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SPARC enhances chemosensitivity by activating the extrinsic pathway of apoptosis Tang, Michelle Jocelyn 2009

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SPARC ENHANCES CHEMOSENSITIVITY BY ACTIVATING THE EXTRINSIC PATHWAY OF APOPTOSIS  by MICHELLE JOCELYN TANG B.Sc. Hon., The University of British Columbia, 2004  A THESIS SUBMI'T'TED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in  THE FACULTY OF GRADUATE STUDIES (Experimental Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  December 2009 © Michelle Jocelyn Tang, 2009  Abstract Colorectal cancer (CRC) is the second leading cause of cancer death in Canada. Treatment failure in advanced disease is due to the development of chemotherapy resistance. Using a genomics approach, the Tai laboratory previously found that Secreted Protein Acidic and Rich in Cysteine (SPARC), a matricellular protein, was down-regulated in chemotherapy resistant CRCs. SPARC has tumour suppressor properties in ovarian and pancreatic cancers, where higher SPARC expression enhances apoptosis in vitro and tumour regression in vivo. In CRCs, it was found that exogenous SPARC re-sensitized CRCs to chemotherapy by inducing apoptosis and inhibiting tumour growth. In this thesis, the mechanisms of SPARC-mediated apoptosis, the fragment of SPARC responsible for SPARC-mediated apoptosis and the interaction between SPARC and collagen IV and its effect on chemosensitivity were examined. I demonstrated that SPARC can be internalized and interacts with the N-terminus of the procaspase 8 death effector domain (DED)-containing domain, activating the extrinsic pathway of apoptosis. This pro-apoptotic activity is mediated through SPARC’s N-terminus domain, where the N-terminus not only enhanced apoptosis in vitro, but also augmented tumour regression in vivo in combination with chemotherapy. The N-terminus domain of SPARC also interfered with the interaction between Bcl-2 and procaspase 8, decreased cell viability and increased apoptosis. I also found that higher SPARC levels correlated with higher caspase 8 expression. Further, I provided evidence that collagen IV and SPARC work in a co-operative manner to enhance apoptosis. This thesis focuses on the interplay between SPARC/Bcl2/procaspase 8 and collagen IV which may be important in modulating a tumour’s response to chemotherapy in CRCs, suggesting that SPARC may be a potential cancer therapeutic.  ii  Table of Contents Abstract........................................................................................................................................... ii Table of Contents........................................................................................................................... iii List of Tables ................................................................................................................................. vi List of Figures............................................................................................................................... vii List of Abbreviations ...................................................................................................................... x Acknowledgments ....................................................................................................................... xiii Co-Authorship Statement ............................................................................................................ xiv 1. Introduction ................................................................................................................................ 1 1.1 Cancer ................................................................................................................................... 1 1.2 Colorectal cancer .................................................................................................................. 1 1.3 Chemotherapy....................................................................................................................... 3 1.4 Drug resistance ..................................................................................................................... 5 1.5 The extracellular matrix........................................................................................................ 7 1.7 SPARC................................................................................................................................ 10 1.7.1 The domains of SPARC .............................................................................................. 10 1.7.2 SPARC expression ...................................................................................................... 11 1.7.3 Functions of SPARC ................................................................................................... 12 1.7.4 SPARC in cancer ......................................................................................................... 12 1.7.5 SPARC in colorectal cancer ........................................................................................ 13 1.8 Mechanistic aspects of SPARC action in cancer biology................................................... 15 1.8.1 Hypermethylation of the SPARC promoter ................................................................. 15 1.8.2 Apoptosis ..................................................................................................................... 15 1.8.3 Angiogenesis, invasion and metastasis........................................................................ 16 1.9 Apoptosis, autophagy, and necrosis ................................................................................... 16 1.9.1 What is apoptosis? ....................................................................................................... 17 1.9.2 Mechanism of apoptosis .............................................................................................. 18 1.9.3 The extrinsic pathway of apoptosis ............................................................................. 19 1.9.4 The intrinsic pathway of apoptosis .............................................................................. 20 1.9.5 Regulators of apoptosis ............................................................................................... 21 1.9.6 The pro-survival pathways .......................................................................................... 21 1.10 Research objectives .......................................................................................................... 22 2. Materials and methods.............................................................................................................. 32 2.1 Cell lines, reagents.............................................................................................................. 32 2.2 Site-directed mutagenesis ................................................................................................... 33 2.3 Coating of plates with collagen IV ..................................................................................... 33 2.4 ELISA for quantification of SPARC or related peptides ................................................... 34 2.4.1 ELISA to quantify the localization of SPARC ............................................................ 34 2.5 RT-PCR .............................................................................................................................. 35 2.6 Chromatin immunoprecipitation (ChIP) assay ................................................................... 36 2.7 RNA interference................................................................................................................ 36 2.8 Cell viability ....................................................................................................................... 37 2.8.1 RNA Interference ........................................................................................................ 37 2.8.2 Caspase 8/9 Inhibition ................................................................................................. 37 2.8.3 SPARC domains .......................................................................................................... 37 2.8.4 SPARC localization ..................................................................................................... 38 2.8.5 Collagen IV.................................................................................................................. 38 iii  2.8.6 Cellular number ........................................................................................................... 39 2.9 Cell proliferation................................................................................................................. 39 2.10 Caspase 3/7 assays............................................................................................................ 39 2.10.1 RNA interference....................................................................................................... 39 2.10.2 SPARC localization ................................................................................................... 40 2.10.3 Collagen IV studies ................................................................................................... 40 2.11 TUNEL Assays................................................................................................................. 40 2.12 Immunoblot analysis......................................................................................................... 41 2.13 Subcellular fractionation and immunoprecipitation ......................................................... 41 2.13.2 SPARC:caspase 8 interaction .................................................................................... 41 2.13.2 Specificity of the SPARC:caspase 8 interaction........................................................ 42 2.14 Animal studies .................................................................................................................. 43 2.14.1 Tumor xenografts ...................................................................................................... 43 2.14.2 Immunohistochemistry .............................................................................................. 44 2.15 Statistics............................................................................................................................ 44 3. A novel interaction between pro-caspase 8 and SPARC enhances apoptosis and ................... 53 potentiates chemotherapy sensitivity in colorectal cancers .......................................................... 53 3.1 Introduction ........................................................................................................................ 53 3.2 Results ................................................................................................................................ 55 3.2.1 Over-expression of SPARC in MIP101 cells augment the expression of genes involved in the extrinsic pathway of apoptosis .................................................................... 55 3.2.2 Cells over-expressing SPARC have enhanced activation of the extrinsic pathway of apoptosis ............................................................................................................................... 56 3.2.3 In the presence of SPARC, caspase 8 decreases cell viability in response to 5-FU.... 57 3.2.4 Absence of caspase 8 diminishes apoptosis following chemotherapy in high SPARCexpressing cells..................................................................................................................... 59 3.2.5 Is the activity of SPARC intracellular or extracellular? .............................................. 60 3.2.6 SPARC interacts with pro-caspase 8 ........................................................................... 62 3.2.7 Specificity of the SPARC: caspase 8 interaction......................................................... 63 3.2.8 SPARC in combination with 5-FU increases caspase 8 expression in tumour xenografts ............................................................................................................................. 64 3.3 Discussion........................................................................................................................... 64 4. SPARC’s N-terminal domain interferes with Bcl-2 and procaspase 8 interaction and promotes apoptosis and tumour regression in vivo ...................................................................................... 87 4.1 Introduction ........................................................................................................................ 87 4.2 Results ................................................................................................................................ 89 4.2.1 Effect of different SPARC domains on cell viability .................................................. 89 4.2.2 SPARC-N confers increased sensitivity to chemotherapy .......................................... 89 4.2.3 Effect of SPARC domains on cell proliferation and apoptosis ................................... 90 4.2.4 Tumour xenografts over-expressing SPARC and SPARC-N are more chemosensitive...................................................................................................................... 92 4.2.5 The N-terminus of SPARC interacts with procaspase 8.............................................. 93 4.2.6 SPARC prevents the interaction between Bcl-2 and caspase 8................................... 93 4.2.7 Mutations in the DED domains of procaspase 8 prevent interactions with Bcl-2 and SPARC.................................................................................................................................. 94 4.3 Discussion........................................................................................................................... 96 5. SPARC interacts with collagen IV to re-sensitize resistant colorectal cancer cells ............... 122 5.1 Introduction ...................................................................................................................... 122 5.2 Results .............................................................................................................................. 124 iv  5.2.1 Collagen IV increases sensitivity of colorectal cancer cells to chemotherapy by decreasing cell viability and cell proliferation.................................................................... 124 5.2.2 Collagen IV increases chemosensitivity through the mitochondrial pathway of apoptosis. ............................................................................................................................ 125 5.2.3 Collagen IV further enhances chemosensitivity of SPARC biological fragments .... 126 5.2.4 SPARC enhances the chemosensitizing effects of collagen IV through the extracellular domain of SPARC ......................................................................................... 129 5.3 Discussion......................................................................................................................... 130 6. Discussion............................................................................................................................... 145 6.1 Review of findings............................................................................................................ 145 6.2 Future directions ............................................................................................................... 148 6.3 Significance ...................................................................................................................... 150 6.4 Conclusions ...................................................................................................................... 151 7. References .............................................................................................................................. 152 Appendix I: Biosafety Certificate............................................................................................... 173 Appendix II: Ethics Certificates ................................................................................................. 174  v  List of Tables Table 2-1. Primer sequences for site-directed mutagenesis primers .............................................46 Table 2-2: Amino acid mutation sequences used in this thesis .....................................................47 Table 2-3. Primer sequences used for RT-PCR.............................................................................49 Table 2-4.ChIP Assay buffers. ......................................................................................................51 Table 3-1 Homology between caspase 8 and other initiator caspases...........................................83  vi  List of Figures Figure 1-1. Illustration of colorectal tumour progression in sporadic and high-risk genetic syndromes. .............................................................................................................................24 Figure 1-2. The progression of colorectal cancer with chromosomal instability (CIN)................25 Figure 1-3. The progression of colorectal cancer with CpG Island methylator phenotype (CIMP).26 Figure 1-4. Intracellular activation of 5-Fluorouracil (5-FU)........................................................27 Figure 1-5. The functional domains of SPARC are represented. ..................................................28 Figure 1-6. The structural domains of SPARC are represented ....................................................29 Figure 1-7. Characteristic morphological features of apoptosis....................................................30 Figure 1-8. A simplified schematic representation of the apoptotic cell signaling pathways. ......31 Figure 2-1. Diagram of constructs used in this thesis...................................................................45 Figure 2-2. Levels of SPARC, SPARC related peptides used in this study. ................................50 Figure 2-3. Efficacy of caspase 8 siRNA on gene silencing. ........................................................52 Figure 3-1. The presence of higher levels of SPARC is associated with greater expression of genes involved in the extrinsic pathway of apoptosis following exposure to 5-FU..............69 Figure 3-2. The presence of higher levels of SPARC is associated with greater levels of apoptosis. ...............................................................................................................................71 Figure 3-3. Inhibition of caspase 8 increases cell viability in CRC cells with greater SPARC expression. .............................................................................................................................72 Figure 3-4. Inhibition of caspase 8 gene expression with siRNA enhances survival of cells expressing higher levels of SPARC following exposure to 5-FU. ........................................75 Figure 3-5. ELISAs of intracellular and extracellular SPARC clones. .........................................78 Figure 3-6. Intracellular SPARC enhances chemosensitivity.......................................................79 Figure 3-7. Extracellular SPARC does not enhance chemosensitivity..........................................80 vii  Figure 3-8. Interaction between pro-caspase 8 and SPARC is detected in the cell membrane fraction...................................................................................................................................81 Figure 3-9. The interaction between pro-caspase 8 and SPARC occurs at the N-terminus of procaspase 8. ...............................................................................................................................82 Figure 3-10. SPARC interacts specifically with caspase 8 ...........................................................84 Figure 3-11. Caspase 8 is up-regulated in response to higher levels of SPARC in tumour xenografts following exposure to 5-FU in vivo.....................................................................85 Figure 3-12: SPARC-induced apoptosis: A model........................................................................86 Figure 4-1. Over-expression of the N-terminal domain of SPARC diminished cell viability in cancer cell lines. ..................................................................................................................101 Figure 4-2. Mutations in the N-terminal domain of SPARC inhibit the chemosensitizing effects of SPARC colorectal cancer cell lines.................................................................................103 Figure 4-3 The effect of SPARC mutations on cell viability. .....................................................105 Figure 4-4. Effect of SPARC-domains on cell proliferation and apoptosis in response to chemotherapy.......................................................................................................................106 Figure 4-5. The effect of caspase 8 silencing in colorectal cancer cells over-expressing the SPARC-domains on apoptosis in response to chemotherapy..............................................108 Figure 4-6. Xenografts over-expressing SPARC-N have greater tumour regression in response to 5-FU.....................................................................................................................................109 Figure 4-7. MIP/SP-N xenografts exhibit greater levels of apoptosis and lower levels of CD31110 Figure 4-8. The site of SPARC:pro-caspase 8 interaction is at the N-terminus of SPARC ........112 Figure 4-9. SPARC and Bcl-2 both interact with pro-caspase 8 with opposing effects..............114 Figure 4-10. Immunoprecipitation of MIP/5FU cells with SPARC peptides..............................116 Figure 4-11. The DEDI domain of caspase 8 is critical for its interaction with Bcl-2................117 Figure 4-12. The DEDI domain of caspase 8 is critical for its effect on cell viability................118 Figure 4-13. Interaction between the N-terminus of both pro-caspase 8 and SPARC are required to reduce cell viability and enhance caspase 3/7 activity. ...................................................119 viii  Figure 4-14: A model: SPARC-mediated apoptosis....................................................................121 Figure 5-1. Collagen IV decreases cellular viability of MIP101 colorectal cancer cells ............135 Figure 5-2. Collagen IV increases the doubling time of MIP101 colorectal cancer cells ...........136 Figure 5-3. The effect of collagen IV on apoptosis. ....................................................................137 Figure 5-4. The effect of collagen IV on the cell signaling pathway of apoptosis......................138 Figure 5-5. Collagen IV enhances chemosensitivity in MIP101 cells over-expressing the various biological domains of SPARC.............................................................................................139 Figure 5-6. Collagen IV increases the doubling time of MIP101 cells over-expressing the various biological domains of SPARC.............................................................................................140 Figure 5-7. Collagen IV enhances chemosensitivity by up-regulating the apoptotic pathway of MIP101 cells over-expressing the various biological domains of SPARC. ........................141 Figure 5-8. The SP-C domain of SPARC influences collagen IV's effect on chemosensitivity. 142 Figure 5-9. The SP-C domain of SPARC and collagen IV co-operatively decrease apoptosis. .143 Figure 5-10. A schematic representation of the cellular signaling events involved when grown on collagen IV. .........................................................................................................................144  ix  List of Abbreviations 5-FU Aa ABC Apaf-1 bFGF ChIP CIMP CIN CIS CIV co-IP CPT-11 CRC CSF DEDI DEDII DEDIIm DEDIm DISC DMEM DNA DR EC ECM ELISA ER Eto FADD FGF Fmk FS GM HNPCC IAP IFN IL IMS IRF LPA MCF7/CIS MDR  5-fluorouracil Amino acid ATP binding cassette Apoptosis protease-activating factor 1 Basic fibroblastic growth factor Chromatin immunoprecipitation CpG island methylator phenotype chromosomal instability Cisplatin Collagen IV co-immunoprecipitation Irinotecan Colorectal cancer colony-stimulating factor Death effector domain I of caspase 8 Death effector domain II of caspase 8 Mutant of the death effector domain II of caspase 8 Mutant of the death effector domain I of caspase 8 Death-inducing signaling complex Dulbecco’s modified eagle’s medium Deoxyribonucleic acid Death Receptor Extracellular domain of SPARC extracellular matrix Enzyme-Linked ImmunoSorbent Assay Endoplasmic Reticulum Etoposide Fas-associated death domain Fibroblast growth factor Fluoromethylketone Follistatin-like domain of SPARC granulocyte macrophage Hereditary Non-polyposis colorectal cancer Inhibitor of Apoptosis Interferon Interleukin intermembrane space interferon regulatory factor lysophosphatidic acid MCF7 cells resistant to cisplatin multi-drug resistance x  MiaPaca/CPT MIN MIP/5FU MIP/SP MIP/SP-C MIP/SP-F MIP/SP-N MIP/ZEO MMP MMR mRNA NER NSAID NT PAGE PB PBm PBS PCR PDGF PI3K polII PVDF RKO/5FU RKO/CPT RLU RNA rSPARC RT s.e. SAM SDS siRNA SMOC SP SP1 SPARC SPARC-/SPARC+/+ SP-C SP-F  MiaPaca cells resistant to irinotecan microsatellite instability MIP101 cells resistant to 5-fluorouracil MIP101 cells over-expressing SPARC MIP101 cells over-expressing the EC-domain of SPARC MIP101 cells over-expressing the follistatin-like domain of SPARC MIP101 cells over-expressing the N-terminal domain of SPARC MIP101 cells stably-transfected with empty vector Matrix metalloproteinases mismatch repair messenger RNA nucleotide excision repair Non-steroidal anti-inflammatory drug N-terminal domain of SPARC polyacrylamide gel electrophoresis Putative binding domain of caspase 8 Mutations in the putative binding domain of caspase 8 Phosphate buffered saline Polymerase Chain Reaction platelet derived growth factor Phosphatidylinosital 3-Kinase RNA polymerase II Polyvinylidene Fluoride RKO cells resistant to 5-Fluorouracil RKO cells resistant to irinotecan Relative Luminescence unit(s) ribonucleic acid Recombinant human SPARC reverse transcription standard error S-adenosylmethionine sodium dodecyl sulfate Small interfering RNA SPARC-related modular calcium binding Full length SPARC specificity factor 1 Secreted protein acidic and rich in cysteine SPARC knock-out SPARC wild-type Plasmid expressing the extracellular domain of SPARC Plasmid expressing the follistatin-like domain of SPARC  xi  SP-Fmut1 SP-Fmut2 SP-N SP-Nmut1 SP-Nmut2 SRP STAT TNF TNM TRAIL TUNEL VDR VEGF Z  Mutant 1- plasmid expressing a mutant follistatin-like domain of SPARC Mutant 2 - plasmid expressing a mutant follistatin-like domain of SPARC Plasmid expressing the N-terminal domain of SPARC Mutant 1- plasmid expressing a mutant N-terminal domain of SPARC Mutant 2 - plasmid expressing a mutant N-terminal domain of SPARC Signal Recognition Peptide Signal Transducers and Activators of Transcription tumour-necrosis factor Tumour-node-metastasis TNF-related apoptosis-inducing ligand Deoxynucelotidyltransferase-mediated dUTP nick end labeling Vitamin D Receptor Vascular Endothelial Growth Factor Benzyloxycarbonyl  xii  Acknowledgments I would like to thank every single person that supported me through my challenging experience. I am extremely grateful to my supervisory committee, Dr. Vince Duronio, Dr. Michael Cox, Dr. Keith Humphries, and Dr. Marianne Sadar for all of their support. I am extremely grateful for salary and travel funding for the Canadian Digestive Health Foundation, Canadian Institutes for Health Research, and National Cancer Institute of Canada. I would like to thank Dr. Isabella Tai for allowing me the opportunity to perform this research in her laboratory. I would not have been able to do the research in my thesis without the help and advice of others at the Genome Science Centre, University of British Columbia, and the BC Cancer Research Centre, and thank all that have impacted my research. I would also like to thank my undergraduate students Ariella Zbar, Johnny Nie, Justin Chan, Winnie So, and Shirley Ho for their hard work and for the opportunity for me to help guide them in their studies. I would also like to thank all members from the Tai laboratory, for who have undoubtedly helped me throughout the years. On a personal note, I would like to thank David Tiedje for being such a supportive and understanding person. I also would like to thank my family for supporting me, teaching me, and helping me though my journey. Most importantly, I thank my parents for whom without them; I could not have achieved this goal.  xiii  Co-Authorship Statement Chapter 1 contains modified information from the following review: Tai, I.T. and Tang, M.J. “SPARC in cancer biology: its role in cancer progression and potential for therapy.” Drug Resistance Updates (2008) 11 (6): 231-46. I was responsible for generating an initial draft, data and figures. I.T. Tai was responsible for the writing, editing, and initial drafts of the figures. Chapter 3 contains portions from a manuscript published in the Journal of Biological Chemistry: Tang, M.J. and Tai, I.T. “A novel interaction between procaspase 8 and SPARC enhances apoptosis and potentiates chemotherapy sensitivity in colorectal cancers.” Journal of Biological Chemistry 2007; 282 (47): 34457-67. I was the primary author of Tang and Tai (2007) and was responsible for generating all of the experimental data, analysis, text, figures and tables. IT Tai contributed to experimental design and intellectual input of this manuscript. Also, SM Cheetham contributed by aiding in the performance of blinded animal studies in Chapter 4.  xiv  1. Introduction 1.1 Cancer Cancer is often described as the progression of normal cells to malignant derivatives. The development of cancer is a multi-step process that involves genetic alterations which affect cellular proliferation and homeostasis. There are many characteristics of cancer cells that include: self-sufficiency in growth signals, insensitivity to growth-inhibitory signals, evasion of apoptosis, limitless replicative potential, sustained angiogenesis, and tissue invasion and metastasis. Each of these changes are acquired during tumor development and involve changes in gene expression (1). Genes that are involved in cancer are often defined as oncogenes. An oncogene is a gene, where when it is mutated or highly expressed, aids in the progression of a normal cell into a tumor cell (2). A proto-oncogene is a normal gene that can become an oncogene due to mutations or increased expression. Proto-oncogenes often code for proteins that regulate cell growth and differentiation, and are often involved in signal transduction (3). Alternatively, a tumor suppressor gene is a gene that protects a cell from the progression to cancer. If this gene is mutated and the mutation causes a loss of function, the cell can progress to cancer. Tumor suppressor genes usually have a repressive effect on the regulation of the cell cycle or promotes apoptosis, or a combination (4). In this thesis, we extensively study colorectal cancer, which will be discussed in greater detail below. 1.2 Colorectal cancer Colorectal cancer (CRC) is the second leading cause of cancer death in Canada, with an estimated 21,500 diagnosed cases and 8,900 deaths in 2008 (5). There are three main categories of CRC: sporadic (~80%, caused by environmental factors including diet, exercise, and smoking) and inherited (hereditary nonpolyposis, <10% and familial adenomatous polyposis, 6%) (6). In 1  the majority of CRC cases, the transformation from normal colonic epithelium to colon adenocarcinoma is a result of numerous genetic events in key oncogenes and tumour suppressors (Figure 1-1) (7). These genetic events cause mutations that create a growth advantage, leading to an increase in malignant cells and eventually invasive adenocarcinoma. CRC development typically involves the following key genes: adenomatous polyposis coli (APC), K-ras, deleted in colon cancer (DCC) genes, and p53 (7). Mutations of the APC gene often initiate CRC tumourigenesis; however, multiple mutations are required for carcinoma development. The various genes involved affect many different pathways that influence CRC development. Examples of genetic pathways involved the development of CRC include the chromosomal instability (CIN) and microsatellite instability (MIN) pathways. The CIN pathway, considered typical of sporadic CRC development, exhibits chromosomal abnormalities (eg. aneuploidy) and sequential inactivation of tumour suppressor genes (eg. APC, p53, and SMAD4) (Figure 1-2) (8-10). Unlike the CIN pathway, CRCs developed via the MIN pathway do not have extensive chromosomal abnormalities and are instead, characterized by defects in mismatch repair (MMR) genes during DNA replication. (11-13). CRCs also arise from regional hypermethylation at CpG islands, causing transcriptional inactivation of genes involved in tumour progression. Examples of genes involved in CpG island methylator phenotype (CIMP) are p16, THBS1, IGF2, hMLH1 (Figure 1-3). After diagnosis of CRC, the tumour is staged by the tumour-node-metastasis (TNM) system. The TNM system characterizes the disease based on the depth of invasion in the bowel wall (T1: invasion of the submucosa to T4: invasion into adjacent structures), regional lymph node involvement (N0: no lymph nodes involved, N1: 1-3 lymph nodes involved, N2: >3 lymph nodes involved (14-18)), and presence of disease at distant sites (M1: presence of metastases at distant site) (19-21). The most common site of CRC metastasis from the colorectum is the liver; other organs affected include the lungs, peritoneum, and intra-abdominal lymph nodes (22). 2  Early detection of CRC continues to improve due to the promotion of screening tests, polypectomies, and changes in diet and lifestyle (23). Screening tests are recommended to patients based on age, malignancy in family history, CRC or colorectal adenoma diagnosis, and medical history of non-specific inflammatory bowel disease (24). After CRC diagnosis, treatment classically involves surgery, chemotherapy and radiation therapy. The treatment regimen of CRC is dependent on many factors such as the stage of the disease and the genetics of the tumour (25). Treatment regimens for malignancies generally include one of the following options: a combination of chemotherapy and surgery, chemotherapy and radiation, surgery and/or radiation followed by chemotherapy (adjuvant therapy) or chemotherapy followed by surgery and/or radiation (neoadjuvant therapy). This thesis focuses on the use of chemotherapy drugs in CRC treatment and will be discussed in further detail below. 1.3 Chemotherapy Chemotherapy, sometimes referred to as systemic therapy, is defined as treatment with drugs. These cytotoxic chemotherapeutic agents induce DNA damage and disrupt cellular targets within specific stages of the cell cycle. Thus, the efficacy of chemotherapy often corresponds to the proliferation rate of the tumour. Chemotherapy drugs can be divided into cell cycle phase specific and non-cell-cycle-phase specific categories. Cell cycle phase specific are for those that target a specific cell cycle phase, such as the G1 or S phase, where non-cell-cycle phase specific drugs are effective at any point of the cell cycle. The vast number of cells present in a rapidly dividing tumour allows for greater efficacy of cell cycle phase-specific drugs, while non-cellcycle-phase-specific drugs are more commonly used for large tumours and those with a low growth fraction. (26). Cell cycle phase-specific and non-cell-cycle-phase specific chemotherapy drugs are further categorized by their specific mechanism of action. Examples of the more specific categories include alkylating agents, anti-metabolites, anti-tumour antibiotics, 3  topoisomerase inhibitors, mitotic inhibitors, and monoclonal antibodies. The three categories discussed below are commonly used in CRC treatment (26). 1) Alkylating agents: Alkylating agents are highly reactive electrophiles that form covalent bonds with nucleophilic groups (eg. amino, carboxyl, phosphate, and sulfhydroxyl). The resulting covalent bonds cause alterations in DNA structure (eg. inter- and intra-strand cross-links), disrupting normal DNA replication, and ultimately leading to cell death. Examples of drugs in this category include mechlorethamine, cisplatin, mitocyin C, altretamine and busulfane (26). 2) Anti-mitotic agents: Anti-mitotic agents are further divided into two sub-categories: i) Topoisomerase inhibitors: As the name indicates, these drugs prevent the function of topoisomerases. DNA topoisomerases bind to supercoiled DNA, during replication or repair of the DNA to maintain cellular viability in the cell cycle. Topoisomerase inhibitors stabilize the topo:DNA complex which disrupts regulation of the cell cycle. Irinotecan and etoposide are examples of topoisomerase inhibitors (26, 27). ii) Anti-microtubule agents: These drugs de-polarize microtubules, which are major cytoskeletal structures of the cell. Drugs included in this group are: vincristine, vinblastine, and vinorelbine (26). 3) Anti-metabolites: Unlike the previous two chemotherapy groups, anti-metabolites differ require metabolic activation before being able to exert its effects. After activation, the drug enters the targeted metabolic pathway and disrupts RNA and DNA synthesis. Many of these drugs resemble substrates that are part of the normal metabolic processes. An example of a drug in this category is a pyrimidine analogue, 5-fluorouracil (5-FU). The intracellular activation of 5FU allows for incorporation into DNA and RNA resulting in the disruption of DNA or RNA synthesis, respectively (Figure 1-4). Other examples include methotextrate and hydroxyurea (26). 4  There are many other categories of drugs used for cancer therapy, such as hormone and enzymatic therapies. New therapeutics including differentiation agents, angiogenesis inhibitors (eg. Bevacizumab (25)), matrix metalloproteinase inhibitors, and signal transduction agents (eg. Cetuximab (25)) are also currently under study (26). Although there are a variety of cancer therapies, drug resistance still remains an obstacle in the path of cancer treatment. 1.4 Drug resistance Treatment of cancer with chemotherapy is only effective if the cytotoxic agents can inhibit cancer cell growth. Unfortunately, drug resistance – which can be inherited (intrinsic resistance) or acquired (extrinsic resistance) strongly hinders the efficacy of chemotherapies (28, 29). Several cellular mechanisms can lead to drug resistance including: 1) Altered membrane transport: Proteins belonging to the ATP Binding Cassette (ABC) transport protein family, alter absorption, distribution, and excretion of chemotherapy drugs, which lowers the intracellular levels of chemotherapy drugs below cell-killing threshold. Although a few transport proteins have been identified, the exact mechanisms of chemotherapy drug uptake is unknown (28) ABC proteins have also been associated with multidrug resistance (MDR). An intriguing characteristic of MDR related proteins is their loose substrate specificity, something that is not seen in other mammalian transporters. This loose substrate specificity causes a cross-resistance to several chemically unrelated compounds, a hallmark of the MDR phenotype. MDR is commonly observed with anti-metabolite drugs (28, 30, 31). 2) Genetic Responses: There are several genetic events that contribute to drug resistance. A few examples include altering genes involved in regulating the ABC transport family, 5  gene rearrangements to increase or decrease gene targets, or defects in the apoptotic pathway. Additionally, as anti-metabolites require metabolic activation prior to exerting their cytotoxic effects, enzymes that inhibit activation decrease their cytotoxic efficacy and activity. This decreases the level of active drug and allows the cancer cell to overcome the cytotoxic effects (28, 31, 32). 3) Enhanced DNA Repair: In this mechanism, the cancer cell is able to repair DNA damage induced by chemotherapy drugs through DNA repair proteins, ultimately reversing the cytotoxic effect. Examples of DNA repair genes involved in drug resistance are DNA MMR genes (eg. hMLH1, hMSH2), xeroderma pigmentosum group E binding factor and excision repair cross-complementing protein (28, 31). 4) Metabolic effects: The two main mechanisms that lead to inactivation of chemotherapy drugs by enzymes decrease the amount of free drug available and levels of cell death. The first mechanism works by over-expressing drug-metabolizing enzymes, which in turn, cause cells to rapidly metabolize and detoxify the cytotoxic drug. The second mechanism works with drugs that require metabolic activation; under-expression of drugmetabolizing enzymes leads to a reduction of the levels of active drug within the cell. Examples that decrease the level of cytotoxic drug include cytochrome P450 and glutathione S-transferase (28, 31). This thesis focuses on the use of 5-FU, an anti-metabolite, as a cytotoxic. The mechanism of action of 5-FU is thought to occur through its inhibition of thymidylate synthase (TS) (Figure 1-4) (31). Numerous studies have demonstrated that TS expression is a key determinant of 5-FU sensitivity (33). Interestingly, a proof-of-principle study showed that MIP101 cells resistant to 5-FU expressed various isoforms of several genes compared to its parental counterpart (34). It would be interesting to determine if these various isoforms affected TS activity; thus contributing to the resistant phenotype. 6  5) Alterations in target proteins: Some cytotoxic drugs exert their effects by targeting specific proteins. Alteration of target protein expression levels or mutation of the target protein can have a major impact on drug resistance. For example, during the course of anti-estrogen treatment for breast cancer, some patients experience a decrease in the level of the targeted estrogen receptor protein; thus, developing a resistant phenotype. Alternatively, mutation of the target protein allows it to retain its activity, but no longer binds to the drug. Thus, the drug no longer inhibits the protein. Drug resistance may also occur through loss of target molecule, which can occur through chromosomal losses (28, 31). 6) Growth factors: Drug resistance can also be attributed to extracellular factors in tumourbearing organs, where production of growth factors provides a growth advantage (26, 31, 35). Increased serum levels of growth factors that contribute to tumour growth have been found in various cancers. For example, human fibroblast growth factor 1 (FGF-1) contributes to tumourigenesis due to its function in development, morphogenesis, and angiogenesis (36). Also, the autocrine production of growth factors from cancer cells helps to overcome the effect of anti-cancer drugs. 1.5 The extracellular matrix The extracellular matrix (ECM), can also affect the efficacy of chemotherapy and drug resistance. As described above, the efficiency of cytotoxics is influenced by various biological functions. The ECM similarly affects specific processes including cell growth, differentiation, motility, proliferation, and apoptosis (26, 37). Proteins present in the ECM such as laminin, collagen, vitronectin, and fibronectin influences these biological processes (38); however, many other proteins are also found in the ECM, since it functions as a reservoir for secreted molecules  7  (eg. growth factors, cytokines, matrix metalloproteinases (MMPs)).  All of these proteins  contribute to tissue organization and homeostasis (37). The ECM affects cellular activity through cell surface receptors, called integrins. Intracellular signals trigger the interaction of the ECM with integrin receptors. The ECMintegrin interaction which transforms the ECM mechanical forces into cellular signals, affects specific signaling pathways, including proliferation, survival, migration, and invasion (37). The ECM is composed of many different proteins, including fibronectin, laminin, vitronectin, and collagen IV. Collagen IV (CIV) is a major component of the basement membrane (39) and has been shown to play a role angiogenesis (40), cell adhesion and migration (41, 42). CIV has also been shown to reduce carcinogenesis. A fragment of CIV, the NC1 peptide, promoted adhesion and inhibited proliferation and invasion, resulting in the inhibition of growth of various human cancer cell lines (43-47). This effect of CIV has also been demonstrated in vivo. Injecting CIV in xenograft models of prostate and renal cell carcinoma decreased tumour growth by decreasing angiogenesis, proliferation, and invasion, and increasing apoptosis (48). Another group of ECM proteins that interacts with cell-surface receptors, ECM proteins, growth factors, cytokines and proteases are matricellular proteins. This group of proteins is unique as they modulate cell function, but do not serve as a scaffold for cells. Matricellular proteins are primarily expressed during development and growth, and in response to injury. Proteins in this family include thrombrospondins, secreted protein acidic and rich in cysteine (SPARC), and the CCN (acronym derived from the names of the first three members to be discovered: cysteine-rich protein 61, connective tissue growth factor, and nephroblastoma overexpressed gene) family of proteins (49).  8  1.6 Protein Synthesis Regulation of protein synthesis can play a key role in cellular function, and thus a brief summary of protein synthesis is presented here. Protein synthesis is a process where the cell builds proteins and involves the translation of mRNA to protein. In eurokaryotic cells, this can occur on free and bound ribosomes. As the mRNA molecule slides through a ribosome, codons are translated into amino acids one by one on a growing polypeptide chain. During and after its synthesis, the polypeptide begins to coil and fold spontaneously forming a functional protein with a specific conformation. Post-translational modifications, such as attachment of carbohydrates, lipids, or phosphate groups, may be required before the protein can fully function. Proteins that are destined for secretion are marked by a signal peptide, which targets the protein to the endoplasmic reticulum (ER). The signal peptide is recognized by a signal recognition particle (SRP). The SRP functions as an adaptor and brings the ribosome to the receptor protein on the ER membrane. Protein synthesis continues there, and the polypeptide crosses the membrane into the cisternal space via a protein pore. The signal peptide is usually removed by an enzyme, and the completed polypeptide is released into the cisternal space via exocytosis (50). Proteins can also enter the cell after being secreted. There are many possible mechanisms of how a protein can enter a cell. Some possible trafficking mechanisms are via bulk endocytosis or receptor-mediated endocytosis (51). The protein of interest for this thesis is SPARC, a matricellular protein that has been shown in our studies to have both intracellular and extracellular roles. However, to date there is no clear understanding of how SPARC, which is a secreted protein, may re-enter the cell and perform its intracellular functions. SPARC will be discussed in greater detail below.  9  1.7 SPARC SPARC (also termed osteonectin, or basement membrane-40, BM-40) was first found as osteonectin as a bone-specific phosphoprotein in mice (40). The same protein was later identified as a serum albumin-binding glycoprotein secreted by endothelial cells (52). Thus, SPARC is a glycoprotein in the matricellular family, which mediates cell-matrix interactions, but not in a structural manner (53). SPARC, a single-copy gene on human chromosome 5q31-q33, contains 10 exons, spans 34.5kb and is highly conserved across species (54-59). The SPARC promoter lacks a classical TATA box, but contains functional repeat CGA boxes, as well as cAMP-, heat shock-, and glucocorticoid-response elements (54, 59). The SPARC gene has two start sites, a major (2.1kb) and minor (3.0kb) mRNA SPARC transcript (54). Both transcripts have identical coding sequences, but the longer isoform utilizes a downstream polyadenylation signal (60). Linkage analysis revealed that SPARC is in close proximity to acidic fibroblast growth factor (FGF), colony-stimulating factor (CSF)-1, granulocyte-macrophage (GM)-CSF, interleukin (IL-3), CSF1 receptor (c-fms) and platelet derived growth factor (PDGF)-β receptor (53). 1.7.1 The domains of SPARC SPARC belongs to the SPARC-like family which contains seven members: SPARC, testican-1, -2, -3, SPARC-like 1 (ie. hevin or Mast9), and SPARC-related modular calcium binding (SMOC)-1, and -2. These members share similar biological domains: the N-terminal acid domain, the follistatin-like domain, and the extracellular domain. Each domain contains its own modular function and will be discussed in greater detail below. (Figure 1-5 and 1-6) (61, 62). Full length SPARC consists of a 17 amino acid (aa) signal sequence, which is cleaved prior to secretion of the protein, with a final protein size of 286 amino acids (32kDa) (62). The 10  N-terminal (NT) region (the first 52 aas), is an acidic region rich in Asp and Glu. It contains the major immunological epitopes and is the most divergent from other members of the SPARC-like family (62). It also contains two adjacent N-terminal Glu3 and Glu4 sites, which can act as amine acceptor sites in transglutaminase-catalyzed crosslinking modifications (63) and is the most divergent from the other members of the SPARC-like family. The follistatin-like (FS) domain (the next 85 aas), contains 10 cysteine residues that are sulfide-bonded, an N-linked complex carbohydrate at N99 and two copper binding sites (64). The sequence encodes a structure homologous to a repeated domain in follistatin, and consists of a N-terminal β-hairpin and a small core of mixed α/β structures (65). The FS-domain contains bioactive peptides that inhibit proliferation of endothelial cells (66); however, another peptide (KGHK) in the same region stimulates endothelial cell proliferation and angiogenesis (66, 67). The final domain, the extracellular Ca2+-binding (EC) domain (the final 149 aas), is mainly α-helical and contains a pair of EF-hand motifs that bind Ca2+ with high affinity (68, 69). The EC domain also interacts with the FS domain, stabilizing the binding of Ca2+ (65). This domain binds to collagen I, III, IV, and V by a mechanism dependent on Ca2+ (70, 71). The EC domain also contains peptide sequences to inhibit endothelial cell proliferation (72, 73). 1.7.2 SPARC expression In adults, expression of SPARC is abundant in tissues undergoing repair, remodeling, wound healing, or disease. Expression of SPARC is limited in normal cells such as steroidogenic cells, chondrocytes, placental trophoblasts, vascular smooth muscle, and endothelial cells (67, 74-77). SPARC expression is increased under stress-like conditions such as tissue repair of fibrocytes and endothelial cells (78), fibroblastic wound-edge (79), and keratinocyte differentiation. It is also up-regulated in disease state cells such as hepatic stellate cells in chronic hepatitis (80). 11  Differential SPARC expression is also observed between normal and neoplastic samples (81). SPARC is up-regulated in the malignant tissues of gastrointestinal, breast, lung, kidney, adrenal cortex, ovary, and brain tumours (78) and can be used as a prognostic marker for various cancers. Recently in a TRAMP model of prostate cancer, it has been found that SPARC expression causes a decrease in proliferating cells and an overall decrease in tumourigenesis. In SPARC -/- mice, an increase in incidence and aggressiveness of prostate cancer was observed, shown by increased distant metastases and invasion (82). Its evolutionary conservation and differential expression in normal versus disease states implies that SPARC may play an important physiological role. 1.7.3 Functions of SPARC SPARC has many functions including disruption of cell adhesion (83), changes of cell shape (84, 85), inhibition of cell cycle (66, 86), cell differentiation (87), inactivation of cellular responses to growth factors (72, 88, 89), angiogenesis (90), tumourigenesis (91-93), cataractogenesis (87, 94, 95), wound healing (77) and regulation of ECM and matrix metalloproteinase production (96). The protein also binds to albumin, thrombospondin, platelet cell membranes, endothelial cells (52, 97-99), and components of the ECM, including collagens, laminin, fibronectin, and vitronectin (100-102). 1.7.4 SPARC in cancer With SPARC’s ability to function in so many basic biological processes, it is not surprising it plays a role in cancer. To date, the exact role of SPARC in the development and progression of cancer has been unclear. The pattern of SPARC expression in tumours and surrounding stroma compared to normal tissue is cancer type dependent. In breast cancer (78, 103-105), melanoma (92), and glioblastomas (106) higher SPARC expression has been observed in tumour tissue when compared with normal cells. Based on this expression pattern, one would 12  hypothesize a potential role of SPARC in tumor promotion or progression. However, lower levels are seen in ovarian cancer (107), CRC (108, 109), pancreatic cancer (110) and acute myelogenous leukemia (111). This expression pattern of decreased SPARC levels would suggest an inhibitory role for SPARC in tumor formation. To date there is growing evidence for an important role for SPARC in various cancers; however, there is no unifying model which explains its contribution to cancer progression. It appears that the role of SPARC in cancer is tumor-specific. The role of SPARC in CRC is the focus of this thesis and will be discussed in detail in the next section. The involvement of SPARC in other cancers is summarized in a recent article by Tai and Tang (112). 1.7.5 SPARC in colorectal cancer SPARC expression is down-regulated in MIP/5FU, MIP/CPT, MIP/CIS, and MIP/Eto CRC cells that are resistant to chemotherapies such as 5-Fluorouracil (5-FU), irinotecan (CPT11), cisplatin (CIS), and etoposide (Eto) respectively, compared to the sensitive parental MIP101 CRC cells (113). It has also been shown that SPARC is down-regulated in various colorectal cancer cell lines, such as RKO, SW620 and HT29 (113). Two studies show that SPARC is down-regulated by methylation of the SPARC promoter (108, 109). SMAD-4 also suppresses SPARC expression (114) which would reduce the availability of SPARC to adhere to the ECM, as seen by gamma-linolenic acid, thus decreasing cell adhesion and cell motility (115). In animal studies, using SPARC-/- mice intercrossed with Apc/Min/+ (which are prone to developing intestinal tumors), the absence of SPARC results in the development of fewer adenomas, but no effect on the size of adenomas (116). No adenocarcinomas or metastases were detected, suggesting that in this animal model, SPARC may contribute to adenomatous formation, but not its cancer progression.  13  SPARC also enhances chemosensitivity of CRC cells. Treatment of SPARC alone did not affect the chemosensitivity of CRC cells; however, the combination of SPARC and chemotherapy treatment re-sensitized therapy-refractory CRC cells. In vitro, SPARC resensitized MIP/5FU cells to 5-FU by enhancing apoptosis. In vivo, the enhancement of apoptosis from the combination treatment of SPARC and 5-FU, results in greater tumour regression of both sensitive and resistant subcutaneous tumours when compared to control treatment or 5-FU treatment alone. Also, over-expression of SPARC in MIP101 cells (MIP/SP) results in increased apoptosis when treated with 5-FU, CPT-11, CIS, and Eto (113). Previous microarray results showed that SPARC expression was down-regulated in therapy-refractory CRC cell lines also demonstrated a down-regulation of the vitamin D receptor (VDR) in the resistant CRC cell lines. It has been shown that vitamin D up-regulates VDR expression. Interestingly, treatment of the CRC tumours with 62.5nmol/L 1α,25dihycroxyvitamin D(3) (1,24-D3) restored both VDR and SPARC expression; thus, increasing the sensitivity of resistant CRC cells, by decreasing cell viability (117). This sensitivity was in part due to increased apoptosis and inactivation of the pro-survival pathway, through Akt. The restoration of chemosensitivity was also seen in vivo, where exogenous SPARC treatment increased VDR expression of tumour xenografts (117). SPARC also affects other biological functions in CRC. SPARC expression delays cell cycle progression at the G1/S phase (113) and 1,25-D3 further halts cell cycle progression in response to SPARC (117). Increased SPARC expression also results in decreased cell proliferation and blood vessel formation (113). These studies support SPARC’s ability to enhance chemosensitivity through multiple pathways in CRC.  14  1.8 Mechanistic aspects of SPARC action in cancer biology Due to SPARC’S ability to affect many biological functions, it is not surprising that SPARC plays various roles in cancer progression. The mechanisms mediating these roles may provide a greater understanding of the functions of SPARC and will be outlined below. 1.8.1 Hypermethylation of the SPARC promoter Cancer cells regularly exhibit an imbalance of cell survival and apoptosis genes. One mechanism that commonly modulates the expression of these particular types of genes is methylation of the promoter (118, 119). Methylation of the SPARC promoter has been observed in various cancer types including lung (120), prostate (121), endometrial (122), pancreas (123), colorectal (108, 109) and leukemia (111). Methylation of the SPARC promoter and subsequent down-regulation of SPARC expression has also been observed in chemotherapy-resistant CRC cells (113). Interestingly in SPARC -/- mice, there was an increase in tumourigenesis of pancreatic (110) and prostate cancers (82). These results suggest that methylation of the SPARC promoter may down-regulate SPARC expression and promote tumourigenesis. Additionally, exposure to a demethylating agent, 5-aza-2’deoxycytidine, up-regulated SPARC expression in pancreatic (123), cervical (124) and CRC cell lines (109). In CRC cells, this up-regulation of SPARC expression correlated to decreased cell viability and an increase in apoptosis in response to chemotherapy (109). 1.8.2 Apoptosis The ability of SPARC to induce apoptosis has been well documented. In various ovarian cancer cells, exogenous treatment with SPARC induced apoptosis (107) and increased cleaved caspase 3 in human ovarian carcinoma cells (125). Similarly, pancreatic (110) and ovarian cancers (125) grown in SPARC-/- mice demonstrate enhanced growth and reduced apoptosis.  15  In CRC cells, overexpression of SPARC reduced cell viability and enhanced apoptosis when exposed to various chemotherapeutic agents (113). Apoptosis can be augmented further following exposure to vitamin D and chemotherapy, by reducing phosphorylation of Akt and subsequent inactivation of the pro-survival pathway through an undefined mechanism (117). 1.8.3 Angiogenesis, invasion and metastasis Additional biological processes are needed for cancer cells to metastasize. Major processes include invasion and angiogenesis. Interestingly, studies have shown that SPARC plays a role in invasion, metastasis, and angiogenesis. As a tumour suppressor, SPARC inhibited adhesion and invasion to collagen IV, vitronectin, fibronectin, collagen I, hyaluronic acid or laminin in ovarian cancer cells (125) and also inhibits cell migration and invasion in pancreatic cancer cells (both in vitro (126) and in vivo (110, 126, 127)). Alternatively, as a proto-oncogene, SPARC promotes the metastasis of melanoma cells (128). SPARC’s role in angiogenesis (a process of neovascularization) is varied (128). In the case of endothelial cells, SPARC directly binds to various growth factors (vascular endothelial growth factor (VEGF), platelet derived growth factor (PDGF), and basic fibroblast growth factor (bFGF)) resulting in reduced interaction with their respective receptors (72, 88, 89) and inhibited angiogenesis. While in ovarian cancer, the absence of SPARC promotes angiogenesis and the tumour obtains metastatic potential (125, 129). As illustrated above, SPARC affects many biological functions. This thesis focuses on apoptosis, which will now be discussed in greater detail. 1.9 Apoptosis, autophagy, and necrosis The most widely used classification of cell death recognizes apoptosis and necrosis. Apoptosis is also known as type I cell death and generally provides an advantage for the organism as it can selectively kill cells that pose a threat (eg. cancer or redundant cells). It is a 16  highly controlled, energy dependent process and targets individual or clusters of cells. In contrast, necrosis is defined as the premature death of cells and is detrimental to the organism. Necrosis is typically caused by external factors (eg. infection and trauma) and causes swelling of the cell and organelles. It is a passive process that often affects large fields of cells. The difference between apoptosis and necrosis lies in the plasma membrane. In apoptosis, the plasma-membrane integrity remains until late in the process. However, in necrosis, the plasma membrane loses its integrity early, which results in the swelling of the cell and organelles (130, 131). A third form of cell death is autophagy, a process where cells digest their own organelles and macromolecules for energy. Similar to apoptosis, autophagy is highly controlled, energy dependent, and affects individual or clusters of cells. It allows for a cell that is deprived of growth factors to survive, thus providing a growth advantage for the cell. However, there is a form of autophagic cell death, type II cell death, where all available substrates are digested and the cell eventually dies. The process of autophagy is the primary mechanism for turnover of long lived proteins, resulting in the maintenance of the quality of proteins and organelles. The differences in these three modes of cell death differ in their morphological and molecular attributes (130, 131). Other cell death mechanisms have been discovered, such as anoikis, which is a form of apoptosis caused by loss of cell adhesion (132), but this thesis largely focuses on the signaling pathway of apoptosis, which will be further discussed below. 1.9.1 What is apoptosis? Apoptosis, a form of programmed cell death, plays a major role in the growth and maintenance of the human body (133). In some cases, during development, apoptosis eradicates unwanted host cells in order to maintain stasis of the system without interfering with the overall 17  function of the tissue (134-137). For example, during development of the nervous system, neurons that do not form proper connections will undergo apoptosis (138). As one might imagine, apoptosis plays a very important role in normal human development; thus, dysregulation of the process has been implicated in several human diseases. Neurodegenerative diseases can be caused by excessive apoptosis, while insufficient apoptosis results in autoimmune syndromes or cancer (137, 139). Many stimuli, both threatening (pathogenic invasion) and non-threatening (redundant or unnecessary cells), can activate or trigger apoptosis (137). After activation, detection and transduction of the signal results in the effector stage, activates specific signaling events and then proceeds to the final stage (140, 141). The final stage is identified by characteristic cell morphology, such as cell shrinkage, reorganization of the cell nucleus, membrane blebbing, and fragmentation of the cell into membrane-enclosed vesicles (134, 140, 141) surface markers that allow rapid phagocytosis by neighboring cells and macrophages (Figure 1-7) (142). The entire execution of apoptosis is brief (approximately 15 minutes to 1 hour) and irreversible. 1.9.2 Mechanism of apoptosis Apoptosis activates specific signaling pathways. CED-3 is one of the first genes identified as part of the apoptotic pathway and originally described in Caenorhabditis elegans (C. elegans) (143). Full length CED-3 undergoes proteolytic cleavage to generate cysteine protease CED-3 in C. elegans (144). Further studies uncovered various mammalian homologues of CED-3, based on their ability to promote apoptosis. Interleukin 1β converting enzyme (ICE), later defined as caspase (cysteine-dependent aspartate specific proteases) 1 was one of the first CED-3 mammalian homologues identified. Studies showed that various stimuli led to over-expression of ICE resulting in cell death, while inhibitors of ICE suppressed apoptosis (145-150). Price (protease resembling ICE), yet another 18  homologue of CED-3, was originally identified as a protease responsible for the cleavage of PARP in an in vitro system (151). Partial inhibition of cell death due to expression of anti-sense Nedd2 identified Nedd2 as another mammalian CED-3 homologue (152, 153). After the identification of many CED-3 mammalian homologues (145, 154-158), CPP32 (later defined as Yama/apopain and finally caspase 3) was identified as the main protease responsible for mammalian apoptosis. CPP32 is derived from a common proenzyme that is related to both ICE (caspase 1) and CED-3. Initial studies found that a peptide inhibitor of apopain/CPP32 prevented apoptosis, suggesting that apopain/CPP32 is important for the initiation of cell death (159, 160). Many other homologues have been identified for genes involved in apoptosis. In metazoans, the specific apoptotic signaling pathway is conserved; however, it is more complex in mammals. Mammalian apoptosis is divided into two arms: the extrinsic and the intrinsic. In both pathways, caspases, which are peptide bond hydrolases with specificity for tetrapeptidemotifs-containing aspartate, play a central role. They are synthesized as inactive proenzymes and comprised of large and small subunits, which upon proteolytic cleavage are activated (161). Caspases are further divided into initiator caspases and effector caspases. Initiator caspases (with large prodomains, eg. caspase 8 or 9) are able to self-activate in addition to activating the effector caspases (with short prodomains, eg. caspase 3 or 7) (162) to execute the final stages of apoptosis as described above. 1.9.3 The extrinsic pathway of apoptosis Apoptosis in mammalian cells involves the extrinsic and intrinsic pathways.  Both  pathways differ in the method of activation and gene involvement. The extrinsic pathway typically involves death receptors, which are members of the tumour necrosis factor (TNF) receptor superfamily. Death receptors are further divided into subfamilies based on the varying 19  intracellular death domains. Examples of death receptors include: CD95 (APO-1/Fas), TRAIL (TNF-related apoptosis-inducing ligand)-R1, TRAIL-R2, TNF-R1, DR3 and DR6 (163). Typically, the extrinsic pathway of apoptosis is activated by death receptors at the cell surface (164-166). After they bind a specific ligand, followed by recruitment of intracellular adaptor proteins and pro-caspases (eg. caspase 8 or 10). All of these components (the receptor, adaptor protein, and pro-caspase) form the death-inducing signaling complex (DISC) (167). After the DISC is formed, caspases self-process and activate effector caspases (Figure 1-8). For example, caspase 8 forms a self-activating heterotetramer of two small and two large subunits. Active caspase 8 then activates effector caspases, such as caspase 3 (168), thus executing the final stages of apoptosis and leading to characteristic apoptotic changes, such as chromatin condensation and blebbing (Figure 1-7). Although the extrinsic and intrinsic pathway can function independently, the extrinsic pathway can also be linked to the intrinsic through the cleavage of Bid (169). After activation of the death receptors, caspase 8 cleaves Bid, producing t-Bid, which translocates to the mitochondria and activates the intrinsic pathway of apoptosis (Figure 1-8) (170). 1.9.4 The intrinsic pathway of apoptosis As the extrinsic pathway is activated by death receptors, the intrinsic pathway of apoptosis is regulated by the mitochondria. Permeabilization of the mitochondrial outer membrane, which is triggered by various stimuli such as calcium, reactive oxygen species, DNA damage, UV radiation, gamma radiation, and ceramide derivatives, is the critical event that commits cells to apoptosis (161, 170-173). During permeabilization, cytochrome c is released from the mitochondrial intermembrane space (IMS), forming the apoptosome (a complex with apoptosis protease-activating factor 1 (Apaf-1), pro-caspase 9, and cytochrome c) (168, 174-  20  176). Self-activation of pro-caspase 9 cleaves effector caspases 3, 6, and 7 (177), which in turn cleave cellular substrates in the final steps of apoptosis (Figure 1-8). 1.9.5 Regulators of apoptosis Regulation of apoptosis is very important because it maintains the equilibrium between cell survival and cell death. Disequilibrium in the apoptotic pathway leads to various human diseases, such as cancer, neurodegeneration, and autoimmune disorders (178, 179). Thus, many regulatory mechanisms are in place to ensure proper maintenance of the intrinsic pathway. As primary regulators of the intrinsic pathway, the Bcl-2 family plays a major role in maintaining the balance between cell survival and apoptosis (180). The members of this important family are classified into two groups: 1) pro-survival (Bcl-2, Bcl-XL, Bcl-w, and Mcl1), and 2) pro-apoptosis (eg. Bax, Bak, Bok), and the “BH3-only” proteins (Bid, Bim, Bik, HRK, Puma, and Noxa) (161, 181, 182). Regulation of apoptosis by the Bcl-2 family occurs by controlling the relative levels of these two groups within each cell. Other proteins that regulate apoptosis are inhibitors of apoptosis (IAP) (183, 184). IAPs bind directly to caspase 3, 7 and/or 9 to impair their activity (185) (Figure 1-8). Examples of human IAP members include X-chromosome-linked IAP (XIAP), cellular IAP-1 (c-IAP1), cIAP2, BIRC1, survivin and Apollon (184, 186). 1.9.6 The pro-survival pathways The regulation of apoptosis can also occur through other signaling pathways. Survival signaling pathways are closely linked to the regulation of the apoptotic pathway. The survival pathways can promote survival by cellular proliferation and the inhibition of apoptosis (Figure 18). Examples of these pathways include phosphatidylinosital 3-kinase (PI3K)-Akt, NFkB, and Ras-Raf-MEK-ERK pathways.  21  Together, the survival and apoptotic signaling pathways contribute to a complex signaling network that regulates proper cell growth (187). In cancer cells, all of the signaling pathways contribute to tumour growth and are therefore important in the treatment of cancer. 1.10 Research objectives High mortality rates continue to be observed in advanced CRC because of therapy-refractory disease. In the last decade, the use of newer agents, such as irinotecan (CPT-11) (188, 189) and oxaliplatin (190, 191) have led to significant improvements in response rates, from ~20% to ~50%. However, the median survival has remained poor at <17 months, partly due to resistance to chemotherapy. Therefore, novel approaches are needed that can re-sensitize unresponsive tumours to chemotherapy to achieve better overall survival in patients with advanced cancers. The effectiveness of chemotherapy can be assessed by its ability to enhance tumour cell death. This can be accomplished by activating signaling pathways involved in cell death (192). We have shown that SPARC is capable of promoting tumour regression by activating apoptosis, which may explain its ability to re-sensitize therapy-refractory cancer cells to chemotherapy. Our earlier findings that SPARC enhances chemosensitivity by increasing apoptosis led us to investigate the signaling events involved in SPARC-mediated apoptosis in this thesis. SPARC has three distinct biological domains, each of which has its own modular function. The main hypothesis of this thesis is that SPARC, which re-sensitizes resistant CRC cells by inducing apoptosis in response to the combination treatment of exogenous SPARC and chemotherapy, enhances chemosensitivity by activation of the extrinsic or intrinsic pathway of apoptosis and through the modulation of the ECM. Therefore, the aims of this thesis are three-fold: 1) To determine the specific signaling pathway of SPARC-mediated apoptosis; 2) To identify the specific domain of SPARC that induces apoptosis; and 3) To determine the function of collagen IV and SPARC in colorectal carcinogenesis. 22  Determining the specific signaling mechanisms of SPARC-mediated apoptosis will provide further insight in the mechanism of how SPARC re-sensitizes therapy-refractory CRC cells. Determining which biological fragment of SPARC is involved will also help distinguish the specificity from the other members of the SPARC-like family. Also, the specific signaling mechanism may then be utilized to promote tumour regression and will help further understand the use of SPARC as a potential therapeutic agent for advanced diseases.  23  Figure 1-1. Illustration of colorectal tumour progression in sporadic and high-risk genetic syndromes. The general paradigm of tumour initiation occurs from sustained genetic damage over time due to the environment (sporadic) and any germline (FAP: familial adenomatous polyposis) mutation that has been inherited. Damaged DNA provides a growth advantage for tumour progression, ultimately forming carcinoma. Adapted from Grady and Carethers, Gastroenterology, 2008.  24  Figure 1-2. The progression of colorectal cancer with chromosomal instability (CIN). CIN is characterized by aneuploidy. Initiation of tumour formation occurs with interruptions of the Wnt signaling pathway, including the APC gene (ACF, aberrant crypt foci). Progression to cancer is driven by cellular clonal expansion that acquires enhanced growth characteristics, including mutational activation of proto-oncogene KRAS and mutation of TP53. Mutational activation of PI3KCA occurs late in the progression of cancer. Adapted from Grady and Carethers, Gastroenterology, 2008.  25  Figure 1-3. The progression of colorectal cancer with CpG Island methylator phenotype (CIMP). CIMP is characterized by abnormal methylation of several promoter sequences of tumour suppressor genes. Methylated promoters of MGMT, EVL, HLTF, SFRP2, SLC5A8, MGMT, MINT, MLH1, TSP1, and TIMP3 occur during the initiation phase of colorectal tumour development. Adapted from Grady and Carethers, Gastroenterology, 2008.  26  Figure 1-4. Intracellular activation of 5-Fluorouracil (5-FU). (dR-1-P, deoxyribose-1phosphate; dUTP, deoxyridine triphosphate; FdUDP, fluorodeoxyuridine diphosphate; FdUMP, fluorodeoxyuridylate; FdURD, 5-fluoro-2’-deoxyuridine; FdUTP, fluorodeoxyuridine triphosphate; FUDP, fluorouridine diphosphate; FUMP, fluorouridine monophosphate; FUrd, 5fluorouridine; FUTP, fluorouridine triphosphate; PPRP, phosphoribosyl phosphate; R-1-P, ribose-1-phosphate). Adapted from Cancer Chemotherapy and Biotherapy: Principles and Practice. Third Edition.  27  Figure 1-5. The functional domains of SPARC are represented.  28  Figure 1-6. The structural domains of SPARC are represented. Reprinted from Matrix Biology, 19 (8), Rolf A. Brekken and Helene E. Sage, SPARC, a matricellular protein: at the crossroads of cell-matrix communication, 816-827, Copyright (2001), with permission from Elsevier.  29  Figure 1-7. Characteristic morphological features of apoptosis. This diagram represents events that occur after the execution of apoptosis. Cytomorphological features include cell shrinkage, chromatin condensation, formation of cytoplasmic blebs and apoptotic bodies, and the phagocytosis of the apoptotic bodies by adjacent parenchymal cells, neoplastic cells, or macrophages.  30  Figure 1-8. A simplified schematic representation of the apoptotic cell signaling pathways. There are many other players and pathways that function in the regulation of apoptosis. Only players pertaining to this thesis are represented. The two main pathways of apoptosis are the extrinsic and intrinsic pathways. Each requires specific activation signals to begin a caspase (Csp)-dependent cascade of molecular events. Each pathway activates its own initiator caspases (8, 9, 10) which will then activate effector caspases (3, 6, 7). The apoptotic pathway can be regulated by members of the Bcl-2 family (anti-apoptotic: Bcl-2, Bcl-XL; pro-apoptotic: Bax, PUMA, Bad), inhibitors of apoptosis (IAPS), and/or pro-survival pathways (PI3K/Akt).  31  2. Materials and methods 2.1 Cell lines, reagents MIP101 cells were used for the majority of the studies. MIP101 cells are an undifferentiated colon carcinoma cell line that was established from a patient with Dukes D colorectal carcinoma that metastasized to the liver (193). Colorectal cancer cells: MIP101, HCT116 (ATCC), RKO; pancreatic cancer cells: MiaPaca; and breast cancer cells: MCF-7, were maintained in DMEM media supplemented with 1% penicillin-streptomycin, 1% kanamycin (Invitrogen) and 10% newborn calf serum at 37°C and 5% CO2. For resistant cell lines, media were also supplemented as follows: MIP101 cells resistant to 5-FU (MIP/5FU), 500 µM 5-FU; CPT-11 (MIP/CPT), 10 µM CPT-11; HCT116 cells resistant to 5-FU (HCT116/5FU), 10 µM 5FU; RKO cells resistant to 5-FU (RKO/5FU), 25 µM 5-FU; CPT-11 (RKO/CPT), 18 µM CPT11; cisplatin (CIS) (RKO/CIS), 30 µM CIS; MiaPaca cells resistant to CPT-11 (MiaPaca/CPT), 100 µM CPT-11; MCF-7 cells resistant to cisplatin (MCF7/CIS), 30 µM CIS. For MIP101 cells stably-transfected with empty vector (MIP/ZEO) or MIP101 cells stably transfected with an expression vector for SPARC (MIP/SP), media were also supplemented with 0.01% zeocin (Invitrogen) (Figure 2-1) (113). MIP/SP cells were similar to morphology as parental MIP101 cells, but were slightly flatter. Mammalian expression vectors containing the N-terminal (SP-N, aa18-69, 5.8kDa), follistatin-like (SP-F, aa70-154, 9.6kDa), and extracellular (SP-C, aa155-303, 16.6kDa) domains were generous gifts from Dr. W. Schiemann (National Jewish Medical and Research Centre; Denver, CO) and tagged with myc and His6–tagged (Figure 2-1) (194). The biological domains of SPARC were expressed in psectag expression vectors, which include the CMV promoter, the T7 promoter/priming site, and the Ig κ-chain leader sequence to allow for secretion (Figure 2-1). All constructs were verified by DNA sequencing. Previous studies utilizing tagged-SPARC (or related peptides) have demonstrated similar biological activity as the 32  native, un-tagged proteins (194, 195). MIP101 cells stably-transfected with SP-N (MIP/SP-N), SP-F (MIP/SP-F), or SP-C (MIP/SP-C), all with similar morphology to MIP101 cells overexpressing SPARC (MIP/SP), were generated and supplemented with 0.02% hygromycin (Sigma). Expression levels of stable and transient transfectants were quantified by ELISA assay to His. Recombinant human SPARC (rSPARC), generously provided by Dr. Neil Desai (Abraxis BioScience Inc., USA), was produced in a G418 selected 293 cell line. rSPARC-containing Cterminal His6 tag was purified from hollow fiber cartridge bioreactors (1.2 m2) by Ni-Sepharose affinity chromatography on an AKTA-FPLC, with a purity of 95% to 99%, and endotoxin levels less than 0.5 EU/mg of protein. 2.2 Site-directed mutagenesis SPARC (SP), SP-N, SP-F, and caspase 8 containing plasmids were used for site-directed mutagenesis, using GeneTailor Site-Directed Mutagenesis System (Invitrogen) as per manufacturer’s protocol. These mutants were expressed in the MIP101 cell line. Please see Table 2-1 for specific primers used to generate the mutations. After mutagenesis of plasmids, mutant constructs were verified by DNA sequencing. Please see Table 2-2 for the specific amino acids mutated and the experimental usage. 2.3 Coating of plates with collagen IV Pre-coated with collagen IV (CIV) plates (BD Falcon) were used for most studies (~50 µg/cm2). For the remainder of the studies, plates were incubated with mouse CIV (BD Falcon) that was dissolved in 0.05 N HCl for 1 hour at room temperature for a final concentration of 50 µg/cm2. Excess CIV was then removed, the plates were washed with 1X PBS to remove the acid and used immediately. All control plates were coated with 0.05 N HCl for 1 hour at room temperature as described above.  33  2.4 ELISA for quantification of SPARC or related peptides To test for expression of stable and transient transfectants, ELISA's were performed. Stable cell lines (MIP/ZEO, MIP/SP, MIP/SP-N, MIP/SP-F and MIP/SP-C) or transiently transfected cells (plasmids as listed above) were grown to 90% confluence and isolated as previously described (113). Microwells were coated with anti-SPARC antibody (1:5000) in coating buffer (0.1M sodium carbonate, pH 9.5) overnight at 4ºC, washed 3 times with PBS-T (0.05% Tween-20), and blocked with 2% BSA in PBS for 1 hour at room temperature. Samples (10 µg/mL) were added to the wells, centrifuged at 2000g for 10 minutes at 4ºC, and incubated for 4 hours at 4ºC. Wells were washed 3 times with PBS and then incubated with either anti-HIS or anti-myc (1:5000) antibodies for 2 hours at room temperature, washed 5 times with PBS-T, and incubated with appropriate secondary-HRP antibodies (1:10 000) for 1 hour at room temperature. Wells were then washed 5 times with PBS-T and 100 µL of TMD chromagen substrate was added to each well for 15-30 minutes. Absorbance was read at 650 nM (Figure 22). Controls used were blank, PBS, and PBS plus antibody wells. The results from these controls were negligible to the results found. 2.4.1 ELISA to quantify the localization of SPARC MIP101, MIP/5FU, RKO, or RKO/CPT cells were transiently transfected with plasmids (full length SPARC: SP, SPARC clones without the intracellular sequence: SPc1, SPc4, SPc5, or empty vector, EV) or incubated with exogenous proteins (full length SPARC: SP, biotinylated SPARC at 1:20 ratio of SPARC:biotin : 20Xb-SP; biotinylated SPARC at a ratio of 1:50 SPARC:biotin 50Xb-SP; or empty vehicle, EV) and grown to 90% confluence. Biotin and SPARC were covalently linked through a N-hydroxysuccinimide ester. The maximum number of lysines that can be coupled is twenty; however, we did not measure the level of lysines coupled to biotin. The above mentioned ratio of 1:20 or 1:50 of SPARC:biotin refers to the ratio of the 34  reagents used. Avidin beads were introduced into the cell culture to bind biotin and thus keep the biotin:SPARC complex extracellular. Prior to harvesting, cells treated with SPARC were slightly flatter, but were similar to cells not treated with SPARC. Conditioned media and cell lysates were isolated in 1X CHAPS. Microwells were coated with anti-SPARC antibody (1:5000) in coating buffer (0.1 M sodium carbonate, pH 9.5) overnight at 4ºC, washed 3 times with PBS-T (0.05% Tween-20), and blocked with 2% BSA in PBS for 1 hour at room temperature. Samples (10 µg/mL) were added to the wells, centrifuged at 2000 g for 10 minutes at 4ºC, and incubated for 4 hours at 4ºC. Wells were washed 3 times with PBS and then incubated with either anti-HIS or anti-myc (1:5000) antibodies for 2 hours at room temperature, washed 5 times with PBS-T, and incubated with appropriate secondary-HRP antibodies (1:10 000) for 1 hour at room temperature. Wells were then washed 5 times with PBS-T and 100 µL of TMD chromagen substrate was added to each well for 15-30 minutes. Absorbance was read at 650 nM. 2.5 RT-PCR Cells were seeded at 150,000 cells/well in 6-well plates. After 24-48 hours, cells were incubated with 0-1000 µM 5-FU for 4-48 hours and RNA isolated with Trizol (Invitrogen) (113). 1 µg of total RNA was used to generate cDNA (Superscript III, Invitrogen). To assay for mRNA stability, cells were incubated with actinomycin D (10 µg/mL) (Sigma) for 0-20 hours, and RNA were isolated for RT-PCR. Specific primers were used as previously described as listed in Table 2-3. PCR products were separated on a 1.5% agarose gel using electrophoresis and expression levels were quantified using ImageJ analysis. The values were normalized to β-actin expression values as previously described (196).  35  2.6 Chromatin immunoprecipitation (ChIP) assay Monolayers of MIP/ZEO and MIP/SP cells were fixed by adding formaldehyde to a final concentration of 1% for 10 minutes at 37°C. Cells were rinsed and collected in 1X PBS. Cell pellets were resuspended in 200 µL SDS lysis buffer (Table 2-4) and lysed on ice for 10 minutes, followed by sonication and centrifugation at 13,000 rpm for 5 minutes. Supernatant was diluted in ChIP buffer (Table 2-4). Cell lysates were pre-cleared and input fractions were put aside. Immunoprecipitations were carried out using anti-interferon regulatory factor (IRF)-1 and IRF-2 antibodies (Santa Cruz) with overnight incubation at 4°C. 100 µL of Protein A Sepharose beads (Sigma) were added and incubated for 1 hour at 4°C, washed with low salt wash buffer, and TE buffer (Table 2-4). Beads were extracted with 50 µL of elution buffer (Table 2-4) and 15 µL of 1M NaHC03 was added to the input samples. Eluates and input fractions were incubated overnight at 65°C. DNA was purified with PCR purification kit (Qiagen), and used for PCR as described above. 2.7 RNA interference Initially, to assess the efficiency of caspase 8 gene expression knock-down by siRNA, high SPARC-expressing MIP/SP and HCT116 cells were used. 24 hours post-seeding, cells were transiently transfected with 20-60 nM scramble oligonucleotide sequence (control), or caspase 8 siRNA (Stealth RNAi, Invitrogen) and cells collected at various time intervals following transfection. 40 nM of siRNA yielded the most efficient knock-down (14-fold decrease in caspase 8 expression at 48-96 hours, Figure 2-3). For all subsequent experiments, 40 nM of siRNA or scramble control was used, and transient transfections carried out for at least 72 hours. Following caspase 8 siRNA transfection, cells were assessed for cell viability, and apoptosis using either Caspase 3/7 assay or TUNEL assay.  36  2.8 Cell viability 2.8.1 RNA Interference To assess the effect of RNA interference on cell viability, cells were seeded in 96-well plates. 24 hours after seeding (~60% confluence), cells were transiently transfected with caspase 8 siRNA for 36-48 hours before incubation with 0-2500 µM 5-FU or 100 µM CPT-11 for 36-72 hours. Cell viability was assessed by MTS assay (Promega) at 490 nm. MTS assays are standard colorimetric assays that measure the activity of enzymes that reduce MTT to formazan by changes in color. These color changes can then be measured through absorbance at specified wavelengths. The percentage of viable cells was calculated based on the A490 reading of nontreated control cells. 2.8.2 Caspase 8/9 Inhibition To determine the effect of caspase 8/9 inhibition, cells were seeded (96-well plates), and incubated 24 hours later (~60% confluence) with 10-50 µM of caspase 8-like inhibitor (z-IETDfmk, Sigma) or caspase 9-like inhibitor (z-LEHD-fmk·TFA, Sigma) for 30 min, followed by incubation with 1000 µM 5-FU for an additional 24 hours. Cell viability was assessed by MTS assay as described above. 2.8.3 SPARC domains To assess if the smaller peptides representing the different domains of SPARC had an effect on cell viability, cells were transiently transfected with expression constructs containing: full length SPARC (SP), NT-domain of SPARC (SP-N), FS domain of SPARC (SP-F), EC domain of SPARC (SP-C), mutant NT-domain of SPARC (SP-Nmut1, SP-Nmut2), mutant FSdomain of SPARC (SP-Fmut1, SP-Fmut2); wild-type caspase 8, or mutants of the DEDI domain (DEDIm), putative binding domain (PBm), or DEDII domain of caspase 8 (DEDIIm), or empty 37  vector control (ZEO) for 48 hours. Cells were then exposed to 100 ng/mL rSPARC (Abraxis BioSciences Inc.), and either 500-1000 µM 5-FU (MIP101 lines), 50-100 µM CPT-11 (MIP101 lines), 18-36 µM CPT-11 (RKO lines), 100-200 µM CPT-11 (MiaPaca lines), or 60 µM CIS (MCF7 lines) for 24-48 hours. For stable overexpression clones, MIP/ZEO, MIP/SP, MIP/SP-N, MIP/SP-F, MIP/SP-C, and MIP/5FU cells were seeded at ~50% confluence. 24 hours postseeding, cells were treated with 1000 µM 5-FU for 48 hours. Cell viability was assessed by WST assay (Roche) at 450nM. WST assays are standard colorimetric assays that measure the metabolic activity of viable cells. Cells reduce WST to a soluble formazan salt, which can then be measured through absorbance at specified wavelengths. The percentage of viable cells was calculated based on the A450 reading of non-treated control cells. 2.8.4 SPARC localization To determine if SPARC exerted its effects intracellularly or extracellularly, cells were transiently transfected with expression constructs containing: full length SPARC (SP), SP rSPARC (SP, biotinylated at 20X (20X b-SP) or 50X concentrations (50X b-SP) or empty vehicle control for 48 hours. Cells were incubated with 1000 µM 5-FU (MIP101 cells lines or 36 µM CPT-11 (RKO lines) for an additional 24 hours and assessed for cell viability by WST assay as described above. 2.8.5 Collagen IV To assess the effect of CIV on cell viability, cells were seeded on CIV or control plates. 24 hours post-seeding (~60% confluence), cells were transiently transfected with SPARC or ZEO control for another 24 hours before incubation with 0-1200 µM 5-FU for an additional 24 hours. Cell viability was assessed by WST reagent as described above. The percentage of viable cells was calculated based on the A450 reading of non-treated control cells using the following  38  calculation: [A450nm(unknown)/A450nm (nontreated control cells on control plates) x 100%] = % cell viability. 2.8.6 Cellular number The number of viable cells was assessed using trypan blue exclusion assay (1:1) on a hematocytometer. Cells were seeded on CIV or control plates with equal numbers (~30% confluence) and counted for 2 consecutive days, starting 3 days after seeding reaching ~70-80% confluency at the end of the assay. 2.9 Cell proliferation Cell proliferation was assessed by determining the percentage of cells positively stained with Ki-67. Cells grown on coverslips were subjected to cell cycle synchronization with double thymidine block as previously described (113). After the final thymidine block, cells were cultured for an additional 48 hours and fixed with 4% paraformaldehyde for 20 minutes, washed in PBS, and incubated with 1% Triton X-100 (Sigma) for an additional 5 minutes. Cell were incubated with primary antibody overnight (1:100, Oncogene) at 4°C, rinsed in PBS, followed by incubation with secondary antibody (1:500, Invitrogen) at 37°C for 20 minutes, rinsed in PBS, and counterstained with 4’,6-diamidino-2-phenyl-indole as previously described (117). The number of Ki-67-positive cells were counted and averaged from 3 different fields (n=3 independent experiments). 2.10 Caspase 3/7 assays 2.10.1 RNA interference Cells were transiently transfected with 40 nM of caspase 8 siRNA for 48 hours, and incubated with 1000 µM 5-FU for another 48 hours. Total cell lysates were isolated and protein concentrations were measure by Bradford assay (BioRad). 20 µg of total protein/sample were 39  used in Caspase-Glo 3/7 Assay (Promega), using a 1:1 dilution of Caspase-Glo 3/7 Substrate. Relative luminescence units (RLU) were quantified using a Viktor2 1420 Multilabel counter (Perkin Elmer). Values were read as per manufacturer’s protocol and normalized to untreated MIP101 cells (baseline of 100%). 2.10.2 SPARC localization To determine if SPARC exerted its effects intracellularly or extracellularly, cells were transiently transfected with expression constructs containing: full length SPARC (SP), SP rSPARC (SP, biotinylated at 20X (20X b-SP) or 50X concentrations (50X b-SP) or empty vehicle control for 48 hours. Cells were incubated with 1000 µM 5-FU (MIP101 cells lines or 36 µM CPT-11 (RKO lines) for an additional 24 hours and used for caspase 3/7 assay as described above. 2.10.3 Collagen IV studies Cells were seeded on CIV or control plates. 24 hours later, cells were transiently transfected with SPARC, SPARC's biological domains and its mutants (or ZEO control) for 24 hours. Cells were incubated with 1000 µM 5-FU for another 0-4 hours. Total cell lysates were isolated and used for caspase 3/7 assay as described above. 2.11 TUNEL Assays Cells were seeded (24-well plates) to achieve ~60% confluence 24 hours later for transient transfection with caspase 8 siRNA. 36-48 hours later, cells were incubated with 1000 µM 5-FU for 36-48 hours, harvested (suspension and attached cells) and fixed onto glass slides with Shandon cytospin at 2000 rpm for 10 min and stained as per manufacturer’s instructions (Promega). The percentage of apoptosis was calculated by counting the number of TUNEL-  40  positive vs. the number of control DAPI cells. The counts were averaged from four different fields by two individuals in a blinded fashion (n=4 independent experiments). 2.12 Immunoblot analysis 48 hours after seeding, cells were incubated with 1000 µM 5-FU, and collected at 0-12 hours for protein as previously described (113, 117). For CIV experiments: 24 hours after seeding on CIV or control plates, cells were transfected with SPARC or ZEO control (Polyethylenimine, Sigma) for an additional 24 hours prior to incubation with 1000 µM 5-FU, and collected at 0-12 hours for protein as previously described (113, 117). Total proteins concentrations were measure by Bradford assay (BioRad). 40 µg total protein/sample was loaded, separated on a 12% SDS-PAGE, then transferred to PVDF membranes (Bio-Rad). Immunodetection was performed using antibodies against caspase 8, cleaved caspase 8, FADD, p-FADD, caspase 10, BID, caspase 9, and caspase 3 (all 1:1000, Cell Signaling Technologies) followed by incubation with the appropriate secondary antibody. All immunoblots were also probed with antibodies to β-actin (1:5000, Abcam) as loading control. Proteins were detected with SuperSignal West Dura (Pierce). 2.13 Subcellular fractionation and immunoprecipitation 2.13.2 SPARC:caspase 8 interaction MIP/SP and HCT116 cells were grown until ~80% confluence, incubated with 1000 µM 5-FU and isolated at 4 hours. Cells were separated into nuclear, cytosolic, and membrane fractions using ProteoExtract Subcellular Proteome Extraction Kit (EMD Biosciences Inc.). Fractionation protocols were validated performing immunoblots and incubating with antibodies for proteins found specifically in the various fractions. Total proteins concentrations were measure by Bradford assay (BioRad). To verify the site of interaction of caspase 8 with SPARC, 41  MIP/SP and MIP/ZEO, cells were incubated with antibodies against caspase 8 targeting its Cterminus (Cell Signaling) or N-terminus (Abcam) in vitro for 24 hours at 1.5-3.0 µg prior to collecting and fractionating the cell lysates, for immunoprecipitation, and as well, caspase 3/7 assay (as described previously). 250 µg of the individual cellular fractions were incubated with antibodies against SPARC (10 µg/mL, Haematologic Technologies), caspase 8 (1:100, Cell Signaling Technology (C-terminus) or Abcam (N-terminus) or a non-specific anti-mouse IgG antibody as control (Cell Signaling Technologies), in PBS overnight (4°C) with gentle agitation. Protein:Antibody mixture was then incubated with 30 µL of Protein A: Protein G (Sigma) (1:1) beads for 4 hours (4°C). Proteins were also incubated with EZView Red His-Select HC Nickel affinity gel (Sigma) for immunoprecipitation of His-tagged SPARC protein. For all complexes, beads were washed 5X with PBS, eluted with 40 µL of 2X SDS-Loading Buffer, and used for immunoblotting against SPARC and caspase 8. Samples of the total protein used for the immunoprecipitations were run in a SDS-PAGE gel and Coomasie stained. Representative images were used to demonstrate the equal amount of protein used per immunoprecipitation. 2.13.2 Specificity of the SPARC:caspase 8 interaction MIP/SP and HCT116 cells were grown until ~80% confluence and incubated with 02.3% glutaraldehyde for 2-5 minutes at 37ºC. The reaction was terminated with 1 M Tris-HCl (pH 8.0) and then cell lysates were isolated. Total proteins concentrations were measure by Bradford assay (BioRad). Lysates were incubated with antibodies against caspase 9, caspase 10 (1:100, Cell signaling), or SPARC (1:100, Hematologic Technologies) in PBS overnight (4°C) with gentle agitation. Protein:Antibody mixture was then incubated with 30 µL of Protein A: Protein G (Sigma) (1:1) beads for 4 hours (4°C). For all complexes, beads were washed 5X with PBS, eluted with 40 µL of 2X SDS-Loading Buffer, and used for immunoblotting against caspase 9, caspase 10, SPARC, caspase 8, FADD, or APAF-1. MIP/SP cells treated with 1000 42  µM 5-FU for 4 hours served as positive controls. Samples of the total protein used for the immunoprecipitations were run in a SDS-PAGE gel and Coomasie stained. Representative images were used to demonstrate the equal amount of protein used per immunoprecipitation. 2.14 Animal studies 2.14.1 Tumor xenografts Tumour xenografts harvested from NIH nude mice (6 weeks old, Taconic Laboratories) were used for histology, RT-PCR or immunoblot. 2 x 106 MIP101 cells were injected into the left flank. Once tumours reached 100 mm3, mice were treated with chemotherapy intraperitoneally using 3-week cycle regimen (x2 cycles) as previously described (113). Experimental groups (6 animals/group) for this study included treatment with: SPARC, SPARC + 5-FU, 5-FU only, and saline. In addition, tumour xenografts of MIP/ZEO and MIP/SP cells from mice treated intraperitoneally with either 5-FU (three consecutive days) or saline, were collected after the 1st cycle of treatment and homogenized (Kinematica, POLYTRONAggregate). Lysates were then prepared for immunoblot or RT-PCR. Analysis in this thesis were conducted on 2 of the animals/group. To assess the influence of the various fragments of SPARC on tumor progression and response to chemotherapy in vivo tumour xenograft animal models were also used. Nude mice (6 weeks old, Taconic) were implanted with 1 x 106 (MIP/ZEO, MIP/SP, MIP/SP-N, MIP/SP-F, or MIP/SP-C) cells into the flanks of each animal. Each experimental group consisted of 14 animals/group. Treatment regimens were initiated once the average tumour size reached 75-100 mm3 (V = (width x height2)/2). Tumours were measured using a hand-held caliper (VWR) with concurrent body weight measurements until the completion of the study. 5-FU chemotherapy was provided (intraperitoneally) using a 3-week cycle regimen (6 cycles) as previously described  43  (113). Control animals received saline (also intraperitoneally). All studies were approved by the Animal Care Committee at the University of British Columbia. Doubling time was averaged from all tumour measurements and calculated as: t1/2 = (t2t1)*ln(2)/ln(volume2/volume1). 2.14.2 Immunohistochemistry Tissue sections were processed for immunohistochemistry based on previously established protocols (113, 117). For CD31 staining, tissue sections were embedded in OCTmedia. Sections were fixed in acetone for 10 minutes at 4°C, washed in 1X PBS, blocked in 2% NCS for 20 minutes at room temperature, and incubated in CD31 antibody (1:100, DAKO) overnight at 4°C. Sections were then washed in 1X PBS, incubated with appropriate secondary (1:500) for 1 hour at RT, and counterstained with DAPI (Molecular probes). A Zeiss Axioplan 2 fluorescence microscope was used for imaging. The number of CD31-positive cells were counted and averaged from three different fields (n=3 independent experiments). 2.15 Statistics Statistical difference between experimental groups were calculated and analyzed using Student’s t-test. Statistical significance was defined as p < 0.05, using Smith’s Statistical Package.  44  Figure 2-1. Diagram of constructs used in this thesis. The figure below summarizes the features of the constructs used in this thesis.  45  Table 2-1. Primer sequences for site-directed mutagenesis primers Sense Anti-sense N-terminal domain of 5’ tccaccttca tcctcttaaa 5’ tttaagagga tgaaggtgga SPARC AATCGGTGTC cctgtcctaa 3’ CCATTTCCAT gcttctcctc 3’ FS-like domain of SPARC 5’ gtcgttactg ttgttctgga 5’ tccagaacaa cagtaacgac CGTGTCGAGG gtgtggaaga 3’ AGGTTCGAGT aaacggtgtt 3' EC domain of SPARC 5’ ttggaagact gactccttcgt 5’ acgaagactc agtcttccaa TCCATCTAA caacaggagt 3’ TCAGGTCCAT aggtactcc 3’ DEDI domain of caspase 8 5’ tctttagaaa tactataacc 5’ ggttatagta tttctaaaga AGGTACCATC cgacttcagg 3’ AGGTACCGTG tagaccggag 3’ Putative binding domain of 5’ ttgac ctgtcacttc 5’ tctagaagtgacag caspase 8 tagaAATAGC ggagttcaag 3’ gtcaacaagg ggtt 3’ DEDII domain of caspase 8 5’ gagatagtct aaagtcttct 5’ agaagacttt agctatctc CCTAGTACAG gtactgggac 3’ GAGCTAACTC ccagaaaatt 3’  46  Table 2-2: Amino acid mutation sequences used in this thesis. Name  SP-Nmut1  Mutant (wild type amino acids sequence Δ mutant amino acids sequence) (amino acid numbers) DGAE Δ NRCP (57-70)  SP-Nmut2  AEET Δ PISI (69-62)  SP-Nmut3  EA Δ SS (68-69)  SP-Nmut5 SP-Nmut6 SP-Nmut7 SP-Nmut8 SP-Fmut1  EA Δ SS (68-69) EVG Δ VHL (51-53) EEV Δ RLQ (64-66) AP Δ DA (18-19) FDSSCH Δ SCRGEY (119-124)  SP-Fmut2  SSCHF Δ SSGSS (121-125)  SP-Fmut3  MCVC Δ DHHH (92-95)  SP-Fmut5 SP-Fmut6 SP-Fmut7 SP-Fmut8 SP-Cmut1  MCVC Δ DHHH (92-95) SNDNKT Δ F-FPKT (113-118) KGH Δ --- (137-139) FATKD Δ PVPSW (126 – 130) VTL Δ --- (166-168)  SP-Cmut2  TEKQ Δ GPPL (189 – 192)  SP-Cmut3  LARDF Δ RINRG (216-220)  SP-Cmut5 SP-Cmut6 SP-Cmut7 DEDIm  VTL Δ --- (166-168) TEKQ Δ GPPL (189 – 192) LARDF Δ RINRG (216-220) YLNTRKE Δ AKRTEFQ (137-143)  PBm  ELQ Δ KEP (148-150)  Experimental Usage (Chapter X)  Cell Viability assays (Chapter 4 and 5) Immunoprecipitations (Chapter 4) Caspase 3/7 assays (Chapter 4 and 5) Cell Viability assays (Chapter 4 and 5) Immunoprecipitations (Chapter 4) Caspase 3/7 assays (Chapter 4 and 5) Cell Viability assays (Chapter 4 and 5) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4 and 5) Immunoprecipitations (Chapter 4) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 4 and 5) Immunoprecipitations (Chapter 4) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 4 and 5) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 5) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 5) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 5) Caspase 3/7 assays (Chapter 5) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Cell Viability assays (Chapter 4) Caspase 3/7 assays (Chapter 4) Immunoprecipitations (Chapter 4) Cell Viability assays (Chapter 4) Caspase 3/7 assays (Chapter 4) Immunoprecipitations (Chapter 4)  47  DEDIIm  YEEFSKE Δ DDINLSF (237-243)  Cell Viability assays (Chapter 4) Caspase 3/7 assays (Chapter 4) Immunoprecipitations (Chapter 4)  48  Table 2-3. Primer sequences used for RT-PCR  Gene  Forward Primer  SPARC  5'- CAT CTT CCC TGT ACA CTG - 3' 5’ATCACAGACTTTGGACAAAG TTTA-3’ 5’AAGCTTCTGATTATTGATTCA AACC-3’ 5’TGTGCAGCATTTAACGTCAT ATGT-3’ 5' TGTGTGATAAACGGTGGAGA A 3' 5'- GCCACGGCTGCTTCCAG 3'  caspase 8 caspase 10 FADD Csp8promoter (for ChIP) β-actin  Reverse Primer  Cycles  Size (bp)  5'- ATG GGG ATG AGG GGA G - 3' 5’TCTGAATCAGTCTCAACAGG TATA-3’ 5’TTCTCTATGTTTCTCAAAAGT TTA-3’ 5’ACGCAGCTTGAGTTCAGAA3’ 5' CAGCCACCAGTCACCTTCTT 3' 5'- GGCGTACAGGTCTTT C - 3'  40  104  36-40  440  36-40  470  36-40  160  40  242  25-30  204  49  Figure 2-2. Levels of SPARC, SPARC related peptides used in this study. Cell lysates from A,B) stable transfectants (in vitro and in vivo) or C-I) transiently transfected cells were isolated 120 hours post-transfection and levels of SPARC and SPARC-related peptides were assayed by ELISA to His. (SP: full length SPARC; SP-N: plasmid expressing the N-terminal domain of SPARC; MIP/SP-Nmut1 or 2: mutant 1 or 2 of plasmid expressing a mutant in the N-terminal domain of SPARC; MIP/SP-F, plasmid expressing the FS-terminal domain of SPARC; MIP/SPFmut1 or 2: mutant 1 or 2 of plasmid expressing a mutant in the FS-terminal domain of SPARC; MIP/SP-C, plasmid expressing the EC domain of SPARC; ZEO: empty vector control; control: no plasmid used). Results represent the mean±s.e. (n=2-3 independent studies). *, statistical difference compared to control, where p < 0.05, Student’s t-test.  50  Table 2-4.ChIP Assay buffers.  Caspase 8 promoter was used for ChIP assay and the sequence is listed in Table 2-3.  51  Figure 2-3. Efficacy of caspase 8 siRNA on gene silencing. To determine the efficacy of genespecific silencing with siRNA, MIP/SP or HCT116 cells were transiently transfected with 40nM of caspase 8 siRNA for 48 hours (similar results seen at 96 hours). Controls included cells exposed to transfection reagents alone (Control) or transfected with scramble sequence (Scramble). Gene silencing was assessed by RT-PCR from purified total RNA extracted from MIP/SP and HCT116 cells.  52  3. A novel interaction between pro-caspase 8 and SPARC enhances apoptosis and potentiates chemotherapy sensitivity in colorectal cancers 3.1 Introduction SPARC is a secreted matricellular glycoprotein (56) involved in development, remodeling, and tissue repair (197). It inhibits cell proliferation, adhesion, and cell-cycle progression (86). Differential expression of SPARC has been observed in various human cancers, and it is unclear why it has variable effects on tumour growth in different tissues (198). However, higher SPARC expression has been observed in human CRCs that are sensitive to chemotherapy, in comparison to therapy-refractory tumours (113). SPARC appears to function as a tumour suppressor in ovarian (107), pancreatic cancers (110), and acute myeloid leukemia cells (111). Moreover, in tumour xenograft models, greater growth of pancreatic cancer cells are observed in SPARC-/mice in comparison to wild type SPARC +/+ (110). One proposed mechanism responsible for retarding the growth of tumours is its ability to enhance apoptosis. This has been demonstrated in studies where exogenous exposure to SPARC resulted in enhanced apoptosis in ovarian cancer cells (107), while its absence endogenously diminished this event (110). Our own laboratory investigated the chemosensitizing properties of SPARC and found it to induce apoptosis in the presence of significantly lower concentrations of chemotherapy in CRCs overexpressing SPARC (113). The signaling events involved in SPARC-mediated apoptosis are the focus of this chapter. The effectiveness of chemotherapy can be assessed by its ability to enhance tumour cell death. This can be accomplished by activating signaling pathways involved in cell death (192). We know that SPARC is capable of promoting tumour regression by activating apoptosis, which may explain its ability to re-sensitize therapy-refractory cancer cells to chemotherapy. A variety 53  of stimuli can trigger apoptosis and two major signaling pathways, “extrinsic” and “intrinsic”, converge biochemically leading to its execution (199-201). The extrinsic pathway is triggered by the activation of death receptors, such as Fas; the tumour necrosis factor-related apoptosisinducing ligand (TRAIL) receptors, DR4 or DR5; or tumour necrosis factor receptor, following binding with their natural ligands (202). This recruits adaptor proteins, such as Fas-associated death domain (FADD), which recruits pro-caspase 8 to form death-inducing signaling complexes (DISCs) (203, 204). Caspase 8 is activated at DISCs, leading to downstream pro-apoptotic events (205). Activation of the initiator caspases 8 and 10 of the extrinsic pathway is associated with the release of its prodomain from catalytically active fragments, which is also observed in other caspases involved in the apoptotic cascade (206, 207). The intrinsic pathway is centered around the mitochondria which is key in regulating the balance between pro- and anti-apoptotic factors, such as anti-apoptotic members Bcl-2, Bcl-XL and pro-apoptotic members Bax, Bak and Bok (208). It can be triggered by a number of stimuli, including agents that cause DNA damage (209) or growth factor deprivation (210). This leads to the permeabilization of the mitochondrial membrane, the release of cytochrome c into the cytosol (211), which then interacts with APAF-1 to recruit caspase 9, resulting in cleavage of executioner caspases and apoptosis (212). The convergence of the extrinsic and intrinsic pathways occurs when caspase 8 activates Bid, a Bcl-2 family member that can trigger downstream targets to initiate the intrinsic apoptotic pathway (169). Our earlier findings that SPARC enhances chemosensitivity by increasing apoptosis led us to investigate the signaling events involved in SPARC-mediated apoptosis in this study. Here, we examined the involvement of SPARC in relation to the extrinsic pathway of apoptosis and showed that there was enhanced activation of this pathway when CRC cells were exposed to chemotherapy in the presence of high levels of SPARC. Even more interesting is our finding of  54  an interaction between SPARC and pro-caspase 8 that augments apoptosis in cancer cells, which begins to explain SPARC’s ability to reverse chemotherapy resistance. 3.2 Results 3.2.1 Over-expression of SPARC in MIP101 cells augment the expression of genes involved in the extrinsic pathway of apoptosis Over-expression of SPARC in MIP 101 CRC cells (MIP/SP) leads to increased sensitivity to 5-FU and CPT-11 chemotherapy by diminishing cell survival and enhancing apoptosis (113). To understand the mechanisms involved in this SPARC-mediated effect, we began by examining the relative contribution of genes involved in apoptosis at the transcriptional level. We noted that caspase 8, 10 and FADD appeared significantly higher in two different clones of MIP/SP cells (Figure 3-1A). The greatest difference in gene expression was observed with caspase 8 and 10, which were ~3.4 and 5.9-fold higher in MIP/SP than MIP/ZEO respectively, thereby suggesting that differential expression of SPARC positively influenced genes involved in the extrinsic pathway of apoptosis. In an earlier study, we observed a greater number of cells undergoing apoptosis when either sensitive MIP101 or 5-FU resistant MIP101 cells (MIP/5FU) were exposed to SPARC in combination with 5-FU in vitro and in vivo (113). Based on these findings, we proceeded to assess the effect of 5-FU exposure on the extrinsic pathway in cells with variable levels of SPARC (highest in MIP/SP, moderate in MIP/ZEO, lowest in MIP/5FU) (113). Higher levels of caspase 8 and 10 gene expression were observed in MIP/SP cells and this increased following exposure to 1000 µM 5-FU (Figure 3-1B). In cells with low SPARC expression, caspase 8 and 10 were not observed either basally or following exposure to 5-FU in either MIP/ZEO or MIP/5FU cells. FADD gene expression increased in all cells following treatment with 5-FU. It was interesting to note that by reducing caspase 8 gene expression with siRNA, there was also a 55  demonstrable reduction in FADD gene expression, but no change was observed with caspase 10 (Figure 3-1C). The elevation in caspase 8 mRNA in cells over-expressing SPARC was interesting, yet the mechanisms involved were unclear. The higher basal levels of caspase 8 mRNA indicated a transcriptional up-regulation in SPARC-over-expressing cells as a potential mechanism. Upregulation of caspase 8 transcription has been shown to occur following binding of transcription factors IRF-1 and IRF-2 to caspase 8 promoter (Figure 3-1D) (213-216), and indeed, we show by ChIP assays that in MIP/SP cells, there was significantly more binding of IRF-1 and IRF-2 to the (-162/+76) DNA region at the 5’-end of caspase 8 gene (Figure 3-1E), suggesting a SPARCmediated effect in the up-regulation of caspase 8 transcripts. We also examined whether some of this effect could be due to a decrease in mRNA degradation by harvesting cells at various timedintervals following incubation with actinomycin D, a RNA polymerase II inhibitor. We noted that in lower SPARC-expressing MIP/ZEO cells, a reduction in basal caspase 8 mRNA levels was detected at 4 hours, while in two different clones of MIP/SP and the higher-SPARC expressing HCT 116 cells, a reduction in caspase 8 mRNA level only began to occur after 8-12 hours of exposure. This also suggested the involvement of SPARC in improving the stability of caspase 8 mRNA (Figure 3-1F). 3.2.2 Cells over-expressing SPARC have enhanced activation of the extrinsic pathway of apoptosis This heightened basal expression of caspase 8 and 10 at the transcriptional level in cells overexpressing SPARC translated to the protein level, where MIP/SP cells again showed an abundance of pro-caspase 8 and 10, in comparison to MIP/ZEO and MIP/5FU cells prior to exposure to 5-FU (Figure 3-2A). Conversion of pro-caspase 8 to its cleaved products occurred following exposure to 1000 µM of 5-FU. There were also higher basal levels of Bid in MIP/SP 56  cells, which peaked at 4 hours after incubating with 5-FU, followed by a gradual decline over the next 8 hours. Interestingly, activation of caspases 9 and 3 was prominently observed in MIP/SP cells following 5-FU incubation and to a lesser degree in MIP/ZEO, and even less in MIP/5FU cells. FADD was basally expressed in all cell lines, however, significantly greater phosphorylated FADD was seen in MIP/SP as early as 2 hours after incubation with 5-FU in comparison to either MIP/ZEO (Figure 3-2B) or MIP/5FU cells (Figure 3-2A). This observation that MIP/SP cells were more likely to undergo apoptosis following incubation with 5-FU than MIP/ZEO cells was further supported by significantly higher levels of caspase 3/7 activity in MIP/SP cells at 12 hours after incubation with 5-FU than in MIP/ZEO cells (16397.0±2787.6 vs. 9954.0±1104.8, p<0.05) (Figure 3-2C). 3.2.3 In the presence of SPARC, caspase 8 decreases cell viability in response to 5-FU The relative contribution of the extrinsic pathway in SPARC-mediated apoptosis was examined by reducing caspase 8 expression using siRNA. Following transient transfection with caspase 8 siRNA we noted that caspase 8 gene expression knock-down in MIP/ZEO cells did not affect their response to 1200 µM 5-FU, as cell viability decreased from 87.8±5.8 to 52.1 ±1.7% (p<0.05) in comparison to control cells transfected with scramble siRNA (Figure 3-3A). Interestingly, caspase 8 gene silencing in MIP/SP abolished the effect of 5-FU by preventing a decrease in cell viability after exposure to 5-FU (97.2±1.5% viable untreated cells vs. 97.6±1.4% after 5-FU treatment, p= 0.8783) (Figure 3-3A). This dramatic effect of increasing cell viability after reducing caspase 8 by siRNA persisted following longer exposure to even higher concentrations of 5-FU in MIP/SP and the intrinsically SPARC-expressing HCT116 cells in comparison to cells not transfected with caspase 8 siRNA (Figure 3-3B). However, even in the presence of caspase 8 siRNA, a significant decrease in cell viability could now be observed in  57  both MIP/SP and HCT116 cells following 5-FU, suggesting that a caspase 8-independent mechanism could now be contributing to this reduction in cell viability. These initial results suggested a caspase 8-mediated effect in promoting apoptosis in cells over-expressing SPARC, and that, in the absence of caspase 8, the possibility that the intrinsic pathway could be contributing to this event was raised. Using chemical inhibitors that display some specificity against caspase 8-like (z-IETD-fmk) and caspase 9-like (z-LEHD-fmk·TFA) activities, we again observed that inhibition of caspase 8-like activity affected MIP/SP cells more dramatically than control MIP/ZEO cells. In response to an exposure to 1000 µM of 5-FU for 24 hours only, cell viability decreased in both control MIP/ZEO cells and MIP/SP cells (Figure 33C). However, in MIP/SP cells, pre-incubation with 10 µM of a caspase 8-like inhibitor abolished this decrease in cell viability observed following exposure to 5-FU (Figure 3-3C), as cell viability remained unchanged in the presence or absence of 5-FU, while a decrease in cell viability of 52.2±14.6% (p<0.05) could still be observed in the 5-FU-treated MIP/ZEO cells in the presence of caspase 8-inhibitor. This increase in cell viability in MIP/SP cells despite the presence of 5-FU could be seen following inhibition with as low as 10 µM of caspase 8-like inhibitor, while no such effect could be demonstrated in MIP/ZEO cells despite a higher concentration of the inhibitor (20 µM). Inhibition of the intrinsic pathway with 50 µM caspase 9like inhibitor desensitized both MIP/ZEO and MIP/SP cells to the effects of chemotherapy by preventing a significant decrease in cell viability in response to 5-FU (Figure 3-3D). These results further support a caspase 8-dependent activation of apoptosis in SPARC over-expressing cells, and suggest that the involvement of caspase 9 occurs downstream of this event. To further assess whether the effect of caspase 8 gene silencing was dependent on SPARC expression, we examined the effect of a different agent, CPT-11 on several CRC cell lines expressing variable levels of SPARC: intrinsically high SPARC-expressing HCT116 cells (113) and high SPARC-expressing MIP/SP cells; and compared them to low SPARC expressing 58  MIP/ZEO and even lower MIP/CPT cells. Again, we noted that MIP/SP cells had a reduction in cell viability after treatment with CPT-11 100 µM (100.0±0.0001 % vs. 63.4±4.9%, p<0.05), which was again abolished after transfection with caspase 8 siRNA despite the presence of CPT11 (98.4±3.1% vs. 97.2±4.0%, p=0.8092) (Figure 3-3E). The most interesting finding was that a similar effect of caspase 8 gene silencing was observed with high SPARC-expressing HCT116 cells, with decreased sensitivity to CPT-11 in comparison to untreated cells (Figure 3-3E). For example, 104.8±1.8% viable cells in the presence of caspase 8 siRNA + CPT-11 vs. 108.6 ±3.0% in untreated controls (p=0.48). 3.2.4 Absence of caspase 8 diminishes apoptosis following chemotherapy in high SPARCexpressing cells Given that knockdown of expression of caspase 8 lessened the effect of 5-FU and CPT-11, and enhanced cell viability in MIP/SP cells, we next examined whether this resulted from a reduction in apoptosis. Caspase 3/7 activity was similar in MIP/ZEO cells following incubation with 5-FU regardless of whether cells were transfected with caspase 8 siRNA (2321.3±661.3 RLU) or not (scramble control: 1915.0±661.3 RLU; p=0.27) (Figure 3-4A). However, in MIP/SP cells, an increase in caspase 3/7 activity was only observed in control MIP/SP cells (untreated 327.7± 30.9 RLU vs. 5-FU-treated 5501.0±800.0 RLU, p<0.05), and this effect was abrogated when treated cells were initially transfected with caspase 8 siRNA (untreated 371.0±58.9 RLU vs. treated 291.0±34.7 RLU, p=0.31). No caspase 3/7 activity was observed in MIP/5FU cells following incubation with 5-FU, either following transfection with caspase 8 siRNA or scrambled control (Figure 3-4A). This reduction in apoptosis was further confirmed by TUNEL assay. In the absence of any caspase 8 inhibition, the highest percentage of cells to undergo apoptosis was observed with MIP/SP cells exposed to 5-FU, with 20.7±6.9 % apoptotic cells, in comparison to the untreated group (5.1±2.0%, p<0.05) (Figure 3-4B). However, following 59  caspase 8 gene knock-down, there was a decrease in the sensitivity of MIP/SP cells to 5-FU (Figure 3-4B), with only 0.5±0.5% apoptotic MIP/SP cells detected. A larger percentage of MIP/ZEO cells continued to undergo apoptosis regardless of the presence or absence of caspase 8 gene expression following 5-FU treatment (no transfection with caspase 8 siRNA: 12.2±0.7% apoptotic cells vs. transfection with caspase 8 siRNA: 12.5±2.4%, p<0.05). Therefore, in complete contrast to MIP/SP cells, knock-down of caspase 8 did not inhibit apoptosis in MIP/ZEO cells (Figure 3-4B, C). Resistant MIP/5FU cells did not respond to 5-FU treatment significantly and this was not influenced by changing the expression of caspase 8 gene expression (Figure 3-4B,C). 3.2.5 Is the activity of SPARC intracellular or extracellular? Based on the above results, there appeared to be recruitment of the extrinsic pathway in SPARC-mediated apoptosis, and in particular, this appeared to be associated with a prominent role for caspase 8. It is unclear how this occurs, but we wondered if this effect was mediated through an interaction between SPARC and members of the death-receptor/caspase 8/DISC complex. However, SPARC is a secreted protein, which leads us to the question the localization of SPARC. SPARC has been reported to be found in various components of the cell (both intraand extra-cellular), depending on the cell cycle phase (217). Our results showed that SPARC seems to be associated with caspase 8, an intracellular protein. Therefore, identification of whether the chemosensitizing effect of SPARC was intracellular or extracellular was needed. To answer this question, the signal sequence was deleted from full length SPARC to produce intracellular SPARC clones (SPc1, SPc4, SPc5). First, we verified the localization of the intracellular clones by ELISA assay. Cell lysates and conditioned media were separated to determine levels of SPARC in each fraction. All the intracellular forms of SPARC indeed only expressed SPARC in the cell lysates as shown by ELISA assay (Figure 3-5). These intracellular 60  SPARC clones were transiently transfected in CRC cell lines and assessed for cell viability and caspase 3/7 activity. Transient transfections with SPc1, SPc4, or SPc5 clones (intracellular SPARC) resensitized therapy-refractory CRC cells (MIP/5FU and RKO/CPT), shown by a decrease in cell viability and an increase in caspase 3/7 activity after 5-FU or CPT-11 treatment (Figure 3-6A-D). In sensitive cells, intracellular SPARC still resulted in a further decrease in cell viability (MIP101: 82.60 ± 4.01 (ZEO + 5-FU) vs. 61.63 ± 1.69% (SPc1 + 5-FU), p<0.05; RKO: 83.09 ± 1.70 (ZEO + 5-FU) vs. 51.26 ± 1.87% (SPc1 + 5-FU), p<0.05) (Figure 3-6A, B) and greater levels of caspase 3/7 activity (MIP101: 4851.5 ± 400.69 (ZEO + 5-FU) vs. 7840.5 ± 633.3871 RLU (SPc1 + 5-FU), p<0.05; RKO: 513 ± 10.02 (ZEO + 5-FU) vs. 914 ± 5.0149 RLU (SPc1 + 5-FU), p<0.05) (Figure 3-6C, D). Similar results were also seen in resistant CRC cells (Figure 36A-D). These results show that intracellular SPARC still exhibited its chemosensitizing abilities. Next, we determined if extracellular SPARC showed similar effects. To obtain a form of SPARC that remains extracellular, a biotinylation reaction was performed to covalenty bind rSPARC to biotin. The reagents were used at two different ratios, either 1:20 or 1:50 (rSPARC:biotin), although the number of biotin molecules coupled to SPARC was not measured. To test the chemosensitizing effects of extracellular SPARC, we added the biotinylated SPARC complex (b-SP) with avidin beads to assess cell viability and apoptosis. We found that the ratio of biotin to SPARC of 20 biotin:SPARC (20x b-SP) was insufficient to keep rSPARC completely extracellular. However, the ratio of 50 biotin:SPARC (50X b-SP) was able to keep rSPARC completely extracellular as shown by ELISA assay (Figure 3-5). We used these various forms of biotinylated SPARC to assess for cell viability and caspase 3/7 assays in CRC cells. Our results showed that SPARC must be completely extracellular (i.e. 50X b-SP) to prevent SPARC's chemosensitizing effects in CRC cells (Figure 3-5). In sensitive cells, no 61  further reduction of cell viability was observed after chemotherapy and 50X b-SP exposure compared to control (MIP101: 70.14 ± 1.36 (ZEO + 5-FU) vs. 73.82 ± 1.42% (50X b-SP+ 5FU), p = 0.09; RKO: 74.68 ± 1.95 (ZEO + 5-FU) vs. 83.46 ± 2.99% (SPc1 + 5-FU), p = 0.03) (Figure 3-7 A, B). Changes in caspase 3/7 activity were also not observed after chemotherapy and 50x b-SP treatment compared to control (Figure 3-7C, D). In resistant MIP/5FU and RKO/CPT cells, the entirely extracellular SPARC was unable to re-sensitize cells to chemotherapy (i.e. cell viability was not decreased or apoptosis was not increased compared to empty vehicle) (Figure 3-7A-D). Interestingly, the 20X b-SP, which still expressed SPARC intracellularly, was able to re-sensitize cells to chemotherapy, similar to SP (unmodified extracellular SPARC) (Figure 3-7A-D). These results showed that SPARC's chemosensitizing abilities are only functional when it is intracellular. 3.2.6 SPARC interacts with pro-caspase 8 As we just determined, SPARC's activity to re-sensitize cells only occurs when SPARC is intracellular. Additionally, SPARC-mediated apoptosis appears to involve caspase 8, which led us to determine if SPARC and caspase 8 interacted. We examined this possibility by assessing binding interactions with SPARC by coimmunoprecipitation (co-IP) studies using antibodies to SPARC, and pro-caspase 8. Different subcellular fractions were examined and interestingly, pro-caspase 8 co-IP’d with SPARC in a reciprocal fashion from the MIP/SP (Figure 3-8A) and intrinsically SPARC-over-expressing HCT116 (Figure 3-8B) cell membrane fractions. Moreover, this interaction between pro-caspase 8 and SPARC disappeared when MIP/SP cells were exposed to 1000 µM 5-FU (Figure 3-8A), when a caspase 8 antibody that recognizes the carboxy-terminal sequence of the p18 fragment of the protein was used for the co-IP. However, using a caspase 8 antibody recognizing the Nterminal region of this protein, the interaction between SPARC and pro-caspase 8 could again be 62  detected despite 5-FU exposure (Figure 3-8A). Similar results were obtained with intrinsically high SPARC-expressing HCT116 cells (Figure 3-8B). These results showed that SPARC interacts with the N-terminus of pro-caspase 8 and with 5-FU treatment, SPARC and the Nterminus of caspase 8 still interact (Figure 3-9). 3.2.7 Specificity of the SPARC: caspase 8 interaction Results from this study showed a specific interaction between SPARC and the Nterminus of pro-caspase 8. The specificity of this interaction was in question. Proteins that bind to caspases to inhibit their activity have been identified, but not many have been shown to interact with caspases to promote apoptosis. The known activators of apoptosis are APAF-1 and FADD, which interact with caspase 9 and caspase 10, respectively (218). As there are limited proteins identified to bind with caspases to promote apoptosis, this makes the identification of a caspase 8 interacting partner highly interesting. I questioned if other proteins with a high homology sequence to caspase 8 could interact with SPARC. BLAST identified the amino acid sequences of caspase 10, caspase 7, caspase 3, and caspase 9 to be the most similar to caspase 8. ClustalW alignments of the amino acid sequences showed that caspase 10 has the highest overall degree of homology with caspase 8; however, this was not within the N-terminus, which is functionally the part of SPARC that is of interest (Table 3-1). To determine if there was a specific interaction occurring, immunoprecipitation experiments were performed. Immunoprecipitation experiments of MIP/ZEO and MIP/SP cell lysates with SPARC and caspase 9 (Figure 3-10A) or 10 (Figure 3-10B) did not show any interaction. To exclude the possibility of a transient interaction that is not captured by cell lysates, protein-protein interactions were stabilized by cross-linking prior to use in immunoprecipitations. Results showed that SPARC and caspase 8 still interacted (Figure 3-10E),  63  but SPARC does not bind with either initiator caspase 9 (Figure 3-10D) or 10 (Figure 3-10C). These results show that SPARC specifically interacts with caspase 8 (Figure 3-10E). 3.2.8 SPARC in combination with 5-FU increases caspase 8 expression in tumour xenografts We previously demonstrated that the combination of SPARC and 5-FU treatment conferred the greatest tumour regression in mouse xenografts of MIP101 cells and this correlated with higher numbers of apoptotic cells (113).  We next examined whether the interaction  between SPARC and caspase 8 could also be detected in vivo. Tumour xenografts bearing MIP/SP cells showed higher levels of caspase 8 gene expression than MIP/ZEO cells (Figure 311A). Moreover, in keeping with our in vitro observations, full length and cleaved caspase 8 protein expression was highest in MIP/SP tumours harvested from animals treated with 5-FU (Figure 3-11B). We also examined tumours from MIP 101 mouse xenografts that had been previously treated with a combination of SPARC and 5-FU, or as single agents, for caspase 8 expression and again, only observed higher levels of caspase 8 in tumours harvested from mice that were administered SPARC, and even more significantly following combination treatment with SPARC and 5-FU (Figure 3-11C). This up-regulation of caspase 8 expression in animals exposed to both SPARC and 5-FU appears to be restricted to the tumour xenografts, since livers harvested from these same mice did not have elevated caspase 8 expression (Figure 3-11C).  3.3 Discussion SPARC, best known for its role as a matricellular protein, is expressed in many different cell types (219). Its role in tumourigenesis has been examined in a variety of cancers (106, 113, 123, 220, 221) including hematological malignancies (111), with diverging actions which may be attributed to tissue-related differences. Previously, we demonstrated that SPARC gene and 64  protein expression changes the sensitivity of therapy-refractory MIP101 CRC cells to chemotherapy in vitro and in vivo by enhancing apoptosis and tumour regression (113). Induction of apoptosis in the presence of SPARC has also been observed in ovarian cancer cell lines (107), lung and pancreatic tumours implanted in SPARC+/+ mice (110), and kidney tumour xenografts (222). Our previous observations, together with others, led us to assess the apoptotic signaling events mediated by SPARC, and in particular, the potential involvement of the extrinsic pathway. Several mechanisms, including inhibition of apoptosis, can contribute to tumour progression in therapy-unresponsive tumours.  Most chemotherapies induce apoptosis via the intrinsic  pathway (223). However, there is growing evidence of the involvement of the extrinsic pathway of apoptosis in chemotherapy-induced cell death. In particular, there have been reports of caspase 8 activation in both a death-receptor dependent (224) and independent fashion (225). In non-small cell lung cancer and leukemia, caspase 8 can be activated by chemotherapies such as cisplatin, etoposide, gemcitabine and topotecan, independent of FADD and death receptors (226). Absence or reduced expression of caspase 8 has been correlated with resistance to apoptosis in a variety of tumours, including neuroblastomas, and small cell lung cancers (227). In laryngeal squamous cell carcinoma, cisplatin-resistant HEp-2 cells had reduced caspase 8 activation (228), while doxorubin-resistant MCF-7 breast cancer cell line had defects in caspases 8, 9 and 10 (229). In our study, we show that in SPARC-over-expressing MIP 101 cells (MIP/SP) that were more sensitive to chemotherapy, higher levels of expression of the genes involved in the extrinsic pathway of apoptosis (caspase 8, 10, FADD) were noted, which increased even more significantly following exposure to 5-FU. This elevated expression at the mRNA level translated to the protein level, where similar increases were seen in MIP/SP cells. Interestingly, higher caspase 8 levels was also observed in tumour xenografts that experienced the most dramatic 65  regression in mice treated exogenously with SPARC and 5-FU, further supporting our in vitro observations. Activation of caspase 8 appears to involve and converge downstream with the mitochondrial pathway through activation of Bid, as higher levels of the full-length protein were observed in MIP/SP cells, which gradually declined following incubation with 5-FU, as one would expect with activation and truncation of Bid. The downstream effects can be seen through a significantly greater activation of caspase 9 and 3 in MIP/SP cells than in MIP/ZEO, while no significant activation is seen in MIP/5FU. Therefore, our findings indicate that in the presence of SPARC, apoptosis occurs as a caspase 8-dependent event which later converges with the intrinsic pathway to involve caspase 9 (Figure 3-12A). However, in the absence of SPARC, apoptosis occurs by activating the intrinsic pathway independently of caspase 8 (Figure 3-12B). This recruitment of caspase 8 in the presence of SPARC, in addition to the involvement of the intrinsic pathway further augments and amplifies the death signal induced by chemotherapy, and explains in part, the chemosensitizing effect of SPARC. In this chapter, we also identified that SPARC's ability to re-sensitize CRC cells only occurs when SPARC is internalized. To date, the mechanism of how SPARC crosses the cell membrane has not been identified. Recently, α5β1 in adipose stromal cells (230) and β1 integrin in lens epithelial cells (231) have been identified as potential SPARC receptors. The identification of how SPARC enters a cell would provide great insight into the biological functions of SPARC. We did not test for the functionality of SPARC crosslinked to biotin. The functionality of SPARC may be greatly perturbed by the biotin; thus, inhibiting its functionality. Additionally, we did not test the effects of avidin beads on cell growth. These controls are necessary to fully interpret the data. Therefore, these results showing that biotinylated SPARC did not carry out the same functions could be interpreted to suggest that SPARC must be intracellular to have its effect. However, it is possible that SPARC has lost functionality after cross-linking to biotin.  66  We further demonstrated that this caspase 8-dependent event may occur because of an interaction between SPARC and pro-caspase 8, taking place at the p43/p41 fragment of caspase 8 containing the N-terminal DED-domains and the catalytic site, since this interaction can no longer be observed upon cleavage of caspase 8 following exposure to chemotherapy when a caspase 8 antibody recognizing the C-terminus was used. However, members of the deathreceptor/caspase 8/DISC are often associated with each other and must be further dissected to determine if other members are present with the SPARC:caspase 8 interaction. As well, in vitro exposure to caspase 8 antibody targeting the N-terminus also interferes with its ability to interact with SPARC, and thereby abrogating its effect on apoptosis. These observations lend further support that the interaction between the N-terminus of caspase 8 and SPARC is important in mediating apoptosis. We questioned if SPARC interacts with a larger complex that contains pro-caspase 8. This is particularly relevant, as studies have shown that Bcl-2 can inhibit caspase 8 mediated apoptosis by binding and inhibiting pro-caspase 8 (232). Interestingly, exposure to SPARC can induce autosomal dominant polycystic kidney disease (ADPKD) cyst-lining epithelial cells to undergo apoptosis as a result of a reduction in the ratio of Bcl-2 and Bax (219). Therefore, one wonders if an interaction between SPARC and pro-caspase 8 may prevent its sequestration by Bcl-2, thereby enhancing apoptosis, as observed in this chapter. The presence of higher caspase 8 gene expression in cells over-expressing SPARC was also particularly interesting. The transcriptional regulation of caspase 8 involves binding of transcription factors IRF-1 and IRF-2 to the interferon sensitive response element site within its promoter (215, 216). Interestingly, our ChIP assay showed greater binding of IRF-1/2 to the promoter regions of caspase 8 gene in cells over-expressing SPARC, thereby indicating that, in the presence of higher levels of SPARC, there is transcriptional up-regulation of caspase 8 in these cells. These results are intriguing, as we also showed that SPARC's chemosensitizing 67  effects occur only when SPARC is intracellular. Thus, as intracellular SPARC may affect the chemosensitivity of cells in several ways: by increasing caspase 3/7 activity and reducing cell viability, or it can also result in regulating caspase 8 transcription. In addition, there also appeared to be diminished caspase 8 mRNA stability in cells with reduced SPARC expression, as higher SPARC-expressing cells only began to show a decrease in basal caspase 8 mRNA levels after 8-12 hours after exposure to actinomycin D. This time frame is comparable to those observed in MCF-7 cancer cells, with caspase 8 mRNA levels diminishing after 6-12 hours of actinomycin D exposure (215). These results suggest a SPARCmediated effect in the regulation of caspase 8 mRNA, and further studies are required to further elucidate these interesting findings. Our current observations support a caspase 8-dependent event in SPARC-mediated apoptosis (Figure 3-12A) and we show that this occurs as a result of its interaction with procaspase 8. This mechanism may also prove to be important in other malignancies where SPARC has also been shown to enhance tumour regression and to induce apoptosis (107, 110, 111, 233). The results presented in this study are exciting as they provide an initial insight into a potential mechanism by which SPARC mediates its chemosensitizing effect in cancers refractory to therapy. The caspase 8-dependent signaling events mediated by SPARC to promote tumour regression not only supports its therapeutic potential in cancer, but also allows us to identify potential downstream targets that can be similarly exploited to enhance tumour regression and overcome chemotherapy resistance in patients with advanced, therapy-refractory CRC. From this chapter, SPARC-mediated apoptosis was shown to occur by activating the extrinsic pathway and enhancing the intrinsic pathway of apoptosis. Since SPARC contains three biological domains, each with its own modular function (62), chapter 4 examined if one of these domains was responsible for SPARC-mediated apoptosis.  68  Figure 3-1. The presence of higher levels of SPARC is associated with greater expression of genes involved in the extrinsic pathway of apoptosis following exposure to 5-FU. A) Basal levels of expression of genes involved in the extrinsic pathway of apoptosis were assessed in MIP/ZEO and MIP/SP cells (2 independent clones) by RT-PCR. B) Changes in the expression of genes involved in apoptosis were assessed following treatment with 5-FU after 4 hours at the RNA level. For caspase 8, PCR was performed at 40 cycles in (A), or (B) 36 cycles. C) The effect of caspase 8 gene silencing in MIP/SP cells on caspase 10 and FADD mRNA levels were determined by RT-PCR following transient transfection with 40 nM caspase 8 siRNA. D) Schematic representation of the 5’- region (-162/+76;) spanning the IRF-1/2 binding regions of the caspase 8 promoter and part of exon 1. Primer positions (forward-p, reverse-p) and amplification products for chromatin immunoprecipitation (ChIP) assays are also indicated. E) ChIP assays to detect binding of transcription factors IRF-1 and IRF-2 to caspase 8 promoter. F) Assessment of caspase 8 mRNA stability following incubation with actinomycin D for 0-20 hours (PCR, 40 cycles). All results representative of 3-4 experiments.  69  70  Figure 3-2. The presence of higher levels of SPARC is associated with greater levels of apoptosis. A, B) Changes in the expression of genes involved in apoptosis were assessed following treatment with 5-FU after various timed intervals at the protein level. Total proteins were isolated from cells harvested at 0-12 hours following treatment with 1000 µM 5-FU, and probed for various proteins shown by immunoblots (A – activated caspases are represented by their cleaved fragments, with the exception of caspase 10 as antibody does not recognize the cleaved fragment), and also caspase 3/7 activity (C) (all results representative of 3-4 experiments). * p<0.05, Student’s t-test, in comparison to untreated controls, or otherwise noted.  71  Figure 3-3. Inhibition of caspase 8 increases cell viability in CRC cells with greater SPARC expression. The relative contribution of caspase 8 was assessed using caspase 8 siRNA. A) MIP/ZEO and MIP/SP cells were transiently transfected with 40 nM caspase 8 siRNA (or scramble control) for 48 hours, followed by treatment with 500 (+), 1000 (++), or 1200 (+++) µM 5-FU for an additional 48 hours. B) MIP/SP and HCT116 cells were also transiently transfected with 40 nM caspase 8 siRNA (or scramble control) for 48 hours, followed by treatment with 1.5, 2.0, or 2.5 mM 5-FU for 48-72 hours. Inhibitors with (C) caspase 8-like (zIETD-fmk) or (D) caspase 9-like (z-LEHD-fmk·TFA) activity were used to determine the relative contribution of the extrinsic (C) or intrinsic pathway (D) in SPARC-mediated apoptosis. C) Cell viability of MIP/ZEO and MIP/SP cells treated with caspase 8-like (0-20 µM) or D) caspase 9-like (0-50 µM) inhibitors; and 1000 µM 5-FU for 24 hours was measured with an MTS assay. E) In addition, the effect of caspase 8 gene silencing on cells with variable SPARC expression (MIP/ZEO, medium; MIP/SP, high; MIP/5FU, low; and intrinsically high SPARC expressing HCT116 cells), was assessed following exposure to CPT-11 (100 µM) for 36 hours. The percentage of viable cells was calculated based on the OD490 reading of non-treated control cells. All results represent mean±s.e (n=3-4 independent studies). * p<0.05, Student’s t-test, in comparison to untreated controls, or otherwise noted.  72  73  74  Figure 3-4. Inhibition of caspase 8 gene expression with siRNA enhances survival of cells expressing higher levels of SPARC following exposure to 5-FU. MIP/ZEO, MIP/SP, and MIP/5FU cells were transiently transfected with 40 nM scramble sequence (control), or caspase 8 siRNA for 36 hours followed by incubation with 1000 µM 5-FU for 36-48 hours. The effect of caspase 8 gene silencing on apoptosis was assessed by (A) caspase 3/7 assay, and (B, C) TUNEL assay. The percentage of apoptotic cells was determined by counting the number of TUNELpositive apoptotic nuclei (green) relative to the number of DAPI-positive nuclei (blue) from 4 different fields, with a minimum of 100 cells/field, by microscopy. Images are representative of 4 independent experiments. Statistical difference is represented by “*”, where p < 0.05, Student’s t-test.  75  76  77  Figure 3-5. ELISAs of intracellular and extracellular SPARC clones. To verify the subcellular localization of the intracellular and extracellular SPARC forms, ELISA assays were performed. A) MIP101 cells were transiently transfected with the full length SPARC (SP), SPARC clones without the signal sequence (three independent clones: SPc1, SPc4, SPc5), or empty vector control (EV) (all pcDNA 3.1 vectors) and grown to 90% confluence. B) MIP, MIP/5FU, RKO, and RKO/CPT cells were also treated with exogenous full length SPARC (SP) biotinylated SPARC (20x b-SP, 50x b-SP), or empty vehicle (EV) and grown to 90% confluence in the presence of avidin beads. Conditioned Media (□) and cell lysates (■) were isolated from both studies and used for ELISA assay with anti-SPARC antibody (1:5000) and quantification was determined based on the A650 reading of control cells. All of the results represent the mean±s.e. (n=2-3 independent studies). *, statistical difference, where p<0.05, Student’s t-test.  78  Figure 3-6. Intracellular SPARC enhances chemosensitivity. The contribution of intracellular SPARC to enhance chemosensitivity was determined using cellular viability and apoptosis assays. A, B) MIP101 and MIP/5FU (A) and RKO, RKO/CPT (B) CRC cells were transiently transfected with full length SPARC (SP), SPARC clones without a signal sequence (SPc1, SPc4, SPc5), or empty vector control (EV) for 48 hours, followed by treatment with 0 (■) or 1000 µM (□) 5-FU (for MIP101 derived cells) or 0 (■) or 36 µM (□) CPT-11 (for RKO derived cells) for an additional 24 hours. Cellular viability was measured by MTT assay. The percentage of viable cells was calculated based on the A450 reading of nontreated controls. C, D) Cell lysates were also isolated after chemotherapy exposure and apoptotic levels were assessed through by caspase 3/7 assay in MIP101 and MIP/5FU (C) or RKO and RKO/CPT (D) cells. Results represent the mean±s.e. (n=3-4 independent studies). *, p<0.05, Student's t-test, in comparison to untreated controls, or otherwise noted.  79  Figure 3-7. Extracellular SPARC does not enhance chemosensitivity. The contribution of extracellular SPARC on chemosensitivity was determined using cellular viability and apoptosis assays. A, B) MIP101 and MIP/5FU (A) and RKO, RKO/CPT (B) CRC cells were exposed to full length SPARC (SP), biotinylated SPARC (20X b-SP, 50X b-SP) or control (empty vehicle) for 48 hours, followed by treatment with 0 (■) or 1000 µM (□) 5-FU (for MIP101 derived cells) or 0 (■) or 36 µM (□) CPT-11 (for RKO derived cells) for an additional 24 hours. Cellular viability was measured by MTT assay. The percentage of viable cells was calculated based on the A450 reading of nontreated controls. C, D) Cell lysates were also isolated after chemotherapy exposure and apoptotic levels were assessed through by caspase 3/7 assay in MIP101 and MIP/5FU (C) or RKO and RKO/CPT (D) cells. Results represent the mean±s.e. (n=3-4 independent studies). *, p<0.05, Student's t-test, in comparison to untreated controls, or otherwise noted.  80  Figure 3-8. Interaction between pro-caspase 8 and SPARC is detected in the cell membrane fraction. Co-localization of SPARC and pro-caspase 8 was determined by coimmunoprecipitation (co-IP) studies following fractionation of (A) MIP/SP or (B) HCT116 cells into nuclear (N), cytoplasmic (C) and membrane fractions (M) with our without exposure to 1000 µM of 5-FU for 4 hours. 250 µg of protein from each cellular fraction were immunoprecipitated with antibodies to SPARC, caspase 8 (C-terminus and N-terminus), 6XHis (to detect His-tagged recombinant SPARC), or IgG (non-specific control), and immunoblotted for SPARC or caspase 8. To determine the effect of chemotherapy on the interaction between SPARC and pro-caspase 8, cellular fractions obtained from cells treated with 5-FU were co-IP with antibodies against SPARC and the western blot probed for pro-caspase 8. With exposure to 5-FU, no interaction between SPARC and the C-terminus of caspase 8 is observed, and only the interaction between the N-terminus of caspase 8 and SPARC is seen following cleavage of procaspase 8. Input = protein loaded per immunoprecipitation. All results representative of n = 34 independent experiments.  81  Figure 3-9. The interaction between pro-caspase 8 and SPARC occurs at the N-terminus of pro-caspase 8. Results provide a model for the site of interaction between SPARC and procaspase 8 .  82  Table 3-1 Homology between caspase 8 and other initiator caspases. Score from amino acid sequence alignments by ClustalW. Caspase 10 14 9 2 1 4 5  Score 34 27 26 17 17 14 13  83  Figure 3-10. SPARC interacts specifically with caspase 8. To determine if SPARC interacted with any other members of the caspase family immunoprecipitation experiments were performed. Co-localization of SPARC with caspase 9 or caspase 10 was determined by co-IP studies using whole protein extracts. A, B) No interaction was observed in either MIP/ZEO or MIP/SP cell lysates with A) caspase 9 or B) caspase 10. C, D) To determine if the interaction was transient, MIP/ZEO and MIP/SP cells were treated with glutaraldehyde to stabilize any transient interactions and an interaction with D) caspase 9 or C) caspase 10 was still not observed. E) Co-localization studies with the glutaraldehyde-fixed MIP/ZEO and MIP/SP cell lysates with known interacting partners (SPARC and caspase 8, caspase 10 and FADD, and caspase 9 and APAF-1) was determined by co-IP studies and interactions with the known interacting partners was observed. Positive controls were used for all studies to verify immunoblotting procedure. Input = protein loaded per immunoprecipitation. Results representative of n = 2-3 experiments.  84  Figure 3-11. Caspase 8 is up-regulated in response to higher levels of SPARC in tumour xenografts following exposure to 5-FU in vivo. Tumour xenografts of MIP/ZEO and MIP/SP cells were harvested after the first cycle of 5-FU treatment (intraperitoneally) and assessed by (A) RT-PCR and (B) immunoblot. C) Paraffin-embedded tumour xenografts of MIP 101 cells harvested from mice following treatment with either saline (control), 5-FU, SPARC alone or in combination with 5-FU. Tissue sections were stained for caspase 8 (Sections 6 µm thick, magnification 40X). Results representative of n = 2-3 independent experiments.  85  Figure 3-12: SPARC-induced apoptosis: A model. In this study, we demonstrate how SPARC influences apoptosis. A) In the presence of SPARC, apoptosis occurs as a caspase 8 dependent event through an interaction (*) between SPARC (yellow) and pro-caspase 8 (green) that results in cleavage of pro-caspase 8 (blue) following exposure to chemotherapy (5-FU). This leads to the activation of the mitochondrial pathway of apoptosis via Bid. This SPARC-mediated event enhances apoptosis and augments chemosensitivity in CRC cells. B) At lower SPARC levels, apoptosis only occurs through the intrinsic pathway, independent of caspase 8.  86  4. SPARC’s N-terminal domain interferes with Bcl-2 and procaspase 8 interaction and promotes apoptosis and tumour regression in vivo 4.1 Introduction Many pathological conditions arise because of abnormal regulation in cellular activities, such as apoptosis, that disrupt the fine balance between cell survival and death.  This  deregulation can contribute to cancer initiation, progression, and even influence a tumour’s response to chemotherapy. Key to the initiation and execution of apoptosis are caspases and members of the Bcl-2 family of proteins. Caspases, as important mediators of apoptosis, belong to a family of cysteine proteases that cleave their substrates with specific Asp-containing tetrapeptide motifs (234). Caspase 8 is a member of this family better known for its involvement in the extrinsic or death-receptor pathway of apoptosis. Upon binding of ligands to cell surface death receptors (members of the TNF family of receptors), caspase 8 is recruited to the intracellular domain of death receptors and proteolytically activated. This, in turn, begins a cascade that involves the direct activation of effector caspases (caspase-3, 6, or 7) leading to apoptosis. The extrinsic pathway, which is classically activated following ligand binding to cell surface death receptors, is distinct from the intrinsic pathway because of the involvement of the mitochondria and Bcl-2 family of proteins in the latter. The intrinsic pathway can be activated by cellular cues, for example, following DNA damage, which then affects the Bcl-2 family of proteins, leading to changes in mitochondrial membrane permeability, release of cytochrome-c and caspase-9 activation. Members of the Bcl2 family have similar structural homology and conserved sequences (BH-containing domains). Examples include anti-apoptotic members Bcl-2, Bcl-Xl; pro-apoptotic proteins BAX, BAK; and BH-3 only proteins that promote apoptosis by binding to Bcl-2 proteins (BAD, BID, and PUMA) (235). Recent studies have shown that in certain cells, activation of caspase 8 can involve the 87  intrinsic pathway by mediating cleavage and activation of BID (236). In chapter 3, it was demonstrated that the convergence of these two pathways can also be mediated by SPARC (secreted protein and rich in cysteine) (196). In a cellular environment rich in SPARC, and in response to cytotoxic influences, this protein interacts directly with the N-terminus of procaspase 8 to enhance apoptosis. SPARC, as a matricellular protein, does not directly contribute to the structural scaffold of the extracellular matrix, but instead, modulates cell-cell and cell-matrix interactions during development and tissue remodeling (237). SPARC can inhibit cell proliferation, adhesion, and cell-cycle progression (86). It also has the ability to promote apoptosis in cancers of the ovary, pancreas, lung and the colorectum (107, 110, 113). The SPARC family of proteins contains seven members, all of which share three similar domains: (1) N-terminus, (2) follistatin-like, and (3) C-terminus (61, 238). In SPARC, the N-terminus contains the immunodominant epitopes and binds to hydroxyapatite (238); the follistatin-like domain contains cysteine-residues and an N-linked complex carbohydrate (67, 86); while the C-terminus contains the extracellular Ca2+binding module (102). We hypothesize that one of the biological domains is responsible for the pro-apoptotic activity of full length SPARC. In the current chapter, we show that a smaller peptide of SPARC is pro-apoptotic and its over-expression augments the sensitivity of tumour xenografts to chemotherapy in vivo. Moreover, we demonstrated that its dramatic effect in enhancing chemosensitivity in therapy-refractory cancers lies in its ability to interfere with the interaction between caspase 8 and Bcl-2.  88  4.2 Results 4.2.1 Effect of different SPARC domains on cell viability We previously showed that exposure to higher levels of SPARC enhances apoptosis and significantly reduces cell viability in colorectal cancer (CRC) cells (113, 196). In order to determine if a specific region within SPARC may be responsible for this pro-apoptotic activity, sensitive and resistant colorectal (MIP), pancreatic (MiaPaca), and breast (MCF7) cancer cells were transiently transfected with vectors containing only the N-terminus (SP-N), the follistatinlike (SP-F), or the C-terminus (SP-C) domains of SPARC (Figure 4-1A). In all chemotherapy naïve and resistant cells examined, transient over-expression of SP-N reduced cell viability (Figure 4-1B) in response to chemotherapy. In sensitive cells, such as MIP, this represented an additional decrease in viability of 37.18±3.65 % (p<0.005) following transfection with SP-N (Figure 4-1B, i), in comparison to empty vector controls. Even more significant is that SP-N also decreased cell viability in all resistant cells examined (Figure 4-1B ii, iv, vi, vii). Interestingly, we also noted an effect following SP-F transfection (Figure 4-1Bii, vi, vii), with the exception of MIP/CPT (Figure 4-1Biv). In contrast, there was no effect with SP-C in resistant cell lines. 4.2.2 SPARC-N confers increased sensitivity to chemotherapy Based on these initial results suggesting that a recombinant protein containing only the N-terminus or the FS-domains of SPARC was capable of diminishing viability in not only sensitive colorectal, pancreatic and breast cancer cells, but also in their chemo-resistant counterparts, we decided to further evaluate this by mutating the N-terminus or FS-domains. We performed random site-directed mutagenesis, within the various SPARC domains, to screen potential amino acids that are involved in SPARC's chemosensitizing abilities (Table 2-2). Our results show that the mutant forms of the NT-domain no longer promoted a chemosensitizing effect. In resistant MIP/5FU cells, transient over-expression of two different mutants of SP-N 89  (SP-Nmut1, SP-Nmut2) failed to increase their sensitivity to 5-FU (Figure 4-2), while overexpression of wild-type SP-N was able to decrease cell viability by 24.48±0.82 % (p<0.005) in response to 5-FU. Similar observations were made with other resistant cells (Figure 4-2). The amino acids 56-69 of SP-N domain appeared to be crucial for its chemosensitizing ability, as mutations outside of this region (eg. amino acids 51-53, 18-19) did not affect its biological function as these other mutant forms of SP-N continued to be effective in augmenting chemosensitivity in therapy-refractory cells (Figure 4-3). Transient over-expression of SP-F consistently conferred greater chemosensitivity to therapy-resistant cells, but not mutant SP-F (SP-Fmut1, SP-Fmut2) (Figure 4-2). Mutations in the other areas of SP-F (amino acids 92-139) also failed to increase their chemosensitivity to 5-FU (Figure 4-3). Consistent with our previous results, the SP-C domain and its mutants did not enhance the chemosensitivity of CRC cells (Figure 4-3). Theses results support that the chemosensitizing ability if through the SP-N domain, specifically amino acids 56-69. 4.2.3 Effect of SPARC domains on cell proliferation and apoptosis This negative effect of the NT- and FS-domains on cell viability led us to further evaluate them using stable clones of MIP101 cells over-expressing either the NT- (MIP/SP-N), the follistatin-like (MIP/SP-F), or the extracellular (MIP/SP-C) domains. MIP/SP, MIP/SP-N, and MIP/SP-F cells were the most sensitive to 5-FU with viability decreasing by 59.2±2.2% (p<0.001), 60.2±1.2% (p<0.001), and 61.8±3.0% (p<0.001) respectively, in comparison to untreated controls. Control MIP/ZEO cells responded to 5-FU by decreasing cell viability by 47.3±4.7% (p<0.001) only, in comparison to untreated cells. MIP/SP-C cells had a reduction in viability following 5-FU of only 50.7±3.7% (p<0.001), which was not significantly different from MIP/ZEO treated cells (Figure 4-4A).  90  This enhanced response to chemotherapy in cells over-expressing either the NT- or the FS-domains of SPARC was interesting. We and others had previously reported that SPARC could inhibit cell proliferation (66, 86, 239), increase apoptosis (107, 110, 113), and delay cell cycle progression (86). We therefore began to further examine the biological effects of these different fragments. We noted that unlike cells over-expressing SPARC, cell proliferation (as measured by Ki-67 positivity) did not change in cells over-expressing the individual domains in comparison to control MIP/ZEO cells (Figure 4-4B).  However, in comparison to control  MIP/ZEO cells, there was a 1.62-fold increase in the percentage of MIP/SP-N cells undergoing apoptosis in response to 5-FU, a response very similar to MIP/SP cells (Figure 4-4C). No significant increase in apoptotic cells were observed in MIP/SP-F or MIP/SP-C following exposure to 5-FU. We previously reported that SPARC-mediated apoptosis involved the activation of the extrinsic pathway, via caspase 8, with subsequent convergence with the intrinsic pathway following activation of BID (196). The possibility that the NT-domain may engage in a similar mechanism to enhance apoptosis was entertained. Interestingly, caspase 8 activity was only observed in MIP/SP-N cells, prior and following exposure to 5-FU (Figure 4-4D). There was also higher expression of cleaved BID and cleaved caspase-3 (after 8 hours of 5-FU) in MIP/SPN cells relative to MIP/SP-F and MIP/SP-C cells. These results suggest involvement of the extrinsic pathway in cells over-expressing the NT-domain, that is consistent with a reduction in caspase 3/7 activity following caspase 8 knock-down by siRNA in MIP/SP-N cells, but not in MIP/SP-F or MIP/SP-C. MIP/ZEO, MIP/SP-F, and MIP/SP-C cells remained responsive to 5-FU even after transfection with caspase 8 siRNA, with an increase in caspase 3/7 activity by: 71.41±1.57% (p<0.001); 93.13±2.95% (p<0.001); and 90.92±4.15% (p<0.001) respectively, in comparison to untreated controls. However, MIP/SP, MIP/SP-N, and HCT116 cells were no longer responsive to 5-FU after caspase 8 gene silencing (p>0.05) (Figure 4-5). 91  4.2.4  Tumour xenografts over-expressing SPARC and SPARC-N are more  chemosensitive Thus far, the results of the in vitro studies indicate that the NT-domain was capable of conferring greater sensitivity to chemotherapy. Only cells over-expressing the NT-domain, and not others, underwent more extensive apoptosis following cytotoxic exposure. Next, we examined if MIP/SP-N cells were also more sensitive to chemotherapy in vivo. Xenografts bearing cells stably transfected with either the NT-domain or SPARC were the most responsive to 5-FU treatment, as tumours remained <400 mm3 after 41 days of treatment, while salinetreated animals harbored tumours greater than 1144.1±181.3 mm3 (Figure 4-6A, B). Tumour xenografts of cells stably expressing other domains of SPARC or empty vector control did not differ in size between the treatment and control groups. Results show significantly longer tumour doubling times of MIP/SP and MIP/SP-N xenografts following treatment with 5-FU, representing a reduction of 76% (p<0.001) and 42% (p<0.001), respectively. In control animals, a longer doubling time was also observed in xenografts of MIP/SP (p<0.001) and MIP/SP-N (p<0.001) in comparison to MIP/ZEO. Xenografts of MIP/SP-C had a similar tumour doubling time as MIP/ZEO, while MIP/SP-F had a faster growth rate (p<0.001) (Figure 4-6C). These observations correlated with a greater percentage of cells undergoing apoptosis in xenografts of MIP/SP (22.2+2.4% vs. 4.2+3.1% in control, p=0.01) and MIP/SP-N (31.3+3.2% vs. 4.3+0.9% in control, p<0.005) cells following treatment with 5-FU in comparison to their saline-treated counterparts (Figure 4-7A, B), while this increase in apoptosis was not observed in xenografts of other cell lines despite treatment with 5-FU. Angiogenesis also appeared to be affected, as xenografts of MIP/SP and MIP/SP-N cells showed significantly fewer CD31+ staining than xenografts of MIP/ZEO, MIP/SP-F or MIP/SP-C (p<0.05, Figure 4-7C). Although SPARC has been previously shown to promote apoptosis and inhibit angiogenesis (113, 222),  92  these results demonstrated that the N-terminus domain of SPARC alone has similar biological activities by also promoting apoptosis and inhibiting angiogenesis. 4.2.5 The N-terminus of SPARC interacts with procaspase 8 The ability of SPARC to promote greater apoptosis in vitro and in vivo, following exposure to cytotoxic agents (113) results, in part, from its interaction with procaspase 8 leading to the activation of the extrinsic pathway of apoptosis (196).  We examined if a similar  interaction could be identified between the different SPARC domains and found only the NTdomain to interact with procaspase 8 in a reciprocal fashion using antibodies against caspase 8 and His6 tag of SPARC constructs by co-immunoprecipitation studies (Figure 4-8A). This interaction was only detected in the membrane fraction of MIP/SP-N cell lysates (Figure 4-8B), and persisted in the presence of 5-FU, thereby confirming that this region interacts with procaspase 8 (196). As further evidence, the presence of mutations within the NT-domain abolished this interaction (Figure 4-8C). 4.2.6 SPARC prevents the interaction between Bcl-2 and caspase 8 Previous studies have reported that apoptosis can be inhibited as a result of an interaction between Bcl-2 and caspase 8 in CHO cells (240) and we wondered if such an interaction also occurred in CRC cells, and in particular, those resistant to chemotherapy. Our findings revealed that in chemotherapy resistant MIP/5FU cells, an interaction between caspase 8 and Bcl-2 could be observed, while it was absent in sensitive MIP/ZEO and MIP/SP cells (Figure 4-9A). This pattern of interaction was replicated in other chemotherapy-resistant cancer cells of various origins (Figure 4-9B), and together with our findings of a SPARC:procaspase 8 interaction, led us to question whether SPARC could influence the interaction between Bcl-2 and caspase 8, to affect chemosensitivity. Interestingly, we noted that exposure of resistant RKO/5FU cells to exogenous rSPARC eliminated this Bcl-2:caspase 8 interaction (Figure 4-9B, *). Similarly, a 93  purified peptide spanning the N-terminus region of SPARC was also able to interfere with the Bcl-2:caspase 8 interaction, based on immunoprecipitation studies (Figure 4-10). However, coincubation with rSPARC in the presence of anti-SPARC antibodies again restored this interaction (Figure 4-9C). 4.2.7 Mutations in the DED domains of procaspase 8 prevent interactions with Bcl-2 and SPARC Results so far have shown that caspase 8 interacts with Bcl-2 and SPARC interferes with the Bcl-2:caspase 8 interaction. We wondered if Bcl-2 interacted within the N-terminus of caspase 8, which is also the site of interaction for SPARC and caspase 8 (196). This led us to perform site-directed mutagenesis studies. To identify the specific site within procaspase 8 that interacts with Bcl-2 or SPARC, specific mutations were introduced in three different regions within the N-terminus of procaspase 8: (1) the DEDI domain (DEDIm), (2) the putative binding region (PBm), and (3) DEDII domain (DEDIIm), which were not tested for activity (Table 2-2). Co-immunoprecipitation studies revealed that transient overexpression of DEDIm and DEDIIm caspase 8 mutants interfered with the ability of Bcl-2 to interact with caspase 8 in MIP/5FU and MiaPaca/CPT cells (Figure 4-11A). In MIP/SP cells, overexpression of DEDIm- and PBm caspase 8 abrogated the interaction between SPARC and caspase 8, while mutations in the DEDII domains of procaspase 8 failed to interfere with its ability to interact with SPARC (Figure 4-11B). These results suggest that the DEDI domain of procaspase 8 is the site of interaction for SPARC and Bcl-2 as it abrogates the Bcl-2:caspase 8 interaction in both the sensitive and resistant cell lines. Interestingly, we noted that over-expression of procaspase 8 mutants abolished the reduction in cell viability and augmentation in apoptosis that is normally conferred by caspase 8 in the presence of SPARC and 5-FU, in resistant cells (Figure 4-12). In the resistant cells, exposure to rSPARC in combination with 5-FU resulted in decreased cell 94  viability following transfection with EV-control: MIP/5FU: 90.14±2.80 (SPARC) vs. 69.89±4.64%  (SPARC+5-FU)  (p<0.005);  MiaPaca/CPT:  108.77±6.85  (SPARC)  vs.  73.98±4.46% (SPARC+5-FU) (p<0.005). Over-expression of wild-type caspase 8 in our resistant cells further decreased cell viability following exposure to rSPARC and 5-FU, in comparison to EV-controls: MIP/5FU: 69.89±4.64 (SPARC+5-FU) vs. 59.21±3.66% (SPARC+5-FU+caspase 8) (p=0.0375); MiaPaca/CPT: 73.98±4.46 (SPARC+5-FU) vs. 46.78±2.01% (SPARC+5FU+caspase 8) (p<0.005). In the resistant cell line RKO/5FU, exposure to exogenous rSPARC in combination with 5-FU resulted in decreased cell viability following transfection with empty vector control: 45.35 ± 4.26 (SPARC) vs. 32.33 ± 2.45% (SPARC+5-FU) (p<0.005). Overexpression of wild-type caspase 8 in this resistant cell line further decreased cell viability following exposure to SPARC and 5-FU, in comparison to empty vector (EV) controls: 32.33±2.45 (SPARC+5-FU) vs. 21.98 ± 1.78% (SPARC+5-FU+caspase 8) (p<0.005). However, over-expression of mutant forms of caspase 8 abolished the reduction in cell viability seen in the presence of rSPARC (Figure 4-12). Similar observations were noted with chemosensitive MIP/SP and MIP/SP-N cells (Figure 4-13). Cell viability was further decreased in MIP/SP and MIP/SP-N after transfection with wild-type caspase 8 and incubation with 5-FU, relative to control empty vector (EV): MIP/SP: 77.22±3.80 (EV) vs. 59.06±1.52% (caspase 8) (p<0.005); MIP/SP-N: 68.79±4.85 (EV) vs. 50.57±3.56 % (caspase 8) (p=0.04). Transfection with any of the caspase 8 mutants eliminated the response of these cells to 5-FU, not only in terms of cell viability, but also decreased caspase 3/7 activity (Figure 4-13A-D). Also, MIP/SP-Nmut cells failed to respond to 5-FU following transfection with either wild-type or mutant caspase 8 (Figure 4-13E-G), thus supporting the N-terminus of SPARC as the site of interaction with procaspase 8. The results reported in this study reveal how an interplay between SPARC, procaspase 8 and Bcl-2 may modulate apoptosis in response to cytotoxic agents. 95  4.3 Discussion Apoptosis occurs as a result of a series of well coordinated events that classically involve either activation of the extrinsic or intrinsic pathways and members of the Bcl-2 protein family. We and others have previously shown higher levels of SPARC, a matricellular protein known to influence cell growth and apoptosis, to be associated with increased apoptosis in ovarian, pancreatic and CRCs (107, 110, 113). Exogenous exposure to rSPARC in vivo promotes greater tumour regression in CRCs that had become refractory to conventional chemotherapies (113). Recently, we identified a potential mechanism that allows SPARC to augment the actions of cytotoxic agents by enhancing the apoptotic cascade (196): SPARC promotes the activation of the extrinsic pathway of apoptosis by interacting with the N-terminus of procaspase 8, with subsequent involvement of the intrinsic pathway to enhance the apoptotic effect.  As an  extension to these earlier observations, the current chapter demonstrates that in cancer cells that have become resistant to chemotherapy, this same N-terminus region of procaspase 8 interacts instead with Bcl-2 to block apoptosis. However, this inhibitory effect on apoptosis can be reversed in the presence of higher levels of SPARC, either from exogenous exposure to SPARC or forced ectopic expression. We further demonstrated that the N-terminus domain of SPARC is the site of interaction with procaspase 8. It must be noted that while the co-immunoprecipitation data supports an interaction between the N-terminus of SPARC and caspase 8, the expression of caspase 8 or SPARC may be due to an artifact found in the experiment. The possibility that the caspase 8:SPARC interaction may only be an artifact causes reservations with the interpretation and conclusions of the results. To validate this data, co-immunoprecipitation experiments can be verified with mass spectrometry to identify proteins that are interacting. However, to fully validate a binary interaction, yeast-two hybrid or transcription-translation experiments in wheat germ cell lysates would need to be performed to eliminate the possibility that the interaction is the result of common intermediary interactions rather than direct interactions. 96  Even more interesting is our finding that not only is the N-terminus responsible for SPARC’s pro-apoptotic activity, but that a recombinant protein of this region alone (54aa in length) can induce apoptosis in the presence of cytotoxic agents in colorectal, pancreatic and breast cancer cells. In fact, tumour xenografts of cells over-expressing only the N-terminus domain of SPARC, and not other domains, experienced the most dramatic decrease in tumour growth in response to chemotherapy. Expression of full length SPARC and its domains after transfection were tested; however, we did not test for the internalization of SPARC peptide or SPARC domain peptides. Verification of the internalization of the peptides would have been highly beneficial for analysis of the results. Also, it is unknown if full length SPARC and the Nterminal domain of SPARC compete for binding to caspase 8 to mediate their chemosensitizing effects. Results supporting such interactions would be interesting to pursue in cells with high SPARC expression. SPARC is expressed in many different cell types (219) and plays an intricate role in tumourigenesis. While some studies report a positive association between high SPARC and more aggressive malignancies (92, 241), several studies, including our own, support the view that it functions in part as a tumour suppressor in neuroblastomas, leukemia, colorectal, ovarian, pancreatic, lung, and breast cancers (106, 107, 110, 113, 123, 220). These divergent actions of SPARC are puzzling, but previous studies have demonstrated that SPARC undergoes proteolysis by a variety of proteases, such as metalloproteinases (MMPs), elastases, cathepsins, and serine proteases (242). These degradation products are also biologically active (238), and differences in tumour-specific protease expression may account for the differences in SPARC’s biological behavior in tumourigenesis. For example, a small peptide containing only 20 amino acids (aa2140) of the N-terminus domain of SPARC had previously been shown to inhibit endothelial cell spreading and bFGF-induced cell migration (66). The native SPARC protein is also known for its ability to inhibit angiogenesis (238), yet release of a proteolytic product containing the Cu2+ 97  binding sequence KGHK (lysine-glycine-histidine-lysine) opposes the activity of the full-length protein by stimulating angiogenesis in vitro and in vivo (67, 76). In our study, we found that xenografts of MIP/SP and MIP/SP-N exhibited fewer CD31+ staining. This is consistent with previous results that the N-terminus domain affects endothelial cell spreading (66) and that the native SPARC protein can inhibit angiogenesis (238). These results may help explain our findings that the effect of the recombinant peptide of SPARC’s N-terminal domain on inhibiting tumour growth was superior to SPARC, and prolonged the tumour doubling time by 46.1% in comparison to SPARC-over-expressing tumours exposed to 5-FU. Our findings indicate that while SPARC’s N-terminus domain inhibits tumour growth, the follistatin-like (FS-) domain alone may have growth-promoting properties, as xenografts over-expressing recombinant proteins of this fragment had significantly shorter doubling times than control xenografts.  These tumours were also unresponsive to 5-FU treatment. These  observations indicate that the efficacy of the native SPARC protein on tumour growth may be altered when its proteolysis results in the release of different fragments with opposing effects. These observations again reflect SPARC’s intricate biology, as potential proteolytic products may have opposing effects on tumour growth, progression and response to therapy. This is particularly interesting as in vitro studies showed that SP-F decreased cell viability. However, the exact mechanism of how the SP-F domain increased in vitro chemosensitivity was not determined. This discrepancy between the in vitro cell viability assays and the in vivo studies is an area that requires further study, in order to allow a better understanding of SPARC’s effect in different types of cancers. Based on the results of the current study, another dimension has been added to the multifaceted biological behavior of SPARC, by demonstrating the pro-apoptotic activity of the N-terminus domain of SPARC (aa18-70). More importantly, we show that the ability of either SPARC or its smaller N-terminus domain to promote apoptosis results from their interference in 98  Bcl-2’s interaction with procaspase 8, which subsequently allows SPARC to interact with procaspase 8, thereby leading to the activation of the extrinsic pathway (Figure 4-14). This assessment of a potential triad involving SPARC, procaspase 8 and Bcl-2 in regulating apoptosis was based on earlier observations that Bcl-2 binds to procaspase 8 to inhibit caspase 8-mediated apoptosis (232); reports of SPARC’s ability to promote apoptosis by decreasing the ratio of Bcl2 and BAX in autosomal dominant polycystic kidney cells (219), together with the recent observations of SPARC’s interaction with caspase 8 (196). Bcl-2 has long been associated with drug resistance since its discovery as a proto-oncogene in non-Hodgkin’s B-cell lymphomas (243), and over the years, it has been extensively studied as a potential target for cancer therapeutics (244). Recently, the differential compartmentalization of Bcl-2 and NRAS has been reported to influence disease states, such as myelodysplastic syndromes and acute myeloid leukemia (245). In their mouse model, the physical interaction of Bcl-2 and mutant NRAS affected the apoptotic machinery. These novel interactions between Bcl-2 and other proteins, highlight the complex mechanisms utilized by cancer cells to evade cell death, leading to disease progression and drug resistance. In this chapter, we demonstrated that Bcl-2 and procaspase 8 interact in cancer cells that have become refractory to therapy, thus providing another mechanism in which Bcl-2 promotes survival and drug resistance.  In addition, our in vitro and in vivo results demonstrated that  SPARC and its smaller fragment containing the N-terminus domain are highly efficacious in modulating and enhancing apoptosis, by its ability to dissociate the interaction between Bcl-2 and procaspase 8. These exciting findings point to the possibility that SPARC (and its smaller N-terminus peptide), and its interference with the Bcl-2:procaspase 8 interaction, can be exploited as a potential therapeutic in disease states, such as cancer, where the promotion of the apoptotic cascade could prove to be clinically beneficial.  99  Results from this thesis so far have shown that SPARC enhances chemosensitivity of CRC cells by activating the extrinsic pathway of apoptosis through the N-terminal domain of SPARC and the N-terminal domain of pro-caspase 8. The ability of cancer cells to metastasize is through its interactions with the extracellular matrix (ECM). SPARC interacts with the various members of the ECM (62), particularly collagen IV through the extracellular domain of SPARC. Collagen IV has also been shown to play a role in inhibiting tumourigenesis (43-45, 47). We therefore assessed the role of collagen IV and SPARC on chemosensitivity in CRC cells in the next chapter.  100  Figure 4-1. Over-expression of the N-terminal domain of SPARC diminished cell viability in cancer cell lines. A) The biological domains of SPARC (SP): the N-terminus (SP-N), follistatin-like (SP-F), and extracellular (SP-C) domains are represented. B) The effect on cell viability following over-expression of the three SPARC domains were assessed by transient transfection. Sensitive and resistant colorectal (MIP, RKO), breast (MCF-7) and pancreatic (MiaPaca) cancer cells with transiently transfected with vectors expressing the full length SPARC (SP), N-terminal domain of SPARC (SP-N), follistatin-like domain of SPARC (SP-F), extracellular domain of SPARC (SP-C), and the vector alone (ZEO). 48 hours after transfection, cells were treated with empty vehicle (■) or chemotherapy (□), for an additional 24 hours. Cell viability was assessed by WST assay. Transfection with the empty vector (ZEO) served as control for all experiments; “Control No txfn”=non-transfected cells. Results represent the mean ± s.e. (n=3-4 independent studies). *, statistical difference compared to control unless otherwise stated, where p < 0.05, Student’s t-test.  101  102  Figure 4-2. Mutations in the N-terminal domain of SPARC inhibit the chemosensitizing effects of SPARC colorectal cancer cell lines. The biological domains of SPARC (SP) and sites of mutations within the N-terminus (SP-N), follistatin-like (SP-F), and extracellular (SP-C) domains are represented. A-D) Cell viability assays were used to determine if mutations in the SPARC-N or SPARC-F domains affected chemosensitivity. Mutations in both the SPARC-N and SPARC-F no longer decreased the survival of chemoresistant cells exposed to chemotherapy. Resistant colorectal (MIP, RKO), breast (MCF-7) and pancreatic (MiaPaca) cancer cells with transiently transfected with vectors expressing the full length SPARC (SP), Nterminal domain of SPARC and its mutants (SP-N, and SP-Nmut1, SP-Nmut2), follistatin-like domain of SPARC and its mutants (SP-F and SP-Fmut1, SP-Fmut2), and the vector alone (ZEO). 48 hours after transfection, cells were treated with empty vehicle (■) or chemotherapy (□), for an additional 24 hours. Cell viability was assessed by WST assay. Transfection with the empty vector (ZEO) served as control for all experiments; “Control No txfn”=non-transfected cells. Results represent the mean ± s.e. (n=3-4 independent studies). *, statistical significance compared to control unless otherwise stated, where p < 0.05, Student’s t-test. Please see Table 22 for information on the specific mutations.  103  104  Figure 4-3 The effect of SPARC mutations on cell viability. A) The biological domains of SPARC (SP) and sites of mutations within the N-terminus (SP-N), follistatin-like (SP-F), and extracellular (SP-C) domains are represented. B) The effect on cell viability following overexpression of the three SPARC domains and its various mutations were assessed. Resistant colorectal cancer cells to 5-FU (MIP/5FU) were transiently transfected with vectors expressing the full length SPARC (SP), N-terminal domain of SPARC and its mutants (SP-N, and SPNmut1, SP-Nmut2, SP-Nmut5, SP-Nmut6, SP-Nmut7, SP-Nmut8), follistatin-like domain of SPARC and its mutants (SP-F and SP-Fmut1, SP-Fmut2, SP-Fmut5, SP-Fmut6, SP-Fmut7, SPFmut8), extracellular domain of SPARC and its mutants (SP-C and SP-Cmut5, SP-Cmut6, SPCmut7) and the vector alone (ZEO). 48 hours after transfection, cells were treated with empty vehicle (■) or chemotherapy (□), for an additional 24 hours. Cell viability was assessed by WST assay. Transfection with the empty vector (ZEO) served as control for all experiments; “Control No txfn”=non-transfected cells. Results represent the mean ± s.e. (n=3-4 independent studies). *, statistical significance compared to untreated control, where p < 0.05, Student’s t-test. Please see Table 2-2 for information on the specific mutations.  105  Figure 4-4. Effect of SPARC-domains on cell proliferation and apoptosis in response to chemotherapy. Stably transfected MIP101 cells with SPARC (MIP/SP); SPARC-N (MIP/SPN); SPARC-F (MIP/SP-F); SPARC-C (MIP/SP-C); empty vector control (MIP/ZEO); or resistant MIP/5FU were used to for A) cellular viability, B) cellular proliferation, and C) TUNEL assays, as well as for D) immunoblots. A) Cells were seeded for 48 hours and then treated with 0 (■) or 1000 (□) µM 5-FU for an additional 48 hours. Cellular viability was assessed by WST assay. B) All cells were synchronized by a double-thymidine block and were grown to 80% confluence after the second release. Cell proliferation was assessed by on Ki-67 staining; C) Cells were grown on coverslips until 80% confluence. Cells were treated with 1000 µM 5-FU for 24 hours and used to assess for apoptotic activity by TUNEL assay. D) Cells were grown until 80% confluence and treated with 1000 µM 5-FU for 0-12 hours. Cell lysates were isolated and changes in the expression of proteins involved in apoptosis were assessed by immunoblots. Results represent the mean±s.e. (n=3-4 independent studies). * p<0.05, Student’s t-test, in comparison to untreated controls, or otherwise noted.  106  107  Figure 4-5. The effect of caspase 8 silencing in colorectal cancer cells over-expressing the SPARC-domains on apoptosis in response to chemotherapy. The relative contribution of caspase 8 in cell lines stably transfected with SPARC (MIP/SP); SPARC-N (MIP/SP-N); SPARC-F (MIP/SP-F); SPARC-C (MIP/SP-C); empty vector control (MIP/ZEO); or resistant MIP/5FU cells were assessed using caspase 8 siRNA. MIP-related cells and endogenous high SPARC-expressing HCT116 cells were transiently transfected with caspase 8 siRNA (or scramble control) for 48 hours, followed by incubation with 0 (■) or 1000 (□) µM 5-FU for an additional 24 hours. Cell lysates were isolated and apoptosis was assessed by caspase 3/7 assay. Results represent the mean±s.e. (n=3-4 independent studies). *, p<0.05, Student’s t-test, in comparison to untreated controls, or otherwise noted.  108  Figure 4-6. Xenografts over-expressing SPARC-N have greater tumour regression in response to 5-FU. A, B) Nude mice with were implanted with MIP/ZEO, MIP/SP, MIP/SP-N, MIP/SP-F, and MIP/SP-C cells subcutaneously. Mice were intraperitoneally treated with saline (-) or 5-FU (+) and tumor growth was assessed. Mice with xenografts of tumours of MIP/SP and MIP/SP-N exhibited the greatest tumour regression in response to 5-FU. C) Doubling time of tumor xenografts were extrapolated from tumor growth measurements. Results represent mean±s.e (n=14 mice/group).  109  Figure 4-7. MIP/SP-N xenografts exhibit greater levels of apoptosis and lower levels of CD31. Xenografts of tumours of MIP/ZEO, MIP/SP, MIP/SP-N, MIP/SP-F, and MIP/SP-C cells were sectioned and used to assess the percentage of A,B) apoptosis or C) CD31 positive blood vessels. A,B) Tumor xenografts sections of (■) saline or (□) 5-FU treated animals were used to determine the number of apoptotic cells, detected by TUNEL staining. C) Tumor xenograft sections of (■) saline or (□) 5-FU treated animals were stained to determine the number of CD31 positive stained blood vessels in the tumour xenografts. * p<0.05, Student’s ttest, ** p < 0.05 in comparison to MIP/SP-F and MIP/SP-C groups. Results representative of n=3 experiments.  110  111  Figure 4-8. The site of SPARC:pro-caspase 8 interaction is at the N-terminus of SPARC. Co-localization of the N-terminus of SPARC with procaspase 8 was determined by co-IP studies either using whole protein extracts or following fractionation of cells into nuclear (N), cytoplasmic (C), and membrane (M) fractions. A) Whole protein extracts of stably transfected MIP101 cells over-expressing the N-terminal domain (MIP/SP-N), follistatin-like domain (MIP/SP-F) and extracellular domain (MIP/SP-C) were used in co-immunoprecipitation studies. 250 µg of protein were immunoprecipitated with antibodies to 6XHis (to detect His-tagged SPARC domains)and caspase 8, and immunoblotted for 6XHis or caspase 8. Only the recombinant N-terminus containing fusion protein from MIP/SP-N cells co-immunoprecipitated with procaspase 8 in a reciprocal fashion. B) Co-localization of SPARC and pro-caspase 8 was determined by co-immunoprecipitation (co-IP) studies following fractionation of MIP/SP-N cells into nuclear (N), cytoplasmic (C) and membrane fractions (M) with our without exposure to 1000 µM of 5-FU for 4 hours. 250 µg of protein from each cellular fraction were immunoprecipitated with antibodies to 6XHis (to detect His-tagged SPARC domains), caspase 8 or IgG (non-specific control), and immunoblotted for 6XHis or caspase 8. This interaction between the N-terminus of SPARC with procaspase 8 occurs at the cell membrane and not the cytoplasmic or nuclear fractions, and is not affected by exposure to 1000 µM 5-FU. C) Whole protein extracts of stably transfected MIP101 cells over-expressing the N-terminal domain and its mutants (MIP/SP-N and MIP/SP-Nmut1, MIP/SP-Nmut2) and follistatin-like domain and its mutants (MIP/SP-F and MIP/SP-Fmut1, MIP/SP-Fmut2) were used in co-immunoprecipitation studies. 250 µg of protein were immunoprecipitated with antibodies to 6XHis (to detect Histagged SPARC domains) and caspase 8, and immunoblotted for 6XHis or caspase 8. Mutations in the N-terminus domain of SPARC prevents its interaction with pro-caspase 8. Neither the wild-type nor the mutated FS domain interacted with caspase 8. Input = protein loaded per immunoprecipitation. All results representative of n = 3-4 independent experiments. Please see Table 2-2 for information on the specific mutations.  112  113  Figure 4-9. SPARC and Bcl-2 both interact with pro-caspase 8 with opposing effects. Colocalization of the Bcl-2 with procaspase 8 was determined by co-IP studies either using whole protein extracts or following fractionation of cells into nuclear (N), cytoplasmic (C), and membrane (M) fractions. A) Whole protein extracts of MIP/ZEO, MIP/SP, and MIP/5FU cells were used in co-immunoprecipitation studies. 250 µg of protein were immunoprecipitated with antibodies to Bcl-2 and caspase 8, and immunoblotted for Bcl-2 or caspase 8. Only MIP/5FU cells co-immunoprecipitated with Bcl-2 and caspase 8 in a reciprocal fashion. B) Whole protein extracts of MIP/SP-N, MIP/SP-F, MIP/SP-C, RKO/5FU, RKO/CPT, RKO/CIS, MiaPaca/CPT, and MCF7/CIS cells were used in co-immunoprecipitation studies. 250 µg of protein were immunoprecipitated with antibodies to Bcl-2 and caspase 8, and immunoblotted for Bcl-2 or caspase 8. All chemotherapy resistant cells co-immunoprecipitated with Bcl-2 and caspase 8 in a reciprocal. The Bcl-2:caspase 8 interaction can be eliminated following exposure to rSPARC (100 ng/mL) (*). C) MIP/5FU and MiaPaca/CPT cells were fractionated into nuclear (N), cytoplasmic (C), and membrane (M) fractions. 250 µg of protein from each fraction were immunoprecipitated with antibodies to Bcl-2 and caspase 8, and immunoblotted for Bcl-2 or caspase 8. In both cell lines, co-immunoprecipitation of Bcl-2 and caspase 8 were observed in the membrane. The Bcl-2:pro-caspase 8 interaction is abrogated after exposure to 100 ng/mL rSPARC in resistant MIP/5FU and MiaPaca/CPT cells. Input = protein loaded per immunoprecipitation. All results representative of n = 3-4 independent experiments.  114  115  Figure 4-10. Immunoprecipitation of MIP/5FU cells with SPARC peptides. MIP/5FU cells were seeded and incubated with a purified peptide spanning the N-terminus region of SPARC (SP-N (1) or (2)), a scramble peptide (peptides at 100 ng/mL) or no peptide for 48 hours. Cells were then isolated in 1X CHAPS lysis buffer and used for immunoprecipitation with Bcl-2 or Caspase 8. Eluates were then run on a 12.5% SDS-PAGE gel, transferred to a PVDF membrane and used for immunoblotting against caspase 8 or Bcl-2, respectively. Detection was determined by chemiluminescence. Input = protein loaded per immunoprecipitation. Results representative of n = 2-3 experiments.  116  Figure 4-11. The DEDI domain of caspase 8 is critical for its interaction with Bcl-2.A,B) Co-localization of the Bcl-2 or SPARC with mutant forms of caspase 8 (death effector domain I (DEDIm), putative binding (PBm), and death effector domain II (DEDIIm) was determined by co-IP studies using whole protein extracts A) Whole protein extracts of MIP/5FU and MiaPaca/CPT cells transiently transfected with vectors containing mutations in the specific domains were used in co-immunoprecipitation studies. 250 µg of protein were immunoprecipitated with antibodies to Bcl-2 and caspase 8, and immunoblotted for Bcl-2 or caspase 8. Mutations in both the DEDIm and DEDIIm domains abrogated the Bcl-2:caspase 8 interaction in resistant cell lines. B) Whole protein extracts of MIP/SP cells transiently transfected with vectors containing mutations in the specific domains were used in coimmunoprecipitation studies. 250 µg of protein were immunoprecipitated with antibodies to SPARC and caspase 8, and immunoblotted for SPARC or caspase 8. Mutations in both the DEDIm and PBm domains abrogated the SPARC:caspase 8 interaction in MIP/SP cells. C) Basal levels of caspase 8 gene expression between various cancer cell lines. Input = protein loaded per immunoprecipitation. All results representative of n = 3-4 independent experiments. Please see Table 2-2 for information on the specific mutations.  117  Figure 4-12. The DEDI domain of caspase 8 is critical for its effect on cell viability. A-D) MIP/5FU, RKO/5FU, and MiaPaca/CPT Cells were transiently transfected with wild-type caspase 8, vectors containing mutations in specific domains (DEDIm, PBm, and DEDIIm), or control empty-vector (EV) for 48 hours. Cells were exposed to 0 (-) to 100 (+) ng/mL rSPARC for 48 hours, and treated with 0 (■) or 500 (□) µM 5-FU for an additional 24 hours. Cells were used for A-C) cell viability assays and D) caspase 3/7 activity. Results represent mean±s.e. (n=3 independent studies). * p < 0.05, Student’s t-test. Please see Table 2-2 for information on the specific mutations.  118  Figure 4-13. Interaction between the N-terminus of both pro-caspase 8 and SPARC are required to reduce cell viability and enhance caspase 3/7 activity. A-D) MIP/SP, MIP/SP-N, MIP/SP-Nmut1, and MIP/SP-Nmut2 cells were transiently transfected with wild-type caspase 8, vectors containing mutations in specific domains (DEDIm, PBm, and DEDIIm), or control empty-vector (EV) for 48 hours and treated with 0 (■) or 500 (□) µM 5-FU for an additional 24 hours. Cells were used for A,C,E,F) cell viability assessed by WST assays and B,D,G) apoptosis by caspase 3/7 assays. A, C) Cell viability was further decreased in MIP/SP and MIP/SP-N after transfection with wild-type caspase 8 and incubation with 5-FU relative to control empty vector (EV). (B, D). In cells over-expressing mutant SP-N (MIP/SP-Nmut1), transfection with caspase 8 was incapable of decreasing cell viability (E-F) or increasing caspase 3/7 activity (G). Results represent mean±s.e. (n=3 independent studies). *, statistical difference where p < 0.05, Student’s t-test. Please see Table 2-2 for information on the specific mutations.  119  120  Figure 4-14: A model: SPARC-mediated apoptosis. A schematic of SPARC and Bcl-2 interacting with caspase 8 to influence apoptosis. In this study, we demonstrate that in an environment low in SPARC, Bcl-2 interacts with pro-caspase 8 to inhibit apoptosis. However, in the presence of SPARC, the N-terminus of SPARC can interact with pro-caspase 8 to inhibit its interaction with Bcl-2, leading to apoptosis.  121  5. SPARC interacts with collagen IV to re-sensitize resistant colorectal cancer cells 5.1 Introduction CRC remains the second leading cause of cancer death in Canada (5). A large obstacle in decreasing the high mortality rate of CRC is the development of chemotherapy resistance, which decreases the efficacy of conventional cytotoxics. The extracellular matrix (ECM) has important implications in the failure of conventional cancer therapies and subsequent tumour progression. The ECM has been shown to modulate the development of tumours through various biological processes such as cellular proliferation, differentiation, and viability. For example, in osteosarcomas, the ECM has been shown to increase cell proliferation and migration (246, 247). In addition, laminin, a component of the ECM, increased levels of cisplatin induced apoptosis in testicular germ cell lines (248). These examples also demonstrate that each component of the ECM can affect various biological functions. The ECM is composed of many different proteins, including fibronectin, laminin, vitronectin, and collagen IV. Collagen IV (CIV) is a major component of the basement membrane (39) and has been shown to play a role in angiogenesis (40), cell adhesion and migration (41, 42). CIV has also been shown to reduce carcinogenesis. A fragment of CIV, the NC1 peptide, promoted adhesion and inhibited proliferation and invasion, resulting in the inhibition of growth of various human cancer cell lines (43-47). This effect of CIV has also been demonstrated in vivo. Injecting CIV in xenograft models of prostate and renal cell carcinoma decreased tumour growth by decreasing angiogenesis, proliferation, and invasion, and increasing apoptosis (48). The mechanisms by which CIV and the ECM affect these biological functions in cancer are largely unknown. However, the cell signaling properties of the ECM are highly dependent on its interactions. Specifically, cellular adhesion to the ECM has been shown to modulate the 122  expression of several cell-cycle regulators, including cyclin-dependent kinases (CDK2 and CDC2), cyclins (A, B, D and E), cyclin-dependent kinase inhibitors (p21 and p27) and Rb (249, 250). The interaction between the cell and ECM is mediated by integrins, which are cell surface receptors. Integrins function as heterodimers composed of α and β subunits that recognize and bind to specific components of the ECM, transducing information that affects cellular behaviour and gene expression (251, 252). One of the many biological functions that integrins mediate is apoptosis; however, experiments investigating the effect of integrin signaling on apoptosis have shown contradictory results. In the majority of cases, the ECM protected cells from chemotherapeutic drug-induced apoptosis (253-258); conversely, there are some cases where the ECM promoted apoptosis (259261). The mechanistic differences between ECM-mediated cell survival and cell death pathways are not well understood. Various ECM interactions and cell signaling pathways have been suggested to account for this discrepancy. For instance, integrin signaling has been shown to modulate the Ras-MAP kinase (Ras-MAPK), Rho-effector, cyclic AMP-dependent protein kinase (PKA), and PI3K/Akt pathways (262, 263). Another protein that binds to various ECM components is SPARC. SPARC is a secreted, phosphorylated and calcium-binding glycoprotein (62). It functions in various biological processes such as cellular adhesion, proliferation, migration, and tissue remodeling. SPARC is expressed during development and in processes that require ECM turnover such as wound healing and tumour progression (53, 86, 100, 197, 264). The interaction between SPARC and the ECM is thought to contribute to the remodeling of the ECM. Tumours grown in SPARC-/- mice display altered production and organization of ECM components with changes in proliferation and apoptosis (127). Previously, we showed that SPARC re-sensitizes chemotherapy resistant CRC cells by activating apoptosis (113). This is supported by other studies where SPARC  123  expression increased apoptosis in ovarian cancer cells (107, 125), autosomal dominant polycystic kidneys (219), neuroblastomas (222) and pancreatic cancers (110). Both SPARC and CIV have been shown to affect apoptosis. Further, CIV has been shown to interact with the extracellular (SP-C) domain of SPARC (102, 265) and SPARC-/mice with lobular tumours have defective CIV deposition in the stroma (266). Based on this preliminary evidence for interaction between CIV-SPARC, we hypothesize that the addition of CIV would increase the sensitizing effects compared to SPARC alone. Thus, we decided to investigate the effect of CIV on CRC growth and apoptosis in conjunction with SPARC and chemotherapy treatment. 5.2 Results 5.2.1 Collagen IV increases sensitivity of colorectal cancer cells to chemotherapy by decreasing cell viability and cell proliferation The tumour microenvironment and its constituents have been shown to influence tumour growth (246, 267-269). Specifically, tumstatin and canstatin (both fragments of CIV) decreased in vivo tumour growth by decreasing proliferation and angiogenesis and increasing apoptosis (270-274). SPARC, a matricellular protein, has also been shown to influence the progression of tumourigenesis (112). We have previously shown that SPARC induces apoptosis in resistant CRC cells (196) and others have shown an interaction between SPARC and CIV (265). The observations suggest that the SPARC-CIV relationship may influence tumourigenesis through various biological mechanisms. We therefore investigated the effect of CIV on cell viability and apoptosis in conjunction with SPARC in CRC. We initially assessed the effects of CIV on cell viability of MIP101 CRC cells by plating these cells on CIV or control matrices and treating with 5-FU chemotherapy. CIV matrices increased the chemosensitivity of MIP/ZEO and MIP/SP cells in response to 500-1000 µM 5-FU 124  treatment relative to control matrices (MIP/ZEO: 500 µM 5-FU: 78.93 ± 3.43 (control) vs. 59.41 ± 5.20% (CIV), p <0.05, 1000 µM 5-FU: 73.33 ± 3.15 (control) vs. 55.81 ± 8.45% (CIV), p<0.05; MIP/SP: 500 µM 5-FU: 66.52 ± 7.08 (control) vs. 48.44 ± 9.78% (CIV), p <0.05, 1000 µM 5-FU: 58.30 ± 8.24 (control) vs. 46.87 ± 9.33% (CIV), p<0.05) (Figure 5-1A,B). Interestingly, resistant MIP/5FU cells, decreased cell viability in response to 5-FU treatment on CIV plates only (Figure 5-1C). These results indicate that CIV increases the sensitivity of CRC cells to chemotherapy. Further, for all cells lines, doubling time was increased in the presence of CIV (MIP/ZEO: 35.61 ± 2.35 (control) vs. 60.21 ± 0.30 (CIV) hours, p <0.05; MIP/SP: 37.07 ± 1.10 (control) vs. 78.87 ± 0.07 (CIV) hours, p<0.05; MIP/5FU: 61.02 ± 0.16 (control) vs. 130.03 ± 4.01 (CIV) hours, p<0.05) (Figure 5-2D), which is indicative of decreased proliferation (Figure 5-2A-C). These results show that CIV decreases cellular number. 5.2.2 Collagen IV increases chemosensitivity through the mitochondrial pathway of apoptosis. Having determined that CIV increases the chemosensitivity of CRC cells, we assessed the effect of CIV and 5-FU on apoptosis in an attempt to recapitulate the previously demonstrated phenotype of increased apoptosis in response to CIV (270-274). When compared to CRC cells grown on control matrices, those grown on CIV matrices demonstrated increased caspase 3/7 levels after treatment with 5-FU for 4 hours (MIP/ZEO: 4 hours post-5-FU treatment: 13000 ± 79 (control) vs. 16372 ± 316 (CIV) RLU, p<0.05; MIP/SP: 4 hours post-5FU treatment: 18814 ± 209 (control) vs. 36178 ± 169 (CIV) RLU, p<0.05; MIP/5FU: 4 hours post-5-FU treatment: 854 ± 27 (control) vs. 14572 ± 158 (CIV) RLU, p<0.05) (Figure 5-3A). This increase in apoptosis was also seen after CPT treatment, although not to the same degree, assayed using the TUNEL assay (CPT treated: MIP/ZEO: 8.98 ± 2.71 (control) vs. 17.12 ± 2.65 125  (CIV)%, p<0.05; MIP/SP: 15.40 ± 3.81 (control) vs. 32.75 ± 5.13 (CIV)%, p<0.05; MIP/CPT: 2.74 ± 1.47 (control) vs. 21.28 ± 6.36 (CIV)%, p<0.05) (Figure 5-3B). Interestingly, while SPARC over-expressing MIP/SP cells remained the most chemosensitive overall, previously resistant MIP/5FU cells showed apoptotic activity in response to chemotherapy when on CIV matrices (i.e. these cells now had restored chemosensitivity) (Figure 5-3A, B). The specific apoptotic signaling cascade by which CIV induces apoptosis has not yet been fully elucidated. Both the intrinsic and the extrinsic pathways have been shown to be evoked by chemotherapy (196). Having confirmed that CIV does, in fact, increase apoptosis in CRC cells in combination with chemotherapy, we proceeded to investigate the specific signaling cascade involved. To do this, we assessed the levels of several caspases. Caspase 8 protein expression did not change when CRC cells were plated on CIV or control matrices, indicating that the extrinsic pathway of apoptosis is not activated. However, when grown on CIV, all cell lines exhibited higher expression of t-Bid, cleaved caspase 9 and cleaved caspase 3, which are players in the intrinsic pathway (Figure 5-4). Thus, our results indicate that CIV enhanced chemosensitivity of CRC cells occurs through the intrinsic and not the extrinsic pathway of apoptosis. 5.2.3 Collagen IV further enhances chemosensitivity of SPARC biological fragments So far, it appears that there is a co-operative effect with CIV and SPARC to affect chemosensitivity of CRC cells (Figure 5-3 and 5-4). We next wanted to determine if this cooperative effect can be localized to a specific biological domain of SPARC. SPARC contains three biological domains, each with unique modular functions (61). MIP101 cells stably overexpressing the biological domains of SPARC (N-terminus: MIP/SP-N; follistatin-like: MIP/SPF; and extracellular: MIP/SP-C) were grown on control or CIV matrices and assessed for cellular viability, cellular proliferation, apoptosis, and changes in gene expression levels. 126  We first assessed changes in cellular viability in response to CIV and chemotherapy. MIP/SP-F cells exhibited an ~18% decrease in cell viability on CIV plates compared to control plates, similar to MIP/SP cells at 500 µM 5-FU exposure. Cells over-expressing the other two domains of SPARC, MIP/SP-N and MIP/SP-C cells, only exhibited a ~11% decrease in cell viability on CIV plates compared to control plates at this lower concentration. In contrast, at 1200 µM 5-FU treatment, MIP/SP-N, MIP/SP-F and MIP/SP-C cells all showed lower cellular viability in response to 5-FU treatment on CIV than on control plates (MIP/SP-N: 1200 µM 5FU: 64.81 ± 7.83 (control) vs. 57.85 ± 13.89% (CIV), p <0.05; MIP/SP-F: 1200 µM 5-FU: 65.24 ± 7.95 (control) vs. 54.50 ± 14.03% (CIV), p <0.05; MIP/SP-C: 1200 µM 5-FU: 73.78 ± 10.20 (control) vs. 68.99 ± 11.18% (CIV), p <0.05); however, this was not as efficient as full length SPARC which exhibited a greater decrease in cell viability. Most interestingly, MIP/SP-C cells did not decrease cell viability in response to 500-1000 µM 5-FU on control plates, but were sensitive to the same concentrations of 5-FU on CIV plates, shown by a decrease in cell viability (Figure 5-5). This decrease in cell viability due to CIV and 5-FU treatment, in MIP101 cells overexpressing the SPARC domains, correlated with increased doubling time (MIP/SP-N: 35.47 ± 0.56 (control) vs. 67.61 ± 0.41 (CIV) hours, p<0.05; MIP/SP-F: 76.21 ± 0.30 (control) vs. 168.94 ± 4.64 (CIV) hours, p<0.05; MIP/SP-C: 26.03 ± 2.58 (control) vs. 94.91 ± 1.61 (CIV) hours, p<0.05) (Figure 5-6). MIP/SP-F cells exhibited a 2.2 fold increase in doubling time when grown on CIV matrices, similar to full length SPARC (2.1 fold). Where MIP/SP-N cells only had a 1.9 fold increase in doubling time on CIV plates and MIP/SP-C cells exhibited a greater fold of 3.6 compared to control plates. The cells over-expressing the SPARC domains also exhibited greater caspase 3/7 levels after 5-FU treatment when grown on CIV plates compared to control (4 hours post-5-FU treatment: MIP/SP-N: 3087.34 ± 75.98 (control) vs. 4183.98 ± 59.52 (CIV) RLU, p<0.05; 127  MIP/SP-F: 3204.04 ± 203.32 (control) vs. 5486.42 ± 210.78 (CIV) RLU, p<0.05; MIP/SP-C: 1246.28 ± 169.25 (control) vs. 3452.33 ± 383.95 (CIV) RLU, p<0.05) (Figure 5-7A). MIP/SP cells exhibited a 1.9 fold increase on CIV plates compared to control plates. A similar increase was found with MIP/SP-F cells, whereas MIP/SP-N cells only exhibited a 1.4 fold increase and MIP/SP-C cells exhibited a 2.7 fold increase. These results show that the changes in cellular viability are due to decreased cellular proliferation and increased caspase 3/7 activity in response to CIV and 5-FU. MIP/SP-F cells exhibit similar changes as full length SPARC when grown on CIV matrices, while MIP/SP-C cells exhibited even greater differences on CIV plates and MIP/SP-N cells showed smaller changes. Next, to determine if the changes in cellular viability, cellular proliferation and caspase 3/7 activity were mediated through the same cell signaling pathways as shown above, we examined the gene expression of components of the apoptotic pathway. These experiments revealed that cells expressing a single domain of the protein showed greater expression of components of the apoptotic pathway, namely cleaved caspase 9, Bid, and caspase 3, when grown on CIV as compared to control plates (Figure 5-7B). Interestingly, both MIP/SP-F and MIP/SP-C cells show greater expression of cleaved caspases on CIV plates compared to MIP/SP-N cells. These results show that MIP/SP-F exhibit similar changes in cell viability, cell proliferation, and caspase 3/7 activity, when grown on CIV plates, as full-length MIP/SP cells. However, MIP/SP-N cells exhibited smaller changes and MIP/SP-C cells showed greater changes, in cell proliferation and caspase 3/7 activity, compared to MIP/SP and MIP/SP-F cells.  128  5.2.4 SPARC enhances the chemosensitizing effects of collagen IV through the extracellular domain of SPARC We have shown that CIV varies in its effect on the chemosensitivity of CRC cells expressing the various fragments of SPARC. Others have shown that SPARC interacts with CIV through the SP-C domain of SPARC (102, 265), and interestingly, results have shown that MIP/SP-C exhibit greater changes when grown on CIV plates in cell proliferation and caspase 3/7 activity compared to MIP/SP, MIP/SP-N and MIP/SP-F cells. We thus wanted to determine whether a specific domain of SPARC co-operatively acts with CIV to affect chemosensitivity. To determine which domain was responsible for the cooperative effect, we initially did a screening process with random mutations. Site-directed mutagenesis was performed on the biological domains of SPARC (Table 2-2) and MIP101 cells stably transfected with these mutated plasmids were plated on CIV (or control) and assessed for cellular viability after 5-FU treatment. Cells carrying mutations in the SP-N and SP-F domains grown on control plates did not decrease cell viability in response to 5-FU (MIP/SPNmut1: 103.53 ± 5.47 (0 µM 5-FU) vs. 96.15 ± 5.18% (1000 µM 5-FU), p = 0.3488; MIP/SPFmut1: 107.18 ± 5.48 (0 µM 5-FU) vs. 92.76 ± 4.85% (1000 µM 5-FU), p = 0.0773). However, these cells did show decreased cell viability with 5-FU treatment when grown on CIV (MIP/SPNmut1: CIV: 99.43 ± 6.90 (0 µM 5-FU) vs. 83.77 ± 3.83% (1000 µM 5-FU), p <0.05; MIP/SPFmut1: 102.87 ± 5.73 (0 µM 5-FU) vs. 77.28 ± 3.82% (1000 µM 5-FU), p<0.05). This decrease in cell viability of MIP/SP-N and its mutants as well as MIP/SP-F and its mutants on CIV plates was also exhibited with full length SPARC cells. However, MIP/SP-N and it mutants did not decrease to the same extent as MIP/SP cells, whereas MIP/SP-F cells showed a similar decrease in cell viability as full length SPARC. Interestingly, cells carrying mutations in the SP-C domain were slightly protected from 5-FU  129  treatment, as shown by increased cellular viability after 5-FU treatment on CIV matrices (MIP/SP-Cmut1, control: 104.80 ± 4.81 (0 µM 5-FU) vs. 82.77 ± 4.35% (1000 µM 5-FU), p <0.05; CIV: 98.78 ± 6.50 (0 µM 5-FU) vs. 93.97 ± 5.09% (1000 µM 5-FU), p=0.5733) (Figure 5-8). These changes in cellular viability from the mutations in SPARC’s biological domains correlated with changes in caspase 3/7 activity. Cells carrying mutations in the SP-N or SP-F domain, which did not respond to 5-FU on control plates, showed greater caspase 3/7 activity on CIV plates (MIP/SP-Nmut1, control: 1379 ± 20 (0 µM 5-FU) vs. 1389 ± 13 (1000 µM 5-FU) RLU, p=0.6902, CIV: 1363 ± 9 (0 µM 5-FU) vs. 2347 ± 64 (1000 µM 5-FU) RLU, p<0.05; MIP/SP-Fmut1, control: 1407 ± 4 (0 µM 5-FU) vs. 1395 ± 20 (1000 µM 5-FU) RLU, p=0.5811, CIV: 1415 ± 10 (0 µM 5-FU) vs.2397 ± 30 (1000 µM 5-FU) RLU, p<0.05). This increase was comparable to the 1.2x fold change exhibited by MIP/SP cells. However, wild-type MIP/SP-N and MIP/SP-F cells were still responsive on both control and CIV plates indicating that the SP-N or SP-F domains do not contribute to the CIV-induced chemosensitivity (Figure 5-9A-C). We next examined the SP-C domain to determine if this domain was involved in the effects of SPARC and CIV. Cells carrying mutations in the SP-C domain, which were responsive on control plates, showed reduced caspase 3/7 activity on CIV (control: 1399 ± 2 (0 µM 5-FU) vs. 2189 ± 173 (1000 µM 5-FU) RLU, p<0.05, CIV: 1407 ± 2 (0 µM 5-FU) vs. 1753 ± 74 (1000 µM 5-FU) RLU, p<0.05) (Figure 5-9D). These changes in caspase 3/7 levels indicate that CIV or SPARC alone enhance the chemosensitivity of CRC cells, and that the SP-C domain and CIV enhances CIV's chemosensitizing effects in a co-operative manner. 5.3 Discussion SPARC, a matricellular protein, is expressed in many different cell types and plays a role in many biological functions and tumourigenesis. SPARC has been shown to interact with CIV 130  (62) and CIV has also been shown to inhibit tumour growth (48). Previously, we demonstrated that SPARC expression enhances CRC tumour regression (113). These previous results and others, which showed that CIV affects tumour growth and interacts with SPARC, led us to assess the effects of CIV on the response of MIP101 resistant CRC cells to chemotherapy in conjunction with the apoptotic pathway. Others have shown that CIV inhibits the growth of tumour cells. Specifically, fragments of CIV inhibited tumour growth in keratinocytes, mammary, colon, and Lewis lung carcinoma as well as endothelial cells by suppressing angiogenesis, decreasing proliferation, enhancing apoptosis, and inhibiting invasion of tumour cells (270, 275-279). In this chapter, we showed that CIV increases the chemosensitivity of CRC cells by decreasing cell viability and proliferation and increasing apoptosis. CRC cells grown on CIV-coated plates exhibit greater caspase 9 cleavage, which is involved in the intrinsic pathway of apoptosis. This increased cleavage is even more significant following exposure to 5-FU. The elevated caspase 9 cleavage also translates to increased levels of caspase 3/7 activity. This CIV mediated activation of the intrinsic pathway of apoptosis is supported by previous work, which showed that canstatin, a non-collagenous domain of CIV, triggers the intrinsic apoptotic pathway in endothelial and tumour cells (271). We also showed that cells grew slower on CIV plates, shown by an increase in doubling time. The potential significance of the longer doubling time in sensitivity of the cells to 5-FU is difficult to explain based on the known actions of the drug and thus would require further study. For example, the cells in this study were not synchronized, and this may be one approach that could be used to fully test this effect. The findings from this chapter were intriguing as the ECM has typically been thought to confer chemotherapy resistance to apoptosis (255, 280) and promote cell survival (281-284). However, our studies have shown that CIV inhibits tumour growth. Our studies were performed on plates coated with CIV, yet we observed an enhanced response to chemotherapy. This could 131  be due to digestion of CIV, producing fragments to promote apoptosis. Product manufacturers have found that CIV produces smears rather than one distinct band during electrophoresis, which might indicate that many fragments are present when commercially acquired CIV is used for experiments. Identification of the specific fragments would help determine why we observed enhanced chemosensitivity and further validate the claims made in this chapter. We also investigated if SPARC contributes to the increased sensitization from CIV. Cleavage of proteins involved in the apoptotic pathway (caspase 9, Bid, caspase 3) is increased with greater SPARC expression and further increased when grown on CIV. These results showed that SPARC and CIV both enhance apoptosis. It was also observed that CIV enhanced levels of caspase 3/7 activity in all cells relative to those grown on control plates; however, baseline levels of caspase 3/7 were higher in MIP/SP cells compared to MIP/ZEO cells. This discrepancy could be due to the differing number of cells to obtain the same confluency at the end of the experiment. Also, the induction change compared to control cells were higher in MIP/SP cells (~1.9 times) compared to MIP/ZEO cells (~1.3 times). Thus, MIP/SP cells on CIV indeed were more sensitive to chemotherapy treatment than on control plates in a co-operative manner. We have shown that this co-operative effect occurs through the SP-C domain, which others have shown to interact with CIV (62). We observed that cells over-expressing the SP-F domain decrease cell proliferation and increase caspase 3/7 activity most similarly to full length SPARC. Cells over-expressing the SP-N domain are not as chemosensitive on CIV plates as full length SPARC, shown by a faster proliferation time and lower caspase 3/7 levels. However, cells over-expressing the SP-C domain are more greatly affected by the combination of CIV and chemotherapy, relative to full length SPARC, shown by an even greater decrease in cell proliferation and a greater increase in caspase 3/7 activity. We also showed that MIP/SP-C cells decrease cell viability at lower concentrations (1000 µM) compared to MIP/SP-N and MIP/SP-F  132  cells. This could be because MIP/SP-N and MIP/SP-F cells are more chemosensitive and reach a plateau earlier than the more resistant MIP/SP-C cells. Although the results have shown that the SP-F domain exhibits the most similar changes in response to CIV and 5-FU, as full length SPARC, mutations in the SP-F domain did not inhibit its chemosensitizing effects. However, mutations in the SP-C domain of SPARC abrogate the chemosensitizing effects of CIV, shown by increased cellular viability and decreased caspase 3/7 activity. This indicates that the SP-C domain of SPARC influences the chemosensitizing effects of CIV. These results are particularly interesting as we also observed in chapter 3 that SPARC's chemosensitizing effects occur only when SPARC is intracellular. However, in chapter 4, we have also observed that the SP-N domain of SPARC is responsible for its chemosensitizing abilities. In this chapter, we show another role for the SP-C domain. This discrepancy could be due to the mechanism that CIV utilizes to increase chemosensitivity. Does CIV interact with an integrin to trigger the signaling cascade? Studies have shown that CIV interacts with the β1 integrin to affect apoptosis (285, 286). If this is the mechanism of action in MIP cells, overexpressing the SP-C domain could facilitate the interaction or alternatively the SP-C domain could be responsible for transcriptional control of the specific integrin. Another possibility is that it is the full length CIV that exerts its chemosensitizing effect and not CIV fragments. If this is the case, perhaps the SP-C domain of SPARC is responsible for the maturation of CIV, as seen in kidney cells (222). To determine the discrepancy between the SP-N and the SP-C domain in increasing chemosensitivity, the mechanisms of how CIV enhances chemosensitivity must be determined. Then, the assessment of how the SP-C domain co-operatively acts with CIV can be found. It would also be interesting to determine why SP-F is not affected after mutagenesis. In this chapter we showed that CIV enhances the chemosensitivity of CRC cells and the SP-C domain of SPARC influences CIV's chemosensitizing effects. Our current observations supports a CIV-dependent event enhancing the chemosensitivity of CRC cells and SPARC and 133  CIV jointly enhances chemosensitivity by further up-regulating apoptosis (Figure 5-10). The use of CIV and SPARC to inhibit tumour growth seems to be multifactorial. However, further experiments are necessary to determine if the in vitro effects demonstrated in this chapter will be recapitulated in vivo. Additionally, we only investigated the effects of collagen IV on colorectal cancer cells. The extracellular matrix is comprised of many other types of collagen and SPARC has been shown to interact with other forms (62); thus, determining if other collagens have similar effects would be very interesting. The results presented in this chapter are exciting because they provide initial insight into a potential mechanism by which CIV and SPARC work to mediate their chemosensitizing effect in tumors that are refractory to therapy. These results, if recapitulated in vivo, may support the use of both CIV and SPARC in the treatment of chemotherapy resistance in patients with advanced therapy-refractory CRC.  134  Figure 5-1. Collagen IV decreases cellular viability of MIP101 colorectal cancer cells. A-C) MIP/ZEO, MIP/SP, and MIP/5FU cells were plated on control or CIV-coated (50 µg/cm2) plates. 24 hours later, cells were treated with 0 (■), 500 ( ), 1000 (□), or 1200 ( ) µM 5-FU for an additional 24 hours and assessed for cell viability through WST assays. The percentage of viable cells was calculated based on the A450 reading of non-treated control cells. *, statistical difference compared to untreated counterpart unless otherwise specified, where p<0.05, Student’s t-test. All of the results represent mean ± s.e. (n = 3-4 independent studies).  135  Figure 5-2. Collagen IV increases the doubling time of MIP101 colorectal cancer cells. AC) MIP/ZEO, MIP/SP, and MIP/5FU cells were plated on control or CIV-coated (50 µg/cm2) plates. Total cellular number was assessed 3-4 days after seeding on control ( ) or CIV-coated (■) plates. *, statistical difference compared to CIV counterpart, where p<0.05, Student’s t-test. D) Doubling time of CRC cells on control (■) or CIV-coated (□) plates were calculated from changes in cellular numbers. *, Statistical difference, where p < 0.05, Student’s t-test. All of the results represent mean ± s.e. (n = 3-4 independent studies).  136  Figure 5-3. The effect of collagen IV on apoptosis. A) MIP/ZEO, MIP/SP, and MIP/5FU cells were plated on control or CIV-coated plates for 24 hours. Cells were then treated with 1000 µM 5-FU for 0 (■), 2 ( ), or 4 (□) hours and assayed for caspase 3/7 activity. All of the results represent mean ± s.e *, statistical difference to cells at t = 0 hours unless otherwise specified, where p<0.05, Student’s t-test. B) Cells were additionally grown on control or CIV-coated coverslips for 48 hours and treated with 0 (□) or 100(■) µM CPT-11 for an additional 24 hours and used for TUNEL assay. The percentage of apoptotic cells was determined by counting the number of TUNEL-positive apoptotic nuclei relative to the number of DAPI-positive nuclei from four different fields, with a minimum of 100 cells/field by microscopy. All of the results represent mean ± s.e *, statistical difference to untreated counterpart unless otherwise specified, where p<0.05, Student’s t-test. All of the results representative of n = 3-4 experiments.  137  Figure 5-4. The effect of collagen IV on the cell signaling pathway of apoptosis. MIP/ZEO, MIP/SP, and MIP/5FU cells were plated on control or CIV-coated plates. Changes in the expression of proteins involved in apoptosis following incubation with 1000 µM 5-FU after various time intervals. Total proteins were isolated from cells harvested at 0-4 hours following treatment with 1000 µM 5-FU and probed for various proteins (activated caspases and Bid are represented by their cleaved fragments). All of the results representative of n = 3-4 experiments.  138  Figure 5-5. Collagen IV enhances chemosensitivity in MIP101 cells over-expressing the various biological domains of SPARC. A) MIP/SP-N, MIP/SP-F, and MIP/SP-C CRC cells were plated on control or CIV-coated (50 µg/cm2) plates. 24 hours later, cells were treated with 0 (■), 500 ( ), 1000 (□), or 1200 ( ) µM 5-FU for an additional 24 hours and assessed for cell viability through WST assays. The percentages of viable cells were calculated based on the A450 reading of non-treated control cells. All of the results represent mean ± s.e. *, statistical difference to untreated counterpart unless otherwise specified, where p<0.05, Student’s t-test. BAll of the results represent n = 3-4 independent studies.  139  Figure 5-6. Collagen IV increases the doubling time of MIP101 cells over-expressing the various biological domains of SPARC. A-C) MIP/SP-N, MIP/SP-F, and MIP/SP-C CRC cells were plated on control or CIV-coated (50 µg/cm2) plates. Cellular number was assessed 3-4 days after seeding on control ( ) or CIV-coated (■) plates. *, statistical difference compared to CIV counterpart, where p<0.05, Student’s t-test. D) Doubling time of CRC cells on control (■) or CIV-coated (□) plates were calculated based on the changes in cellular number. All of the results represent mean ± s.e. *, statistical difference to control counterpart unless otherwise specified, where p<0.05, Student’s t-test. All of the results represent n = 3-4 independent studies.  140  Figure 5-7. Collagen IV enhances chemosensitivity by up-regulating the apoptotic pathway of MIP101 cells over-expressing the various biological domains of SPARC. MIP/SP-N, MIP/SP-F, and MIP/SP-C CRC cells were plated on control or CIV-coated (50 µg/cm2) plates for 24 hours. Cells were then treated with 1000 µM 5-FU for 0 (■), 2 (□) or 4 ( ) hours and assayed for caspase 3/7 activity. All of the results represent mean ± s.e. *, statistical difference to untreated counterpart unless otherwise specified, where p<0.05, Student’s t-test. B) Changes in the expression of proteins involved in the apoptotic pathway following incubation with 1000 µM 5-FU after various time intervals. Total proteins were isolated from cells harvested at 0-4 hours following treatment with 1000 µM 5-FU and probed for various proteins (activated caspases and Bid are represented by their cleaved fragments). All of the results represent n = 3-4 independent studies.  141  Figure 5-8. The SP-C domain of SPARC influences collagen IV's effect on chemosensitivity. A-D) MIP101 cells stably-expressing full length SPARC (MIP/SP), SPARC’s various biological domains (MIP/SP-N, MIP/SP-F, MIP/SP-C), or their mutants (MIP/SP-Nmut1, MIP/SP-Nmut2, MIP/SP-Fmut1, MIP/SP-Fmut2, MIP/SP-Cmut1, MIP/SP-Cmut2) were plated on control or CIV-coated (50 µg/cm2) plates. 24 hours later, cells were treated with 0 (■) or 1000 (□) µM 5FU for an additional 24 hours and assayed for cell viability with WST assays. The percentages of viable cells were calculated based on the A450 reading of non-treated control cells. All of the results represent mean ± s.e. *, statistical difference, where p < 0.05, Student’s t-test. All of the results are representative of 3-4 independent experiments. Please see Table 2-2 for information on the specific mutations.  142  Figure 5-9. The SP-C domain of SPARC and collagen IV co-operatively decrease apoptosis. A-D) MIP101 cells stably-expressing full length SPARC (MIP/SP), SPARC’s various biological domains (MIP/SP-N, MIP/SP-F, MIP/SP-C), or their mutants (MIP/SP-Nmut1, MIP/SP-Nmut2, MIP/SP-Fmut1, MIP/SP-Fmut2, MIP/SP-Cmut1, MIP/SP-Cmut2) were plated on control or CIV-coated (50 µg/cm2) plates. 24 hours later, cells were treated with 0 (■) or 1000 (□) µM 5FU and assessed for caspase 3/7 activity after treatment with 0 (■) or 1000 (□) µM 5-FU for an additional 24 hours. All of the results represent mean ± s.e. *, statistical difference, where p < 0.05, Student’s t-test. All of the results are representative of 3-4 independent experiments. Please see Table 2-2 for information on the specific mutations.  143  Figure 5-10. A schematic representation of the cellular signaling events involved when grown on collagen IV. A schematic of the SPARC-CIV relationship that enhances apoptosis. In this study, we demonstrated that SPARC and CIV collectively increase chemosensitivity in CRC cells.  144  6. Discussion This thesis examined the mechanisms by which SPARC mediates apoptosis and resensitizes chemotherapy resistant colorectal cancers (CRCs). This included the specific signaling mechanisms of SPARC-mediated apoptosis, the biological fragment of SPARC responsible, and the effect collagen IV (CIV) on chemosensitivity. In this discussion, I will review the findings of the research, future directions, and significance of this research. 6.1 Review of findings Results from this thesis shed light on the mechanism by which SPARC re-sensitized therapy-refractory CRC to chemotherapy. In chapter 3, the specific mechanism of SPARCmediated apoptosis was identified. It showed that following chemotherapy treatment, SPARC interacts with the N-terminus of caspase 8. This interaction then activates the extrinsic and enhances the intrinsic pathway of apoptosis. After the identification of the specific signaling cascade, we then investigated which biological domain of SPARC was responsible. SPARC contains three biological domains, each with its own modular function (238). In chapter 3, we found that the N-terminus of SPARC, the most divergent among the SPARC-like family (62), activates apoptosis in a similar manner as full-length SPARC. The N-terminal domain of SPARC interacted with pro-caspase 8 to activate apoptosis and promote tumour regression. The discovery of a SPARC:pro-caspase 8 interaction led to further questions about other proteins that interacted with caspase 8. It has previously been established that Bcl-2, an anti-apoptotic protein, interacts with caspase 8 and inhibits caspase 8 mediated apoptosis in neuroblastomas (232). We showed that the SPARC:caspase 8 interaction augmented apoptosis in chemotherapy sensitive cells; however, in therapy-refractory cells, apoptosis was inhibited through a Bcl-2 and caspase 8 interaction. This led to additional experiments to determine if  145  SPARC and Bcl-2 competed to bind with caspase 8, and we showed that SPARC and Bcl-2 indeed compete to interact with caspase 8. Finally, we investigated how the ECM influenced SPARC’s ability to re-sensitize CRC cells. SPARC is a matricellular protein that interacts with components of the extracellular matrix (ECM), such as collagen IV, but does not serve a structural role (61, 62). In chapter 5, we investigated the effect of collagen IV (CIV) on CRC cells and how this effect is changed in the presence of SPARC. Results showed that CIV sensitized CRC cells to chemotherapy, by enhancing the intrinsic pathway of apoptosis. Finally, we showed that SPARC enhanced CIVinduced chemosensitizing effects through the SP-C domain of SPARC. This thesis contributes to the understanding of SPARC’s role in tumourigenesis. It identified various mechanisms that SPARC utilizes to augment apoptosis and tumour regression. The mechanisms identified were the specific signaling pathways regulated by SPARC, the specific interaction between SPARC and caspase 8, and the competition between SPARC and Bcl-2 to bind to caspase 8. The SPARC, Bcl-2 and caspase 8 potential triad could potentially be exploited for treatment as inhibiting the Bcl-2:caspase 8 interaction, with SPARC, would allow for the SPARC:caspase 8 interaction and subsequent activation of the extrinsic pathway of apoptosis and greater tumour regression. As well, we found that CIV and SPARC collectively augmented apoptosis. This identifies CIV as another component that could be added to a combination therapeutic regimen for the treatment of colorectal cancer. Results have also shown that the combination treatment of exogenous SPARC and chemotherapy decreased cell viability, increased apoptosis, and inhibited the Bcl-2:caspase 8 interaction. As well, in vivo treatment with exogenous SPARC and chemotherapy inhibited CRC tumour growth (113). Yet, as shown in chapter 3, only intracellular SPARC was able to exert its chemosensitizing effects. The biotin/avidin complex could have affected SPARC and inhibited  146  its activity, or it is necessary that SPARC enters the cell to exert its effects. This leads to the question of how exogenous SPARC enters the cell. There are many possible mechanisms of how a protein can enter a cell. Some possible trafficking mechanisms include bulk endocytosis or receptor-mediated endocytosis (51). To date, the mechanism that SPARC utilizes to cross the cell membrane has not been determined. The mature protein is only 32kDa, making any of the above options feasible. However, recent progress has been made in identifying potential cell surface receptors for SPARC. These include α5β1 in adipose stromal cells (230) and β1 integrin in lens epithelial cells (231). Additional research is needed to identify how SPARC enters a cell, allowing SPARC to exert its chemosensitizing effects. This discovery, that SPARC needs to be intracellular to effectively enhance apoptosis, may explain why SPARC augments tumour growth in particular cancer types. Possible mechanisms SPARC utilizes to promote tumour growth may be due to the localization of SPARC or perhaps protein trafficking is dysregulated; thus, SPARC remains extracellular and unable to bind to pro-caspase 8 to activate apoptosis. Given the complex role of SPARC in cancer, it is also possible that the dysregulation of a SPARC receptor would contribute to the functionality of SPARC and possibly the efficacy of SPARC treatment. The identification of the specific mechanism(s) of SPARC-entry into the cell could also be critical in the use of SPARC as a small-molecule inhibitor of Bcl-2. Also, there have been recent technologies identified to improve cellular uptake to the cell, increasing the efficacy of potential therapies. These include using Vitamin B12 as a carrier (287), inhibiting efflux pumps (288), and the use of cell-penetrating peptides (289). The use of these technologies could also help the efficacy of SPARC as a potential therapy. In chapter 5, it was found that collagen IV (CIV) further sensitized CRC cells to chemotherapy. However, these results are preliminary and based only on in vitro experiments 147  using CIV-coated plates. Others have shown that specific fragments of collagen have greater apoptotic activity than others (275, 279). In this thesis, the most effective collagen fragment to re-sensitize resistant CRC cells to chemotherapy was not determined. The use of specific CIV fragments and in vivo experiments are needed to validate the in vitro co-operative effects of CIV and SPARC on cell viability and apoptosis. Also, the ECM is known to be dysregulated in cancers. It is possible that other components of the ECM may protect CRC cells from chemotherapy treatment. Based on the complexity of the ECM, questions arise regarding whether specific components of the ECM modulate the efficacy of chemotherapy drugs. For example, the microenvironment between the primary and metastatic site can differ, varying its response to chemotherapy. Identification of the specific components of the ECM that influence its response to chemotherapy, and whether these components differ between different organs would help validate the effect of the ECM and SPARC on tumour growth. 6.2 Future directions Further study of SPARC biology is still required to fully understand the complexity of this highly interesting protein. As discussed above, the mechanism(s) by which SPARC enters the cell could provide great insight into how SPARC modulates tumourigenesis. If SPARC entry is receptor-mediated, immunoprecipitation and mass spectrometry experiments will identify potential receptor. Another approach is to fluorescently-label SPARC and use live microscopy to determine if entry occurs by endocytosis or diffusion through the cell membrane. One also wonders about the effect of SPARC on metastasis. In this thesis, metastasis does not occur in the tumour xenograft model used in the in vivo studies. Many biological processes are required for cells to metastasize. Examples include, but are not limited to, invasion of the ECM, movement to the site of interest, re-implantation, and vascularization. In various 148  cancers, SPARC has shown the ability to affect these metastasis-related processes (290). For example, SPARC is known to function as a de-adhesive protein, which leads to speculation about its role in metastasis. This thesis largely focuses on apoptosis. Studies in the Tai laboratory have also shown decreased expression of CD34, a marker for vasculature, in SPARC overexpressing CRC cells (113). Determining whether SPARC mediates processes that affect metastasis such as invasion or angiogenesis would be of great significance. Previous results from the Tai laboratory showed that SPARC expression is decreased in resistant CRC cells (113). There are various mechanisms that can attribute to the downregulation of SPARC. Two studies in CRC showed that this down-regulation is a result of methylation of the SPARC promoter (108, 109). Further, 5-aza-2’-deoxycytidine up-regulated SPARC expression (109). Other mechanisms of transcription regulation include chromatin remodeling, histone acetylation, repressors, activators, enhancers, transcription factors, or even post-translational modifications (51). Further studies to determine if dysregulation of any of these mechanisms results in decreased SPARC expression in advanced cancers could also be of great therapeutic significance. In chapter 5, I observed that CIV re-sensitizes CRC cells to chemotherapy. In CRCs, it is not known whether a specific fragment of CIV is responsible for this effect and/or if SPARC specifically interacts with a certain type of CIV. Additional in vitro and in vivo studies using specific CIV fragments would provide greater detail about the chemosensitizing effect of CIV. SPARC is also known to interact with other ECM matrices (62). Marastoni et al. have shown that alteration of the ECM is responsible for diseases such as cancer (37). Thus, if SPARC interacts with other ECM matrices, the knowledge of whether these matrices promote or inhibit tumour growth may provide insight on why SPARC acts as a proto-oncogene or a tumour suppressor in various cancers.  149  Additionally, SPARC is a de-adhesive protein (62). This leads to further questions on whether SPARC affects anoikis, which is defined as apoptosis induced by incorrect cell/ECM attachment. Metastatic cancer cells have the ability to evade anoikis (291). As SPARC can prevent cell attachment, it could also affect the ability of cancer cells to attach to its metastatic site through anoikis. Specific experiments to investigate this would include prevention of cell attachment (eg. using poly-HEMA), variation of SPARC levels, and assays for cell viability and/or apoptotic activity. 6.3 Significance In this thesis, I showed that SPARC affects many biological processes that contribute to tumour growth. These processes include cellular proliferation and apoptosis. The ability to regulate multiple processes makes SPARC an attractive candidate to use as treatment for malignancies. Various studies have supported the use of SPARC as a cancer therapy. Most interestingly, exogenous treatment with SPARC reduced the growth of pancreatic cancers (123), CRCs (113), neuroblastomas (292), and leukemia cells (111). In melanoma, SPARC enhanced the chemosensitivity of cells to cisplatin treatment (195). The Tai laboratory has also shown that SPARC can enhance sensitivity of resistant CRC cells to chemotherapy (113) and vitamin D (117). The use of SPARC treatment has also been observed in vivo. In animal studies, SPARC, when used as a purified protein (113, 292) or in gene therapy using viral expression systems (113, 293, 294), has been shown to be an effective cancer therapeutic. Results from these studies and this thesis support the use of SPARC, more specifically, the N-terminal of SPARC, as a therapeutic agent. Also, as identified in this thesis, SPARC competes with Bcl-2, which points to its use as a potential small molecular inhibitor.  150  6.4 Conclusions SPARC is a complex protein as it is both up- and down-regulated in different cancers. It is known to regulate many biological processes such as proliferation, angiogenesis, adhesion, apoptosis, ECM remodeling, tumourigenesis, and invasion. 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