloendopeptidase, thermolysin, preferentially cleaves peptides and proteins on the N-terminal side of leucine, phenylalanine, isoleucine and valine residues (Heinrikson, 1977). Importantly, Ryu and colleagues have successfully used thermolysin to convert zymogen porcine KLK4 to its mature form (Ryu et al., 2002). Accordingly, thermolysin was used to activate the recombinant pro-KLK4-V5-His used in this study.

Thermolysin and pro-KLK4-V5-His in molar ratios 1:5 and 1:20 thermolysin to KLK4 were incubated for 1 h at 37˚C before the reaction was terminated with the addition of 10 μM phosphoramidon, a specific inhibitor of thermolysin (Suda et al., 1973). A reduction in size of pro-KLK4-V5-His from 35 kDa to 34 kDa, consistent with loss of the pro-region and formation of the mature form, was observed when resolved by SDS-PAGE on a 10% gel run under reducing conditions and visualised with a silverstain (Figure 3.3A). An overnight incubation caused a reduction in size to 30 kDa, suggesting further thermolysin mediated cleavage over this time period, or autocatalysis by KLK4-V5-His once activated by thermolysin (Figure 3.3A).

As downstream assays required the presence of minimal amounts of thermolysin and its inhibitor phosphoramidon, the activation protocol was adjusted to a 1:80 molar ratio of thermolysin to KLK4 for 1 h at 37˚C before the addition of 10 μM phosphoramidon. SDS-PAGE on a 12% gel run under reducing conditions and visualised with a Commassie stain displayed a similar size reduction from 35 kDa to 34 kDa, suggestive of pro-region removal (Figure 3.3B). To confirm the actual site of cleavage, the bands apparent in untreated and thermolysin treated KLK4, were excised from the gel and sequenced by Edman degradation N-terminal sequencing. This analysis indicated that pro-KLK4-V5-His had been cleaved between

58 glutamine30 and isoleucine31 giving a mature protein predicted to be catalytically active (Figure 3.3B).

Figure 3.3 Removal of pro-KLK4-V5-His pro-region by thermolysin. A. SDS-PAGE on a 10% gel run under reducing conditions with protein visualisation by silverstain of 1 μg of pro-KLK4-V5-His incubated with thermolysin in the molar ratios 1:20 and 1:5 thermolysin to KLK4 for 1 h or overnight at 37˚C before the reaction was ended with 10 μM phosphoramidon. B. SDS-PAGE on a 12% gel run under reducing conditions with protein visualisation with a Commassie stain of 4.0 μg of 1:80 thermolysin to KLK4 molar ratio incubated for 1 h at 37˚C before the reaction was ended with 10 μM phosphoramidon. The protein band for the thermolysin treated pro-KLK4-V5-His was excised and N-terminally sequenced using Edman degradation sequencing to confirm pro-region removal. The resulting N-terminal sequence is displayed beside the 34 kDa band.

59 3.3.4 Characterisation of thermolysin activated KLK4

Having successfully removed the pro-region of pro-KLK4-V5-His to generate a protease predicted to be catalytically active, active site titration was performed to determine the actual amount of active KLK4-V5-His, with kinetic analysis against the previously published KLK4 tri-peptide substrate F-V-R-pNA to determine the level of KLK4-V5-His activity.

Active site titration of KLK4-V5-His was performed with methylumbelliferyl p- guanidinobenzoate (MUGB). The release of 4-methylumbelliferone in a 1:1 stoichiometric ratio (Figure 3.4A) enables the quantitation of the conversion, by thermolysin, of KLK4 zymogen (pro-KLK4-V5-His) to mature KLK4 (KLK4-V5- His) when correlated to a standard curve generated from known quantities of 4- methylumbelliferone (Figure 3.4B). The input of 8.35 μM pro-KLK4-V5-His into an 80:1 molar ratio with thermolysin at 37˚C for 1 h before the reaction was stopped with 10 μM phosphoramidon generates 2.5 μM of KLK4-V5-His (30% conversion) (Figure 3.4C). Importantly, the same amount of untreated pro-KLK4-V5-His as well as the activating amount of 10 μM phosphoramidon inhibited thermolysin both showed no activity (Figure 3.4C).

To determine the kinetics of proteolytic cleavage, the pNA conjugated tri-peptide F- V-R-pNA identified by Matsumura et al as having the greatest hydrolysis by KLK4 was used (Matsumura et al., 2005). Monitoring release of pNA from F-V-R-pNA by KLK4-V5-His over time determines the reaction velocity (Figure 3.5A). Before in depth kinetic analysis to determine the kinetic parameters of KLK4-V5-His activity could be performed, the optimal pH for hydrolysis of F-V-R-pNA was first determined. By analysing the velocity of pNA release from 750 μM F-V-R-pNA by KLK4-V5-His at a pH range from 3-11, an optimal pH for KLK4-V5-His activity was determined as pH 8, with greater than 50% of maximal activity observed between pH 6-10 (Figure 3.5B).

60

Figure 3.4 Active site titration of KLK4-V5-His. A. Schematic depicting active site complex formation between KLK4-V5-His with MUGB causing the release of the fluorescent product 4-methylumbelliferone which can be measured using an excitation at 365 nm and monitoring emission at 445 nm. B. Quantitation of the released 4-methylumbelliferone, which represents the amount of active KLK4-V5- His due to the 1:1 stoichiometric ratio between KLK4-V5-His and MUGB, is possible when correlated to a standard curve generated from known quantities of 4- methylumbelliferone C. Conversion percentage of pro-KLK4-V5-His to active KLK4-V5-His from a 1:80 thermolysin to KLK4 molar ratio incubation. The negative controls pro-KLK4-V5-His only and phosphoramidon inactivated thermolysin only are also displayed. Data is representative of the average of triplicate experiments +/- SEM.

61

Figure 3.5 pH spectrum of KLK4-V5-His activity. A. Schematic depicting KLK4- V5-His cleavage of the F-V-R-pNA tri-peptide, after the arginine residue in the P1 position, to liberate pNA for spectrophotometric measurement at 405 nm. Monitoring release of pNA from F-V-R-pNA over time determines the reactions velocity (activity). B. Activity levels of 40 nM KLK4-V5-His against 750 μM F-V-R-pNA at increasing pH levels. Data points represent velocity over 3 minutes at each pH interval relative to the maximal velocity for the experiment; obtained at pH 8.0. Measurements are represented by the mean values of triplicate experiments +/-SEM.

As this pH range agreed closely with data from the study by Matsumura et al (Matsumura et al., 2005) the kinetic analysis of KLK4-V5-His was performed at the same pH used by these workers (pH 8.8), thereby permitting ready comparison of data. Kinetic parameters were determined by assaying velocities of a fixed amount of KLK4-V5-His (40 nM) and trypsin (2 nM), quantitated by active site titration, against increasing concentrations of F-V-R-pNA (0 μM, 32.5 μM, 75 μM, 150 μM, 300 μM, 600 μM). Fitting the velocities to the Michaelis-Menten equation yielded -1 the kinetic constants of: Km=47 +/- 6.3 μM, kcat=1.8 +/- 0.12 s , kcat/ Km=38 +/-1.9

62 -1 -1 (s. mM) for KLK4 (Figure 3.6A) and Km=74 +/- 3.4 μM, kcat=248 +/- 1.6 s , kcat/ -1 Km=3351 +/- 470 (s. mM) for trypsin (Figure 3.6B).

Figure 3.6 Velocities of KLK4-V5-His and trypsin hydrolysis of the F-V-R-pNA tri-peptide. The Lineweaver-Burke plots of KLK4-V5-His (A)(40 nM) and trypsin (B)(2nM) derived from the double reciprocals of velocities (y-axis; 1/V(μM s-1) of the enzyme at increasing concentrations (32.5 μM, 75 μM, 150 μM, 300 μM, 600 μM) of the tri-peptide F-V-R-pNA (x-axis; 1/[S](μM)). Kinetic constants calculated from the graphs are displayed beside the respective plot. Velocities represented are the means of triplicate experiments +/- SEM.

3.3.5 Defining the importance of the catalytic serine residue in KLK4 activity

Having determined the activity of KLK4-V5-His following removal of the pro- region by thermolysin, the importance of the catalytic serine for the activity of this

63 enzyme was next examined by determining the activity levels of a mutant KLK4-V5- His containing a substituted alanine residue for the catalytic serine. To generate the cDNA encoding the mutant pro-KLK4-V5-His, site directed mutagenesis was utilised (Figure 3.7A). The mutant pIB:KLK4-V5-His plasmid was expressed in Sf9 insect cells and affinity purified similarly to wild-type pro-KLK4-V5-His. Like wild- type pro-KLK4-V5-His, removal of the pro-region of mutant pro-KLK4-V5-His by thermolysin in a molar ratio of 1:80 thermolysin to mutant KLK4 resulted in a size reduction from 35 kDa to 34 kDa (Figure 3.7B) and PNGase F mediated carbohydrate removal reduced protein size from 35 kDa to 33 kDa (Figure 3.7C), both identical characteristics to wild-type KLK4.

Assessment of the activity of equal amounts of mutant KLK4-V5-His (30 nM), based on conversion efficiency of thermolysin determined for wild-type KLK4-V5-His, in comparison to wild-type KLK4-V5-His (30 nM) was determined by the hydrolysis of increasing concentrations (32.5 μM, 75 μM, 125 μM, 250 μM, 500 μM and 750 μM) of the peptide F-V-R-pNA. Wild-type KLK4-V5-His showed increasing levels of activity from 32.5 μM to 500uM F-V-R-pNA, where maximal activity was achieved (Figure 3.8B). In contrast, mutant KLK4-V5-His showed no appreciable activity at any of the substrate concentrations (Figure 3.8B), due to an inability to liberate pNA from the F-V-R-pNA tri peptide substrate, highlighting the importance of the catalytic serine for proteolytic activity. The minimal velocity rates recorded at all substrate concentrations for mutant KLK4-V5-His are attributable to spontaneous hydrolysis of the F-V-R-pNA peptide as the vehicle control, consisting of phosphoramidon inhibited thermolysin proportional to that at each interval, displayed similar velocities. Due to limited amounts of mutant pro-KLK4-V5-His, only one replicate could be performed for this comparative experiment.

64

Figure 3.7 Comparison of insect cell expressed recombinant mutant pro-KLK4- V5-His and wild-type pro-KLK4-V5-His. Schematic representation of the full- length 274 amino acid (aa) pro-KLK4-V5-His protein produced from stable incorporation of the pIB:KLK4-V5-His vector into Sf9 insect cells. Structural features marked on the schematic include; 26 amino acid pre-region, 6 amino acid pro-region, the catalytic triad residues; histidine57, aspartate102 and alanine195 (substitution from serine195), disulphide bridges (designated by dashed line), N- linked glycosylation of asparagine169, V5 epitope for detection by Western blot analysis and Histidine x 6 (His) tag for protein purification. B. SDS-PAGE on a 10% gel run under reducing conditions and Commassie stain of 1 μg of a 1:80 molar ratio of thermolysin to mutant pro-KLK4-V5-His incubated for 1 h at 37˚C before the reaction was terminated with 10 μM phosphoramidon. C. Western blot analysis and detection with an anti-V5 antibody following SDS-PAGE on a 10% gel run under reducing conditions of mutant and wild-type pro-KLK4-V5-His and thermolysin activated mutant and wild-type KLK4-V5-His before and after deglycosylation by PNGaseF.

65

Figure 3.8 Comparison of wild-type and mutant KLK4-V5-His velocities. A. Schematic depicting a lack of hydrolysis by mutant KLK4-V5-His of the tri-peptide F-V-R-pNA, resulting in an inability to liberate pNA from the tri-peptide for spectrophotometric measurement at 405 nm. B. Velocity comparison of (30nM) wild-type and (30nM) mutant KLK4-V5-His at increasing F-V-R-pNA (32.5 μM, 75 μM, 125 μM, 250 μM, 500 μM and 750 μM) substrate concentrations. The vehicle control represents phosphoramidon inhibited thermolysin proportional to the amount present at each substrate interval. Data points represent velocity at each substrate concentration interval relative to the maximal velocity for the experiment; obtained at 500 μM for wild-type KLK4-V5-His. Measurements are represented by the mean values of triplicate replicates of a single experiment.

3.3.6 Inhibition of KLK4 activity

The ability of thermolysin activated KLK4 to form a stable complex with the serine protease inhibitor aprotinin was examined by incubating protease and inhibitor in the molar ratio 4:1 for 1 h at 37˚C. Reactions products were analysed by SDS-PAGE on a 10% gel run under non-reducing conditions.

As shown in Figure 3.9 aprotinin was able to form a stable complex with KLK4-V5- His, as indicated by the shift under non-reducing conditions from 29 kDa to 35 kDa;

66 consistent with the molecular weight of aprotinin (6 kDa) (Figure 3.9; lane 4). Importantly, aprotinin was unable to complex with pro-KLK4-V5-His in the 4:1 molar ratio aprotinin to KLK4 zymogen (Figure 3.9; lane 2). Further, only a proportion of KLK4-V5-His (Figure 3.9; lane 4, lower band) was able to complex with aprotinin (Figure 3.9; lane 4, upper band), consistent with active site titration results that showed a proportion (30%) of the zymogen was converted to mature enzyme (Figure 3.4B). Interestingly, pro-KLK4-V5-His and thermolysin activated KLK4-V5-His migrated to 31 kDa and 29 kDa respectively (Figure 3.9; lane 1 and 3), a smaller molecular weight than when pro-KLK4-V5-His and KLK4-V5-His are run under reducing conditions (35 kDa and 34 kDa respectively; Figure 3.3B). This is likely due to the compact nature of the protein when the disulphide bonds are left intact under non-reducing conditions.

Figure 3.9 Aprotinin complex formation with zymogen and mature form KLK4- V5-His. Aprotinin incubated for 1 h at 37˚C in the 4:1 molar ratio aprotinin to pro- KLK4-V5-His or KLK4-V5-His and 1 μg of untreated and aprotinin treated pro- KLK4-V5-His and KLK4-V5-His analysed by SDS-PAGE on 10% gel run under non-reducing conditions with protein visualisation with a commassie stain. Aprotinin is able to form a complex with KLK4-V5-His (lane 4, upper band), but not with pro- KLK4-V5-His (lane 2). The displayed gel is representative of duplicate experiments.

Aprotinin inhibits several serine proteases including trypsin and plasmin (Hewlett, 1990). Further, aprotinin has been used to inhibit the catalytic activity of bacterial recombinant KLK4 in in vitro experiments (Takayama et al., 2001b), and was assessed as an inhibitor of insect produced recombinant KLK4 activity against the F- V-R-pNA tri-peptide in this study (Figure 3.10A). The inhibition of KLK4-V5-His trypsin-like activity by aprotinin was assessed by incubating enzyme and inhibitor in a 20:1 molar ratio aprotinin to mature KLK4 (30 nM; based on MUGB active site

67 titration results) for 5 minutes at 37˚C before reaction velocities were determined against 750 μM F-V-R-pNA in comparison to 30nM uninhibited KLK4-V5-His. Setting the velocity recorded for uninhibited KLK4-V5-His at 100% activity, aprotinin caused a ~85% inhibition of KLK4-V5-His activity when present in a 20 fold molar excess (Figure 3.10B). The residual KLK4-V5-His activity observed after inhibition could be attributable to spontaneous hydrolysis of the tri-peptide, as seen in the mutant KLK4-V5-His and vehicle control activity assays in the concentration range of 32.5 μM, 75 μM, 125 μM, 250 μM, 500 μM and 750 μM of the peptide F- V-R-pNA (Figure 3.8B).

Figure 3.10 Inhibition of KLK4-V5-His activity by aprotinin. A. Schematic depicting lack of catalytic activity of KLK4-V5-His due to its proteolytic inhibition by aprotinin resulting in no pNA release from the tri-peptide F-V-R-pNA for spectrophotometric measurement at 405 nm. B. Activity levels due to pNA release from cleavage of 750 μM FVR-pNA tri-peptide by 30 nM KLK4-V5-His without and with aprotinin inhibition in a 20:1 molar ratio aprotinin to mature KLK4. The velocity obtained for uninhibited KLK4-V5-His was set at 100% activity and aprotinin inhibited KLK4-V5-His velocity displayed relative to this value. Data is represented by the mean of triplicate experiments +/- SEM.

68 3.4 Discussion

Previous work in our laboratory had generated purified recombinant pro-KLK4-V5- His (Figure 3.1). Analysis of this protein as a part of this PhD program of research, revealed a molecular weight of 35 kDa, significantly larger than the predicted 29.3 kDa (KLK4 =27.0, V5/His= 2.3 kDa) for the zymogen enzyme, due to post translational modifications (Figure 3.1). Part of the additional molecular weight of the zymogen KLK4 protein is attributable to the denatured state of the protein when run under reducing conditions. Under non-reducing conditions zymogen KLK4 resolved to a size of 31 kDa (Figure 3.9; lane 1), due likely to the compact nature of the protein when the disulphide bridges of the molecule remain intact. Bacterial KLK4 showed a similar 4 kDa difference between folded (non-reduced; 21 kDa) and unfolded (reduced; 25 kDa) (Takayama et al., 2001b). Glyosylation most likely represents the remainder of the additional size, as when PNGase F removed carbohydrate moieties, presumably from the putative N-linked glycosylation site residue asparagine169, the protein reduced in size to 34 kDa (Figure 3.2). A method which in future studies could greatly improve the analysis of the different post- translational KLK4 isoforms is mass spectrometry. Mass spectrometry enables the exact covalent composistion of a protein to be determined and has been used extensively to characterise protein post-translational modifications (Guerrera and Kleiner, 2005). Utilising mass spectrometry may potentially identify KLK4 isoform(s) which have enhanced enzymatic activity.

Glycosylation is the most common post-translational modification and is essential for protein folding, stability, degradation, trafficking, secretion and plasma membrane localisation (Helenius and Aebi, 2004). Interestingly, in work from our laboratory, a translated exon 1 deleted isoform of KLK4 lacking the signal peptide and pro-region, localised to the nucleus of the LNCaP prostate cancer cell line (Dong et al., 2005). Consistent with this observation, this KLK4 isoform lacks N-linked glycosylation at the putative asparagine120 residue.

In eukaryotes protein secretion is linked to protein glycosylation within the endoplasmic reticulum and the Golgi apparatus (Helenius and Aebi, 2004). Upon emergence from the large ribosomal subunit during translation, the signal peptide

69 directs contact of the ribosomal complex with the membrane of the ER through binding with the signal recognition particle (Walter et al., 1981) and subsequent complex formation with the docking protein (Meyer et al., 1982). The nascent protein is then co-translationally translocated across the membrane of the ER and the signal peptide removed by signal peptidases (Kreil et al., 1980). Accordingly, KLK4 lacking a signal peptide would accumulate intracellularly, which was consistent with the nuclear localisation of the exon 1 deleted isoform of KLK4 (Dong et al., 2005).

In several genetic disorders, including Alzheimer’s, Parkinson’s, Huntington’s and Type 1 diabetes, aberrant intracellular accumulation of mutant proteins has been reported (Dobson, 1999; Horwich, 2002). In a study by Klokk and colleagues, an exon 1 deleted KLK4 cDNA, which they mistakenly referred to as full-length KLK4, was over-expressed in PC-3 and DU145 prostate cancer cell lines, accumulating within the nucleus (Klokk et al., 2007). Over-expression of the truncated KLK4 increased the proliferation, colony formation and motility of each cell line. Consistently, several proliferative factors were up regulated, including cyclin B1, E2F1, PCNA and Ki67, co-comitant with a decrease in the transcription of cycle cell inhibitory genes, including; p21, p15 and p16 (Klokk et al., 2007). Although the intracellular exon 1 deleted KLK4 isoenzyme has functional implications for prostate cancer development and progression, full length secreted KLK4 remains a central enzyme in KLK studies due to its potential in extracellular events.

Classical serine protease function involves extracellular proteolysis to modulate a variety of cellular processes by selective cleavage of specific substrates to influence cell behaviour (Turk, 2006). Well known physiological examples include the proteases of the blood coagulation (eg. thrombin) (Stubbs and Bode, 1993) digestive (trypsin) (Yamashina, 1956) and wound healing (plasmin) (Romer et al., 1996) cascades. Consistent with an extracellular physiological role, the secretion of the glycosylated full-length KLK4 protein is detected in seminal fluid (Dong et al., 2005). Recombinant KLK4 from insect cells is secreted as a catalytically inert zymogen with the active site of the enzyme sequestered intramolecularly, demonstrated in this study by a lack of interaction between the recombinant zymogen enzyme and the active site inhibitor aprotinin (Figure 3.9; lane 2).

70 The in vivo conversion mechanism of KLK4 zymogen to the catalytically active mature enzyme remains unknown. In contrast, it is known that the six amino acid pro-region of trypinsogen, zymogen of the structurally similar S1 family member trypsin, is removed by limited proteolysis by the type II transmembrane serine protease (Hooper et al., 2001) enterokinase (Kitamoto et al., 1994), resulting in formation of the active enzyme (Yamashina, 1956). However, unlike trypsin and many other serine proteases which have a pro-region ending in a lysine or arginine, the pro-region of KLK4 ends in a glutamine (Nelson et al., 1999). Thus, pro-KLK4 cannot be activated by other serine proteases with trypsin-like specificity or by auto- activation. The carboxyl residue of the scissile bond cleaved for pro-region removal is an isoleucine (Nelson et al., 1999). This may suggest removal of the pro-region of KLK4 by a metalloprotease, due to the preference of these enzymes for hydrophobic resides in the P1’ site (Turk et al., 2001). Indeed, removal of the pro-region of recombinant porcine KLK4 zymogen, in which the P1’ isoleucine is present, was achieved by cleavage with the metalloendopeptidase thermolysin (Ryu et al., 2002) and was successfully applied in this study to convert zymogen KLK4 to mature KLK4 (Figure 3.3).

A crystal structure study on KLK4 has shown that removal of the pro-region creates a conformational change within the protein, facilitating the formation of the oxyanion substrate binding site (Debela et al., 2006a). Stability of the newly formed active site is achieved through the formation of an internal salt bridge between the nascent N-terminal residue isoleucine31 (designated isoleucine16 in Figure 3.11; due to alternate residue numbering as a result of a substituted trypsinogen pro-region) and asparagine194 (Figure 3.11; (Debela et al., 2006a); a mechanism in common with other trypsin-like serine proteases (Page and Di Cera, 2008). In this study, the observation of complex formation between mature KLK4 and aprotinin indicated that the active site had been correctly formed following activation with thermolysin (Figure 3.9; lane 4). Additionally, a catalytically active KLK4 following thermolysin treatment was demonstrated by the release of 4-methylumbelliferone from MUGB (Figure 3.4B).

The catalytic efficiency of mature KLK4 in this study, as assessed by kinetic parameters derived from cleavage of a tri-peptide substrate (FVR-pNA) (Km=47 +/-

71 -1 -1 6.3 μM, kcat=1.8 +/- 0.12 s , kcat/ Km=38 +/-1.9 (s.mM) ), was approximately 1000 -1 fold less than that observed for trypsin (Km=74 +/- 3.4 μM, kcat=248 +/- 1.6 s , kcat/ -1 Km=3351 +/- 470 (s. mM) ) (Figure 3.6). Unlike trypsin, which cleaves indiscriminately at any accessible lysine or arginine residue (Hinman et al., 1976), KLK4 requires specific residues flanking the scissile peptide bond (Takayama et al., 2001b; Matsumura et al., 2005).

A structural feature that imparts specificity between various serine proteases is the shape and the width of the substrate pocket (Perona and Craik, 1995). The crystal structure of KLK4 highlights a number of structural determinates around the active site cleft that would result in narrow substrate specificity, including a negatively charged region that spans an identical area to that of the thrombin positively charged anion binding exosite, a region known to regulate substrate specificity, a notched negatively charged S2 pocket formed by the 99 loop active site “roof”, and a rigid active site floor, formed by 148 loop of KLK4 (Figure 3.11; (Debela et al., 2006a).

Figure 3.11 Crystal structure of KLK4. Ribbon plot of KLK4 shown in standard orientation with loops 37, 70-80, 99 and 148 in orange. The active site residues histidine57, aspartate102 and serine195, the nickel binding residues histidine25 and glutamic acid77, the salt bridge forming residues isoleucine16 and asparagine194, as well as the inhibitor PABA and aspartate189, both located in the S1 pocket, are shown as stick models (Taken from (Debela et al., 2006a).

The derived enzyme kinetics of mature KLK4 in this study agreed well with the findings of Matsumura et al (Km = 48.3 μM, kcat = 1.28 s-1 and kcat/Km = 38.3

72 (s.mM)-1) using the same substrate and Drosophila melanogaster S2 cell expressed recombinant pro-KLK4 that contained a synthetic trypsinogen pro-region that was activated by enterokinase (Matsumura et al., 2005). Interestingly, mature KLK4 generated in Escherichia coli cells exhibited greater catalytic activity towards the identical F-V-R-pNA peptide substrate (Km = 20.7 μM, kcat = 3.64 s-1 and kcat/Km = 175.7 (s.mM)-1) in experiments by Debela and colleagues (Debela et al., 2006a). This protein, while containing an identical peptide backbone, would lack N-linked glycosylation, a post-translational modification bacteria lack due to the absence of eukaryotic glycosylation machinery – the ER and golgi apparatus (Brooks, 2004).

The modulation of serine protease activity due to differences in N-glycosylation is exemplified by proteases of the coagulation cascade. For example, the rate of fibrin dependent activation of plasminogen by tissue plasminogen activator (tPA) depends on the occupancy of the glycosylation sites in Kringle 2 in tPA and on Kringle 3 in plasminogen. The combined effect of glycoforms of both the tPA and plasminogen molecules results in a 4-fold range of activities (Kaufman, 1998). Furthermore, the amidolytic and anticoagulant activites of human activated protein C are significantly increased by the elimination of glycosylation sites on the heavy chain of this protease (Grinnell et al., 1991). In addition, the activity of mouse KLK8 is dependent upon N- linked glycosylation of the kallikrein loop. Deletion of this glycosylation site leads to an alteration in substrate specificity (Oka et al., 2002). Collectively these data support the hypothesis, that glycosylation may be an important post-translational control for mature KLK4 enzyme activity. Furthermore, use of bacterial derived KLK4, may not comprehensively reflect the physiological specificities and interactions of this glycoprotein.

Another condition that is critical for enzyme activity is pH. A pH range, in which optimal activity occurs, reflects the physiological environment the enzyme originates from. For example a pH of 2.0 is optimal for the digestive enzyme pepsin, isolated from the stomach (Sri Ram and Maurer, 1957). KLK4 is predominately found in the ejaculate, which has a physiological pH of 7.5 to 8.5 (Makler, 2003). This study has shown an optimal pH of 8.0 for mature KLK4 with high levels of activity between pH 6.0 and 10.0 (Figure 3.5), consistent with a physiological role in seminal fluid. Also sharing a pH optimum of approximately 8.0 is PSA, one of the most abundant

73 proteins in the secretion from human prostate epithelium cells into seminal fluid (Christensson et al., 1990).

The known in vivo physiological function of PSA is to degrade the seminal vesicle proteins seminogelins I and II for liquidification of the seminal clot (Lilja, 1985). Like PSA, KLK4 is known to degrade a component of the seminal fluid, prostatic acid phosphatase (PAP), although the consequences of this interaction are unclear (Takayama et al., 2001b). In vitro protein substrate studies suggest physiological roles for KLK4 not just in seminal fluid. Degradation of the extracellular matrix (ECM) components type I and type IV collagen are indicative of involvement in ECM remodelling (Obiezu et al., 2006) and consistent with KLK4 homology to EMSP-1, a protease that degrades the extracellular matrix surrounding enamel crystallites, a critical step in tooth development (Hu et al., 2000). Furthermore, in vitro KLK4 cleaves pro-PSA (Takayama et al., 2001a), fibrinogen (Obiezu et al., 2006), pro-urokinase type plasminogen activator (uPA) (Beaufort et al., 2006) and the uPA receptor (Beaufort et al., 2006), illustrating the potential for KLK4 to influence a variety of cellular behaviours. Identification of relevant in vivo substrates of KLK4 is critical to understanding the patho-physiological roles of this enzyme. Members of the PAR family have great potential as KLK4 substrates, due to their almost exclusive activation by serine proteases with trypsin-like specificity. This hypothesis will be explained and tested in subsequent chapters.

In summary, this chapter shows the successful generation and purification of catalytically inert pro-KLK4 from an insect cell expression system. Removal of the pro-region by thermolysin at the correct activation site generates a catalytically active mature enzyme with similar kinetic parameters to a previously published insect cell expressed recombinant KLK4 (Matsumura et al., 2005). The activity of the glycosylated KLK4 utilised in this study is significantly less than that of a published bacterial derived non-glycosylated KLK4 (Debela et al., 2006a), highlighting the influence of glycosylation for the catalytic activity of KLK4. This illustrates the importance of using glycosylated enzymes in in vitro studies to better reflect their activity/specificity in vivo. Furthermore, mature KLK4 shared an optimal pH of 8.0, similar to that of the prostatic-derived seminal fluid enzyme, PSA.

74

CHAPTER 4

ANALYSIS OF PROTEASE ACTIVATED RECEPTOR

ACTIVATION BY KALLIKREIN-RELATED PEPTIDASE 4

75 4.1 Introduction

As described in Chapter 1 KLK4 is associated with prostate cancer. However, the mechanism(s) by which KLK4 mediates its effects on cells are not known. Work described in this chapter examines the ability of this protease to initiate cell signaling via PARs.

Recently, several members of the KLK family have been shown to initiate trans- plasma membrane signal transduction via PARs. Oikonomopoulou and co-workers demonstrated that KLK14 activates PAR-2 and PAR-4, but inactivates (or “disarms”) PAR-1 (Oikonomopoulou et al., 2006). This group also showed that KLK5 and KLK6 activate PAR-2 (Oikonomopoulou et al., 2006). Consistent with this finding, Angelo et al have shown that KLK6 is capable of cleaving a peptide spanning the PAR-2 activation site but not peptides spanning the activation site of the other PARs (Angelo et al., 2006). KLK5 and KLK14 signaling via PAR-2 has been demonstrated independently in a study that also showed that KLK7 and KLK8 are not capable of signaling through this receptor (Stefansson et al., 2008). As reviewed in Chapter 1, two of the main signaling events initiated by PAR activation are cytosolic release of the second messenger Ca2+ and activation of the ERK signaling cascade (Macfarlane et al., 2001). Accordingly, both of these signaling events were assessed in this study to determine if KLK4 is able to activate PAR family members.

Changes in intracellular concentrations of free Ca2+ in response to cell surface receptor activation by neurotransmitters, hormones and other molecular messengers, constitutes one of the major events of intracellular communication (Clapham, 2007). Stimulation of GPCRs activates phospholipase C (PLC), catalysing the breakdown of phosphatidylinositols 4, 5-bisphosphate (PIP2) into the second messenger molecules inositol 1, 4, 5-triphosphate (IP3) and diacylglycerol (DAG). IP3 mediates the rapid 2+ release of stored Ca by activating IP3 receptors in the endoplasmic reticulum (ER), while DAG activates protein kinase C (PKC) and generates a host of bioactive molecules (Figure 1.5, grey filled) (Clapham, 2007). Transient increases in cytsolic calcium can be monitored with the fluorescent calcium indicator FURA-2 acetoxymethyl (AM) (O'Connor and Silver, 2007). FURA-2 AM enters the cells as the uncharged ester which is hydrolysed intracellularly by esterases to the free

76 carboxylate, which in turn sequesters calcium and undergoes an excitation optimum shift between 380 nm (calcium free) and 340 nm (calcium bound). Monitoring the emission at 510 nm allows detection of the excitation shift. The more calcium present in the cytoplasm the higher the absorption shift, towards 340 nm from 380 nm, when monitored at 510 nm. Utilising FURA-2 AM in this study will enable examination of transient releases of intracellular calcium in response to PAR activation by KLK4.

Although PARs are able to strongly stimulate polyphosphoinositide hydrolysis and the activation of PKC, this is insufficient to initiate the various cellular effects elicited by agonists, such as thrombin (McKenzie et al., 1992). This implicates activation of other intracellular signaling cascades to mediate the variety of cellular changes that occur following PAR activation. One of the well described intracellular cascades following PAR activation is the ERK cascade. Activation of the highly similar ERK 1 and 2 isoforms requires phosphorylation on both tyrosine and threonine residues within a specific TEY motif (Anderson et al., 1990), by members of the ERK kinase family (Nakielny et al., 1992). ERK kinase family activation is in turn regulated by serine phosphorylation by members of the RAF family of proto- oncogene products (Figure 1.5, black filled) (Kyriakis et al., 1992). In this study, by using Western blot analysis with antibodies that specifically recognise the phosphorylated ERK isoforms, initiation of the ERK cascade will be assessed in response to PAR activation by KLK4.

Described in this chapter are experiments examining intracellular Ca2+ concentration in response to KLK4 activation of PAR-1, -2 or -4. Due to the results obtained from these experiments, the remainder of this chapter focussed on PAR-2. The ability of KLK4 to activate ERK 1/2 and the specificity of PAR-2 activation by KLK4, using siRNA knock-down and synthetic N-terminal cleavage approaches is demonstrated.

4.2 Materials and Methods

These are comprehensively described in Chapter 2. The reagents used specifically in experiments performed in this chapter are described below.

77 The lung myofibroblast cell line derived from PAR-1 -/- mice (N1LF), the PAR-1- lung myofibroblast PAR-1 -/- (LMF), PAR-2-LMF or PAR-4-LMF cell lines used for signaling assays were a generous gift from Dr. Andrade-Gordon (R. W. Johnson Pharmaceutical Research Institute, Pennsylvania). The N1LF cells line was generated by immortalisation of lung myofibroblasts cells derived from PAR-1-deficient mice, created by homologous recombination to disrupt the PAR-1 gene (Darrow et al., 1996). PAR-1-LMF, PAR-2-LMF and PAR-4-LMF cells were generated by stably transfecting N1LF cells with expression constructs encoding either human PAR-1, PAR-2, or PAR-4 (Andrade-Gordon et al., 1999). These cell lines, due to their lack of murine Par-1, Par-2, Par-3 or Par-4 and their stable reconstitution with a single human PAR (PAR-1, PAR-2 or PAR-4), provide a valuable cell model to study activation of each PAR in isolation, eliminating complex multi-PAR activation. Cells stably expressing PAR-3 are not available and this receptor in mouse does not mediate transmembrane signaling leading to the proposal that PAR-3 acts as a co- receptor for signaling via PAR-4 (Nakanishi-Matsui et al., 2000).

The mammalian siRNA expression vector pSilencer 3.1-H1 puro was used to reduce expression of PAR-2 in the PAR-2-LMF cell line. Three siRNA sequences; 5- GATCCAGGAAGAAGCCTTATTGGTTTCAAGAGAACCAATAAGGCTTCTTC CTTTTTTTGGAAA-3; 5-GATCCAGTAGACTTGGTGTGAAGATTCAAGAGAT CTTCACACCAAGTCTACTTTTTTTGGAAA-3; and 5-GATCCGTAGTCGTGAA TCTTGTTCATTCAAGAGATGAACAAGATTCACGACTATTTTTTGGAAA-3), were inserted into the pSilencer 3.1-H1 puro vector according to the instructions of the manufacturer. PAR-2-LMF cells were transfected with the PAR-2 pSilencer 3.1- H1 puro constructs or the supplied pSilencer 3.1-H1 puro negative control and selected for stable incorporation of each construct for 2 weeks with 2 μg/mL of puromycin.

4.3 Results

4.3.1 Cytosolic calcium release in response to PAR activation by KLK4

The ability of the arginine/lysine specific serine protease KLK4 to initiate intracellular signaling through PAR activation has not previously been examined. In

78 the experiments described here the ability of recombinant active KLK4 (described in Chapter 3 and referred to as KLK4-V5-His) to induce changes in intracellular [Ca2+] in PAR-1-LMF, PAR-2-LMF and PAR-4-LMF cells was assessed by Ca2+ flux assays.

4.3.1.1 Determination of PAR-1 activation by KLK4

As shown in Figures 4.1A, in the PAR-1-LMF cell line active KLK4 (300 nM) initiated a prompt transient Ca2+ mobilisation through PAR-1 activation. Ca2+ flux was also induced by the positive controls for PAR-1 (10 nM thrombin (Figure 4.1B) and 100 μM PAR-1 AP (Figure 4.1C)). Treatment of the PAR-1-LMF cells with 100 μM PAR-2 AP resulted in no calcium mobilisation, confirming that only PAR-1 is present in the cells and the calcium mobilisation by KLK4 is due exclusively to PAR-1 activation (Figure 4.1D). In other experiments, 1 μM KLK4 zymogen (equivalent to amount used to generate 300 nM active KLK4) did not initiate PAR-1 signaling (Figure 4.1E), confirming that activation was not due to a contaminating protease carried over from the insect cell media. Further, the vehicle control (phosphoramidon (4 μM) inhibited thermolysin (4.6 nM)-equivalent to amounts used to generate 1 μM active KLK4) also failed to mobilise Ca2+, eliminating the possibility of thermolysin activation (Figure 4.1F).

4.3.1.2 Determination of PAR-2 activation by KLK4

In addition to PAR-1 activation, active KLK4 (300 nM) was also able to initiate prompt transient Ca2+ mobilisation through PAR-2 activation (Figure 4.2A). Interestingly, the magnitude of the Ca2+ release, defined as efficacy (Kenakin, 2005), by KLK4 activation was higher through PAR-2 than PAR-1 (compare peak height of Figure 4.2A to Figure 4.1A). The positive controls for PAR-2 activation (10 nM 2+ trypsin (Figure 4.2B) and 100 μM PAR-2 AP (Figure 4.2C) also mediated Ca mobilisation, while no mobilisation resulted in response to 100 μM PAR-1 AP confirming that only PAR-2 is present in the PAR-2-LMF cells, and the calcium mobilisation by KLK4 is due exclusively to PAR-2 activation (Figure 4.2D). Further, 1 μM KLK4 zymogen (pro-KLK4-equivalent to the amount used to generate 300 nM active KLK4) (Figure 4.2E) and vehicle control (phosphoramidon (4 μM)

79 inhibited thermolysin (4.6 nM)-equivalent to amounts used to generate 1 μM active

KLK4) (Figure 4.2F) did not initiate PAR-2 signaling.

Figure 4.1 PAR-1 mediated calcium mobilisation. PAR-1-LMF cells were incubated with either (A) active KLK4 (300 nM), (B) thrombin (10 nM), (C) PAR-1 AP (100 μM), (D) PAR-2 AP (100 μM), (E) KLK4 zymogen (1 μM - equivalent to amount used to generate 300 nM active KLK4) or (F) phosphoramidon (4 μM) inhibited thermolysin (4.6 nM) (equivalent to amounts used to generate 1 μM active KLK4) as indicated and fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm (Em510(340/380)) using a fluorescent plate reader. The ratio of Em510(340/380) is proportional to intracellular Ca2+ ion concentration. Arrow indicates time of treatment. Data are representative of experiments performed in triplicate and repeated 3 times.

4.3.1.3 Determination of PAR-4 activation by KLK4

In contrast to PAR-1 and PAR-2, no calcium mobilisation was observed in cells expressing PAR-4 in response to 300 nM active KLK4 (Figure 4.3A). This lack of Ca2+ flux in the PAR-4-LMF cell line serves to further support the observation from

80 Figures 4.1 and 4.2 that KLK4 specifically signals via PAR-1 and PAR-2. Positive control agonists for PAR-4 (10 nM trypsin (Figure 4.3B) and 500 μM PAR-4 AP (Figure 4.3D)) each induced Ca2+ mobilisation, demonstrating the presence of functional PAR-4, while an absence of PAR-2 was confirmed by 100 μM PAR-2 AP eliciting no response (Figure 4.3C).

Figure 4.2 PAR-2 mediated calcium mobilisation. PAR-2-LMF cells were incubated with either (A) active KLK4 (300 nM), (B) trypsin (10 nM), (C) PAR-2 AP (100 μM), (D) PAR-1 AP (100 μM), (E) KLK4 zymogen (1 μM - equivalent to amount used to generate 300 nM active KLK4) or phosphoramidon (4 μM) inhibited thermolysin (4.6 nM) (equivalent to amounts used to generate 1 μM active KLK4) as indicated and fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm (Em510(340/380)) using a fluorescent plate reader. The ratio of Em510(340/380) is proportional to intracellular Ca2+ ion concentration. Arrow indicates time of treatment. Data are representative of experiments performed in triplicate and repeated 3 times.

81

Figure 4.3 PAR-4 mediated calcium mobilisation. PAR-4-LMF cells were incubated with either (A) active KLK4 (300 nM), (B) trypsin (10 nM), (C) PAR-2 AP (100 μM) or (D) PAR-4 AP (500 μM) as indicated and fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm (Em510(340/380)) using a fluorescent plate reader. The ratio of Em510(340/380) is proportional to intracellular Ca2+ ion concentration. Arrow indicates time of treatment. Data are representative of experiments performed in triplicate and repeated 3 times.

4.3.1.4 Characterisation of KLK4 mediated calcium mobilisation through PAR- 1 and PAR-2

Having established KLK4 as an activator of PAR-1 and PAR-2, the difference in KLK4 efficacy of calcium mobilisation through PAR-1 and PAR-2, suggested by differences in flux magnitudes in Figures 4.2A and 4.1A, was examined further. Figures 4.4A and 4.4B illustrate the inability of 300 nM active KLK4 to completely activate all surface PAR-1 and PAR-2, respectively, as a second treatment with the appropriate AP induced increases in cytosolic calcium in the PAR-1-LMF and PAR-

82 2-LMF cell lines. Treatment with 300 nM active KLK4 was unable to elicit even minimal activation, illustrated by full Ca2+ mobilisation when cells were exposed to 500 μM PAR-4 AP 3 minutes after 300 nM KLK4 exposure (Figure 4.4C). Due to limited amounts of recombinant KLK4, an appropriate concentration range to further study PAR-4 activation was not possible and PAR-4 was not examined further.

Figure 4.4 Functional PAR-1,-2 or -4 remaining after KLK4 treatment. (A) PAR-1-LMF, (B) PAR-2-LMF or (C) PAR-4-LMF cells were incubated with active KLK4 (300 nM; ) for a 3 minute period before a second treatment with either PAR- 1 AP (100 μM; ), PAR-2 AP (100 μM; ) or PAR-4 AP (500 μM; ) as indicated. Fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm (Em510(340/380)) using a fluorescent plate reader. The ratio of Em510(340/380) is proportional to intracellular Ca2+ ion concentration. Arrow indicates time of treatments. Data are representative of experiments performed in triplicate, repeated 3 times.

83 In order to determine saturating KLK4 concentrations for PAR-1 and PAR-2 mediated calcium signaling, a concentration response experiment was performed by incubating PAR-1-LMF and PAR-2-LMF cells with increasing concentrations of enzyme (0.1nM to 1 μM) and examining changes in Ca2+ mobilisation. As shown in Figure 4.5A, increasing concentrations of active KLK4 stimulated increased Ca2+ mobilisation via both PARs. The maximum response via PAR-1 was achieved at ~300 nM of active KLK4 with a concentration to stimulate half-maximal response calculated as pEC50 (-logEC50) = 7.49 +/- 0.34 from a nonlinear regression curve using Graphpad Prism 4 (Figure 4.5A; left panel). In contrast, a maximum concentration response via PAR-2 was not achieved at the maximum concentration of active KLK4 currently available to us. However, using the concentration response achieved at 1 μM of active KLK4 as the maximum response, a concentration to stimulate half-maximal response was calculated as pEC50 = 6.53 +/- 0.17 (Figure 4.5A; right panel), which demonstrates that KLK4 has higher potency for PAR-1 than PAR-2.

To compare the efficacy of KLK4 initiated Ca2+ mobilisation via PAR-1 and PAR-2 expressing cells, fluorescence values obtained at each concentration of active KLK4 were divided by the highest response observed for each of these PARs; which was via PAR-2 at 1μM active KLK4 (marked by in Figure 4.5B). Graphical comparison in Figure 4.5B shows that whereas the response to KLK4 reached a maximum at ~300 nM ( )) in PAR-1-LMF cells, the response in PAR-2-LMF cells increased over the entire KLK4 concentration range, showing that the efficacy of active KLK4 mediated Ca2+ mobilisation is higher through PAR-2 than PAR-1. Due to limited amounts of recombinant KLK4 a saturating concentration for PAR-2 activation was unable to be obtained.

4.3.2 Cleavage of the PAR-2 N-terminus by KLK4

As concentration-effect curves indicated that KLK4 initiated Ca2+ mobilisation displayed higher efficacy via PAR-2 than via PAR-1 (Figure 4.5B), KLK4 activation of PAR-2 was analysed further. A fluorescence quenched peptide spanning the PAR-

84 2 activation site (Abz-SKGR↓SLIGK-Dnp; ↓ marks the activation site), which is efficiently activated by low concentrations of trypsin, tryptase and acrosin but not by high concentrations of thrombin (Smith et al., 2000), was used to examine the kinetics of direct PAR-2 cleavage by active KLK4 (40 nM).

Figure 4.5 KLK4 concentration responses of PAR-1 and PAR-2 activation and effect on calcium mobilisation. A. PAR-1-LMF and PAR-2-LMF cells were incubated with active KLK4 (0.1 nM to 1 μM) and fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm (Em510(340/380)) using a fluorescent plate reader. [Ca2+] is displayed graphically as a percentage of the highest recorded response. Experiments were performed in triplicate and repeated 3 times. B. Displays a graphical comparison of responses of PAR-1-LMF and PAR-2- LMF cells to active KLK4 ( marks the saturation concentration of active KLK4 PAR-1 mediated calcium release and the marks incomplete saturation of PAR-2 mediated calcium release by active KLK4). Data is the mean value of triplicate experiments +/- SEM.

85 Cleavage of the scissile peptide bond following the arginine residue within the fluorescence quenched substrate leads to the separation of the intramolecular donor- acceptor pair and thus, an increase in the fluorescence (Yaron et al., 1979). Consequently, the fluorescence increase is proportional to the amount of peptide hydrolysed allowing quantitation of enzyme kinetics. Identical enzyme concentrations as used in F-V-R-pNA tri-peptide kinetic assays in chapter 3, 40 nM active KLK4 or 10 nM trypsin, were incubated with varying concentrations (0 μM, 0.5 μM, 2.5 μM, 5 μM, 10 μM, 25 μM, 50 μM and 100 μM) of Abz-SKGRSLIGK- Dnp substrate and fluorescence monitored over time with initial velocities fitted to the Michaelis-Menten equation to determine kinetic parameters. KLK4 was able to efficiently cleave the PAR-2 peptide with kinetic parameters of Km=8.0 +/- 0.7 μM, kcat = 7.9 +/- 0.3 s-1 and kcat/Km = 1.0 +/- 0.4 (s.μM)-1 (Figure 4.6A) compared to trypsin with the kinetic parameters of Km=14 +/- 3.4 μM, kcat = 226 +/- 2.3 s-1 and kcat/Km = 16 +/- 0.7(s.μM)-1 (Figure 4.6B).

4.3.3 ERK phosphorylation in response to PAR-2 activation by KLK4

As discussed in the introduction ERK phosphorylation represents another intracellular pathway downstream of PAR activation. The ability of KLK4 to signal via PAR-2 was further examined by analysing ERK1/2 activation. We performed SDS-PAGE on 12% gels run under reducing conditions, followed by Western blot analysis to detect phosphorylated ERK1/2 (present as 44 kDa and 42 kDa proteins), in PAR-2-LMF cells stimulated with 40 nM active KLK4 using total ERK1/2 as a control for relative expression of the kinase. Treatments with 10 nM trypsin and 100 μM PAR-2 AP were used as positive controls for ERK1/2 activation via PAR-2. As shown in representative blots in Figure 4.7A, KLK4 induced ERK1/2 phosphorylation within 5 minutes with loss of activation apparent at 15 minutes. PAR-2 AP and trypsin induced ERK1/2 phosphorylation in a similar time dependent manner (Figures 4.7B and 4.7C respectively).

86

Figure 4.6 Characterisation of KLK4 cleavage of a peptide spanning the PAR-2 activation site. The Lineweaver-Burke plots of (A) active KLK4 (40 nM) and (B) trypsin (10nM) derived from the initial velocities of the enzyme at increasing concentrations of the PAR-2 peptide SKGR↓SLIGK (“↓” indicates activation site). Kinetic constants calculated from the graphs are displayed beside the respective plot. Initial velocities represent the means +/- SEM of triplicate experiments repeated 3 times (note that the SEM for KLK4 velocities are not visible due to small error values).

87

Figure 4.7 Time dependent phosphorylation of ERK1/2 in response to PAR-2 activation. PAR-2-LMF cells were incubated with either (A) active KLK4 (40 nM), (B) PAR-2 AP (100 μM) or (C) trypsin (10 nM) and cell lysates collected at the indicated times were analysed by SDS-PAGE on a 12% gel run under reducing conditions followed by Western blot analysis using anti-ERK and anti-phospho-ERK (p-ERK) antibodies. Each agonist experiment was repeated 3 times with displayed blots representative of all experiments.

The concentration response of active KLK4 initiated signaling via PAR-2 was examined by incubating PAR-2-LMF cells with increasing concentrations of enzyme (0.1 to 300 nM) and examining changes in ERK1/2 activation at 5 minutes. Consistent with Ca2+ mobilisation assays (Figure 4.5), increasing concentrations of active KLK4 stimulated increased ERK1/2 activation following PAR-2 activation (Figure 4.8). Interestingly, unlike Ca2+ mobilisation assays in which a saturating concentration of KLK4 for PAR-2 activation was unable to be determined in a concentration response ranging from 0.1nM to 1 μM, a maximal plateau in ERK

88 phosphorylation was reached at approximately 100 nM active KLK4, with a concentration to stimulate half-maximal response calculated from a nonlinear regression curve using graphpad prism 4 as pEC50 = 7.67 +/- 0.24.

Figure 4.8 KLK4 concentration response of PAR-2 activation and effect on phosphorylation of ERK1/2. Concentration dependence of active KLK4 on ERK1/2 activation. Lysates from PAR-2-LMF cells incubated with active KLK4 (0.1 to 300 nM) for 5 minutes were examined by anti-phospo-ERK (p-ERK) and anti-ERK Western blot analysis following SDS-PAGE on 12% gels run under reducing conditions. The ratio of p-ERK to total ERK was determined by densitometry and is displayed graphically. The data represents mean values of duplicate experiments +/- SEM.

89 4.3.4 Depletion of PAR-2 by siRNA and consequences for signal transduction

RNA interference represents a powerful tool for inhibition of gene expression in exploring protein function (Elbashir et al., 2001). Introduction of siRNA duplexes, or small-hairpin RNAs, targeting PAR-1 and PAR-2 have been effectively used to reduce cellular abundance of these two proteins in studies examining receptor activation mechanisms (Morris et al., 2006; Salah et al., 2007; Salah et al., 2007). We have therefore, utilised siRNA mediated PAR-2 protein depletion in this study to more closely examine the specificity of the signal transduction elicited by KLK4 through PAR-2.

Three PAR-2 siRNA knockdown constructs spanning nucleotides 254-274 (PAR-2 siRNA construct #1), 1635-1655 (PAR-2 siRNA construct #2) and 2190-2210 (PAR- 2 siRNA construct #3) were generated in the Ambion pSilencer 3.1-H1 puro vector. Lysates from PAR-2-LMF cells stably transfected with these constructs were analysed by Western blot analysis using an anti-PAR-2 antibody following SDS- PAGE on a 12% gel run under reducing conditions. As shown in Figure 4.9A it was apparent that construct #2 effectively reduced PAR-2 levels in comparison to a non- specific control with no homology to known human, mouse or rat genes. Densitometric analysis of PAR-2 levels in comparison to the loading control GAPDH indicated that construct #2 significantly reduced (p<0.01) PAR-2 levels by an average of 60% in Western blot analysis of lysates extracted on three separate occasions in comparison to the non-specific control construct (set as 100%), while siRNA constructs #1 and #3 failed to consistently reduce PAR-2 expression levels (Figure 4.9B).

90

Figure 4.9 siRNA depletion of PAR-2 protein in PAR-2-LMF cells. PAR-2-LMF cells were stably transfected with one of 3 PAR-2 siRNA constructs (#1, #2 or #3) or a scrambled sequence siRNA control construct (NS, non-specific siRNA). A. Cell lysates were analysed by Western blot analysis probing with either an anti-PAR-2 (SAM-11) or anti-GAPDH antibody following SDS-PAGE on 12% gels run under reducing conditions. B. The level of PAR-2 expression for each of the siRNA constructs relative to GAPDH was determined by densitometry and displayed graphically, with the non-specific siRNA PAR-2 levels set as 100% and the 3 PAR-2 siRNA as relative values. Data represented is the mean of triplicate experiments +/- SEM. Statistical significance (p<0.01) was determined with student’s t-test with a significance threshold set at p<0.05.

The effect of PAR-2 knock-down on KLK4 initiated intracellular signaling was examined by assessing Ca2+ mobilisation and levels of activated ERK1/2 in PAR-2- LMF cells stably transfected with siRNA construct #2 and the non-specific control. Cells treated with 1 μM active KLK4, to ensure maximal calcium mobilisation, showed that cytosolic Ca2+ release was significantly reduced (p<0.01) by an average of 58% in cells stably transfected with construct number #2 in comparison to cells stably transfected with the non-specific control construct (set as 100%) in two

91 independent experiments (Figure 4.10A). Consistent with this finding, 100 μM PAR-2 AP stimulated Ca2+ mobilisation was significantly reduced (p<0.01) by an average of 61% due to PAR-2 depletion by construct #2 (Figure 4.10B).

Figure 4.10 Reduction of calcium mobilisation by PAR-2 siRNA mediated protein depletion. PAR-2-LMF cells stably transfected with either PAR-2 siRNA construct #2 or non-specific siRNA construct (NS), grey and black traces respectively, were incubated with (A) active KLK4 (1 μM) or (B) PAR-2 AP (100 μM) and fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm (Em510(340/380)) using a fluorescent plate reader. The ratio of Em510(340/380) is proportional to intracellular Ca2+ ion concentration. Arrow indicates time of treatment. To the right of each calcium release trace is a graphical comparison where Ca2+ release (peak height of the trace) is displayed graphically as a percentage of the highest recorded response. Values represent the means +/- SEM of duplicate experiments performed in triplicate. Statistical significance (p<0.01) was determined with student’s t-test with a significance threshold set at p<0.05.

92 In conjunction with Ca2+ mobilisation, reduction in activation of the ERK pathway was assessed following PAR-2 protein depletion. PAR-2-LMF cells stably transfected with siRNA construct #2 or the non-specific control were treated with either 40 nM active KLK4 or 100 μM PAR-2 AP for 5, 10 and 15 minutes, or remained untreated (0 minutes; basal). Protein lysates were extracted and analysed by Western blot analysis with anti-phosphorylated ERK and anti-ERK antibodies. Densitometric analysis of the ERK western blots was used to determine values for total ERK at each time point relative (5, 10 and 15 minutes) relative to untreated cells (0 minute; basal) for the KLK4 and PAR-2 AP treatments. Similarly, densitometric analysis of the anti-phosphorylated ERK Western blot was used to determine a value for phosphorylated ERK at each time point (5, 10 and 15 minutes) relative to untreated cells (0 minute; basal) for each treatment. The ratio of these values is the fold increase of ERK activation (phosphorylation) over basal levels (Figure 4.11; bottom panels); an approach used by Morris and colleagues to assess the magnitude of ERK1/2 phosphorylation following PAR-1 and PAR-2 activation in the MDA-MB-231 cell line (Morris et al., 2006).

To quantitate ERK1/2 phosphorylation reduction levels in the PAR-2-LMF construct #2 cell line for each treatment, the maximal ERK phosphorylation value obtained normalised to total ERK (the non-specific control cell line 5 minute time point) was set as 100% and all other values graphed relative to it. When treated with 40 nM active KLK4, cells transfected with construct #2 showed an average of 56% reduction (p <0.05) in ERK1/2 phosphorylation at the 5 minute time points relative to cells transfected with a non-specific control, with the reduction sustained at the 10 minute and 15 minute time points (p <0.05), in three independent experiments (Figure 4.11; bottom left panel). Similarly, an average 58% reduction (p <0.05) in ERK1/2 phosphorylation at the 5 minute time point was also apparent in cells with reduced PAR-2 levels which had been treated with 100 μM PAR-2 AP, with the reduction sustained at the 10 minute and 15 minute time points (p <0.05), in three independent experiments (Figure 4.11; bottom right panel). Overall, the 60% reduction of PAR-2 protein levels mediated by siRNA knock-down was consistent with the reduction in signal transduction through Ca2+ mobilisation and ERK phosphorylation (average of 56-61%).

93

Figure 4.11 Reduction of ERK1/2 phosphorylation by PAR-2 siRNA mediated protein depletion. PAR-2-LMF cells were stably transfected with PAR-2 siRNA construct #2 or a scrambled sequence siRNA control construct (NS, non-specific siRNA) and treated with either 40 nM active KLK4 (left panels) or 100 μM PAR-2 AP (right panels). Lysates were collected at the indicated time points and analysed by anti-ERK and anti-phospho ERK (p-ERK) Western blot analysis. The ratio of p-ERK to total ERK are represented graphically as the mean (+/- SEM) of three individual experiments, with the fold over basal phosphorylated ERK value for the non-specific control cell obtained at the 5 minute time point set as maximal phosphorylation (100%). The fold over basal phosphorylated ERK values of the other time points for the non-specific control and PAR-2 siRNA construct #2 cells lines are displayed as a percentage of this value for the KLK4 and PAR-2 AP treatments. Statistical significance was determined with student’s t-test with a significance threshold set at p <0.05. The Asterisks above the PAR-2 siRNA #2 trace denotes a statistically significant reduction (p <0.05) at the specified time point in comparison to the non-specific control.

94 4.3.5 Discussion

Intracellular signaling, initiated via activation of members of the PAR family of GPCRs by the action of trypsin-like serine proteases, is important in several physiological processes, including inflammation, cardiovascular responses and skin function (Macfarlane et al., 2001). Furthermore, aberrant cell signaling initiated by trypsin-like serine proteases and transduced via PARs has been shown to subjugate diverse cellular processes including differentiation, proliferation, migration, angiogenesis and evasion of apoptosis (Macfarlane et al., 2001) - processes critical for development and progression of cancer (Hanahan and Weinberg, 2000). The serine protease agonists responsible for PAR activation in these settings, where many proteolytic enzymes can be present, are not well defined. This Chapter has demonstrated the ability of the arginine/lysine specific (Takayama et al., 2001b), prostate cancer associated (Day et al., 2002; Dong et al., 2005; Veveris-Lowe et al., 2005; Gao et al., 2007; Klokk et al., 2007) serine protease KLK4 to initiate intracellular signaling through the activation of PAR-1 and PAR-2, but not PAR-4 (Figure 4.1, 4.2 and 4.3 respectively)

More specifically, experiments demonstrated that KLK4 initiates a prompt, transient Ca2+ mobilisation in PAR-1- and PAR-2-expressing cells. The specificity of the response via PAR-1 and PAR-2 was indicated by the lack of Ca2+ flux in cells expressing PAR-4 in response to KLK4. Furthermore, positive control agonists for PAR-1 (thrombin and PAR-1 AP), PAR-2 (trypsin and PAR-2 AP), and PAR-4 (trypsin and PAR-4 AP) each induced changes in Ca2+ concentration in the respective PAR-expressing cells, confirming that the receptor expressed by each cell line was functional. In other controls, proKLK4 and phosphoramidon-inhibited thermolysin did not initiate PAR signaling, while PAR-4 cells were shown to have intact PAR following treatment with KLK4 as the PAR-4 AP, applied 3 min after protease treatment, induced transient Ca2+ mobilisation.

The ability to signal via more than one PAR is known for other serine proteases. These include thrombin, which signals via PAR-1 and PAR-4 and also cleaves PAR- 3 (Coughlin, 1999), trypsin, which activates PAR-2 (Nystedt et al., 1994) and PAR-4 (Xu et al., 1998), activated Factor X, which, in complex with activated Factor VII

95 and tissue factor, signals via PAR-1 and PAR-2 (Camerer et al., 2000; Riewald and Ruf, 2001) and KLK14, which activates both PAR-2 and PAR-4 and both activates and disarms PAR-1 (Oikonomopoulou et al., 2006). Interestingly, our analysis of Ca2+ mobilisation in response to KLK4 suggests that although signaling via PAR-1 is more potent than via PAR-2, KLK4 displayed greater efficacy for signaling via the latter PAR (Figure 4.5). This contrasts with trypsin IV, which had similar potency as well as efficacy for inducing Ca2+ mobilization via these PARs in receptor over- expressing cells (Knecht et al., 2007).

Higher potency via PAR-1 may be a result of lower levels of expression of the PAR- 1 receptor, or because at higher enzyme concentrations, KLK4 cleaves the PAR-1 N- terminus at sites other than the activation sequence, thereby inactivating or “disarming” the receptor. Although these proposals remain to be tested, the latter has been suggested to explain the observations that Ca2+ mobilisation via PAR-2 in response to KLK6 reached a maximum at a much lower concentration than KLK14, and this maximal response to KLK6 was much lower than observed for KLK14 (Oikonomopoulou et al., 2006). It was proposed by Oikonomopoulou and colleagues that the responses to KLK6 were due to a receptor activating ability at lower enzyme concentrations via cleavage of PAR-2 at the consensus activation site and to a “disarming” ability at higher concentration via cleavage downstream of the activation site, although this group is yet to publish data supporting this hypothesis (Oikonomopoulou et al., 2006). Thus, like the observations made for KLK6 activation of PAR-2, it is possible that KLK4 regulates cellular activity by differential activation of PAR-1 and PAR-2; at lower enzyme concentration signaling via both PARs, and as KLK4 concentration increases via PAR-2 as PAR-1 is potentially disarmed.

This hypothesis parallels observations of thrombin mediated signaling in human platelet cells. Exposure of human platelets, which express both PAR-1 and PAR-4, to thrombin results in a biphasic Ca2+ mobilisation profile, a result of dual receptor activation (Kahn et al., 1998). However, PAR-4 is relatively insensitive to thrombin, with half maximal response approximately 50 fold higher than PAR-1 (Kahn et al., 1998). This suggests that PAR-4 may function as a low affinity thrombin receptor that is activated in conditions where high concentrations of thrombin are achieved,

96 helping to sustain thrombin responses when PAR-1 becomes rapidly inactivated (Kahn et al., 1998).

Due to the ability of KLK4 to signal with higher efficacy via PAR-2, we analysed signaling via this receptor:agonist system in greater detail. Our enzymatic studies, using a fluorescence quenched peptide indicated that KLK4 efficiently cleaves a peptide spanning the PAR-2 activation site, suggesting that intracellular signaling mediated by KLK4 occurs via direct proteolytic activation of this receptor. KLK4 was able to efficiently cleave the PAR-2 peptide with kinetic parameters of Km 8.0, kcat = 7.9 s-1 and kcat/Km = 1.0 (s.μM)-1 (Figure 4.6), which are similar to those previously observed for KLK6 against a similar PAR-2 peptide (Abz- SSKGR↓SLIGQ-Dpn) of Km =5.6 μM, kcat = 7.9 s-1 and kcat/Km = 1.4 (s.μM)-1) (Angelo et al., 2006). The kinetic constants derived from PAR-2 N-terminus cleavage by KLK4 are significantly less than those for trypsin cleavage (Figure 4.6). This is possibly due to the structural determinants, highlighted in the Chapter 3 discussion, which influence a narrowed active site cleft (Debela et al., 2006a), regulating the interaction between KLK4 and the N-terminus of PAR-2 in a more controlled manner than that of trypsin. Although direct cleavage of a PAR-2 peptide by KLK4 has been shown in this study, an indirect cleavage mechanism cannot be ruled out. In a study of PAR-2 activation on endothelial cells, factor VIIa was seen to act both indirectly by generating factor Xa for PAR-2 activation, and directly, by activating PAR-2 following binding to tissue factor expressed on the cell surface (Camerer et al., 2000). The factor VIIa and tissue factor complex highlights the complexity of mechanisms for PAR activation. Further mechanisms of PAR activation by KLK4 other than simple protease binding, for example, complex formation with a cell surface co-factor, support further investigation.

KLK4 is shown in this study to rapidly initiate ERK1/2 activation (Figure 4.7A). Interestingly, ERK1/2 activation began to display a maximal response at 100 nM active KLK4 (Figure 4.8), a markedly different result from the inability to saturate Ca2+ mobilisation, even with concentrations up to 1 uM (Figure 4.5). The ability to saturate ERK1/2 may be a result of multiple intermediate molecules synergistically activating the ERK cascade. For example, stimulation of GTP/GDP exchange on ras p21 by thrombin is mediated through a Gi protein and tyrosine kinase dependent

97 mechanism. Whether the pathways are directly linked, or are separate signal transduction events remains unclear (Kahan et al., 1992). Interestingly, the GPCR β2 adrenergic receptor can activate ERK through both G protein dependent and independent pathways (Sun et al., 2007), highlighting potential for members of the PAR family to behave in a similar manner. The potential for multiple intermediates in ERK cascade activation following PAR-2 activation by KLK4 is potentially quite distinct from intracellular Ca2+ mobilisation, which is primarily mediated through a 2+ single protein, Gq/11 (Clapham, 2007). Furthermore, ERK1/2 activation and Ca mobilisation can both be initiated via disparate events at the cell surface, including GPCR and receptor tyrosine kinase ligand docking (Malarkey et al., 1995; Berridge et al., 2003). Thus it is possible that KLK4 is able to cleave/activate other cell surface proteins in addition to PAR-1 and PAR-2 to initiate the differential induction of Ca2+ ion flux and ERK1/2 phosphorylation observed by us.

To address the specificity of KLK4 signal transduction through cell surface receptors, KLK4 signaling via PAR-2 was assessed by siRNA mediated knock-down of PAR-2 levels and correlated to changes in signal transduction. Depletion of approximately 60% of PAR-2 protein (Figure 4.9) correlated strongly to an approximately 60% reduction in both calcium mobilisation and ERK activation in response to cells being exposed to KLK4 (Figure 4.10 and 4.11), signifying that the initiation of KLK4 signal transduction in this study is largely initiated by PAR-2 activation. However, this result does not rule out KLK4 cleavage/activation of other surface proteins to initiate either Ca2+ mobilisation or ERK1/2 phosphorylation, as the PAR-2-LMF cells are an over-expression system and PAR-2 signaling could mask other transduction events mediated through additional cell surface proteins.

Lack of PAR-4 activation by KLK4 in this study may be a consequence of several factors (Figure 4.3). Firstly, the concentration of trypsin required for half maximal activation of PAR-4 is 5 times greater than that for PAR-2 (Nystedt et al., 1994; Xu et al., 1998). It is therefore possible that higher concentrations of KLK4 than those assayed are required for PAR-4 activation, an experiment unable to be performed due to limited amounts of recombinant KLK4. Furthermore, the presence of a cell surface receptor to act as a co-receptor may be required. In mouse platelet cells, which express only PAR-3 and PAR-4, it has been proposed that full activation of PAR-4

98 requires the presence of PAR-3 to act as a co-receptor. Consistent with this hypothesis, in mice platelet cells deficient in PAR-3, the response to thrombin is significantly delayed and less sensitive (Kahn et al., 1998). Therefore, further studies using higher concentrations of KLK4 are required to determine if this enzyme can activate PAR-4.

The activation of the PAR family elicits alterations in a number of cellular activites including proliferation, morphology, motility and cytokine release (Macfarlane et al., 2001). These activities are essential for physiological processes such as cardiovascular responses, neuronal cell survival, platelet aggregation, inflammation and epidermal function. Further, many of the cellular functions the PARs mediate have critical roles in cancer development and progression, such as apoptosis, proliferation and migration (Macfarlane et al., 2001). The role of PAR activation in prostate cancer will be discussed in further chapters. In summary, work done in this chapter has identified PAR-1 and PAR-2 as substrates of KLK4. Furthermore, KLK4 activates each of these receptors over distinct concentration ranges, suggesting a level of regulation complexity that supports further testing. In addition, KLK4 catalyses the mobilisation of the second messenger calcium and initiation of the signal transduction ERK cascade, primarily through PAR-2, likely utilising a direct cleavage mechanism.

99

100

CHAPTER 5

EXPRESSION AND LOCALISATION OF PROTEASE ACTIVATED RECEPTOR-2

IN PROSTATE CANCER TISSUE AND CELL LINES AND PROCESSING

FOLLOWING KALLIKREIN-RELATED PEPTIDASE 4 ACTIVATION

5.1 Introduction

Transcripts of each of the 15 KLKs have been detected in the prostate. Further, all kallikreins, except KLK8, are expressed at the protein level in normal or cancerous prostate (Clements et al., 2004; Shaw and Diamandis, 2007). KLK4 is highly expressed in normal prostate (Nelson et al., 1999; Yousef et al., 1999; Harvey et al., 2000) and recently this protease has been associated with prostate cancer progression (Veveris-Lowe et al., 2005; Gao et al., 2007). For example, stable over-expression of KLK4 in prostate cancer PC-3 cells resulted in an increased ability of these cells to migrate, accompanied by a transition from an epithelial morphology to a fibroblastic shape and, consistently, a significant decrease in E-cadherin protein levels and an increase in vimentin expression (Veveris-Lowe et al., 2005). In addition, using an inducible expression system, it has been demonstrated that over-expression of KLK4 results in significantly increased colony formation, migration and proliferation of the PC-3 and DU145 prostate cancer cell lines (Klokk et al., 2007). Furthermore, KLK4 protein levels are elevated in malignant prostate compared with normal tissue (Dong et al., 2005; Klokk et al., 2007), while prostate cancer patient sera contains antibodies that bind recombinant KLK4 (Day et al., 2002). Most recently using co- culture systems, it has been shown that KLK4 is a potential mediator of cellular interactions between prostate cancer cells and osteoblasts (bone forming cells) in bone metastases (Gao et al., 2007).

Recently several prostatic KLK enzymes with trypsin-like substrate specificity have been shown to cleave PARs leading either to receptor activation or disarming (Oikonomopoulou et al., 2006; Stefansson et al., 2008). Furthermore, the transcription and expression of PAR family members has been observed in normal and neoplastic prostate tissue, and prostate cancer cell lines. In the first paper reporting the cloning of the human PAR-2 cDNA, Nystedt and colleagues demonstrated high transcription of the mRNA for this receptor in normal prostate tissue (Nystedt et al., 1995). Importantly, members of this receptor family have been demonstrated by a number of studies to be elevated in prostate cancer. In a study by Black and colleagues, PAR expression in prostate cancer samples was elevated in comparison to normal prostate glands for PAR-1 (45%), PAR-2 (42%) and PAR-4 (68%), with elevated expression most notable in cancerous epithelial cells (Black et

102 al., 2007). Importantly, changes in cancer associated cellular functions result as a consequence of PAR-2 activation in prostate cancer derived cell lines. For example, in LNCaP cells, PAR-2 AP induced activation of members of the Rho GTPase family (Greenberg et al., 2003; Black et al., 2007); proteins which are of critical importance in cytoskeletal reorganisation and cell migration (Titus et al., 2005). In addition, AP mediated activation of PAR-2 induced activity of the collagenases MMP-2 and -9 by the prostate cancer cell lines LNCaP, PC-3 and DU145 (Wilson et al., 2004).

Having established that KLK4 is an efficient activator of PAR-2 (Chapter 4), the potential physiological relevance of KLK4 signaling via PAR-2 was explored by immunohistochemical analysis of agonist and receptor expression in primary prostate cancer and bone metastasis lesions. We have also examined cellular consequences of KLK4 mediated signaling via PAR-2 in prostate cancer PC-3 cells.

5.2 Materials and Methods

These are comprehensively described in Chapter 2. The reagents used specifically in experiments performed in this chapter are described below.

Serial sections (4 μm) of archival formalin-fixed paraffin-embedded blocks from primary prostate cancers (n = 6; Gleason scores 3+4 to 4+5) and prostate cancer bone metastases (n = 2) were obtained from Sullivan Nicolaides Pathology (Taringa, Australia) and the Royal Prince Alfred Hospital (Sydney, Australia), respectively, following institutional ethics approval (immunohistochemistry was performed by Dr Ying Dong).

A mammalian expression construct encoding PAR-2 with green fluorescent protein (GFP) at the C-terminus was generated in the pEGFP-N1 vector (Clontech, Mountain View, CA). PCR employing Pfu DNA polymerase (Invitrogen) was used to amplify the PAR-2 coding region with the specific primers 5'- CTCGAGCTTCCAGGAGGATGCGG -3' (Xho1 site underlined) and 5'- GAATTCGATAGGAGGTCTTAACAGTGGTTGAAC -3' (EcoR1 site underlined). Following amplification and purification, the PAR-2 PCR product was restriction

103 digested by Xho1/EcoR1 and ligated into an Xho1/EcoR1 digested pEGFP-N1. The integrity of the construct was verified by dye terminator sequencing.

5.3 Results

5.3.1 Expression of PAR-2 in normal, neoplastic and metastatic prostate tissues Having demonstrated that the prostate cancer associated protease KLK4 efficiently cleaves a peptide spanning the PAR-2 activation site and initiates intracellular signaling via PAR-2 to mobilise cytosolic Ca2+ and phosphorylate ERK, the potential pathological relevance of these observations was examined by performing immunohistochemical analysis of 6 primary prostate cancer tissue specimens. The expression pattern of receptor (PAR-2) and agonist (KLK4) in consecutive tissue sections was analysed using a PAR-2 specific monoclonal antibody (SAM-11) and the previously described anti-KLK4 antibody (Harvey et al., 2003). Representative images are shown in Figure 5.1. PAR-2 and KLK4 had similar expression patterns with both expressed by glandular epithelial cells with little evidence of stromal staining in serial sections (Figures 5.1A-B and 5.1D-E respectively). PAR-2 was detected in benign glands (BNG) of benign prostatic hyperplasia (BPH), prostatic intraepithelial neoplasia (PIN) and cancer (Ca). Expression levels were higher in regions of PIN and Ca than in BNG regions of BPH (comparing BNG and PIN in Figure 5.1A and BNG and Ca in Figure 5.1B). A similar staining pattern was apparent for KLK4. As published previously (Dong et al., 2005), prostate cancer tissue samples showed little KLK4 staining in benign glands, with stronger staining in regions of PIN (comparing BNG and PIN in Figure 5.1D) and Ca (comparing BNG and Ca in Figure 5.1E). Negative controls were free of staining (Figures 5.1D and 5.1H).

Bone metastasis is the cause of significant morbidity and mortality in prostate cancer patients (Shaffer and Scher, 2003). Recently our laboratory demonstrated that KLK4 is expressed by both prostate cancer cells and osteoblasts in the in vivo metastatic bone environment and that osteoblast-like SaOs2 cells induce KLK4 expression in co-culture systems with prostate cancer derived LNCaP and PC-3 cells (Gao et al., 2007). Accordingly, we performed immunohistochemistry to examine the expression pattern of KLK4 and PAR-2 in consecutive sections from prostate cancer bone

104 metastases from 2 patients. As shown in Figure 5.2 receptor and agonist had significant overlap in expression in prostate cancer bone metastasis. In regions of prostate cancer lesions PAR2 and KLK4 were highly expressed by prostate cancer cells (Figures 5.2A and 5.2C respectively). In regions containing prostate cancer lesions and bone, strong staining of osteoblasts lining the bone surface was also apparent for both PAR2 and KLK4 (arrows Figures 5.2B and 5.2D respectively). Negative controls were free of staining (data not shown).

Figure 5.1 Immunohistochemistry of PAR-2 and KLK4 expression in primary prostate cancers. Consecutive sections (4 μm) of primary prostate cancer specimens were stained with either an (A-B) anti-PAR-2, (D-E) anti-KLK4 or (C-F; negative control) secondary only antibody. Prostate intraepithelial neoplasia (PIN), benign glands (BNG) of benign prostatic hyperplasia, and cancer (Ca) are indicated.

105 5.3.2 Transcription and expression of PAR-2 in normal and neoplastic prostate epithelial cell lines

Having established PAR-2 co-expression with KLK4 in primary prostate cancer and prostate cancer bone metastasis, the cellular processing of KLK4 activated PAR-2 in prostate cancer cell lines was analysed. Accordingly, initial experiments first examined the expression of PAR-2 in prostate epithelial derived cell lines. The presence of PAR-2 in 5 prostate cell lines derived from a range of pathologies was examined by RT-PCR and Western blot analysis using PAR-2-LMF cells as a positive control for PAR-2 mRNA and protein. The cell lines were derived from the following sources: RWPE1, virus transformed normal epithelium (Webber et al., 1996); RWPE2, Ki-ras transformed RWPE1 cells (Webber et al., 1996); LNCaP, lymph node metastasis (Horoszewicz et al., 1983); PC-3, bone metastasis (Kaighn et al., 1979); and DU145, brain metastasis (Stone et al., 1978). As shown in Figure 5.3, PAR-2 mRNA was detected in each of these cell lines as indicated by the presence of a specific 583 bp product. Negative control lanes containing no template showed no contaminating DNA was present in the experiment. Consistent with calcium mobilisation experiments in which no flux was observed when PAR-1- and PAR-4- LMF cell lines were exposed to PAR-2 AP, transcription of PAR-2 was detected in PAR-2-LMF cells but not in PAR-1- or PAR-4-LMF cells.

Consistent with PCR results, PAR-2 protein was detected at high levels in each of the analysed cell lines with bands at ~55, 65 and 85 kDa (Figure 5.4). The different bands represent differential post-translational modification of the PAR-2 protein. Consistently, PAR-2 bands at ~65, ~85 and ~105 kDa reduced to the ~55 kDa band on tunicamycin treatment of breast cancer derived cell lines, while the ~85 kDa was also phosphorylated in these cells (Ge et al., 2004). Further, different intensities for the ~65 kDa band were observed in the LNCaP and DU145 cell lines, potentially indicating differing levels of PAR-2 post translational-modifications between the cell lines (Figure 5.4). Interestingly, only the 55 kDa form of PAR-2 was present in the PAR-2-LMF cell line, due possibly to differences in post-translational modifications introduced by mouse LMF cells and the human prostate derived cell lines (Figure 5.4).

106

Figure 5.2 Immunohistochemical analyses of PAR-2 and KLK4 expression in prostate cancer bone metastasis. Consecutive sections (4 μm) were stained with either an anti-PAR2 or an anti-KLK4 antibody. A. PAR-2 staining in prostate cancer cells in a prostate cancer bone metastasis lesion. B. PAR-2 staining in a prostate cancer bone metastasis lesion showing a region of bone. C.KLK4 staining in prostate cancer cells in a prostate cancer bone metastasis lesion. D. KLK4 staining in a prostate cancer bone metastasis lesion showing a region of bone. Bone, cancerous cells (Ca) and osteoblasts lining the bone surface (arrows) are indicated.

Figure 5.3 The transcription of PAR-2 in prostate epithelial derived cell lines. RT-PCR analysis of PAR-2 mRNA expression (30 cycles) in PAR-1-LMF, PAR-2- LMF, PAR-4-LMF and the prostate epithelial derived cell lines RWPE-1, RWPE-2, LNCaP, PC-3 and DU145 using the specific primers 5’-AGAAGCCTTATTGGTAA GGTT-3’ and 5’-AACATCATGACAGGTGGTGAT-3’ to give a 582 bp product. The B-actin house-keeping gene was used as a positive control (30 cycles) with the specific primers; human 5’-TGTCACCTTCACCGTTCCA-3’ and 5’CAAGATCAT TGCTCCTCCTG-3’ and mouse 5’ CGTGGGCCGCCCTAGGCACCA-3’ and 5’- TTGGCCTTAGGGTTCAGGGGGG-3’) to give a 313 bp and 250 bp product respectively. No template controls (control) were included as negative controls for the experiments.

107

Figure 5.4 The expression of PAR-2 in prostate epithelial derived cell lines. SDS-PAGE on a 12% gel run under reducing conditions with Western blot analysis of PAR-2 protein expression in the prostate derived cell lines RWPE-1, RWPE-2, LNCaP, PC-3 and DU145 and the mouse derived PAR-2-LMF cell line using an anti-PAR-2 antibody (SAM11; detects mouse and human PAR-2).

5.3.3 Localisation of endogenous PAR-2 in a prostate cancer cell lines

For serine protease mediated PAR activation to occur, the receptor must be located on the cell surface (Vu et al., 1991). Therefore, experiments to delineate surface localisation of endogenous PAR-2 in PC-3 cells were performed using two approaches: (1) biotinylation of intact cells and (2) confocal microscopy.

In biotinylation experiments, plasma membrane proteins were isolated by treating intact PC-3 cells with a biotinylating agent and biotinylated (cell surface) proteins purified using streptavidin conjugated beads. This technique permits separation of cell surface and cytoplasmic proteins. Western blot analysis using an anti-PAR-2 antibody following SDS-PAGE run on a 10% gel run under reducing conditions of the cell membrane (M) and cytoplasmic (C) fractions showed that no endogenous PAR-2 was detectable at the cell surface (Figure 5.5A; M-lane 2), presumably due to in vitro autocrine/paracrine activation and subsequent internalisation of PAR-2 by endogenous serine proteases expressed by the cell line. Accordingly, to inhibit in vitro activation of PAR-2 during culture, the broad range serine protease inhibitor aprotinin was added to PC-3 cells before plasma membrane and cytoplasmic proteins were isolated from intact cells using cell surface biotinylation. Aprotinin was demonstrated to enrich for cell surface levels of endogenous PAR-2, as detectable levels of PAR-2 resulted from aprotinin treatment (Figure 5.5A; M-lane 4).

108 Furthermore, endogenous PAR-2 was detectable in the cytoplasm of cells with and without aprotinin treatment (Figure 5.5A; C-lane 1 and lane 3). Cell surface biotinylation was determined to be efficient in separating membrane and cytoplasmic proteins due to the presence of the cytoplasmic protein GAPDH only in the cytoplasmic fractions (Figure 5.5B; lane 2 and 4 upper panel) and the presence of the cell surface protein integrin α2 (Figure 5.5C; lane 1 and 2 lower panel), in the membrane fractions.

Figure 5.5 Surface biotinylation of endogenous PAR-2 in PC-3 cells. A. Western blot analysis following SDS-PAGE run on a 10% gel run under reducing conditions of protein fractions collected from PC-3 cells with and without aprotinin treatment partitioned between cytoplasmic (C) and plasma membrane (M) fractions by cell surface biotinylation. Fractions were analysed for PAR-2 using antibody SAM11, and for purity using an antibody against a cytoplasmic marker (GAPDH; B). C. Western blot analysis following SDS-PAGE run on a 10% gel run under reducing conditions of the membrane fractions used in panels A and B for a known cell surface protein, integrin α2, using an anti-integrin α2 antibody.

To confirm the cell surface localisation of endogenous PAR-2 observed in biotinylation experiments, confocal microscopy analysis was employed. PC-3 cells were first transiently transfected with a GFP expression construct (green). This was performed to permit delineation of the plasma membrane/cytoplasmic boundary of cells expressing GFP. As the cell surface biotinylation results demonstrated that aprotinin effectively enriched for surface levels of endogenous PAR-2 (Figure 5.5A;

109 lane 4), PC-3 cells transiently transfected with GFP were aprotinin treated before endogenous PAR-2 was detected with a PAR-2 specific monoclonal antibody (red). Serial Z sections from the apical to the basal surface of GFP expressing PC-3 cells identified the presence of endogenous PAR-2 at high levels within the cytoplasm. Further, a small amount of this receptor was present on the cell membrane (arrowheads), outside the cytoplasm boundary marked by GFP (Figure 5.6). PAR-2 that localised to the surface of PC-3 cells displayed a punctate staining pattern (Figure 5.6; arrowheads), similar to that observed in the breast cancer cell line MDA-MB-231, in which PAR-2 localises to lipid rafts and caveolae (Awasthi et al., 2007).

Figure 5.6 Confocal microscopy analysis of endogenous PAR-2 in PC-3 cells. Confocal microscopy analysis of aprotinin treated PC-3 cells transfected with GFP (green) to delineate the plasma membrane/cytoplasmic boundary. Endogenous PAR- 2 was detected with an anti-PAR-2 antibody (SAM-11) followed by incubation with an anti-mouse secondary antibody conjugated with a fluorescent tag (red). Panels from left to right (i-viii) represent consecutive Z-sections from the apical to basal surface and display large cytoplasmic stores of PAR-2 and a small amount of PAR-2 at the cell surface (arrowheads).

In comparison to the rest of the plasma membrane, lipid rafts and caveolae, which are cholesterol and glycosphingolipid rich micro-domains, are enriched in Src family kinases, various GPCRs, their downstream signaling molecules and many other signaling proteins (Galbiati et al., 2001). Thus, due to their known high density GPCR content and the similarities in staining patterns between confocal microscopy

110 analysis in this study and work by Awasthi and colleagues, lipid raft localisation for PAR-2 was examined. A fluorophore conjugated to cholera-toxin B-subunit, which binds to the lipid raft constituent ganglioside GM1 (green), was used to stain lipid rafts in PC-3 cells that had been aprotinin treated and endogenous PAR-2 detected with an anti-PAR-2 antibody (red). As shown in Figure 5.7 PAR-2 (red) was localised to punctate structures in both the cytoplasm and on the plasma membrane of PC-3 cells as was apparent in Figure 5.6, with the majority of the PAR-2 signal detected in the cytoplasm. Interestingly, there was no overlap between PAR-2 signal and lipid raft localisation (green) in Figure 5.7, indicating that this receptor is not located within lipid rafts on the surface of PC-3 cells.

Figure 5.7 Examination of PAR-2 lipid raft localisation. Confocal microscopy analysis of aprotinin treated PC-3 cells probed for endogenous PAR-2 with an anti- PAR-2 antibody (SAM-11) followed by incubation with an anti-mouse secondary antibody conjugated with a fluorescent tag (red). Lipid rafts were detected with an Alexa fluor-488 conjugated Cholera toxin B subunit (green). Panels from left to right (i-iv) represent consecutive Z-sections from apical to basal surfaces and show that surface PAR-2 (arrowheads) does not co-localise with cholera-toxin B subunit to lipid rafts in the cell membrane.

111 5.3.4 KLK4 activation of endogenous PAR-2 in a prostate cancer cell line

Having demonstrated surface expression of endogenous PAR-2 in PC-3 cells, the ability of KLK4 to initiate signaling in these cells was assessed by examining transient changes in Ca2+ ion mobilisation. As shown in Figure 5.8A, 300 nM active KLK4 initiated a prompt, transient Ca2+ mobilisation in these cells. Further, PAR-1 AP and PAR-2 AP both initiated Ca2+ mobilisation (Figure 5.8B and 5.8C respectively), demonstrating that PC-3 cells have functional PAR-1 and PAR-2 present at the cell surface, and the Ca2+ mobilisation mediated by KLK4 is likely via possible activation of the two receptors (Figure 5.8A). The concentration response of KLK4 initiated Ca2+ mobilisation was examined by incubating PC-3 cells with increasing concentrations of enzyme (0.1 nM to 1 μM). As shown in Figure 5.8D increasing concentrations of active KLK4 stimulated increased Ca2+ mobilisation consistent with levels observed in PAR-2-LMF cells (Figure 4.3) and a concentration to stimulate half-maximal response calculated as pEC50 = 6.59 +/- 0.20 from a nonlinear regression curve using graphpad prism 4. Collectively these data indicate that PC-3 cells express functional PAR-1 and PAR-2 and that KLK4 initiates Ca2+ mobilisation in these cells in a concentration dependent manner.

5.3.5 KLK4 mediated internalisation of endogenous PAR-2 by flow cytometry

A well described event following PAR-2 activation is its internalisation from the cell surface. This involves PAR-2 uncoupling from G proteins and subsequent receptor endocytosis, followed by ubiquitination mediated lysosomal degradation (Bohm et al., 1996; Dery et al., 1999; Jacob et al., 2005). To determine if KLK4 activation induces the loss of PAR-2 from the surface of PC-3 cells, cell surface PAR-2 levels following KLK4 activation was determined by flow cytometry analysis.

As shown in Figure 5.9A, the non-permeabilisation fixation technique employed in this experiment (omission of a post-fixation detergent mediated membrane permeabilisation step, see section 2.2.12), in which the anti-PAR-2 antibody only has access to surface proteins, showed a fluorescence shift (black trace) in comparison to a control in which cells were incubated only with the secondary antibody (grey trace), indicating detection of cell surface PAR-2. To quantitate KLK4 and trypsin

112 mediated internalisation of endogenous PAR-2, PC-3 cell were treated with the respective agonist, subjected to the non-permeabilisation fixation process used in Figure 5.9A, then analysed by flow cytometry using an anti-PAR-2 antibody. A 10 minute exposure to 100 nM mature KLK4 or 10 nM trypsin resulted in a significant loss (p<0.05) of PAR-2 from the surface of PC-3 cells, of an average of 39% and 60% respectively (Figure 5.9B).

Figure 5.8 KLK4 initiated changes in intracellular Ca2+ ion concentration in prostate cancer derived PC-3 cell lines. PC-3 Cells were incubated with either (A) 300 nM mature KLK4, (B) 100 μM PAR-1 AP or (C) 100 μM PAR-2 AP as indicated and fluorescence at 510 nm was measured following alternating excitation at 340 nm and 380 nm(Em510(340/380)) using a fluorescent plate reader. The ratio of Em510(340/380) is proportional to intracellular Ca2+ ion concentration ([Ca2+]i). Experiments were performed in triplicate and repeated 3 times. D. Concentration dependence of KLK4 induced changes in intracellular Ca2+ ion concentration in prostate derived PC-3 cells. Cells were incubated with mature KLK4 (0.1 nM to 1 μM) and fluorescence was measured as described in A. Data is displayed graphically as a percentage of the highest recorded response (1 μM mature KLK4). Data represents mean values +/- SEM of triplicate experiments performed 3 times.

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Figure 5.9 Quantitation of endogenous PAR-2 internalisation in PC-3 cells. A. Flow cytometry analysis for cell surface PAR-2 on PC-3 cells using a non- permeabilising fixation process. Following fixation, cells were incubated with either an anti-PAR-2 (SAM11) antibody followed by a secondary antibody conjugated with a fluorophore (black line) or the secondary antibody only (grey line). B. Amount of surface PAR-2, determined by the flow cytometry method displayed in A, following 100 nM KLK4 or 10 nM trypsin treatments were plotted as a percentage of surface levels of PAR-2 in untreated PC-3 cells. Data represented is the mean of triplicate experiments +/- SEM. Statistical significance was determined with student’s t-test with a significance threshold set at p<0.05.

5.3.6 KLK4 mediated internalisation of exogenous PAR-2 in a prostate cancer cell line

The previous result sections of this Chapter have demonstrated that PC-3 cells express endogenous PAR-2 at the cell surface. PAR-2 at the surface of PC-3 cells is activated by KLK4 to induce Ca2+ mobilisation and subsequently leads to the loss of endogenous PAR-2 from the cell surface. To assess whether the loss of PAR-2 from the cell surface following KLK4 activation is due to the well established receptor internalisation pathway and not due to clearance from the cell surface by other mechanisms, confocal microscopy analysis of PAR-2 tagged with GFP was performed.

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Exogenously expressed PAR-2 tagged with GFP at the carboxyl terminal has been utilised in a number of studies to examine internalisation mechanisms of PAR-2 (Bohm et al., 1996; Dery et al., 1999). Further, GFP at the carboxyl terminal of PAR- 2 has been demonstrated not to interfere in signaling and trafficking of PAR-2 (Dery et al., 1999). As shown in Figure 5.10, in unstimulated PC-3 cells, PAR-2 was detected at the plasma membrane (arrowheads - upper overlay panel) and in cytoplasmic vesicles. Stimulation of PAR-2-GFP expressing cells with active KLK4 (100 nM) for 10 minutes, resulted in internalisation of PAR-2 from the cell surface and endocytosis into intra-cellular vesicle (Figure 5.10 arrow – lower overlay panel).

Figure 5.10 KLK4 treatment of PC-3 cells transiently transfected with a PAR-2- GFP expression construct. Cells, either unstimulated (upper panels) or stimulated with active KLK4 (100 nM; lower panels) for 10 minutes, were stained with DAPI to identify nuclei, fixed then imaged on a Leica SP5 confocal microscope. Cell surface PAR-2 is indicated by arrowheads in an unstimulated cell (upper overlay panel). Endocytosed PAR-2 is indicated by an arrow in a KLK4 stimulated cell (lower overlay panel).

As described in Chapter 2 quantitative analysis employing a modification of a previously described approach (Scherrer et al., 2006) was used to determine the amount of PAR-2 on the cell surface following KLK4 stimulation (Figure 5.11). As shown in Figure 5.11A using ImageJ software, cellular regions of interest were defined for the whole cell, the intracellular region and the nucleus (respectively red,

115 yellow and blue). The signal obtained for the whole cell and the intracellular region was corrected for background signal by subtracting nuclear fluorescence. Cell surface fluorescence was then obtained by subtracting the corrected value for the intracellular region from the corrected value for the whole cell. These values were then divided by the number of pixels contained within each region to give fluorescence density values for the cell surface (Di surface) and cytoplasm (Di cytoplasm). The ratio of Di surface to Di cytoplasm was determined to normalize data across the counted cell population. This analysis revealed that PAR-2 surface levels significantly decreased by ~80% following stimulation with KLK4 (p<0.05). These data indicate that KLK4 induces loss of exogenous PAR-2 GFP from the cell surface and receptor internalisation Figure 5.11B.

Figure 5.11 Quantitation of PAR-2 GFP internalisation in PC-3 cells. A. Using ImageJ software, cellular regions of interest were defined for the whole cell, the intracellular region and the nucleus (respectively red, yellow and blue) and signal intensity values obtained for each. Whole cell and intracellular values were corrected for background by subtracting nuclear fluorescence. Cell surface fluorescence was then obtained by subtracting the corrected value for the intracellular region from the corrected value for the whole cell and divided by the number of pixels contained within each region to give fluorescence density values for the cell surface (Di surface) and cytoplasm (Di cytoplasm). The ratio of Di surface to Di cytoplasm was determined to normalize data across the counted cell population. B. Quantification of cell surface PAR-2 described in A was performed on randomly selected unstimulated and KLK4 stimulated cells (n = 15) from randomly selected fields. Results are displayed graphically as mean +/- SEM. Statistical significance (p<0.01) was determined with student’s t-test with a significance threshold set at p<0.05.

116 5.4 Discussion

As indicated in Chapter 4 the prostate cancer associated enzyme KLK4 is an efficient activator of PAR-1 and PAR-2. In this Chapter, PAR-2 was selected for further analysis to examine the potential relevance of its activation by KLK4 to prostate cancer progression. Further, cellular processing of KLK4 activated PAR-2 was also examined.

To be of physiological relevance, receptor and agonist must be expressed at the same sites. Immunohistochemical analysis of primary prostate cancer and prostate cancer bone lesion samples in this chapter indicated expression of both PAR-2 and KLK4 by cells of glandular epithelium origin (Figure 5.1) as well as by osteoblasts (Figure 5.2), establishing potential for autocrine/paracrine activation interactions in vivo. Our observation of increased PAR-2 expression during prostate cancer progression is consistent with a recent report showing elevated PAR-2 in regions of prostate cancer compared with adjacent normal glandular epithelial cells in 42% of patient samples (n = 40) (Black et al., 2007). These researchers also showed that PAR-2 levels increased with increasing Gleason score (Black et al., 2007). In addition, our finding of PAR-2 expression by osteoblasts in prostate cancer bone metastases (Figure 5.2) is consistent with studies in mice demonstrating that murine osteoblasts also express this receptor in vitro and in vivo (Abraham et al., 2000). The key observation from our immunohistochemical analysis of PAR-2 and KLK4 co-expression in primary prostate cancer as well as bone metastasis lesions, suggests that KLK4 will contribute to PAR-2 activation in these settings.

This is potentially significant as recent reports have demonstrated cancer associated functional consequences of PAR-2 activation in prostate cancer derived cell lines. For example, in LNCaP cells, PAR-2 AP induced activation of members of the Rho GTPase family (Greenberg et al., 2003; Black et al., 2007); proteins which are of critical importance in cytoskeletal reorganisation and cancer (Titus et al., 2005). In addition, AP mediated activation of PAR-2 induced activity of the collagenases matrix metalloprotease 2 and 9 by the prostate cancer cell lines LNCaP, PC-3 and DU145 (Wilson et al., 2004). Further, it has been suggested that activation of PAR-2 in bone mediates inhibition of osteoclast differentiation and bone lytic activity and

117 this effect is likely mediated by osteoblasts (Smith et al., 2004). This is potentially significant as prostate adenocarcinomas have primarily an osteoblastic phenotype in bone metastasis lesions (Ye et al., 2007).

It is possible that KLK4 initiated signaling via PAR-1 will also be relevant in prostate cancer. Recently two reports have consistently identified an increase in expression at mRNA and protein levels of this receptor during prostate cancer progression, although the expression pattern reported by these groups differed (Kaushal et al., 2006; Black et al., 2007). Kaushal et al noted predominant expression in endothelial cells in cancerous regions with cancer cells also showing PAR-1 expression in some early and advanced stage specimens (Kaushal et al., 2006). In contrast, using a different antibody, Black and colleagues reported PAR-1 expression in cancer cells in 45% (n=40) of prostate cancer samples and in periglandular stromal cells in 55% (n=20) of higher grade cancers (Black et al., 2007). Of relevance PAR-1 expression has also been reported in human osteoblast- like cells (Jenkins et al., 1993) and mouse osteoblasts (Abraham et al., 1998) and this GPCR mediates thrombin induced proliferation of primary rat osteoblasts (Abraham and MacKie, 1999). Interestingly, cell lines derived from prostate cancer bone (VCaP) and soft tissue (DuCaP) metastases of the same patient showed increases in PAR-1 mRNA and protein in comparison to non-diseased adjacent tissue (Chay et al., 2002). Furthermore, PAR-1 expression was elevated in the VCaP in comparison to DuCaP cell lines, implicating PAR-1 as a component of metastastic progression (Chay et al., 2002). Therefore, like PAR-2, activation of tumour and osteoblast expressed PAR-1 by KLK4 may potentially be important for the establishment and growth of prostate tumours in the bone microenvironment.

Similarly, cancer associated functional consequences of KLK4 expression have been noted in prostate cancer derived cell lines. For example, KLK4 over-expression induced an increased ability of PC-3 cells to migrate, accompanied by a transition from an epithelial morphology to a fibroblastic shape and, consistently, a significant decrease in E-cadherin protein levels and an increase in vimentin (Veveris-Lowe et al., 2005). In addition, inducible over-expression of KLK4 resulted in significantly increased colony formation, migration and proliferation of both PC-3 and DU145 prostate cancer derived cells (Klokk et al., 2007). Recently, our lab has shown that

118 KLK4 levels increased in LNCaP and PC-3 cells co-cultured with osteoblast-like SaOs2 cells, while KLK4 over-expressing PC-3 cells had increased migration towards SaOs2 cell conditioned media (Gao et al., 2007). Although it is not clear how these KLK4 effects were mediated, KLK4 activation of PAR-2 may have been an integral factor and has potential as an important component of prostate cancer progression in both primary and bone lesions.

In addition to examination of expression of KLK4 and PAR-2 in prostate cancer this chapter also analysed cellular processing of KLK4 activated PAR-2. Before examining processing of PAR-2 it was first necessary to identify PAR-2 expressing prostate derived cell lines. RT-PCR and Western blot analyses indicated that PAR-2 is expressed by cell lines derived from normal and neoplastic prostate epithelium (Figure 5.3 and 5.4 respectively). Furthermore, the expression of PAR-2 in these prostate epithelial derived cell lines shows different forms of the receptor, due presumably to different post-translation modifications of PAR-2 (Figure 5.4). Multiple PAR-2 protein sizes is consistent with the finding of Ge et al in the breast cancer cell line MDA MB-231, where PAR-2 bands at ~65, ~85 and ~105 kDa reduced to the ~55 kDa band on tunicamycin treatment, while the ~85 kDa was also phosphorylated in these cells (Ge et al., 2004). Further, Compton and colleagues examined expression of exogenous PAR-2 in Chinese-hamster ovary fibroblasts and showed that PAR-2 displayed molecular weights that ranged from 55-100 kDa (Compton et al., 2002). Compton et al identified N-glycosylation as the primary source of post-translational modification of PAR-2, as removal of the N-linked glycosylation reduced the protein size range to 33-48 kDa, consistent with the predicted molecular weight of 44 kDa for PAR-2. N-glycosylation is a common post- translational feature of GPCRs that regulates their surface expression. Consistently, mutation of both N-glycosylation sequons has been shown to significantly reduce the surface expression of PAR-2 (Compton et al., 2002).

To transduce extracellular stimuli, PAR-2 must be localised to the cell surface (Bohm et al., 1996). This study has shown, following enrichment, the localisation of endogenous PAR-2 on the surface of PC-3 cells (Figure 5.5 and 5.6). Interestingly, unlike the breast cancer cell line MDA-MB-231, in which PAR-2 localises to lipid rafts and caveolae (Awasthi et al., 2007), PC-3 did not display lipid raft localisation

119 of its endogenously expressed PAR-2 (Figure 5.7). This however, does not eliminate the potential for association with proteins located within lipid rafts, illustrated by interaction between PAR-1 and the lipid raft protein flotillin-2 (Hazarika et al., 2004). Furthermore, the staining pattern observed in Figure 5.6 may represent PAR- 2 endocytosed into clathrin coated pits, due to staining pattern similarities between this study and work done by Bohm and colleagues (Bohm et al., 1996). Endocytosis is an important component of PAR-2 signal termination (Traynelis and Trejo, 2007).

The irreversible nature of PAR activation requires tightly regulated desensitization mechanisms for control of receptor signaling (Traynelis and Trejo, 2007). Both β- arrestin-1 and –2 are critical for termination of PAR-1 and -2 signal transduction, as an absence of both β-arrestins results in sustained phosphoinositide hydrolysis following activation of each receptor in murine fibroblasts (Paing et al., 2002; Stalheim et al., 2005). We have shown that KLK4 induces the rapid release of cytosolic Ca2+ in PC-3 cells, presumably via activation PAR-1 and PAR-2 activation, with a maximal release achieved within 20 seconds of receptor activation, followed by a subsequent termination of signal transduction (Ca2+ mobilisation) and a return to baseline (Figure 5.8A). Importantly, unlike the breast carcinoma cell line MDA-MB- 231 in which thrombin activation of PAR-1 is persistent due to dysregulated receptor trafficking (Booden et al., 2004), PC-3 displayed functional desensitizing mechanisms, illustrated by the rapid termination of Ca2+ mobilisation in response to KLK4 (Figure 5.8A), due presumably to β-arrestin binding. Future studies which include desensitization Ca2+ mobilisation experiments (the saturation of the PAR of interest with its respective AP to induce its removal from the cell surface) would enable the study of PAR-1 and PAR-2 in isolation, potentially identifying different efficacies in KLK4 signaling in the PC-3 cell line.

The termination of GPCR signaling in the continued presence of agonist is orchestrated by a series of events that can be considered as three distinct processes; receptor desensitization, internalisation and downregulation (Luttrell, 2006). The internalisation or endocytosis of GPCRs occurs more slowly than desensitization, taking place over the course of several minutes following receptor activation (Ferguson, 2001). Binding of β-arrestins to the phosphorylated GPCR facilitates the

120 agonist-promoted endocytosis through clathrin coated pits of many receptors including angiotensin II type 1a, endothelin A and monocyte chemoattractant protein-1 (Ferguson, 2001). In this study the internalisation of approximately 39% and 60% of endogenous PAR-2 occurs 10 minutes after KLK4 and trypsin activation respectively (Figure 5.9B). Potentially the discrepancy in internalisation efficiency between KLK4 and trypsin relates to differences in the rate of receptor activation by the enzymes. In Chapter 4 it was shown that every second a trypsin molecule cleaves 226 PAR-2 N-termini peptides (kcat = 226 sec-1), a rate which is approximately 30 times greater than KLK4 (kcat = 7.9 sec-1) (Figure 4.6). As PC-3 cells were exposed to 10 times the amount of molecules of active KLK4 (100 nM) than trypsin (10 nM), it would be expected that trypsin would activate approximately 35% (KLK4 kcat = 79 (7.9 x10) sec-1) / trypsin (kcat = 226 sec-1)) more cell surface PAR-2, correlating strongly to the additional 35% of endogenous PAR-2 internalised from the surface of PC-3 cells by trypsin in comparison to KLK4.

Downregulation of GPCRs, which is the persistent loss from the cell surface following GPCR internalisation, is the least understood of the events controlling GPCR signal termination. GPCR downregulation involves, at least in part, sorting of the internalised receptor through recycling pathways (Luttrell and Lefkowitz, 2002). However, unlike the majority of GPCRs, which are internalised into endosomes where they are dissociated from their agonist, dephosphorylated and returned to the cell surface, activated PAR-2 is mono-ubiquitinated, internalised and sorted directly to lysosomes for rapid degradation (Bohm et al., 1996; Dery et al., 1999; Jacob et al., 2005). As such, internalisation and downregulation collectively regulate PAR-2 signal termination by; removal from agonists and signaling effectors and prevention of continued signaling via recycling of activated receptor to the cell surface.

It was shown in this study that exogenous GFP tagged PAR-2 is rapidly internalised into cytoplasmic vesicles from the cell surface within 10 minutes of KLK4 activation, consistent with endogenous internalisation (Figure 5.10). Unexpectedly, exogenous PAR-2 was more readily internalised from the cell surface (compare Figure 5.9B to 5.11B), possibly due to differences in activation efficiencies between endogenous and exogenous PAR-2 by KLK4. The intracellular vesicles to which internalised PAR-2 localises are likely early endosomes, due to staining pattern

121 similarities between the lower panel in Figure 5.10 and timecourse studies by Bohm et al and Dery et al, in which internalised PAR-2 was shown to traffic to the early endosomes 10-30 minutes post activation (Bohm et al., 1996; Dery et al., 1999).

Interestingly β-arrestins, which are internalised complexed with PAR-2 in the early endosome, can recruit ERK1/2 to PAR-2 in endosomal scaffolds, allowing the endocytosed receptor to continue to signal (DeFea, 2007). Importantly, it has been proposed by DeFea and colleagues that the complex might ensure that ERK1/2 remains in the cytosol and does not translocate to the nucleus to stimulate proliferation (DeFea et al., 2000). Although the mitogenic effects of nuclear ERK1/2 have been commonly examined, cytoplasmic substrates exist for ERK1/2 including cytoskeletal proteins, whose phosphorylation is involved in cell migration and morphological alterations (Klemke et al., 1997). Consistent with KLK4 mediating prostate cancer cellular effects through PAR-2, and formation of a cytoplasmic PAR- 2/ERK scaffold complex similar to that seen in the breast cancer cell line MDA MB- 231 (Ge et al., 2004), the ectopic expression of KLK4 in PC-3 cells leads to a change in cell morphology and an increase in cellular migration (Veveris-Lowe et al., 2005). Furthermore, PC-3 proliferation was decreased as a result of KLK4 over-expression (Veveris-Lowe et al., 2005), providing additional support for the possibility of scaffold formation, that would inhibit ERK1/2 nuclear translocation, and thus eliminating its mitogenic effects, although further experiments would be required to test this hypothesis.

This study has shown for the first time that the receptors PAR-1 and PAR-2 are activated by the serine protease KLK4 and that PAR-2 and KLK4 are co-expressed during prostate cancer progression. Thus, KLK4-mediated cell signaling via PARs may be important in prostate cancer progression.

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CHAPTER 6

GENERAL DISCUSSION AND FUTURE DIRECTIONS

6.1 Key observations

This program of study has examined activation of members of the PAR family by the prostate cancer associated enzyme KLK4. A critical observation from this study is that KLK4 and PAR-2 are co-expressed during prostate cancer progression, establishing the potential importance of this receptor/agonist system in this disease. Importantly, an independent study has also observed increased PAR-2 expression, in addition to PAR-1 and PAR-4, in prostate cancer (Black et al., 2007).

Another critical finding is that KLK4 activates PAR-1 and PAR-2, but not PAR-4. Further, KLK4 activated PAR-1 and PAR-2 over distinct concentration ranges, with KLK4 activation and mobilisation of Ca2+ demonstrating higher efficacy through PAR-2. Thus, the remainder of this study focussed on PAR-2. KLK4 was demonstrated to directly cleave a synthetic peptide that mimicked the PAR-2 N- terminal activation sequence. Further, KLK4 mediated Ca2+ mobilisation through PAR-2 was accompanied by the initiation of the ERK signaling cascade. The specificity of ERK signaling mediated through PAR-2 by KLK4 activation was demonstrated by siRNA mediated protein depletion, with a reduction in PAR-2 protein levels correlating with a reduction in KLK4 mediated Ca2+mobilisation and ERK phosphorylation.

This study also delineated cellular processing of PAR-2 following activation by KLK4. Focusing on endogenous PAR-2, this receptor was found to be expressed by each of the five analysed prostate derived cell lines and, in particular, to be expressed on the surface of PC-3 cells. KLK4 was shown to initiate mobilisation of intracellular Ca2+ in these cells consistent with signaling via PAR-1 and PAR-2. Further, the activation of cell surface PAR-2 on PC-3 cells by KLK4 lead to internalisation of this receptor in a time dependent manner.

6.2 Implications of PAR-1 and PAR-2 activation for prostate cancer tumourigenesis

Consistent with the prostatic expression of PAR-1, androgen regulation of the encoding gene has been recently reported (Salah et al., 2005). These researchers

124 showed that stimulation of LNCaP cells with dihydrotestosterone (DHT) increased the transcription of the PAR-1 gene, whereas PC-3 cells, which lack an androgen receptor, were unresponsive. Furthermore, consistent with PAR-1 androgen responsiveness, neoplastic prostate tissue samples before and after androgen ablation therapy displayed a substantial decrease in PAR-1 mRNA levels after androgen ablation (Salah et al., 2005). Androgen regulation of the PAR-2 gene has not yet been demonstrated.

Like PAR-1, androgen regulation has also been reported for KLK4 in the prostate. KLK4 has been shown to be up-regulated by androgens at both the mRNA (Nelson et al., 1999; Korkmaz et al., 2001) and protein (Xi et al., 2004) level. However, androgen response elements have still to be identified in the KLK4 promoter. Functionally the over-expression of KLK4 has been demonstrated to alter several cellular functions that are associated with prostate cancer progression, including alterations in morphology and increases in cellular migration, colony formation, proliferation and invasion (Veveris-Lowe et al., 2005; Klokk et al., 2007). Activation of PAR-1 and PAR-2 may explain the cellular alterations accompanying KLK4 over- expression, as numerous studies have demonstrated that activation of PAR-1 and PAR-2 in cancer cell lines initiates a variety of cellular processes that are associated with cancer progression including proliferation, secretion of tumour promoting factors, changes in morphology and increased migration (Arora et al., 2007). Thus, future studies incorporating in vitro cellular assays to determine which cellular functions change in response to KLK4 activation of PAR-2 in prostate cancer cell lines would be beneficial.

The breakdown of the basement membrane and ECM components is an essential step for prostate cancer progression as it allows the spread of cancer cells from their primary site (Steeg, 2006). Indicative of a role in these events KLK4 degrades the ECM proteins type I and type IV collagen in vitro (Obiezu et al., 2006) and PAR-2 activation induces the secretion of MMP-2 and MMP-9 from prostate cancer cells (Wilson et al., 2004), proteases with well established roles in matrix remodelling. These observations suggest KLK4 involvement in prostate cancer metastasis through basement membrane degradation either through direct breakdown of ECM components and/or indirectly through PAR-2 activation and subsequent release of

125 MMP-2 and MMP-9. Once a path through the ECM and basement membrane is established, prostate cancer cells move from the prostate to preferentially invade the bone (Kozlowski and Sensibar, 1999).

6.2.2 Potential contributions of KLK4 activation of PARs to prostate cancer bone metastasis

The key observation from our immunohistochemical analysis of PAR-2 and KLK4 co-expression in primary prostate cancer as well as bone metastasis lesions, suggests that KLK4 will contribute to PAR-2 activation in these settings. Importantly, KLK4 has been proposed to be a potential mediator of cellular interactions between prostate cancer cells and osteoblasts in bone metastases (Gao et al., 2007). A striking feature of prostate cancer is its propensity to metastasise preferentially to the bone where it is essentially incurable (Kozlowski and Sensibar, 1999). Once within the bone microenvironment prostate cancer causes bone matrix turnover (Lange and Vessella, 1998), resulting in osteoblastic (bone formation) and osteolytic (bone destruction) lesions, although osteoblastic is the dominant remodelling phenotype, seen in approximately 85% of bone metastases (Yoneda, 1998). Thus, the pathological bone formation apparent in prostate cancer bone metastases is the result of the influx of cancer cells which alter the balance between the normal activities of osteoblasts (bone forming cells), osteoclasts (bone resorbing cells) and other cells of the bone microenvironment (Bonfil et al., 2007). The cancer cells are thought to subjugate normal bone remodelling processes involving differentiation and activity of osteoclasts from haemopoietic precursor cells (osteoclastogenesis) (Logothetis and Lin, 2005). A potentially important role for PAR-2 in these processes is indicated by the observation that in bone marrow cultures, which are rich in haemopoietic precursor cells, PAR-2 activation inhibits osteoclast differentiation induced by parathyroid hormone, 1,25-dihydroxyvitamin D or IL-11 and decreases the expression of the osteoclastic factors IL-6 and cyclooxygenase-2 (Smith et al., 2004). This suggests that PAR-2 functions to inhibit osteoclastogenesis and potentially attenuate the level of bone turnover, which is consistent with the predominant osteoblastic phenotype of prostate cancer bone metastases.

126 In support of this proposal, immunohistochemical analysis in this study has demonstrated PAR-2 expression by osteoblasts, consistent with studies in mice demonstrating that murine osteoblasts also express this receptor in vitro and in vivo (Abraham et al., 2000). Importantly, a potential role has also been indicated for PAR- 1 in pathological settings in bone by a report showing that activation of this GPCR by thrombin induces increased proliferation of primary rat osteoblasts (Abraham and MacKie, 1999). Thus, the demonstration by this study of KLK4 activation of PAR-1 and PAR-2 has potential to be functionally important in prostate cancer bone metastasis. The complexity of autocrine/paracrine interactions between KLK4, as agonist, and PAR-1 and PAR-2, as receptors, is highlighted by our observation that KLK4 and PAR-2 are both expressed by osteoblasts and tumour cells in prostate cancer lesions. Future studies examining the consequences of PAR-1 and PAR-2 activation by KLK4 on the cellular behaviours of osteoblasts and osteoclasts using in vitro assays and mouse models, may potentially delineate a molecular mechanism involved in the osteoblastic phenotype of prostate cancer bone metastases.

6.3 Mechanistic considerations of KLK4 activation of PAR-1 and PAR-2

The ability to signal via more than one PAR, as shown here for KLK4, is known for several serine proteases. These include thrombin, which signals via PAR-1 and PAR- 4 and also cleaves PAR-3 (Coughlin, 1999), trypsin, which activates PAR-2 (Nystedt et al., 1994) and PAR-4 (Xu et al., 1998), activated Factor X, which, in complex with activated Factor VII and tissue factor, signals via PAR-1 and PAR-2 (Camerer et al., 2000; Riewald and Ruf, 2001) and KLK14, which activates both PAR-2 and PAR-4 and both activates and disarms PAR-1 (Oikonomopoulou et al., 2006). This study has demonstrated that KLK4 signals via PAR-1 and PAR-2 and that KLK4 mediated Ca2+ mobilisation via PAR-1 is more potent than via PAR-2. Similar observations have been made about thrombin. Exposure of human platelets, which express both PAR-1 and PAR-4, to this serine protease results in a biphasic Ca2+ mobilisation profile, a result of dual receptor activation (Kahn et al., 1998). However, PAR-4 is relatively insensitive to thrombin, with half maximal response approximately 50 fold higher than PAR-1 (Kahn et al., 1998). The difference in thrombin potency between PAR-1 and PAR-4 is due to the presence of a hirudin–like binding domain in the amino terminal of PAR-1. Although not essential for activation, this structural motif

127 allows the efficient binding to and activation of PAR-1 by thrombin (Bouton et al., 1995).

In contrast to potency, KLK4 displays higher efficacy through PAR-2. This contrasts with trypsin IV, which had similar potency as well as efficacy for inducing Ca2+ mobilisation via PAR-1 and PAR-2 in receptor over-expressing cells (Knecht et al., 2007). The ability of KLK4 to tranduce stimuli at higher concentrations via PAR-2 than PAR-1 may signify that PAR-2 is the primary receptor for KLK4 transduction during pathological events of increased KLK4 release, such as cancer. This hypothesis may mirror the paradigm of thrombin mediated human platelet activation. When exposed to high levels of thrombin as a result of vascular injury, platelets rapidly have their surface PAR-1 activated, after which signaling is sustained by the low affinity thrombin receptor PAR-4, helping to prolong platelet aggregation (Macfarlane et al., 2001). Desensitisation of PAR-4 occurs more slowly relative to PAR-1, due to a delayed phosphorylation of the carboxyl tail of PAR-4 (Shapiro et al., 2000). A direction of future research will be to define the molecular mechanisms regulating differential KLK4 signaling potency via PAR-1 and PAR-2 and the mechanisms regulating the efficacy of the physiological responses.

6.4 Potential in vivo regulation mechanisms

Of importance to the regulation of PAR activation in normal and diseased prostate, in addition to KLK4, seven other prostate expressed serine proteases are known to initiate intracellular signaling via PAR-1 and PAR-2 in in vitro systems. These include the type II transmembrane serine proteases (Hooper et al., 2001) TMPRSS2 (Wilson et al., 2005), human airway trypsin (HAT) (Miki et al., 2003) and matriptase/MT-SP1 (Takeuchi et al., 1999), other KLK serine proteases, KLK5, KLK6 and KLK14 (Oikonomopoulou et al., 2006; Stefansson et al., 2008) and extra- pancreatic trypsin IV (Cottrell et al., 2004). The ability of multiple prostatic serine proteases to signal via PAR-1 and PAR-2 indicates that activation of these receptors in the prostate will require both tight regulation of the activating proteases (for example by the presence of activators and inhibitors) in addition to spatial and temporal expression of receptor and agonist.

128 6.4.1 Activation of the KLK4 zymogen

One of the key regulatory elements in protease function is the proteolytic conversion from inactive precursor to active enzyme (auf dem Keller et al., 2007). Due to the irreversible nature of their activation, serine proteases are produced as inactive zymogens that remain quiescent until external stimuli initiate their conversion to mature enzymes (Amour et al., 2004). The activation of serine proteases can sometimes be mediated by highly orchestrated cascades. Proteolytic cascades function to coordinate the sequential activation of protease zymogens, causing the rapid amplification of signal in response to minimal stimulus (Amour et al., 2004). Importantly, the possibility that KLK4 will activate PAR-1 and/or PAR-2 via a proteolytic cascade reminiscent of the blood coagulation (Davie et al., 1991) and wound healing (Castellino and Ploplis, 2005) serine protease cascades, has not yet been explored. However, proteolytic cascades involving other KLKs have recently been reported. For example, Brattsand and colleagues have implicated KLK5, 7 and 14 in a proteolytic cascade in the skin (Brattsand et al., 2005). In vitro data from that study suggests that KLK5 auto-activates and subsequently activates KLK7 and 14. Interestingly, KLK14 and multiple KLKs (KLK5, 6 and 14), activate PAR-1 and PAR-2 respectively (Oikonomopoulou et al., 2006; Stefansson et al., 2008), illustrating the potential of both receptors as targets of KLK cascades.

Evidence for a KLK cascade also exists in seminal fluid for KLK2, 3, 4, 5 and 14. In vitro experiments of auto-activated KLK5 show activation of KLK3, after which, activated KLK3 is inactivated by a series of internal cleavages by KLK5 (Michael et al., 2006). Furthermore, activated KLK2 (Kumar et al., 1997), KLK4 (Takayama et al., 2001a) and KLK14 (Emami and Diamandis, 2008) are reported to remove the pro-region of KLK3. Further, similar to KLK5, KLK14 regulation is bidirectional with its inactivation of active KLK3 via internal cleavages (Emami and Diamandis, 2008). Interestingly, a study of KLK zymogen activation profiles indicated that no KLK, with the exception of a minimal KLK11 effect, can activate KLK4, although KLK9, 10 and 15 were not tested (Yoon et al., 2007). Furthermore, this study indicated that KLK4 can remove the pro-region from a number of prostate expressed KLKs, including KLK1-3, 5, 6, 9 and 11-15 (Yoon et al., 2007). Collectively these data may indicate, in addition to its activation of PAR-1 and PAR-2, that KLK4 is a

129 potential initiator of a KLK cascade in the prostate, with many of the potential cascade products having the potential for PAR activation. Due to the highly overlapping tissue expression and activation profiles, it is apparent that tight regulation of a prostate KLK cascade would be required.

6.4.2 Regulation of KLK4 activity

Dysregulated serine protease activity in many disease settings requires, quite separately from protease over-expression or post translational modulation, the loss of mechanisms regulating normal activation and cessation of proteolytic activity (Turk, 2006). In prostate cancer, as in other diseases, this dysregulation is mediated by changes in levels of various factors including metal ions (eg. Zn2+) and proteinaceous 2+ inhibitors. For example, in prostate and other tissues Zn ions provide an effective means of regulating the action of several KLK and other trypsin-like enzymes including KLK2 (Lovgren et al., 1999a), KLK4 (Debela et al., 2006a), KLK5 (Michael et al., 2006; Debela et al., 2007), KLK8 (Kishi et al., 2006), KLK12 (Memari et al., 2007a) and KLK14 (Borgono et al., 2007c) as well as prostasin (Shipway et al., 2004) and uPA (Ishii et al., 2001). Of relevance to normal prostate 2+ physiology as well as prostate cancer, Zn levels are up to 10 times higher in normal prostate than in other tissues (Kavanagh, 1985), while there is a 10 to 20 fold decrease in these levels in patients with advanced prostate cancer (Zaichick et al., 2+ 1996; Zaichick et al., 1997). Based on in vitro studies this reduction in Zn levels would be expected to be accompanied by a corresponding increase in the activity of tryptic proteases.

Progression of prostate cancer is also often accompanied by loss of expression of proteinaceous inhibitors of trypsin-like enzymes. For example, the serpin anti thrombin III which complexes with KLK2 (Cao et al., 2002), KLK4 (Obiezu et al., 2006), KLK6 (Magklara et al., 2003), KLK12 (Memari et al., 2007b) and KLK14 (Borgono et al., 2007c) is down-regulated during prostate cancer progression (Frenette et al., 1997; Cao et al., 2002). Also, protein C inhibitor, another serpin which complexes with KLK1 (Ecke et al., 1992; Espana et al., 1995), KLK2 (Deperthes et al., 1995; Heeb and Espana, 1998; Mikolajczyk et al., 1999; Lovgren et al., 1999a), KLK5 (Luo and Jiang, 2006), KLK8 (Luo and Jiang, 2006), KLK12

130 (Luo and Jiang, 2006; Memari et al., 2007b) and KLK14 (Luo and Jiang, 2006), is down-regulated in prostate cancer (Cao et al., 2003). Further, levels of α-2 macroglobulin, which complexes with KLK2 (Frenette et al., 1997; Grauer et al., 1998; Heeb and Espana, 1998; Mikolajczyk et al., 1998), KLK4 (Obiezu et al., 2006), KLK5 (Michael et al., 2005), and KLK13 (Kapadia et al., 2004), also decrease during prostate cancer progression (Kanoh et al., 2001).

Interestingly, in addition to reduced levels of expression of protease inhibitors during prostate cancer progression there is evidence that certain trypsin-like serine proteases also modulate inhibitor levels by inactivation. For example, KLK2 has been reported to inactivate the serpin plasminogen activator inhibitor-1 by direct cleavage of the inhibitor (Mikolajczyk et al., 1999). Overall, future KLK4 studies of PAR-1 and PAR-2 activation that include altering ion levels, particularly Zn2+, and prostate expressed proteinaceous inhibitors, are key for expanding the global understanding of in vivo KLK4/PAR interactions.

6.5 PARs as therapeutic targets in prostate cancer

At present ~30% of all pharmaceuticals target GPCRs (Hopkins and Groom, 2002), such as the highly successful drugs Zyprexa and Zoloft (Schlyer and Horuk, 2006). This group of drugs selectively modifies GPCR behavior through full or partial receptor agonism or receptor antagonism (Hopkins and Groom, 2002). Thus, it may be possible to selectively target PARs in disease processes. In this respect, a number of small non-peptidic antagonists have been developed for PAR-1 (Barry et al., 2006) and have been effective in inhibiting thrombin mediated platelet aggregation (Zhang et al., 2001) and demonstrated antithrombotic effects in an arterial thrombosis injury model (Derian et al., 2003). Importantly, application of PAR-1 antagonists suppressed both in vitro angiogenesis in a matrigel tube formation assay and in vivo in the chick chorioallantoic membrane model (Zania et al., 2006). Further, small peptidic antagonists for PAR-4 have demonstrated effective reduction of thrombin induced endostatin release from platelets (Ma et al., 2001) and reduced the infarct size in rat models of myocardial ischemia/reperfusion (Strande et al., 2008). The success of PAR-1 non-peptidic antagonists in inhibiting angiogenesis in vivo, an in vitro model of thrombin induced colon carcinoma (Heider et al., 2004)

131 and hepatocellular carcinoma cell migration (Kaufmann et al., 2007), as well as a PAR-4 peptidic antagonist reducing in vitro hepatocellular carcinoma cell migration (Kaufmann et al., 2007), support the potential for PARs as a target for antagonism in the treatment of prostate cancer. Although at present no selective antagonist has been developed for PAR-2, preliminary data has indicated limited success with antagonists derived from the partially reversed PAR-1 and PAR-2 AP sequences (Al-Ani et al., 2002). As the PARs display a diverse tissue expression, selective targeting of antagonists to tumour-expressed PARs would be challenging. A phenomenon which may potentially be used to discriminate tumour cells from their healthy counterparts, is alterations in PAR glycosylation due to carcinogenesis

It has been known for decades that the structure of glycans at the cell surface changes with the onset of carcinogenesis. In 1969, Meezan and colleagues undertook comparative analysis of surface sugar chains of mouse 3T3 fibroblasts and their simian virus (SV-40) transformants, reporting a shift to larger molecular size in the surface glycoproteins of the transformed cell line (Meezan et al., 1969). More recent studies have identified a range of glycan alterations amongst cancer-associated cell surface glycoproteins using monoclonal antibodies and mass spectrometry (Dube and Bertozzi, 2005).

Carbohydrate based anti-cancer vaccines represent a therapeutic area gaining increasing amounts of interest. This research is largely focused on two areas; immunotherapy using antibodies specific to cancer associated glycans and prevention of the formation of cancer glycans (Dube and Bertozzi, 2005). Application of these technologies may have relevance to therapeutic development against PARs if carcinogenesis associated glycan alteration occurs. Furthermore, combination of current small molecular weight PAR antagonist development (Andrade-Gordon et al., 1999; Berger, 2000; Ma et al., 2001) and the ability to target cancer specific glycans holds great potential to selectively deliver and disrupt PAR activation on the surface of cancer cells.

132 6.6 Future directions

Future directions for research based on the results of this study have been proposed to address several key areas that hold great potential for delineating further the contribution of KLK4 activation of PARs to prostate cancer progression.

There are a number of experiments that could more fully interrogate the consequences of KLK4 activation of PARs for prostate cancer progression. These would include in vitro assays, employing prostate cancer cells modulated for PAR-1 and/or PAR-2 expression, monitoring changes in the cancer associated cellular functions proliferation, migration, invasion, apoptosis and colony formation following KLK4 activation of PARs. To complement in vitro findings, in vivo experiments utilising mouse models could be performed. For example, using a SCID mouse model to assess primary tumour formation and spontaneous metastasis of subcutaneous matrigel implanted prostate cancer cells following KLK4 treatment would be beneficial. Further, intra-tibial injection of prostate cancer cells into PAR-1 -/- and PAR-2 -/- mouse models and assessment of tumour formation in each knockout model in comparison to wild-type mice would help to determine the importance of these receptors in prostate cancer bone metastasis. PAR-1 -/- and PAR-2 -/- mice would first need to be crossed into an immune-compromised mouse strain. Importantly, use of blocking antibodies and antagonists targeting PAR-1 and PAR-2 in in vitro and in vivo experiments has the potential to discriminate the contribution of each of these receptors to prostate cancer progression.

To more clearly understand the alterations in cellular functions observed in vivo and in vitro following KLK4 activation of prostate cancer cell expressed PARs, parallel studies identifying differential regulation of RNA and proteins should be carried out. One approach to identify changes in RNA expression would be to examine by microarray analysis KLK4 activated (against untreated) prostate cancer cell lines modulated for PAR expression. Further, analysing the same samples by proteomic approaches (eg. two-dimensional gel electrophoresis followed by mass spectroscopy sequence analysis) can relate those changes identified at the RNA level to differential protein expression. Critically, comparative experiments that analyse prostate cancer cells or tissue treated with PAR-1 antagonist/blocking antibody in comparison to

133 samples treated with PAR-2 antagonist/blocking antibody has the potential to define the signaling pathways each receptor initiates and define functional pathways and gene targets not currently known to be modulated by the PARs.

A key observation of this study was the higher potency of KLK4 activation via PAR- 1 than via PAR-2. In order to define the potency differences in greater detail, an important initial experiment would be to establish the amounts of PAR-1 and PAR-2 expressed at the surface of cells and to use these results as a normalisation baseline for further concentration response experiments. Further, peptide cleavage studies, similar to the approach utilised by Oikonomopoulou and colleagues (Oikonomopoulou et al., 2006), to identify PAR-1 and PAR-2 N-terminus cleavage sites at increasing KLK4 concentrations would identify cleavage of each N-terminus at sites which would disarm the receptor and potentially explain differences in KLK4 potency. In addition, bioinformatics approaches to examine the presence or absence of motifs in PAR-1 and PAR-2 that function as structural determinants for KLK4 activation may potentially identify domains that control the potency of KLK4 activation, in a manner similar to the hirudin-like domain of PAR-1 enhancing the potency of thrombin activation.

In contrast to potency differences, the efficacy of KLK4 activation via the receptors, as assessed by Ca2+ mobilisation, was greater through PAR-2 than PAR-1. Similar to potency analyses, using the amounts of each receptor expressed at the cell surface as a normalisation baseline in further concentration response experiments would be an important initial step. Further, assessing the efficacy of KLK4 activation of each receptor in relation to several other intracellular signaling pathways (eg. Rho and Cyclic adenosine monophosphate), would determine if efficacy differences are conserved across multiple signaling pathways. Additionally, an expansive study examining the activation of the PARs by all prostatic expressed trypsin-like KLKs in parallel would determine if activation redundancies and/or differences exist between KLK family members and may provide further insight into a prostate KLK cascade that ends in PAR activation.

134 6.7 Summary

In summary, the work presented in this thesis has defined new signal transduction events initiated by KLK4 and transduced at the cell surface by PARs. These activation events have potential relevance to prostate cancer progression as KLK4 and PAR-1/PAR-2 show increased expression and are co-expressed by similar cell types during progression of this disease. Importantly, further delineation of the consequences of these events may lead to exciting new possibilities in therapeutic approaches to prostate cancer and bone metastasis.

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