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Protein induction and estrogen metabolism in subcellular preparations and tissue slices of the golden Syrian hamster
Beleh, Mustapha Ahmed, Ph.D.
The Ohio State University, 1992
UMI 300 N. ZeebRd. Ann Arbor, MI 48106 PROTEIN INDUCTION AND ESTROGEN METABOLISM IN SUBCELLULAR
PREPARATIONS AND TISSUE SLICES OF THE GOLDEN SYRIAN HAMSTER
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the degree of Philosophy in the Graduate
School of The Ohio State University
By
Mustapha Ahmed Beleh, B.S.
****
The Ohio State University
1992
Dissertation Committee Approved by:
Robert W. Brueggemeier, Ph.D.
Young C. Lin, D.V.M., Ph.D.
Duane D. Miller, Ph.D. Adviser
Dennis R. Feller, Ph.D. College of Pharmacy Dedication
To the sweetest sister, Dalia
ii ACKNOWLEDGMENTS
I would like to express my gratitude and thanks to everyone who aided in making the past five years successful:
My family, especially my late mother, my father, stepmother, sister and brother for all the support and care they offered over the years.
My adviser Dr. Robert Brueggemeier for his guidence, patience and teachings over the course of my graduate career.
The members of my graduate committee, especially Dr. Young Lin for his help and advise.
Denise Donley and Patricia Goetz, both for their assisting in some of the studies performed within this dissertation work and for their friendship, Nancy
Katlic for her help with different aspects in the laboratory, George Chang for aiding in cell culture related techniques and Jack Fowble and John Miller for their various technical help.
Soheila Ebrahimian, my friend and laboratory partner for five years, and my friend and roommate Nader Moawad for all the help and support they supplied.
My fellow graduate students, especially Kimberly Markovich, Allen
Hopper, Meri Slavica and Anne "cool" Quinn.
Yasser AbdelGhany, a friend who is always there, and all my friends over the years, Vimon, Yushi, Becky and Nagy. Finally, I would like to thank my friends who stood by me during the final phase of my graduate studies: Helen, Saroj, Yasmine and Patricia White for their help in preparation of this dissertation, Farida, Michelle and Shamim for their friendship and support. VITA
February 24, 1963 ...... Bom - Egypt
June 1985 ...... B.S. pharmacy, University of Alexandria
School of Pharmacy, Egypt
1987 - 1988 ...... Graduate Teaching Associate
The Ohio State University
Columbus, Ohio
1988 - 1991 ...... Graduate Research Associate
The Ohio State University
Columbus, Ohio
1991 - present ...... Proctor and Gamble fellowship
The Ohio State University
Columbus, Ohio
PUBLICATIONS AND ABSTRACTS
1.Brueggemeier, R. W.; Tseng, K.; Katlic, N. E.; Beleh, M. A.; Lin, Y. C. Estrogen metabolism in primary kidney cell cultures from Syrian hamsters. J. Steroid Biochem. 1990, 36, 325-331.
2.Brueggemeier, R, Tseng, K., Katlic, N., Beleh, M., Lin, Y. Estrogen metabolism in primary kidney cell cultures from Syrian hamsters, American Association of Cancer Research, Washington D.C., May 1990, Abst. No. 1330.
v 3. Beleh, M., Donley, D., Lin, Y., Brueggemeier, Estrogen metabolism in subcellular preparations and tissue cultures of Syrian hamster liver and kidney, Internationa Symposium on Hormonal Carcinogenesis, Cancun, Mexico, March 1991.
4.Beleh, M., Donley, D., Lin, Y., Brueggemeier, Estrogen metabolism in subcellular preparations and tissue cultures of Syrian hamster liver and kidney, The Endocrine Society meeting, Washington D.C., June 1991, Abst. No. 532.
FIELD OF STUDY
Major field : Pharmacy TABLE OF CONTENTS
PAGE
DEDICATION...... ii
ACKNOWLEDGMENTS...... iii
VITA...... v
LIST OF FIGURES...... x
LIST OF TABLES...... xiv
LIST OF PLATES...... xvi
CHAPTER I. INTRODUCTION...... 1
1.1 Cancer ...... 1 1.1.1 Historical perspective ...... 1 1.1.2 The cancer epidemic ...... 3 1.1.3 Cancer terminologies ...... 4 1.1.4 Treatment of cancer ...... 5 1.1.5 Risk factors and cancer ...... 9
1.2 Hormone-dependent cancer ...... 10 1.2.1 Endocrinology and hormones ...... 10 1.2.2 Steroid hormones and their receptors ...... 12 1.2.3 Estrogens and their physiological role ...... 17 1.2.4 Estrogen biosynthesis and metabolism ...... 21 1.2.5 The role of estrogens in carcinogenesis ...... 23
1.3 The golden Syrian hamster : an animal model for estrogen-dependent cancer ...... 26 1.3.1 Induction of renal adenocarcinomas ...... 26 1.3.2 Morphology and histology of the tumors ...... 29 1.3.3 The role of hormonal and carcinogenic activity of estrogens in tumorigenesis ...... 33 1.4 Growth factors, oncogenes and cell transformation ...... 36 1.4.1 Growth factors : general aspects ...... 36 1.4.2 Growth factors : a closer look ...... 41 1.4.3 Oncogenes and tumor suppressor genes ...... 44 1.4.4 The role of growth factors, oncogenes and tumor supressor genes in breast cancer ...... 47
1.5 Estrogen metabolism...... 50 1.5.1 Oxidative metabolism of estrogens ...... 50 1.5.2 Estrogen metabolism in Syrian hamsters ...... 53 1.5.3 The role of DNA and protein adducts and cell death in carcinogenesis ...... 58
CHAPTER II. OBJECTIVES AND EXPERIMENTAL...... 62
2.1 Statement of objectives ...... 62
2.2 Experimental ...... 64 2.2.1 Materials and methods ...... 64 2.2.2 Preparation of primary kidney cell cultures ...... 66 2.2.3 Preparation of kidney and liver microsomes ...... 67 2.2.4 Preparation of kidney and liver slices ...... 68 2.2.5 Protein biosynthesis in primary kidney cell cultures...... 68 2.2.6 Separation of proteins using one and two dimensional electrophoresis ...... 69 2.2.7 Western blotting ...... 71 2.2.8 Immunoprecipitation of TGF-a and bFG F ...... 72 2.2.9 Estrogen metabolism in primary kidney cell cultures ...... 72 2.2.10 Separation of various metabolites ...... 73 2.2.11 HPLC analysis of estrogen metabolites ...... 73 2.2.12 Characterization of water soluble metabolites ...... 74 2.2.13 Subcellular distribution of metabolites in tissue slices ...... 75 2.2.14 3H20 assay of estrogen 4-hydroxylase ...... 75 2.2.15 Bromoestrogens inhibition of estrogen metabolism ...... 76 2.2.16 Approaches to structure elucidation of estrogen metabolism ...... 77
CHAPTER III. RESULTS AND DISCUSSIONS...... 78
3.1 Cell type identification ...... 78
3.2 Protein biosynthesis in kidneys of Syrian hamsters ...... 81 3.2.1 Separation and identification of proteins ...... 81 3.2.2 Identification of specific proteins ...... 83
viii 3.3 Estrogen metabolism in Syrian hamsters ...... 127 3.3.1 Estrogen metabolism in primary kidney cell cultures ...... 127 3.3.2 Estrogen metabolism in microsomal pellets ...... 128 3.3.3 Estrogen metabolism in tissue slices ...... 128
3.4 Kinetics of estrogen 2/4 hydroxylase ...... 159
3.5 Distribution of estrogen metabolites in kidney and liver slices ...... 163
3.6 Characterization of water soluble metabolites ...... 172
3.7 Inhibition studies bromoestrogens ...... 179
3.8 Approaches to the identification of metabolites ...... 189
CHAPTER IV. SUMMARY...... 191
BIBLIOGRAPHY...... 198
ix LIST OF FIGURES
FIGURE PAGE
1. The cell cycle ...... 8 2. Examples from different classes of steroid hormones ...... 13
3. Action of steroid hormones ...... 14
4. The zinc finger structure in the steroid receptor ...... 16
5. Estrogen secretion during the menstrual cycle ...... 18
6. Natural and synthetic estrogens ...... 20
7. Estrogen biosynthesis ...... 22
8. Major routes of estrogen metabolism ...... 23
9. The origin of kidney tumors in Syrian hamsters ...... 31
10. Appearance of dysplasic foci and increase in area of vascularization in kidneys of estrogen treated hamsters ...... 32
11. Different factors participating in the tumorigenesis in Syrian hamsters ...36
12. The action of growth factors on the cell cycle ...... 38
13. Substrates of growth factor receptors ...... 39
14. Amino acid sequence of IG F -I ...... 45
15. The oxidation of catechols to the muconic acid ...... 51
16. Mechanism of catecholestrogens formation ...... 52
17. Metabolism of diethylstilbesterol ...... 55
18. Redox cycle of quinones ...... 57
x 19. Levels of 8-hydroxyguanosine in kidneys and liver of DES-treated hamsters ...... 61
20. Mechanism of cell transformation by estrogen binding to tubulin ...... 61
21. Scan of autoradiogram from kidney cell culture of untreated hamsters on DU-8 spectophotometer ...... 88
22. Scan of autoradiogram from kidney cell culture of three months DES-treated hamsters on DU-8 spectrophotometer ...... 95
23. Scan of autoradiogram from kidney cell culture of six months DES-treated hamsters on DU-8 spectrophotometer ...... 102
24. Scan of autoradiogram from kidney cell culture of nine months DES-treated hamsters on DU-8 spectrophotometer ...... 109
25. Scan of autoradiogram from kidney cell culture of eleven months DES-treated hamsters on DU-8 spectrophotometer ...... 116
26. Biosynthesis of protein band e (34 kDa) and band g (20 kDa) in kidney cell cultures of untreated and DES-treated hamsters ...... 122
27. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney cell cultures of untreated and DES-treated hamsters (full view)...... 131
28. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney cell cultures of untreated and DES-treated hamsters (expanded view) ...... 133
29. The effect of DES treatment on major metabolites formation in kidney cell cultures ...... 135
30. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney cell cultures of untreated hamsters using [4-14C]-estradiol and [6,7-3H]-estradiol ...... 137
31. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney microsomes of untreated and nine months DES-treated hamsters ...... 139
32. Reverse phase HPLC radiochromatogram of organic extractable metabolites from liver microsomes of untreated and nine months DES-treated hamsters ...... 141 33. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters ...... 143
34. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of four months DES-treated hamsters... 145
35. Reverse phase HPLC radiochromatogram of organic extractable metabolites from liver slices of untreated hamsters ...... 147
36. Reverse phase HPLC radiochromatogram of organic extractable metabolites from liver slices of four months DES-treated hamsters 149
37. Polar metabolite formation in kidney and liver slices of untreated and four months DES-treated hamsters ...... 151
38. Major metabolites formation in kidney slices of untreated hamsters 153
39. Major metabolites formation in liver slices of untreated hamsters 155
40. Kinetic parameters of estrogen 2- and 4-hydroxylase in kidney and liver microsomes of untreated hamsters ...... 161
41. Distribution of metabolites in liver and kidney slices of untreated and four months DES-treated hamsters ...... 164
42. Distribution of metabolites in different cellular components in liver slices of untreated hamsters ...... 166
43. Organic soluble, water soluble, and protein bound metabolites in liver slices of untreated hamsters ...... 168
44. Reverse phase HPLC radiochromatogram of organic extractable metabolites from different cellular fractions in liver slices of untreated hamsters ...... 170
45. Reverse phase HPLC radiochromatogram of organic extractable metabolites after cleaving of the water soluble metabolites in formed liver slices of untreated hamsters 175
46. Reverse phase HPLC radiochromatogram of organic extractable metabolites after cleaving of the water soluble metabolites in formed kidney slices of untreated hamsters 177
47. Structures of bromoestrogens ...... 180 48. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters in presence of 2-bromoestradiol ...... 181
49. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters in presence of 4-bromoestradiol ...... 183
50. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters in presence of 2,4-dibromoestradiol ...... 185
51. Inhibition of the formation of polar metabolites, estrone and methoxyestrogens by 4-bromoestrogen in kidney slices of untreated hamsters ...... 187
52. Possible metabolic pathways involved in estrogen metabolism in different preparations from Syrian hamster ...... 196
xiii LIST OF TABLES
TABLE PAGE
1. Correlation of the carcinogenic and hormonal effects of estrogens with their ability to induce tumors in Syrian hamsters ...... 35
2. Pretreatment and treatment of kidney cell culture with 10 nM estradiol ..68
3. Total protein secreted from primary kidney cell cultures of untreated and DES-treated hamsters ...... 85
4. Total protein secreted from primary kidney cell cultures of untreated hamsters subjected to different treatments with estradiol ...... 86
5. Major protein bands from autoradiograms of secreted proteins from kidney cell cultures of untreated hamsters ...... 93
6. Major protein bands from autoradiograms of secreted proteins from kidney cell cultures of three months DES-treated hamsters 100
7. Major protein bands from autoradiograms of secreted proteins from kidney cell cultures of six months DES-treated hamsters ...... 107
8. Major protein bands from autoradiograms of secreted proteins from kidney cell cultures of nine months DES-treated hamsters 114
9. Major protein bands from autoradiograms of secreted proteins from kidney cell cultures of eleven months DES-treated hamsters 121
10. Values of apparent Km ± S.E. of enzymes catalyzing the formation of the major metabolites in liver and kidney slices of untreated hamsters ...... 157
11. Values of apparent Vma)S + S.E. of enzymes catalyzing the formation of the major metabolites in liver and kidney slices of untreated hamsters ...... 158
12. Percent of water soluble metabolites formed in liver slices from untreated hamsters ...... 173
13. Percent of water soluble metabolites formed in kidney slices from untreated hamsters ...... 174
xiv 14. Partial listing of peaks identified in electron ionization mass spectroscopy of metabolite c
xv LIST OF PLATES
PLATE PAGE
I. A microscopical section from primary kidney cell cultures staining positive for the presence of GGT activity ...... 79
II. A microscopical section from primary kidney cell cultures staining positive for the presence of AP activity ...... 80
III. Autoradiogram of secreted proteins from kidney cell cultures of untreated hamsters ...... 87
IV. Autoradiogram of secreted proteins from kidney cell cultures of three months DES-treated hamsters ...... 94
V. Autoradiogram of secreted proteins from kidney cell cultures of six months DES-treated hamsters ...... 101
VI. Autoradiogram of secreted proteins from kidney cell cultures of nine months DES-treated hamsters ...... 108
VII. Autoradiogram of secreted proteins from kidney cell cultures of eleven months DES-treated hamsters ...... 115
VIII. Autoradiogram from two-dimensional electrophoresis of secreted proteins from kidney cell cultures of nine months hamsters ...... 124
IX. Autoradiogram of immunoprecipitated proteins from kidney cell cultures of untreated and DES-treated hamsters using antibodies against T G F-a ...... 125
X. Autoradiogram of immunoprecipitated proteins from kidney cell cultures of untreated and DES-treated hamsters using antibodies against bFGF ...... 126
xvi CHAPTER I
INTRODUCTION
1.1 CANCER
1.1.1 HISTORICAL PERSPECTIVE
In every historical era a certain disease seems to linger as the most fearsome.
In the biblical years leprosy, in the middle ages plague, in the nineteenth century tuberculosis and perhaps in the twentieth century cancer. However, cancer has been known since the early ancient Egyptians who described surgical procedures to remove tumors on some of their papyri as early as 2500 B.C. [1].
Early civilizations attributed the cause of cancer to various gods. The first to use the term carcinoma was Hippocrates back in the fourth century B.C. as a reference to tumors that spread and destroy the patient, in contrast to carcinos which include benign tumors. He postulated that cancer was due to an increase in black bile which is secreted by the spleen, the first theory to relate cancer to natural causes [2].
In the second century A.D., Galen classified tumors to those according to nature, such as breasts in females, those exceeding nature, such as bone proliferation during healing of fractures and those contrary to nature which are neoplastic growth. He also was the first to outline the similarity between the gross shape of a crab and the
1 disease cancer [3]. Writings from the middle ages made reference to cancer houses, families and villages, which were the first suggestion that cancer may be inherited or has an environmental etiology, but it was also thought of as an infectious disease [4].
In the seventeenth and eighteenth century, the theory of the black bile as the cause of cancer began to be disputed. In 1775, Sir Percival Pott was the first to describe occupational cancer due to exposure to carcinogens, when he described the occurrence of scrotum tumors in young chimney sweepers. He not only identified carcinogens as causative agents, but also suggested that cancer has an extended latent period [5].
Ramazzini also reported life style associated cancer when he attributed the high occurrence of breast cancer among nuns to the celibate life of these women [6]. The nineteenth century saw the emergence of theories about the etiology of cancer as we understand it today. Bichat described the anatomy of many cancers and defined it as an accidental formation of tissues [7]. Later, Johannes Muller was able to demonstrate that the cancer tissue was made up of cells using a microscope. In 1863,
Rudolf Virchow not only suggested a relationship between irritation and some cancers, but more importantly declared that every cell is bom from another cell, thus establishing cancer as a cellular disease [8]. However, scientists were divided on the origin of cancerous cells. Some believed that normal cells are converted to cancerous cells, while others thought that cancer cells exist from embryonic life but are not expressed until later in the organism's existence [9]. The term metastasis was introduced in 1829 by Recamier [10], but it wasn't until 1872 that Waldeyer showed that cells from a primary cancer can infiltrate blood and lymphatic vessels [11].
At the beginning of the twentieth century, three theories existed for the etiology of cancer [12]: 3
1. The irritation theory introduced by Virchow, related cancer to the effect of
chemical agents.
2. The embryonic theory introduced by Recamier and Lobstein, that argued
that cancerous cells exist from birth and are activated later in life.
3. The parasitic theory which thought of cancer as an infectious disease.
This was supported by the finding of Doven that a bacterium, micrococcus
neoformans , was present in some neoplasms and thus believed to be the
cause of all cancers. This was later found to be a common staphylococcus.
Now we realize that cancer has various etiologies. It is now accepted that most neoplasms appear to arise from a single cell that undergoes malignant transformation [13]. The first step is called initiation which is the change in DNA induced by a variety of chemical, physical or viral agents. In most cases, the presence of a promoting factor, which may be the same agent or a different factor, is required to complete the carcinogenic process [14]. The susceptibility of DNA to change may vary from one individual to another. This can be illustrated by differences in normal DNA repair mechanisms, variable expression of oncogenes, mutations in tumor suppressor genes and other hereditary factors, and finally the immune function as illustrated in the development of Kaposi’s sarcoma in patients with acquired immune deficiency syndrome (AIDS) [15].
1.1.2 THE CANCER EPIDEMIC
Cancer is now the leading cause of death among women in the United States.
If the trend continues it will be the leading cause of death in the U.S. by the year
2000. This rise is primarily due to the decline in deaths due to cardiovascular 4 diseases rather than a sharp rise in the cancer mortality [16]. As a matter of fact, the age-adjusted death rate from cancer over the past 60 years has increased only slightly.
This doesn't reflect the progress in treatment of all types of cancers, which could be understood when we realize the increase in cancer incidence: one in every three
Americans alive today will develop cancer. In 1991, more than a million individuals will be diagnosed as having cancer and that doesn't include nonmelanoma skin cancers. To be cured of cancer means that a patient has no evidence of the disease and has the same life expectancy as a person that never had cancer. For most types of cancers, five years without symptoms following treatment is accepted to consider a patient cured. In the beginning of this century, few patients had any hope of long term survival. By the thirties, one in every five patients was considered cured; in the forties, one in four; in the sixties, one in three and today four out of every ten patients will be cured, which amounts to 440,000 patients of those diagnosed with cancer this year. Still one of every five deaths in the U.S. are due to cancer [17].
1.1.3 CANCER TERMINOLOGIES
Over the years many terms have been used and then confused. A tumor is a readily defined mass of tissue distinct from normal physiological growth [18]. Cancer is now best defined as the biological characteristics of a malignant neoplasm.
Neoplasia is a heritably altered, relatively autonomous growth of tissues [19]. In addition plasias can be divided into [20]:
- Hyperplasia, referring to an increase in cell number.
- Metaplasia, or the change of one type of cell to another in a specific
organ. - Dysplasia, which is the alteration of cells in shape, size and organization.
- Anaplasia, denoting the alteration in intracellular macromolecule synthesis
such as proteins and nucleic acids.
For a neoplasia to be malignant, it has to possess the following characteristics: nonencapsulated, invasive, poorly differentiated, mitotic, does not exhibit contact inhibition, grows rapidly, is metastatic and anaplastic [21].
Neoplastic tumors are also classified according to the type of tissue they originated from, such as epithelial, connective tissue, hemopoietic and immune
system [22]. But probably the most widely used classification is based on the embryonic origin [23]. In this classification, the cancer is named according to its origin followed by the suffix -oma, which means tumor. Thus sarcoma is a malignant neoplasm arising from the mesodermal embryonic germ layer. A carcinoma, on the other hand, arises from the endoderm or ectoderm. A highly malignant tumor that has the appearance of both types is termed carcinosarcoma, while a tetroma arises from the three germ layers. The suffix -blastoma indicates tumors that have a primitive appearance resembling embryonic structures. Finally, cancer involving the abnormal increase in leukocytes is called leukemia.
1.1.4 TREATMENT OF CANCER
Early in cancer treatment, the one method used was surgery. Today there are four main approaches to the treatment of cancers: (1) Surgery
It is still an effective way of removal of cancerous cells that have not
metastasized. It is a common method of treatment in many cancers such as breast and ovarian cancers, but even if few tumor cells have been dispersed to other organs such as the lungs and the brain it cannot be treated successfully by surgery. One problem associated with surgery is the common recurrences of the cancers within five years of the operation [24].
(2) Irradiation
The discovery of the X-ray back in the nineteenth century by Roentegen, followed by the Curies' discovery of radium, opened a new avenue for treatment of cancers: radiotherapy [25]. This is based on delivering a lethal dose of radiation to the tumor with a minimal dose to the surrounding tissues and organs. Fortunately, the cancerous cells are more susceptible than normal tissues to the damaging effects of radiation. This method is very efficient in localized tumors such as those of the larynx and the prostate. The main drawbacks lie in the damage to the surrounding tissues, the alteration of the immune system of the patient, and the occasional failure of such therapy due to the insensitivity of the cells or the inadequate dose of the irradiation [26].
(3) Immunotherapy
This is probably the least developed method in cancer therapy. It is used to destroy the last few cells after using other forms of therapy. A non-specific antigen such as Bacille Calmette Guerin (BCG), a weakened derivative of a mycobacterium bovis is used to boost the immune system of the patient and stimulates macrophage production. Recently immunization using the patient's own immune cells has been
under investigations [27].
(4) Chemotherapy
This therapy involves the administration of drugs that would kill cancerous
cells. Unfortunately, these drugs also have some effects on normal cells, although
there is a difference in the susceptibility to these agents [28]. However, they have the
advantage of not being limited by the metastasis of tumors as they act on the total
mass of tumors. In general, chemotherapeutic agents can be classified into :
1. Alkylating agents, which cross-link DNA and block its replication.
Examples include nitrogen mustards, thiotepa, chlorambucil and
cyclophosphamide [29].
2. Antimetabolites, which inhibit nucleic acid and protein biosynthesis,
thereby leading to cell death. These include methotrexate,
6-mercaptopurine and 5-flourouracil [30].
3. Miscellaneous agents, such as Vinca alkaloids [31].
New approaches are now being investigated for developing specific agents for
different types of cancers. For example, in breast cancer, agents that inhibit the
action of estrogens (antiestrogens) such as tamoxifen [32] and those that inhibit
estrogen biosynthesis (aromatase inhibitors) such as 4-hydroxyandrostenedione and
aminoglutethimide [33] have been employed. These are designed to have fewer side
effects than the general agents and may be more beneficial in treating these types of cancers. All cells which are synthesizing DNA go through a series of phases known as
the cell cycle (fig. 1). A cell in the resting G0 phase may enter the cell cycle due to many factors. It goes to the Gt phase where it contains diploid amount of DNA. It
can spend varying periods of time in thisphase, until it goes to the S phase which
involves DNA synthesis. During Glf DNA synthesis is absent, while RNA and
protein synthesis continues normally. In the late Glt an unknown signal initiates a burst of RNA synthesis and shortly thereafter S phase begins, and the cell becomes committed to undergo division or remain polyploid. During G2 phase, the cell ceases to synthesize DNA and enters mitosis, while RNA and protein synthesis continues.
When it enters the M phase, RNA and protein synthesis diminishes abruptly, while the genetic material segregates into daughter cells [34].
S
Figure 1. The cell cycle
(Adapted from Pitot [23]) 9
Chemotherapeutic agents can be classified according to where they act in the cell cycle:
(a) Class I, non-specific agents that are toxic to both proliferating and
resting (G0) cells. Examples are nitrogen mustard and gamma
irradiation.
(b) Class II, phase-specific agents. These affect cells in specific parts of
the cell cycle and don't affect cells in the G0 phase if exposure time is
short. Examples include vinblastine, methotrexate and azaserine.
(c) Class III, cycle specific agents. These agents damage both proliferating
and resting cells but cycling cells are more sensitive than resting cells.
These include 5-fluorouracil, actinomycin and cyclophosphamide.
1.1.5 RISK FACTORS AND CANCER
Cancer appears to depend greatly on the life style and personal choices of individuals, and it is now suggested that these factors are far more important than environmental pollution. Exposure to carcinogens, such as many chemicals and radiation can increase the chance of an individual to contract cancer. This was first demonstrated in occupationally-related cancers [35]. It has been estimated that 30% of all cancer mortality is due to smoking and 35% to diet-related causation, while only 20% are due to a variety of exogenous factors including viruses, drugs, exposure to X-rays, ultraviolet light and other radiation, i.e. more than 65% of all cancer deaths are self-preventable [36]. Tobacco and alcohol are the major causes of cancer.
Animal fat especially from red meat appears to play a role and a high fiber, low fat diet appears to be beneficial especially in colon and rectal cancers. Vitamin A, C, 10 and E have been suggested to lower cancer risk, while direct exposure to sunlight increases the risk of skin cancer [37]. Heredity is another risk factor, and some neoplasms are passed through defined Mendelian patterns of heredity [38]. Finally hormones have been shown to be associated with many types of cancers. Endometrial cancer is caused by the cumulative exposure to estrogen in absence of progestins [39], while breast cancer is enhanced by both hormones [40]. Ovarian cancer is also affected by a variety of hormonal changes [41] and prostate cancer is most likely related to the exposure to testosterone or its metabolite, 5-hydroxytestosterone [42].
A high percent of these cancers depend on hormones for their development as well as for their initiation. Drugs that would decrease the levels of these hormones or oppose their actions can be used as potential therapeutic agents against these cancers, which are known as hormone-dependent cancers.
1.2 HORMONE-DEPENDENT CANCER
1.2.1 ENDOCRINOLOGY AND HORMONES
Multicellular organisms function through a complex network of communication. One system is the nervous system, composed of a central control lying in the brain and the spinal cord, and connected by wires or nerve cells to various parts of the body to deliver and receive messages. The other system is the endocrine system which is a wireless system and depends on certain chemicals in communications. These chemical substances fall into two major categories: hormones and growth factors [43]. 11
The endocrine systems have been known for a long period of time. Ancient
Egyptian papyri dating back to 1500 B.C. describe a disease that appears to be related
to diabetes mellitus. Over the years, other endocrine-related disorders have been
described by ancient Chinese and Indian physicians and early Western scientists.
In the nineteenth century, Claude Bernard was the first to postulate that the
different cell fluids must be controlled and serve an important role in the well being
of the organism. He also postulated that the two communication systems, the nervous
and the endocrine systems once thought to be acting independently are in fact very
closely related and concluded that they work in collaboration, or in his own words
serve "to maintain the internal milieu" [44],
The growth of endocrinology was aided by the use of animal models.
Discovery of new substances and functions of glands was usually achieved via the
removal of a gland, and observation of the biological effects. Then the extract of the gland was injected and the recovery from the biological effect, if demonstrated, provided information about the presence of a substance responsible for this effect.
This substance was then isolated and identified [45].
These substances were usually hormones which are defined as chemical
messengers that are secreted directly into the blood stream by specialized cells capable of synthesizing and releasing them in response to specific signals [46]. Hormones
usually act in a telecrine nature, exerting their effects at a distant cell or tissue.
However, in some cases, they act as paracrine factors, acting on neighboring cells or even in autocrine fashion, acting on the same cell. Hormones are secreted at very low concentrations, usually in the nanomolar to picomolar range. Earlier ancestors of the
hormones have been detected in simple organisms, such as E. coli which has a 12 substance very similar to insulin. As organisms become more and more complex, the number of hormones and their functions increase [46].
Hormones are usually classified according to their chemical structures, functions or source. For example, estrogens can be classified as steroidal, sex or ovarian hormones [47]. Steroid hormones also include progestins, androgens, glucocorticoids, mineralocorticoids and cholecalciferols (Figure 2).
1.2.2 STEROID HORMONES AND THEIR RECEPTORS
Steroid hormones are a group of substances having a common steroidal skeleton. They have a common precursor, cholesterol, which through a series of enzymatic reactions leads to the biosynthesis of the different classes of steroid hormones. They are highly lipophilic and circulate bound to carrier proteins, with a minute amount found in the unbound form, the latter being the biologically active form. It is uncertain how they enter cells, but it has been postulated that the free form dissolves in the lipid bilayer of the plasma membrane. In addition, a plasma membrane receptor has been suggested, and may serve as a transport function.
However, the relation between this receptor and the interior receptor is unknown [46].
Once the steroid hormone enters the cell, it binds to a high affinity receptor, which resides largely in the nucleus. This complex then binds to the DNA and chromatin through a DNA binding domain on the receptor, which leads to an increase in gene expression and transcription of specific mRNA and thus the production of a wide range of proteins and enzymes (Figure 3) [48]. Cholesterol Progesterone
CHjOH CHjOH I C = 0 c = o -OH
Corticosterone Cortisol
CH2OH
c = o OH H—C CH3
O O Aldosterone Testosterone
COOH OH
OH HO H
Cholic Acid Estradiol-170
Figure 2 Examples from different classes of steroid hormones
(Adapted from Schulster [49]) 14 . S R ^ e 'f transformation mRNA DNA ® p 7 \ transport s i ? p activation ' d * transcription / 'y P // \mRNA / processing nuclear ^ *wiw» ^yT-dA pore SR Nucleus Cytoplasm Figure 3. Action of steroid hormones
(Adapted from Liao et al [61])
The steroid receptors are closely related with similar molecular weights and
shapes. It was earlier believed that the unoccupied receptor resided in the cytoplasm,
and that when it bound the steroid, migrated to the nucleus [50]. However, it has
recently been shown that due to technical flaws this is not true [51, 52]. It appears
now that the receptors are mostly nuclear. In addition, the receptors can be phosphorylated by kinases and dephosphorylated by phosphatases and that only the unphosphorylated receptors can bind the steroid [53].
Work by Jensen in the 1950s focused attention on the presence of high affinity binding sites for estrogens in estrogen target cells [54]. This was later recognized as the estrogen receptor. Recently, the study of the nature and function of these receptors was aided greatly by the development of antibodies against them and then the cloning of cDNA and genes for these receptors [55, 56]. These studies were directed towards the early events in the binding of steroids to the receptors followed by the binding of this complex to specific DNA sequences. From these
investigations, a very high homology among different steroid receptors, as well as
with the thyroid and the vitamin D receptor and even with the v-erb A, an oncogenic
protein was detected [57, 58]. The receptor can be divided to five domains named
A-E. Regions A and B contain information that enhances transcription as well as
preferential activation of certain genes. While these regions that are present towards
the amino-terminal end vary in size from one type of receptor to another, the regions
at the carboxy end are similar in size [59]. Region C, a 6 6 amino acid sequence, is conserved throughout this family of receptors, and is known as the DNA binding domain, while region E is the steroid binding domain [60]. Fjrther studies showed
the presence of nine cysteine moieties in region C, eight of which form two zinc
fingers each containing one zinc molecule (Figure 4), which may aid in the binding to
DNA and may also act as transcriptional factors, more of which are present at other regions [62]. The zinc fingers also contain a sequence which cognate the specific
steroid response elements (SRE) in the genomic DNA. This was substantiated when the zinc finger region of the estrogen receptor replaced that on a glucocorticoid receptor and the resulting molecule activated genes containing an estrogen response element [63]. It has been suggested that the first zinc finger contains the primary information for sequence specificity of binding while the second stabilizes the binding of the receptor to DNA [64]. The unoccupied receptor appears to exist in association with cellular heat shock proteins (hsp90), which renders the receptor incapable of binding to DNA. When the steroid hormone binds, these hsp90 dissociate, exposing regions important for DNA binding [65]. How the steroid receptor complex then activates specific genes and induces the production of certain proteins and enzymes is less well understood. What is known is that target genes possess short cis elements known as SRE located usually within their 5'-flanking regions, and these confer
hormonal regulation upon receptor binding [ 6 6 ]. Any changes in these SREs can alter
the receptor binding or abolish it. As little as two base changes convert a
glucocorticoid responsive gene to an estrogen responsive gene [63]. Another
important factor is the presence of sequences that participate in transcriptional
activation at regions A, B, and D. These sequences are thought to be accessible upon binding of the steroid causing an allosteric modification of the receptor. As a matter of fact, it appears that steroid antagonists such as tamoxifen act by binding to the receptor but are unable to cause the correct allosteric effect to unveil these factors
[67, 6 8 ].
Qly HI* — Affl - Tyr Ala aiy Lys - - Val Asp Cys Aap “ lls “ Qly Tltf - Ala C ys ,Cya Val Zn Qly A l L»«“ C ys Cy* Cys Cys - Lys Gly Met - P h s l I Qly Tyr
Figure 4. The zinc finger structure in the steroid receptor
(Adapted from O'Malley [59]) 17
1.2.3 ESTROGENS AND THEIR PHYSIOLOGICAL ROLE
Estrogens are the group of steroid hormones that arise from aromatization of
C1 9 steroids to yield C 1 8 steroids through the action of a specific enzyme, aromatase.
Estrogens are the female sex hormones. This group of compounds, by definition, have one common effect: the development and maintenance of the female sex characteristics. Estrogens influence the growth and function of the uterus, the development of the vagina, the vaginal mucosa, the cervix, the fallopian tubes and breast tissues. They also play an important role in the female menstrual cycle. They stimulate the secretion of leutinizing hormone (LH) which leads to ovulation and formation of corpus luteum [69]. In addition, they are produced in the later stages of pregnancy and they sensitize the uterus to the action of oxytocin that stimulates uterine contraction. Also estrogens are responsible for the growth of the mammary glands in readiness for milk production [70]. They also have other secondary effects that include effects on bone maturation and structure, an increase in blood coagulation, water retention and the synthesis of proteins and enzymes [71]. Recently estrogens were detected in the brain and appear to have a function in brain development [72].
The most potent natural estrogen in humans is estradiol. It arises from the aromatization of the C 1 9 testosterone. Studies done by Falck showed that granulosa cells in the ovaries were less efficient than theca interna cells in estrogen biosynthesis
[73]. Short and Ryan demonstrated that granulosa cells had weaker biosynthesic capabilities but that they contained all the necessary enzyme systems [74]. These results agreed with Short's two cell theory of estrogen biosynthesis where he visualized that both cells of the follicle collaborated in the estrogen biosynthesis [75]. 18
The current concept is that theca cells are the target of leutinizing hormone (LH) and
thus are the main source of the C 1 9 steroids and that the granulosa cells are the
primary site for aromatization.
The amount of estradiol secreted depends on the menstrual cycle stage [76].
During the early follicular phase, it is produced at about 60 ^cg/day. Later on, it
increases in amount until it reaches 400 /xg/day at the height of the preovulatory
estrogen surge. During the luteal phase, it is produced mainly by the ovary carrying
the corpus luteum. It is unclear which of the cells of the corpus luteum are
responsible for estrogen biosynthesis, but evidence suggest that the theca interna cells
may play that role. The amount of estradiol produced decreases sharply at the beginning of the luteal phase and then slowly increases as the corpus luteum matures and the levels diminish again until menses (Figure 5). 1 8 CO
O — ---- E 2 0 0 - < t r aO) h - 1 0 0 - C/D J LU J L
-12 -8 -4 0 4 8 12
DAY OF CYCLE
Figure 5. Estrogen secretion during the menstrual cycle
(Adapted from Yen [77]) 19
In general, estrogens are classified as steroidal and non-steroidal estrogens.
Steroidal estrogens are either natural or synthetic. Natural steroidal estrogens include estradiol and its various metabolites such as estrone and estriol. Equilin and equilenin are another group of natural steroidal estrogens. Synthetic steroidal estrogens are
mainly esters and 17a-ethinyl derivatives of the natural estrogens. Non-steroidal estrogens include stilbenes such as diethylstilbestrol and phytoestrogens (Figure 6 ).
Natural steroidal estrogens all have the C 1 8 hydrocarbon skeleton, termed estrane. The stereochemistry of the molecule is determined in relation to the C-18 methyl group which is assigned the B-position. Thus the a-position would be down from the plane of the molecule and the B-position is above the plane of the molecule.
The common structural feature for steroidal estrogens include an aromatic ring and a hydroxyl or hydroxy conjugate at C-3 and an oxygen-containing substituent at C-17.
Estradiol (E£ has a hydroxyl group at the C-17 in a B-position. If the hydroxyl group is in the a-position, the compound, named estradiol 17-a, is inactive. Estrone (Ej), another estrogen where the 17-hydroxyl group is oxidized to the ketone is interconvertible with estradiol in vivo, and has approximately 50-70% of its activity.
Estriol (E 3 ), an estrogen unique to humans, has a 16a-hydroxyl group and possess
10% of the activity of estradiol. Synthetic steroidal estrogens were mainly designed to obtain better oral absorption. 17a-Ethinylestradiol (EE) has proven to be a very potent, orally active estrogen. The addition of a methyl ether at C-3 (mestranol) results in a less potent estrogen while the cyclopentyl ether at C-3 (quinsterol) resulted in a more potent compound, with prolonged activity. Stilbenes are probably the most important non-steroidal estrogens. Diethylstilbestrol (DES) is less expensive and as potent as estradiol and is also orally active. They appear to act by mimicking the action of steroidal estrogens at the estrogen receptors. Stereostructura of rings C and 0 showing the a and 0 position of 18 theC-17 hydroxyl group. Estrane CH.
18 13 16 14 15
H
Q
OH
m J o 6 & HO c * e . h o ^ ° ~ |
HO'
BiochaninA EOUOL ETHINYL ESTRADIOL
OH f " , -TV« *— O t S" rr'r V 7 ACH.CM, u .C M M O 'S / e^«*» C H ,C M >
Nemlnl
Figure 6 . Natural and synthetic estrogens
(Adapted from Henzl [78] and MacLachlan [79]) 21
1.2.4 ESTROGEN BIOSYNTHESIS AND METABOLISM
As previously mentioned estrogens and all other steroids arise from
cholesterol. Free cholesterol can be converted to pregnenolone through the action of an enzyme, side chain cleavage cytochrome P450 (P450scc). This is further converted to a C 1 9 androgen through the action of a 17, 20a-lyase. These androgens
may be aromatized to estrogens by the action of a specific cytochrome P450 enzyme known as aromatase (Figure 7a) through a three step conversion, which requires three moles of oxygen and three moles of NADPH. This enzyme is a heme-containing enzyme that aromatizes ring A of androgens to yield estrogens and formic acid. The reaction proceeds via the hydroxylation of the C-19 carbon twice to the dihydroxy steroid, which loses a mole of water to give the 19-oxo product. The last step is still not fully understood but results in the production of the aromatized estrogen and formic acid (Figure 7b) [80].
As previously mentioned, estradiol and estrone are interconvertible in vivo.
Any of these two estrogens can be hydroxylated at the 16-position by a
16-hydroxylase to give a series of 16a- and 16B-hydroxyestrogens [81], Another important route is the formation of catechol estrogens through hydroxylation at the A ring at C-2 or C-4 catalyzed by an enzyme estrogen 2/4-hydroxylase [82]. These can be further metabolized to form methyl ethers at C-3 to give methoxyestrogens. More recently, other metabolites have been detected such as 15a-hydroxylated compounds
[83]. All of these primary metabolic pathways are catalyzed by cytochrome P450 enzymes. Finally, secondary metabolic pathways include conjugated metabolites such as sulfates, glucuronides and thioethers (Figure 8 ). « CHOLESTEROL 7 6
OH OH
HO
PROGESTERONE ESTRADIOLTESTOSTERONE
b OB NADFB HO K 1 D P H H 0
18, 19-ditiydrsxyinarsatifiidion*IS-hydPSxyindPMtmadlDiM
KADFH
0 2 " x t I
[Enx]-0-00-0 Figure 7. Estrogen biosynthesis, a) General steroidogenesis b) Mechanism of aromatization (Adapted from Tepperman [46] and Brueggemeier [80]) 23 OH hydroxylation 2-hydroxylation % 4 -hydroxylation Conjugation 3-sulfates 3-glucuronides Figure 8 . Major routes of estrogen metabolism 1.2.5 THE ROLE OF ESTROGENS IN CARCINOGENESIS Almost one-third of all new cases of cancer in 1990 were hormone related [17]. Estrogens are associated with a group of these cancers such as breast, uterine and endometrial cancers. The first evidence of the involvement of estrogens in breast cancer came late in the nineteenth century when Beaston reported clinical improvement of breast cancer in women after castration [84]. Then in 1932, Lacassagne induced mammary adenocarcinomas in mice after long-term administration of estrone. In fact, risk factors for breast cancer include early menarche and late menopause. In postmenopausal women, obesity is another risk factor, due to a higher estrogen level in serum. In the first case, premenopausal women, the ovary is the principal source of estrogens. After menopause the ovary 24 produces little estrogen, but androgens are still produced in large quantities and are converted to estrogens by the aromatase enzyme in a variety of peripheral tissues including fats and adipose tissues [85]. This is supported by the relationship of plasma levels of estrogens to body weight and obesity. One of the striking observations is that the concentration of estrogens in premenopausal and postmenopausal women is similar in breast tissues, while there is a sharp decline in serum estrogens in postmenopausal women. Recent data suggests that normal breast epithelial cells may be regulated by progestins rather than estrogens [ 8 6 ]. Progestins are weak stimulators of cell growth in absence of estrogens but are inhibitory in their presence. It has been suggested that development of breast cancer arises from a switch of progestin to estrogen regulation [87]. These tumors may later progress to what are known as estrogen independent tumors, where they lose any steroid hormone sensitivity. In fact, only 30% of human breast cancers regress under endocrine treatment. This regression is usually temporary followed by the estrogen independent state. The presence of estrogen receptors (ER+) has been used as a good indication of whether the tumor is estrogen dependent. However, only 50% of ER+ cells respond to endocrine treatment (compared to about 5% of ER- cells) [85]. Recently other measures such as the presence of epidermal growth factor receptors [ 8 8 ] and progesterone receptors [89] have been used. The mechanism by which estrogens affect the tumor initiation and development is not fully understood. Involvement of estrogen metabolism in tumor initiation will be discussed in details later. Estrogens also induce a large number of enzymes and proteins that are involved in nucleic acid synthesis such as thymidine kinase and synthase, and they also stimulate DNA synthesis [90]. They induce 25 laminin receptors [91] and cathepsin D [92] both causing degradation of extracellular matrix contributing to the invasiveness of tumor cells. Estrogens also trigger the production of growth factors. This group of polypeptides is mostly mitogenic and stimulates cell growth and proliferation [93]. Finally estrogens induce protooncogenes and the production of oncogenic proteins such as c-myc, c-erb B and int-2. The expression of these proteins can cause transformation of cells, and they have been shown to induce tumors in animals [94]. In order to study the effects of estrogens in breast cancer, several systems have been used. One is the use of cancer cell lines such as the hormone-responsive MCF-7 cells [95] and the hormone-independent MDA-MB-231 cells [96]. These cultures facilitate the study of the direct effects on the tumor cells and avoid indirect effects that occur in vivo via other organs. Another system is the use of anchorage dependency as a measure for the tumorigenicity of the cells after various treatments [97]. Finally some animal models have been used. The advantage of using such models is the ability to study the overall picture in vivo and also to study the progression of the tumor using animals treated for various periods of time. One example of an animal model for estrogen-dependent cancer is the golden Syrian hamster. 26 L3 THE GOLDEN SYRIAN HAMSTER: AN ANIMAL MODEL FOR ESTROGEN-DEPENDENT CANCER: 1.3.1 INDUCTION OF RENAL ADENOCARCINOMA In 1944, Vasquez-Lopez reported the presence of a kidney tumor in golden Syrian hamsters treated with estrogens, but this was considered as a metastases from a pituitary tumor. Three years later, Matthews et al were able to demonstrate that these renal adenocarcinomas were a direct result of the estrogen treatment [98]. This was later confirmed by others including Kirkman and Bacon [99, 100, 101]. Spontaneous renal adenocarcinoma in golden Syrian hamsters are extremely rare. But upon treatment with DES for 250 days, these tumors develop in all animals [102]. Golden Syrian hamsters are the only species among rodents that show estrogen-dependent renal neoplasms, except for a hybrid line of mice where the tumor incidence is very low at only two percent [103]. Studies show that a wide variety of estrogens and estrogenic compounds can give rise to tumors, while inactive steroids such as estradiol-17a do not [102]. On the other hand, natural and synthetic steroids vary in their ability to induce tumor formation. Estradiol-178, the most potent natural estrogen, has the ability to induce tumors in all animals treated for at least 250 days. Diethylstilbestrol (DES), a synthetic non-steroidal estrogen, has the same effect and is actually used to induce renal tumors because it is inexpensive and easy to formulate into pellets. 17a-Ethinylestradiol (EE), a potent synthetic steroidal estrogen gives rise to tumors in only about 15% of the animals treated, while the addition of an 118-methoxy group to 27 EE (moxesterol) renders the compound highly tumorigenic, with an incidence rate of 90% [104]. Estrogen is required for both the initiation and the development of the tumor. The animal has to be constantly exposed to estrogen, and if the treatment is stopped at any point, the tumors will regress. If estrogen treatment is then continued even after 200 days, the tumors are regenerated. When the tumors are transplanted into other animals, they produce neoplasms only if the host has been primed with estrogen for at least 45 days [105]. Metastases of the primary tumor usually to the abdominal cavity, occurs in about 45% of the cases, although metastases to other organs have been reported [106]. While age differences did not affect the incidence of tumor formation [107], the sex of the animal was important. Intact males developed renal adenocarcinoma when treated with estrogens, while females had to be ovariectomized to develop these tumors [108]. The reason for this discrepancy was that the circulating progesterone in females reversed the effect of estrogens. In fact, administration of progesterone or other steroid hormones such as androgens and glucocorticoids prevents tumor formation in Syrian hamsters [105]. Antiestrogens were also found to prevent tumor formation when administered with estrogens [109]. Subsequent studies demonstrated that this effect correlates with the ability of the compound to inhibit the binding of estrogens to their receptors, since nafoxidene, an antiestrogen that inhibits estrogen binding to its receptor, prevented tumor formation, while MER-25, another antiestrogen that acts by a different mechanism, has no effect on tumorigenesis [ 1 1 0 ]. Studies have also proved the ability of antiestrogens to cause tumor regression in tumor bearing animals [109]. 28 The kidney is not a target organ for estrogens, but estrogen receptors have been detected in rats and mice kidneys [111]. In the Syrian hamster, the concentration of estrogen receptors is very low, but increases about 3.5-fold with estrogen treatment [112]. The concentration of progesterone receptors is negligible. The levels of progestrone receptors rise with estrogen treatment, reaching 13-17 fold after about two months of treatment and 45-50 fold in the primary tumors [113]. This increase in progesterone receptors is also inhibited by antiestrogens and androgens [114]. The tumors can be transplanted into other hamsters pretreated with estrogen. These tumors remain estrogen-dependent up to at least twelve passages. Like the primary tumors, they are inhibited by antiestrogens, but not androgens. The latency period for the first transplant can vary from 4-12 months and is reduced with successive transplants [101]. The size, as well as estrogen and progesterone receptor concentrations, were reduced during successive transplantation. Recently, Lin and co-workers were able to shorten the latency period to two months and increase the tumor mass 20-30 fold by transplanting the primary tumor beneath the renal capsule with no changes in progesterone receptor levels [115]. Li and Li were able to duplicate this feat by injecting the tumor cells intraperitoneally, but these tumors had higher estrogen and progesterone receptor levels [116]. A permanent cell culture line from the primary renal tumors of Syrian hamsters has been established and called H-310 line [117]. This cell line is an estrogen-induced and estrogen-dependent tumor cell line. The growth of the cell line was not stimulated by estrogens; however, it was stimulated by the addition of fetal calf serum to the media that already contained calf serum. This stimulation was absent if the serum is heated to 90°C [117]. Similar results were obtained when 29 treating the cell line with liver or kidney extract from hamsters treated with estrogen, but not from untreated animals. This stimulation was again inhibited by heating to 100°C [118]. A polypeptide growth factor that is induced by estrogens was suggested as the growth stimulating agent [119]. Another set of experiments showed that charcoal-dextran-stripped Syrian hamster serum inhibited the growth of the cell line H-301, and this effect was reversed by the addition of estrogens. The authors concluded that the tumorigenic property of estrogens in the Syrian hamsters is due in part to inhibiting a growth inhibitory factor present in hamster serum, called estrocolyone [ 1 2 0 ]. 1.3.2 MORPHOLOGY AND HISTOLOGY OF THE TUMORS The morphology, anatomy and cell origin of the renal adenocarcinoma is a controversial issue. Early researchers proposed that the tumor is composed of epithelial cells, derived from the proximal or distal tubules [99, 101]. More recently, this view has been shared by Goldfarb and Pugh [121]. They showed that the dysplasia prevalent in these tumors was greatly concentrated in the inner cortex and outer stripe of the medulla. They finally suggested that stromal cells facilitate the transformation of proximal tubule epithelial cells leading to tumor formation. The second view states that the hamster kidney tumor originates from mesenchymal components [122, 123]. Hacker et al studied the cell of origin histologically and histochemically [124]. They concluded that the proliferating foci are formed of spindle-shaped cells located between tubules and not within the tubules themselves. These foci, the precursors of the larger tumor, matched the larger tumors as well as metastatic tumors [125], histologically and histochemically, as shown by the high 30 activity of alkaline phosphatase, adenyl cyclase and glucose- 6 -phosphate dehydrogenase, and absence of 5-glutamyl transpeptidase and glucose- 6 -phosphatase. Both the tumors and the foci also expressed vimentin and desmin but not cytokeratin. They thus concluded that the tumors had a mesenchymal interstitial cell origin and probably were derived from vascular smooth muscle cells. The third view disagrees with the mesenchymal cell origin theory [126]. Morphological studies of the early lesions and the tumors showed that they were composed of two types of cells, large epithelial cells, and small poorly differentiated cells that still had epithelial characteristics (Figure 9). The larger cells had cilia, microvilli and intracytoplasmic lumens, which are typical epithelial features. These large cells were similar to early metanephric tubules, while the small cells appeared to be related to the blastemal cells of the developing kidney. Finally, they reported that the early lesions were found in the kidney interstitium close to large arteries, leading to tumor formation in both the renal tubules and interstitium. Histochemically, they were able to demonstrate the presence of desmin, vimentin and, in contrast to Hacker's studies, cytokeratin, which means that the intermediate filaments were of epithelial and mesenchymal characteristics. They concluded that the cell of origin is a cell that is committed to an epithelial differentiation pathway. Further studies by the same group using light and electron microscopes and immunoperoxidase analysis demonstrated that DES induced interstitial lesions, while EE, that caused a low incidence of tumor formation, induced a tubular lesion [127]. They deduced that the interstitial, but not the tubular lesions were preneoplastic due to the similarity of the renal tumors to these lesions morphologically and histochemically, and that co-treatment with both estrogens, DES and EE led to the absence of both the early interstitial foci and adenocarcinoma, thus physical injury glomerular cells or Stages in Differentiation estrogen- induced progenitor renal cell damage cortex blastemal-like ( toxicity) stem cell mature reactive reactive proximal tubular cells Renal Tumor Cells small blastemal- large like epithelial tumor cell tumor cell Figure 9. The origin of kidney tumors in Syrian hamsters (Adapted from Li et al [126]) 32 intact castrated castrated+DES intact castrated castrated+DES Figure 10. Appearance of dysplasic foci (a) and increase in area of vascularization (b) in kidneys of estrogen treated hamsters confirming the earlier studies that deduced that the origin of the tumor is the renal interstitium and not the proximal tubules. Dysplasic foci have been reported to appear somewhere around three months of estrogen treatment, where they were detected in 65 % of kidneys from castrated hamsters treated with DES. The same study showed an increase in the area of vascularization after the same period of treatment [128] (Figure 10). 33 1.3.3 THE ROLE OF HORMONAL AND CARCINOGENIC ACTIVITY OF ESTROGENS IN TUMORIGENESIS Early investigations of estrogen-dependent cancer have attributed the tumorigenic property of estrogens solely to its hormonal effects, i.e., binding to its receptor, leading to a cascade of events resulting in cell growth and proliferation. While this is a very important aspect of the process, only recently have researchers appreciated the carcinogenic property of estrogens and their role in cancer, which arises from metabolic activation and the production of reactive intermediates. The Syrian hamster model has enabled the study of both properties and their importance in estrogen-dependent tumors. The hormonal effect can be visualized by the presence of estrogen receptors and the increase in these receptors upon estrogen treatment [ 1 1 2 ]. Another evidence is the effect of antiestrogens, that act by competitive binding to the estrogen receptor, to inhibit tumor formation, and the inability of inactive estrogenic compounds such as estradiol-17a to induce tumors. The first nonhormonal aspect of the tumorigenesis process came from studies that showed that administration of or-naphthoflavone, an inhibitor of monooxygenases leads to prevention of renal adenocarcinoma [129]. Further studies showed that other estrogen metabolism modulators and inhibitors, such as 2 -fluoroestradiol and vitamin C, had the same effect [130]. The presence of DNA modifications and adducts following estrogen treatment of the hamsters were demonstrated [131], and these were specific to kidney tissues [132] and postulated to arise from activated metabolites and/or free radicals formed during metabolic pathways. Enzymes responsible for estrogen metabolism were found to exist primarily in the cortex, the site of tumor formation, and a 7-fold higher rate of catechol estrogens are formed in the kidneys in 34 hamsters as compared to rats. These observations prompted Li and Li to study the ability of different estrogens to induce renal adenocarcinoma in hamsters and compare their competitive binding to the estrogen receptor and their ability to be metabolized through the catechol estrogens pathway in vivo in this model (Table I). These studies showed a good correlation between tumorigenesis of estrogens and their ability to form catechol estrogens. So while EE is a very potent estrogenic compound, it cannot form catechol estrogens in hamster kidneys and produces tumors in only 2 0 % of treated hamsters. The same holds true for other potent estrogens such as a-zearalanol, although it was reported later that a potent estrogen, mexosterol, gives rise to a high incidence of tumors, but cannot form catechol estrogens [133]. Thus, tumorigenicity in Syrian hamsters appears to be a complicated process. It may involve the formation of metabolism of estrogens that may lead to the formation of reactive intermediates and possibly DNA adducts or bind cellular proteins. It also involves hormonal effects of estrogens that lead to cell proliferation directly or indirectly by activation of cellular oncogenes or the production of polypeptide growth factors that may in turn participate in the tumorigenic process (Figure 11). 35 Table I. Correlation of the carcinogenic and hormonal activity of estrogens with their ability to induce tumors in Syrian hamsters (Adapted from Li et al [133]) Combined number oi No. of animals tumor Induction of with tumors/ nodules in Competitive progesterone total no. of SE with both Estrogens* binding4 receptor* animals tumors kidneys * Steroidal estrogens 170-Estradioi 89.8 ±0.7 48.5 ±6.3 6/6 100 18.0 ±4.0 Estrone 85.2 ±1.3 46.9 ±1.2 8/10 80 10.4 ±5.0 Estriol 84.0 ±2.0 35.0 ±2.5 4/7 57 2.7 ±2.0 Z-Hydroxyestradiol 81.2 ±0.8 — 0/6 0 0 2-Hvdroxyestrone 81.5 s 6.9 __ 0/6 0 0 4-Hvdroxyestradiol 87.8 ±2.9 5/5 100 15.6 ±2.0 4-Hvdroxyestrone 79.0 ±2.3 — 2/6 33 1.5s 1.5 Ethinyl estradiol 90.5 s 1.1 56.1 ±3.3 3/15 20 0.6 ±0.5 Equilin 78.4± 1.7 27.1) ±2.0 6/8 75 5.5 s 0.9 d-Equilenin 49.2 ±5.5 11.5 ±2.0 0/9 0 0 Stilbene estrogens DES 90.1 ±0.9 51.7 ±3.2 10/10 100 22.0 ±3.0 d-Dienestrol 8.0 ±0.7 ND* 0/10 0 0 OES 3.4-oxide 90.0 ±1.6 __ 6 /7 86 11.4 ±3.0 a-Dienestro! 88.6 ±1.4 35.6=2.9 7/8 86 10.4 ±3.0 Hexestroi 90.4 ±0.8 48.5 ±1.6 6/6 100 25.8 ±6.0 Estrogen mycotoxins a-Zearaianoi 82.3 ±3.0 45.4 ±5.6 1/8 12 0.2 ±0.0 £-ZearaIanol 48.0 ±4.0 17.7 ±4.0 0/8 0 0 Zearalanone 48.0 ±6.0 7.8 ±1.9 0/8 0 0 Zearalenone 43.0 ±5.0 6.4 ±0.9 0/8 0 0 Phvtoestrogens Coumestrol 51.0 ±2.0 10.2 ±2.0 0/8 0 0 Biochanin A 0 6.7 ±5.3 0/8 0 0 • Duration of estrogen treatment was 9.0-10.0 months. After initial pellet implantation, additional estrogenic compound pellets were implanted every 3.0 months, except for estrone, cstriol. 2-hydroxrest radial, 2-hvdroxv- estrone. ethinyl estradiol, and zeanlenone. which were implanted every 2.0 months. * Competitive binding of radioinen estrogens for estrogen receptor was carried out on cytosols obtained from hamster estrogen- induced renal carcinomas, with 5.0 nM (5H]estradiol alone or in combination with competitive nonradioactive compounds (5.0 X 10*7 m). (sH]EttradioJ concentration in these cytosols, without competitor, corresponded to 0% inhibition. Values are expressed as meanSSCM. ‘ Results are expressed as fcmtomoics per milligram of cytosol protein, meansSEM. * Values represent the mean number of tumor fod per animal in each group ±SEM. ' ND. not detectable. 36 o n c o s c n c . ACTIVATION* ____ (GROWTH FACTORS) — . / ^ u m n s o / HORMONAL » (vti-rrn HORMONAL OES «t5S*l> urfmfO CARLT NEOPLASTIC RINAL RCT TUBULE CELLS " tN ,L TUiULC * * " £ 1 2 ? INFILTRATING AMO r o e ( r 0 C | TUMOR PRIMITIVE • CRRANJION INTERSTITIAL I / CfLL? RgpARATIVE NYPCRPLASIA ' , | nom -> r c c it ic I REACTIVE METABOLITE I I I / \ / FREE RAOICAL FORMATION \ / \ * I 0 TOKicirr KT CCU(OAMAAC) Figure 11. Different factors participating in the tumorigenesis in Syrian hamsters (Adapted from Li et al [134]) 1.4 GROWTH FACTORS. ONCOGENES AND CELL TRANSFORMATION 1.4.1 GROWTH FACTORS: GENERAL ASPECTS In 1962, Stanley Cohen described the isolation of a compound that accelerated eyelid opening and teeth eruption in newly bom mice, and called it epidermal growth factor (EGF) [135]. Now, many polypeptide growth factors are known and being studied in different systems, species and cell types and are implicated to participate in many physiological processes. In very simple terms, a growth factor is a substance that when added to a cell in a test tube causes it to divide once. These growth factors are multifunctional, depending on the species, body site and cell type, and are mostly but not always mitogenic and growth stimulating. They appear to be mainly a way for the cell to communicate with itself or its surroundings. They can be considered as the alphabet of the biological regulating language, the grammar of which we are trying to study [136]. Multicellular organisms have highly coordinated mechanisms to control cellular interactions. In regard to growth control, complex signaling networks mediate normal embryonic development and are responsible for systemic responses to wounding and infections. These effects are achieved by factors that can act in a positive or a negative sense to affect cell proliferation and differentiation. These factors lie in two major categories, hormones and growth factors. Growth factors may act in an autocrine fashion, affecting the same cell that secretes them, in a paracrine fashion affecting neighboring cells, or even in a telecrine manner acting at a distant site from the origin of the secretion. The action of growth factors as mitogenic agents is believed to allow cells in the G 0 phase to enter and proceed through the cell cycle (Figure 12). Accordingly, growth factors are either "competence factors" that help the cell traverse the Gj phase, such as epidermal growth factor (EGF), transforming growth factor-a (TGF-a) and fibroblast growth factors (FGFs), or "progression factors" such as insulin-like growth factor-I (IGF-I) that commit the cell to DNA synthesis. A critical point in the cycle exists where the presence of both factors are essential for the cell to progress towards mitosis. Many oncogenic products can mimic the action of the competence factors, while transforming growth factor-fl (TGF-fl) and interferons can inhibit these proliferative effects even very late in the Gt phase. The disruption of the signal, even for a short period of time, causes the cell to revert to the G 0 state, and the absence of growth 38 factors can even lead to apoptosis or programmed cell death in some types of cells [137]. EGF PDGF FGF IGF-1 INSULIN TGFP Inhlbltlei Figure 12. The action of growth factors on the cell cycle (Adapted from Aaronson [137]) Growth factors have been shown to exert their effects by binding to a cellular receptor. These receptors have an extracellular domain that binds the ligand and an intracellular domain which acts as a kinase, specifically phosphorylating tyrosine moieties, an event that leads to transducing the mitogenic signal [138]. After the ligand binds, the receptor forms dimers or oligomers that lead to the activation of the kinase function [139]. The tyrosine kinase sequence is highly conserved and is 39 absolutely required for receptor signaling as demonstrated by a point mutation on the adenosine triphosphate (ATP) binding site that led to the inability of the receptor to cause phosphorylation and abolished biological activity [140]. The receptor itself can be auto-phosphorylated, causing its own inactivation [141]. To understand how growth factors regulate cell growth, we have to answer an important question: how does the signal that is initiated by the binding of a ligand to a cell surface receptor reach the final target, the DNA in the nucleus? Shedding light on the substrates for the tyrosine activity may help us answer this. The substrates include phospholipase C (PLC) [142], phosphatidylinositol-3'-kinase (PI3K) [143], guanosine triphosphate activating proteins (GAP) [144] and src and src-related tyrosine kinases [145] (Figure 13). P1dln*(3)P g u b ili H i Figure 13. Substrates of growth factor receptors (Adapted from Aaronson [137]) 40 - PLC hydrolyzes phosphatidyl inositol 4,5-bisphosphate (PI-4, 5 P2 ) and generates two secondary messengers rapidly after the ligand binds. The first, inositol triphosphate (IP 3 ), releases stored intracellular calcium and the second, diacylglycerol (DAG), activates protein kinase C [146]. - PI3K phosphorylates the 3'-position on the inositol ring of phosphatidyl inositol (IP) and this has been implicated in transformation. - GAP regulates the function of ras protein [147], a critical component of the mitogenic signaling pathways, which is achieved by the phosphorylation of GAP, that leads to relieving its inhibition of the ras function. - The src and related genes are protooncogenes which encode non-receptor tyrosine kinase proteins [148], that are stimulated by growth factors leading to an increase in the phosphorylation of other kinase substrates. Another protooncogene, ra/is also activated by growth factors. The protein product is a serine-threonine kinase [149] which phosphorylates unidentified substrates that are believed to play a role in cell proliferation. Other reports suggest roles for a guanosine triphosphate binding protein, ion-channels participation, and activation of adenylyl cyclase in the signal transduction process, but these are less well studied [137, 150]. Different growth factors have specific substrates that they phosphorylate. IGF-I, a progression factor in the cell cycle, phosphorylates different substrates from competence factors. And even within this latter group, platelet-derived growth factor (PDGF) has been shown to interact with all of the mentioned substrates, while colony stimulating factor-I (CSF-I) does not phosphorylate PLC or GAP [137]. EGF and TGF-a can not phosphorylate GAP efficiently, while FGFs phosphorylates another substrate, P90 which has not been identified and has not been shown to be 41 phosphorylated by any other growth factors [151]. This variation in substrate specificity has been postulated to be responsible for the differences of the actions of growth factors on the same cell, and the ability of only some to cause cell differentiation. Thus these secondary messengers and oncogenic products appear to function as the signal delivering system to the nucleus leading to the initiation of DNA synthesis. Yet our knowledge is incomplete, and more information must be obtained in order to identify the full spectrum of how these growth factors are able to cause mitogenic signaling and other cellular responses. This is particularly important since growth factors have been implied to play a role in carcinogenesis, and it is postulated that malignant cells arise as a result of the progression of genetic events that include unregulated expression of growth factors and their receptors. 1.4.2 GROWTH FACTORS: A CLOSER LOOK Epidermal growth factor (EGF) was the first polypeptide growth factor isolated. It is a small single chain polypeptide of 53 amino acids and has a molecular weight of about 6 kDa [135]. It stimulates the proliferation of epidermal and epithelial cells in various species and cell cultures [152]. The EGF receptor is a tyrosine kinase that causes phosphorylation of many proteins including phosphoinositol leading to a cascade of unknown events that ends in stimulation of RNA and protein synthesis and committing the cell to DNA synthesis [153]. The kinase activity is absolutely required for activity, since binding of EGF to a mutant receptor that binds the growth factor but possess no kinase activity failed to generate a mitogenic signal [140]. This growth factor appears to act in autocrine or paracrine fashions, since its concentration in the plasma is very low (1 ng/ml) [154]. In 42 humans, the concentration of EGF is 5-fold higher in the first two years of life compared to adults, and may be important in the development of the newborn, and in wound healing [155]. A related molecule is the transforming growth factor-a (TGF-a), that was discovered in the conditioned media of sarcoma virus transformed cells [156]. This polypeptide is synthesized as a large molecule of a molecular weight ranging between 20 and 26 kDa depending on the species and is then processed to smaller polypeptides. In humans, it exists as 25, 21 and 17-19 kDa precursors that are processed to a 7 kDa form. It is structurally related to EGF, and binds to the EGF receptor more tightly than EGF itself, but antibodies against EGF do not neutralize TGF-a [157]. TGF-a has a wide range of physiological activity, that ranges from enhancement of bone resorption to inhibition of gastric acid secretion, and helps in wound healing [158]. It causes cell proliferation and is secreted from many epithelial tumor cells such as MCF-7 cells [159], from which TGF-a mRNA can be detected after 6 hours of estrogen treatment and antibodies directed against TGF-a causes growth suppression in these cells [160]. Transforming growth factor-8 (TGF-8) is a 25 kDa polypeptide found as a dimer of identical chains, linked by disulfide bridges [161]. It binds to a specific receptor that possess tyrosine kinase activity, which differs structurally and functionally according to the cell type and species. This growth factor is produced by cells of nearly every origin [162], and is growth inhibitory in most cells such as many epithelial cell lines in vitro and in developing mammary glands in vivo, although cell growth stimulation has been observed in some cell types including mesenchymal cells and fibroblasts [163]. Another important effect is its induction of changes in the extracellular matrix composition, that may affect the rate of cell proliferation. It 43 stimulates the production of all major matrix proteins such as collagen, fibronectin and elastin [164, 165] via an increase in the transcription rate and the stabilization of mRNA. It also decreases the matrix degradation by inhibition of proteases and antagonizing other factors that induce the production of these enzymes [166]. It stimulates wound healing [167] and bone resorption [168]. The levels of TGF-fi are lower than normal in some tumor cell lines such as MCF-7 cells, while in others TGF-fi receptors are absent supporting the notion that TGF-fi has a role in regulating cell growth and proliferation [169]. Fibroblast growth factors are classified into an acidic form (aFGF) and a basic form (bFGF). They both have the same spectra of activity, but the basic form is 10-100 fold more potent [170]. The acidic form is a 14-16 kDa polypeptide found mainly in the brain. The basic form has a molecular weight of 16-18 kDa, and has a highly conserved sequence, where human and bovine bFGF differ in only two out of 155 amino acids [171]. The FGF receptor binds both forms, leading to an activation of a tyrosine kinase domain and a wide range of biological activity. These growth factors stimulate cell proliferation in many normal cells [172] and modulate cell differentiation, preventing differentiation of cultured myoblasts [173] and stimulating it in neuronal [174] and adipose cells [170]. It appears to play a role in early embryogenesis as evident by the high amounts of bFGF found in early embryos [175], fetal brain [176] and fetal adrenal cortex [177], which then declines with aging. It also stimulates wound healing partially due to its angiogenic property or its ability to stimulate the formation of new capillaries [178], a phenomena that may be important for the well-being of solid tumors that need vascularization to survive and proliferate, thus the implication of bFGF in the tumorigenic process of carcinomas. 44 Platelet-derived growth factor (PDGF) is a glycoprotein with a molecular weight of 30-34 kDa, and is composed of two different polypeptide chains A and B connected by disulfide bridges to give rise to three possible combinations [179]. This growth factor binds to two different but closely related receptors that exhibit tyrosine kinase activity. It is a major mitogen in serum for connective tissue derived cells and stimulates cell proliferation of mesenchymal cells [180], as well as many other effects such as vasoconstriction [181], chemotaxis and promotion of polymorphonuclear leukocytes and monocytes [182]. It plays a role in embryogenesis and wound healing [183] and is secreted from a large number of transformed cells [184]. Insulin-like growth factors (IGFs) are a family of peptides that have a high homology (about 50%) in its amino acid sequence with proinsulin. IGF-I shares 48% of its 70 amino acids with proinsulin (Figure 14), while IGF-II shares about 50% of its 67 amino acids [185]. The B domain in these polypeptides is necessary for binding to the three types of a tyrosine kinase IGF receptor. One type of receptor binds IGF-I more tightly, while IGF-II possess higher affinity to another, and insulin binds to all of these receptors, but with less affinity [186]. IGF-I and II are secreted from many types of cells and like all other growth factors have a variety of biological activities that include cell growth, and insulin-like effects such as lowering blood glucose levels [187]. 1.4.3 ONCOGENES AND TUMOR SUPPRESSOR GENES Cell proliferation is believed to be governed by two genetic effects, growth promotion by protooncogenes and growth constraint by tumor suppressor genes or antioncogenes. Carcinogenesis is thus a deviation from the normal cellular 45 g-Oomoin ^gjASPfLEulARGARG, PROLU A#*"***? A-Oomain \gly PRO V he .YS ASN PRO PROLYS i l / ALA /TR57 V / THR GLY " g l n tyr pro ~ala ^ __^.gly I-A'~ARG__ARG_SER_SER—SER C-Domain Figure 14. Amino acid sequence of IGF-I Boxed amino acids are shared with proinsulin (Adapted from Clemmons [187]) mechanism by overexpression of protooncogenes or mutations converting them to oncogenes and inactivation of tumor suppressor genes, with evidence pointing out that both events have to occur for malignancy to arise [188]. Oncogenes are defined as genes that cause cancer. They were first detected as dominantly acting transforming viral genes that cause tumors following viral infection of animals [189]. It was then discovered that these genes are mutations of normal cellular genes, and that they play an important role in tumorigenesis based on the finding that 30% of human cancers contained activated oncogenes [190] and that the protein products of oncogenes are related to growth factors and their receptors [188]. 46 Tumor suppressor genes are defined as genes related to the suppression of cell proliferation [191]. The first experiment to point out the importance of the loss of the function of a gene in tumorigenesis, showed that malignant cells fused with normal cells resulted in loss of malignancy, thus introducing the theory that tumor suppression arises from genes present in the normal cells that replaced the defective function in the malignant cells [192]. While identification of oncogenes is relatively easy by their transforming ability, tumor suppressor genes are much harder to identify as they have a negative role in transformation. Thus, the presence of tumor suppressor genes are monitored blindly by hybridizing genes, locating those that have no known functions, and then examining their presence or loss in tumors [193]. Many oncogenic products act as tyrosine kinases, a property that leads to mitogenic transduction in many cells, and have a high homology with growth factors and their receptors. For example, the erb-B protein has about a 95% homology to the EGF receptor, and v-sis product is similar to the PDGF B chain [194]. Erb-B is often overexpressed in adenocarcinomas of the breast, stomach, and ovary, and its overexpression can confer transformed phenotype [195]. Ras mutations have been shown in 30-50% of lung and colon cancer cases, while it is present in 90-95% in pancreatic cancers [196]. Myc overexpression was demonstrated in lung cancer, and its mutation in B and T cell lymphomas [197]. These effects usually denote a more aggressive type of cancer. The first tumor suppressor gene identified was the retinoblastoma (RB) gene located on chromosome 13 [198]. Since then, others have been located such as P53 gene and erb A gene. The mechanism of action of these genes is currently unknown, but in general cells are believed to shut down growth via one of three methods: one is a holdup at the end of the Gx phase in the cell cycle, as illustrated by the inhibition 47 caused by TGF-fi, the second is undergoing irreversible postmitotic differentiation and the third is apoptosis or programmed cell death. How the tumor suppressor genes fit within these responses remains to be fully determined. 1.4.4 THE ROLE OF GROWTH FACTORS, ONCOGENES AND TUMOR SUPPRESSOR GENES IN BREAST CANCER The hormonal effects of estrogens include modulation of gene expression and possibly the stimulation of the production of polypeptide growth factors. Receptors for EGF as well as the secretion of TGF-a have been shown to be stimulated by estrogens in vitro [199], and detected in virtually all breast cancer cell lines derived from epithelial tumors including MCF-7 cells [200], and estrogen-independent cancer cells such as MDA-MB-231 [201] at levels much higher than normal. They were also found in biopsies of human and rodent mammary tumors [202] and in urine samples from breast cancer patients [203]. Antibodies against TGF-a or EGF receptors cause growth suppression in MCF-7 cells [160]. TGF-a and EGF are growth inducers even in the absence of estrogens. In mice, the removal of the submandibular gland, the major source of these growth factors, lowered the incidence of tumors and the growth rate of breast tumors in this animal while the injection of EGF returned the incidence rate and the growth rate back to normal levels [86]. IGF-I was also detected in human breast cancer cells [204], Its secretion was increased 3-6 fold after estrogen treatment, and inhibited by antiestrogens [205]. Antibodies against this growth factor have been shown to inhibit the growth of MDA-MB-231 cells in vitro [206] and tumor growth near mammary ducts in vivo [207]. Less information is available on PDGF secretion from breast carcinomas. While it was detected in some breast cancer 48 cell lines [208], PDGF has not demonstrated growth regulation of these cells. FGF is required for the normal growth of mouse mammary cells in culture [209], and most breast cancer cell lines express FGF activity [210]. TGF-fi has been shown to be secreted by breast cancer cells [211] but it is believed to be a growth inhibitory factor. Its secretion is inhibited by estrogens, and stimulated by antiestrogens, and its growth inhibitory effect was reversed by antibodies directed against this factor [169]. Normal breast epithelial cells are inhibited by TGF-fi as well, and implants of TGF-8 inhibit the mammary ductal development, but not the distant mammary glands [212], which implies that it acts in an autocrine or possibly a paracrine fashion. It inhibits the growth of breast cancer cell lines including the estrogen-independent MDA-MB-231 [169]. Growth factors are thus believed to control the growth of normal and cancerous breast cells. The deviation of breast tumors from estrogenic control implies the presence of growth control elements other than estrogens that can take over. This has led to the hypothesis that growth factors may play that role, especially since they control growth in absence of estrogens. Other suggested roles for growth factors include the regulation of extracellular degradation that precedes the invasive and metastatic stage of cancer cells, and helping in the vascularization of the tumors [86]. Three oncogenes have been observed to be amplified in primary breast tumors, c-myc, c-erb-2 and int-2. The c-myc encodes two nuclear proteins [213], that cause mitogenic stimulation of the cell followed by terminal differentiation and is believed to play a role in the transition of the cell from G0 to the Gj phase in the cell cycle [214]. Amplification and overexpression of c -myc is seen in about 30% of primary human breast cancer and correlates with poor prognosis [215]. In mice, its overexpression caused tumors in about 3 months [216]. The oncogene c -erb-2 is 49 activated by a point mutation altering a valine to a glutamic acid [217], expresses a protein that has a high homology with the EGF receptor, and possesses tyrosine kinase activity [218]. It is activated in about 25 % of breast cancers and also correlates with unfavorable prognosis [215]. The activation of this oncogene also leads to a high tumor incidence in mice [219]. Amplified int-2 is found in 10-20% of breast cancer cases [220], and its protein product exhibits a very high homology to bFGF [221]. The inactivation of four tumor suppressor genes have been identified in breast cancer cells: the retinoblastoma gene, P53 gene, the DCC gene and the nm23 gene [94]. These three groups, growth factors, oncogenes and tumor suppressor genes appear to be good candidates for therapeutic targeting. The inhibition of growth factors and their receptors and the subsequent cell growth inhibition was accomplished by using specific antibodies both in vivo and in vitro , and tumors that overexpress receptors can be targeted by toxins linked to the antibodies [222]. Erbstatin, a specific tyrosine kinase inhibitor, and the related tyrphostine caused cell proliferation inhibition in cultures [223, 224]. Suramin, a polyanionic compound, has shown growth inhibition in many breast cancer cell lines [225], with a decrease in the secretion of most growth factors. Other approaches that have yet to be examined include specific receptor antagonists, and genetic therapy by supplying a lost tumor suppressor gene or the activation of a latent gene. 50 1.5 ESTROGEN METABOLISM 1.5.1 OXIDATIVE METABOLISM OF ESTROGENS Estrogens can undergo phase I and phase II metabolism in vivo. The liver is the main site of metabolism, although other organs have been reported to play a role such as the kidneys and the small intestine. Phase I metabolism of estrogens includes oxidation, hydroxylation, and methylation. Phase II leads to the formation of glucuronides, sulfates, and thioethers, in addition to further oxidation of estrogens to polyhydroxylated compounds. Oxidative metabolism is the most important phase I reaction. Oxidation of estradiol to estrone is one pathway, and hydroxylation on ring D at C-15 and C-16 constitutes another pathway. However, the pathway that has been implicated most to play a role in the carcinogenic property of estrogens is the formation of catechol estrogens and their further metabolism. Aromatic hydroxylation on ring A in estradiol can take place ortho to the C-3 phenolic hydroxy group at either C-2 or C-4 to give the 2-hydroxy and 4-hydroxyestradiol respectively, through the action of an enzyme, estrogen 2/4 hydroxylase. This enzyme is a mixed function oxidase from the cytochrome P450 family, and requires NADPH and oxygen as cofactors [82]. These catechols can be further metabolized through various pathways. Formation of glucuronides and sulfates have been reported and these secondary metabolites are very water soluble and can be easily excreted. The enzyme catechol O-methyl transferase (COMT) is present in many tissues and leads to the methylation of the phenolic group to the methyl ether [226]. Reports of further metabolism of catechol estrogens to semiquinones and quinones have been accumulating [227, 228], and it is through this pathway that different reactive metabolites and free radicals are generated and may bind to various cellular proteins and cause DNA damage [229]. Finally, an enzyme catechol 1,2-dioxygenase has been isolated from some species and shown to oxidize catechols to a muconic acid derivative [230], presumably through an addition of an oxygen molecule across the double bond between the hydroxyl groups and the subsequent breakage of the aromatic ring to yield the diacid (Figure 15). ■OH +O2 ■OH *" Figure 15. The oxidation of catechols to the muconic acid (Adapted from Nozaki et al [230]) Catechol estrogens exhibited very weak estrogenic activity and were thus considered a method of inactivation of potent estrogens. This idea was first challenged when it was reported that catechol estrogens existed in high amounts in brain tissues [231] and were later found to act as inhibitors of the enzymatic methylation of catecholamines and may have an important biological role [232]. Estrogen 2/4 hydroxylase has been shown to be specific to aromatic estrogens. Testosterone and progesterone are not substrates for this enzyme, while other synthetic estrogens such as DES are excellent substrates [233]. The mechanism of the 52 reaction has been studied, but is still not totally understood. It has been established that the reaction proceeds without the migration of hydrogens in an NIH shift [234] and has no isotope effect [235]. LeQuesne et al proposed that the reaction goes through the formation of an epoxide by the attack of cytochrome P450 on the aromatic ring, which can tautomerize to the epoxyenone and then rearomatizes to the catechol, presumably through the attack of a base on the proton at C-2 and epoxide opening [236] (Figure 16) Figure 16. Mechanism of catechol estrogen formation (Adapted from LeQuesne et al [236]) The epoxyenone formed in this reaction was reported to be stable enough to leave the active site and exert biological effects [236]. The same researchers also reported the isolation of 1,2-epoxide-enone estradiol when estradiol was incubated with microsomes from MCF-7 cells in presence of I,2-epoxy-3,3,3-trichloropropane, a potent epoxide hydrogenase inhibitor [236], and others reported the isolation of 53 3,4-epoxyenones from other cell lines. LeQuesne et al synthesized different epoxyenones and reported that they have the ability to induce transformation of Balb C/3T3 cells with the 4a,5a-epoxyenone being the most potent transforming compound [237]. Binding of catechol estrogens and their oxidative metabolites to proteins have been reported [229]. The evidence for these adducts comes from experiments showing that cytochrome P450 inhibitors decreased the total amount of estrogens that bind to proteins. The reactive species is believed to be an oxidative metabolite of catechol estrogens, probably quinones or semiquinones [227], the formation of which was demonstrated using electron spin resonance (ESR) [238]. The formation of both 2,3- and 3,4- quinones of estradiol has been shown and it appears that the latter is more susceptible to nucleophilic attack and protein binding, because the electrophillic positions on the 2,3-quinone are sterically hindered [238]. Estradiol metabolites have been shown to irreversibly bind to DNA in rat hepatocytes and liver microsomes [229], and recently DNA damage was shown to be induced by estrone 3,4-quinone in MCF-7 cells [239]. 1.5.2 ESTROGEN METABOLISM IN SYRIAN HAMSTERS The recognition that estrogen metabolism plays a role in the tumorigenic process leading to the formation of renal adenocarcinoma in Syrian hamsters has prompted many researchers to study the different pathways involved. Both the liver and kidneys of the hamster contain the enzyme estrogen 2/4 hydroxylase. In the liver, estradiol is oxidized almost exclusively to the 2-hydroxy derivative, while in the kidneys equal amounts of the 2- and 4- hydroxy estradiol are formed [240]. The kinetics of estrogen 2-hydroxylation in the kidney microsomes were studied and the 54 catalysis was reported to have an apparent Km of 6.43 /*M and apparent Vmax of 0.051 nmol/min/mg. In animals treated with estrogens, the apparent and Vmax were very high with high standard deviations possibly due to a decrease in estrogen 2-hydroxylase [241]. This enzyme has been shown to be inhibited by haloestrogens [242], and by other steroids such as androgens and progestins [243]. The methylation of catechol estrogens is a mean for clearing these metabolites, preventing their availability for undergoing oxidation to more reactive intermediates. In the Syrian hamsters, 4-hydroxyestradiol is cleared at a much lower rate than 2-hydroxyestradiol, which has been attributed to the inhibition of the O-methylation of 4-hydroxyestradiol by 2-hydroxyestradiol [244]. This was viewed as an implication for 4-hydroxyestradiol to be further metabolized into a carcinogenic metabolite, especially regarding the fact that both catechol estrogens are produced in equal amounts in the kidney, the site of tumor formation. DES is a good substrate of estrogen 2/4 hydroxylase, and can form catechol estrogens. Other pathways involve aliphatic hydroxylation of the ethyl side chain or the epoxide formation at the double bond, leading to the formation of a phenolic ketone. But perhaps the most important pathway in carcinogenesis is the formation of a semiquinone which can give rise to DES-4',4"-quinone (DES Q). This species can then rearrange non-enzymatically to z,z-dienestrol (DIES) or reduced enzymatically to the hydroquinone [245] (Figure 17). The oxidation of DES to DES Q has been shown to occur in vitro [246]. Roy and Liehr reported that DES Q is also formed in vivo in Syrian hamsters and that its formation was inhibited by estrogen metabolism modulators such as vitamin C and a-naphthoflavone, both causing the inhibition of tumor formation as well in a manner that correlates well with the inhibition of DES Q production, suggesting a role for this metabolite [247]. Quinones have the capacity to go through a reversible redox cycle, and this may lead to the formation of free radicals [248], such as the superoxide anion radical (O2 '*) (Figure 18). In addition these quinones by themselves are reactive and may bind to cellular nucleophiles such as DNA and proteins [249]. They can be reduced enzymatically through a one-electron or two-electron reactions. Enzymes that are J*>hydroiy - E-0E5 J*>diiMlhoiy >E • OES E-0E5- *'**- Mmlquinon* E -0 E 5 I - hydraay- E-OES l - h y d r a i y - Z . Z - O E 5 3'-m«lhaiy-Z.Z-0lES \ 0 l-hydreiy-E • if-0E5 I - Kydruy - 3’• im IlMay • E -y -O E S Figure 17. Metabolism of diethylstilbesterol (Adapted from Metzler et al [245]) 56 involved in one-electron transfer reactions mostly belong to the cytochrome P450 family such as NADPH-cytochrome P450 reductase, NADH-cytochrome b5 reductase and NADH-ubiquinone reductase. They catalyze the conversion of quinones to semiquinones that are unstable and can autooxidize reducing molecular oxygen to the . superoxide anion radical, which in turn can undergo several fates. It may be reduced in the presence of NADPH and subsequently interact with iron salts to form the more reactive oxygen species such as hydrogen peroxide or the hydroxyl radical [250], or it may be oxidized enzymatically by superoxide dismutase [251]. Any hydrogen peroxide that is formed can be controlled via two classes of enzymes, catalases and glutathione peroxidases that reduce it to water and oxidizes glutathione (GSH) to its disulfide (GSSG) [252], which is then recycled via glutathione reductase and NADPH. Another enzyme is DT-diaphorase which is a two-electron oxidizing enzyme that utilizes NAD(P)H to catalyze the conversion of quinones to hydroquinones. Hydroquinones are stable, water soluble metabolites that can be excreted as such or as their glucuronide derivatives [253]. Their autooxidation is prevented by the enzyme superoxide dismutase [254]. Thus the diaphorase, superoxide dismutase, the catalases and the glutathione-related enzymes are considered the cellular defense mechanism against quinone toxicity [255]. Any imbalance in the levels of these enzymes may lead to an increase in tumor incidence. Roy and Liehr reported the presence of two reductases that use estrogen quinones as their substrate in the Syrian hamsters kidneys [256], a quinone reductase and a cytochrome P450 reductase. Using inhibitors of each of these enzymes and measuring the levels of superoxide anion radical formation, they deduced that while the cytochrome P450 reduced the quinone by a one electron reduction resulting in the OKI H M N iZ -O IH Figure 18. Redox cycle of quinones (Adapted from Roy et al [248]) production of the superoxide anion radical, the quinone reductase was a two-electron transfer reductase and was thus similar to the DT-diaphorase producing the stable hydroquinone. Interestingly, the activity of the quinone reductase decreased by 80% specifically in the kidneys (but not in the liver) of hamsters treated with estrogens for one month. Some other enzymes were affected. There was a significant increase in glutathione peroxidases, and a decrease in catalases activity, while other free radical detoxifying enzymes such as superoxide dismutase, glutathione reductase and transferase were unaffected [257]. A different study disagreed with these results. They found that all protective enzymes including the glutathione-related enzymes and the superoxide dismutase activities were decreased in kidneys of estrogen-treated animals [258]. This discrepancy may have been due to the fact that the first study looked at animal treated up to 4 months, while the second study looked at animals treated for 6 to 8 months. Furthermore, Li et al showed that superoxide dismutase displayed a transient increase in kidneys of animals treated for 1.5 to 3 months, followed by a decrease below control levels after 4.5 months of treatment [259]. 58 They also reported that catalase activity declined steadily after 1.5 months of DES treatment. In tumor tissues, the activity of all the protective enzymes mentioned are decreased, except for the superoxide dismutase that nearly doubled when compared to control levels. The oxidation of DES to DES Q was also markedly decreased, and all these observations were viewed as an adaptation to the neoplastic environment after chronic exposure to estrogens [260, 261]. Modulators of estrogen metabolism have been shown to prevent tumor formation in Syrian hamsters. Administration of 7,8-benzoflavone plus DES did not give rise to kidney tumors, but caused liver tumors instead, which has been attributed to the modification of the flavones metabolism [262]. Vitamin C has also been shown to inhibit renal adenocarcinoma formation by about 50%, with a concurrent decrease in the major DNA adduct formed upon administration of DES [263]. In addition, it caused an inhibition of further metabolism of DES Q to the reactive intermediates in a dose-dependent manner, thus leading to the speculation that its effect is due to a reduction in the estrogen quinone metabolites and their DNA adducts [264]. Other compounds such as 2(3)-t-butyl-4-hydroxyanisol (BHA) and dicumarol caused a decrease in the renal peroxidative activity of cytochrome P450 that correlated well with a similar reduction in tumor induction [265]. 1.5.3 THE ROLE OF DNA AND PROTEIN ADDUCTS AND CELL DEATH IN CARCINOGENESIS Covalent DNA adducts have been shown to be produced by many chemical carcinogens [266] and may lead to point mutations and activation of oncogenes and 59 chromosomal alterations [267] and finally induction of tumors. Early investigations of DES-DNA adducts in Syrian hamsters were unsuccessful. Very low levels of binding of DES to DNA was later observed but not isolated [268], until Liehr and coworkers used 32P-postlabeling assays to demonstrate the presence of covalent binding to DNA in the kidneys of DES-treated animals, but no such binding in the kidneys of untreated controls [131]. Further studies showed that structurally diverse estrogens form the same DNA adducts, while other steroid hormones did not form such adducts. This observation lead to the conclusion that these adducts are a result of common reactive compounds or free radicals formed through the metabolism of estrogens [269]. A role for estrogen metabolism was further aided by studies showing that tamoxifen, an antiestrogen that competitively binds to the estrogen receptor, inhibited tumor formation, but not DNA adduct formation, thus ruling out a receptor mechanism [270]. The adducts formed peaked after about 5 months of treatment, prior to any visible tumors, and are specific to the kidneys, with the highest concentration in the cortex, the origin of the tumor [271]. In the tumor tissues, these adducts were absent, but instead hypomethylation was observed which is not detected in untreated or estrogen-treated animals [272]. Recently, Liehr and coworkers reported specific DES-DNA adducts in kidneys of DES-treated hamsters [273]. The major of these adducts matched the one generated in vitro by the reaction of DES Q with DNA or 2'-deoxyguanosine-3'-monophosphate. The authors reasoned that the inability to detect these adducts earlier was due to their chemical instability and rapid DNA repair. The levels of 8-hydroxyguanosine was measured and found to increase in kidneys, but not livers, of hamsters treated with DES for only 15 days [274], The level of this modified nucleoside is used to measure the production of free radicals in the system, especially hydroxyl radicals, and their interaction with DNA (Figure 19). 60 Binding of carcinogens to proteins has been previously reported. In vitro studies show covalent binding of DES to various proteins [244, 245], usually, but not necessarily, through a thiol group on a cysteine or a glutathione moiety. Metabolism of estrogens to the quinone is not a prerequisite for this binding, and it has been suggested that a phenoxy radical may be the interacting species [275]. However studies by Li and Li showed that the amount of irreversible binding of estrogens to microsomal proteins in hamsters is related to the amount of catechol estrogens formed [240]. Metzler and coworkers were able to demonstrate that estrogen quinones can bind to the a- and C-subunits of tubulin and that this binding was dependent on the estrogen. The inability of these adducts to polymerize microtubules in vivo, and the earlier findings that estrogens inhibit microtubule assembly without any covalent binding, leads to impaired formation of mitotic spindles, nondisjunction of chromosomes and aneuploidy leading to cell transformation [276] (Figure 20). Little attention has been directed to the role of cell death in tumor growth and regression. Cell death can be classified into two categories: necrosis, resulting from a lack of blood supply [277] and apoptosis or programmed cell death, a process by which the cell can self-destruct [278]. The latter process is controlled by intrinsic factors along with hormones and growth factors and is activated in tissues destined to die during embryonic development or normal cell turnover [279]. Failure of cells to undergo apoptosis may lead to tumor formation [280, 281]. In the Syrian hamsters, ceasation of estrogen treatment caused regression of the tumor accompanied by a decrease in the mitotic activity and an increase in the rate of cell death, while retreatment reversed these effects [282]. This single-cell death was defined as apoptosis and thus a role for the inhibition of apoptosis was suggested along with the 61 stimulation of cell proliferation as possible mechanisms for DES-induced tumorigenesis. ' a. Liver b. Kidney Figure 19. Levels of 8-hyroxyguanosine in kidney and liver of DES-treated hamsters (Adapted from Roy et at [274]) |pUIHOHI M«T«»OtlTli] I I CO»AL«NT NON-COVALfNTI i u t BBACTION WITH TU»UHW 1 [ in h ibitio n Of HICNOTUNUH A«H M »IT| , - - J ------1 [ a m bu plo ipt . MICBOHUCLII I i I C l LI tbawifopmation I Figure 20. Mechanism of cell transformation by estrogen binding to tubulin (Adapted from Epe et at [276]) CHAPTER H OBJECTIVES AND EXPERIMENTAL 2.1 STATEMENT OF OBJECTIVES Breast cancer is one of the leading causes of death among women in the United States. About one third of these cancers are controlled by estrogens and estrogenic compounds, thus termed estrogen-dependent cancer. One potential experimental animal model for studying estrogen dependent cancers is the golden Syrian hamster. The goal of the studies is to examine the exact role(s) of estrogens in the tumorigenic process, using this animal model. This will be attempted by studying protein biosynthesis in the kidneys of the Syrian hamsters, the site of tumor formation, and the effect of estrogen treatment on the stimulation or inhibition of certain proteins, as well as the short term effects of estradiol (6-8 hours exposure) on protein production. Various techniques will be used to uncover the nature and properties of these proteins and specifically identify proteins that are affected by estrogen treatment. Since the metabolism of estrogens may play an important role in the tumorigenic process may involve estrogen metabolism, the metabolic profiles of estradiol obtained from Syrian hamsters treated with estrogens for varying periods of time in three different preparations, a primary kidney cell culture, kidney and liver 62 63 microsomal preparations, and freshly prepared tissue slices from the liver and kidneys will also be examined. Comparing the metabolism of estrogens in these preparations at various stages in the tumorigenic process may lead to the identification of metabolites that are affected by estrogen treatment, or that appear or disappear at certain stages in tumor formation. The kinetic parameters of various enzymes in these metabolic pathways, including estrogen 4-hydroxylase in both the liver, the major site of metabolism in the body and the kidneys, the site of tumor formation will be evaluated. Also studied will be the localization of various metabolites within the components of the cell and the extent of protein binding of these metabolites and the effect that estrogen treatment will exert. Water soluble metabolites arise from conjugation with glucuronide, sulfates and thioethers. The formation of these conjugates in both the liver and kidneys, and the effect of estrogen treatment will be evaluated, followed by the identification of the nature of these conjugates. An examination of the effects that bromoestrogens, which inhibit the enzyme estrogen 2/4-hydroxylase and possibly other cytochrome P450 enzymes, will show on the metabolism of estradiol in both the liver and kidneys of the Syrian hamster may provide insights regarding the nature of the metabolites formed. These studies should unveil the complete picture of estrogen metabolism in this animal model and may lead to better understanding of the tumorigenic process in the Syrian hamster. 64 2.2 EXPERIMENTAL 2.2.1 MATERIALS AND METHODS [2-3H]-Estradiol-176 (18.1-25.3 Ci/mmole) and [6,7-3H]-estradiol-17B (49 Ci/mmole) were purchased from New England Nuclear Corp. (Boston, MA). [4-3H]-Estradiol-1713 (39.7 Ci/mmole) and bromoestrogens were previously synthesized in our laboratory. p 5S]-Methionine (860-1112 Ci/mmole) was obtained from ICN radiochemicals (Irvine, CA). Steroids purchased from Steraloids (Wilton, NH), chemicals for electrophoresis and protein determination from BioRad Laboratory (Rockville, NY), NADP+, glucose-6-phosphate and glucose-6-phosphate dehydrogenase and bovine serum albumin from Sigma Chemical Co. (St. Louis, MO), HPLC grade solvents from Burdick and Jackson (Mcgaw, IL). All other chemicals were obtained from Aldrich Chemical Co (Milwaukee, WI). Culture medium RPMI-1640 and trypsin were obtained from Gibco (Grand Island, NY). Fetal calf serum was heat inactivated for 60 minutes at 60°C before use. Sterile culture flasks and centrifuge tubes were obtained from Corning (Coming, NY), while other glassware were sterilized in a Marketforge sterilmatic. A vertical laminar flow type hood was employed to isolate cells and change media. Cells were cultured in a Forma 3029 C02 incubator, examined on an Olympus IMT-2 inverted research microscope. Tissues were homogenized in SDT Tekmar tissumizer. Amberlite XAD-2 ion exchange resins (20-60 mesh) were purchased from Aldrich Chemical Co. HPLC was conducted on a Beckman System Gold using a reverse phase column (Altex Ultrasphere ODS, 5 micron, 4.6 mm I.D. x 25 cm) using a model 757 absorbance UV detector (Applied Biosystems) and a Beckman 171 radioisotope 65 detector, and Beckman Readyflow III. Radioactive samples were counted on a Beckman LS6800 liquid scintillation counter using Budget Solve from RPI (Mount Prospect, IL). A Beckman J2-21 centrifuge and a Beckman L5-50B ultracentrifuge were used to separate different cellular components. Dialysis was performed in a Pierce Microdialyzer System 500 using Spectrum dialysis tubing. Electrophoresis was run in a Hoefer SE600 vertical slab gel unit, while isoelectric focusing was run in a BioRad 155 tube cell using a Buchler 3-1500 power supply. Gels were dried on a BioRad 224 slab gel dryer, and exposed to X-OMAT AR X-ray film from Kodak (Rochester, NY) and the autoradiograms were scanned on a DU-8 spectrophotometer. Western blotting was performed on a TE-50 Hoefer transfor, the nitrocellulose paper was purchased from Hoefer Scientific (San Fransisco, CA), monoclonal antibodies against mouse TGF-a raised in rabbits from Oncogene Science (Manhasset, NY) and polyclonal antibodies against rabbit bFGF raised in rats from Collaborative Research Inc. (Bedford, MA). Protein concentrations were determined using Bradford assay [283] and a Gilford spectrophotometer. Golden Syrian hamsters, 80-120 gm, were purchased from Harland Industries, Inc. (Cumberland, IN). Castrated males were maintained under constant temperature and humidity, with a 14 hour light, 10 hour dark cycle. DES-pellets obtained from Copley Pharmaceuticals (Boston, MA) contained 20 mg/pellet and were implanted subpannicularly in the shoulder region under ether anaesthesia. DES-pellets were replaced every three months. 66 2.2.2 PREPARATION OF PRIMARY KIDNEY CELL CULTURES The following procedure was performed according to Lin's method [113]. Hamsters were sacrificed while under ether anaesthesia, and the kidneys immediatly removed aseptically. The kidneys were rinsed in phosphate buffered saline (PBS) several times, cleaned, the adrenal glands and fat removed and then minced into very small pieces. The minced particles were trypsinized at 37°C for 45 minutes using 0.25% trypsin in RPMI-1640. The remaining debris were then filtered through a 140 micron mesh and the kidney cell suspension was quenched with an equal volume of RPMI-1640 containing 10% fetal calf serum (FCS) and gentamycin (20 pig/ml). The cells were then suspended in RPMI-1640 containing 10% FCS and gentamycin and the viability was about 80-90% using trypan blue exclusion stain method. The cells were inoculated in 75 mm2 cell culture flasks, at approximately 1-10 x 106 cells/flask and 10 ml 10%FCS in RPMI-1640, and incubated at 37°C in a cell culture incubator (95% air, 5% C02, moisture saturated atmosphere). The media was changed after the second day, after the cells are already attached and started to proliferate. The media was then changed every other day, until cells reach 80-90% confluence. For cell type determination, the presence of two enzymes present in epithelial cells but not fibroblasts were investigated, following the procedure of Gibson and D'Ambrosio [284]. First the cells were tested for their gamma glutamyl transpeptidase activity (GGT). Cells, after reaching 80-90% confluence, were washed in PBS, air dried and incubated at room temperature for 45 minutes with 0.08 gm/L of 5-glutamyl 4'-methoxy B-naphthylamide, 0.26 gm/L glycyl glycine and 0. 4 gm/L fast garnet GBC in 25 mM Tris-HCl buffer, pH 7.3. The cells were then washed twice with distilled water, incubated for two minutes in 0.1 M cupric sulfate, followed by two more washings in distilled water. Positive cells for GGT activity 67 showed a bright red precipitate in their cytoplasm under phase contrast, bright field light microscope, while negative cells appeared light brown. In the second method, alkaline phosphatase activity (AP) was tested, where 80-90% confluent cells were washed in PBS, then air dried and incubated at room temperature for three hours in 100 mM sodium chloride, 5 mM magnesium chloride, 0.17 gm/L of 5-bromo-4-chloro-3-indoyl phosphate and 0.33 gm/L nitroblue tetrazolium in 25 mM Tris-HCl buffer, pH 9.5. The cells were then washed in 10 mM EDTA and viewed under a light microscope. Positive cells showed a blue violet precipitate, while negative cells appeared light gray. 2.2.3 PREPARATION OF KIDNEY AND LIVER MICROSOMES The procedure here follows that of Brueggemeier [285]. Hamsters were sacrificed, the kidneys and liver removed immediately and tissues placed in 0.05 M Tris-HCl buffer, pH 7.4, over ice. All the following steps were carried out at 0-4°C. The tissues were weighed, then cut into small pieces and homogenized in 0.05 M Tris-HCl buffer, pH 7.4. The homogenized solution was then centrifuged at 10,000xg for 20 minutes, and the supernatant layer was then centrifuged at 105,000xg for one hour. The resulting pellets were suspended in Tris-HCl buffer, pH 7.4, then centrifuged at 105,000xg for another hour to obtain the microsomal pellets, which were either used fresh or stored at -70°C. 68 2.2.4 PREPARATION OF KIDNEY AND LIVER SLICES After hamsters were sacrificed, liver and kidneys were removed and placed in PBS buffer. Using a sharp blade, the tissues were cut into 0.5-1 mm thick slices, The sections were cut longitudinally to contain all types of cells, and then used fresh. 2.2.5 PROTEIN BIOSYNTHESIS IN PRIMARY KIDNEY CELL CULTURES Cells used to investigate protein production were then divided into four groups (A,B,C,D, see table X). They were incubated with methionine-free Dulbecco's modified Eagle media (DMEM) for one hour, then spiked with [35S]-methionine and incubated for eight hours. The media and the cells were then separated and frozen at -20°C. Table 2. Pretreatment and treatment of kidney cell cultures with 10 nM estradiol A BC D Pretreated with lOnM estradiol + + -- Treated with lOnM estradiol + + 69 2.2.6 SEPARATION OF PROTEINS USING ONE AND TWO DIMENSIONAL ELECTROPHORESIS Secreted proteins, present in the media from the cell cultures, were dialyzed through a 6000-8000 molecular weight cutoff dialysis tubing, using SDS dialysis solution (2 gm SDS, 40 ml of 60% glycerol, 10 ml of 8-mercaptoethanol, 80 ml triple distilled water in 50 ml 0.25 M Tris-HCl buffer, pH 6.8) for twenty four hours at room temperature and gentle stirring. The dialyzed solution was then boiled with an equal volume of SDS sample diluting buffer (1% SDS, 10% glycerol, 0.05% bromophenol blue and 5% B-mercaptoetanol in 0.0625 M Tris-HCl buffer pH 6.8) for fifteen minutes. Cellular proteins were precipitated from cells or homogenized tissue slices using 30% trichloroacetic acid, and centrifuged at 10,000xg for twenty minutes. The protein pellets were solubilized in SDS sample diluting buffer and boiled for fifteen minutes. Proteins were separated using 10-15% SDS-PAGE electrophoresis according to Lammelli's procedure [286]. Equal amounts of protein from each sample was stacked at 30 mA for one hour, then run at 60 mA for two to two and a half hours, until the dye (bromophenol blue) reaches the gel front. The gel was washed in distilled water, stained in 0.05 % commassie blue for one hour, then destained in 10% acetic acid overnight. The gels were washed three times in distilled water, then either directly dried at 80°C for three hours or if autoradiography was required, soaked in 1 M sodium salicylate as a fluor for thirty minutes, then dried. The dried gels were exposed to an X-ray film for 5-7 days at -70°C. The film was then developed for 5-7 minutes and fixed for three minutes in the dark, washed in running water for ten minutes, and finally dried in a dust-free area. The autoradiograms were scanned using a DU-8 spectrophotometer at 486 nm. For further identification of the proteins, two-dimensional electrophoresis was employed following O'Farrell's procedure [287]. The first dimension consisted of 70 running an isoelectric focusing tube gel (13 x 0.5 cm). The gel mixture was made up of 5.5 gm urea, 1.33 ml of 30% acrylamide solution, 2 ml nonidet NP-40, 1.97 ml distilled water, 0.4 ml of ampholines, pH 5-7 and 0.1 ml of ampholines, pH 3-10. The solution was swirled to dissolve the urea, 10 /x 110% ammonium sulfate was added; The solution was degassed for one minute, 7^tl TEMED added and then poured into clean tubes, avoiding trapping air bubbles. Each gel was overlayed with 8 M urea and allowed to stand for two hours, the overlaying solution removed and 20 n 1 lysis buffer (9.5 M urea, 2% nonidet NP-40, 1.6% ampholines, pH 5-7, 0.4% ampholines, pH 3-10, 5% B-mercaptoethanol) added and overlayed with water. The gel was allowed to stand for another two hours, fresh lysis buffer added, and overlayed with previously degassed 0.02 M sodium hydroxide. The tube gels were placed in a BioRad tube cell, with 0.01 M phosphoric acid in the lower reservoir and 0.02 M sodium hydroxide in the upper reservoir. The gels were prerun at 200 V for fifteen minutes, followed by 300 V for thirty minutes and finally 400 V for another thirty minutes. The lysis buffer was then replaced by 10 /xl sample overlaying solution containing 0.8% ampholines, pH 5-7, 0.2% ampholines, pH 3-10 in 9 M urea solution, overlayed with 0.02 M sodium hydroxide and run at 500 V for eighteen hours, followed by 1000 V for one hour. The gels were immediately extruded from the glass tubes, equilibrated with 5 ml SDS sample buffer (10 gm glycerol, 2.3 gm SDS, 5 gm B-mercaptoethanol in 100 ml of 0.0625 M Tris-HCl buffer, pH 6.8) for two hours, changing the buffer twice, and stored indefinitely at -70°C. To run the second dimension, an SDS-PAGE gel was employed, where the tube gel was embedded in a 1 % agarose gel that was poured over the stacking gel, and the slab gel then run as described above. The gel was then stained, destained and exposed to an X-ray film and the autoradiogram developed as described above. 71 2.2.7 WESTERN BLOTTING The procedure for protein blotting followed that of Towbin [288] with some modifications. One and two-dimensional SDS-PAGE gels were run as described above, soaked in blotting media (0.025 M Tris base, 0.193 M glycine in 20% methanol in water) for five minutes, placed over a nitrocellulose paper (NC paper), avoiding trapping any air bubbles then sandwiched between two wet filter papers, covered with scotch-brite pads and secured into a plastic unit, without allowing the assembly to dry. The assembly was lowered into a blotting media in a Transfer system and run at 60 V for twenty hours. The NC paper was left to dry for one hour, then soaked in 5% low-fat milk as a blocking agent for two hours at room temperature with gentle agitation, followed by extensive washing in PBS containing 3% Tween 20, incubation with the primary antibody for two hours at 4°C, and then extensive washing in PBS/Tween. The NC paper was soaked in 8.775 gm sodium chloride and 6.055 gm Tris base in water, pH 7.5 (Wash solution A), for ten minutes with gentle agitation, then incubated with the appropriate dilution of the secondary antibody linked to alkaline phosphatase in a phosphate-free blocking solution (5% non-fat milk in wash solution A) for one hour at room temperature, followed with three washings in wash solution A, each for ten minutes. Finally the paper was shaken with BioRad alkaline phosphatase developing solution, until the bands appeared, and developed to the desired intensity, then washed in 0.5 M EDTA in PBS, pH 8 and stored. 72 2.2.8 IMMUNOPRECIPITATION OF TGF-a AND BFGF Samples were boiled with 2.5 mM EDTA in 1%SDS in water for 8 to 10 minutes, and treated with an equal amount of 0.2% SDS, 0.5 mM EDTA, 0.8% mM dithiotheriotol, 0.8% Triton X-100 and 0.3 M sodium chloride in 40 mM Tris-HCl buffer pH 7.5 (Solution B), and incubated with 4 mg protein A-sepharose CL-4B for five hours at 4°C, to reduce non-specific binding. After centrifugation at 700xg for two minutes, the supernatant layer was incubated with 5 p\ of the specific antibody (100 /xg/ml of monoclonal antibodies against TGF-a and 1 mg/ml for polyclonal antibodies against bFGF) for four hours at 4°C. The antigen-antibody complex was precipitated using 4 mg protein A-sepharose and incubating overnight at 4°C. The pellets collected by centrifugation at 700xg, were washed four times in solution B, and the pellets then dissolved in SDS sample buffer and boiled for ten minutes, and separated on an SDS-PAGE gel electrophoresis and visualized by autoradiography as mentioned above. 2.2.9 ESTROGEN METABOLISM IN PRIMARY KIDNEY CELL CULTURES Cells used for estrogen metabolism studies were incubated for 24 hours with 100 nM, 2.5 [ id [bJ^HJ-estradiol. The media and the cells were separated and stored at -20°C. 73 2.2.10 SEPARATION OF VARIOUS METABOLITES The supernatant layers from different incubations were thawed, 10 gm of ammonium sulfate added to precipitate the proteins, and then centrifuged at 4°C for twenty minutes. The supernatant solution was separated from the protein pellets, the latter washed three times with equal volumes of ethyl acetate, and the washings added to the water layer which was further extracted three more times with equal volumes of ethyl acetate. The organic layers were combined, dried using sodium sulfate, then evaporated to dryness under a stream of nitrogen. The precipitate was dissolved in 200 /il of 75% methanol in water for further studies. 2.2.11 HPLC ANALYSIS OF ESTROGEN METABOLITES Separation of the organic-extractable metabolites was performed by reverse phase HPLC according to the method of Brueggemeier [289], using a Beckman system gold HPLC. Aliquots (20-30 /zl) were separated on a Beckman Ultrasphere ODS column (5 /zm, 4.6 mm x 25 cm) using a gradient of 30% to 50% methanol in water over 35 minutes (to separate the more polar metabolites), followed by another gradient of 50% to 60% methanol in water over 35 minutes (to separate the less polar metabolites), at a flow rate of 1 ml/min. The eluent was monitored with a UV detector at 254 nm, then combined with 3 ml of Ready flow III scintillation cocktail and the radioactive metabolites detected by a Beckman model 171 radioisotope detector. Authentic steroids were coinjected with the samples and detected by a UV detector. 74 2.2.12 CHARACTERIZATION OF WATER-SOLUBLE METABOLITES To investigate the nature of the water-soluble metabolites, the presence of glucuronides, sulfates and thioethers was examined. All procedures were adapted from Ball et al [290], with some modifications. For the presence of glucuronides, 0.2 mg 8 -glucuronidase enzyme [Sigma (EC 3.2.1.31) type L-III] in 100 mM phosphate buffer, pH 3.8 (to inhibit sulfatase activity), was added to 1 ml of the sample. For the presence of sulfate conjugates, 1 mg aryl sulfatase [Sigma (EC 3.1.6.1) type H-l], 20 mM D-glucaro-1,4-lactone (to inhibit glucuronidase activity) in 100 mM acetate buffer, pH 5 was added to 1 ml of the sample. The reaction mixture in both cases were then incubated for 24 hours at room temperature with gentle agitation, the reaction stopped by the addition of 1 ml ethyl acetate, and the water layer extracted three times with equal volumes of ethyl acetate. The organic layers were combined, dried using sodium sulfate and aliquots (0.5 ml) of both the organic and water layers were combined with 4.5 ml of Budget solve scintillation cocktail and counted on an LS6800 liquid scintillation counter. To determine the presence of thioethers, 1 ml of the sample was combined with 25 mg Raney nickel in 10% acetic acid in water, incubated for twenty four hours at 4°C under nitrogen with gentle agitation (to keep the Raney nickel suspended in the solution). The solution was then extracted three times with equal volumes of ethyl acetate and an aliquot (0.5 ml) of both the organic and water layers were counted as described above. Finally, the organic-extractable fraction from each experiment was evaporated to dryness, redissolved in 75% methanol in water and injected on reverse phase HPLC, as previously described to identify the conjugated metabolites. 75 2.2.13 SUBCELLULAR DISTRIBUTION OF METABOLITES IN TISSUE SLICES All steps were carried out at 0-4°C. Slices were homogenized in 10 ml 0.1 M Tris-HCl buffer, pH 7.4, and the homogenate was then centrifuged at 10,000xg for twenty minutes, and the supematent layer was then centrifuged at 105,000xg for one hour. The precipitate from the 10,000xg fraction and the microsomal pellets from the 105,000xg centrifugation were homogenized in 10 ml 0.1 M Tris-HCl buffer, pH 7.4. Aliquots (1 ml) of each fraction (10,000xg fraction, cytoplasmic fraction and the microsomal fraction) were counted using Budget solve scintillation cocktail and an LS6800 counter. Each solution was then subjected to protein precipitation and ethyl acetate extraction, as described above. Aliquots of each fraction (protein pellets, aqueous layer and organic layer) were counted for radioactivity, and then the organic layers were evaporated to dryness and redissolved in 75 % methanol in water and injected on reverse phase HPLC to detect the metabolites formed. 2.2.14 3 H20 ASSAY OF ESTROGEN 4-HYDROXYLASE To determine the kinetic parameters of the enzyme estrogen 4-hydroxylase in liver and kidney microsomal preparation, the studies were carried out under initial velocity conditions, and measured by the 3 H20 assay developed by Brueggemeier [291]. Various concentrations of estradiol (0.5-10 jiM, approximately 250,000 dpm of ^HJ-estradiol/flask) in 100 (A propylene glycol were incubated at 37 °C for fifteen minutes with 0.8 mM NADP+, 2 mM glucose- 6 -phosphate, 2.5 units of glucose- 6 -phosphate dehydrogenase (as cofactors for the enzymatic reaction) and 2 ml of hamster liver or kidney microsomes in 0.1 M Tris-HCl buffer, pH 7.4. The assay 76 was performed in triplicate, with blank samples incubated with boiled microsomes and/or in the absence of cofactors. The incubations were stopped by the addition of 2 ml 2 N HC1 and the samples frozen overnight, then later thawed and the released 3 H20 was separated from the steroids using a non-ionic XAD-2 column (1.2 x 6 cm) with water as the eluent to give a final volume of approximately 20 ml. One ml of each sample was mixed with 4 ml Budget solve scintillation cocktail and counted in an LS6800 liquid scintillation counter. 2.2.15 BROMOESTROGENS INHIBITION OF ESTROGEN METABOLITES Freshly prepared kidney slices were incubated with 1 /xM estradiol (2.5 fiCi, [6,7-3 H]-estradiol) in 5 ml RPMI-1640 media, and different concentrations (0, 2, 5 /xM) of 2-bromoestradiol, 4-bromoestradiol or 2,4-dibromoestradiol at 37°C for 0, 3, 6 hours. The incubations were stopped by the addition of 1 ml ethyl acetate in addition to 1 ml 5 % ascorbic acid and authentic catechol estrogens to protect the oxidative metabolites. Protein precipitation using ammonium sulfate and ethyl acetate extraction of the supernatant layer and the homogenized slices followed, and aliquots from each fraction (protein pellets, aqueous layer and organic-extractable metabolites) were counted on LS6800 counter. Organic extractable fractions were evaporated to dryness and redissolved in 75 % methanol in water and the formed metabolites separated and detected on reverse phase HPLC. The water-soluble metabolites were also studied as previously described. 77 2.2.16 APPROACHES TO STRUCTURE ELUCIDATION OF ESTROGEN METABOLITES Freshly prepared liver slices were incubated with different concentrations of estradiol (1, 5, 10 /xM) in 5 ml RPMI-1640 at 37°C for 6 hours. The reaction was stopped by the removal of the slices from the incubation mixture and the addition of 1 ml ethyl acetate, 1 ml 5 % ascorbic acid and authentic catechol estrogens. The proteins were precipitated using ammonium sulfate, and the resulting protein pellets were washed three times with equal volumes of ethyl acetate. The washings were added to the aqueous layer which was extracted three more times with equal volumes of ethyl acetate and the organic layers combined, dried with sodium sulfate, evaporated to dryness and redissolved in 75 % methanol in water. To monitor the experiment, one sample was run under the same exact conditions except for the addition of 2.5 pCi [6 ,7-3 H]-estradiol prior to the incubation. The metabolites were then separated on reverse phase HPLC to four major fractions. Each fraction was dried down by spinning at 1725 rpm under vacuum in a Savant speedvac concentrator, and redissolved in 100 /xl 75% methanol in water. The mass spectroscopy of metabolites was determined by both electron ionization and desorption ionization. CHAPTER HI RESULTS AND DISCUSSIONS 3.1 CELL TYPE IDENTIFICATION One of the preparations used in examining the role(s) of estrogens in the induction and the development of estrogen-dependent cancer is the primary hamster kidney cell culture. Microscopical examination indicated a cell population made up of epithelial cells and fibroblasts. Renal adenocarcinomas that arise after estrogen treatment in Syrian hamsters are postulated to be of an epithelial cell origin, and the composition of the tumor tissue is mostly epithelial. Moreover, breast carcinoma in humans are epithelial cell tumors. For the cell culture to bear a close resemblence to the environment in vivo, it has to be composed of mainly epithelial cells. We used two histochemical markers to examine the cell population, gamma-glutamyl transpeptidase (GGT) activity and alkaline phosphatase (AP) activity. These two enzymes are present in epithelial cells but not fibroblasts. We used three different cell culture preparations and counted 1 0 0 cells/preparation in three different sections of the glass slide. The GGT activity was positive in 71, 78, 73 cells out of 100, with an average of 74 _+ 5.1 % (Plate I), while AP activity was positive in 79, 73 and 77 cells out of 100, with an average of 76 ± 4.4% (Plate II). Thus the kidney cell 78 79 Plate I. A microscopical section from a primary kidney cell culture staining positive for the presence of GGT (x400). Positive cells showed a bright red precipitate in their cytoplasm under the light microscope 80 Plate II. A microscopical section from primary kidney cell culture staining positive for the presence of AP (xlOO). Positive cells showed a blue violet color on their cytoplasm under the light microscope 81 culture is composed of a mixed cell population that contains about 75 +. 3.64% epithelial cells. 3.2 PROTEIN BIOSYNTHESIS IN KIDNEYS OF SYRIAN HAMSTERS 3.2.1 SEPARATION AND CHARACTERIZATION OF PROTEINS Protein biosynthesis was examined by p 5 S]-methionine incorporation in primary hamster kidney cell cultures from untreated and DES-treated Syrian hamsters. The total protein secreted from cell cultures were determined by Bradford assay. The total protein production increased only slightly with DES treatment, where untreated hamsters produced about 6 8 /xg/ml, which increased by about 3 fold in hamsters treated for eleven months with DES. In tumor-bearing hamsters, a sharp increase in protein production was observed, reaching about 1.64 mg/ml (Table 3). In addition, short term exposure to estradiol (groups A-D, page 67) stimulated protein production by about 2-3 folds (Table 4). This stimulation was acheived using 10 nM estradiol, and higher concentrations did not affect total protein production. This increase may reflect the hormonal effects of estrogens on protein synthesis. Secreted proteins were then separated on 10% SDS-PAGE electrophoresis, visualized by fluorography, then the resulting autoradiograms scanned on a DU - 8 spectrophotometer. Pictures of autoradiograms from secreted proteins from cell cultures of 0, 3, 6 , 9, and 11 month-treated animals are shown on Plates III - VII, and scans from these autoradiograms are shown in Figures 21-2 5 . The major bands that appeared in these autoradiograms had molecular weights of 75-77 kDa (band a), 64-66 kDa (band b), 56-57 kDa (band c), 45-46 kDa (band d), 34-36 kDa (band e), 29-30 kDa (band./) and 20-22 kDa (band g). The effects of short term estradiol exposure (groups A-D, see page 6 8 ) appears to be only significant in cell cultures from untreated animals, while cultures pretreated and treated with estradiol (group A) and those pretreated only (group B) or treated only (group C) showed no differences. The effect of DES treatment was obvious on two protein bands e and g, both showing an increase in amount with an increase in DES treatment (Table 5-9), compared to protein band d which does not change with DES treatment (Figure 26). Protein band e showed an increase from 6.25% of total protein produced in untreated hamsters, to nearly 10% in six months DES-treated hamsters, to over 14% in eleven months DES- treated hamsters. Protein band g represented 3.26% of total protein in untreated hamsters, 7.64% in six months DES-treated hamsters and reached 11.77% in eleven months DES-treated hamsters. Other protein bands did not show consistent stimulation or inhibition by DES treatment. Further characterization of these proteins was attempted by using two-dimensional electrophoresis. We were unable to consistently produce reliable autoradiograms, probably due to the low amounts of proteins produced by cell cultures. Plate VIII shows a two-dimensional autoradiogram of secreted proteins from a nine-month treated animal. The first dimension was run on a 7% urea- acrylamide gel with ampholytes ranging from pH 3-10, to separate proteins based on their isoelectric point. The second dimension was run on a 15 % gel to seperate the lower molecular weight proteins (less than 20 kDa). Two protein spots appeared on this autoradiogram that were absent on all those from untreated hamsters, one with a molecular weight of 20-22 kDa, and the other of 16-18 kDa. The lower molecular 83 weight spot appeared towards the basic side of the gel, indicating an isoelectric point of 8.5-9. 3.2.2 IDENTIFICATION OF SPECIFIC PROTEINS The presence of two growth factors, TGF-a and bFGF was investigated. TGF-a has been identified in many breast tumor cell lines and in biopsies and urine samples of breast cancer patients [202] and has been reported to exist as a 20-25 kDa, which coincides with protein band g. Properties of bFGF include a molecular weight of 16-18 kDa and an isoelectric point of around 9, both seen in one of the two spots appearing in Plate VIII. In addition, bFGF is an angiogenic factor and studies show the increase in the area of vascularization in kidney of Syrian hamsters after DES treatment [128]. To investigate the presence of TGF-a and bFGF, a monoclonal antibody against mouse TGF-a and a polyclonal antibody against rabbit bFGF were utilized. Both growth factors have an amino acid sequence that is highly conserved among different species, as illustrated by the amino acid sequence of bovine and human bFGF that has 155 amino acids and differed by only two [171], thus the antibodies used might crossreact with the hamster proteins. Results from Western blots were inconclusive, probably due to the low sensitivity of the technique coupled with the small amounts of proteins produced from cultures of hamsters. Immunoprecipitation of the growth factors, followed by separation on SDS-PAGE electrophoresis, showed that cell cultures from five month-treated animals produced a TGF-a-like protein (Plate IX), that was unaffected by short term exposure to estradiol, and a bFGF-like protein (Plate X) that was stimulated by eight hours exposure of the cell culture to 10 nM estradiol. Cell cultures from untreated animals 84 did not produce these proteins. Thus DES-treatment of Syrian hamsters caused an overall increase in production of total proteins, and in the stimulation of at least three protein bands, that possessed molecular weights of 34-36 kDa, 20-22 kDa and 16-18 kDa. The last two protein bands contain a TGF-a-like protein and a bFGF-like protein respectively. The induction of these proteins and growth factors may play an important role in the tumorigenic process in Syrian hamsters, including cell proliferation and vascularization of the tumor tissue. 85 Table 3. Total protein secreted from primary kidney cell culture of untreated and DES-treated hamsters. Cells were pretreated and treated with 10 nM estradiol and the secreted protein concentrations were determined by Bradford assay and the results expressed in /xg/ml DES treatment Animal 1 Animal 2 Animal 3 Mean ±_ S.E. in months 0 90 50 65 68.33 ±28.58 3 80 110 105 98.33 ±22.73 6 140 100 170 136.66 ± 49.67 9 175 190 210 191.66 ± 16.66 11 220 180 260 220.00 ± 56.56 Tumor-bearing 1610 1680 1645.00 ± 49.50 86 Table 4. Total protein secreted from primary kidney cell culture of untreated hamsters, subjected to different treatments with estradiol. The cultures were subjected to pretreatment and treatment (A), pretreatment only (B), treatment only (C) or no treatment (D) with 10 nM estradiol as described in experimental. Protein concentrations were determined by Bradford assay and results expressed in /xg/ml Experiment AB c D 1 90 90 60 30 2 50 70 80 12 3 65 50 60 35 4 80 80 90 45 Mean 71.25 72.50 72.50 30.50 S.E. 15.15 14.79 12.99 11.97 87 Plate III. Autoradiogram of secreted proteins from kidney cell cultures of untreated hamsters. The lanes represent pretreatred and treated (A), pretreated only (B), treated only (C) and untreated (D) with 10 nM estradiol as described in experimental. Each lane recieved 40,000 dpm of radiolabeled proteins. ' ! A BCD Figure 21. Scan of autoradiogram from kidney cell culture of untreated hamster on DU- 8 spectrophotometer. Each scan represents a treatment group A-D. Major bands appear as following: Band a (70-73 mm), band b (88-92 mm), band c (115-119 mm), band d (126-129 mm), band e (140-144 mm), b a n d /(148-152 mm) and band g (168-171 mm) 88 89 GROUP A Figure 21 90 Figure 21 (cont.) GROUPB © Q O ■S N* OJ ID ■S CD '3 •S O •S iS •S iS 91 Figure 21 (cont.) GROUPC ■3 13 <3 <3 j J >3 >3 rf ■■t iT . in T ro in ■ in -3 >3 <3 •3 '3 <3 ■ © 8.0000 i.8800 2.0000 iue2 (cont.) 21 Figure GROUPD re <30 oo rt £ rfo I i 92 93 Table 5. Major protein bands from autoradiograms of secreted proteins from kidney cell culture of untreated hamsters. The cultures were subjected to pretreatment and treatment (A), pretreatment only (B), treatment only (C) or no treatment (D) with 10 nM estradiol as described in experimental. Results expressed in percent of total protein and is the mean +. S.E. of three experiments Band A B c D a 3.22 ± 1.23 2.61 ± 1.31 2.38 ± 1.05 0.95 ± 0.42 b 5.61 ±2.14 3.22 ± 1.42 3.21 ±0.87 3.20 ±2.21 c 6.14± 1.54 5.93 ± 2.12 4.27 ± 1.22 1.08 ± 0 .3 2 d 21.43 ±5.47 19.09 ± 2.33 21.09 ±3.19 18.91 ±2.98 e 6.25 ±0.52 3.16 ± 1.12 4.07 ± 1.62 N/D f 7.75 ±2.52 8.05 ± 3.21 12.48 ±2.17 1.44 ±0.68 g 3.26 ± 1.47 2.88 ± 1.21 3.19 ± 1.22 1.18 ± 0.32 94 Plate IV. Autoradiogram of secreted proteins from kidney cell cultures of three months DES-treated hamsters. The lanes represent cultures that were pretreatred and treated (A), pretreated only (B), treated only (C) and untreated (D) with 10 nM estradiol as described in experimental. Each lane recieved 40,000 dpm of radiolabeled proteins. Figure 22. Scan of autoradiogram from kidney cell culture of three months DES-treated hamster on DU - 8 spectrophotometer. Each scan represents a treatment group A-D. Major bands appear as following: band a (73-75 mm), band b (8 6 - 8 8 mm), band c (117-119 mm), band d (135-138 mm), band e (148-152 mm), band/(156-159 mm) and band# (162-165 mm) 95 96 GROUP A r * © >3 © OJ © © © © I I © © © © © Figure 22 3.0080 i.3300 2.0000 Figure 22 (cont.) 22 Figure GROUPB m m 3 1 s- Is PO 97 98 ' 0 Figure 22 (cont.) rf • 0 © © GROUP C © •0 © in© p.] © © © © © © © © ■ © 99 in © 0000’S 0000’T 0000 100 Table 6 . Major protein bands from autoradiograms of secreted proteins from kidney cell culture of three months DES-treated hamsters. The cultures were subjected to pretreatment and treatment (A), pretreatment only (B), treatment only (C) or no treatment (D) with 10 nM estradiol as described in experimental. Results expressed in percent of total protein and is the mean +. S.E. of three experiments Band A B C D a 3.75 ± 1.12 3.45 ± 1.18 1.92 ±1.12 2.81 ± 1.20 b 17.11 ±3.24 9.88 ± 1.85 15.88 ±3.15 17.39 ±5.28 c 7.80+1.72 8.32 ±2.14 8.40 ±3.01 7.13 ±1.60 d 22.99 ± 3.47 22.56 ±5.12 17.88 ± 5.36 16.07 ± 3.65 e 8.07 ± 2.54 6.61 ± 1.58 8.64 ±2.15 9.36 ±3.21 f 9.32 ±2.14 8.21 ± 1.25 7.45 ± 2.04 7.23 ± 1.19 g 5.21 ± 1.02 5.32 ± 1.75 4.21 ±0.82 3.89 ± 1.27 101 Plate V. Autoradiogram of secreted proteins from kidney cell cultures of six months DES-treated hamsters. The lanes represent cultures that were pretreatred and treated (A), pretreated only (B), treated only (C) and untreated (D) with 10 nM estradiol as described in experimental. Each lane recieved 40,000 dpm of radiolabeled proteins. A B C D Figure 23. Scan of autoradiogram from kidney cell culture of six months DES-treated hamster on DU - 8 spectrophotometer. Scans represent treatment groups A-D. Major bands appear as following: band a (70-74 mm), band b (84-88 mm), band c (122-126 mm), band d (135-138 mm), band e (148-151 mm), band/(162-164 mm) and band g (176-180 mm) 102 GROUP A Figure 23 0 . 0 0 0 0 i . 0 0 0 0 2 . 0 0 0 0 J -J n i © iue2 cn. r jn ro (cont.) 23 Figure © r\i r-._ ro . •0 . • -) © i^i GROUP B GROUP / D - - -3 ij-. -rH ID i\| ' © © © 'JD ro IT' © © © 0 1 105 H GROUP C Figure 23 (cont.) 0000’S 0000’T 0000 « 106 Figure 23 (cont.) GROUP D oz I El ro ■H 107 Table 7. Major protein bands from autoradiograms of secreted proteins from kidney cell culture of six months DES-treated hamsters. The cultures were subjected to pretreatment and treatment (A), pretreatment only (B), treatment only (C) or no treatment (D) with 10 nM estradiol as described in experimental. Results expressed in percent of total protein and is the mean +. S.E. of three experiments Band AB c D a 6.74 ± 2.56 2.33 ± 1.52 5 .1 2 ± 1.29 7.87 ± 2 .4 1 b 6.70 ± 2.04 7.15 ± 1.25 5.01 ± 0 .7 0 5.28 ± 1 .4 3 c 11.17 + 3.21 11.31 ± 2 .4 8 12.68 ± 4 .0 9 15.56 ± 3.82 d 20.88 ± 3.42 19.07 ± 2.84 16.21 ± 4 .5 7 17.74 ± 3 .1 0 e 9.72 ± 1.58 8.65 ± 1.29 10.24 ± 3 .1 7 9.65 ± 1.26 f 5.34 ± 1 .8 5 5.83 ± 1.62 3.85 ± 1.42 5.59 ± 1.52 g 7.64 ±1.77 7.69 ±2.54 6.75 ± 1.69 7.59 ± 2 .9 1 108 Plate VI. Autoradiogram of secreted proteins from kidney cell cultures of nine months DES-treated hamsters. The lanes represent cultures that were pretreatred and treated (A), pretreated only (B), treated only (C) and untreated (D) with 10 nM estradiol as described in experimental. Each lane recieved 40,000 dpm of radiolabeled proteins. A B C O 1 Figure 24. Scan of autoradiogram from kidney cell culture of nine months DES-treated hamster on DU - 8 spectrophotometer. Scans represent treatment groups A-D. Major bands appear as following: band a (72-76 mm), band b (81-83 mm), band c (124-128 mm), band d (130-134 mm), band e (145-150 mm), b a n d /(155-160 mm) and band g (168-172 mm) 109 110 rTi GROUP A © © © © i-4 w ■ in oo - t © © © ■ ' 0 Figure 24 I l l Figure 24 (cont.) © © GROUP B © © r- -’f Ifi AJ m a j >.£| rt ■’3’ iT . ■ r-- r-- r-o AJ CO ' t - t A uo■ i > /I & I \ AJ r-- AJ ■o © •H — ■ © • 0 -S: © © ro © • J © © •»—4 r.O t •T—• ■ # \ i A \ i -1 V- l \ u© • I.T1 *£i >0 V r - / \ 1T1 j1 \ A \l\ '0 \ v / l / A a / S © © © © ■ \ © 1 © © © © © © % ■ ■ © ■ * © © © 1A 'i> 0 . 0 0 0 0 i . 0 0 0 0 2 . 0 0 0 0 — 1 ■ Figure 24 (cont.) 24 Figure / GROUP C GROUP CT-ro CO Figure 24 (cont.) r- ro co £3 •vD © © '. j 3 © GROUP D © ■ •M -rH ro •jj ©■ © ro in ID IT, r- 0-, © © © ro © '"0 ©5> 'D ■ ■:o © © © ©■ © © ©■ t-H © '•£» 0 1 114 Table 8 . Major protein bands from autoradiograms of secreted proteins from kidney cell culture of nine months DES-treated hamsters. Cultures were subjected to pretreatment and treatment (A), pretreatment only (B), treatment only (C) or no treatment (D) with 10 nM estradiol as described in experimental. Results expressed in percent of total protein and is the mean +. S.E. of three experiments Band A B C D a 4.56 ± 1.02 2.63 ±0.51 2.36 ±0.83 2.68 ±1.20 b 8.06 ± 1.28 8.11 ± 2 .3 1 8.40 ± 1 .5 6 7.73 ± 2 .4 1 c 8.66 ± 1.89 9.81 ± 2 .1 6 7.87 ± 1 .4 9 9.96 ± 2.64 d 27.34 ±5.24 27.01 ± 3.22 26.86 ±3.50 26.34 ±5.81 e 1 1 .1 2 ± 1.46 11.27 ± 3.11 11.55 ± 2 .4 5 10.07 ± 2.74 f 7.65 ±2.18 7.57 ± 1.67 6.42 ± 0.43 5.86 ± 1 .0 1 g 10.65 ± 2 .4 1 9.22 ± 2.02 10.25 ± 1 .2 1 9.80 ± 1 .8 4 115 Plate VII. Autoradiogram of secreted proteins from kidney cell cultures of eleven months DES-treated hamsters. The lanes represent cultures that were pretreatred and treated (A), pretreated only (B), treated only (C) and untreated (D) with 10 nM estradiol as described in experimental. Each lane recieved 40,000 dpm of radiolabeled proteins. Figure 25. Scan of autoradiogram from kidney cell culture of eleven months DES-treated hamster on DU - 8 spectrophotometer. Scans represent treatment groups A-D. Major bands appear as following: band a (81-84 mm), band b (91-93 mm), band c (132-135 mm), band d (141-143 mm), band e (155-158 mm), b a n d /(165-168 mm) and band g (180-184 mm) 116 0.3000 i.0000 2.0000 in © © A © © © © Figure 25 Figure GROUP A GROUP CO Ti t ' er-. © in © © . in in UJ oj ■H © <:<■} © © T. IT © 117 118 Figure 25 (cont.) C'J TO © r-- © © a GROUP B C'i '•£< IT, ro I.U IO co f? iX , •Im r - - < 3 ■ r-- i n a •H • 3 •s-l ■ - t t **4 t - i • 0 TT" A CO j \ fO 7 ** • |T*I TO ■ • ro i 1 .0 IO 0.0000 1.0000 2.0000 iue2 (cont.) 25 Figure GROUP C GROUP r-- 00 ID *j c?j rp Q 119 |J. ■ 120 r-- in H fij i-i GROUP D i-i Figure 25 (cont.) {v! 0900 * 3 0000' T 0000'0 121 Table 9. Major protein bands from autoradiograms of secreted proteins from kidney cell culture of eleven months DES-treated hamsters. Cultures were subjected to pretreatment and treatment (A), pretreatment only (B), treatment only (C) or no treatment (D) with 10 nM estradiol as described in experimental. Results expressed in percent of total protein and is the mean +. S.E. of three experiments Band A B c D a 4.33 ± 1.45 6.01 ± 2 .1 5 6.42 ± 1.34 3.66 ± 1 .1 8 b 4.67 ± 1.33 5.38 ± 1.85 6.11 ± 1 .8 8 3.39 ± 1 .0 5 c 7.91 ± 1.24 11.37 ± 2 .8 7 11.20 ± 3 .3 3 9.17 ± 1.14 d 22.78 ±3.16 24.18 ±3.11 23.96 ± 2 .4 8 21.81 ± 4 .0 8 e 14.10 ± 2 .3 8 12.30 ± 1.28 15.74 ± 3 .6 2 14.03 ± 3.55 f 10.08 ± 3 .2 1 13.03 ± 1.52 13.49 ± 3 .7 6 11.41 ± 2 .8 2 g 11.77 ± 1.72 9.31 ± 1.49 10.34 ± 1.12 10.79 ± 3.37 Figure 26. Biosynthesis of protein band e (34 kDa) and protein band g (20 kDa) in kidney cell cultures of untreated and DES-treated hamsters. Results expressed as the mean of the percent of total protein from three experiments. 122 % % of Total Protein 5 - 25 30 20 5 - 15 10 - - o. t o ertd Proteins Secreted of Wt. Mol. 20 kDa 34 kDa 46 kDa 46 kDa 34 kDa 20 I I Untreated hamster Untreated I I Figure 26 9 month treated hamster hamster treated month 9 hamster treated month 6 hamster treated month 3 11 month treated hamster treated 11 month 123 124 Plate VIII. Autoradiogram from two-dimensional electrophoresis of secreted proteins from kidney cell culture of nine month DES-treated hamster. A and B denote the acidic and basic side respectively. V A B 125 Plate IX. Autoradiogram of immunoprecipitated proteins from kidney cell cultures of untreated and DES-treated hamsters using a monoclonal antibody against TGF-a. Lanes 1 and 2 are samples from cultures of an untreated hamster, while lane 3 and 4 are from cultures of a five months DES-treated hamster. Samples ran in lanes 1 and 3 were not exposed to estradiol, while those in lane 2 and 4 were exposed to 10 nM estradiol for 8 hours. - /• r - p - r « 2 ’3 ' i'. • if'1 - ' • > V 126 Plate X. Autoradiogram of immunoprecipitated proteins from kidney cell cultures of untreated and DES-treated hamsters using a polyclonal antibody against bFGF. Lanes 1 and 2 are samples from cultures of an untreated hamster, while lane 3 and 4 are from cultures of a five months DES-treated hamster. Samples ran in lanes 1 and 3 were not exposed to estradiol, while those in lane 2 and 4 were exposed to 10 nM estradiol for 8 hours. 1 2 3 4 127 3.3 ESTROGEN METABOLISM IN SYRIAN HAMSTERS To study estrogen metabolism in Syrian hamsters, three preparations, the primary kidney cell culture, microsomal preparations from liver and kidneys and freshly prepared liver and kidney tissue slices were utilized. 3.3.1 ESTROGEN METABOLISM IN PRIMARY KIDNEY CELL CULTURES The metabolic profiles in the primary kidney cell cultures of untreated hamsters were compared to those of hamsters treated with DES for different periods of time, and are shown in Figure 27 (full view) and Figure 28 (expanded view). The metabolites were separated by reverse phase HPLC, using a gradient of 30% to 75% methanol in water over 50 minutes, and detected by a Beckman model 171 radioisotope detector. In primary kidney cell cultures, the isolation of catechol estrogens (peaks e and J) decreased by increasing DES treatment period, and completely disappeared after about six months treatment. This decrease is not a result of formation of less amounts of catecholestrogens, but rather reflects the presence of the enzyme systems to further metabolize any formed catechol estrogens, since the amount of catechol estrogens formed as detected by 3 H20 release is unchanged. The polar metabolites a, b and c may be some of these metabolites. They increased with DES treatment, and metabolite c appeared only after DES treatment. Estriol (peak d) and estrone (peak g) were detected, but were not affected by DES treatment, while no methoxyestrogens (peaks * and j) were isolated. A summary of these results is presented in Figure 29. 128 Metabolism of estradiol was also compared in kidney cell cultures from untreated hamsters using [6,7- 3 H]-estradiol and [4- 1 4 C]-estradiol as substrates to detect formation of metabolites which would release the tritium at positions 6 and 7. The metabolic profiles from these cell cultures were similar and there were no peaks that appeared by using one substrate, and not the other (Figure 30). 3.3.2 ESTROGEN METABOLISM IN MICROSOMAL PREPARATIONS Studies of estradiol metabolism in microsomal preparations showed a very low rate of metabolism, compared to the primary kidney cell cultures. In kidney microsomes, metabolite c appeared only in preparations from DES-treated animals, and the overall polar metabolites were of higher quantities in these preparations. No estrone, catecholestrogens or methoxyestrogens were detected, while estriol was unaffected by DES treatment (Figure 31). In liver microsomes, the same results were obtained, except that metabolite c appeared in preparations from untreated animals, and increased by DES treatment, and low amounts of catecholestrogens were detected as well (Figure 32). The appearance of polar metabolite c in liver preparations from untreated animals but not from kidneys of the same hamsters implies that it may have a role in tumorigenesis. 3.3.3 ESTROGEN METABOLISM IN TISSUE SLICES For these studies, a different gradient was used to better separate the region of polar metabolites. The gradient used was a biphasic one; from 30% to 50% methanol in water over 35 minutes, then 50% to 60% methanol in water over another 35 129 minutes. We studied the metabolism in both liver and kidneys and compared untreated hamsters to 4 month DES-treated hamsters. In kidney slices from untreated hamsters, approximately 30% of the substrate estradiol remained unmetabolized after 6 hours of incubation. The polar metabolite a was the main polar metabolite detected, with very little of metabolite b and no metabolite c. While no catecholestrogens were detected, a small quantity of estriol, and a large amount of estrone and methoxyestrogens were isolated (Figure 33). In kidney slices from 4 month DES-treated hamsters, a much higher amount of polar metabolites was detected, and metabolite c appeared after 6 hours incubation (Figure 34). In liver slices, the rate of metabolism was even higher, with only about 10 -15% of the substrate remaining after 6 hours of incubation. The polar metabolites a, b and c were detected in slices from untreated animals (Figure 35). These polar metabolites showed a quantitative increase in slices from DES-treated animals (Figure 36), especially metabolite c, while amounts of estrone and methoxyestrogens decreased slightly. Comparison of the amounts of the major metabolites formed in both tissue slices from untreated and DES-treated animals is shown in Figure 37. These results confirm the findings from experiments in cell cultures and microsomal preparations, but the results also provide a more complete picture of estrogen metabolism in the Syrian hamsters. The amounts of different metabolites produced by kidney slices (Figure 38) and liver slices (Figure 39) from untreated animals represent an average of three experiments. From triplicate incubations at 5 different concentrations of estradiol (10 nM -1 0 /xM), the kinetic parameters of the enzyme(s) that catalyze the formation of these metabolites were calculated using Cleland's weighted regression computer 130 program [292],and are reported in Tables 10 and 11. Enzymes catalyzing the formation of metabolites b and c were not determined in kidney slices due to the very small quantities produced. The apparent Km for the formation of estrone and methoxyestrogens were similar in liver and kidney, but the enzyme system that catalyzes the formation of polar metabolite a had an apparent K,^ that was 6-fold lower in kidney than in liver. The apparent Vmax for formation of estrone and methoxyestrogens was 100-fold lower in kidney compared to liver, while for the formation of polar metabolite a was less than 10-fold lower. These results indicate that the enzyme system responsible for the formation of polar metabolite a is much more active than other estrogen metabolism enzymes in the kidney and may thus account for the rapid conversion of any formed catecholestrogens to the more polar metabolites. Figure 27. Reverse phase HPLC radiochromatograms of organic extractable metabolites from kidney cell cultures of untreated and DES-treated hamsters (full view). Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak * and j). 131 cpm ek: b c b a peaks: 10,000 8,000 4,000 6,000 2,000 -— j "- 20 f h j i h g f e 40 Figure 27 60 80 ie (min.) time estradiol untreated 3 month Rx month 3 6 month Rx month 6 9 month Rx month 9 DES Treatment 13 month Rx month 13 Length of of Length to Figure 28. Reverse phase HPLC radiochromatograms of organic extractable metabolites from kidney cell cultures of untreated and DES-treated hamsters (expanded view). Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h ) and methoxyestrogens (peak i and j). 133 cpm 2,000 1,500 1,000 500 b c b a time(min.) Figure 28 h 9 l e 13 month Rx month 13 9 month Rx month 9 6 month Rx month 6 3 month Rx month 3 untreated estradiol Figure 29. The effect of DES treatment on major metabolites formation in kidney cell cultures 135 H untreated 3 month Rx H | 6 month Rx 9 month Rx jliijii 13 month Rx peaks a&b peak c estriol catechols estrone metabolite Figure 29 O n Figure 30. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney cell cultures of untreated hamsters, using [4-1 4 C]-estradiol and [6,7- 3 H]-estradiol. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and/), estrone (peak g)t estradiol (peak h) and methoxyestrogens (peak i and 7 ). 137 2,500 2,000 1,500 Substrate E Q. O 1,000 500 [6,7-H]-E2 r v V/l 0 40 60 80 time (min.) peaks: e f g h ij Figure 30 Figure 31. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney microsomes of untreated and 9 months DES- treated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h ) and methoxyestrogens (peak i and j). 139 2,500 2,000 1,500 E Q. Length of O DES Treatment 1,000 9 month Rx 500 untreated 0 20 40 60 80 peaks: a b c d e f g h i j time (min.) Figure 31 o Figure 32. Reverse phase HPLC radiochromatogram of organic extractable metabolites from liver microsomes of untreated and 9 months DES-treated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and/), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and j). 141 2,500 r 2,000 1,500 E Q. Length of O DES Treatment 1,000 9 month Rx 500 untreated 0 20 40 60 80 peaks: a b c d e f g h i j time (min.) Figure 32 K> Figure 33. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamster. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and/). 143 peaks: cpm 14.000 10,000 12.000 12.000 2.000 4.000 6,000 8,000 Time (min.)Time Figure 33 Figure g i j i h g f 0 hour 0 3 hour3 6 hour 6 Incubation Incubation Period Figure 34. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of 4 months DES-treated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and j), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and j). 145 12,000 r 10,000 8,000 Incubation Period q . 6,000 / ; u / 6 hour 4,000 VV-J 3 hour 2,000 0 hour 40 60 80 Time(min) peaks: a b c d e f g h i j Figure 34 0\ Figure 35. Reverse phase HPLC radiochromatogram of organic extractable metabolites from liver slices of untreated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and/). 147 cpm 12,000 10,000 ek: f e d c b a peaks: 8,000 4,000 6,000 2,000 0 20 Figure 35 Figure 40 Time(min) 60 ghij 80 Ohour 3 hour 3 Incubation 6 hour 6 Period oo Figure 36. Reverse phase HPLC radiochromatogram of organic extractable metabolites from liver slices of 4 months DES-treated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and j). 149 cpm 400 r 14,000 12,000 10,000 ek: f e d c b a peaks: 8,000 4,000 6,000 2,000 0 20 40 Time(min) Figure 36 Figure 60 ghij 80 0 hour 0 3 hour 3 Incubation 6 hour 6 Period Figure 37. Polar metabolite formation in kidney and liver slices of untreated and 4 months DES-treated hamsters. Slices were incubated for six hours with IfxM estradiol 151 Amount produced (pmol/mg/min) 0.4 0.1 0.2 - 0.3 - - POLAR A Liver-untreated Kidney-untreated Figure 37 Figure POLAR B POLAR C POLAR B POLAR I Kidney-treated I I Liver-treated U\ 3 S Figure 38. Major metabolite formation in kidney slices from untreated hamsters. Slices were incubated for six hours with 10 nM - 5 estradiol 153 E i -B- MeO-E’s - - 0 ...... polar A ----- 3+S------polar B polar C [estradiol] //M Figure 38 4^ Figure 39. Major metabolite formation in liver slices from untreated hamsters. Slices were incubated for six hours with 10 nM - 10 /*M estradiol 155 pmol metabolite/mg/min o 2 4 [estradiol] [estradiol] 6 fiM 8 Figure 39 Figure 10 12 MeO-E’s - ■ B - 3 I - polar C polar polar B polar A polar E 1 E ----- ...... O n 157 Table 10. Values of apparent Km +. S.E. of enzymes catalyzing the formation of the major metabolites in liver and kidney slices from untreated hamsters Metabolite Liver slices Kidney slices Estrone 5.40 ± 3.50 7.43 +. 1.61 Estriol 10.97 + 3.80 N/D 2-Methoxyestrone 10.61 ± 4.10 7.20 +. 0.49 Peak a 17.37 + 6.40 2.81 + 0.93 Peak b 4.69 + 2.00 N/D Peake 9.12 + 9.90 N/D 158 Table 11. Values of apparent Vmax ± S.E. of enzymes catalyzing the formation of the major metabolites in liver and kidney slices from untreated hamsters Metabolite Liver slices Kidney slices Estrone 1.30 ± 0 .3 9 0 . 0 1 0 2 ± 0 . 0 0 1 2 Estriol 0.58 ± 0 .1 1 N/D 2-Methoxyestrone 0.34 ± 0,08 0.0042 ± 0.0001 Peak a 0.02 ± 0.005 0.0031 ±0.0003 Peak b 0 . 0 1 ± 0 . 0 0 1 N/D Peak c 0.01 ± 0.005 N/D 3.4 KINETICS OF ESTROGEN 2/4 HYDROXYLASE Previously in our laboratory, the estrogen 2-hydroxylase enzyme activity in liver and kidney microsomes from Syrian hamster was characterized using the 3 H20 assay, and the reported apparent Km for kidney microsomes was 6.43 ± 3.22 /xM with an apparent Vmax of 0.051 ± 0.016 nmol/min/mg, and in the liver microsomes, an apparent Km of 2.86 ± 1.40 /xM and an apparent Vmax of 0.129 ± 0.015 nmol/min/mg [293]. These studies were extended to determine the same parameters for the estrogen 4-hydroxylase enzymatic activity, using the 3 H20 assay and Cleland's weighted regression computer program. The apparent Km for this enzyme in freshly prepared liver microsomes is 11.42 ± 2.1 M, with an apparent Vmax of 0.017 ± 0.002 nmol/min/mg, while in kidney microsomes the apparent Km was 13.9 ± 7 .1 HM and the apparent Vmax of 0.061 ± 0.016 nmol/min/mg. Figure 40 compares these values along with values obtained from frozen microsomes. While the apparent Vmax for estrogen 2 -hydroxylase in the liver is almost 1 0 -fold higher than that of estrogen 4-hydroxylase, in the kidney the apparent Vmax of these enzymes are equal. This agrees with reports that the amount of 2-hydroxy and 4-hydroxyestrogens formed are equal in the kidneys, while in the liver the 2 -hydroxylated product is dominant. When comparing results obtained from fresh and frozen preparations, the apparent Vmax is about 10 fold higher in fresh preparations. This is probably due to the partial degradation of the enzyme upon freezing. The apparent Km values were different in fresh and frozen preparations, as well, but the standard errors were high. Dannan et al reported the presence of at least eleven different estrogen 2/4-hydroxylase isozymes in rat liver [294]. The reason for the differences in apparent Km values between fresh 160 and frozen preparations and the high standard errors obtained may be due to the presence of similar different isozymes in hamster kidney and liver responsible for the hydroxylation and/or differences in isozyme degradation upon freezing. Figure 40. Kinetic parameters for estrogen 2- and 4-hydroxylase in kidney and liver microsomes of untreated hamsters 161 2-Hydroxylase app. Km (-fc-S.E.) microsomes fresh liver 2.8 fiM (1.4) frozen liver 2.5 /iM (1.0) fresh kidney 6.2 /iM (3.2) frozen kidney 10.2 juM (4.8) 4-Hydroxylase fresh liver 11.4 /iM (2.1) frozen liver 8.4 /iM (5.2) fresh kidney 13.9 juM (7-1) frozen kidney 4.5 juM (2.2) ” 1 i I i I 1 1 i I ~ 0.00 0.02 0.04 0.06 0.08 0.10 0.12 0 .1 4 0 .1 6 app. V max(nmol/mg/mln) Figure 40 0to\ 163 DISTRIBUTION OF ESTROGEN METABOLITES IN KIDNEY AND LIVER SLICES The distribution pattern of various metabolites produced from liver and kidney slices were examined. In liver slices, the amount of steroids retained within the slices increased gradually, reaching 40-45 % of total steroids after 6 hours, and was unaffected by DES treatment. In kidney slices, the amount retained in the slices from DES-treated hamsters was higher after 3 hours, but then increased slowly to reach the same levels as in kidney slices from untreated hamsters after six hours (Figure 41). Within the slices, the distribution to different cellular compartments in both liver and kidney slices revealed equal amounts of steroids in each fraction after 6 hours, although the cytoplasmic fraction contained much higher amounts after 3 hours in liver slices (Figure 42). In the kidney slices, very small amounts of water soluble metabolites (about 3-5%) and protein bound steroids (3-4%) were formed, while organic soluble materials were more than 90%. In the liver slices, about 20-25% of the metabolites are water soluble, while 15-20% are bound to proteins (Figure 43). Detection of the metabolites that were retained in the cellular compartments from liver slices showed that while the 1 0 ,0 0 0 xg fraction, containing nuclear fragments and large pieces of the plasma membrane, and the microsomal pellets retained mainly estrone and estradiol, metabolites from the cytoplasmic fraction included polar metabolites, estriol, estrone and methoxyestrogens, but no catecholestrogens (Figure 44). Slices from DES-treated animals showed the same patterns as discussed above. Figure 41. Distribution of metabolites in liver and kidney slices from untreated and 4 months DES-treated hamsters. Slices were incubated for 6 hours with 1 jtiM estradiol 164 100 LIVER SLICES KIDNEY SLICES 160 0 4 0 0 4 0 OL time(hour) time(hour) Supernatant-untreated Slice-untreated Supernatant-treated Slice-treated Figure 41 Figure 42. Distribution of metabolites in different cellular compartments in liver slices of untreated hamsters. Slices were incubated for 0, 3 and 6 hours with 1 /aM estradiol 166 100 0 hr. Incubation 3 hr. incubation 6 hr. incubation Supernatant Layer | Cytoplasmic Fraction §j§ 10,000g Fragment II Microsomal Fraction Figure 42 Figure 43. Organic soluble, water soluble and protein bound metabolites in liver slices of untreated hamsters. Slices were incubated for 0, 3, and 6 hours with 1 and 10 /tM estradiol 168 PERCENT OF STEROID 100 80 60 20 TIMEINCUBATION OF (HOURS) Figure 43 Figure ORGANIC SOLUBLE ORGANIC SOLUBLE [ESTRADIOL]=1 C^uM40 [ESTRADIOL]=1 G^M [ESTRADIOL] =1 tyiM [ESTRADIOL]=1//M [ESTRADIOL]=1/iM [ESTRADIOL]=1/ WATER SOLUBLE WATER SOLUBLE PROTEINBOUND PROTEINBOUND — . 0 — j M vo as Figure 44. Reverse phase HPLC radiochromatogram of organic extractable metabolites from different cellular fractions in liver slices of untreated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and,/). 170 Peaks: a b b a Peaks: cpm 1,200 1,000 400 600 800 200 0 20 c c d e f f e d Figure 44 Figure time(min) 40 080 60 g g hi j 10,000g fragment 10,000g i v i~ i fri ^ cytopl microsomal Cellular fraction ilasmic action pellets 172 3.6 CHARACTERIZATION OF WATER SOLUBLE METABOLITES Conjugation of estrogens to water soluble metabolites occurs via formation of glucuronides, sulfates and thioethers. The presence of these metabolites were detected in liver and kidney slices of Syrian hamsters (Tables 13 and 14). The amounts of these metabolites were not changed in slices from hamsters treated with DES. Cleavage of glucuronides and sulfates was accomplished using specific enzymes, while thioethers were reduced by Raney nickel to liberate the free steroids. The chemical nature of the liberated metabolites was further examined and separated on reverse phase HPLC. In the liver slices, metabolites that form sulfates and thioethers are mainly estrone and estradiol, while glucuronidated metabolites include catecholestrogens and polar metabolite a as well (Figure 45). In the kidney slices, the evaluation of thioether metabolites was not accomplished due to very low amounts isolated, but sulfates were mainly conjugated estradiol, and glucuronides resulted from polar metabolite a, catecholestrogens, estrone and estradiol (Figure 46). 173 Table 12. Percent of water soluble metabolites formed in liver slices from untreated hamsters. Slices were incubated for 3 and 6 hours with 1 piM estradiol Percent of Water Percent of total Conjugate Soluble Metabolites Metabolites Glucuronides 3 hour 20.57 ± 1.21 3.20 + 0.009 6 hour 27.07 ± 1.63 4.75 ± 0.062 $ulfate$ 3 hour 12.76+ 1.32 1.99 + 0.01 6 hour 23.06 + 2.88 3.96 ±0.11 Thioethers 3 hour 19.60 ± 1.58 3.06 ±0.012 6 hour 22.20 + 4.38 3.81 ± 0 .1 7 174 Table 13. Percent of water soluble metabolites formed in kidney slices from untreated hamsters. Slices were incubated for 3 and 6 hours with 1 /iM estradiol Percent of Water Percent of total Conjugate Soluble Metabolites Metabolites Glucuronides 3 hour 29.99 ± 2.25 0.23 ± 0.0001 6 hour 32.77 + 0.88 0.33 + 0.00004 Sulfates 3 hour 2 2 .11+ 0.70 0.17 + 0.0004 6 hour 25.75 ± 1.02 0.26 ± 0.005 Thioethers 3 hour 31.69 + 0.42 0.25 + 0.0002 6 hour 32.91+0.92 0.33 + 0.0004 Figure 45. Reverse phase HPLC radiochromatogram of organic extractable metabolites after cleaving the water soluble metabolites formed in liver slices of untreated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and/), estrone (peak g), estradiol (peak h ) and methoxyestrogens (peak i and j). 175 Peaks: a b c d d c b a Peaks: cpm 1,000 200 400 600 800 0 0 20 iemn ^ . . . . ^ time(min) 40 Figure 45 Figure efghij 080 60 Glucuronides Sulfates \ Thioethers conjugate 0\ -4 Figure 46. Reverse phase HPLC radiochromatogram of organic extractable metabolites after cleaving the water soluble metabolites formed in kidney slices of untreated hamsters. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and f), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and j). 177 ek:ab a Peaks: cpm 1,000 200 400 600 800 20 time(min) Figure 46 Figure 6040 80 Glucuronides Sulfates conjugate 00 - j 179 3.7 INHIBITION STUDIES BY BROMOESTROGENS The effects of the estrogen 2/4-hydroxylase inhibitors, 2-bromoestradiol, 4- bromoestradiol and 2,4-dibromoestradiol (Figure 47), on estradiol metabolism in kidney slices were investigated. These three inhibitors showed similar overall inhibition of estradiol metabolism, where twice the amount of unmetabolized estradiol was isolated from the slices that recieved 5 /xM of the inhibitor, compared to preparations recieving no inhibitors. The amounts of polar metabolites a and b were decreased significantly, along with levels of estrone and methoxyestrogens. Figures 48, 49 and 50 show HPLC separation of metabolites formed in slices with or without inhibitors, and Figure 51 summarizes the effect of 4-bromoestradiol on the formation of the major metabolites. The distribution of metabolites was affected only slightly, with fewer metabolites retained in the slices incubated with the inhibitor, while amounts of water soluble metabolites were unchanged. These results confirm that the polar metabolites arise through oxidative metabolism of estrogens, probably by further metabolism of catecholestrogens, through pathways that may form reactive intermediates and free radicals that may interact with proteins and DNA and cause cell damage. 180 OH OH Br HO HO Br 2-Bromoestradiol 4-Bromoestradiol OH Br HO Br 2,4-Dibromoestradiol Figure 47. Structures of bromoestrogens Figure 48. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters in presence of 2-bromoestradiol. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and f), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and/). 181 1,0,000 8,000 g- 6,000 4,000 J!L / V 5 m 2,000 2 m o m __ i______40 60 80 time (min) peaks: e f g h ij Figure 48 00N> Figure 49. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters in presence of 4-bromoestradiol. Metabolites are polar metabolites (peak a, b and c), estriol (peak d), catecholestrogens (peak e and J), estrone (peak g), estradiol (peak h) and methoxyestrogens (peak i and j). 183 10,000 8,000 E g - 6 , 0 0 0 4.000 5/M 2.000 0 40 60 80 time (min) peaks: e f g h ij Figure 49 00 Figure 50. Reverse phase HPLC radiochromatogram of organic extractable metabolites from kidney slices of untreated hamsters in presence of 2,4-dibromoestradiol. Metabolites are polar metabolites (peak a, b and c), estriol (peak rf), catechol estrogens (peak e and J), estrone (peakg), estradiol (peak h) and methoxyestrogens (peak i and j). 185 8,000 r 6,000 E Q. O 4,000 2,000 20 40 time (min) peaks: a b e f Figure 50 Figure 51. Inhibition of the formation of polar metabolites, estrone and methoxyestrogens by 4-bromoestradiol in kidney slices of untreated hamsters 187 0 .8 ffi c* = 1 I f 0.6 Q . S =3 8 C O Q . E 0.4 < 0.2 JL Polar A Estrone Methoxy E’s ■ [l]=0jUM □ [l]=2yUM [i]=5^M Figure 51 00 oo 189 3.8 APPROACHES TO THE IDENTIFICATION OF METABOLITES Investigations to identify polar metabolite c were initiated. This metabolite appears only after DES treatment in kidneys of Syrian hamsters, while it appears in the liver of untreated hamsters. It appears in the kidney after three months of DES treatment, which coincides with the appearance of other morphological changes such as the appearance of dysplasic foci and the increase in area of vascularization [128], thus might play a role in tumorigenesis. Using electron ionization mass spectroscopy, an M + peak with a mass of 288 was identified. The exact mass revealed that the compound is a trihydroxylated derivative of estradiol. The relative intensities and the molecular formulae of the M + peak and other peaks obtained from fragmentation of the parent compound is shown in Table 14. Desorption ionization mass spectroscopy was inconclusive. For further identification of this metabolite, several trihydroxylated estrogens were injected on reverse phase HPLC, including hydroxylated derivatives of estradiol at 2, 4, 6 a, 11a and 16a. None of these compounds co-eluted with metabolite c. Accordingly, metabolite c was tentatively assigned 15a-hydroxyestradiol. This is consistent with the fragmentation patterns from electron ionization mass spectroscopy and relative retention time on reverse phase HPLC as reported in recent literature [83, 295], Further identification of the exact chemical structure of this metabolite may unravel more information about its role in tumorigenesis. 190 Table 14. Partial listing of peaks identified in electron ionization mass spectroscopy of metabolite c, with their relative intensities and calculated molecular formulas Molecular Weight Relative intensity Molecular formula 288 7 C1 8 H2 4 O3 261 4 C1 7 H2 5 O2 217 16 C is^iO x 175 89 ^12^150! 149 72 CioH^Oj 147 1 2 2 CioHnOj CHAPTER IV SUMMARY The golden Syrian hamster develop renal adenocarcinomas upon exposure to estrogens for a prolonged period of time [98]. The tumorigenic process has been postulated to involve two estrogenic effects: the hormonal effects and the carcinogenic toxicity [134]. The hormonal effects arise from receptor binding, followed by a cascade of events leading to an increase in gene expression, DNA synthesis and stimulation of protein production. Evidence for the importance of the hormonal effect in tumorigenesis in the Syrian hamster has been indirectly demonstrated. Antiestrogens inhibit tumor formation act by competitive inhibition of receptor binding [110], and estrogens that possess weak potency, such as estradiol 17-a fail to induce tumors [102]. However, very little direct evidence exists in the literature about the stimulation of protein production in estrogen treated animals, the induction of specific growth factors, or oncogene activation in this animal model. DES treatment caused an overall increase in protein production, with a 3 fold increase after nine months treatment and a 2 0 fold increase in tumor-bearing hamsters. Exposure of kidney cell cultures from these hamsters to 10 nM estradiol showed about 2-3 fold increase in total protein production. 191 192 Three protein bands showed a consistent stimulation by DES treatment, whereas a 46 kDa protein band that was produced in similar quantities in both untreated and DES-treated hamsters for varying periods of time. These DES-stimulated protein bands had molecular weights of 34-36 kDa, 20-22 kDa and 16-18 kDa. Short term exposure of the kidney cell cultures to 10 nM estradiol showed an increase in the production of these proteins only in cultures of untreated hamsters, but not DES- treated ones. The 16-18 kDa protein band possessed a high isoelectric point of approximately 8-9. Growth factors have been implicated to play a role in carcinogenesis in many tumors including breast cancer [ 8 6 ]. To explore if these stimulated proteins belong to this family of polypeptides, specific antibodies to two of these growth factors, TGF-a and bFGF, were used in immunoprecipitation studies. The results demonstrated the presence of a TGF-a-like protein with a molecular weight of 20-22 kDa, and a bFGF- like protein that had a molecular weight of 16-18 kDa. Both of these proteins were absent in kidney cell cultures of untreated hamsters, but were produced from cultures of 5 month DES-treated hamsters. While the TGF-a-like protein was not stimulated by short term exposure to 10 nM estradiol, bFGF-like protein was induced by such treatment. These growth factors may play a role in cell proliferation in the kidney of the Syrian hamster. The bFGF-like protein may be responsible, at least in part, for the large increase in the area of vascularization observed in kidney of 3 month DES-treated hamsters [128], an important phenomena for the well-being of solid tumors. Further studies on the time course for appearance of these proteins during the DES treatment of the Syrian hamster may provide information on potential on potential roles in tumorigenesis. 193 The carcinogenic toxicity of estrogens is postulated to arise from the formation of reactive metabolites and free radicals, that can bind to proteins and form DNA adducts leading to cellular damage, possibly mutations and tumor formation [273]. Estrogen metabolism was investigated in the kidney of the Syrian hamsters, the site of tumor formation, and compared to the results obtained from the same studies in the liver, the main site of metabolism in the body and the organ that shows no estrogen- dependent tumor formation. The present studies showed that the amounts of catechol estrogens formed from estradiol in kidney of Syrian hamsters diminished by prolonging DES treatment. This decrease reflected the further metabolism of these catechol estrogens to other highly polar metabolites. Polar metabolites were separated to three compounds a, b and c, which showed a quantitative increase after DES-treatment. While metabolite a did not change with estrogen treatment, metabolite b showed a significant increase in hamsters treated with DES compared to untreated hamsters. Metabolite c was not isolated in untreated hamsters, but appeared in DES-treated hamsters. This metabolite may be important in tumorigenesis, due to its absence in kidneys of untreated hamsters, but was present in livers of these hamsters. It also appears in kidneys after 3 months of DES treatment, which coincides with other changes that are related to tumor formatiom such as increase in area of vascularization and appearance of dysplasic foci [128]. Preliminary studies to identify this metabolite showed that it is a trihydroxylated compound that has been tentatively assigned as 15a-hydroxyestradiol. The complete elucidation of the chemical structure of this metabolite may shed some light on its role in tumorigenesis, and whether it is metabolite c itself or some reactive intermediates and/or free radicals that may arise during the pathway leading to its formation that may bind cellular proteins or form DNA adducts and cause cell 194 damage. Among the other metabolites formed in the kidneys of Syrian hamsters are estriol and other 16-hydroxylated compounds, estrone and methoxyestrogens. These metabolites are affected only minimally by DES treatment. The study of the kinetics of the microsomal enzymes that catalyze the formation of these metabolites demonstrated that the apparent Vmax of these enzymes was higher in liver than in kidney of Syrian hamsters, while the apparent Km was similar. One exception was the enzyme system catalyzing the formation of metabolite a, where it had a 6 -fold lower apparent Km in kidney than in liver. Moreover, while the apparent Vmax f°r enzymes catalyzing the formation of estrone and methoxyestrogens was 1 0 0 - fold lower in kidney than in liver, the apparent Vmax f°r the enzyme system responsible for metabolite a formation was less than 1 0 -fold lower, indicating that this enzyme system is very active in the kidney compared to other metabolic enzymes. Estrogen 4-hydroxylase activity catalyzes the conversion of estrogens to the 4-hydroxylated catecholestrogen. The evaluation of the kinetics of this enzyme in the Syrian hamsters revealed that it has a 2-fold higher apparent Km than estrogen 2-hydroxylase in kidney, with a similar apparent Vmax. In liver, the apparent Km of estrogen 4-hydroxylase was about 4-5 fold higher than estrogen 2-hydroxylase, and the apparent Vmax was about 1 0 -fold lower. These results agree with other reports that both the 2- and the 4-hydroxylated catechol estrogens are formed in equal amounts in the kidney, but that the 2 -hydroxyestrogens is the predominant in the liver of Syrian hamsters [240]. Other reports indicate that the O-methylation of 4-hydroxyestrogens is inhibited by 2-hydroxyestrogens [244]. This may lead to the accumulation of this catechol estrogen and its availiability to undergo further metabolism and may be the precursor of the damaging reactive compounds. 195 Binding of estrogens to cellular proteins and formation of water soluble metabolites are very low in the kidney of Syrian hamsters compared to the liver. However, a significant amount of these metabolites were found to be retained in the nuclear and microsomal fractions of the cells (about 30% of total steroids after 6 hours incubation). Estrone and estradiol were detected as the only organic extractable metabolites from these cellular fractions, while the whole spectrum of metabolites were isolated from the cytoplasmic fraction. This suggests that lipophilicity is an important factor in cellular distribution of metabolites. The formation of water soluble conjugates was demonstrated in both the liver and the kidney of Syrian hamsters. Glucuronide, sulfate and thioether conjugates of estrone and estradiol were detected, and glucuronides of catechol estrogens and metabolite a were also isolated. The amount of these metabolites formed in the kidney constituted a very low percentage of total metabolites while higher amounts were formed in the liver, consistent with a higher capacity of liver to conjugate metabolites. These observations suggest that the decreased steroid conjugation in kidney results in greater availability of estrogen metabolites which can participate in the formation of damaging reactive species. Estrogen metabolism in kidney of Syrian hamsters was inhibited in the presence of the estrogen 2/4 hydroxylase inhibitors, the bromoestrogens, with a significant drop in the amount of polar metabolites formed. This supports the hypothesis that these polar metabolites arise through oxidative metabolism of estrogens, probably resulting from further metabolism of catecholestrogens. Figure 52 summarizes the possible metabolic pathways involved in estradiol metabolism in the different preparations from Syrian hamsters. Figure 52. Possible metabolic pathways involved in estradiol metabolism in different preparations from Syrian hamsters. 196 c s m c ^HO HO 15-aOH-E m c s m c s 2-MeO-E 2-OH-E OH 4-OH-Ea MeO 4-MeO-E g (2-MeO-E,) (2-OH-E) (4-OH-E,) (4-MeO-E,) m c s m c s OH - i polar -mcs mcs polar metabolites metabolites protein- m c s m c f protein- bound bound Detected in: m = microsomal assays c = primary kidney cultures s = tissue slices Figure 52 VO - j BIBLIOGRAPHY 1. Breasted, J. H. 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