BIOACTWATION OF BY HUMA. NADPH-CYTOCHROME P450 REDUCTASE

Shairoj Ramji

A thesis submitted in confonnity with the requirements for the degree of Master of Science Graduate Department of Phannacology University of Toronto

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Bioactivation of doxorubicin by human NADPH-cytochrome P450 reductase. Master of Science, 1998. Shairoj Ramji, Department of Pharmacology, University of Toronto.

Doxorubicin (DOX) is a useful antineopIastic dmg with multiple mechanisms of cytotoxicity. One such mechanism involves the reductive bioactivation of the quinone ring to a semi-quinone radical, and subsequent redox-cycling. It is known that rat NADPH- cytochrome P450 reductase (CYPRED)catdyzes DOX reduction and sensitizes human cancer cells to DOX in vitro. The purpose of this investigation was to detemine whether human CYPRED also catalyzes DOX reduction. DOX reduction occurred in a human

CYPFED cDNA heterologous expression system and also in human liver microsomes.

Using a bank of 17 human Iivers, the rate of microsomal DOX reduction was correlated with

CYPRED activity and immunoreactive protein expression. DOX reduction was inhibited by diphenyliodonium chloride, an inhibitor of CYPRED. DOX reduction was not inhibited by carbon monoxide, suggesting that the cytochromes P450 did not catalyze DOX reduction.

These results suggest that human CYPRED catalyzes DOX reduction. A marnmalian expression vector containing human CYPRED cDNA was constmcted for future transfection into breast cancer cells to assess whether sensitivity to DOX will be increased. ACKNOWLEDGEMETS

1 foremost would like to thank my supervisor, Dr. D. S. Riddick, for providing me with the opportunity to pursue this research and for being such a wonderful advisor. 1 am especially grateful for his encouragement, knowledge and expertise, approachability, understanding, and general day-to-day guidance and support.

1 also would like to thank Dr. T. Inaba, not only for providing the hurnan liver microsomes, but also for his scientific counsel and discussions.

For their scientific and technical contributions, and especially for their great encouragement, advice, support, generosity and friendship, I would like to thank my lab mates, Chunja Lee and Anahita Bhathena. Also, for scientific guidance and technical support, 1 thank Peter Beaumont, Judy Wong, Dr. Peter McPherson, Yoav Timsit, and John

Giamone who are al1 from the Department of Pharmacology.

Several individuals have provided equipment and materiais which have made this research possible. Human NADPH-cytochrome P450 reductase cDNA was a generous gift from Dr. Frank Gonzalez (National Cancer Institute, Bethesda, MD). 1 would like to thank

Dr. Gerald Batist (Jewish General Hospital, Montreal) for providing the MCF-7/ADR breast cancer ce11 line. For their assistance with the breast cancer ce11 lines, thanks also to Dr.

James Ballinger and Pat Firby (Princess Margaret Hospital/Ontario Cancer Institute). For assistance in preparing the human liver microsomes, 1 am grateful to Dr. Xu, Nancy Fischer, and Tommy Cheung from Dr. Inaba's Laboratory. Thanks are also due to Dr. A.B. Okey and Dr. G.J. Goldenberg for use of Iaboratory space and equipment.

iii Thanks especially to Alison Feniak, Sanya Petrovic, Linda Toy and Lois Haighton for their encouragement, advice, and constant great belief in my abilities. And, of course, thanks to my Murn and Dad for always being there.

1 also gratefully acknowledge the Canadian Breast Cancer Foundation and the

University of Toronto School of Graduate Studies for financial support to conduct this research. TABLE OF CONTIZNTS

ACKNOWLEDEMENTS ...... iii

LISTOFTABLES ...... viii

LIST OF FIGURES ...... ix

LIST OF ABBREVIATIONS ...... x

Introduction ...... 1 1.1 Statement of Research Problem ...... 1 1.2 Breast Cancer and Breast Cancer Treatment ...... 1 1.3 Doxorubicin ...... 3 1.3.1 Mechanisms of action ...... 5 1.3.2 Mechanisms of resistance ...... 5 1.4 Doxorubicin Biolansformation and Mechanism of Free Radical Production ...... 13 1.4.1 Doxorubicin metabolism ...... 13 1.4.2 Enzymatic fiee radical production ...... 15 1.4.3 Other mechanisms of free radical production ...... 22 1.5 Evidence of Toxicity from Doxorubicin-Induced Free Radicals and Redox-Cycling ...... 24 1.5.1 DNA damage ...... 24 1S.2 Lipid peroxidation ...... 24 1.5.3 Ce11 toxicity ...... 25 1.5.4 Cardiotoxicity ...... 32 1.5.5 Toxic effects of DOX semi-quinone and other DOX radicals ...... 34 1.5.6 The relative importance of free radicals in the mechanism of cytotoxicity of doxorubicin ...... 37 1.6 Enzymology of Free Radical Production from Doxorubicin ...... 39 1.6.1 Evidence for involvement of NADPH-Cytochrome P45O reductase in doxorubicin reduction ...... 39 1.6.1.1 Enzyme kinetic studies ...... 40 1.6.1.2 Transfection studies ...... 40 1.6.2 Other enzymes involved in doxorubicin reduction ...... 41 1.6.3 Other bioreductive antineoplastic agents ...... 42 1.7 NADPH-Cytochrome P450 Reductase ...... 43 1.7.1 Function. structure and regdation ...... 43 1.7.2 Tissue distribution ...... 48 1.7.3 Occurrence in normal and tumour human breast tissue ...... 48 1.8 Gap in Knowledge Leading to the Research Hypothesis ...... 54 1.9 Research Hypotheses ...... 55 1.10 Experimental Approach and Rationale ...... 56

2 . Methods ...... 58 2.1 Chemicals. Reagents and Source of Materials ...... 58 2.2 SampIe Preparation ...... 59 2.2.1 Rat liver microsornes ...... 59 2.2.2 Human liver microsornes ...... 60 2.2.3 Protein analysis ...... 61 2.3 Enzyme Assays and Other Analytical Techniques ...... 63 2.3.1 CYPRED activity ...... 63 2.3.2 DOX reduction ...... 64 2.3.3 Inhibition studies ...... 66 2.3.4 Irnmunoblot analysis ...... 68 2.4 Ce11 Culture ...... 71 2.4.1 Ce11 lines and culture techniques ...... 71 2.4.2 Growth curves ...... 73 2.4.3 MIT cytotoxicity assay ...... 73 2.5 Molecular Biology Techniques ...... 75 2.5.1 Construction of a mammalian expression vector containing human CYPRED cDNA ...... 75 2.5.1.1 Small-scale preparation of vector DNA ...... 77 2.5.1.2 Restriction enzyme digestion and agarose gel electrophoresis ...... 82 2.5.1.3 Purification of DNA from agarose gels ...... 84 2.5.1.4 DNA ligation ...... 85 2.5.1.5 Bacterial transformation ...... 86 2.5.2 DNA sequencing ...... 87 2.5.3 Optimization of antibiotic concentration for selection of stable transfectants ...... 89 2.6 Statistical Analysis ...... 90 3. ResuIts ...... 91 3.1 Hypothesis #l ...... 91 3.1.1 Optimization of enzyme assays and immunoblots ...... 91 3.1.1.1 CYPRED activity ...... 91 3.1.1.2 DOX reduction ...... 93 3.1.1.3 Immunoblots ...... 96 3.1.2 CYPRED activity and DOX reduction in microsomes from human lymphoblastoid cells ...... 98 3.1.3 Correlation analysis using a bank of human liver microsornes ....100 3.1.3.1 DOX reduction with CYPRED activity ...... 100 3.1.3.2 DOX reduction w ith immunoreactive protein ...... 102 3.1.4 Inhibition studies ...... 104 3.1.4.1 Chernical inhibitor ...... 104 3.1.4.2 Antibody inhibitor ...... 106 3.1.4.3 Carbon monoxide ...... IO8 3.1.4.4 Inhibition of CYPRED activity by DOX ...... 109 3.2 Hypothesis #2 ...... 111 3.2.1 Ce11 culture ...... 111 3.2.1.1 Growth properties ...... 111 3.2.1.2 MTT assay ...... 113 3.2.1.2.1 Initial characterization ...... 113 3.2.1.2.2 DOX toxicity ...... 117 3.2.2 Construction of a mammalian expression vector containing human CYPRED cDNA ...... 119 3.2.3 Optimization of antibiotic concentration for selection of stable transfectants ...... 126 4 . Discussion ...... 128 4.1 Hypothesis #1 .Role of Human CYPRED in DOX Bioactivation ...... 128 4.2 Hypothesis #2 .Sensitization of Human Breast Cancer Cells to DOX by Overexpression of CYPRED ...... 135 4.3 Future Research Directions ...... 139 4.4 Surnrnary and Implications of Research ...... 141 5 . References ...... 143 List of publications and abstracts arising from thesis ...... 168 APPENDIX A List of Chernicals and Reagents With Their Sources ...... 169 APPENDIX B Characteristics of Liver and Kidney Donors ...... 172

vii LIST OF TABLES

Table 1: Cytochromes P450 Detected in Normal and Turnour Tissue from Human Breast Samples by Immunoblotting or Immunohistochemistry ...... 50 Table 2: Control Groups Used To Measure DOX-stimulated NADPH Oxidation . . 65 Table 3: Standard Setup for the MTT Assay in a 96-Well Plate ...... 75 Table 4: Effect of CO on CYPRED Activity and DOX Reduction in Rat and Hunan Liver Microsornes ...... 108 Table 5: Methods for Determinhg the Role of an Enzyme in the Biotransformation of a Compound ...... 128

viii LIST OF FIGURES

Figure 1: The Chemical Structure of Doxorubicin (DOX) ...... 4 Figure 2: Biotransformation of Doxorubicin to Less Toxic Metabolites ...... 14 Figure 3: Redox-cycling of Doxorubicin. Toxicity and Antioxidant Defences ...... 16 Figure 4: Formation. Metabolic Fate. and Potential Toxicity of the Doxorubicin Semi-Quinone ...... 20 Figure 5: The Catalytic Cycle for a Cytochrome P450-mediated Hydroxylation Reaction ...... 44 Figure 6: Representation of the Functional Domains of NADPH-Cytochrome P450 Reductase ...... 46 Figure 7: Construction of Mammalian Expression Vector with Human CYPRED cDNA ...... 78 Figure 8: Map of pUC9 Vector with CIoned CYPRED cDNA ...... 81 Figure 9: Map of pcDNA3.1 (+) Expression Vector into which the CYPRED cDNA was Subcloned ...... 81 Figure 10: CYPRED Activity in Human Liver Microsornes ...... 92 Figure 1 1: Optimization of DOX Reduction in Rat Liver Microsornes ...... 94 Figure 12: DOX Reduction in Human Liver Microsomes ...... 95 Figure 13: Imrnunoblot of CYPFED Protein Expression in Human Liver .Standard Curve of Protein Concentration Versus Immunoreactivity ...... 97 Figure 14: CYPRED Activity and DOX Reduction in Microsomes from Human Lymphoblastoid Cells ...... 99 Figure 15: Correlation of CYPRED Activity and DOX Reduction in a Bank of HumanLivers ...... 101 Figure 16: Correlation of CYPRED Protein Expression with CYPRED Activity and DOX Reduction in a Bank of Hurnan Livers ...... 103 Figure 17: DPIC Inhibition of CYPRED Activity and DOX Reduction in Rat (A) and Hurnan (B) Liver Microsornes ...... 105 Figure 18: Antibody Inhibition of CYPRED Activity and DOX Reduction in Rat (A) and Human (B) Liver Microsornes ...... 107 Figure 19: Inhibition of CYPRED Activity by DOX in Rat (A) and Human (B) Liver Microsomes ...... 110 Figure 20: Growth Curve of MCF-7lWT and MCF-7/ADR Ce11 Lines ...... 112 Figure 2 1: Deterrnination of Relationship between Absorbance and Ce11 Number in the MTT Assay ...... 114 Figure 22: Assessrnent of Ce11 Growth in Microculture Over Tirne ...... 116 Figure 23: Sensitivity of MCF-7lWT and MCF-7/ADR Cells to DOX ...... 118 Figure 24: Restriction Enzyme Digestion of pUC9 to Isolate the CYPRED cDNA ...... 120 Figure 25: Restriction Enzyme Digestion of pcDNA3.1 (+) from Ligation Reaction with CYPRED cDNA ...... 122 Figure 26: Sequence of Hurnan CYPRED cDNA and the Portion of the Sequence Identified by DNA Sequencing ...... 124 Figure 27: Optimization of Antibiotic Concentration for Selection of Stable Transfectants ...... 127 LIST OF ABBREVIATIONS

ANOVA one-way analysis of variance ATCC American Type Culture Collection ATP adenosine triphosphate BIS N, N'-methylenebis acrylarnide BSA bovine senuri albumin BSO buthionine sulfoximine cDNA complementary DNA CHO Chinese hamster ovary CIAP calf intestinal alkaIine phosphatase CMV cytomegalovirus promoter CO carbon monoxide CO2 carbon dioxide CTP cytidine triphosphate CYPRED NADPH-cytochrome P450 reductase dATP deoxyATP dCTP deoxy CTP DDT dichlorodiphenyl trichloroethane dGTP deoxyGTP DMPO 5,s-dimethyl- 1-pyrrolidine-N-oxide dm deoxy TïP DOX doxorubicin DPIC diphenyliodonium chloride DMSO dirnethyl sulfoxide DNA deoxyribonucleic acid DTPA diethylenetriaminepentaacetic acid E. coli Escherichia coli EDTA ethylenediaminetetraacetic acid ESR electron spin resonance FBS fetal bovine (or calf) serum GSH reduced glutathione GSHPx glutathione peroxidase GSSG oxidized glutathione GST glutathione S-transferase GTP guanosine triphosphate H+ hydrogen ion (proton) H20 water H202 hydrogen peroxide HCI hydrogen chloride (hydrochloric acid) HEPES N-(2-hydroxyethyl]piperazine-N'-(2-ethanesulfonic acid) HRP horseradish peroxidase Go concentration inhibiting 50% of growth or enzyme activity KCl potassium chloride Km the Michaelis constant LB Luria-Bertani medium MDR multidrug resistance a-MEM a-minimal essential medium MgCl2 magnesium chloride MRP multidrug resistance associated protein MTT [3-(4,5-dimethylthiazo1-2-yl)-2,5-diphenyteoli bromide] NADP+ nicotinamide adenine dinucleotide phosphate (oxidized form) NADPH nicotinamide adenine dinucleotide phosphate (reduced form) NAD+ nicotinamide adenine dinucleotide (oxidized form) NADH nicotinamide adenine dinucleotide (reduced form) NaOH sodium hydroxide NQOl NAD(P)H: quinone oxidoreductase 0;. superoxide OH* hydroxyl radical P450 cytochrome P450 PBS phosphate buffered saline RNA ribonucleic acid rPm revoIutions per minute SD standard deviation SDS sodium dodecyl sulfate SDS-PAGE sodium dodecyl sulfate polyacrylarnide gel electrophoresis SV40 simian virus 40 promoter TAE 40 mM Tris-acetate/l rnM EDTA, pH 8.0 TBE 0.09 M Tris-borate12 mM EDTA, pH 8.3 TCDD 2,3,7,8-tetrachlorodibenzo-p-dioxin TE 10 rnM Tris-HC1/1 rnM EDTA, pH 8.0 TEG 25 mM Tris110 mM EDTAJSO rnM glucose TEMED N,N,N' ,N'-tetrarnethylethylenediamine TNT 20 mM Tridl37 mM NaCl/O. 1% Tween-20, pH 7.6 Tris Tris(hydroxymethy1)aminomethane TTP thymidine triphosphate UV ultraviolet vm maximal velocity of an enzymatic reaction 1 INTRODUCTION

1.1 Statement of Research Problem

Breast cancer is a major heakh concern for many Canadian women. For adjuvant

and treatment of metastatic disease, doxorubicin is one of the drugs of choice.

However, irs effectiveness is lirnited by severe toxicity and drug resistance. One way to

enhance the effectiveness of doxorubicin is to enhance its mechanisrn of action at the level of

the tumour cells. This research project evaluates the role of the drug biotransformation

enzyme, NADPH-cytochrome P450 reductase, in bioactivating doxorubicin and sensitizing

breast cancer cells to the drug.

1.2 Breast Cancer and Breast Cancer Treatment

Breast cancer is the most fiequently diagnosed form of cancer in Canadian and

Arnerican women, accounting for approximately 30% of al1 new cancer cases (Landis et al.,

1998; NCI, 1998). Breast cancer also is the second leading cause of cancer death in these

women. The National Cancer Institute of Canada (NCI, 1998) estirnates that in 1998, 19,300

new cases of breast cancer will be diagnosed with 5,300 breast cancer deaths occurring. The

Arnerican Cancer Society estimates that in the United States, 178,700 new breast cancer

diagnoses wilI be made with 43,500 deaths (Landis et al., 1998). This is quite significant,

since current and predicted incidence rates of breast cancer in Canada and the United States

are the highest in the world (Gaudette et al., 1996; NCI, 1998).

Treatment of breast cancer involves a number of therapies depending on the tumour and nodal stages of the disease as well as the hormonal receptor status of the tumour and menopausal status of the patient (Allum and Smith, 1995; Dawson and Taylor, 1995; 2 Overmoyer, 1995). Generally, treatment of the cancer at the local level involves surgical removal of the primary tumour and radiation therapy. This may be followed by adjuvant therapy, which is administration of chemotherapeutic drugs to patients who may have a high risk of occult micro-metastases but no clinical or radiologic evidence of metastatic disease

(Tamock and Hill, 1992). Occasionally, drug treatment is also given prior to treatment of the primary tumour (neoadjuvant therapy). For advanced cases of breast cancer, particularly if there is significant metastases, chemotherapy is the mainstay of treatment. Depending on the hormone receptor status of the tumour cells, anti-estrogenic drugs (eg. tamoxifen) may also be admidstered. Quite often the chemotherapeutic regirnen is a combination of antineoplastic drugs, which for breast cancer prirnarily include doxorubicin, , , and/or (Seeger and Woodcock, 1995; Medical

Letter, 1996). Doxonibicin (adriarnycin) is one of the most effective drugs against breast cancer and is the basis for many combination-based chemotherapeutic treatments (Seeger and

Woodcock, 1995).

Chemotherapy, however, does have its limitations. Firstly, since the dnigs distinguish poorly between normal and malignant cells, systemic toxicity is the most cornmon and most serious consequence of chemotherapy. In breast cancer patients, toxic effects include bone marrow suppression leading to increased risk of infection, skin changes (eg., alopecia, hyperpigmentation), gastrointestinal side effects (nausea, vomiting, diarrhea), amenorrhea, cardiotoxicity, and the risk of second malignancies (Overrnoyer, 1995; Seeger and Woodcock, 1995). While the doses for chemotherapy are chosen to maximize the therapeutic usefulness of treatment with tolerable toxicity, it remains that drug toxicity is a major negative aspect for breast cancer chemotherapy.

Secondly, the successful use of chemotherapy rnay be limited by drug resistance.

Drug resistance rnay be a result of physiological factors (eg., tumour blood flow, pH and oxygenation) or biological factors (ie., tumour ce11 heterogeneity). However, quite often drug resistance refers to biochemical resistance in which specific cellular macrornolecules are responsible for the drug resistance (Ringborg and Platz, 1996). Tumours rnay be intrinsically resistant to drugs; that is they display oniy a minima1 response, or none at all from initiation of treatment (Tannock, 1992b). Some tumours rnay also display acquired resistance; that is the tumour responds to initial treatment but rnay soon become unresponsive to the drug.

Therefore, both drug resistance and systemic toxicity limit the usefulness of chemotherapy in breast cancer treatment. It would be desirable to enhance the effectiveness of antineoplastic drugs such that lower doses rnay be used resulting in less systemic toxicity.

One way of enhancing the effectiveness of a drug is by maxirnizing its possible mechanism(s) of action selectively in tumour cells, and identifying and minirnizing its possible rnechanisms of resistance. This will be considered for the drug doxorubicin in this thesis.

1.3 Doxorubicin

Doxorubicin (DOX) consists of a water-insoluble planar four-ring

(structures ABCD) linked to the water-soluble sugar daunosamine (Figure 1). Other similar anthracycline drugs are and . DOX is produced by the fungal species

Streptomyces and has been used clinically since the 1970s for a wide variety of solid tumours and leukemias (Keizer et al. , 1990; Erlichrnan, 1992; Tamock, l992a). Figure 1: The Chemïcal Structure of Doxorubicin @OX) DOX consists of a water-insoluble planar four-ring anthracycline (structures ABCD) linked to the water-soluble sugar daunosamine at Carbon 7. Several parts of the molecule are involved in the metabolism of DOX. Ring C is the quinone group which can be activated into a semi- quinone radical after one-electron reduction. The iron-chelation site is probably fonned by the oxygen atoms of Carbons 11 and 12. Iron at this site can be reduced by the doxorubicin molecule either by oxidation of the hydroquinone (Ring B) or by oxidation of the side chain at carbon 9 (Keizer et al. , 1990). 1.3.1 Mechanisms of action

A number of mechanisrns of action appear to contribute to the cytocidal effect of

DOX (Erlichman, 1992). DOX intercalates DNA between base pairs perpendicular to the long axis of the double helix, leading to partial unwinding of the DNA helix. DOX also binds covalently to the 'enzyme topoisomerase II which may lead to single- and double-strand

DNA breaks. Topoisornerase II is an enzyme involved in the cleavage, unwinding and rejoining of segments of DNA during DNA and RNA synthesis. DOX appears to stabilize the cleavage in the DNA strands by binding to the topoisomerase II-DNA complex, thereby preventing re-ligation of the DNA and leading to ce11 death (Tewey et al., 1984; Erlichman,

1992; Tannock, 1992b). Therefore, the drug inhibits both RNA and DNA synthesis and has preferential toxicity for cells in the of the ce11 cycle.

DOX may also be biotransformed to a free radical (ie., containing an unpaired electron) which may directly produce toxic effects or whicli may, in turn, react rapidly with oxygen to produce superoxide (O,-.) causing oxidative stress and ultimately cell death.

This mechanism is discussed further in Section 1.4. Studies have also indicated that DOX may bind to ce11 membranes and rnay kill cells through membrane-related effects, at times without entering the ce11 (Murphree et al., 1981; Tritton and Yee, 1982; Tritton, 1991). The mechanisms of action of DOX are complex and only partly understood. Furthemore, it is not clear which particular mechanism or combination of mechanisms actually kills the cells

(Speth et al., 1988; Erlichman, 1992).

1A2 Mechanisms of resistance

Dmg resistance to DOX is known to occur and this is mediated by many possible mechanisrns. Ce11 Iines selected to be resistant to DOX often have cross-resistance to a

number of other drugs (eg., , , puromycin). These dmgs rnay have very

different chemical structures and usually are natural cornpounds. Ce11 lines expressing this

type of cross-resistance are considered to have the multidrug resistance (MDR) phenotype

which is associated with overexpression of a 170 kD transrnembrane glycoprotein, the P-

glycoprotein encoded by the MDRl gene. This protein is found in plasma membranes and

functions as an energy-dependent efflux pump which lowers the cellular accumulation of

DOX and other cross-resistant drugs (Tannock, 1992b). Clinically, in untreated breast

cancer patients MDRl expression is low; however, there are indications that some patients

treated with may develop tumours with increased expression of the P-

glycoprotein (Goldstein et al., 1989; Merkel et al., 1989; Schneider et al., 1989).

In other in vitro studies, resistance to DOX has been associated with decreased

activity of type II topoisomerases and/or the presence of different forms of the enzyme whose

activity does not facilitate drug-induced DNA damage (Morrow and Cowan, 1990; Bugg et

al. , 1991; Evans et al. , 1994; Gudkov et al., 1993). Currently , the clinical relevance of this

mechanism of drug resistance is unknown (Ringborg and Platz, 1996). Another possible

mechanism of resistance involves the p53 tumour suppressor gene. This gene codes for a

nuclear phosphoprotein (p53) which binds to specific DNA sites and stimulates the

downstream expression of genes that control growth. Wild-type p53 plays an essential role

in the negative regulation of ce11 growth and apoptosis, and acts as a tumour suppressor protein in a ce11 cycle-dependent manner (Ogretrnen and Safa, 1997). In several ce11 Iines

including breast cancer ce11 lines, loss of normal p53 function has been associated with 7 resistance to DOX and other drugs (Ogretman and Safa, 1997). This loss of p53 function may occur through loss of p53 alleles, cytoplasmic sequestration or point mutations (Chen et al., 1990a; Lowe et al. , 1994; Bergh et al., 1995; Jansson et al., 1995; Lim et al. , 1996;

Ogretman and Safa, 1997). In clinical studies approximately 24% of breast cancer patients had alterations in the p53 gene and this may affect the prognosis of the disease (Soussi et al.,

1994; Bergh et al., 1995; Linn et al., 1996).

Other possible mechanisms of resistance to DOX may involve the glutathione S- transferases (GST), glutathione peroxidases (GSHPx) and the multidrug resistance associated protein (MRP), al1 of which relate to glutathione metabolism. GSTs are a family of detoxification enzymes which catalyze the conjugation of electrophilic substances with glutathione (GSH). In humans five classes of cytosolic GSTs referred to as alpha, mu, pi, theta, and zeta have been identified. The premise of this mechanism of resistance is based on the free radicalhedox-cycling mechanism of action of DOX. Currently, there is no evidence that glutathione conjugates of DOX or its primary metabolites are fonned in the presence of ce11 constituents or whole cells (Dixon et al., 1995). However, DOX may undergo redox-cycling leading to the production of reactive oxygen species which in turn may lead to lipid peroxidation. DOX has been shown to increase production of toxic lipid peroxides and hydroxyalkenals, which may be detoxified by GST-catalyzed glutathione peroxidase activity and glutathione conjugation, respectively (Alin et al, 1985 ; Singhal et al. ,

1992; He et al., 1996; Hubatsch et al., 1998). Therefore, one would expect that high expression of GSTs wou1d be associated with BOX resistance. However, whether this is a significant mechanism of DOX resistance in breast cancer is debatable. A number of 8 investigators have reported that breast tumours have elevated GST expression or activity

compared to normal peritumoral tissue (Di Ilio et al., 1985; Forrester et al., 1990; Albin et

al. , 1993; Kelley et al. , 1994).

The extent of GST expression has also been characterized in the human breast cancer

ce11 line MCF-7,which has played an important role in studies of drug response in breast

cancer (Levenson and Jordan, 1997). This is an estrogen-responsive ce11 line derived from a

fernale patient with a breast adenocarcinorna. A DOX-resistant derivative of the ce11 line

(MCF-7lADR; 192-fold more resistant) was originally isolated by growing MCF-7lWT cells

in the presence of stepwise increasing concentrations of DOX (Batise et al., 1986). The

MCF-7IADR ce11 line was characterized by a typical MDR phenotype, decreased intracellular

drug accumulation (Cowan et al., 1986), overexpression of the P-glycoprotein drug-effiux

pump (Fairchild et al., 1987), elevated GST pi (Batist et al., 1986) and selenium-dependent

glutathione peroxidase activity (Sinha et al., 1987a; 1987b).

Although increased GST expression (especially GST-pi) has been reported to occur in

the MCF-7/ADR ce11 line (Batist et al., 1986; De La Torre et al. , 1993), its role in drug

resistance is uncertain. In another study using MCF-7/ADR cells, increased expression of

GST was not reported (Chen et al., 1990b). Additionally , a drug-sensitive revertant from a

multidrug-resistant derivative of MCF-7 cells continued to express the high level of GST-pi

that was induced during drug selection (Yusa et al., 1988). Also, no consistent change in

DOX resistance was reported in drug-sensitive breast cancer ce11 lines transfected with GST cDNAs resulting in high GST expression (Moscow et al., 1989; Fairchild et al. , 1990;

Townsend et al., 1992). With the exception of one study in which the results were 9 inconclusive (Millward et al., 1990), no clinical studies were identified in the literature that related DOX therapy or resistance in breast cancer patients with GST expression or modulation.

Glutathione peroxidases (GSHPx)are selenium-dependent enzymes which utilize glutathione to reduce both hydrogen peroxide and more complex organic hydroperoxides (eg. fatty acid and phospholipid hydroperoxides) (Cheeseman and Slater, 1993). As with the GST enzymes, it has been hypothesized that increased expression of GSHPx may contribute to

DOX resistance by catalyzing the detoxification of the drug-induced fatty acid and lipid hydroperoxides (Dixon et al., 1995). In breast tumour tissue, increased activity of selenium- dependent GSHPx compared to normal peritumoral tissue was reported (Forrester et al.,

1990). Increased activity was also reported in multidrug-resistant MCF-7 cells which persisted in vivo when transplanted in mice (Cowan et al., 1986; Akman et al., 1990;

Mimnaugh et al., 1991; Townsend et al., 1991). AIso, in multidmg-resistant MCF-7 cells, purified GSHPx was reported to reduce DOX-induced free radical formation (Sinha et al.,

1989). Decreased cellular sensitivity to DOX was reported to occur, when purified GSHPx was introduced into cells by mechanical disruption of ce11 membranes (Doroshow et al.,

1991). Results from these in vitro studies suggest that increased GSHPx levels rnay contribute to DOX resistance.

A final mechanism which relates to glutathione metabolism and which may produce increased DOX resistance is the presence of the multidrug resistance associated protein

(MRP). MRP is a 180-195 kD membrane glycoprotein which is a member of the ATP- binding cassette transporter gene superfamily (Cole et al., 1992; Ringborg and Platz, 1996). 10 Overexpression of MRP in some drug-sensitive cancer ce11 lines resulted in increased resistaxice to several naturally occurring drugs, including DOX (Cole et al., 1992; Grant et al., 1994; Zaman et al., 1994; Breuninger et al., 1995; Lautier et al., 1996). Studies indicate that similar to P-glycoprotein, MRP prevents intracellular accumulation of these drugs. How MRP does this is not known. One theory is that MRP is a high-affinity transporter of glutathione conjugates and glucoronide conjugates at the plasma membrane and also into intracellular secretory vesicles (Muller et al., 1994; Jedlitschky et al., 1996; Van

Luyn et al., 1998). Therefore regarding DOX, once again resistance to DOX rnay be due to the efflux of glutathione conjugates of lipid peroxidation products fonned via GST-catalyzed reactions. However, this theory does not provide an explanation of the decreased cellular accumulation of DOX. Another theory is that MRP functions by effluxing xenobiotics and glutathione fi-om the intracellular cornpartment into the extracellular medium by a

CO-transportmechanism; ie., the glutathione and drug are not necessarily conjugated (Rappa et al., 1997). This would explain resistance to dmgs such as etoposide, DOX and which are not known to produce glutathione conjugates in cells. In vivo studies conducted in mice with complete loss of MRP expression, indicate that MWexerts a role in drug detoxification and glutathione metabolism (Lorico et al., 1997). Data are lirnited on the specifïc role of MRP in breast cancer cells. In MCF-7 cells increased expression of both

MRP and GST pi protected cells from DNA adduct formation and cytotoxicity associated with the carcinogen, 4-nitroquinoline 1-oxide (Morrow et al., 1998). The rate of conjugate formation of the carcinogen was also increased as was the MRP dependent efflux of the conjugate. In clinical studies, MW expression has been shown to be an important prognostic 11 factor in patients with breast cancer (Giaccone et al., 1995; Nooter et al., 1997).

As with the GSTs, drug resistance to a number of antineoplastic agents may be due to the presence, absence or varying amounts of other drug metabolizing enzymes, such as the cytochromes P450. Several anticancer drugs are adrninistered as which must undergo biotransformation to yield pharmacologically active compounds. Conversely several active drugs are administered which are biotransfonned to inactive pxoducts and are rapidly excreted. Dmg metabolizing enzymes are most concentrated in the liver, but are also found at lower levels in other tissues. The route of administration, phannacokinetics and biotransformation of a drug affect the amount of active drug that reaches the target and non- target tissues. This in turn determines the effectiveness and toxicity of the drug. Although substantial biotransformation occurs in the liver, the presence of P450s and other drug metabolizing enzymes in other tissues including tumours suggests that biotransformation at the local level rnay be significant in terms of drug response. In tumour cells, limited response or resistance to a drug may be due to metabolism of the drug into inactive compounds, or to the presence of low levels of bioactivating enzymes. For exarnple, growth of pancreatic carcinoma ceIl lines in the presence of daunorubicin led to the induction of carbonyl reductases, with the greatest degree of induction occurring in the ce11 line resistant to daunorubicin (Figure 2) (Soldan et al., 1996). Carbonyl reductases catalyze the reduction of daunorubicin to daunorubicinol, which has significantly lower antineoplastic potency. The induction of carbonyl reductases may be a contributing factor to daunorubicin resistance.

Another exarnple involves the drug cyclophospharnide which requires activation by liver cytochrome P450 enzymes for antitumour activity. The therapeutic effectiveness of the drug, bowever, is limited by the hematopoietic, renal and cardiac toxicity that accompanies systernic distribution of liver-derived activated drug metabolites. To limit this toxicity and to increase the therapeutic effïciency, it would be ideal to administer lower doses of the drug and/or to enhance its bioactivation at the site of action in the tumour cells.

Cyclophophosarnide has been shown to be bioactivated by rat CYP2B1 (Clarke and Waxman,

1989). In in vitro studies, transfection of the CYP2B1 cDNA into a panel of human tumour ce11 lines, including a breast cancer ce11 Iine (MCF-7),resuIted in increased sensitization of the cells to cyclophospharnide, compared to the corresponding parental ce11 line (Chen and

Waxman, 1995b; Chen et al., 1996). Growth of MCF-7 tumours overexpressing CYP2B1 in rnice administered cyclophosphamide produced significant tumour growth delay, without any apparent increase in host toxicity. Transfection of the CYP2B1 cDNA into the breast tumour cells increased the capacity for inkatumoral activation of the drug which had the beneficial effect of concentrating the active drug at the site of action, while at the same tirne decreasing systemic exposure to reactive metabolites. The intrinsic inability of the tumour cells to bioactivate the drug limited its therapeutic efficiency.

Therefore, the presence of drug metabolizing enzymes in tumour tissue is a significant factor in tumour response to the drug (Guengerich, 1988; LeBlanc and Waxman, 1989;

Graham et al. , 1991). Knowledge of which drug biotransformation enzymes occur in normal and tumour breast tissue may provide an indication of the tumour response to a drug.

Modulation of expression of drug biotransformation enzymes in tumour tissue would be a means of increasing the effectiveness of a drug. However, to date limited research fias been conducted on the expression of drug metabolizing enzymes in various types of tumour tissue. 13 Also, the enzymes that bioactivate or detoxia antineoplastic drugs particularly at sites other than the liver are not well characterized. The focus of this thesis is to evaluate the biotransformation of DOX for the purposes of increasing its therapeutic efficiency in breast tumour cells. As such, the rnetabolism of DOX and how this might relate to its free radical and redox-cycling mechanism of action will be discussed.

1.4 Doxorubicin Biotransforrnation and Mechanism of Free Radical Production

1.4.1 Doxorubicin metabolism

The major pathway of DOX biotransformation for the purposes of inactivating and elirninating the drug is ,stereospecific reduction of the ketone on Carbon 13 of the molecule to produce doxorubicinol, a compound which still retains some cytotoxic activity (Figure 2).

This reaction is catalyzed by the cytoplasmic aldoketoreductases (including carbonyl reductase) which occur in most ce11 types but particularly in red blood cells, liver and kidney cells (Loveless et al., 1978; Ballet et al., 1986; Speth et al., 1988; LeBot et al., 1991;

Robert and Gianni, 1993). There is some indication that doxorubicinol further undergoes phase II conjugation reactions (sulfation and glucoronidation) prior to being excreted

(Takanashi and Bachur, 1976; Powis, 1987). However, this is uncertain and probably represents a minor pathway of metabolism in humans (Powis, 1987). O -qar Doxorubicin aldoketoreductases NAD(P)H-quinone oxidoreductase \ (DT-diaphorase)

O -sugar IIADPH-Cytochrome O supr Doxorubicinol '450 Reductase Doxorubicin Dihydroquinone

Doxoru bicin 7-Deoxyaglycone Figure 2: Biotransfomation of Doxonibicin to Less Toxic Metabolites 15 A minor pathway of DOX metabolism is reductive deglycosylation of the sugar group to produce DOX 7-deoxyaglycones which have no cytotoxic activity. However, this pathway cm also be considered a bioactivation pathway, since some reactive intermediates are produced in this pathway (see Section 1.4.2). DOX aglycones are present in the biological fluids of only some patients treated with DOX (Curnmings et al., 1986; Curnmings &

McArdle, 1986). They occur only transiently and at very low concentrations compared with the parent drug and its major metabolite, doxorubicinol.

Another possible detoxification pathway of DOX involves two-electron reduction of the quinone moiety to a DOX hydroquinone as catalyzed by NAD(P)H-quinone oxidoreductase, which is a cytosolic flavoprotein also known as DT-diaphorase (Figure 2).

Whether ant&acyclines, including DOX, are substrates for NAD(P)H-quinone oxidoreductase is uncertain with some studies suggesting they are (Fisher et al., 1992; Kasahara et al.,

1994), whereas other studies indicate that they possibly are not (Wallin, 1986; Cummings et al., 1992b; Wakusawa et al., 1997). In clinical studies, the DOX hydroquinone metabolite was not reported to occur in biological fluids (Powis, 1987); however, this may sirnply be due to the fact that detection of this metabolite was not specifically conducted at the time.

1.4.2 Enzymatic free radical production

DOX can also be biotransfonned enzymatically to a reactive interrnediate. As shown in Figure 3, the quinone moiety of DOX is reduced in a one-electron step to a semi-quinone radical (Svingen and Powis, 1981; Bachur et al., 1982; Doroshow, îY83a). A number of flavoproteins have been reported to catalyze this reaction by accepting an electron from

NADPH or NADH and donating it to DOX. Under aerobic conditions, the semi-quinone

17 radical is highly unstable and will quickly donate its unpaired electron to oxygen (O,)

fomiing superoxide (O;.). By reducing oxygen to superoxide the parental DOX is regenerated. This sequence of reactions is known as redox-cycling and can continue

indefinitely. The toxicity of DOX via redox-cycling is thought to occur from the subsequent generation of the hydroxyl radical (OH*), which is the most reactive and damaging of the oxygen free radicals and is known to cause tissue darnaging effects such as lipid peroxidation, and DNA and protein darnage (Cheeseman and Slater, 1993; Kehrer, 1993;

Stohs, 1995).

The proposed sequence of reactions that would result in the formation of hydroxyl radicals is as follows. Superoxide dismutates to hydrogen peroxide (E40,)and oxygen.

20,. + 2H+ + %O2 + O, [equation 11

This can occur spontaneously (albeit rather slowIy) or it is catalyzed by the enzyme superoxide dismutase which occurs in two main fonns in mammalian cells: the copper-zinc forrn in the cytoplasm and the manganese forrn in mitochondria. Superoxide can then react with hydrogen peroxide to produce the hydroxyl radical in a reaction known as the Haber-

Weiss Reaction.

H202+ O,-* * OH* + OH-+ O2 [equation 21

This spontaneous reaction would be unlikely to occur under normal physiological conditions due to the low steady-state concentrations of the reactants (Cheeseman and SIater, 1993).

The reaction does occur significantly, however, in the presence of transition metai ions such as iron and copper and is referred to as the iron-catalyzed Haber-Weiss Reaction (equations 3 and 4). 02-• + Fe3+ + O2 + Fe2+ [equation 31

H202 + Fe2+ + OH. + OH' + Fe3+ [equation 41

The end product is the formation of the hydroxyl radical.

The ce11 has some defences against these reactive oxygen species (Figure 3). As described above superoxide dismutase removes superoxide. This is the only known enzyme whose substrate is a free radical (Cheeseman and Slater, 1993). In addition, catalase and

GSHPx are enzymes that detoxify peroxides. Catalase occurs primarily in peroxisomes and mediates the following reaction.

2H202 + 2H20 + O, [equation 51

GSHPx occurs in the cytoplasm of most cells and will react with both hydrogen peroxide and fatty acid hydroperoxides, if the hydroperoxides are first cleaved frorn membrane phospholipids by a phospholipase (Wendel, 1980). The reaction catalyzed by GSHPx is represented in the following equations, where GSH represents reduced glutathione and GSSG represents oxidized glutathione.

H202 + 2GSH + 2H,O + GSSG [equation 61

ROOH + 2GSH 4 ROH + H,O + GSSG [equation 71

Other compounds present in the ceIl act as 'scavengers' or interceptors of free radicals. In ce11 membranes, this includes a-tocopherol which by reacting with lipid peroxyl radicals will terminate lipid peroxidation chain reactions. Other scavengers include ascorbic acid (vitamin C) in the aqueous phase of cells and plasma, uric acid in plasma and glutathione in the cytoplasm (Cheeseman and Slater, 1993).

Redox-cycling is rapid in the presence of oxygen but is slow under hypoxic conditions

(Keizer et al., 1990). In the absence of oxygen, the DOX semi-quinone free radical accumulates and rnay potentially bind covalently to nucleic acids and proteins. However, whether this actually occurs in cells is uncertain. The DOX semi-quinone is quite unstable with an estirnated diffusion radius of 0.6 or O. 1 pm under anaerobic and aerobic conditions, respectively (Svingen and Powis, 1981). Since the average ce11 diameter is 5 to 20 Pm, oniy a small proportion of the semi-quinone radicals produced in the endoplasmic reticulum or cytoplasm would be expected to reach the nucleus (Keizer et al., 1990). However, reactive metabolites of the semi-quinone rnay be formed which rnay interact with DNA and proteins.

Moore (1977) and Sinha and Gregory (1981) have proposed that the semi-quinone undergoes further biotransforrnation to active or inactive species. As shown in Figure 4, the semi- quinone free radical rnay rearrange chemically by eliminating the daunosamine sugar group

(reductive deglycosylation) to produce a series of reactive aglycone intermediates which are proposed to bind to DNA covalently. Two major pathways of semi-quinone rearrangement have been proposed (Moore, 1977; Sinha and Gregory, 1981). Firstly , the semi-quinone rnay degrade directly to the 7-deoxyaglycone metabolite via a C7-centred aglycone radical.

Although, the 7-deoxyaglycone metabolite is inactive (Henry, 1979; Averbuch et al., l985), the C7-centred aglycone radical has been reported to have the potential to bind to DNA

(Sinha and Gregory, 1981). However, other investigators believe that the C7 radical is not adequately reactive to alkylate DNA (Abdella and Fisher, 1985). In the second pathway, anaerobic bioreductive deglycosylation rnay occur via disproportionation of the semi-quinone Doxorubicin 9 OH

Drug-macromolecule

C7 Quinone Methide C7 Fret Radical (Tautornert A)

Figure 4: Formation, Metabolic Fate, and Potentiai Toxicity of the Doxorubicin Semi-quinone free radical after one electron reduction, resulting in the formation of C7-quinone methide.

The C7-quinone methide occurs as two tautomers (a substance that exists as two mutually convertible isomers in equilibrium). One tautomer of the C7-quinone methide (tautomer B in

Figure 4), is reported to have the potential to alkylate DNA (Moore, 1977). Moreover, it has a half-life of several seconds which is long enough for diffusion through the ceIl and alkylation of DNA (Kleyer and Koch, 1983). But whether it is an alkylating agent is debatable, since other investigators have reported that the C7-quinone methide has Iimited capability to react with sulfydryl groups of proteins and is probably not sufficiently reactive to bind to DNA (Ramakrishan and Fisher, 1986). Rather, it preferentially abstracts a solvent proton to fonn the 7-deoxyaglycone metabolite (Rarnaskrishan and Fisher, 1986). Gaudiano et al. (1994) reported that the C7-quinone methide can be conjugated to reduced glutathione

(GSH) forming 7-S-glutathionyl-7-deoxyaglycone. However, whether this occurs in whole cells or in the presence of ce11 constituents is not known. These DOX-GSH conjugates were less toxic to MCF-7/WT cells and reportedly they prevented DOX access to the nuclei by being predominantly located in the cytoplasm and Golgi region (Serafino et al., 1998).

Toxicity of the DOX-GSH conjugates to MCF-7 cells resistant to DOX (MCF-7lADR) did not change suggesting that resistance to DOX may be due to an increased ability of the cells to form DOX-GSH conjugates. However, as stated before, the formation of the DOX-GSH conjugates in cells or with ce11 constituents has not been shown to occur.

'Fherefore, it has been proposed that under hypoxic conditions, the toxicity of DOX may be a result of the DOX semi-quinone radical and some of its aglycone metabolites (the

C7-free radical and the C7-quinone methide) which may interact with DNA. On the other hand, it has also been suggested that formation of the semi-quinone and its aglycone intermediate metabolites represents deactivation of DOX and is not a significant pathway of toxicity (Cummings et al., 1992a). Studies evaluating DOX-induced toxicity via its metabolism to reactive intermediates and redox-cycling are discussed in Section 1.5.

1.4.3 Other mechanisms of free radical production

A number of other possible mechanisms of free radical production from DOX has been proposed. For example, at low partial pressures of oxygen, DOX semi-quinone was reported to react with hydrogen peroxide resulting in subsequent breakdown of deoxyribose

(Bates and Winterbourn, 1982). The authors reported that a sirnilar reaction with deoxyribose in DNA may be responsible for the DNA strand scission observed in the presence of DOX. The reaction was catalyzed by submicromolar concentrations of iron and occurred at low oxygen tensions, conditions which may occur in tumour cells.

Another mechanism of DOX-induced free radical production involves the DOX-iron complex (Keizer et al. , 1990; Cummings et al. , 19% b). Iron (Fe3+)complexes to DOX between the oxygen atoms of Cl1 and Cl2 (see Figure 1). The ferric iron (Fe3+)is then reduced to ferrous iron (Fe2+)enzymatically by NADPH-cytochrome P450 reductase, nonenzymatically by glutathione or by intramolecular autooxidation of the DOX B ring hydroquinone. DOX-Fe2+can then react with oxygen to form superoxide which may then dismutate to form hydrogen peroxide; or the complex can react with hydrogen peroxide to produce hydroxyl radical. For both reactions DOX-Fe3+ is regenerated. This system is comparable to redox-cycling of DOX in that no DOX metabolites are formed and the reaction can proceed indefinitely. In the absence of a reducing system, autooxidation of the DOX-iron complex can also produce hydrogen peroxide and superoxide, with the final metabolite being dehydroxyacetyl-9-carboxyl doxorubicin (9-COOH-DOX) .

In in vitro studies, the DOX-iron complex has been shown to bind tightly to DNA and also to catalyze the formation of oxygen free radicals in the presence of double-stranded

DNA (Eliot et al. , 1984; Muindi et al., 1984; 1985). This is quite an interesting finding since DOX alone bound to double-stranded DNA does not undergo one electron reduction or mediate free radical production either enzymatically or chemically (Sato et al., 1977;

Kalyararaman et al. , 1980; Berlin and Haseltine, 1981; Rowley and Halliwell, 1983;

Youngrnan et al., 1984). The DOX-iron complex has also been shown to damage membranes and cause lipid peroxidation in the presence or absence of a reducing system

(Myers et al. , 1982; Sugioka and Nakano, 1982; Gutteridge, 1983; 1984).

It is uncertain whether the proposed DOX-iron complex mechanism of action is a significant mechanism of DOX toxicity in whole cells or in vivo. There are several reasons why it may not be a significant mechanism (Gelvan and Samuni, 1988; Keizer et al., 1990;

Cummings et al. , 1991b). Firstly , the 9-COOH-DOX metabolite has not been detected in patients treated with DOX. Secondly, the occurrence of DOX-iron complexes has not been demonstrated in intact cells or clinical specimens. Finally, the binding of iron to DOX is pH dependent and at the expected DOX concentrations at physiological pH, binding of DOX to iron is not expected to occur (Gelvan and Sarnuni, 1988). 1.5 Evidence of Toxicity from DOX-Induced Free Radicals and Redox-Cycling

1.5.1 DNA damage

In a number of studies, DNA damage or deoxyribose breakdown by redox-cycling of

DOX was reported to occur (Lown et al. , 1977; Berlin and Haseltine, 1981; Gutteridge and

Toeg, 1982; Potmesil et al. , 1983; 1984; Rowley and HalIiwell, 1983; Youngman et al. ,

1984; Feinstein et al., 1993). These were in vitro studies conducted with DNA or nuclear fractions or whole cells from which DNA was later isolated. The DNA damage was mediated by oxygen and usually occurred in the presence of an activating system (for exarnple, purified rat NADPH-cytochrome P450 reductase). In some studies, the DNA damage was inhibited by superoxide dismutase, catalase, iron chelators and hydroxyI radical scavengers, which suggests that the DNA darnage was mediated via hydroxyl radicaIs. The most cornrnon type of DNA damage reported was Strand scission. Concentrations of DOX used were 2.3 to 400 PM.

Low level covalent binding of DOX to DNA in the presence of chernical reducing agents, microsomes, isolated nuclei or cells in culture was also reported to occur (Cummings et al., 1991a). However, the relevance of these studies has been questioned, since high concentrations of DOX (50 PM to 1 rnM) were required to produce detectable resuIts and distinct, individual adducts could not be identified (Curnmings et al., 1992a).

1.5.2 Lipid peroxidation

In the presence of a metal catalyst, unsaturated fatty acids undergo lipid peroxidation when exposed to hydroxyl radicals (Gutteridge, 1982). Numerous in vitro studies have been conducted in which DOX-mediated lipid peroxidation occurred after metabolic activation by heart, liver, kidney or tumour ce11 microsomes, mitochondria or nuclei (Goodman and

Hochstein, 1977; Mimnaugh et al., 1981; 1982; 1983; 1985a; 1985b; Muliawan et al., 1982;

Sterrenberg et al., 1984; Goeptar et al., 1993). Hydroxyl radicals generated by the iron- catalyzed Haber-Weiss reaction appears to be the main mechanism since in the presence of oxygen, the lipid peroxidation was inhibited by superoxide dismutase, catalase, glutathione, iron chelators and hydroxyl radical scavengers. The concentrations of DOX used in these studies ranged from 10 pM to 200 PM.

Lipid peroxidation was also evaluated in vivo in rats with a subcutaneously growing rat mammary carcinoma (Sp 107) (Curnmings et al., 1992a). Following intratumoural injection of 70 pg DOX/animal, there was no evidence of increased lipid peroxidation in liver or tumour tissue. In other animal studies, free radical scavengers were adrninistered with DOX to determine whether a reduction in cardiotoxicity would occur. The cardiotoxicity of DOX is thought to be due to redox-cycling of DOX with its associated lipid peroxidation (see section 1.5.4). Administration of the scavengers (ascorbate, N- acetylcysteine, cysteine, glutathione and a-tocopherol) resulted in decreased cardiotoxicity in the animals (Myers et al. , 1977; Freeman et al. , 1980; Doroshow et al. , 1981; Fujita et al. ,

1982; Yoda et al., 1986). However, whether this was due to decreased lipid peroxidation is uncertain. Interestingly, while reducing cardiotoxicity, this treatment did not reduce the turnouricidal effect of DOX.

1.5.3 CeII toxicity

A major drawback of some of the above in vitro studies is that they were conducted with subcellular fractions and not related to ce11 toxicity. Several studies have been conducted which evahate the role of DOX redox-cycling in relation to cell toxicity and

resistance. Comrnon parameters evaluated were the production of hydroxyl radical and the

effect of free radical scavengers on ce11 toxicity.

In MCF-71WT cells exposed to 20-30 pM DOX, the hydroxyl radical was detected,

whereas it was not detected in cells resistant to DOX (MCF-7/ADR), even at high

concentrations of DOX (330 PM) (Sinha et al., 1987a,b). NADPH added extracellularly to

both ce11 Iines, strongly stirnulated hydroxyl radical production. Superoxide dismutase,

catalase and iron chelators inhibited hydroxyl radicaI formation. However, the doses of

DOX used to detect the hydroxyl radical in the above experiments were quite high cornpared

to the IC,, values of the ceIl lines (0.025 pM and 4.8 pM for MCF-71WT and MCF-7/ADR,

respectively). Formation of hydroxyl radicals may have been occurring at lower

concentrations, but nlay not have been detected due to the detection method used. Hydroxyl

radicals are highly reactive and they may have reacted with cellular constituents too rapidly

to be detected by the relatively insensitive method of electron spin resonance (ESR)spin

trapping (Keizer et al., 1990; Sinha and Mimnaugh, 1990). Therefore, the effect of free

radical scavengers on cellular toxicity is an alternate approach to evaluate redox-cycfing

toxicity. For both ce11 Iines, addition of superoxide dismutase and catalase increased ce11

survival at DOX concentrations of 0.01 and 1 pM for MCF-7/WT and MCF-7lADR cells,

respectively. Since neither NADPH, superoxide dismutase or catalase can cross the ce11

membrane, these data suggest that hydroxyl radicak were forrned outside the cell, perhaps

catalyzed by reductive enzymes in the plasma membrane. The much lower hydroxyl radical production in MCF-7lADR cells appeared to be due to increased expression of GSHPx, 27 which reacts with hydrogen peroxide and would therefore limit the production of hydroxyl

radical. In boîh ce11 lines, there was no difference in the activities of NADPH-cytochrome

P45O reductase, NADH-cytochrome b, reductase, xanthine oxidase, superoxide dismutase and catalase. In a subsequent study by the sarne investigators, hydroxyl radical formation was decreased by the addition of purified GSHPx to ce11 lysate from MCF-7/WT cells (Sinha et al., 1989).

Therefore, the studies conducted by Sinha et al. (1987a; 1987b; 1989) suggest that the cytotoxicity of DOX in MCF-7IWT and MCF-7IADR cells may be due to the formation of hydroxyl radicals, and that a possible mechanism of resistance in MCF-7IADR cells is increased expression of GSHPx, which reduces the formation of liydroxyl radicais. A number of other studies support these findings. The cytotoxicity of 0.5 pM DOX to

MCF-7IWT cells was greatly reduced in the presence of superoxide dismutase, catalase, iron chelators penneable to the plasma membrane, the hydroxyl radical scavenger N- acetylcysteine, and the organo-selenium compound 2-phenyl- 1,2-benzisoselenazol-3(2H)-one, which possesses GSHPx-like activity (Doroshow, l986a). Cytotoxicity of 1.75 pM DOX to

MCF-7/WT cells also was decreased when GSHPx and superoxide dismutase were introduced into cells by a "scrape loading" method (Doroshow et al., 1991). Increased activity of these enzymes in the cells was confirmed and correlated with the degree of DOX resistance acquired. Sensitivity to DOX increased in MCF-7/WT and MCF-7/ADR cells exposed to exogenously produced superoxide and hydrogen peroxide (Mimnaugh et al.,

1989). The effect was both concentration and tirne-dependent. The MCF-7/ADR cells which were 500-fold more resistant to DOX,tolerated the superoxide and hydrogen peroxide approxirnately 4-fold more than the MCF-71WT cells. As in the studies by Sinha and colleagues, the MCF-7IADR cells were reported to have greatly increased GSHPx activity and a slight increase in superoxide dismutase activity. The concentrations of DOX used in these studies were 10 to 20 nM and 5 to 10 pM in MCF-7lWT and MCF-7lADR cells, respectively. DOX toxicity to MCF-7IADR cells in the presence of buthionine sulfoximine

(BSO), which reduces cellular glutathione (GSH) levels by inhibiting of y-glutamylcysteine synthetase, was also investigated (Dusre et al., 1989). Minirnally cytotoxic doses of BSO reduced GSH levels by 80 to 90% and also increased ce11 sensitivity measured by two separate assays with a dose modifying factor of 5 to 7. (A dose-modifying factor is the ratio of the IC,, without BSO to the IC5, with BSO.) This increase in sensitivity was apparent at

DOX concentrations of 2.5 to 5 PM. BSO treatrnent also resulted in 2-fold greater production of hydroxyl radical by the whole cells. As in the studies by Sinha and colleagues, high concentrations of DOX (200 pM) were required to detect hydroxyl radicals by ESR spin-trapping .

Results from another study conducted in Ehrlich turnour ceIls also agree with the results of Sinha and colleagues (Doroshow, 1986b). DOX (1.5 PM) cytoxicity was greatly reduced in the presence of superoxide dismutase, catalase, various hydroxyl radical scavengers and various iron chelators. Cytoxicity was not produced by an analog of DOX,

5-iminodaunorubicin, which has a modified quinone structure that prevents redox-cycling. In addition, the hydroxyl radical was detected in intact ceIls in the presence of 250 pM DOX, hydrogen peroxide and iron. The hydroxyl radical was not detected with

5-irninodaunorubicin. In Chinese hamster ovary cells, depletion of GSHPx by transfection of 29 a vector that resulted in production of antisense GSHPx mRNA, resulted in an approximate

2-fold decrease in resistance to DOX (Taylor et al., 1993).

The results of some studies, however, contradict the findings of Sinha and colleagues.

Transfection of GSHPx cDNA into MCF-7IWT cells resulting in 40-fold elevation of activity did not produce resistance to DOX (Liebman et al., 1995). In another study, using spin- trapping and ESR spectroscopy, the formation of the DOX semi-quinone and the hydroxyl radical was detected under anaerobic and aerobic conditions, respectively, using MCF-7/WT and MCF-7/ADR cells and a DOX concentration of 1 rnM (Alegria et al., 1989). Although superoxide dismutase elirninated the hydroxy1 radical signal, catalase had no effect , which according to the authors suggested that the hydroxyl radical was fonned from the superoxide and not from hydrogen peroxide. An iron chelator (not permeable to the plasma membrane) had no effect on hydroxyl radical and semi-quinone formation. Also, although the MCF-

7lADR cells were approximately 500-fold more resistant to DOX, comparable levels of the radicals were generated in both ce11 lines. MCF-7/WT cells had higher levels of glutathione and the authors cited other studies indicating that MCF-7lADR cells had higher levels of

GSTs and GSHPx. Also, in the same study, similar results were obtained with DOX-sensitive and DOX-resistant Chinese hamster ovary ceIls. Alegria et al. (1989) concluded that oxygen derived radicals do not play a role in DOX antitumour activity. Cytotoxicity in the presence of superoxide dismutase, catalase and the iron chelators was not evaluated.

Other studies which contradict the findings of Sinha and colleagues have been conducted in different ce11 lines and may reflect biochemical differences in the ce11 lines

(Cumrnings et al., 1991b). In one study by Keizer et al. (1988), Chinese hamster ovary 30 cells (CHO? were selected by long-term adaptation to step-by-step increased oxygen tensions

(up to 99 % 0,/1%CO,), resulting in these cells expressing higher levels of glutathione, superoxide dismutase, catalase and GSHPx compared to the wild-type cells (CHOS).The sensitivity of both ce11 lines to DOX (O to 20 PM) was then assessed. Based on the extracellular concentration of DOX, CHOr cells were more sensitive to DOX; however they also had more cellular uptake of DOX. Sensitivity of both ce11 lines was considered to be essentially equal based on the intracellular concentration of DOX. Therefore, despite the increased levels of antioxidant defenses in CHOr cells, sensitivity to DOX was not changed.

In sensitive and multidrug resistant human ovarian cancer cells, cytotoxicity to DOX was decreased in the presence of hydroxyI radical scavengers (N-acetylcysteine, sodium benzoate and dimethyl sulfoxide) at DOX concentrations of 0.2 to 0.5 pM and 20 to 50 pM for sensitive and resistant cells, respectively (Cervantes et al. , 1988). The degree of protection by the hydroxyl radical scavengers was similar in both sensitive and resistant cells. However, cytotoxicity was not affected by the presence of ascorbic acid (a superoxide scavenger), superoxide dismutase, and catalase. Introduction of superoxide dismutase and catalase into the cells by the "scrape-loading" method also did not affect the cytotoxicity. The authors concluded that hydroxyl radicals were involved in the cytotoxicity of DOX, as opposed to superoxide or hydrogen peroxide.

An alternate method of evaluating DOX-mediated toxicity via redox-cycling is to inhibit reductive metabolism of DOX. This was achieved by using artificial electron acceptors which would compete with DOX for electrons frorn the reduced flavoproteins

(Keizer et al. , 1989). These artificial electron acceptors were phenazine methosulfate, 31 menadione and methylene blue. The cytotoxicity to SW- 1573 human lung tumour cells with the electron acceptors at nontoxic concentrations did not change with 0.5 to 4 pM DOX, although it decreased for another drug, whose toxicity may also be mediated by flavoprotein-catalyzed metabolism. These results suggest that fiavoprotein-catalyzed metabolism was not required for DOX induced cytotoxicity.

The role of NADPH-cytochrome P450 reductase (CYPFED) in the cytotoxicity of

DOX to MCF-7/WT cells was assessed by Bartoszek and Wolf (1992). In the presence of purified rat CYPRED, NADPH and DOX at concentrations greater than 5 PM, cytotoxicity was increased. This was dependent on the concentration of both DOX and CYPRED.

Increased cytotoxicity was not observed when CYPIUED, NADPH and DOX were incubated for a period of tirne before exposure to the cells, implying that a short-lived metabolite was responsible for the cytotoxicity. To determine whether redox-cycling was involved, cytotoxicity assays were conducted in the presence of glutathione, superoxide dismutase, a-tocopherol (which inhibits formation of lipid hydroperoxides), mamit01 and DMPO (53- dimethyl-1-pyrrolidine-N-oxide) which are hydroxyl radical scavengers. With the exception of glutathione, none of these free radical scavengers reduced the cytotoxicity of DOX. In addition, increased cytoxicity was not associated with lipid peroxidation. These data suggest that oxygen radicals did not play a primary role in CYPRED-mediated DOX toxicity in the

MCF-71WT cells. However, at DOX concentrations of 10 PM and above, CYPRED- dependent binding of radio-labelled DOX to cellular proteins was detected. And at a concentration of 50 pM DOX, CYPRED-dependent covalent binding of DOX to DNA was detected. Therefore, these resuits indicate that exogenous CYPRED was associated with 32 increased cytotoxicity. The cytotoxicity was Iikely not due to reactive oxygen species, but to

a reactive metabolite which binds to DNA and proteiris.

In summary, studies assessing the role of redox-cycIing in DOX-induced cytotoxicity

in whole cells are contradictory. In MCF-7breast cancer ce11 lines and Ehrlich tumour cells, redox-cycling of DOX does appear to be a mechanism of cytotoxicity and increased expression of GSHPx may be a rnechanism of resistance to DOX (Doroshow, 1986a; 1986b;

Sinha et al., 1987a,b; 1989; Dusre et al. , 1989; Mimnaugh et al., 1989; Doroshow et al. ,

1991). However, CYPRED-dependent cytotoxicity of DOX in MCF-7 cells did not appear to be mediated by redox-cycling and was perhaps due to the presence of an unidentified reactive metabolite (Bartoszek and Wolf, 1992). In other ce11 lines (Chinese hamster ovary cells, human ovarian cancer cells and human lung tumour cells), redox-cycling of DOX did not appear to mediate its cytotoxicity, nor were changes in antioxidant defense enzyme expression associated with DOX resistance (Cervantes et al., 1988; Keizer et al., 1988;

Alegria et al., 1989; Keizer et al., 1989). This may be due to sorne biochemical differences between these ce11 lines and the MCF-7 ce11 line (Keizer et al., 1990; Cummings et al.,

1991b).

1S.4 Cardiotoxicity

A major limiting side effect of DOX treatment is a potentially irreversible cumulative dose-related cardiomyopathy that manifests itself as congestive heart faiIure (Speth et al.,

1988; Overrnoyer, 1995; Shan et ai., 1996). The risk of congestive heart failure is 0.1 to

1.2 % up to a maximal cumulative dose of 550 mg/m2. During adjuvant chemotherapy the total dose received is usually 240 to 400 mg/m2. In addition to this chronic cardiomyopathy 33 which is usually apparent within a year of treatment, treatment with anthracyclines has also

been associated with late-onset cardiotoxicity (ventricular dysfunction and arrhythrnias) which

manifests years to decades after treatment has been completed. Although the data are

limited, this appears to occur at doses commonly used for cancer treatment in approxirnateIy

5 to 7 % of treated patients (Shan et al. , 1996).

The selective toxicity of DOX to heart cells is generally considered to be the result of free radical darnage (Smith et al., 1985; Keizer et al., 1990; Shan et al., 1996). The heart may be particularly susceptible due to a number of reasons. Firstly, heart muscle cells are rich in mitochondria where DOX is reduced to the serni-quinone via catalysis by NADH dehydrogenase. Unlike in the liver or tumour tissue, it appears that the semi-quinone reacts with hydrogen peroxide to directly produce hydroxyl radicals, even in the presence of oxygen

(Nohl and Jordan, 1983). Therefore, site-specific and more efficient production of the hydroxyl radical occurs in a major organelle of the heart muscle cell. This has also been shown to occur in the sarcoplasmic reticulum (Ferrans, 1978). Microscopic examination of heart tissue shows that DOX-induced cardiac damage starts at the mitochondrion and sarcoplasmic reticulum (Olson and Capen, 1977). Secondly, the heart has been reported to have relatively low levels of antioxidant defenses which rnay result in DOX-induced lipid peroxidation. For example, low levels of superoxide dismutase and catalase, and a low rate of glutathione turnover have been reported (Griffith and Meister, 1979; Doroshow et al.,

1980). In addition, DOX treatment tends to deplete GSHPx levels (Doroshow et al., 1980).

In in vivo animal studies, CO-administrationof DOX with free radical scavengers reduced cardiotoxicity without significantly affecting tumour response (Myers et al., 1977; Freeman et al., 1980; Doroshow et al. , 1981; Fujita et al. , 1982; Yoda et al. , 1986). Thus the

oxidative stress produced by DOX in the heart may be far greater than in other better-

protected tissues, such as liver and kidney, thereby explaining its selective toxicity. DOX

has also been shown to affect calcium influx across the sarcoplasrnic reticulum by activating

the calcium release charnel (Holmberg and Williams, 1990; Kusuoka et al. , 1991). Free

radical-induced ce11 membrane damage has been observed with altered calcium influx,

suggesting that these two cardiotoxic pathways may be linked (Burton et al., 1990; Holmberg

et al. , 1991).

In the United States, an iron chelator, dexrazoxane (ICRF-187), has been approved

for use in women with metastatic breast cancer adrninistered a cumulative anthracycline dose

of 350 mg/m2 (Seifert et al., 1994; Shan et al., 1996). Bowever, studies indicate that

dexrazoxane may interfere with the antitumour efficacy of the anthracyclines (Sehested et al.,

1993; Seifert et al., 1994).

Significant cardiotoxicity associated with the free radical rnechanisrn of DOX,

indicates that this mechanism has the potential to cause significant cellular damage, given the right biochemical conditions.

1.5.5 Toxic effects of DOX semi-quinone and other DOX radicals

A few studies have been conducted in which the covalent binding of DOX serni- quinone and its reactive metabolites (ie., the DOX C7 free radical and the DOX C7-quinone methide) to cellular macromolecules was evaluated. Anaerobic incubation of rat liver microsomes and DNA with DOX (1700 PM) was reported to produce covalent binding of

DOX to DNA (Sinha and Gregory, 1981). This reaction was inhibited by glutathione, which 35 does not react directiy with the semi-quinone. The authors concluded that a metabolite of the semi-quinone was formed that bound to the DNA and which also may react with glutathione.

They reported that this active species was the DOX C7 free radical. Sinha and colleagues have shown in a nurnber of other similar studies that under anaerobic conditions and in the presence of an activating system such as liver microsornes or nuclei (which contain reductases), DOX was bound to DNA (Sinha and Chignell, 1979; Sinha, 1980; Sinha et al.,

1984). This reaction was NADPH-dependent, inhibited by glutathione and not inhibited by dicoumarol (an inhibitor of DT-diaphorase). Under aerobic conditions, binding of DOX to

DNA was not detected. Although not specifically identified, Sinha and colleagues proposed that the reactive metabolites are the DOX semi-quinone, DOX C7 free radical and DOX

C7-quinone methide.

The binding of DOX metabolites to DNA in the presence of rat liver microsomes was shown in another study by Wallace and Johnson (1987). However, in contrast to the studies by Sinha and colleagues, the degree of DNA binding was much greater in the presence of oxygen than under anaerobic conditions. The concentration of DOX used in these studies was 1.0 mM.

Binding of DOX metabolites to proteins has also been proposed and shown to occur under aerobic conditions in the presence of rat liver and heart microsomes (Ghezzi et al.,

1981; Scheulen et al. , 1982). This was dependent on oxygen, NADPH and microsomal protein concentration. It was inhibited by reduced glutathione and other sulfhydryl compounds. The presence of superoxide dismutase had no effect on the results.

Therefore, these in vitro studies suggest that metabolites of DOX or the DOX semi- quinone can covalently bind to DNA and proteins. WhiIe possible metabolites under anaerobic conditions have been proposed, the possible metabolite binding to DNA and protein under aerobic conditions was not identified. The major limitation of these studies, however, is that they were conducted with isolated DNA or protein, and required high concentrations of DOX. In the absence of a whole ce11 system, it is uncertain whether the extent of DOX metabolite binding to DNA or protein will result in cytotoxicity. One such study was conducted by Bartoszek and Wolf (1992). In the presence of purified rat

CYPRED, NADPH and DOX at concentrations greater than 5 PM, cytotoxicity to MCF-

7lWT cells was increased under aerobic conditions, most likely by a short-lived reactive metabolite. Glutathione inhibited this cytotoxicity. At DOX concentrations of 10 pM

CYPRED-dependent binding of radio-labelled DOX to cellular proteins was detected, and at a concentration of 50 pM DOX, CYPRED-dependent covalent binding of DOX to DNA was detected. The possible DOX metabolite binding to DNA and protein was not identified.

DNA adduct formation from interrnediates of DOX reductive rnetabolism was evaluated in vivo in rats with a subcutaneously growing rat mammary carcinoma (Sp 107)

(Cumrnings et al., 1992a). Following intratumoural injection of 70 pg DOXIanimal, low levels of adducts were formed (approxirnately 1 adduct per 10' nucleotides) with four putative adducts detected. However, DNA adduct formation was not detected when the formation of DOX 7-deoxyaglycones were stimulated 155-fold. Since formation of the deoxyaglycones may resuIt from reductive metabolism of DOX with the production of reactive interrnediates (the semi-quinone, the C-7 free radical or the C7-quinone methide), these results suggest that the formation of anaerobic DOX-derived reactive species was not associated with DNA adduct formation in vivo.

Therefore, these in vitro studies indicate that a DOX metabolite formed in the

presence of an activating system under anaerobic conditions can covalently bind to DNA.

However, this has not been shown to occur in vivo. Additionally, there is evidence of an

unidentified DOX metabolite formed under aerobic conditions which may bind to DNA and proteins in vitro.

1.5.6 The relative importance of free radicals in the mechanism of cytotoxicity of DOX

The relative importance of free radicals in the mechanism of cytotoxicity of DOX is unknown. Obviously, the extent of free radical production, redox-cycling and cytotoxicity is dependent on the biochemical environment of the tumour cell. This includes the presence of free radical defence enzymes and scavengers, the presence and subcellular location of bioactivating enzymes, cellular iron and oxygen content, the subcellular location of free radical formation and the size of the cell.

The inhibition of topoisomerase 11 by DOX is considered to be a significant mechanisrn of cytotoxicity. Many investigators report that this is the main mechanism of action of DOX based primarily on the following reasons (Keizer et al., 1990; Cumrnings et al., 1991b).

1. DOX at clinically relevant concentrations binds preferentially and with high efficiency

to DNA. This includes its intercalation to DNA and its interaction with

topoisomerase II. Protein-associated single-strand and double-strand DNA breaks

have been reported to occur at DOX concentrations of 0.5 to 2.5 pM (Curnrnings et

al., 1991b). 38 2. Free radical associated damage of tumour cells by redox-cycling occurs at much

higher concentrations, which would not occur clinically.

These are both valid arguments. DOX tends to accumulate in tissues. Approximately 10 to

500 tirnes the plasma DOX concentration has been reported to occur in tissue samples from

various carcinomas and liver taken from patients administered DOX 30 minutes before

surgical removaf (Cumrnings and McArdle, 1986; Speth et al., 1988). Based on a plasma

concentration of 0.02 pM (10 pg/L), approximate tissue concentrations of DOX would range

from 0.2 to 10 pM (Speth et al., 1988). The literature indicated that in in vitro studies using

subcellular fractions, DNA damage and lipid peroxidation from DOX-induced redox-cycling

occurred at concentrations ranging from 2.3 to 1000 pM. However, in studies evaluating

redox-associated cytotoxicity in MCF-7 breast cancer cells, DOX at concentrations ranging from 10 nM to 5 pM (MCF-7lWT cells) and 1 to 5 pM (MCF-7IADR cells) were used, which is within the range of expected clinical concentrations. Nonetheless, results fkom the cytotoxicity studies did not consistently indicate that redox-cycling was associated with DOX cytotoxicity.

Data are Iirnited about whether DOX metabolism in hypoxic conditions is associated with cytotoxicity. Although in vitro studies indicate the potential of DOX metabolites to bind to proteins and DNA, this has not been shown in whole cells or in vivo.

However, DOX metabolism and its associated redox-cycling, is a significant mechanism of toxicity in cardiac cells which is also apparent clinically. The biochemical conditions within the heart tissue are ideal for redox-cycling and indicate that this mechanism can cause severe cytotoxicity . 39 Most likely cytotoxicity of DOX in cancer cells is multifactorial, including al1 of

DOX's proposed mechanisms from the ce11 membrane to the nucleus. Although it is unlikely

that DOX metabolism and redox-cycling is the main mechanism of cytotoxicity of cancer

cells, the potential of redox-cycling to cause severe cellular darnage is apparent in cardiac cells. Enhancing a particular mechanism of DOX toxicity in tumour cells, would be an ideal way to increase its efficacy. Manipulating DNA or topoisornerase II to increase DOX response is not feasible at this tirne. However, manipulating the biochemical environment of the tumour ce11 to increase free radical production or the rate of redox-cycling is a distinct possibility. One way this can be achieved is by increasing free radical production through increased metabolism of DOX. As such, the enzymology of free radical production needs to be assessed.

1.6 Enzymology of Free Radical Production from DOX

1A. 1 Evidence for involvement of NADPH-Cytochrome P450 reductase in DOX

reduction

Nurnerous studies have been conducted that demonstrate the involvement of CYPRED in the bioreductive activation of DOX. Generally, three categories of studies have been conducted. These are:

1. studies conducted with whole cells or ce11 constituents, and in which increased

cytoxicity, DNA damage, lipid peroxidation or hydroxyl radical formation was shown

to occur in the presence of purified rat or rabbit CYPRED (Berlin and Haseltine,

1981 ; Komiyama et al., 1982; Cummings et al., 1991a; Bartoszek and Wolf, 1992;

Feinstein et al., 1993); 40 2. enzyme kinetic studies conducted with animal liver microsomes, subcellular fractions

of tumour cells or punfied animal CYPRED;

3. cytotoxiciîy studies in cells with cDNA-expressed CYPRED.

Studies from the first category were discussed in Section 1.5 and will not be discussed further. Studies from the second and third category are discussed below.

1.6.1.1 Enzyme kinetic studies

DOX was shown to be reduced in microsomes from mouse and rat liver, heart, lung and spleen and mouse L1210 and P388 tumours or with purified rat or rabbit CYPRED

(Handa and Sato, 1976; Goodman and Hochstein, 1977; Bachur et al., 1978; 1979; Kharasch and Novak, 1981; 1983; Komiyama et al., 1986; Vile and Winterbourn, 1989; Goeptar et al., 1993). This reduction was dependent on the consumption of NADPH and oxygen and was linear with respect to microsomal protein concentration. ESR spectroscopy showed that a free radical intermediate was produced, consistent with generation of the DOX semi- quinone (Bachur et al., 1978; 1979; Komiyarna et al. , 1986; Goeptar et al. , 1993). With purified rat liver CYPRED, the Kmvalue for DOX-stimulated oxygen consumption was 960 pM and the V, was 2.69 pmol/min/mg protein (Bachur et al., 1979).

1.6.1.2 Transfection Studies

Following transfection of human CYPRED cDNA into Chinese hamster ovary cells resulting in approximately a 10-fold increase in CYPRED activity, increased sensitivity to

DOX (up to 3 Sfold) was reported in one study (Sawamura et al., l996), wliereas there was no change in DOX sensitivity in another study in which human CYPRED was stably expressed in V79 Chinese hamster fibroblasts (Schmalix et al., 1996). 41 Human CYPRED cDNA was also transfected into the human breast cancer ce11 line,

MDA-23 1 (Patterson et al. , 1997); however, ce11 sensitivity to DOX was not evaluated.

1.6.2 Other enzymes involved in DOX reduction

Other enzymes shown to catalyze one-electron reductive bioactivation of DOX and

their subcellular locations are:

xanthine oxidase in cytoplasm (Pan and Bachur, 1980; Doroshow, 1983a; Komiyama

et al., 1986)

NADH dehydrogenase in mitochondria (Davies et al., 1983; Doroshow, l983b)

ferredoxin reductase in mitochondria (Gutteridge and Toeg, 1982; Rowley and

Halliwell, 1983)

NADH-cytochrorne b, reductase in the endoplasmic reticulum at pH 6.6, but not at

pH 7.6 (Hodnick and Sartorelli, 1994)

nitric oxide synthase in macrophages and endothelium (Hodnick and Sartorelli, 1998)

rat cytochrome P450 2B1 (in the endoplasmic reticulum) (Goeptar et al. , 1993).

Therefore, redox-cycling of DOX can occur in the cytoplasm, mitochondrion and endoplasmic reticulum.

In addition, free radical production from DOX has been shown to occur in the presence of nuclei, possibly catalyzed by flavoproteins in the nuclear stroma (Bachur et al.,

1982).

The presence of DOX deoxyaglycones in the biological fluids of some patients, suggests that reductive bioactivation of DOX may also occur clinically (Mross et al., 1988;

Keizer et al., 1990). DOX deoxyaglycones are considered to be metabolites of the semi- quinone fomed under anaerobic conditions.

1.6.3 Other bioreductive antineoplastic agents

A number of other dmgs require reductive bioactivation for their cytocidal effect.

The prototype of a bioreductive alkylating agent is mitomycin C (Sartorelli et al., 1994).

One electron or two electron reduction of the quinone group of mitomycin C produces the semi-quinone or hydroquinone metabolites, which may be further transforrned to compounds containing electrophilic sites that alkylate or cross-link with DNA (Rauth et al., 1993). In the presence of oxygen, the serni-quinone form can undergo redox-cycling similar to DOX.

However, since mitomycin C was shown to be selectively cytotoxic towards celIs cultured under hypoxic conditions, relative to cells grown aerobically, it appears that redox-cycling may not be a significant mechanism of mitomycin C cytotoxicity, with inhibition of the formation of the highly allcylating metabolites (Kennedy et al., 1980a; 1980b; Rauth et al.,

1993; Ross et al. , 1996; Tomasz and Palom, 1997). In recent studies evaïuating the toxicity in mammalian and bacterial ce11 transfectants over-expressing a certain reductase,

NAD(P)H:quinone oxidoreductase (NQO1, DT-diaphorase) and CYPRED have been shown to increase cytotoxicity via enhanced catalysis of the two or one electron reduction of mitomycin C, respectively (Bligh et al., 1990; Belcourt et al., 1996a; 1996b; Mikami et al.,

1996; Sawamura et al., 1996). The exception to this is one study in which cytotoxicity to mitomycin C did not increase in Chinese hamster ovary cells transfected with DT-diaphorase cDNA (Gustafson et al. , 1996).

The selective toxicity of mitomycin C to hypoxic cells has lead to the development of other bioreductive dmgs selective for hypoxic cells, which in vivo cm be quite resistant to 43

ionizing radiation and some chemotherapeutic agents (Ross et al., 1996). These drugs include

newly discovered antibiotics related to mitomycins (KW-2149, BMS-181174, FR66979,

FR900482) (Tomasz and Palom, 1997), and other quinone-containing compounds

(indoloquinone E09, diaziquone, and streptonigrin) (Ross et al., 1996). Heterocyclic di-N-

oxides (eg., tirapazamine, also known as SR4233) are simi1ar to DOX in that one-electron

reduction bioactivates the drug, and two-electron reduction inactivates the drug (Patterson et

al., 1995; 1997). Tirapazamine is currently undergoing Phase II and 111 clinical trials (Evans

et al., 1998). Studies are currently being conducted to identify which reductive enzymes bioactivate these drugs .

1.7 NADPH-Cytochrome P450 Reductase

1.7.1 Function, structure and regulation

NADPH-cytochrome P450 reductase (CYPRED) (NADPH: ferrihemoprotein oxidoreductase, EC 1.6.2.4) along with the cytochromes P450 are components of the microsomal mixed Eunction oxidase system. The structure and function of CYPRED has been assessed in a number of studies in which CYPRED has been cloned, sequenced, expressed and mutated (Backes, 1993; Shen and Kasper, 1993; Strobel et al. , 1995).

CYPRED is located in the endoplasmic reticulum and the nuclear membrane, and its major function is to catalyze electron transfer from NADPH to cytochrome P450 during oxidative metabolism of endogenous and exogenous compounds (Backes, 1993; Strobel et al., 1995).

Specifïcally, during a cytochrome P450-mediated hydroxylation reaction of a substrate

(Figure 5), the reducing cofactor NADPH donates an electron to CYPRED, which in turn reduces the heme iron of the substrate-bound cytochrome P450 enzyme from the ferric (Fe3') I ,Substrate (RH)

NADPH-cytochrorne - Adapted from Parkinson, 1996 P450 reductase lcvtochrornei NADH-Cvtochrome- bs ] b5 reductase I

Figure 5: The Catalytic Cycle for a Cytochrome P45O-mediated Hydroxylation Reaction Following binding of the substrate (RH) to the P450 enzyme (Step A), the heme iron is reduced from the-femc (Fe3+) to the ferrous (Ft?) state-by the addition of a single electron from NADPH-cytochrome P450 reductase (CYPRED)(Step B). Oxygen binds to cytochrome P450 in its ferrous state (Step C) and the Fe2+-O2complex is converted to an Fe2+-OOHcomplex by the addition of a proton (H+) and a second electron, which is derived from CYPRED or NADH-cytochrome b, reductase (Step D). Introduction of a second proton cleaves the Fe2+-OOHcomplex to produce water and an (Fe0)3+ complex (Step E), which transfers one oxygen atom to the substrate (Step F). Release of the oxidized substrate @OH) returns cytochrome P450 to its original state (Step G). to the ferrous (Fe2+)state (step B in Figure 5) (Parkinson, 1996). The hydroxylation

reaction continues with oxygen binding to the ferrous iron forming a Fe2+-0, complex, which

is then converted to a Fe2+-OOH complex by the addition of a proton (H+)and a second

electron (step D in Figure 5). This second electron may be derived from CYPRED or

NADH-cytochrome b, reductase. Therefore, the function of CYPRED is closely associated

with that of the cytochrome P450 enzymes.

In addition to the cytochromes P450, a large number of other electron acceptors from

CWRED have been identified (Backes, 1993; Shen and Kasper, 1993). For example,

CYPRED reduces other microsomal proteins such as cytochrome b,, heme oxygenase and fatty acid elongase, as well as nonphysiologic acceptors such as cytochrorne c and ferricyanide. CYPRED has also been reported to be involved in nitroreduction and azoreduction of xenobiotics, and in one eIectron reduction of a number of quinones (O'Brien,

1991; Backes, 1993). Additionaily , CYPRED may be involved in microsomal lipid peroxidation in conjunction with EDTA-Fe2+, cytochromes P450 and active oxygen (Sevanian et al., 1990).

CYPRED is a 77 kDa protein containing flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD). It is composed of at least five functional domains: an amino- terminal membrane binding domain, binding domains for FMN, FAD and NADPW, and substrate binding domains (see Figure 6). The electron flow through the reductase follows the pathway from NADPH to FAD to FMN to electron acceptors (Strobel et al., 1995; Wang et al., 1997). The arnino acid sequence of CYPRED in humans (Haniu et al., 1989;

Yamano et al., 1989; Shephard et al. , l992), rats (Porter and Kasper, 1985) and a number From Strobel et al., 1995

Figure 6: Representation of the Functional Domains of NADPH-Cytochrome P450 Reduct ase

Indicated are regions for binding NADPH, for binding to the endoplasmic reticulum, for interacting with cytochrome P450, and for interacting with other substrates, as well as regions for binding both of the flavin cofactors, FAD and FMN. 47 of other species has been determined. Generally, mammalian reductases are about 90%

homologous (Shen and Kasper, 1993; Yamano et al. , 1989). In humans, two polymorphic

sites within the protein sequence have been identified (Haniu et al., 1989). At position 499,

a valine or an alanine is present and at position 551, an arginine or a glutamine is present.

The sequence of the CYPRED cDNA and chromosomal localization of the

corresponding gene have also been determined in rats (Porter and Kasper, 1985; Murakami

et al. , 1986) and humans (Yamano et al. , 1989; Shephard et al. , 1992). Unlike the

cytochromes P450, CYPRED appears to be encoded by a single gene and the resulting protein is able to interact with al1 the cytochromes P450 as well as its other substrates

(Shephard et al., 1989; Shen and Kasper, 1993). Based on the protein polymorphisms, at

least three alleles for the CYPRED gene within the human population have been identified

(Haniu et al., 1989; Yamano et al., 1989; Shephard et al., 1992). Although, cytochrome

P450 function is closely associated with CYPRED function, CYPFCED gene expression is regulated independently of cytochrome P450 gene expression. For exampIe, CYPRED expression is induced by many, but not all, cytochrome P45O inducers, and usually the magnitude of CYPRED induction is smaller. Inducers of CYPREI) and cytochromes P450 include phenobarbital, trans-stilbene oxide (Gonzalez and Kasper, 1982) and dexamethasone

(Simmons et al., 1987). Dichlorodiphenyl trichloroethane (DDT)at high doses was reported to induce CYPRED without significant induction of total rat liver cytochromes P450

(Balakrishnan et al., 1985). One interesting aspect of CrPRED regulation is its differential inducibility. For example, phenobarbital induces microsomal CYPRED expression without significantly inducing its expression in the nuclear envelope (Gonzalez and Kasper, 1982). 48 The mechanism of CYPRED induction by phenobarbital appears to be through transcriptional

activation of the gene (Hardwick et al., 1983). Recent studies also indicate that thyroid

hormone status affects CYPRED expression in rats through transcriptional and post-

transcriptional pathways (Waxman et al., 1989; OYLearyet al., 1997).

1J.2 Tissue distribution

The cytochrome P450-dependent drug metabolism system (CYFTGD and cytochromes

P450) is widely distributed in mammalian species and in the various tissues of each species

(Strobel et al., 1995). In mammals the highest concentration is in the liver, but expression also occurs in lung, small intestine, kidney , colon, brain, marnrnary glands, heart, adrenals, skin and other tissues. In al1 tissues, CYPRED within a species has been shown to be essentially identical to hepatic CYPRED in terms of its irnrnunoiogic, physical, kinetic and catalytic properties. CYPRED levels are usually lower than that of total cytochrome P450.

In most tissues, for each molecule of CYPRED, there are 10-20 molecules of cytochrome

P450 (Reed et QZ., 1986; Parkinson, 1996). However, in the olfactory epithelium, the

CYPRED: P45O ratio is 1: 3 (Reed et al., 1986).

1.7.3 Occurrence in normal and tumour human breast tissue

In human breast tissue, expression of CYPRED and cytochromes P450 is not well characterized due to the limited arnount of accessible tissue and difficulties associated with the detection of low P450 levels (Hellrnond et al., 1998a; Spin. et al., 1998). Table 1 sumrnarizes the available information on cytochrome P450 protein expression in normal and tumour breast tissue in humans. In general, much lower levels of cytochrome P45O are detected in normal and tumour breast tissue compared to known levels in the liver (Smith et 49 al., 1993; Campaigne Larsen et al., 1998; Hellmold et al., 1998a). In normal breast tissue the foflowing cytochromes P450 have been detected by immunoblotting or immunohistochemistry: CYP2E1, CYPSAG, CYP4A11, CYP2C, CYP19, CYP2D6

(Forrester et al., 1990; Hellmold et al., 1998a). In a primary breast epithelial ceIl line,

CYPlBl and very Iow amounts of CYPlAl were detected (Campaigne Larsen et al., 1998).

In tumour tissue, CYPlB1, CYP2C, CYP2A, CYPlA and CYP3A were detected (Forrester et al., 1990; Murray et al., 1993; 1997; Smith et al., 1993; McKay et al., 1995). However, it appears that for both normal and tumour breast tissue, there is some variability in cytochrome P450 expression. For example, CYPIA1 /2, CYP2C8-10, CYP2E1 and CYP3A4 were not detected in normal and tumour tissue by Albin et al. (1993), although they were detected in the other studies. Also, mRNA of the following enzymes were detected in Iow levels in normal and tumour breast tissue: CYPlAl, CYPlBl, CYP2C, CYP2D6, CYP3A4,

CYP3A5 (Huang et al., 1996). In rats the cytochrome P450 content of breast tissue is approxirnateIy 0.1 % of that in the liver, and several specific forms of cytochrome P450 have been detected, which are regulated in the breast as a function of age and hormone status

(Hellmold et al., 1995; 1998b). Variability in human tissues may also be due to age, hormone status and other factors. 50

Table 1: Cytochromes P450 Detected in Normal and Tumour Tissue from Human Breast Samples by ImmunobIotting or Immunohistochemistry Breast Source of Tissue 1 Number of 1 CYP450 enzymes detected in Reference Tissue Type donor majority of samples some not detected or weak detection

- normal' reduction 1 CYP2E1, CYP2A6, 1 CYP19, Hellmold et al., 1998a

normal Murray et al., 1993

normal biopsy 1O - - Mway et al. , 1997 normal Albin et al., 1993

- -- - normal Forrester et al,, 1990

Ii Source of Tissue Number of CYP45O enzymes detected in Reference Tissue Type donor samples majority of samples some not detected or samples weak detection normal primary CYPlBl Campaigne epithelial cells Larsen et al., isolated from 1998 reduct ion mammoplasty tissue tumour biopsy - CYPlA Murray et al., CYP3A 1993 tumour biopsy CYPlBl Murray et al. , 1997 tumour biopsy CYPlBl McKay et al., 1995 tumour surgical tumour Albin et al., 1993 removal

tumour biopsy Forrester et al. , 1990 Breast Source of Tissue Number of CYP450 enzymes detected in Reference Tissue Type donor samples rnajority of samples-lLne 1 not detected or 1 1 samples ( weak detection 1 I turnour Smith et al. , 1993

1 total membrane fraction of tissue 2 Primary human breast tumour was grown as xenografts in mice and challenged with CYP450 inducers. Data sumarized in table are from control mice not admiminstered any inducers. ~munohistoche&al studies confhmed that protein expression was localized within malignant human epithelial cells rather than mouse-derived tissues. 53 One study was located in which CYPRED activity was measured in 14 samples of

hurnan breast tumour biopsies (Patterson et al. , 1997). The activity (presented in graphical form) ranged from approximately 20 to 140 nmol reduced cytochrome clrnidmg total membrane protein, with most values being close to 30 nmol/min/rng membrane protein.

This is approximately 3 to 8 times lower than CYPRED activity in human liver which ranged from 107 to 232 nmol reduced cytochrome c/min/mg microsomal protein (McManus et al.,

1987; Pearce et al. , 1996).

Expression of CYPRED also has been assessed in hurnan breast cancer ce11 lines.

Although breast cancer ce11 lines cannot be considered representative of normal or tumour breast cells since constitutive forms of cytochrome P450 may not be maintained (Hellmold et al., 1998a), nonetheless they provide a mode1 for assessing the role of drug-metabolizing enzymes in ce11 sensitivity to dmgs. In MCF-7/WT cells, CYPRED activity was reported to be approxirnately 18, 16 or 10 nmol reduced cytochrome c/min/mg protein in the total membrane fraction, in the ce11 lysate and in the S9 fraction (representing both microsomal and cytosolic proteins), respectively (Patterson et al., 1995; 1997; Fitzsimmons et al., 1996).

In MCF-7 cells selected to be resistant to DOX (MCF-7/ADR), the reported activity in the

S9 fraction was simiIar at 12.5 nmol reduced cytochrome c/min/mg protein (Fitzsimmons et al., 1996). In a number of other studies, slightly iower CYPRED activity was reported in

MCF-7IADR cells (32 to 49 nmol/rnin/mg microsomal protein) compared to MCF-7/WT cells (52 to 66 nmollmin/mg microsomal protein) (Sinha et al., 1987a; 1989; Mimnaugh et al., 1989; 1991). In general, cancer ce11 lines derived from breast tissue, including MCF-7, display below average CYPRED activity compared to other cancer ce11 lines, and usually 54 have lower activity than the two electron reducing enzyme DT-diaphorase (Fitzsimmons et

al., 1996).

MCF-7/WT and MCF-7/ADR cells have low CYPl Al activity (Cowan et al., 1986;

Ivy et al., 1988). However, in the presence of 2,3,7,8-tetrachlorodibenzo-p-dioxin(TCDD)

or other aromatic hydrocarbons, CYPlA1 and CYPlBl activity and expression are greatly

induced in MCF-7lWT cells (Spink et al., 1992; 1998; Arellano et al., 1993; Christou et al.,

1994; Dohr et al. , 1995), but not in MCF-7/ADR celIs (Cowan et al. , 1986; Ivy et al. ,

1988), a difference related to the estrogen receptor statu of the ce11 lines. CYPlAl and

CYPlBl are known to catalyze the metabolic activation of a number of procarcinogens (eg. polyaromatic hydrocarbons) and also the hydroxylation of 17P-estradiol (EJ (Spink et al.,

19%). Therefore, induced expression of these enzymes, via the aromatic hydrocarbon (Ah) receptor and other possible mechanisms, has a marked effect on procarcinogen and estrogen metabolism. Peters and Roelofs (1997) have suggested that CYPlAl activity/inducibility in

MCF-7lWT cells may be due to a point mutation in the CYPlAl gene (exon 7: isoleucine + valine) which leads to enhanced enzyme activity. The mutated form of CYPlAl was detected in MCF-7/WT cells, but not in MCF-7lADR cells. Perhaps the deficient inducibility of CYPlAl enzyme activity in MCF-7/ADR cells could lead to failure in activation of anti- cancer drugs and thus contribute to drug resistance (Peters and Roelofs, 1997).

1.8 Gap in Knowledge Leading to the Research Hypothesis

Based on the preceding literature review, it is apparent that reductive rnetabolism of

DOX to the DOX semi-quinone and subsequent redox-cycling has the potential to be a significant mechanism of cellular toxicity. Although in tumour cells, this mechanism of DOX toxicity does not appear to be a major mechanism relative to its topoisomerase II

inhibition mechanism, nonetheless modulation of drug biotransformation enzymes to increase

the extent of redox-cycling is a potential means of enhancing the effectiveness of DOX at the

site of action. Enhancing the effectiveness of DOX at the site of action would be desirable,

in order to allow use of lower DOX doses and limit its toxicity or to overcome drug

resistance caused by increased expression of antioxidant enzymes such as GSHPx. Enhanced

cytotoxic activity of DOX may be achieved by increasing the expression of bioactivating

enzymes in tumour cells. CYPRED from rats and rabbits catalyzes the reductive bioactivation of DOX; however, no studies have been identified that show human CYPRED also catalyzes this reaction. Since mammalian CYPREDs have greater than 90% homology, it is expected that human CYPRED would catalyze DOX reduction. However, extrapolation of rodent models to human systems may be misleading, since, for example, the catalytic rate of reduction can be very different in rodent versus human enzyme systems (Rass et al.,

1996). Therefore, it needs to be determined whether human CYPRED catalyzes DOX reduction. If this is the case, then it can be determined whether increased expression of human CYPRED in tumour cells would result in increased DOX cytoxicity. To date, no studies were located in which the cytotoxicity of DOX was evaluated in tumour cells, particularly breast cancer cells , overexpressing human CYPRED .

1.9 Research Hypotheses

1, The reductive bioactivation of DOX is catalyzed by human CYPRED.

2. Transfection of human CYPRED cDNA into human breast cancer cells results in

greater sensitivity to DOX . 1.O Experimental Approach and Rationde

Hv~othesis#1

A number of approaches were used to deterrnine whether human CYPRED catalyzes the reduction of DOX. These approaches were:

1. to deterrnine whether DOX reduction occurs in human liver microsomes and compare

rates to rat liver microsomes;

2. to deterrnine whether DOX reduction occurs in microsomes of cells overexpressing

human CYPRED (ie., Gentest heterologous expression systern);

3. to determine whether there is a correlation of DOX reduction with CYPRED activity

and immunoreactive protein in a bank of human liver microsomes; and

4. to conduct the following inhibition studies in human liver microsomes

a) inhibition of DOX reduction by:

i) chemical inhibitor ii) antibody inhibitor

iii) carbon monoxide

b) inhibition of CYF'RED activity by DOX.

Hypothesis #2

In order to determine whether transfection of human CYPRED cDNA into human

MCF-7breast cancer cells results in enhanced DOX cytotoxicity, the following objectives were established:

1. characterization of the human breast cancer ce11 Iines MCF-7/WT and MCF-7/ADR,

in ternis of growth and DOX cytotoxicity; 57 2. construction of a mamrnalian expression vector containing human CYPRED cDNA;

3. transfection of the vector containing human CYPRED cDNA into breast cancer cells

(MCF-7lWT and MCF-7IADR) and selection of stable transfectants;

4. confimation of increased CYPRED expression in transfected cells by CYPRED

immunoblots and CYPRED activity; and

5. conduct comparative DOX cytotoxicity assays in ce11 lines expressing varying levels

of CYPRED activity .

For the purposes of this thesis, the fnst two objectives under hypothesis #2 were included and the remaining objectives will be continued by others in the laboratory at a later tirne. 2. Methods

2.1 Chemicals, Reagents and Source of Materials

Doxorubicin hydrochloride, diethylenetriaminepentaacetic acid (DTPA) , cytochrome

c, 3-(4,S-dimethylthiazol-2-yl)-2,5-dipheny1te01i bromide (MTT)and NADPH were

obtained from Sigma Chemical Co. (St. Louis, MO). Diphenyliodonium chloride (DPIC) was

obtained from Aldrich Chemical Co. (Milwaukee, WI) . Geneticin was purchased from

Gibco/BRL Life Technologies (Gaithersburg, MD). A complete Iist of al1 other chemicals

used in the course of this study is given in Appendix A.

Microsornes from human lymphoblastoid cells transfected with human CYPRED

cDNA or vector were obtained from Gentest Corporation (Woburn, MA). A goat polyclonal

antibody raised against rat CYPRED and accompanying goat serum were also obtained from

Gentest Corporation. Anti-goat IgG horseradish peroxidase (HRP) conjugate was obtained

from Sigma Immuno Chemicals (St. Louis, MO).

The human breast adenocarcinorna ce11 line, MCF-7/WT, and the corresponding ce11

Iine selected for resistance to DOX, MCF-7/ADR, were obtained fiom Dr. James Ballinger

(Ontario Cancer Institute/Princess Margaret Hospital) who had originally obtained them from

the American Type Culture Collection (ATCC, RockviIle, MD) and from Dr. G. Batist

(Jewish General Hospital, Montreal), respectively .

Human CYPRED cDNA was a gift from Dr. Frank Gonzalez (National Cancer

Institute, National Institutes of Health, Bethesda, MD). The mammalian expression vector, pcDNA3.1 (+) was obtained from Invitrogen Corporation (Carlsbad, CA). Al1 DNA restriction enzymes, calf intestinal alkaline phosphatase (CIAP), T4 DNA ligase, RNase and ATP were obtained from Pharmacia Biotech Products (Uppsala, Sweden). Competent

Escherichia coli bacteria (strain DH5a) were a gift from Dr. J. Peter McPherson (Dr.

Goldenberg's Laboratory, Department of Pharmacology, University of Toronto). For DNA

sequencing reactions, [CY-~~SI~ATPwas obtained fiom Amershm (Arlington Heights, IL) and

T7 primer was synthesized by ACGT Corporation (Toronto, Ontario).

2.2 Sarnple Preparation

2.2.1 Rat liver microsornes

Four male Fischer 344 rats approximately nine weeks old and weighing 190 to 225 g were obtained from Harlan Sprague Dawley Inc (Indianapolis, IN). The rats were maintained in the Division of Comparative Medicine at the University of Toronto. Animals were cared for in accordance with the principles and guidelines of the Canadian Council on

Animal Care and al1 animal experirnentation was approved by the University of Toronto

Animal Care Cornmittee. Following an acclirnation period of 2 days, the rats were sacrificed by decapitation. The livers, perfused in situ with cold 1.15 % (w/v) potassium chloride

(KCI), were removed, weighed, chopped with scissors and homogenized in 4 volumes of cold phosphate-buffered KCI [l-15 % (w/v) KC1, 10 mM potassium phosphate, pH 7.4) using a motor-driven Potter-Elvehjem tissue grinder (Wheaton, Millville, NJ). The homogenate was then centrifuged at 9000 x g for 20 minutes at 4°C in a Beckrnan J2-21M centrifuge

(Fullerton, CA). The resulting pellet was discarded and the supernatant was further centrifuged at 106,000 x g for 60 minutes at 4°C in a Beckman L-80 ultracentrifuge. The resulting supernatant (cytosolic fraction) was discarded and the pellet (microsomal fraction) was resuspended in a small volume of phosphate-buffered KCl by homogenization. This 60 microsomal fraction was washed by further centrifugation at 106,000 x g for 60 minutes at

4°C in a Beckman L-80 ultracentrifuge. The resulting microsomal pellet was resuspended in storage buffer (approxirnately 1.5 ml) consisting of 10 mM Tris, 20% (vtv) glycerol and 1 mM EDTA at pH 7.4. Aliquots (approximately 1 ml) of this washed microsomal fraction were frozen by immersing in liquid nitrogen and then transferred to -70°C for long-term storage.

2.2.2 Human liver microsornes

Human liver samples were generously provided by Dr. Ted Inaba (Department of

Pharmacology, University of Toronto) who had obtained the livers in collaboration with Dr.

W.A. Mahon, Dr. B. Taylor (Toronto General Hospital), and Dr. M. Robinette (Metro

Organ Retrieval and Exchange).

The livers were obtained from individuals who were donating their kidneys for other purposes. The available information on the donors is listed in Appendix B. The donors, once designated as brain dead, were maintained on a circulation support system until their kidneys and livers were removed. The livers were then immersed in 1.15% (wlv) KC1 at O to 4OC and transported to a nearby laboratory, where they were immediately sliced into 1 cm thick pieces, frozen in dry ice and stored at -70°C in sealed plastic containers. These livers were designated with a code K (as for kidney donor) plus a number (eg., K22).

Microsornes were isolated from these donated livers as follows. Small pieces of liver

(approximately 10 g in totaI) were chipped from the frozen liver sIices and allowed to partially thaw on ice. The tissue was minced with a razor btade until it had the consistency of a thick paste and then added to one volume of buffer [SO mM Tris-HC1 and 1.15 % (wtv) 61 KCl, pH 7.41. The tissue was then homogenized for about 15 seconds using a Brinkrnam

Instruments Polytron (Westbury, NY), followed by furîher homogenization in a Potter-

Elvehjem glass tissue grinder. The remaining procedures were similar to procedures used

for isolating rat liver microsomes (section 2.2.l), with the following exceptions:

a Sorval1 RC2-B centrifuge (Norwalk, NY) was used for the first centrifugation (ie.,

9000 x g for 20 minutes);

the second centrifugation was conducted at 100,000 x g for 60 minutes;

washing of the microsomal fraction or the third centrifugation (ie., 100,000 x g for 60

minutes) was not done;

the microsomal pellet was resuspended in storage buffer (approximately 1.O ml)

consisting of 50 rnM Tris-HC1 and 10% (w/v) sucrose at pH 7.4;

aliquots of approximately 500 pl of the resuspended microsomal fraction were frozen

by placing in a freezer at -70°C.

Protein analysis

The protein concentration of the rat liver microsomes was determined by the method of Lowry et al. (1951). The assay was conducted at room temperature. Bovine serum albumin @SA) was used to prepare 0.5 ml protein standards at concentrations of O, 0.08,

0.16, 0.24, 0.32 and 0.4 mglml. In order to be within the range of the standard protein concentrations, triplicate aliquots of 5 pl of the rat liver microsomal sample were added to

495 pl water. To al1 standards and sarnples, 0.5 ml of 0.5 M NaOH was added. After 30 minutes, 5 ml of a solution containing 100 parts 2 % (w/v) sodium carbonate, 1 part 1 %

(w/v) cupric sulfate and 1 part 2% (w/v) sodium potassium tartrate was added to al1 62 standards and sarnples. After 10 minutes, 0.5 ml of 1.O M Folin phenol reagent was added.

After 30 minutes, the absorbance of the standards and sarnples was rneasured at 670 nm on a

Beckman DU-65 spectrophotometer (Fullerton, CA) using the first standard (O mg proteinhnl) as a calibration blank. The Quant II Quadratic Soft-Pac module (Beckman) was used to fit the standard curve via non-linear regression.

The protein concentration of the hurnan liver microsomes was determined using the

BCA protein assay reagent kit from Pierce Chernical Company (Rockford, IL). The kit included a BSA stock solution (2 mg/ml) for preparation of protein standards. Briefly, BSA protein standards were prepared at concentrations of 0, 100, 200, 300, 400 and 500 pglml.

Aliquots of human liver microsomal sarnples were diluted to be within the range of the standard protein concentrations (usually 1 in 100 or 1 in 150 dilutions). To O. 1 ml of each protein standard and unknown protein sample, 2.0 ml of working reagent was added. The working reagent was prepared from solutions included in the kit and contained sodium carbonate, sodium bicarbonate, BCA detection reagent, sodium tartrate, NaOH and copper sulfate. The standards and samples were then incubated in a shaking water bath at 37°C for

30 minutes. After the standards and samples were cooled to room temperature, the absorbance of each was measured at 562 nm on a Beckman DU-7 spectrophotorneter

(Fullerton, CA) using the first standard (O pg proteidml) as a calibration blank. A standard curve was plotted using linear regression analysis with the graphing program Prism v2.0 for

Power Macintosh (GraphPad Software Inc., San Diego CA). 2.3 Enzyme Assays and Other Analytical Techniques

2.3.1 CYPRED activity

CYPRED activity in microsomal preparations was assayed under aerobic conditions at

30°C in 1 ml incubation mixtures consisting of 300 rnM potassium phosphate buffer (pH 7.7)

and 70 pM cytochrome c. The incubation mixture also contained approximately 0.03, 0.1,

or 0.025 mg/ml microsomal protein from rat liver, hurnan Iiver or human lymphoblastoid cells, respectively. Reactions were initiated by addition of 1 mM NADPH and the rate of cyctochrome c reduction was determined spectophotometrically (Beckman DU-65 spectrophotometer) at 550 nrn based on an extinction coefficient of 21 mM-'cm-' (Phillips and

Langdon, 1962; Strobel and Dignam, 1978). The blank reaction consisted of al1 reagents without addition of microsomal protein. The rate of enzyme-catalyzed reaction was detennined by subtracting the rate of reaction occurring in the absence of microsomes.

Actual microsomal protein concentrations used in the CYPRED activity assays were determined by the Lowry assay (see Section 2.2.3). Aliquots of the original microsomal preparations from rat liver, human liver or human lymphoblastoid cells were diluted to approximately 0.3, 1, or 0.25 mg proteidml in 300 rnM potassium phosphate buffer pH 7.7, respectively. To confirm these protein concentrations, triplicate 100 or 200 pl aliquots of the diluted protein preparations were further diluted with water to a volume of 500 pl and then treated as samples for the Lowry assay. This further dilution of the protein samples was necessary to prevent precipitation during the Lowry assay due to the presence of potassium phosphate buffer (Peterson, 1979). 2.3.2 DOX reduction

A DOX stock solution at a concentration of 2 rng/ml (3.45 mM) was prepared in PBS

at pH 2.5 to 3.5. The stock solution was stored in a foil-wrapped glass container at 4°C.

For al1 experiments with DOX, a working solution of 1 mM DOX in 50 mM Tris HC1 buffer

was prepared from the stock solution.

DOX reduction was measured indirectly as the DOX-stimulated microsomal oxidation

of NADPH based on the method of Goeptar et al. (1993). NADPH oxidation in the

microsomal preparations was assayed under aerobic conditions at 37OC in 1 ml incubation

mixtures consisting of 50 mM Tris HC1 buffer (pH 7.4), 5 rnM magnesium chloride, 0.5

rnM diethylenetriaminepentaacetic acid (DTPA, a metal chelator), and 0.1 mM DOX. The

incubation mixture also contained approximately 0.2, 0.5, or 0.2 mg/ml microsomal protein from rat liver, human liver or human lymphoblastoid cells, respectively. Reactions were initiated by addition of 0.125 mM NADPH and the rate of NADPH oxidation was determined spectophotometrically (Beckman DU-65 spectrophotometer) at 340 nm based on an extinction coefficient of 6.22 mM"'cm''. c

In order to determine the rate of DOX-stimulated consumption of NADPH, three control groups with the absence/presence of microsomes and/or DOX were used for each test reaction, as outlined in Table 2. Control Groups Used To Measure DOX-stimulated NADPH Oxidation II Microsomes DOX NADPH (

Test Reaction 1 + + + + or - refers to the presence or absence of the various components

Microsomal consurnption of NADPH in the absence of DOX was calculated by subtraction of

Control #1 from Control #2. Microsomal consumption of NADPH in the presence of DOX was calculated by subtraction of Control #3 from the Test Reaction. DOX-stimulated microsomal NADPH consurnption was calculated by subtraction of microsomal NADPH consumption in the absence of DOX from microsomaI NADPH consurnption in the presence of DOX. DOX-stimulated microsomal NADPH consumption is referred to as DOX reduction in this thesis.

Actual microsomal protein concentrations used in the DOX reduction assays were determined by the Lowry assay (see Section 2.2.3). Aliquots of the original microsomal preparations from rat liver, hurnan liver or human lymphoblastoid cells were diluted to approxirnately 2, 5, or 2 mg proteidml in 50 mM Tris HCI buffer pH 7.4, respectively. To confirm these protein concentrations, 10 pl triplicate aliquots of the diluted protein preparations were further diluted with water to a volume of 500 pl and then treated as samples for the Lowry assay. 2.3.3 Inhibition studies

Diphenyliodonium chloride (DPIC) is a mechanism-based chemical inhibitor of

CYPRED (Tew, 1993; 07Donnellet al. , 1994). To assess DPIC inhibition of CYPRED

activity and DOX reduction, DPIC was added to the appropriate incubation mixtures at

concentrations of O, 0.16, 0.3 1, 0.63, 1.25, 2.5 and 5 rnM, prior to the initiation of the

reactions by addition of NADPH.

Carbon monoxide (CO) is an inhibitor of the cytochrornes P450 (Gibson and Skett,

1994). To assess CO inhibition of CYPRED activity and DOX reduction, CO was bubbled

through the appropriate incubation mixtures for one minute, just prior to the addition of

NADPH.

Antibody inhibition of CYPRED activity and DOX reduction was also conducted.

Polyclonal antibody that recognizes both rat and human CYPRED was produced by the manufacturer (Daiichi mire Chernicals Co. Ltd, Tokyo; supplied by Gentest) by irnnlunizing a goat with purified rat liver CYPRED. To measure inhibition of CYPRED activity and

DOX reduction, undiluted microsornes were incubated for 1 hour on ice with increasing concentrations of antibody in goat serum. Additionally, normal goat serum was added to the incubations in order to keep the amount of goat serum in each incubation constant afthough the amount of antibody for each incubation was different. Because only limited quantities of antibody were available, most antibody inhibition assays were conducted witll only two replicates and at slightly lower microsomal protein concentrations. These microsomal protein concentrations were expected (based on initial optimization experiments) to still produce high enough enzyme activity that would aliow detection and quantitation following inhibition by 67 antibody. For inhibition of rat microsomal CYPRED activity, the concentrations of antibody used were approximately 0, 10, 22.2 and 50 pl antibody1100 ~g microsomal protein; the amount of microsomal protein in the 1 ml reaction mixture was approximately 30 pg. For inhibition of human microsomal CYPRED activity, the concentrations of antibody used were approxirnately 0, 5, 15, and 40 pl antibody1100 pg microsomal protein; the amount of microsomal protein in the 1 ml reaction mixture was approximately 25 pg. For inhibition of rat rnicrosomal DOX reduction, the concentrations of antibody used were approximately 0,

9.8, 25, and 50 pl1100 pg microsomal protein; the amount of microsomal protein in the 1 ml reaction mixture was approximately 100 pg. For inhibition of human microsomal DOX reduction, the experiments were originally planned to have antibody concentrations of 0, 10,

20, and 50 pl1100 pg rnicrosomal protein with the amount of microsomal protein in the 1 ml reaction mixture being approximately 400 pg. However, based on the Lowry assay conducted subsequently, the microsomal protein concentration in the reaction mixture was actually 0.3 mgld which would make the antibody concentrations 0, 13, 26, and 65 pl1100 pg microsomal protein.

Inhibition of CYPRED activity by DOX was also assessed. The 1 ml incubation mixtures consisted of 50 mM Tris HCl buffer (pH 7.4), 30 pM cytochrome c, 1 rnM

NADPH and 0.07 or 0.04 mg/ml microsomal protein from rat liver or human liver, respectively. The amount of DOX added to the reaction mixtures was 0, 0.2, 0.3, 0.4, 0.5 or 0.6 mM, starting from a 1 rnM DOX solution in 50 mM Tris HC1 (pH 7.4) prepared from the original DOX stock solution (Section 2.3.2). Each reaction mixture had the same amount of PBS at pH 2.8 as would occur with the highest concentration of DOX added. It was 68 previously detennined that the addition of the PBS did not significantly affect the pH of the

reaction mixture.

2.3.4 Immunoblot analysis

Immunoblot analysis was conducted to detect and quantify the amount of

immunoreactive CYPRED protein in human liver microsomal samples. As a positive control

to identify the CYPRED Ummunoreactive band, microsornes from hurnan lyrnphoblastoid celIs

overexpressing human CYPRED were also used. Liver microsomal samples were diluted to

an approxirnate protein concentration of 5 to 10 mg/ml which was verified by the Lowry assay. These samples were further diluted to protein concentrations ranging from 0.25 to 5 mg/ml, following which they were diluted 1: 1 in 2X sarnple buffer [O. 125 M Tris-HC1, 20%

(v/v) glycerol, 4% (w/v) sodium doecyl sulfate (SDS), 10% (v/v) P-mercaptoethanol,

0.004 % (w/v) bromophenol blue, pH 6.81. Microsomes from human Iymphoblastoid cells were directly diluted in 1X sample buffer (0.0625 M Tris-HC1, 10% (vlv) glycerol, 2 %

(w/v) SDS, 5% (v/v) P-rnercaptoethanol, 0.002% (w/v) bromophenol blue, pH 6.8) to produce a protein concentration of O. 1 mg/ml. Al1 samples were then denatured by heating in boiling water for 5 minutes. These samples were then ready to load into the eIectrophoresis gel in equal volumes (10 pl) with: a) varying protein concentrations (1.25 to

15 pg protein/well) from one human liver (K27); or b) equal protein concentrations (10 pg proteidwell) from a bank of human Iivers. Sarnples from human lymphoblastoid cells were loaded at 5 to 10 pl or 0.5 to 1 pg proteidwell.

The microsomal proteins were separated by molecular mass using SDS- polyacrylamide gel electrophoresis (SDS-PAGE) (Laemmli, 1970). A Bio-Rad Mini-Protean 69 II gel apparatus attached to a Bio-Rad Model 1000/500 Power Supply was used for the gel

electrophoresis. Discontinuous polyacrylarnide gels measuring 70 x 80 x 0.75 mm were made. The gel consisted of a 10% separating gel and a 4% stacking gel. The percent value refers to the total percentage (wlv) of acrylarnide monomer and N,NY-methylenebis acrylarnide (BIS) crosslinker present in the gel. Both gels were 2.7 % in terms of the arnount of BIS crosslinker as a percent of the total amount of acrylarnide monomer and BIS crosslinker. The separating gel also consisted of 0.375 M Tris-HCl (pK 8.8), 0.1 % (w/v)

SDS, 0.05 % (w/v) ammonium persulfate and 0.05 % (vlv) N,N,N' ,N7-tetramethyl- ethylenediamine (TEMED). The stacking gel also consisted of 0.125 M Tris-HCl (pH 6.8),

0.1 % (w/v) SDS, 0.05 % (wh) ammonium persulfate and 0.1 % (v/v) TEMED.

Electrophoretic separations were conducted at 200 V constant voltage for approximately 45 minutes. The electrophoresis tank buffer consisted of 0.025 M Tris-HC1, 0.192 M glycine, and 0.1 % (wlv) SDS at pH 8.3. To determine the approximate molecular masses of the immunoreactive protein, 7 to 10 pl of Bio-Rad Prestained Low Range molecular mass markers were loaded into one or more lanes of the gel.

The separated proteins were transferred fiom the gel to nitrocellulose membranes

(Hybond ECL, Amersham) electrophoretically according to the method of Towbin et al.

(1979) and using a Bio-Rad Mini TransbIot electrophoretic transfer ce11 attached to a Bio-Rad

Model 200j2.0 Constant Voltage Power Supply . Gel transfer was conducted at 100 V constant voltage for 1 hour with cold transfer buffer and a Bio-Ice cooling unit. The transfer buffer consisted of 25 mM Tris, 192 mM glycine and 20% (vlv) methanol at pH 8.3.

To assess efficiency of transfer, the nitrocellulose membranes were covered with 70 0.2% (v/v) Ponceau S stain and placed on an orbital shaker (Bellco Glass Inc., Vineland

N.J.) for 10 minutes and then rinsed with distilled water. To block nonspecific binding sites,

the membranes were incubated in "Blotto" overnight at 4°C with gentle agitation on the

orbital shaker. Blotto consisted of 5%(w/v) skim milk powder, 20 mM Tris, 137 mM

NaCl, O. 1% (v/v) Tween-20, pH 7.6 and a pinch of thimerosal as preservative. Prior to use,

the Blotto was centrifuged at 600 x g for 10 minutes and the supernatant was used for al1

incubations. After the overnight incubations in Blotto, the membranes were washed by

shaking at room temperature in large volumes of TNT [20 rnM Tris, 137 mM NaCI, 0.1 %

(v/v) Tween-20 at pH 7.61 as follows: 3 quick rinses followed by a change of TNT every 10

minutes for a full 45 minutes. The primary antibody used was goat polyclonal anti-rat

CYPRED (Gentest), which according to the manufacturer also cross-reacts with human

CYPRED. The membranes were incubated with primary antibody dilutions in Blotto of

1:5000 for 1.5 hours with shaking at room temperature. After incubation with the primary antibody, the membranes were washed with TNT as described above. The secondary antibody was anti-goat Ig horseradish peroxidase (HRP)conjugate. The membranes were incubated with secondary antibody dilutions in Blotto of 15000 for 1.5 hours with shaking at room temperature and subsequently washed with TNT.

Chemiluminescence detection was carried out by mixing equal volumes of Amersharn

ECL detection reagents 1 and 2 and immersing the membranes in the mixture for 1 minute with shaking. The membranes were drained, wrapped in plastic wrap and placed in a Fisher

Biotech cassette (8 x 10 inch) (no intemiQing screens) with Kodak X-OmatTMAR film for the appropriate time period. The film was developed in a Kodak M35A X-omatic processor in the Department of Molecular and Medical Genetics, University of Toronto.

For relative quantitative analysis of the imrnunoblot bands, the films were first scanned using an UMAX SuperVista S-12 scanner with the aid of the prograrn Adobe

Photoshop v4.0 for Power Macintosh. Quantitative analysis was performed using the program IPLab Gel Scientific Image Processing v1 Se for Power Macintosh (Signal

Analytics, Vienna VA).

2.4 Ce11 Culture

2.4.1 Ce11 lines and culture techniques

The human breast adenocarcinorna ce11 line, MCF-7/WT, and the corresponding ceII line, MCF-7/ADR, selected for resistance to DOX were chosen for this study. The ce11 lines were received frozen and were thawed by rapid agitation in a 37°C water bath and diluted into medium in 80 cm2 (T-80) tissue culture flasks.

The ce11 Iines were maintained in a-minimal essential medium (a-MEM) supplemented with 10% fetal caIf semm in T-80 flasks. Cells were grown in a Queue Ce11

Culture Incubator (Parkersburg WV) at 37OC with a humidified atmosphere containing 5 %

CO,. Al1 ce11 culture work was conducted under sterile conditions in a SterilGARD (Baker

Co., Sanford MA) larninar flow tissue culture hood. The ce11 lines were subcultured after approximately five days of growth at dilutions ranging from 1: 10 to 1 :75, depending upon their growth characteristics and pending ce11 culture experiments. Subculture involved aspirating al1 growth medium from the near-confluent flasks with a sterile glass Pasteur pipette. The cells were washed twice with approximately 10 ml sterile PBS which was aspirated with each wash. To allow the cells to detach from the surface of the flask, 2 ml of sterile trypsin (0.25% w/v) was added. After an incubation period of approximately 3 minutes (MCF-7/ADR cells) or 10 minutes (MCF-7/WT cells), 8 to 15 ml of medium was added to stop trypsin activity. The ce11 suspension was mixed up and down in a sterile pipette to loosen large ce11 clumps and to dislodge any cells still remaining on the surface of the flask. Aliquots of 0.1 to 1.5 ml were dispensed into T-80 flasks already containhg approximately 13 ml of medium. During their growth, the medium of the cells was replaced as necessary .

The MCF-7IWT cells had a tendency to clump, even with vigourous up and down pipetting and long trypsinization times (up to 15 minutes). Therefore, for those assays requiring single ce11 suspensions of MCF-7/WT cells, the cells were subcultured with the following extra steps. Once the cells were in suspension with medium, the suspension was drawn up into a sterile syringe attached to a stainless steel Spinal Quincke 18G x 3.5" needle

(Becton Dickinson Company, Franklin Mes, NJ). The suspension was drawn up and down through the needle several times. While the suspension was in the syringe, the steel needle was then replaced with a sterile 25G x 518" needle (Becton Dickinson) and the suspension was then passed through the needle into a sterile falcon tube. The last step involved passing the ce11 suspension through a 40 pm ceIl strainer (Falcon) to remove any possible remaining clumps. This procedure increased ce11 counts as measured in the Coulter Counter by about

20% and most of the cells appeared to be singlets when viewed under the microscope

(Nikon, Mode1 TMS-F),although a few doublets and triplets were still visible. 2.4.2 Growth curve

The growth curve of each ce11 line in the absence of DOX was assessed. Rapidly growing cells from each ce11 line were harvested by trypsinization and counted in a Coulter

Counter (Mode1 ZM with Sampling Stand II). The cells were seeded in triplicate at both lxlOS and 5x104 cells per 35 mm diameter well, in 6-well tissue culture plates (Corning).

For each cell line, the cells from three culture wells were harvested every 24 hours for 5 days and counted in the Coulter Counter in triplicate. Mathematical manipulation of the raw data was performed using Microsoft Excel (Version 5.0a) for Power Macintosh on an Apple

Power Macintosh 8500/180 cornputer. The doubling tirne of the ce11 lines during the exponential growth phase was detemined.

2.4.3 MTT cytotoxicity assay

To assess the sensitivity of the ce11 lines to DOX, the MTT [3-(4,5-dimethylthiazol-2- y1)-2,5-diphenyltetrazoliuni bromide] assay was used, based on the method of Alley et al.

(1988). The premise of the assay is that MT',a water-soluble yellow tetrazolium salt which is readily taken up by the cells, is reduced by mitrochrondrial and microsomal enzymes in viable cells, to its corresponding purple water-insoluble formazan product. After an incubation period with the MTT, the forrnazan product is solubilized in dimethyl sulfoxide

(DMSO) and quantitated using a scanning multiwell spectrophotometer. The MTT assay is widely used for drug screening with a nurnber of human tumour ce11 lines and has produced results showing good correlation with other cytotoxicity assays such as the clonogenic assay and the dye exclusion assay (Carmichael et al., 1987; Alley et al., 1988). 74

The M'IYI; assay was conducted as follows. Rapidly growing cells were harvested by trypsinization and counted with a haemocytometer (la fontaine NEUBAUER irnproved Bright line). Using 96-well flat bottom microculture plates (Nunc) and a 12-channel pipetter

(Brinkmann 12-tramferpette 50-200 pl), each well was inoculated with 5000 cells in medium in a volume of 200 pl. Table 3 illustrates the standard plate setup for the MTT assay, including the blanks and controls. After 24 hours, when the cells were in a logarithmic growth phase, 100 pl of medium was removed from each well and replaced with 100 pl of medium containhg DOX at various concentrations or medium alone. Plates were then incubated for a further 72 hours to allow sufficient time for ce11 death to occur. To complete the assay, 50 pl of MTT at a concentration of 1 mglm1 (prepared as a 5 mg/ml solution in

PBS and diluted 1 in 5 in culture medium) was added to each well. To allow viable cells to metabolize MTT, the plates were incubated for 4 hours at 37°C. Then 200 pl of medium was removed from each well and replaced with 150 pl DMSO to solubilize the forrnazan crystals. To ensure thorough mixing, plates were placed on an orbital shaker (Bellco Glass

Inc., Vineland NJ) set at 300 rpm for 5 minutes. Absorbance at 540 nm was measured on a

Labsystems Multiskan ILS microplate reader. Data were collected using Delta Soft v3 software on a Macintosh Classic II cornputer and electronically transferred to a Microsoft

Excel v5.0a spreadsheet for manipulation. Table 3: Standard Setup- for the MTT Assay in a 96-We11 Plate ------1 2 3 4 5 6 7 8 9 10 11 12 A pb rned rned rned rned med rned rned rned rned rned rned B pb rned bl con ex1 ex2 ex3 ex4 ex5 ex6 ex7 rned C pb rned bl con ex1 ex2 ex3 ex4 ex5 ex6 ex7 rned D pb rned bl con ex1 ex2 ex3 ex4 ex5 ex6 ex7 rned E pb med bl con ex1 ex2 ex3 ex4 ex5 ex6 ex7 rned F pb rned bl con ex1 ex2 ex3 ex4 ex5 ex6 ex7 rned G pb rned bl con ex1 ex2 ex3 ex4 ex5 ex6 ex7 rned H pb rned rned rned rned rned rned rned med rned rned rned

Legend: pb = plate blank; rned = medium only (not used for assay); bI = experiment blank containing medium only; con = control for experiment, contains ceIls but no drugs; ex# = wells used for experirnent, contains cells with drug at differing concentrations. Al1 wells were treated equally in terms of medium removal and replacement, and addition of MTT. For a typical assay, the plate blank @b) gave essentially zero absorbance. The medium blanks (bl) in lane 3 (rows B to G) were averaged and this value, the absorbance of the medium alone, was subtracted from al1 other lanes. The value remaining in lane 4, the control lane, after these subtractions, represents the absorbance for cells unaffected by drug treatrnent and is assigned the value of 100%, to which al1 other drug treatment lanes are compared.

2.5 Molecular Biology Techniques

2.5.1 Construction of a mammalian expression vector containing human CYPRED

cDNA

The CYPRED cDNA (Yarnano et al., 1989) was received as a bacterial culture and stored at -80°C. Infornation concerning the bacterial strain, the type of vector containing the CYPED cDNA and the specific DNA restriction sites were not provided. However, a literature search was conducted to identify other investigators who used the CYPRED cDNA 76 from the same source. Based on these studies (Tamura et al., 1992; Belcourt et al., 1996a) and subsequent restriction digestion analysis, it was concluded that the cDNA was cloned into the EcoRI site of the phagemid pUC9. For selection purposes, this vector contains an ampicilIin resistance gene .

The following steps were involved in the construction of a mamrnalian expression vector construct containing the full-length human CYPRED cDNA (Figure 7):

isolation of single bacterial colonies containing the pUC9 vector (with the CYPRED

cDNA) by streaking the original bacterial inoculum ont0 an LB agar plate with

ampicillin;

growth of an overnight liquid culture of these bacteria from a single colony for small-

scale preparation of pUC9 DNA;

restriction enzyme analysis of the pUC9 DNA including cutting the DNA with EcolRI

to isolate the CYPRED cDNA (see Figure 8);

transformation of competent E. coli bacteria (strain DHSa) with the mammalian

expression vector pcDNA3.1(+) (see Figure 9) and growth of the transformed

bacteria on an LB agar plate with ampicillin;

growth of an overnight liquid culture of these transformed bacteria from a single

colony for small-scale preparation of pcDNA3.1 (+) DNA;

restriction enzyme analysis of the pcDNA3.1 (+) DNA including linearizing the

DNA at the EcoRI site and dephosphoryIating the 5' termini of the DNA with calf

intestinal alkaline phosphatase (CIAP) to minimize self-ligation;

ligation of CYPRED cDNA with pcDNA3.1 (+) at the EcoRI site; 8. transformation of competent E. coli bacteria (strain DHSa) with DNA frorn the

ligation reaction and growth of transforrned bacteria on an LB agar plate with

ampicillin;

9. screening colonies of these transformed bacteria by growing overnight liquid cultures

from a single colony for srnaIl-scale preparation of vector DNA and conducting

restriction digestion analysis to determine if the CYPRED cDNA is present and

ligated in the correct orientation.

Unless otherwise indicated, al1 procedures for the construction of the expression vector with the hwnan CYPRED cDNA were conducted according to the methods of

Sarnbrook et al. (1989) or Ausubel et al. (1997) under sterile conditions.

2.5.1.1 Smalbscale preparation of vector DNA

Single bacterial colonies were isolated by scraping the surface of frozen bacterial cultures with an inoculating loop and then immediately streaking the loop ont0 the surface of an LB agar plate with ampicillin [1%(wlv) bactotryptone, 0.5 % (wlv) bacto-yeast extract,

172 mM NaCI, 1.5% bacto-agar, 50 pg arnpicillin/rnl]. The plates were inverted and placed in a Fisher Scientific Isotemp Incubator (Mode1 630D) at 37°C overnight. Following colony formation, the plates were stored at 4°C until required.

For small-scale preparations of vector DNA, isolated colonies were picked from agar plates with an autoclaved toothpick and used to inoculate 5 ml of LB medium 11% (wlv) bactotryptone, 0.5% (w/v) bacto-yeast extract, 172 mM NaCl] containing 50 pg ampicillixdml. This liquid culture was incubated overnight in a G24 Environmental Incubator

Shaker (New Brunswick Scientific Co. Inc., Edison, NJ) set at 37°C with vigourous shaking. Figure 7: Construction of Mammalian Expression Vector with Human CYPRED cDNA

pcDNA3.'1(+) Exoressio n Vector

original bacterial inoculum (bacteria with WC9 containing CYPRED cDNA)

transform bacteria 4isolate single cdonies 4

ovemight liquid overnight Iiquid culture fmm 1 ctdony C culture fmm 1 colony

1 ornaIl scsle preparation of pK9 DNA 1 small scale prepantion of pcDNA3.l(+) DNA

cDNA

'ECORI site pUC9 DNA EcoRl restriction digest and EcoRl Restriction Digest dephosphorylation with ClAP -1 - 1

mal site 7.8 kb mal site pcDNA3.1(+) coRl site CORIsite religated pcDNA3.1(+) with cDNA pcDNA3.1(+) with cDNA in correct orientation in incorrect orientation (Figure continued on next page) 79 i Figure 7: Construction of Mammalian Expression Vector with Human CYPRED cDNA (contrd)

EcoRl site mal site 0 mal site cdsite CORI site pcDNA3.1(+) religated pmNA3.1 (+) with cDNA pcDNA3.1(+) with cDNA in correct orientation in incmect orientation

transfm baaeria with 1DNA ligation mixture

screen colonies by preparing 1ovemight liquid cultures

small scale vector DNA preparations

I EcoRl site mal site

mal site CORIsite GORIsite ~cDNA3.1(+1-7 kligated pcDNA3.1(+) with cDNA pcDNA3.1(+) with cDNA in correct orientation in incorrect orientation

restriction digest and gel etectrophoresis

vector 5.4 kb a 2.4 kb

restriction enzyme analysis and sequencing to determine orientation of cDNA insert (Figure continued on next page) 80 Figure 7: Construction of Mamrnafian Expression Vectot with Human CYPRED cDNA (cont'd)

Smal

pcDNA3.1(+) with cDNA pcDNA3.1(+) with cDNA in incorrect orientation in correct orientation

To detemine orientation of cDNA insert restriction digest with Smal and gel electrophoresis lac gene [*RI 263

pUC9 with CYPRED cDNA gene ampicillin resistance CYPREO

EcoRl 2661

Figure 8: Map of pUC9 Vector with Cloned CYPRED cDNA

oning Site

Afill Hindlll Asp7181 Kpn l BamHl BstXl EcoRl EcoRV BstXl Neomycin gene Notl resistance IXhol CMV = cytomegalovirus SV40 = simian virus 40

Figure 9: Map of peDNA3.1 (+) Expression Vecîor into which the CYPRED &NA was Subcloned 82 A small amount (1.5 ml) of rhis bacterial culture was placed into an Eppendorf microfuge tube and then centrifuged at 14,000 x g in a tabletop Eppendorf microcentrifuge (mode1

5415). The supernatant (medium) was removed, leaving the bacterial pellet as dry as possible. The pellet was resuspended in 100 pl of cold TEG (25 mM Tris, 10 mM EDTA,

50 rnM glucose). Following incubation at room temperature for 10 minutes, 200 pl of a solution containing 0.2 M NaOH and 1% (wlv) SDS was added. The tube was gently inverted 5 times and stored on ice for 5 minutes. Then, 500 pl of cold potassium acetate solution (3 M with respect to potassium, 5 M with respect to acetate, pH 4.8) was added.

The tube was mixed in an inverted position for 10 seconds, stored on ice for 5 minutes and then centrifuged at 14,000 x g for 5 minutes. The resulting supernatant was transferred to a fresh tube and 500 pl phenol:chloroform (1:l) was added. The tube was vortexed and centrifuged at 14,000 x g for two minutes. The top phase was removed from the tube to a fresh tube and 1 ml of 100% ethanol was added and mixed by vortexing. After 2 minutes at room temperature, the tube was centrifuged at 14,000 x g for 5 minutes. The supernatant was removed and al1 fluid was drained ont0 a paper towel. The remaining DNA pellet was washed by the addition of 1 ml of 70% ethanol and then recentrifuged for 5 minutes. Al1 the supernatant was rernoved and any remaining moisture was blotted with a Kim-Wipe. The

DNA pellet was resuspended in 30 to 50 pl of TE (10 mM Tris-HC1, 1 mM EDTA, pH 8) containing RNase (20 pglml). These vector DNA preparations were stored at 4°C until required . 2.5.1.2 Restriction enzyme digestion and agarose gel electrophoresis

Vector DNA was digested with various restriction enzymes under conditions 83 recommended by Pharmacia Biotech Products. Usually a 20 pl digestion reaction contained

1 pl of DNA, 1 pl of enzyme, and the appropriate amount of water and 10X One-Phor-Al1

Buffer Plus (Pharrnacia Biotech) to make the reaction 1X or 2X with respect to buffer.

Digestion tirnes for most enzymes were 2 to 3 hours at 37°C. The restriction enzyme SmaI

was used at 30°C. Uncut samples of DNA were also treated to the same conditions but

without the addition of restriction enzyme.

Following restriction digestion, DNA bands of various sizes were separated

electrophoretically using a Bio-Rad Mini- or Maxi-Sub Ce11 DNA Electrophoresis unit

attached to a Bio-Rad Model 20012.0 Constant Voltage Power Supply. Mini- (7~10~0.3cm)

or maxi- (15~15~0.3cm) gels containing 0.8 to 1.5% (wlv) agarose, 0.5 pglml ethidium

bromide, 40 rnM Tris-acetate and 1 rnM EDTA at pH 8.0 were prepared. Approximately

2.5 pl of 10x sample loading bufier (50% glycerol, 1 rnM EDTA pH 8.0, 0.25 % bromophenol blue, 0.25% xylene cyan01 FF) was added to the restriction digest mixtures and the entire sarnple was loaded onto the gel. To determine the approximate size of the DNA bands, 5 to 10 pl of a 1 kb DNA ladder (Gibco/BRL) was also loaded into one or more lanes of the gel. The buffer used during the electrophoresis was 1X TAE (40 rnM Tris-acetate/l mM EDTA, pH 8.0). The gels were run at 50 to 70 V until the dyes in the sample loading buffer had mn an appropriate distance (usually 45 minutes to 1.5 hours). The gels were viewed under ultraviolet (UV) light using a UV illumination box (Vilber Lounnat,

Intersciences Inc., Markam, ON). Photographs of the gels were taken using a Polaroid

Direct Screen Instant Camera DS34 with a Model QSP #14 Hood (International

Biotechnologies Inc., New Haven, CT.) and using Polaroid Professional Coaterless B&W Instant Pack film (8.5x10.8 cm). If a separated DNA band was required for further

manipulations, the band was cut from the rest of the gel using a razor blade and stored at

-20°C until required for gel purification.

2.5.1.3 Purification of DNA from agarose gels

The hurnan CYPRED cDNA was cloned into the EcoRI site of pUC9 vector (see

Figure 8). Restriction digestion of the vector with EcoRI (to isolate the 2.4 kb band representing the CYPRED cDNA) and ScaI (to further digest the 2.7 kb vector DNA into 1.8 and 0.9 kb fragments) was conducted, followed by gel electrophoresis. ScaI was included in the restriction digestion to enhance the resolution of the band representing CYPRED cDNA, since without ScaI, two bands of very similar sizes (2.4 versus 2.7 kb) were produced. The approximately 2.4 kb band representing the cDNA was cut from the rest of the gel and purified from the gel using a Qiagen (Chatsworth, CA) Qiaex II Kit for DNA Extraction from Agarose Gels. With this kit, DNA fragments are extracted and purified by solubilization of agarose and selective, quantitative adsorption of nucleic acids to the QIAEX

II silica particles in the presence of high salt. Briefly, the DNA band was excised from the agarose gel, placed in a 1.5 ml microfuge tube and weighed. To solubilize the agarose and allow adsorption of nucleic acids to the silica particles, a high salr buffer (QXI) and silica particles (QIAEX II) (both provided with the kit) were added. The mixture was incubated at

50°C for 10 minutes with mixing every two minutes to keep the silica particles in suspension. Following centrifugation at 14,000 x g for 30 sec, the supematant containing al1 non-nucleic acid impurities was rernoved. The pellet (containing the silica particles with adsorbed nucleic acids) was washed with the high salt buffer (QXI) to remove any residual 85 agarose contaminants. The pellet was then washed twice with an ethanol-containing buffer

(PE, provided with the kit) to remove residual salt contaminants. The pellet was then air-

dried at room temperature for 10 to 15 minutes. The DNA was then eluted by resuspending

the pellet in 20 pl of water and incubating the mixture at room temperature for 5 minutes.

Following centrifugation, the supernatant containing DNA was put into a clean tube. To

increase the yield of DNA, a second elution step was conducted using the remaining pellet

(the pellet is the silica particles) with 10 pl of water. In the end, the DNA was eluted into a

final volume of 30 pl of water and stored at -20°C until required for ligation reactions.

2.5.1.4 DNA ligation

The expression vector, pcDNA3.1 (+) (see Figure 9) was Iinearized with EcoRI and

prior to ligation with CYPRED cDNA, the expression vector DNA was treated with calf

intestinal alkaline phosphatase (CIAP). CIAP is an enzyme that catalyzes the hydrolysis of the 5'-phosphate residues in linearized DNA, replacing it with 5'-hydroxyl termini (Ausubel

et al., 1997). This prevents recircularization of the vector during ligation, while still allowing the insert to be ligated at the 3'-termini of the vector. Following restriction digest of pcDNA3.1 (+) with EcoRI, the reaction mixture (25 pl) was diluted 1: 1 with water such that the One-Phor-AI1 Buffer Plus concentration was lx. CIAP was diluted 20-fold in 10X

One-Phor-Al1 Buffer Plus and then 1 pl was added to the diluted digest reaction mixture.

This mixture was approximately 2 CIAP unitslml. The reaction mixture was incubated at

37°C for 0.5 hour. CIAP was then inactivated by heating the reaction mixture for 10 minutes at 8S°C. To precipitate the DNA from the reaction mixture, 5 pl of 3 M sodium acetate (pH 5.2) was added, followed by addition of 110 pl 100% ethanol. The mixture was stored on ice for 15 minutes and then centrifuged at 14,000 x g for 10 minutes. The supernatant was discarded and the resulting DNA pellet was resuspended in 100 pl of 10 mM

Tris (pH7.5). It was then stored at -20°C until required for the ligation reaction.

The DNA ligation reaction consisted of 6 pl of CYPRED cDNA purified from the agarose gel, 1 pl of pcDNA3.1 (+) cut with EcoRI and treated with CIAP, 1 pl of T4 ligase, 1 pl of 10x One-Phor-Al1 Buffer Plus and 1 pl of 10 mM ATP. The ligation reaction mixture was incubated in a 16°C waterbath for 22 hours.

2.5.1.5 Bacterial transformation

Both pcDNA3.1 (+) DNA and DNA from the ligation reaction with pcDNA3.1 (+)

DNA and human CYPRED cDNA were transformed into competent E. coli bacteria (strain

DHSa). To 50 pl of thawed competent bacterial cells, 1 pl of pcDNA3.1 (+) (ie., 1 ng

DNA) or 5 pl of DNA ligation mixture was added. The cells were incubated on ice for 30 minutes. They were then heat-shocked by placing them in a 37°C water bath for 30 seconds followed by immersion in wet ice for 2 minutes. To increase the volume of the ce11 suspension, 450 pl of SOC medium [2% (wh) bacto-tryptone, 0.5% (w/v) bacto-yeast extract, 8.6 rnM NaCl, 2.5 mM KC1, 20 rnM glucose, 10 mM MgCl,, pH 7.01 was added.

The ce11 suspension was then incubated for 1 hour at 37OC shaking at 225 rpm. To reduce the volume of the ce11 suspension, the cells were then centrifigecl for approximately 20 seconds at 14,000 x g and 300 pl of the supernatant was removed. The cells were resuspended gently in the remaining 200 pl of medium. Using a glass spreader, 50 and 150 pl of the ce11 suspension was spread over LB agar plates with ampicillin (50 pglml). The plates were inverted and incubated overnight at 37°C. Following colony formation, the 87 plates were stored at 4°C until required for small-scale vector DNA preparations of the

bacterial colonies.

2.5.2 DNA sequencing

In addition to restriction enzyme analysis, DNA sequencing of a portion of the

expression vector ligated with hurnan CYPRED cDNA was conducted to confirrn that the

correct DNA construct was made. DNA sequencing was conducted using the Phamacia

Biotech ?3equencing Kit, which is based on the Sanger-Coulson method of chain-tenninating

nucleotides (Sanger et al. , 1977). Briefly, using methods described in Section 2.5.1.1,

small-scale vector DNA preparations were made from bacteria carrying the ligated vector.

Single-stranded DNA is required as a template for the sequencing and for annealing of the

primer to the DNA. Therefore, as per the sequencing kit, 10 pl of the double-stranded

vector DNA from the mini-prep was denatured using 2 M NaOH. The denatured DNA was ethanoI-precipitated, washed and then resuspended in 10 pl distilled water. In the expression vector LpcDNA3.1 (+)], a T7 priming site (ie., TAATACGACTCACTAT) occurs just upstream of the multiple cloning site in which the CYPRED cDNA was ligated (see Figure

9). The annealing reaction was conducted by combining 3 pl of T7 primer, 10 pl of resuspended single-stranded DNA, and 2 11 of Annealing Buffer included in the kit. The reaction was incubated at 65°C for 5 minutes and then quickly transferred to a 37°C water bath for 10 minutes. It was placed at room temperature for 10 minutes before proceeding to the sequencing reactions. The sequencing reactions consist of labelling reactions and temination reactions. The labelling reaction involves the enzyme-catalyzed synthesis of the complementary DNA strand starting from the primer. The reaction consists of T7 DNA polymerase and limiting concentrations of al1 four deoxynucleotides (ie., dATP, dTTP, dGTP, dCTP), one of which is radio-labelled. In this case, [CY-~'S]~ATP( > 1000 Cilmmol) was used. A11 other reagents were provided in the kit. The reaction consisted of the annealed DNA template (15 pl), the labelling mix with the 4 deoxynucleotides (3 pl), 1 pl

[CX-~~SI~ATPand 2 pl of diluted (1 in 5) T7 DNA polymerase. The reaction was incubated at room temperature for 5 minutes before proceeding to the terrnination reaction. The terrnination reaction consists of al1 four deoxynucleotides in non-limiting concentrations and also one of four dideoxynucleotides. The dideoxynucleotides are incorporated as nucleotides into the DNA, but block further synthesis. Therefore, the reaction with dideoxyATP would consist of DNA strands al1 terrninating with an adenine residue. Four termination reactions were conducted using each of the four dideoxynucleotides. Al1 reagents for the termination reactions were provided in the kit. The reactions consisted of 4.5 pl of the labelling reaction and 2.5 pl of the appropriate "Mix Short" reagent (ie., deoxynucleotides + specific A, C, G, or T dideoxynucIeotides). The reactions were incubated at 37°C for 5 minutes and then 5 pl of the stop solution containing bromophenol blue and xylene cyan01 FF (provided in the kit) was added.

For each sequencing reaction, DNA of varying base pair lengths were separated by size electrophoretically using a thin polyacrylamide gel under denaturing conditions and using the Stratagene (La Jolla, CA) BaseAce Sequencer apparatus. The gel was 36.5 x 40 cm and consisted of 8 M urea, 6% (wlv) acrylamideIBIS, 0.09 M Tris-borate, 2 mM EDTA, 0.05%

(v/v) TEMED and 0.04 % (w/v) ammonium persulfate. The buffer in the electrophoresis apparatus was lx TBE (0.09 M Tris-borate12 mM EDTA, pH 8.3). The gel was pre-mn for 89 1 hour at 40 to 50 watts. Prior to loading, the DNA samples were incubated for 3 minutes at 90°C. Then 2 pl of each sample was loaded beside each other (ie., samples ending with adenine, cytosine, guanine and thymine residues). The next four Ianes were left empty. The gel was run at 50 watts for approximately 3 hours. To increase the arnount of readable sequence, the samples were re-loaded in the four empty wells and the gel was run for an additional 3 hours. The gel was removed, blotted ont0 filter paper and covered with plastic wrap. The gel was dried for 1 hour on the Bio-Rad Gel Dryer (Mode1 583). The plastic wrap was removed and the gel was placed in a Fisher Biotech Autoradiography FBxC 1417 cassette (14 x 17 inches) with Kodak BioMax MR Film (35 x 43 cm) for the appropriate time period. The film was developed in a Kodak M35A X-ornatic processor.

2.5.3 Optimization of antibiotic concentration for selection of stable transfectants

Prior to transfection of the ce11 lines with vector containing human CYPWD cDNA, selection conditions for each parental ce11 line were deterrnined. The expression vector, pcDNA3.1 (+) contained a gene coding for bacterial aminoglycoside phosphotransferase, which would confer resistance to arninoglycoside antibiotics such as neomycin, kanomycin and geneticin (G418) (Sarnbrook et al., 1989). To determine the optimal antibiotic concentration to use when establishing and selecting for a stable ce11 line, untransfected

MCF-7lWT and MCF-7lADR cells were seeded in triplicate at 10,000 cells/well in 6-well tissue culture plates containing medium with geneticin at concentrations of 0, 200, 400, 600,

800 or 1000 pglrnl. The geneticin was prepared in 100 mM HEPES (pH 7.3) so that addition of the antibiotic would not significantly alter the pH of the medium (Ausubel et al.,

1997). The cells were incubated for 6 days and were fed with the selective medium every few days as necessary. The plates were then stained with a dye containing 0.5% (wlv) methylene blue and 50% (vlv) methanol and the degree of ce11 growth was assessed by the presence and density of the blue stain. The relative density of the stain was quantitated by scanning the culture plates using an UMAX SuperVista S-12 scanner with the aid of the program Adobe Photoshop v4.0 for Power Macintosh. Quantitative analysis was performed using the prograrn IPLab Gel Scientific Image Processing v 1.5e for Power Macintosh (Signal

Analytics, Vienna VA). The minimum concentration of geneticin that resulted in cornplete ce11 death was to be used to select stable transfectants.

2.6 Statistical Analysis

Where appropriate, data are presented as mean f standard deviation (SD). Comparisons of two independent treatment groups were carried out by Student's t-test. Comparisons of three or more independent treatment groups were carried out by randomized design one-way analysis of variance (ANOVA) followed by post-hoc Newman-Keuls test. Statistical significance of linear correlation data was determined by the Pearson correlation test. In al1 cases, a result was considered to be statistically significant at pS0.05. Al1 statistical analyses were performed on original raw data (not percentages) with the program InStat

(GraphPad) . 3.1 Hypothesis #1: The Reductive Bioactivation of DOX is Catalyzed by Human CYPRED

3.1.1 Optimization of enzyme assays and immunoblots

3.1.1.1 CYPRED activity

In rat hepatic microsomes, the assay for CYPRED activity was optimized previously

by others in the laboratory and it was found that optimal enzyme activity occurred at concentrations of 70 pM cytochrorne c, 0.03 mg microsomal proteidml and 1 rnM NADPH.

Under these conditions, CYPRlED activity in the four rats was 353.5 f 17.9 nmol/min/rng protein.

For human liver microsomes, the same concentrations of cytochrome c and NADPH that were used with rat liver microsomes were used. The assay was optimized by varying the concentration of human liver microsomal protein to determine the linear range of protein concentrations for this enzyme assay (Figure 10). The data presented in Figure 10 indicate that with increasing protein concentration, CYPRED activity increased linearly and that at the concentrations of cytochrome c and NADPH used, the protein concentration was rate- limiting. Therefore, it was decided to use protein concentrations of O. l mg/rnl for subsequent assays using human liver microsomes. Although the data were fit by linear regression, it does appear that CYPRED activity is begiming to level off at the higher protein concentrations used. Figure 10: CYPRED Activity in Human Liver Microsornes CYPRED activity was assayed spectrophotometrically at 550 nm in 1 mi incubation mixtures consisting of 300 mM potassium phosphate buffer (pH 7.7),70 pM cytochmme c, 1 mM NADPH and varying microsomal protein concentrations from human liver K26. Each data point is the mean f SD of triplicate determinations. The equation of the line of best fit was generated by least-squares linear regression analysis. 93 3.1.1.2 DOX reduction

To assay DOX reduction in rat liver microsomes, the optimal concentrations of DOX,

NADPH and microsomal protein were determined by conducting assays in which

concentrations of two of the three substances were kept constant, while the third was varied

(Figure 11). Concentrations of NADPH (0.125 mM) and DOX (O. 1 mM) that produced

readily detectable DOX reduction activity without causing spectrophotometric interference

and a microsomal protein concentration from within the linear range (0.2 mgfml) were chosen for subsequent assays using rat liver microsomes.

To assay DOX reduction in human liver microsomes, the same concentrations of

DOX and NADPH that were used with rat liver rnicrosomes were used. The assay was optirnized by varying the concentration of human liver rnicrosomal protein to determine the linear range of protein concentrations for this enzyme assay (Figure 12). The data presented in Figure 12 indicate that with increasing protein concentration, DOX-stimulated NADPH consumption increased Iinearly and that at the concentrations of DOX and NADPH used, the protein concentration was rate-limiting. Therefore, it was decided to use protein concentrations of 0.5 rng/ml for subsequent assays using humôn liver microsomes. These results suggest that an enzyme or enzymes present in the human liver microsomes catalyzed

DOX reduction. I I I I I 1 0.00 0.05 0.10 0.15 0.20 0.25 0.30 NADPH Concentration DOX Concentration (mM) (mM)

Figure Il: Optimization of DOX Reduction in Rat Liver Microsornes To determine optimal concentrations of DOX, NADPH and microsomal protein for the assay, 2 of the 3 substances were kept constant, while the third was varied. DOX reduction was measured spectrophotometrically at 340 nm in 1 ml incubation mixtures which also contained 50 rnM Tris HCI buffer (pH 7.4), 5 rnM magnesium chloride -n- without DOX and 0.5 mM DTPA. Each data point is the mean f SD of triplicate determinations. A. Measurement of DOX reduction with varying NADPH concentrations, 0.1 mM DOX and 0.18 mg microsomal proteinhl from rat liver. NADPH concentrations of 0.5 and 0.75 rnM were also used but did not produce usehl results due to extremely high absorbance values. B. Measurement of DOX reduction with varying DOX concentrations, 0.125 mM NADPH and 0.15 mg microsornai proteidml from rat liver. C. Measurement of DOX reduction with varying rat liver microsornai protein concentrations, 0.125 mM NADPH and 0.1 mM DOX. Protein Concentration mcirm Figure 12: DOX Reduction in Human Liver Microsornes DOX reduction was assayed spectrophotometrically at 340 nm in 1 ml incubation mixtures consisting of 50 mM Tris HCl buffer (pH 7.4), 5 mM magnesium chloride, 0.5 rnM DTPA, 0.1 rnM DOX, 0.125 mM NADPH and varylng microsoma1 protein concentrations from human liver K27. Each data point is the mean f SD of triplicate determinations. The equation of the line of best fit was generated by least-squares Iinear regression analysis. 3.1.1.3 Immunoblots

Prior to conducting immunoblots for the detection and relative quantitative analysis of

CYPRED protein in a badc of hurnan liver microsomal samples, an immunoblot was conducted using one human liver sample (K27) at various protein concentrations (Figure 13).

This was used to plot a standard curve of protein concentration versus imrnunoreactivity. As shown in Figure 13, with increasing protein concentration used, the CYPRED immunoreactivity increased linearly. A protein concentration of 10 pgIweII, which falls within the standard curve, was selected for subsequent imrnunoblots with human liver microsomal samples .

Hurnan liver CYPRED is approximately 77 kDa (Yarnano et al., 1989), which was approximately where the major imrnunoreactive band from the human liver microsomes rnigrated on the immunoblot. As a positive control, microsomes from human lymphoblastoid cells overexpressing human CYPRED were also used in the immunoblot. The immunoreactive band fiorn the human lyrnphoblastoid cells migrated slightly fur-ther than the immunoreactive bands from the human liver microsomes. This discrepancy was also noted in independent immunoblots conducted by the suppliers of the hurnan lymphobIastoid ce11 rnicrosomes (Gentest Corporation) and they are currently investigating further to detennine why the lymphoblastoid-expressed CYPRED has a different mobility than that seen in human liver microsomes. In other immunoblots (data not shown) using higher Iymphoblastoid microsomal protein concentrations, two immunoreactive bands were visible: a) a very faint band which had the same mobility as that seen in the human liver microsomes and which most likely represents the endogenous lymphoblastoid CYPRED, and b) the very dark band Protein . Concentration . (~g/well) kDa 1 .zs 2.5 5 IO 1s .A

* microsornes from human lymphoblastoid cells overexpressing human CYPRED (0.5 pg)

Prote in Concentration

Figure 13: ~imoblotof CYPRED Protein Expression in Human Liver - Standard Curve of Rotein Concentration Versus ~unoreactïvity hunoblot was produced by loading varying concentrations of microsomal protein from human liver sample K27 or 0.5 pg microsomal proteinfwell from human lymphoblastoid cells overexpressing human CYPRED on a 10%acrylamide gel. The Hybond ECL membrane (Amersham) was incubated with 1:Sûûû primary aati-CYPRED antibody (Gentest) and 1:50 secondary anti-goat Ig HRP conjugate (Sigma Unmunochemicals). Using enhanced cherniluminescence detection the membrane was exposed to Kodak film for 3 minutes and then developed. The equation of the line of best fit was generated by least-squares linear regression analysis. (Film exposure times of 1, 5 and 10 minutes similarly produced a linear standard cuve of protein concentration versus immunoreactivity. Data not shown) which migrated slightly Merand which represents the overexpressed CYPRED from

transfection of a vector encoding the human liver CYPRED cDNA as cloned by Yamano et al. (1989). Therefore, the immunoreactive bands from the hurnan liver microsomal sarnples

at approximately 77 kDa represent the hepatic CYPRED protein.

3.1.2 CYPRED activity and DOX reduction in microsomes from human lymphoblastoid

cells

A number of approaches were used to test the first hypothesis that human CYPRED catalyzes the reduction of DOX. The first approach was to detennine whether significant

DOX reduction occurred in rnicrosomes of lymphoblastoid cells overexpressing human

CYPRED. As shown in Figure 14, CYPRED activity and DOX reduction were more than

10-fold higher in microsomes from lymphoblastoid cells transfected with cDNA encoding human CYPRED compared to microsornes from cells transfected with vector alone. Since the only apparent difference between the vector microsomes and the CYPICED microsornes is the overexpression of CYPRED and increased CWRED activity, the data therefore suggest that CYPREiD catalyzed the increased DOX reduction in this system. -DOX Reduction 1

" Vector Microsornes

Figure 14: CYPRED Activity and DOX Reduction in Microsornes from Human Lymphoblastoid Cells CYPKED activity was assayed spectrophotometrically at 550 nm in 1 ml incubation mixtures consisting of 300 mM potassium phosphate buffer (pH 7.7), 70 pM cytochrome c, 1 mM NADPH and 0.025 mg microsomal proteinhnl from human lymphoblastoid cells. DOX reduction was assayed spectrophotometrically at 340 nm in 1 ml incubation mixtures consisting of 50 mM Tris HC1 buffer (pH 7.4), 5 mM magnesiun chloride, 0.5 mM DTPA, 0.1 mM DOX, 0.125 mM NADPH and 0.2 mg microsomal proteinlm1 fiom human lymphoblastoid cells. Vector microsomes refers to microsomes fiom cells transfected with vector ody. CYPRED microsomes refers to rnicrosomes from ceIls transfected with vector containhg human CYPRED cDNA. Each data point is the mean f SD of triplicate determinations. *Significantly different from vector microsomes, based on Student's t-test @ I; O. OS). 3.1.3 Correlation analysis using a bank of human liver microsornes

The second approach was to determine whether the rate of DOX reduction was correlated with CYPRED activity and protein expression in a bank of 17 hurnan livers.

3.1.3.1 DOX reduction with CYPRED activity

Bot. DOX reduction and CYPRED activity in the human liver microsomal sarnples are presented in Figure 15. CYPRED activity varied approxirnately 2.4-fold (range: 66 to

155 nmol/rnin/mg protein; median = 91 nmol/min/mg protein). DOX reduction varied approxirnately 4.4-fold (range: 2.04 to 8.94 nrnol/min/mg protein; median = 3.98 nmol/min/mg protein).. A statistically significant correlation was observed between DOX reduction and CYPRED activity (r = 0.84; p < 0.0001). The coefficient of determination

(3)was 0.70, suggesting that about 70% of the variability in DOX reduction could be attributed to the variability in CYPRED activity. A CYPRED Actlvity ' DOX Reduction

Human Liver Sample Number Human Livsr Sample Number

C Correlation Analysis 10.o-r

1 1 1 i 50 100 150 200 CYPRED Actlvity (nrnoUmlnlmg protein)

Figure 15: Correlation of CYPRED Activity and DOX Reduction in a Bank of Human Livers CYPRED activity (A) was assayed spectrophotometrically at 550 nm in 1 mi incubation mixtures consisting of 300 mM potassium phosphate buffer (pH 7.7),70 pM cytochrome c, 1 mM NADPH and 0.1 mg microsomal protein/&. DOX reduction (B) was assayed spectrophotometrically at 340 nm in 1 ml incubation mixtures consisting of 50 mM Tris HC1 buffer (pH 7.4), 5 mM magnesium chloride, 0.5 rnM DTPA, 0.1 mM DOX, 0.125 mM NADPH and 0.5 mg microsomal protein/ml. Each bar is the mean f SD of triplicate determinations. (C) The equation of the line of best fit was generated by least-squares linear regression analysis. Statistical significance of the correIation was determined by the Pearson correlation test. 3.1.3.2 DOX reduction with immunoreactive protein

To determine whether DOX reduction correlated with CYPRED protein expression in

the human liver microsomal samples, immunoblots were conducted (Figure 16, panels A and

B). Due to the number of liver samples, two blots were required. To compare relative

immunoreactivities of al1 liver sarnples, the immunoreactivities were normalized to liver

sample K21 which was run on both blots. As a positive confirmation of a predicted relationship, CYPRED immunoreactivity correlated significantly with CYPRED catalytic activity (r = 0.86; p < 0.0001). Therefore, CYPRED protein expression as measured by immunoblots is related to CYPRED activity. Furthermore, a statistically significant correlation was observed between DOX reduction and CYPRED immunoreactivity (r =

0.76; p = 0.0004), providing further evidence that CYPRED contributes significantly to

DOX reduction in human liver microsornes.

3.1.4 Inhibition studies

The third approach to test the hypothesis that CYPRED catalyzes DOX reduction was

to determine whether inhibitors of CYPRED activity would also inhibit DOX reduction and

whether DOX would inhibit CYPRED activity. Both chemical and antibody inhibitors of

CYPRED were used.

3.1.4.1 Chemical inhibitor

DPIC is a mechanism-based chemical inhibitor of CYPRED (Tew, 1993; O'Domeil et al., 1994). Results of inhibition studies conducted with DPIC using rat and human liver microsomes are presented in Figure 17. In both rat and hurnan liver microsomes, DPIC inhibited CYPRED activity in a concentration-dependent marner. In human liver microsomes, DPIC also inhibited DOX reduction to a similar extent as it inhibited CYPRED activity (up to approximately 80% inhibition), and the sensitivity of both reactions to DPIC was similar (IC,, = 0.6 mM). In rat liver microsomes, DPIC also inhibited DOX reduction, but a maximum of only 60% inhibition was achieved, whereas CYPRED activity was inhibited by up to 80%. Since an inhibitor of CYPRED also inhibited DOX reduction, these data suggest that both human and rat CYPRED may catalyze DOX reduction. The data also indicate that in rat liver microsomes in particular, other enzymes in addition to CYPRED may catalyze DOX reduction. A Rat Llver Microsornes

-c- CYPRED Activity -t- DOX Reduction * 40 * 30 * 20 * * 10

O! I I I I 1 O 1 2 3 4 5 DPIC Concentration (mW Human Liver Microsomes

+CY PRED activity +DOX reduction

O 1 1 I I 0.0 0.5 1.0 1.5 2.0 2.58 3.01 DPIC Concentration (mM)

Figure 17: DPIC Inhibition of CYPRED Acüvity and DOX Reduction in Rat (A) and Human (B) Liver Microsomes CYPRED activity was assayed spectrophotometrically at 550 nm in 1 ml incubation mixtures consisting of 300 mM potassium phosphate buffer (pH 7.7), 70 pM cytochrome c, 1 rnM NADPH and 0.03 (rat) or 0.1 (human) mg microsomal protein/ml. DOX reduction was assayed spectrophotometrically at 340 nm in 1 ml incubation mixtures consisting of 50 mM Tris HCl buffer (pH 7.4), 5 mM magnesium chloride, 0.5 rnM DTPA, 0.1 mM DOX, 0.125 mM NADPH and 0.2 (rat) or 0.5 (human) mg microsomal proteinlml. Human liver sarnpie K15 was used for the assays. DPIC was added to the incubation mixtures at concentrations of 0, 0.16, 0.3 1, 0.63, 1.25, 2.5 or 5 (rat only) mM, prior to the initiation of the reactions by addition of NADPH. Each data point is the rnean SD of tripliate detedmtions. 100% control activites for CYPRED activity in rats and humans were 359.85 and 101.65 nmol/minlmg protein, respectively. 100% control activites for DOX reduction in rats and humans were 12.50 and 5.O7 nmol/min/mg protein, respectively . *Significantly different from enzyme activity in the absence of DPIC, based on randomized design one-way ANOVA and post-hoc Newman-Keuis test @ 5 0.05). 106 3.1.4-2 Antibody inhibitor

Results of inhibition studies conducted with a goat polyclonal antibody raised against rat CYPRED (and that cross-reacts with human CYPRIED) are presented in Figure 18. In both rat and human liver microsomes, the antibody inhibited CYPRED activity in a concentration-dependent marner. In rat liver microsomes, DOX reduction appeared to be inhibited at 9.8 and 25 pl antibody/100 pg microsomal protein (32 and 62% inhibition, respectively). However, this inhibition was not strictly concentration-dependent, since at the highest concentration tested (50 pl antibody/100 pg microsomal protein), the extent of inhibition was only 27%. In human liver microsomes, marginal inhibition appeared to occur at 13 and 26 pl antibody/100 pg microsomal protein (8 and 27% inhibition, respectively); however, as in the rat, this inhibition was not strictly concentration-dependent since no inhibition occurred at the highest concentration of 65 pl antibody/100 pg microsomal protein.

These results indicate that while antibody inhibition of CYPRED activity occurred in rat and human liver microsomes, inhibition of DOX reduction was marginal and concentration- independent. Rat Liver Microsomes

Antibody Concentration bU100 pg microsomal protein) Human Lier Microsomes

-t DOX Reduction

I 1 1 I o IO 20 30 40 5b 6'0 Antibody Concentration @MO0pg rnicrosomal protein)

Figure 18: Antibody Inhibition of CYBRED Activity and DOX Reduction in Rat (A) and Human (B) Livw Microsomes CYPRED activity was assayed spectrophotometrictilly at 550 nm in 1 ml incubation mixtures consisting of 300 mM potassium phosphate buffer @W 7.7), 70 pM cytochrome c, 1 mM NADPH and 0.03 (rat) or 0.025 (hm) mg microsomal protein/ml. DOX reduction was assayed spectrophotometncaily at 340 nm in 1 ml incubation mixtures consisting of 50 mM Tris HCl buffer (pH 7.4), 5 rnM magnesium chloride, 0.5 mM DTPA, 0.1 mM DOX, 0.125 mM NADPH and 0.1 (rat) or 0.3 (human) mg microsomal protein/ml. Human fiver sample K15 was used for the assays. Undiluted microsornes were incubated with antibody against CYPRED for 1 hour on ice prior to conducting the assays. Antibody concentrations were as follows: for CYPRED activity in rat liver, 0, IO, 22.2 and 50 pl/ 100 pg microsomal protein; for CYPRED activity in human liver, 0, 5, 15 and 40 y1/100 pg microsomal protein; for DOX~eductionin rat liver, 0, 9.8, 25 and 50 p1/100 pg microsomal protein; and for DOX reduction in huma. liver, 0, 13, 26 and 65 pi/100 pg microsomal protein. Ail reactions contained equal amounts of goat sem. Each data point is the mean of duplicate determinations, with the exception of CYPRED activity in rats in which each point is the mean f SD of triplicate determinations. 100% wntrol activity of CYPRED in rats was 253.16 f 3.14 nmol/min/mg protein. The range of CYPRED activity in humans was 120.32-123.60, 74.91-83.11, 44.26-45 .go, and 28.85-32.62 nmol/min/mg protein for antibody concentrations of 0, 5, 15 and 40 p1/100 pg microsorna1 protein, respectively. The range of DOX reduction in rats was 28.45-28.45, 14.23-24.19, 4.27-17 .O7, 18.50-22.76 nmoUmin/mg protein for antibody concentrations of 0, 9.8, 25 and 50 yU100 pg microsomal protein, respectively. The range of DOX reduction in humans was 6.81-7.36, 4.63-8.45, 4.09-6.27, 8.45-8.45 nmoYminhng protein for antibody concentrations of 0, 13, 26 and 65 p1/100 pg microsornai protein, respectively. *Significantiy different from CYPRED activity in the absence of antibody, based on randomized design one-way ANOVA and pst-hoc Newman-Keds test @ S 0.05). 3.1.4.3 Carbon monoxide (CO)

Although CYPRED may directly catalyze the reduction of DOX, it may a1so be

indirectly involved in the reaction if the cytochromes P45O catalyze DOX reduction also, as

suggested by Goeptar et al. (1993). Therefore, to distinguish the possible direct or indirect

role of CYPRED in catalyzing DOX reduction, inhibition of the reaction by CO in human

and rat liver microsomes was investigated. The results are presented in Table 4.

II Table 4: Effect of CO on CYPRED Activity and DOX Reduction in Rat and II Human Liver Microsornes

(nmol/min/mg protein) Assay -CO (control) Rat Liver Microsomes CYPRED Activity DOX Reduction Human Liver Microsomes CYPRED Activity DOX Reduction

CYPRED activity was assayed spectrophotometrica1Iy at 550 nrn in 1 ml incubation mixtures consisting of 300 mM potassium phosphate buffer (pH 7.7), 70 pM cytochrome c, 1 mM NADPH and 0.03 (rat) or 0.1 (human) mg microsomal proteidml. DOX reduction was assayed spectrophotometrically at 340 nm in 1 ml incubation mixtures consisting of 50 rnM Tris HCl buffer (pH 7.4) , 5 mM magnesium chloride, 0.5 rnM DTPA, 0.1 rnM DOX, 0.125 mM NADPH and 0.2 (rat) or 0.5 (human) mg microsomal proteidml. Human liver sample KI5 was used for the assays. CO was bubbled into the incubation mixtures for one minute, just prior to the addition of NADPH. Data are presented as the mean * SD of triplicate deterrninations with the exception of DOX reduction in human liver microsomes in which data are presented as the mean f SD of sextuplicate deterrninations. *Significantly different frorn enzyme activity in the absence of CO, based on Student's t-test (p 50.05). 109 In rat and human liver microsomes, CO did not inhibit CYPRED activity, as expected since

CO binds directiy to heme and inhibits cytochromes P450 but not CYPRED. However, CO did inhibit DOX reduction by approximately 36% in rat liver microsomes, suggesting that cytochromes 1.450 may contribute to DOX reduction activity in rat liver. However, in human liver microsomes, CO did not inhibit DOX reduction, which indicates that under these experimental conditions, cytochromes P450 are not major catalysts of DOX reduction. In fact, a consistent small increase in DOX reduction in the presence of CO was observed. The reason for this is unknown.

3.1.4.4 Inhibition of CYPRED activity by DOX

The final inhibition experiment to test the first hypothesis was to determine whether

DOX would inhibit CYPRED activity in rat and human liver microsomes. As shown in

Figure 19, some inhibition of CYPRED activity by DOX did occur with increasing DOX concentrations. Although the extent of inhibition was small and high concentrations of DOX were required, these data do provide additional evidence that DOX interacts with CYPRED. -- " I Rat Liver Microsomes

DOX Concentration (m Ml I Human Liver Microsomes

DOX Concentration (mM)

Figure 19: Inhibition of CYPRED Activity by DOX in Rat (A) and Human (B)Liver Microsomes CYPRED activity was assayed spectrophotometrically at 550 nm in 1 ml incubation mixtures consisting of 50 rnM Tris HC1 buffer (pH 7.4), 30 pM cytochrome c, I mM NADPH and 0.07 (rat) or 0.04 (human) mg microsomal protein/d. Human liver sample K15 was used for the assays. DOX was added to the reaction mixtures at concentrations of 0, 0.2, 0.3, 0.4, 0.5 or 0.6 mM. Each data point is the mean ISD of triplicate detenninations. 100% control activities were 47.43 and 35.52 nmol/rnin/mg protein for rat and human liver microsomes, respectively. *Significantly different from CYPRED activity in the absence of DOX,based on randomized design one-way ANOVA and post-hoc Newman-Keuls test @ 50.05). 3.2 Hypothesis #2: Transfection of Human CYPRED cDNA into Human Breast Cancer Cells Results in Greater Sensitivity to DOX

3.2.1 Cell culture

To test the second hypothesis, the human breast cancer ce11 lines, MCF-7/WT and

MCF-7/ADR, will be used as recipients for transfection of CYPRED cDNA. Therefore,

prior to conducting the transfections, both ce11 lines were characterized in terms of growth

properties and sensitivity to DOX.

3.2.1.1 Growth properties

MCF-7IWT and MCF-71ADR ce11 lines grew in a-MEM supplemented with 10%

FBS, with each ce11 Iine displaying characteristic and unique growth properties (Figure 20).

The MCF-7lWT cells had a faster growth rate than the MCF-7IADR cells. While MCF-

7lWT cells reached confluency at approximately 1.5 x 106 cellslwell, MCF-71ADR celIs

reached confiuency at approximately 2.5 x 10' cellslwell. Under the microscope, it appeared

that while MCF-7lWT cells required very little space for growth (sometirnes growing on top of each other), MCF-71ADR cefls required more space and high confluency cultures were maintained only with frequent changes of growth medium. Also compared to MCF-7/WT cells, MCF-7/ADR cells displayed a lower plating efficiency, as observed by the lower ce11 counts after 24 hours compared to the number of cells initially plated (Figure 20). The doubling times of the ce11 lines were detennined during the exponential growth phase to be

20 and 40 hours for MCF-7/WT and MCF-7lADR ceIls, respectively. 10%

.I

I I-

I

1os

I I -m- MCF-7MT Cells I +MCF-7lADR Cells

Time

Figure 20: Growth Curve of MCF-7lWT and MCF-7IADR CeIl Lines Each cell line was seeded at 1@ cells per well in triplicate in 6 well plates. For each line, five plates were prepared and one plate was counted per day for 5 days. Each data point represents the mean f SD of triplicate determinations. 3.2.1.2 MTT assay

3.2.l.2.l Initial characterization

The MTT assay is an indirect measurement of growth. As opposed to directly

counting the number of living cells, the MTT assay measures the ability of a population of

cells to metabolize a dye (Ailey et al., 1988). Accordingly, a number of preparatory

experiments were conducted to ensure that the end absorbance values of the MTT assay were

relateci to the number of ce1Is present.

The first preparatory experiment was to determine whether MCF-7lWT and MCF-

7/ADR cells metabolize MTT and whether the extent of this metabolism was related to ce11

number. Cells were counted and plated at differing densities in 96-well plates. The plates

were placed in an incubator for 2 hours during which time it was expected that the cells

would adhere to the bottom of the well without significant ce11 replication occurring. MTT

was then added to the wells and the remainder of the assay was conducted as outlined in

Section 2.4.3. Figure 21 illustrates the relationship between ce11 number and absorbance for

the two ce1 lines. For both ceIl lines, insufficient dye is being metabolized below lx104

cells per well to be accurateiy read by the plate reader. However, at ce11 densities greater than 1x10' cells per well, increasing ce11 density resulted in an increase in MTT absorbance.

Therefore, both MCF-7lWT and MCF-7/ADR cells were able to metabolize MTT and the extent of metabolism (as measured by the absorbance) was related to the ce11 number at ce11 densities of lx104 celIs/well or greater. Number of cells per well

Figure 21: Determination of Relationship between Absorbance and Ce11 Number in the MTT Assay Predetennined numbers of MCF-7/WT and MCF-7fADR cells were seeded into 96-well plates and allowed to adhere for 2 hours. MTT dye was added and incubated for 4 hours and then developed according to the standard protocol. Each point represents the mean f SD of duplicate 96-well microplates with sextuplicate determinations per plate. 115 The second preparatory experiment was to determine the growth characteristics of the

two ce11 lines in microculture over the. The cells need to be maintained in microculture for

an adequate period of time, such that cytotoxicity from DOX (which occurs primarily when

cells are replicating) is alIowed to occur. Also, mitochondria require tirne to stop actively

respiring after the point of loss of cellular viability has been reached (Cannichael et al.,

1987). Therefore, it was necessary to determine the maximum amount of tirne cells could be

sustained in culture without medium replacement. Specifically, it was necessary to detennine

optimum seeding density of the cells required to maintain the cells in the exponential growth phase for at least a few days and which would produce high absorbance values

(approximately 1.O units). The high control absorbance values were necessary since dmg treatment would produce lower absorbances that still need to be within the linear detection range of the assay (starting at approxirnately 0.1 units). To detennine these conditions, four densities of cells (2,500; 5,000; 10,000; and 20,000 cells per well) were seeded into quintuplicate 96-well plates and one plate was developed every day over a 5 day period.

Figure 22 illustrates the increasing absorbance readings associated with higher seeding densities and the changes in absorbance over the 5 day culture period. For both MCF-7/WT and MCF-7/ADR cells, the plots indicate the following: 1) the absorbance was associated with ce11 number; 2) within 24 hours of plating, the cells were in an exponential growth phase, the duration of which was dependent on the initial seeding density; and 3) the maximum absorbance readings achieved were approxirnately 1.0 to 1.2 units . For subsequent cytotoxicity assays, it was decided that for both ce11 lines 5,000 cells/well would be seeded, since at this ce11 density the cultures were in an exponential growth phase for four 0.0 : O $0 4b do do 160 740 Time (hours)

0,o ! 1 1 1 I 1 1 O 20 40 60 80 100 120 Time (hou rs)

Figure 22: Assessrnent of Cell Growth in Microculture Over Time MCF-71WT (A) and MCF-71ADR (B) ceIls were plated in quintuplicate 96-weH plates at four densities (2,500; 5,000; 10,000; and 20,000 cells per well). One plate was developed at approximately 24 hour intervals over 5 days. Each data point is the mean i: SD of sextuplicate deterrninations. 117 days and produced high absorbances on the fourth day. The drug (DOX)would be added at

24 hours after plaîhg, which is the beginning of the exponential growth phase, and the MTT assay would be conducted after four days of ce11 growth when peak absorbances occurred.

3.2.1.2.2 DOX toxicity

Prior to stable transfection of hurnan CYPRED cDNA into MCF-7/WT and MCF-

7lADR cells, the toxicity of DOX to untransfected cells was determined (Figure 23). Based on two independent experiments, the MCF-7/ADR ce11 line was found to be 30 to 65-fold more resistant to DOX than the MCF-7IWT line, with the following IC,, values observed:

MCF-7lWT (68 to 72 nM), MCF-7/ADR (2.4 to 4.3 PM). This relative sensitivity of MCF-

7/WT and MCF-7/ADR cells to DOX was essentially unchanged after the cells had been passaged in the absence of DOX for more than a year (data not shown). DOX Concentration (PM)

Figure 23: Sensitivity of MCF-7/WT and MCF-7/ADR Cells to DOX Concentration-response curves for MCF-7/WT and MCF-7/ADR ce11 lines following treatment with DOX. Cells were treated for approxirnately 72 hours with the concentrations of DOX indicated on the abscissa. Cytotoxicity was assessed using a microplate MTT assay. Fractional absorbance is expressed as a percentage of the absorbance displayed by cells not exposed to DOX (see Table 3). Each data point represents the mean f SD of duplicate 96- well microplates with sextuplicate determinations per plate. 119 3.2.2 Construction of a rnammalian expression vector containing human CYPRED

cDNA

The hurnan CYPRED cDNA was cloned into the EcoM site of the phagemid pUC9.

To isolate the cDNA, the pUC9 vector was cut with either EcoRI or EcoRI and ScaI DNA

restriction enzymes (see Figure 24). Digesting the vector with EcoN aIone produced two

bands of very similar sizes: 2.4 kb representing the CYPRED cDNA and 2.7 kb representing the pUC9 vector. To enhance the resolution of the band representing the CYPRED cDNA, a

second restriction digestion reaction was conducted with both EcoRI and ScaI, where ScaI further digested the 2.7 kb vector DNA into 1.8 and 0.9 kb fragments. This allowed easier purification of the cDNA band fiom the gel. To subclone the approximately 2.4 kb

CYPRED cDNA into the EcoRI site of the mamrnalian expression vector pcDNA3.1 (+), a ligation reaction was conducted consisting of the CYPRED cDNA and pcDNA3.1 (+) DNA which was previously linearized by restriction digestion with EcoRI. As shown in Figure 7, the ligation reaction would produce three types of vectors which would allow bacterial growth on LI3 plates with ampicillin. These vectors would be pcDNA3.1 (+) ligated to itself (5.4 kb), pcDNA3.1 (+) ligated with CYPFZED cDNA in the correct orientation (7.8 kb), and pcDNA3.1 ( +) ligated with CYPRED cDNA in the incorrect orientation (7.8 kb). pUC9 with CYPRED cDNA ED Scal 356

Agure 24: Restriction Enzyme Digestion of pUC9 to Isolate the CYPRED cDNA

The restriction digest reaction consisted of 1 pl pUC9 DNA -Lane Tvoe of Restriction Digest Exnected Band Sizes from srna11 scale vector DNA preparations, 1 pl of the Mo appropriate restriction enzyme(s), 4 pl 10x One-Phor-Al1 1 uncut 5.1 (supercoileci) Buffer Plus (Pharmacia) and water for a total volume of 20 pl. - The reactions were incubated ai 37°C for approximately 2.5 2 EcoRI 2.7 (pUC9 DNA) hours. The DNA bands were separated on a 1.5%agarose gel. 2.4 (CYPRED cDNA) Each well was loaded with 12 pl of the reaction mixture. The 3 ScaI digestion reactions conducted and the DNA band sizes expected (as illustrated in the pUC9 map) were as follows: 4 EcoRl + ScaI 2.4 (CYPRED cDNA) 1.8 (pUC9 DNA) 0.9 (pUC9 DNA) 121 Bacterial colonies that were transformed with DNA from the ligation reaction were screened

for the presence of vector containing the cDNA insert. The screening process involved

small-scale preparation of vector DNA and restriction digestion with EcoRI. Those vectors

with the cDNA insert would produce two bands of approximately 5.4 kb (expression vector)

and 2.4 kb (cDNA) which would be visible on DNA gel electrophoresis. If no insert was

present, oniy one band of 5.4 kb would be present. Of the 24 bacterial colonies screened in

this mariner, one colony yielded the 2.4 kb band (data not shown). To further confirm that

the vector in this colony did indeed contain the CYPRED cDNA and additionally to confirrn

that the cDNA was in the correct orientation, fûrther restriction digestion analysis of the

vector was conducted (Figure 25).

From the results of the restriction digestion analysis of the ligated pcDNA3.1 (+), as

shown in Figure 25, the following interpretations were made:

1. cornparison of Lanes 2 and 4 indicate that the ligated vector is approximately 8 kb

when linearized or at least larger than the uniigated vector by approximately 2 kb, as

expected if the CYPRED cDNA was ligated into the vector;

2. digestion of the ligated vector by five restriction endonucleases produced DNA bands

with the sizes expected if the CYPRED cDNA was ligated into the vector; and

3. restriction enzymes (ie., SmaI and StuI, Lanes 7 and 8) which cut the insert

asymmetrically indicated that the cDNA was ligated into the pcDNA3.1 (+) vector in

the correct orientation. Sall

Stul 4453 pcDNA3.1(+) with CYPRED CIMA In Correct Oikntith 7830 bp

Figure 25: Restriction Enzyme Digestion of pcDNA3.1 (+) froin Ligation Reaction with CYPRED cDNA

of 1 pl pcDNA3. I (+) Expccied Band dm The restriction digest reaction consisted Typt of (kb) DNA from small scale vector DNA preparations of colonies Vector Type Restriction ' Lanc Digest Correct Incorrect transformed witli DNA from the ligation reaction or non- Orientation Orientation ligated DNA as a control, 1 pl of the appropriate restriction ' gaicd pcDNA3,1 (+) digest enzyme, 4 or 2 CI 1Ox One-Phor-Al1 Bufier Plus ------supercoiled supcrcoileâ unllgaledpcDNA3.j (+) EcoRj 5.4 5.4 (Pharmacia) and water for a total volume of 20 pl. The , reactions were incubated at 37OC or 30°C for approxhately 3 ligrled pc~~~3.1(+) uncul supercoiled supercoilai 2.5 hours. The DNA bands were separated on a 0.8% agarose ligated pcDNA3.1 (+, BamHI 7.8 7.8 gel. Esch well was loaded with 20 pl of the reaction mixture. 5 ligated pcDNA3.1 (+) EcoRl 5.4, 2.4 The digestion reactions conducted and the DNA band sizes . 5.4, 2.4 expected (as illustrated in the above map) were as follows: 6 ligated pc~~~3.1(+) Sall 5.6, 2.2, 0.034 , 5.6, 2.2, 0.034 7 ligaied pcDNA3.1 (+) SmaI 6.1, 1.7 2.9,4.9 8 ligaied pcDNA3.1 (+) Stul 6.6, 1.2 3.4, 4.4 123 As a final step to confîrm that the correct construct of the expression vector with

CYPRED cDNA was made, a portion of the ligated expression vector was sequenced using a

T7 priming site, which is just upstream of the multiple cloning site in the expression vector

(Figure 9). In Figure 26, the results of the sequencing reaction along with the entire sequence of the human CYPRED cDNA as reported by Yamano et al. (1989) are presented.

Approximately 200 bp of the CYPRED cDNA sequence were confirmed by the sequencing reaction. The orientation of the insert was also confirmed to be correct, since the 5' end of the confinned sequence corresponded to the 5' end of the originally cloned CYPRED cDNA

(Yamano et al. , 1989).

Therefore, based on the restriction digestion analysis and the DNA sequencing, it was concluded that the correct DNA construct of the expression vector containing the human

CYPRED cDNA was made. This constnrct wiI1 be transfected into MCF-7/WT and MCF-

7/ADR cells to test if elevated CYPRED activity can sensitize cells to the cytotoxic effects of

DOX. Figure 26 Sequence of human CYPRED cDNA and the Portion of the Sequence Identifid by DNA Sequenchg The expression vector ligated with human CYPRED cDNA was sequenced using the Sanger-Coulson method of chah-terminatingnucleotides and the Phannacia Biotech T7SequencingKit. A ïï primer was used to start the sequence at the T7 prirning site (ie., TAATACGACTCACTAT)located just upstream of the EcoRI site into which the human CYPRED cDNA was subcloned. The complete sequence of the CYPRED cDNA as reported by Yamano et al. (1989) is presented and the portion of the sequence that was confirmed is underlined below the Yamano et al. (1989) sequence. S'end CYPREDcDNA atgatcaaca tgggagactc ccacgtggac accagctcca ccgtgtccga ggcggtggcc - ccacataaac accaactsca rcatatccaa aacaatamc 61 gaagaagtat ctcttttcag catgacggac atgattctgt tttcgctcat cgtgggtctc ac atnattctat tttcactcat catg.gfTtct€

121 ctaacctact ggttcctctt cagaaagaaa aaagaagaag tccccgagtt caccaaaatt a eccccaaat t cawaatt

181 cagacattga cctccectgt cagagagagc agctttgtgg aaaagatgaa gaaaacgggg ac aactttataa aaggaataa? çLaaaa2

241 aggaacatca tcgtgttcta cggctcccag acggggactg cagaggagtt tgccaaccgc

301 c tgtccaagg acgcccaccg c tacgggatg cgaggcatgt cagcggaccc tgaggagtat

36lgacctggccg acctgagcag cctgccagag atcgacaacg ccctggtggt tttctgcatg

421gccacctacg gtgagggaga ccccaccgac aatgcccagg acttctacga ctggctgcag

481 gagacagacg tggatctctc tggggtcaag t tcgcggtgt t tggtcttgg gaacaagacc

541 tacgagcact tcaatgccat gggcaagtac gtggacaagc ggctggagca gctcggcgcc

601 cagcgcatct ttgagctggg gttgggcgac gacgatggga acttggagga ggact tcatc

661 acc tggcgag agcagt tctg gccggccgtg tgtgaacact t tggggtgga agccactggc

72lgaggagtcca gcattcgcca gtacgagctt gtggtccaca ccgacataga tgcggccaag

781 gtgtacatgg gggaga tggg ccggctgaag agctacgaga accagaagcc cccctt tgat

84lgccaagaatc cgttcctggc tgcagtcacc accaaccgga agctgaacca gggaaccgag

901 cgccacc tca tgcacctgga a t tggacatc tcggactcca aaa tcaggta tgaatc tggg

961gaccacgtgg ctgtgtaccc agccaacgac tctgctctcg tcaaccagct gggcaaaatc

1021 ctgggtgccg acctggacgt cgtcatgtcc ctgaacaacc tggatgagga gtccaacaag 108laagcacccat tcccgtgccc tacgtcctac cgcacggccc tcacctacta cctggacatc Figure 26 Sequence of human CYPRED cDNA and the Portion of the Sequence Identifid by DNA Sequencing (cont'd)

114laccaacccgc cgcgtaccaa cgtgctgtac gagctggcgc agtacgcctc ggagccctcg

1201 gagcaggagc tgc tgcgcaa ga tggcctcc tcc tccggcg agggcaagga gctgtacc tg

1261agctgggtgg tggaggcccg gaggcacatc ctggccatcc tgcaggactg cccgtccctg

1321cggcccccca tcgaccacct gtgtgagctg ctgccgcgcc tgcaggcccg ctactactcc

1381 a tcgcctcat cctccaaggt ccaccccaac tctgtgcaca tctgtgcggt ggt tgtggag

1441 tacgagacca aggccggccg catcaacaag ggcgtggcca ccaactggct gcgggccaag

1501 gagcctgccg gggagaacgg cggccgtgcg ctggtgccca tgttcgtgcg caagtcccag

1561ttccgcctgc ccttcaaggc caccacgcct gtcatcatgg tgggccccgg caccggggtg

162lgcacccttca taggcttcat ccaggagcgg gcctggctgc gacagcaggg caaggaggtg

1681 ggggagacgc tgctgtac ta cggc tgccgc cgc tcagatg aggactacct gtaccgggag

174lgacgtggcgc agttccacag ggacggtgcg ctcacccagc tcaacgtggc cttctcccgg

180lgagcagtccc acaaggtcta cgtccagcac ctgctaaagc aagaccgaga gcacctgtgg

1861aagttgatcg aaggcggtgc ccacatctac gtctgtgggg atgcacggaa catggccagg

1921gatgtgcaga acaccttcta cgacatcgtg gctgagctcg gggccatgga gcacgcgcag

1981gcggtggact acatcaagaa actgatgacc aagggccgct actccctgga cgtgtggagc

2041taggggcctg cctgccccac ccaccccaca gactccggcc tgtaatcagc tctcctggct

2101 ccctcccgta gtctcctggg tgtgtttggc ttggccttgg catgggcgca ggcccagtga

216lcaaagactcc tctgggcctg gggtgcatcc tcctcagccc ccaggccagg tgaggtccac

2221 cggcccctgg cagcacagcc cagggcctgc atgggggcac cgggctccat gcctctggag

2281gcctctggcc ctcggtggct gcacagaagg gctctttctc tctgctgagc tgggcccagc

2341ccctccacgt gatttccagt gagtgtaaaL aattttaaat aacctctggc ccttggaa 3' end 126 3.2.3 Optimization of antibiotic concentration for selection of stable transfectants

Prior to transfection of the ce11 lines with the expression vector containing human

CYPRED cDNA, antibiotic (geneticin) selection conditions for each parental ce11 line were detennined. Ce11 growth was assessed by the presence and density of methylene blue-stained colonies. Figure 27 presents the relative density of the stain when cells were grown in the presence of various concentrations of geneticin. As shown in Figure 27 (panel B), for the

MCF-7lADR cells, staining intensity was essentially reduced to background levels by a geneticin concentration of 400 pglml. Also, no colonies were visible upon microscopic examination at this dose. Therefore, 400 pglml was considered to be the minimum concentration of geneticin that resulted in complete ce11 death of MCF-7IADR cells. For the

MCF-7lWT cells, staining intensity was reduced to background levels by a geneticin concentration of 600 pglml (Figure 27, panel A); however upon microscopic examination, a few small colonies were visible. Therefore, 800 pglml was considered to be the minimum concentration of geneticin that resulted in complete ce11 death of MCF-7/WT cells. These geneticin concentrations were to be used to select stable transfectants. A MCF-7M Cells '1 MCF-7lADR Cells

Geneticin Concentration Geneticin Concentration @@ml) l (cLs/ml)

Figure 27: Optirnization of Antibiotic Concentration for Selection of Stable Transfectants To determine the optimal antibiotic concentration to use when establishing and selecting for a stable ce11 line, untransfected MCF-7/WT and MCF-7lADR cells were seeded in triplicate at 10,000 cellslwell in 6-well tissue culture plates containing medium with geneticin at concentrations of 0, 200, 400, 600, 800 or 1000 pglml. The cells were incubated for 6 days and were fed with the selective medium every few days as necessary. The plates were then stained with a dye containing 0.5% (wlv) methylene blue and 50% (vlv) methanol and the degree of cell growth was assessed by the presence and density of the blue stain. The relative density of the stain was quantitated by scaming the culture plates using an UMAX SuperVista S-12 scanner with the aid of the program Adobe Photoshop v4.0 for Power Macintosh. Quantitative analysis was performed using the program IPLab Gel Scientific Image Processing v1 .Se for Power Macintosh (Signal Analytics, Vienna VA). Based on ce11 density and microscopic examination, the minimum concentration of geneticin that resulted in compiete.cel1 death and that was to be used to select stable transfectants was 800 and 400 pglml in MCF-7/WT and MCF-7lADR cells, respectively. Each point represents the mean ISD of triplicate determinations. 4. Discussion

4.1 Hypothesis #1 - Role of Human CYPRED in DOX Bioactivation

Table 5 presents the standard strategy required to demonstrate in vitro that a specific enzyme catalyzes the biotransformation of a dmg (Friedberg and Wolf, 1996; Guengerich,

1996; Meyer, 1996).

Table 5: Methods for Determining the Role of an Enzyme in the Biotransformation of a Compound II 1. Detection of biotransformation reaction with purified enzyme or in a heterologous II expression system 2. Detection of biotransformation reaction in appropriate tissue (eg ., human liver microsomes) Inhibition of biotransformation by chemical and antibody inhibitors II 3* 4. Correlation of biotransformation reaction with enzyme activity or protein expression in a collection of human livers

(Friedberg and Wolf, 1996; Guengerich, 1996; Meyer, 1996)

The strength of the data presented in this thesis relates to the use of these multiple approaches to show that human CYPRED catalyzes the reduction of DOX. DOX reduction was detected in a heterologous expression system (microsomes from human lyrnphoblastoid cells overexpressing human CYPRED), which suggests that the human CYPRED enzyme has the inherent capacity to carry out the reaction (Figure 14). However, this approach does not indicate the importance of CYPRED in the overall metabolism of DOX, particularly in relation to other enzymes that occur in drug-metabolizing tissues. As such, the second approach was used in which DOX reduction was shown to occur in hurnan liver microsomes. 129 To verify that CYPRED in the microsomes was catalyzing the reaction, a third approach was used in which a series of inhibition studies was conducted, including inhibition by a chemical inhibitor (DPIC) and an antibody inhibitor.

DPIC is a mechanism-based chemical inhibitor of CYPRED. The proposed mechanism of action of DPIC is as follows (Tew, 1993; OYDonnellet al., 1994). In the presence of NADPH, CYPRED reduces DPIC to a reactive phenyl radical, which covalently binds to either the flavin cofactor or adjacent amino acid side chains important in catalysis.

Inhibition of CYPFED activity is irreversible and is both the and concentration-dependent

(Tew, 1993). Additionally, DPIC is a potent inhibitor of CYPRED activity with inhibition occurring at concentrations of 0.2 mM (Tew, 1993). This is sirnilar to Our results in which

CYPRED activity was inhibited at 0.16 rnM. In terms of its specificity, studies indicate that it inhibits only those flavin enzymes that catalyze one-electron reduction, such as CYPRED and NADPH oxidase, but not glutathione reductase or glucose oxidase which transfer two electrons during catalysis (OYDomeiiet al., 1994). Our results show that DPIC inhibited

CYPRED activity and DOX reduction in the human liver microsomes in a concentration- dependent manner (Figure 17). This suggests that in the microsomes, CYPRED was catalyzing DOX reduction. Although DPIC may inhibit other flavoenzymes that might possibly also catalyze DOX reduction (ie., NADH dehydrogenase, xanthine oxidase and nitric oxide synthase), these enzymes are not located in the endoplasmic reticulum of cells.

In terms of antibody inhibition, DOX reduction was not inhibited in the human liver microsomes by a polyclonal antibody against CYPRED (Figure 18). Although in rat liver microsomes, inhibition by the antibody appeared to be occurring at the lower antibody 130 concentrations, inhibition at the highest concentration was not detected (Figure 18). Since

the antibody clearly inhibited CYPRED catalytic activity toward cytochrome c (Figure 18), it

is not known why inhibition of DOX reduction by the antibody did not occur. One possible

reason is that for the antibody protein concentrations to be achieved in the assays,

particularly with the human sarnples and the rat sample with the highest antibody

concentration, large volumes of antibody solution were required which may have diluted the

reactions to such a degree that adequate interaction of antibody to microsomal protein did not

occur. It has been reported in other investigations, that demonstration of inhibition of an

enzyme by monoclonal or polyclonal antibodies is relatively difficult with poor reproducibility (Meyer, 1996).

Another possible reason for the observed antibody inhibition of cytochrome c reduction, but not DOX reduction, may be due to the presence of multiple substrate binding sites on the CYPRED protein (see Figure 6). For example, in other studies, specific antipeptide antibodies to regions of CYPFED inhibited P4501A1 and P4502Bl activities, but not cytochrome c reduction, indicating differences in the interaction of cytochromes P450 and cytochrome c with CYPRED (Strobel et al., 1995). The polyclonal antibody used in Our experirnents inhibited cytochrome c reduction and is also reported by the suppliers (Gentest) to inhibit reduction of P450s. However, it is not known exactly where DOX interacts with

CYPRED. Perhaps DOX interacts with CYPRED at a site not inhibited by the antibody (ie., not the cytochrome c site nor the cytochrome P450 site) which may possibly explain why inhibition of DOX reduction by the antibody did not occur.

In another inhibition assay, the activity of CYPRED was shown to be inhibited by DOX in the human liver microsomes (Figure 19), which may suggest that DOX and cytochrome c interact with the same enzyme. However, since relatively high concentrations of DOX (0.2 to 0.5 mM) were required and only partial inhibition of CYPRED activity was observed, this might also indicate that cytochrome c and DOX interact with CYPRED at different sites and only at high concentrations can inhibition be observed. The high concentration of DOX required in this assay may not be clinically relevant, since the substrate used to measure CYPRED activity, cytochrome c, is an artificial non-physiologie substrate for CYPRED.

Assuming that the antibody experiments are technically Sound, then to understand why

DOX reduction was not inhibited by the antibody and also to elucidate the interaction of

DOX and cytochrome c with CYPRED, a number of further experiments may be conducted.

Firstly, the antibody experiments couId be repeated using CYPRED antibodies obtained from other sources. Secondly, to determine whether DOX and cytochrome c interact at different sites on the enzyme, classical enzyme kinetic studies could be conducted in which CYPRED activity (ie., the rate of cytochrome c reduction) is measured as a function of cytochrome c concentration in the absence and the presence of increasing concentrations of DOX. If DOX and cytochrome c are competing for the same site on the enzyme, then the maximal velocity of cytochrome c reduction (V-) would not change with increasing concentrations of DOX; however, the Michaelis constant (KJ for cytochrome c (ie., the concentration of cytochrome c that gives 50% of the maximum rate of the reaction) would increase with increasing concentrations of DOX. If DOX and cytochrome c are not competing for the same active site, then the V,, of cytochrome c reduction would decrease with increasing concentrations of DOX, while the Km values for cytochrome c would remain the sarne. Thus further experimentation is required to clariQ the interaction of DOX with CYPRED,particularly in relation to cytochrome c.

In the fourth approach to determine whether human CYPED catalyzes DOX reduction, a statistically significant correlation was shown between DOX reduction and both

CYPRED activity and protein expression using the microsomes from a bank of 17 hurnan livers (Figures 15 and 16). With the exception of one study in which relatively low levels of CYPRED activity in a human liver bank were reported and which varied 8-fold (Placidi et al., 1993), our results agree with those of other investigators who reported very little variation (ie., less than 3-fold) in the range of activity, protein expression and mlWA levels of CYPRED in hurnan populations (McManus et al., 1987; Yarnano et al., 1989; Shephard et al., 1992; Pearce et al., 1996). In practice, it is difficult to establish correlations if the range of activities is less than 5-fold (Guengerich, 1996). However, we were able to show a correlation in spite of the fact that CYPRED activity and DOX reduction only varied 2.4-fold and 4.4-fold, respectively .

Although CYPRED may directly catalyze the reduction of DOX, it may also be indirectly involved in the reaction if the cytochromes P450 catalyze DOX reduction also.

Therefore, to distinguish the possible direct or indirect role of CYPRED in catalyzing DOX reduction, inhibition of the reaction by CO in human and rat liver microsornes was investigated (Table 4). In rat liver microsornes, DOX reduction was inhibited by approximately 36%, which suggests that cytochromes P450, in addition to CYPRED catalyze

DOX reduction in rats. This is in agreement with another study in which rat CYP2BI and 133 CYPRED were shown to make approxirnately equal contributions to DOX reduction in the

liver microsomes of phenobarbital-treated rats (Goeptar et al., 1993). AIso in rats, DOX

reduction was inhibited by only 60% in the presence of the highest concentration of the

chernical inhibitor (DPIC), cornpared to 80 % inhibition of CYPRED activity . This suggests that enzymes in addition to CYPRED were catalyzing DOX reduction in the rats. In contrast to this, in the human liver microsomes, DOX reduction was not inhibited by CO, which suggests that the cytochromes P450 were not major catalysts of DOX reduction. It will be of interest to determine whether human cytochromes P450 play a more important role in

DOX reduction under conditions of reduced oxygen tension.

Therefore, based on a combination of complementary approaches, these results indicate that human CYPRED catalyzes the reduction of DOX under aerobic in vitro conditions. Although, DOX reduction has been shown to occur in the presence of rat or rabbit liver rnicrosomes and with purified rat or rabbit CYPRED (Handa and Sato, 1976;

Goodman and Hochstein, 1977; Bachur et al., 1978; 1979; Kharasch and Novak, 1981;

1983; Komiyama et al. , 1986; Vile and Winterbourn, 1989; Goeptar et al., 1993), to my knowledge this is the first demonstration that DOX reduction is catalyzed by human

CYPRED. Compared to rats, however, the results indicate that the catalytic rate of DOX reduction is approximately 2- to 5-fold Iower in human liver microsomes.

A limitation of these results is the fact that DOX reduction was rneasured indirectIy by NADPH oxidation. Therefore, it is not entirely certain that DOX metabolism by human

CYPRED results in the production of the semi-quinone free radical. NADPH oxidation has been used routinely by others in animal models to measure DOX reduction, and in some 134 studies the rate of DOX-stimulated NADPH oxidation has been used as a simple surrogate for measurement of the DOX semi-quinone radical by ESR spectroscopy under anaerobic conditions. It is assumed that Our experiments using human and rat liver rnicrosomes are comparable in this regard to the other animal studies and that the semi-quinone was produced. However, it would be ideal to directly confirrn that human CYPRED biotransforrns DOX to the semi-quinone radical.

DOX can act as a substrate for a number of cellular enzymes. In addition to

CYPRED, reductive activation of DOX to the DOX semi-quinone has been reported to be catalyzed by xanthine oxidase, NADH dehydrogenase, ferredoxin reductase, NADH- cytochrome b, reductase and nitric oxide synthase (Pan and Bachur, 1980; Davies et al.,

1983; Doroshow, 1983a; 1983b; Rowley and Halliwell, 1983; Komiyarna et QI., 1986;

Hodnick and SartorelIi, 1994; 1998). In addition, DOX can be reduced to less toxic metabolites by carbonyl reductase and NAD(P)H-quinone oxidoreductase (DT-diaphorase)

(Robert and Gianni, 1993; Kasahara et al., 1994). We are fairly confident that in the human liver microsomes, CYPRED was catalyzing DOX reduction. The following enzymes are located in the cytoplasm or mitochondria and would not be expected to significantly contribute to rnicrosomal DOX reduction: xanthine oxidase, NADH dehydrogenase, ferredoxin reductase , carbonyl reductase and NAD(P)H-quinone oxidoreductase . Nitric oxide synthase is found primarily in macrophages and endothelium as opposed to hepatocytes. NADH-cytochrome bS reductase is located in the endoplasmic reticulum and may have catalyzed DOX reduction along with CYPRED; however, the extent of its contribution is expected to be lower than that of CYPRED for two reasons. Firstly, it prefers NADH as a cofactor compared to CYPRED which prefers NADPH. Secondly,

NADH-cytochrome b, reductase has been reported to catalyze DOX reduction at pH 6.6, but

not at pH 7.6 (Hodnick and Sartorelli, 1994), which is close to the pH (pH 7.4) of the buffer

used in our assays.

Although CYPRED-catalyzed reduction of DOX was shown to occur in human liver

microsomes, it needs to be detemined whether this reduction would occur in a cellular system, particularly in human tumour ceIIs, and whether increased expression of human

CYPRED would result in increased sensitivity of the tumour cells to DOX. This is considered in the second hypothesis.

4.2 Hypothesis #2 - Sensitization of Human Breast Cancer Cells to DOX by

Overexpression of CYPRED

Several studies have shown that one-electron reduction of DOX results in the formation of the DOX semi-quinone and subsequent redox-cycling leading to the formation of the hydroxyl radical which in turn leads to oxidative damage and cytotoxicity. This has been shown to be a significant mechanism of toxicity in cardiac tissue. However, whether DOX- induced free radical production is a significant mechanism of toxicity in cancer cells is uncertain. Other studies indicate that another unidentified metabolite of aerobic CYPRED- catalyzed DOX reduction may cause cytotoxicity through covalent binding to DNA and proteins (Ghezzi et al., 1981; Scheulen et al. , 1982; Wallace and Johnson 1987; Bartoszek and Wolf, 1992). Therefore DOX reduction as catalyzed by CYPRED results in the formation of hydroxyl radicals and/or reactive metabolites that bind to macromolecules.

The purpose of the second component of this research project was to determine 136 whether increased expression of human CYPRED in breast cancer cells would increase their

sensitivity to DOX. In addition, this mode1 would allow one to elucidate the possible role of

free radicals or other possible metabolites in the cytotoxicity of DOX in breast cancer cells.

Increased expression of human CYPRED in breast cancer cells (MCF-71WT and

MCF-7/ADR) was to be accomplished through stable transfection of the human CYPRED cDNA. Prior to doing this, however, bot. cell lines were characterized in terms of growth and cytoxiciîy to DOX. The doubling times of the ce11 lines were 20 and 40 hours for MCF-

7/WT and MCF-7lADR cells , respectively (Figure 20). Additionally , MCF-7lADR cells were approxirnately 30 to 65-fold more resistant to DOX than MCF-7IWT cells, as deterrnined by the MTT microplate colorimetric assay (Figure 23). The initial report on the

MCF-7/ADR ce11 line indicated that this line was 192-fold more resistant to DOX than the

MCF-7lWT cells, as determined by a ce11 counting assay (Batist et al., 1986). As compared to our IC,, values of 68 to 72 nM (MCF-7!WT) and 2.4 to 4.3 PM (MCF-7lADR), the original study reported IC,, values of 25 nM (MCF-7lWT) and 4.8 pM (MCF-7/ADR)

(Batist et al., 1986). There are two factors that likely contribute to the differences between our results and the original investigation. First, differences in absolute IC,, values may be due to the fact that sensitivity of MCF-7-derived ce11 lines to DOX is often lower in MTT assays than in assays based on cell number or clonogenicity (Chen and Waxrnan, 1995a).

Second, the decreased relative resistance of the MCF-7lADR ce11 line in Our study may be due to the fact that we had passed this ce11 line in DOX-free medium for about six months prior to perfonning the cytotoxicity assays. When MCF-71ADR cells are passed for prolonged periods in drug-free medium, these cells dispIay increased sensitivity to DOX 137 along with decreased expression of P-glycoprotein (Budworth et al., 1997). Our results are

similar to hivo studies that found that passage of MCF-7lADR cells in drug-free medium for

48 to 60 weeks resulted in a cell line that was 50 to 68-foId more resistant to DOX than

MCF-7lWT ceils (Batist et al. , 1986; Fairchild et al, , 1987).

For subsequent transfections, a marnrnalian expression vector containing the hurnan

CYPRED cDNA was constructed and the sequence of a portion of the insert was deterrnined.

The expression vector containing the insert was pcDNA3.1 (+) (Invitrogen) which was chosen due to its reported ability to produce stable clones in mammalian cells with relatively high expression of the insert which is controlled by the cytomegalovirus promoter. The actual transfection procedure, antibody selection of stable transfectants, screening of ce11 lines for increased CYPRED expression, and cytotoxicity assays will be conducted by others in the laboratory and is not included as part of this thesis. The goal is to isolate and characterize transfectants derived frorn both the MCF-7/WT and MCF-7lADR cells that express varying levels of CYPRED activity.

When the breast cancer ce11 lines with increased expression of liurnan CYPRED are available, there may be a number of possible outcornes of the cytotoxicity assays. Firstly, in both transfected cell lines (MCF-7/WT/RED and MCF-7/ADR/RED), cytotoxicity to DOX may be increased compared to cells transfected with vector alone (MCF-7/WT/VEC and

MCF-7lADRNEC). This would suggest that in these cells, CYPRED-catalyzed DOX reduction led to the increased cytotoxicity. Addition of a CYPRED inhibitor (eg., DPIC) should decrease the sensitivity of the CYPRED-overexpressing cells to that of the vector- transfected counterparts. On the other hand, increased cytotoxicity may not be observed in both ce11 lines with increased hurnan CYPRED expression. This rnay be because the

sensitivity of the cytotoxicity assay to be used (MTT assay) rnay not be sufficient to detect

small changes (likely less than 5-fold) in sensitivity to DOX. In that case, a more sensitive

assay that is able to detect several orders of magnitude of ce11 killing, such as the colony

forming assay, will be required. Another reason for lack of increased cytotoxicity in the

cells with increased CYPRED expression, is sirnply that increased DOX reduction will not

result in increased cytotoxicity. Perhaps the Ievel of expression of CYPRED wili not be high

enough to produce significant quantities of DOX metabolites leading to cytotoxicity. Or

perhaps the levels of antioxidant defenses in the cells will be able to counteract the increased

levels of DOX metabolites and free radicafs. Another reason is that although significant

DOX reduction rnay be occurring, the formation of metabolites and free radicals simply have

no effect on ce11 survival due to the subcellular location of metabolism (smooth endoplasmic

reticulum) in relation to critical cellular targets (such as the nucleus and plasma membrane).

A final interpretation of these data would be that CYPRED-catalyzed DOX metabolism does

not contribute significantly to DOX cytotoxicity in MCF-7cells. A third possible outcome

of the cytotoxicity assays in the transfected ce11 lines is that increaied cytotoxicity will occur

in the transfected wildtype ce11 line (MCF-7/WTIRED), but not in the transfected DOX-

resistant ce11 line (MCF-7/ADR/RED). This rnay simply be due to fower levels of CYPRED

expression in MCF-7lADFURED cells compared to MCF-7/WT/RED cells. Or, more likely,

increased expression of CYPRED rnay have no effect due to the mechanisms of resistance known to be present in the MCF-7/ADR cells. For exarnple, there rnay be decreased cellular accumulation of DOX due to the presence of P-glycoprotein which serves as a DOX efflux 139 pump. Or perhaps, as reported in several studies, the resistant cells express higher levels of

GSHPx which would inhibit the formation of the cytotoxic hydroxyl radicals.

Overexpression of CYPRED in the MCF-7lADR ce11 line may help to determine the role that

processes related to DOX redox-cycling play in the resistance of this ce11 line. In addition, it

will be interesting to determine whether CYPRED overexpression can overcome other drug

resistance mechanisms (P-glycoprotein, GST-pi, GSHPx) that are known to CO-existin MCF-

7IADR celIs.

4.3 Future Research Directions

Assuming that increased expression of human CYPRED results in increased

sensitivity to DOX in at least one of the breast cancer ce11 lines, a number of future experirnents should be conducted to elucidate the mechanism of CYPRED-dependent toxicity.

There appear to be two general mechanisms of toxicity from CYPRED-generated metabolites of DOX under aerobic conditions. These are the redox-cycling mechanism of toxicity and toxicity from the covalent binding of a DOX metabolite to DNA and protein. To detemine whether DOX cytotoxicity is dependent on redox-cycling, cytotoxicity assays should be conducted with the transfected ce11 lines in the presence of free radical scavengers or antioxidant enzymes. If redox-cycling is a significant mechanism of toxicity, then cytotoxicity should decrease in the presence of free radical scavengers or antioxidant enzymes. Although superoxide dismutase, catalase and GSHPx are ideal antioxidant enzymes to use, they cannot traverse the plasma membrane. However, certain iron chelators and hydroxyl radical scavengers do cross the plasma membrane. Also, several processes associated with redox-cycling, such as lipid peroxidation and hydrogen peroxide formation, can be measured and compared to rates in the control ce11 line. If redox-cycling is an

important mechanism of toxicity, we would predict that the CYPRED-overexpressing cells

should show greater production of reactive oxygen species and lipid peroxidation.

To determine whether cytotoxicity is due directly to a CYPRED-generated DOX

metabolite, covalent binding of radiolabeled DOX to proteins and DNA in a whole ce11

system should be assessed using the methods of Bartoszek and Wolf (1992). The degree of

binding should be compared in transfected and control ce11 lines (ie., vector transfected ce11

line). The effect of glutathione (which rnay interact with the reactive metabolite) on the

degree of DOX binding to protein or DNA should also be assessed. However, since

glutathione does not pass the plasma membrane, glutathione precursors (eg., glutathione

diethyl ester or N-acetylcysteine), which do pass the plasma membrane may be used.

If increased cytotoxicity is not observed in the resistant transfectant cells (MCF-

7/ADR/RED), then the biochemical differences between the sensitive (MCF-7/WT/RED) and resistant transfectants may be determined. Dmg resistance is a multifactorial process and although the P-glycoprotein is prirnarily associated with drug resistance in MCF-7/ADR cells, this does not preclude other biochemical rnechanisms of resistance that rnay affect the redox-cycling or reactive metabolite mechanisms of action of DOX. Possible biochemicaI parameters to assay for include the level of P-glycoprotein and DOX efflux, glutathione,

GSHPx, GSTs, superoxide dismutase and catalase.

Although sensitivity to DOX may be increased in cells with increased CYPRED expression in vitro, it remains to be seen whether this effect will also be observed in vivo.

As such, another future experiment invoIves transplanting the CYPRED-transfected breast cancer cells into an immune-incompetent nude mouse model. Once the tumours have reached a standard size, DOX would be administered at therapeutic doses and the extent of turnour regression measured. One would compare the response of tumours derived from

CYPRED-transfected celk to tumours derived from vector-transfected cells. To assess whether the tumour regression (if observed) is due to DOX redox-cycling, the effect of iron chelators in these animals can also be evaluated. Iron catalyzes the formation of hydroxyl radicals from superoxide and hydrogen peroxide, and the presence of iron chelators would be expected to reduce hydroxyl radical formation and its associated cytotoxicity, thereby reducing the extent of tumour regression.

These breast cancer ce11 lines overexpressing CYPRED can also be used to evaluate in cancer cells the role of CYPRED in the cytotoxicity of other bioreductive antineoplastic drugs, such as mitomycin C, tirapazamine, and other dmgs being developed (eg., indoloquinone E09, diaziquone, streptonigrin). This model would be particularly useful if the cytotoxicity assays were conducted under hypoxic conditions, since these drugs are being developed primarily for hypoxic tumour cells which occur in solid tumours in vivo and which have displayed resistance for radiation and standard chemotherapy.

4.4 Summary and Implications of Research

The main purpose of this research project was to conduct a series of preparatory experiments prior to modulating CYPRED expression in human breast cancer ce11 lines to increase their sensitivity to DOX. The first preparatory experiments were to determine whether DOX reduction was catalyzed by hurnan CYPRED. Using multiple approaches, Our results lead us to conclude that human CYPRED catalyzes the reduction of DOX, most likely 142 to a reactive intermediate which rnay contribute to DOX toxicity. The second preparatory

experiments were to characterize DOX-sensitive and resistant breast cancer ce11 lines (MCF-

7/WT and MCF-7/ADR) in terms of growth and DOX cytotoxicity. The resistant ce11 line

was approxirnately 30 to 65-fold more resistant to DOX compared to the sensitive ce11 Iine

and had a much slower growth rate. The final preparatory experiments involved the construction of a marnmalian expression vector containing human CYPRED cDNA which will be used for subsequent transfection experiments.

Following the transfection of human CYPRED cDNA into the breast cancer ce11 lines, one can determine whether CYPRED-catalyzed DOX reduction will sensitize the cells to

DOX and one can also elucidate the mechanism of CYPRED-dependent DOX toxicity. For example, is redox-cycling of DOX a significant mechanism of toxicity in cancer cells?

The fmdings of this study and studies to be conducted subsequently are significant for a nurnber of reasons. These studies may provide evidence that modulation of CYPRED expression in breast cancer cells can sensitize the cells to DOX in vitro and will set the stage for determining whether this will also occur in vivo. This suggests a possible future treamient of breast cancer in which more selective tumour ce11 kill may be achieved by combining conventional chemotherapy with drug-metabolizing enzyme gene therapy. In this manner, drug bioactivation may be seIectively enhanced within tumour cells, leading to a requirement for lower drug doses, decreased systemic toxicity, and improved therapeutic effect. 5. References

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Abstracts

1. S. Ramji, T. Inaba, and D.S. Riddick. Bioactivation of doxorubicin by human NADPH-cytochrome P450 reductase. Fifth Annual Symposium with University of Toronto Drug Safety Research Group - Drug Information Association (1998). (Toronto, Ontario; February , 1998).

2. S. Ramji, T. Inaba, and D.S. Riddick. Bioactivation of doxorubicin by human NADPH-cytochrome P450 reductase. FASEB Journal 12: A147 (1 998). (Experimental Biology '98; San Francisco, California; April 1998).

3. S. Ramji, T. Inaba, and D.S. Riddick. Bioactivation of doxorubicin by human NADPH-cytochrome P450 reductase. Visions in Phannacology, p. 37 (1998). (Toronto, Ontario; May, 1998). hblications

1. K. Wang, S. Ramji, A. Bhathena, C. Lee, and D.S. Riddick. Glutathione S-transferases in wild-type and doxorubicin-resistant MCF-7 human breast cancer ce11 lines. Xenobiotica, In Press, October, 1998. APPENDIX A List of Chernicals and Reagents With Their Sources

Chemical 1 source acetic acid (glacial) Mallinckrodt acrylamide BioShop agarose 1 GibcolBRL ------ammonium persulfate ICN ampicillin Sigma bac to-agar Difco bacto-tryptone 1 Difco bacto-y east extract Difco boric acid 1 BDH bovine semm albumin Sigma bromophenol blue Bio-Rad carbon rnonoxide (CO) Linde chloroforrn Mallinckrodt cupric sulfate pentahydrate Anachemia - - --- cytochrome c Sigma diethylenetriaminepentaacetic acid (DTPA) Sigma dimethyl sulfoxide (DMSO) Caledon diphenyliodonium chloride (DPIC) Aldrich doxorubicin hydrochloride Sigma ethanol (1 00 %) Commercial Alcohols ethanol (95%) Materials Distribution Centre (University of Toronto) ethidium bromide Sigma ethylenediaminetetraacetic acid (EDTA) BDH fetal calf serum GibcoIBRL Source Pharmacia Biotech ------Sigma Gibco/BRL BDH glycerol ACP chernicals glycine BioShop ' HEPES Sigma LN-(2-hydroxyethyllpiperazine-N'-(2- ethanesulfonic acid)] hydrochloric acid J.T. Baker magnesium chloride Caledon P-mercaptoethanol Bio-Rad methanol Caledon 1 N ,N'-methylenebis acrylamide (BIS) Bio-Rad methylene blue Bio-Rad - . -- a-minimal essential medium (a-MEM) University of Toronto Media Preparation Service MTT 13-(4,5-dimethylthiazoI-2-y1)-2,5- Sigma diphenyltetrazolium bromide] NADPH (nicotinamide adenine dinucleotide Sigma phosphate - reduced form) potassium acetate Fisher -- 10X One-Phor-Al1 Buffer Plus Pharmacia Biotech Sigma phosphate buffered saline (PBS) University of Toronto Media Preparation Service

pp -- Ponceau S stain Sigma 1 -- - potassium chlonde @Cl) J.T. Baker 1 -- 1 potassium phosphate (dibasic) BDH 11 Chemical 1 source potassium phosphate (monobasic) Anachernia skim milk powder 1 Carnation I sodium acetate BDH sodium carbonate BDH - sodium chloride BDH sodium dodecyl sulfate (SDS) Caledon sodium hydroxide (NaOH) Fisher sodium potassium tartrate tetrahydrate Sigma sucrose BDH N,N, N' ,N' -tetramethylethyienediamine Mallinckrodt (TEMED) thimerosal Sigma Tris [Tris(hydroxymethy1)amino- methane] Sang on ------WPS~~ Difco Tween 20 Sigma urea BDH xylene cyan01 FF Bio-Rad APPENDM B Characteristics of Liver and Kidney Donors

Date Obtained Physical Description Cause of Known Drug Intake I Death Nov. 17/82 male, 21 yrs seizures amitriptyline cirnetidine Nov. 17183 1 fernale, 23 yrs, cocaine epileptic -- - Mar. 6/84 female, 35 yrs phenytoin phenobarbital Apr. 12/84 male, 39 yrs motor dopamine, Pen G vehicle levophed colIision furosemide -- --- subarachnoid mannitol hemorrhage dopamine Jul. 15/85 male, 20 yrs drowning insu lin lasix Feb. 7/86 female, 37 yrs cerebral dopamine nonsmoker bleeding hep arin mannitol Feb. 10/86 female, 50 yrs meningioma- diazepam nonsmoker craniotomy , phenobarbital postoperative nitroprusside cerebral dexamethasone ederna Mar. 13/86 female, 50 yrs cardiac propanol01 nonsmoker arrest methyldopa amitriptyline chlorodiazepoxide May 10/86 male, 22 yrs motor nonsmoker vehicle collision May 22/86 female, 21 yrs suicide nonsrnoker 11 Liver # 1 Date Obtained Physical Description Cause of Known Drug Intake Death Jun. 10/86 male, 26 yrs motor dopamine vehicle collision female, 39 yrs subarachnoid dopamine hemorrhage lidocaine male, 15 yrs motor dopamine vehicle collision female, 55 yrs subarachnoid antibiotics hemorrhage K28 hl. 15/86 male, 21 yrs bilateral dopamine cerebral vasopressin infarction K29 Aug. 13/86 male, 19 yrs ~ motor dopamine vehicle vasopressin collision phenylephrine Furosemide 2efazoline mannitol heparin note: al1 donors were of Cau