User:Drbogdan/BogdanDennis-PhD-Dissertation-1973-TEXT

The exact mechanism of chemical carcinogenesis remains unclear, although it is generally agreed that modification of genetic expression constitutes an essential step in the process. The possibility that DNA itself represents the crucial biological target of chemical carcinogens is supported by in vivo studies, involving various polycyclic hydrocarbons, demonstrating a positive correlation between carcinogenicity and covalent binding to DNA. Polycyclic hydrocarbons are relatively unreactive chemically and, apparently, require biological activation for covalent bond formation with DNA in vivo. The overall objectives of this research are: first, to study the biotransformation of the polycyclic hydrocarbon carcinogen benzo(a)pyrene in mammalian systems; also, to study in the presence of such biotransformation the interaction between benzo(a)pyrene and polynucleotides, including mammalian DNA; and finally, to study the biological significance of the above interactions.

A comparison of characteristics of benzo(a)pyrene biotransformation processes in various tissues of the human and rat fetal-placental unit was attempted and was based on known features of aryl hydrocarbon hydroxylase (a NADPH-dependent mixed-function oxidase system able to transform polycyclic hydrocarbons) including inducibility, subcellular localization, requirements for optimal activity and the effect of various agents. Although differences were

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observed between enzyme systems of the various tissues studied, the similarities appeared more striking and provided a basis for choosing the adult rat liver system as a model suitable for studies dealing with the remaining objectives of the present research.

Inclusion of calf thymus DNA in incubation mixtures containing benzo(a)pyrene and the model biotransformation system substantially reduced the recovery of BaP products as compared to controls. Similar decreases were observed with Poly G and, to lesser extents, with Poly I and the alternating copolymer Poly (I-C). No effects were observed with the double-stranded Poly I-Poly C, Poly A, Poly U nor with DNA modified with nitrogen mustard; increases were observed with yeast RNA and denatured DNA. The decrease in recoverable benzo(a)pyrene products was directly correlated with covalent binding of benzo(a)pyrene and polynucleotides as determined by sedimentation studies. Neither DNA nor Poly G enhanced the enzymatic disappearance of recovered benzo(a)pyrene products. The above findings suggest that benzo(a)pyrene, as a result of biotransformation, interacts chemically and specifically with base residues on the native DNA molecule in a manner which, apparently, involves a reactive BaP intermediate product.

DNA treated with benzo(a)pyrene or with the noncarcinogenic benzo(e)pyrene in the presence of the rat liver aryl hydrocarbon hydroxylase system exhibited decreases in template abilities as measured in an E. Coli DNA-dependent

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RNA polymerase system when compared to template abilities of DNA controls. These findings suggest that DNA, treated enzymatically with benzopyrenes, is biologically modified but in a manner which does not appear to be carcinogen-specific.

Apparently, as a result of the described studies, benzo(a)pyrene can biologically and chemically modify mammalian nucleic acid in the presence of a mixed-function oxidase system found in rat and human tissues in a reaction which appears to involve an enzyme generated benzo(a)pyrene intermediate product, to be dependent on the secondary or tertiary structure of nucleic acids and to be base-specific. These in vitro findings appear to be consistent with the general hypothesis that initiation of carcinogenic processes in mammalian systems in vivo can be a result of chemical reactions between biologically activated chemical carcinogens and critical sites on cellular macromolecules.

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ACKNOWLEDGEMENT edit

I wish to express my deep appreciation to Dr. Zdzislaw F. Chmielewicz, Dr. Lorne K. Garrettson, Dr. Peter M. Hebborn, and Dr. Thomas I. Kalman for their help, criticism and encouragement during the course of this research, without which this thesis could not have been completed in its present form.

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For my wife, Jan, without whom . . .

TABLE OF CONTENTS edit

INTRODUCTION-------------------------------------------------------------------------------------1-19

MATERIALS------------------------------------------------------------------------------------------20-22

METHODS

PREPARATION OF HOMOGENATES IN FETAL-

PLACENTAL UNIT STUDIES-------------------------------------------------------------------23-25

PREPARATION OF RAT LIVER MICROSOMES------------------------------------------25-26

ENZYME ASSAYS IN FETAL-PLACENTAL UNIT STUDIES---------------------------26-27

ASSAY FOR ARYL HYDROOCARBON HYDROXYLASE------------------------------28-30

PREPARATION OF FLUORESCENT BaP

HYDROXYLATED PRODUCTS---------------------------------------------------------------30-31

SUCROSE GRADIENT SEDIMENTATION STUDIES----------------------------------31-32

PREPARATION OF DNA MODIFIED WITH

NITROGEN MUSTARD------------------------------------------------------------------------32-33

PREPARATION OF DNA FOR TEMPLATE STUDIES---------------------------------33-35

ASSAY FOR DNA TEMPLATE ACTIVITY-------------------------------------------------35-36

PREPARATION OF RNA POLYMERASE------------------------------------------------36-40

DETERMINATION OF DNA-----------------------------------------------------------------40

DETERMINATION OF RNA------------------------------------------------------------------40-41

DETERMINATION OF PROTEIN-----------------------------------------------------------41

RESULTS

BIOTRANSFORMATION OF BaP IN VARIOUS

TISSUES OF THE HUMAN AND RAT FETAL-

PLACENTAL UNIT----------------------------------------------------------------------------42-50

EFFECT OF CALF THYMUS DNA ON THE APPEARANCE

OF FLUORESCENT PRODUCTS OF BaP BIOTRANSFORMATION-----------50-57

EFFECT OF VARIOUS MONONUCLEOTIDES AND SYNTHETIC

POLYNUCLEOTIDES ON THE APPEARANCE OF FLUORESCENT PRODUCTS

OF BaP BIOTRANSFORMATION--------------------------------------------------------57-60

EFFECT OF CALF THYMUS DNA MODIFIED

BY NITROGEN MUSTARD ON THE APPEARANCE

OF FLUORESCENT PRODUCTS OF BaP BIOTRANSFORMATION---------60-61

EFFECT OF YEAST RNA AND DENATURED

CALF THYMUS DNA ON THE APPEARANCE

OF FLUORESCENT PRODUCTS OF BaP

BIOTRANSFORMATION-----------------------------------------------------------------61-63

BIOLOGICAL ACTIVITY OF CALF THYMUS

DNA CHEMICALLY MODIFIED BY BaP

BIOTRANSFORMATION-----------------------------------------------------------------63-65

DISCUSSION----------------------------------------------------------------------------154-167

BIBLIOGRAPHY-------------------------------------------------------------------------168-177

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INTRODUCTION edit

Cancer has been defined as a growth of progressively dividing, undifferentiated cells which possesses a nearly complete independence as a biological entity from the host organism (1). Agents which are capable of producing cancers in man and other animals in one or several organs or tissues, regardless of the route of exposure and the dose and physical state of the agent used, are considered carcinogens (2). It is now appreciated that many biological, physical or chemical agents are able to provoke the carcinogenic process, the more notable of these agents being viruses, radiation and chemical substances.

Among the causes of human malignancies, chemicals are the most important. According to a recent estimate, chemical substances, either endogenous or environmental, are thought responsible for 90% of the cancers occurring in man (3). Furthermore, the World Health Organization concluded in 1965 that at least one-half of all cancer in man is due to environmental factors (4). Although many chemical carcinogens are well-known environmental contaminants (5,6), experimental proof that human cancer results from exposure to chemical carcinogens is not available.

The relationship between chemicals and carcinogenesis was first noted as early as 1775 when Sir Percivall Pott, a prominent eighteenth-century English surgeon, recognized a high incidence of scrotal cancer in London chimney sweeps and presumed the cause to be due to the chronic exposure of

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such workers to chimney soot (7). In the following century other examples of skin cancer produced by occupational exposure to coal tar and related industrially used materials appeared in the literature (8). But it was not until the studies of Yamgiwa and Ichikawa (9) in 1915 that cancer was produced experimentally, for the first time, by applying coal tar to an animal’s skin.

Later, during the 1920’s and early 1930’s, fundamental studies were performed which revealed the carcinogenic potency of coal tars (10), identified the major active components of coal tar (11) and characterized the carcinogenic properties of a pure hydrocarbon, 1,2,5,6-dibenzanthracene (12). Besides polycyclic hydrocarbons, chemicals now known to elicit strong carcinogenic responses in animals include certain aromatic amines, azo dyes and nitrosamines (13,14,15). Alkylating agents, such as nitrogen mustards, bisulfan, ethionine, and the lactones, β-propiolactone, ethylenimines, epoxides and the fungal product, aflatoxin, also have demonstrated carcinogenic potency (16,17,18).

Whereas the bulk of earlier studies involving chemical carcinogens dealt with the isolation, synthesis and chemical characterization of active substances, studies in more recent years have been concerned with the interaction of carcinogenic chemicals with biological systems. The finding that certain chemical carcinogens depend on biological transformation for activity (19) has promoted considerable research interest in the biological handling of carcinogenic

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substances, primarily to identify ultimate carcinogenic substances as well as to elucidate essential steps in their production.

The manner by which carcinogenic substances initiate malignant growth is far from understood at the moment. However, as stated by Miller and Miller in 1966 (19), “Cellular macromolecules appear to be the fundamental targets of carcinogens since only macromolecules appear capable of storing, replicating and transforming the information needed in the growth of a cell and in its control.” Further, in the same review, “Conversion of a normal cell to a cancer call and propagation of the latter state, indefinitely through numerous cell divisions, demands that there be some change in the informational content of the cell. On the basis of modern molecular genetics and the ability to induce mutations by chemical modification of DNA, reaction with DNA is a particularly attractive model for chemical carcinogenesis.” Correlations between the carcinogenicity of chemicals and the binding of such carcinogens to cellular macromolecules, including DNA, in vivo (20,21) and in vitro (22,23), have encouraged attempts to identify the crucial biological receptor with which the ultimate carcinogen interacts to initiate the carcinogenic process.

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In order to provide insights into the mechanism of chemical carcinogenesis, the overall objective of the present research is to study the biological transformation of BaP in mammalian tissues and the interaction of the products of such transformation with DNA and other polynucleotides.

A. CARCINOGENIC PROPERTIES OF BENZO(A)PYRENE edit

BaP (Fig.1), a polycyclic aromatic hydrocarbon, is a planar molecule containing a phenanthrene nucleus and is relatively inert chemically. Besides being found in coal tar (26,27), BaP is now known to be present in certain cooked and smoked foods (28, 29, 30, 31), polluted city air (32,33) and tobacco smoke (34, 35, 36, 37). With regard to tobacco smoke, BaP is considered the most potent of the coal tar carcinogens present and is found in the greatest amount (38).

The carcinogenic properties of BaP have been extensively studied since 1932 when Cook (39) found the major active substance in coal tar to be BaP by studying skin tumor formation in mice treated topically with coal tar, BaP derived from coal tar and synthetic BaP. Subsequent research extended these initial findings and demonstrated BaP-induced malignant tumor growth in most animal species studied, including mice (40,41), rats (42,43), rabbits (44) and guinea pigs (45). One early study involving 26 human volunteers reported the absence of skin tumor development arising from the application of a 1% BaP solution in benzene to the skin daily over a four month period (46). Follow-up studies on these subjects, however, have not been published.

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BaP, like most carcinogenic polycyclic hydrocarbons, produces cancers at the site of application. Other than skin cancer induced by painting animal skin with a solution of BaP, gliomas and sarcomas have been observed when pellets containing BaP were implanted in the brain of mice (47). Furthermore, carcinomas developed in the salivary gland of rats (48) and vagina of mice (49) when these organs were implanted with pellets containing BaP. Parenteral administration of BaP solutions have elicited neoplasms of lung and spleen in mice (50) and liver and bone marrow in rats (51,52). Adenocarcinoma of the stomach developed in mice fed a diet containing BaP (53).

Based on several indices designed to measure the biological response to carcinogenic substances, BaP is among the most potent carcinogens known and often has been regarded as a standard in determining the carcinogenic potency of other agents. One established carcinogenic index is that of Iball (54) and is defined as 100 times the number of animals bearing tumors divided by the number of animals living at the day of appearance of the first tumor and by the mean number of days from the application of substance to the appearance of tumors. Based on data published by Heiger (55), the carcinogenic index of BaP when applied to the skins of C57 mice is 65. Compared to the indices derived from the use of the other 14 possible hydrocarbons containing five fused rings, BaP is the most potent carcinogen. 1,2;7,8-dibenzanthracenes and 5,6- and 1,2-benzo-

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chrysenes are less carcinogenic; the other compounds, including benzo(e)pyrene (BeP) (Fig.1) are inactive.

B. BIOLOGICAL TRANSFORMATION OF BENZO(A)PYRENE edit

The intrinsic properties of a drug which allow it to interact with a given receptor site are not the only features which determine the intensity and duration of drug action. In order for a drug to exert its in vivo action it must be capable of transgressing a series of membrane barriers permitting its absorption, distribution as well as excretion. The latter is of major importance since rarely is it desirable that the drug action be permanent. Generally, drugs exist partly in non-polar form at physiological pH and thus are readily reabsorbed into the circulation through kidney tubules. Previous studies (56) have demonstrated that most drugs are metabolized in the liver to more polar compounds resulting in an increase in kidney excretion; the hepatic enzymes responsible are intimately associated with microsomes, subcellular particles derived from membranes of the endoplasmic reticulum; and, finally, many drugs stimulate this common enzyme system while others inhibit the system.

On the basis of considerable studies performed since the 1940’s, the metabolic fate of BaP, in some respects, is well-known. in vivo studies reported by Falk and others (57,58) investigated the biological transformation of 14C-BaP in mice and rats after subcutaneous administration, intravenous injection and intratracheal instillation.

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Maximal metabolic alteration occurred in the liver, with subsequent excretion through the hepatoliliary system and elimination through the feces. After characterizing the sulfate and glucuronide metabolic conjugates found in the bile, the authors concluded that the rat liver oxidized BaP to the following products: 3-monhydroxy BaP: 6-monohydroxy BaP: 1,6-dihydroxy BaP: and 3,6-dihydroxy BaP. These and other possible metabolites of BaP were tested by other investigators (59,60,61,62) and did not induce tumor formations regardless of whether metabolites were tested alone, in combination with each other or in combination with croton oil, a carcinogenic promoter substance. Therefore, as some concluded (57,63), the metabolism of BaP represents a detoxification process in the basic pharmacological sense and unaltered BaP apparently acts as the essential biologic stimulus. However, as discussed more fully in the next section, this conclusion may not be fully justified and, at least in part, the metabolism of BaP actually may represent a crucial process in the initiation of carcinogenesis.

The in vitro oxidation of BaP in rat liver homogenates was first studied by Conney and others (64). After incubating BaP, homogenate and buffer, these investigators isolated and quantitatively characterized the metabolic products as follows: 30% 3-monohydroxy BaP; 16% 1-monohydroxy BaP; 3% 3,6-dihydroxy BaP; 6% 3,6 BaP-quinone and 4% 1,3 BaP-quinone. The oxidation reactions were assayed

8

by measuring the fluorometric disappearance of extractible BaP from reaction mixtures. Boiled and trypain-treated homogenates contained no activity, whereas dialyzed homogenates retained full activity. Therefore, the oxidation of BaP was enzyme-mediated and did not depend on dialyzable cofactors.

Cell fractions obtained by differential centrifugation did not have much activity when assayed alone, but the combined microsomes and supernatant fractions had 80% of the activity of the original homogenate. Further experiments demonstrated that the microsomes contained the enzyme system and the supernatant fraction, functioned as a generator of NADPH. In addition, maximal activity resulted when homogenates were incubated under a saturated oxygen atmosphere. Therefore, the enzyme was an NADPH-dependent liver microsomal mixed-function oxidase. Since the enzyme derived from rat liver microsomes transforms not only BaP, but a number of other polycyclic hydrocarbons including the noncarcinogen, BeP (65,66), to phenolic derivations, Gelboin and others (67) have suggested the enzyme be called arylhydrocarbon hydroxylase (AHH).

Recently, AHH has been reportedly solubilized (68,69), and the mechanism of enzyme activity (Fig. 2) is now thought to include the transport of reducing equivalents from NADPH through NADPH-dependent cytochrome C reductase, a flavoprotein, to the heme protein, cytochrome P-450. Molecular oxygen then reacts with the reduced P-450 to produce an

9

active oxygenated complex which is apparently responsible for the oxidation of drug substrates.

When Conney et al, (64) pretreated rats with a single intraperitoneal injection of BaP 24 hours before sacrifice, they observed a 5-to 10-fold increase in AHH activity. Also effective in producing induction of AHH activity were other carcinogenic polycyclic hydrocarbons, such as 3-methylcholanthrene (MC) and 1,2:5,6-dibenzanthracene. Subsequently, however, Arcos (70) demonstrated that the induction phenomenon is not always related to polycyclic hydrocarbon carcinogenicity. For example, whereas the noncarcinogens 2’- and 3’-methyl 1,2-benzanthracene do enhance activity, the carcinogen 3,4-benzphenanthrene does not enhance activity. Nevertheless, maximally increasing activity by pretreating animals with inducer substances permits more accurate studies to be performed. This potential for an amplified expression of enzyme activity has been utilized by a number of investigators in their studies on the oxidation of BaP by microsomal enzyme systems.

More than 200 substances, including various drugs, insecticides and carcinogens, are known capable of stimulating the activities of liver microsomal drug metabolizing enzymes (71). It has been suggested by previous studies that these substances can be divided into two groups: broad spectrum inducers and narrow spectrum inducers (72). Phenobarbital (Pb), an example of the broad spectrum group, stimulates a large number of drug metabolizing enzymes.

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increases microsomal NADPH-cytochrome C reductase and cytochrome P-450 and causes a substantial proliferation of hepatocellular smooth endoplasmic reticulum (72,73,74,75). In contrast, BaP, representative of the narrow spectrum agents, induces only a limited number of microsomal enzymes. Although a hepatic microsomal cytochrome, cytochrome P-448, is increased, BaP, unlike Pb, does not induce a significant increase of either NADPH-cytochrome C reductase or hepatocellular smooth surfaced membranes (72,76,77).

Evidence suggests that the administration of inducers enhance enzyme activities by increasing the amounts of enzyme protein produced. For example, inducer pretreated rats show a greater liver protein weight (78,79,80) and ability to incorporate amino acids into microsomal proteins (81,82) than noninduced animal controls. Furthermore, simultaneous pretreatment of inducers and inhibitors of protein biosynthesis, such as puromycin (83), and inhibitors of DNA-dependent RNA biosynthesis, such as actinomycin D (84), prevents the induction response. Recently, increases in the template efficiency of liver chromatin has been shown to occur after the administration of MC to rats (85). The difference between control and MC systems is eliminated by extraction of the chromatin with 2M NaC1, a procedure which removes many of the nuclear proteins. These findings suggest that inducers may stimulate enzyme biosynthesis primarily at the level of gene transcription.

AHH activity has been found in numerous tissues in a

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variety of species (84,86). Besides homogenates derived from rat and human liver (87), enzyme activity occurs in homogenates from rat skin, kidney, lung, small intestine, adrenal gland, spleen and thymus (88,89). No detectable AHH activity was observed or could be induced in rat heart or brain tissue homogenates.

When human immature (9-12 weeks gestation) placental homogenates were assayed by Juchau et al. (90), no AHH activity could be detected. However, Welch et al. (91) found activity in human term placental homogenates. These investigators assayed activity by measuring the fluorometric appearance of hydroxylated BaP products. When placentas derived from 17 nonsmoking and 17 smoking mothers were assayed, Welch et al. Observed that mothers who smoked cigarettes had high activity, whereas nonsmokers had little or no activity. Similar results were reported by Nebert et al. (92) using placentas from 51 nonsmoking and 46 smoking mothers. Further studies by Welch’s group (93) revealed induced activity in placenta as well as maternal and fetal lung in rats exposed to cigarette smoke 5 hours daily for 3 days. When pregnant rats were pretreated orally 24 hours before sacrifice with any one of the polycyclic hydrocarbons known to be present in cigarette smoke (carcinogens and noncarcinogens), placental AHH activity was induced in every case. Apparently, the BaP (and/or other constituents) present in cigarette smoke induced the hydroxylase enzyme.

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In other studies, Bulay and Wattenberg (94) injected BaP or vehicle only subcutaneously in pregnant mice during the latter half of gestation. Progeny were delivered by ceasarean section, placed with foster mothers and, beginning at 4 weeks of age, treated topically with the carcinogenic promoter, croton oil, twice a week for 24 weeks. Increased tumor incidence and tumor count per animal in skin and lung were found in progeny of mothers pretreated with BaP compared to progeny of control mothers pretreated with vehicle alone. These findings suggest that exposure to BaP could have a carcinogenic effect on fetuses in utero.

Because exposure to carcinogenic substances, such as those present in cigarette smoke including BaP, alters the metabolism of BaP in organs important to fetal development and because exposure to BaP apparently can produce carcinogenic effects in utero, the fetal-placental unit (comprising maternal liver, placenta and fetal liver) is a worthy subject to study with regard to the problem of BaP-induced carcinogenesis. For these reasons and, in order to provide a more established basis for assessing the role of biological transformation in carcinogenic processes generally, one objective of the present research is to study the biological transformation of BaP in various tissues of the human and rat fetal-placental unit.

C. INTERACTIONS BETWEEN BENZO(A)PYRENE AND POLYNUCLEOTIDES edit

Much of the fundamental knowledge concerning the interaction between DNA and BaP has been provided by the detailed

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studies of Ts’o and his collaborators. In 1968, Ts’o et al. (99) reported physical binding between BaP and DNA and presented evidence to support the suggestion that this weak interaction involved the intercalation of the planar BaP molecule between the base stacks of nucleic acid. This binding was not carcinogen-specific, since binding also occurred, to the same extent (1 molecule/2000 nucleotide bases) and with similar affinity (as determined by binding constants), with the noncarcinogenic structural isomer, BeP. Further, BaP, physically bound to DNA, could be completely extracted with hexane, indicating that no chemical linkage was formed between BaP and DNA.

In later work Ts’o et al. (95) found that a covalent linkage, between BaP and DNA, resulted from reacting, at neutral pH in aqueous solution at room temperature, the physical complex with such oxidizing agents as iodine and hydrogen peroxide. Under comparable conditions, the noncarcinogen, BeP, was unable to interact covalently with DNA suggesting a positive correlation between such findings and carcinogenicity. Evidence suggests that BaP radical cations, generated in the iodine-induced reaction, interacts specifically with guanine residues on the DNA molecule. Recently, DNA, modified in this manner, has been shown to be mutagenic and to be less able to transform bacterial cells on the basis of studies with an in vitro B. subtilis system (96). In addition, template activity was reduced by nearly 80% in the in vitro transcription of the BaP-

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modified DNA by highly purified RNA polymerase derived from Micrococcus lysodeikicus. These findings demonstrate that modification of DNA, resulting from the oxidation of BaP, may be of considerable biological importance.

In 1964, Brookes and Lawley (20) showed, on the basis of in vivo studies, that polycyclic hydrocarbons, applied to the skin of mice, become covalently bound to DNA. More importantly, the degree of binding between the polycyclic hydrocarbons and the mouse skin DNA, but not RNA or protein, was reported to be directly correlated to their carcinogenic potency. Subsequent studies further supported these initial observations using mouse embryo cells in culture (22,97). It was found that for a number of polycyclic hydrocarbons tested, the binding index (defined as the extent of hydrocarbon binding to DNA in μmoles/mole P resulting from the metabolism of 1 mμmoles of hydrocarbon per ml. of medium) had values of less than one for noncarcinogens and in the range 15-150 for a series of known carcinogens.

Because covalent binding between polycyclic hydrocarbons and DNA requires the breaking and reforming of chemical bonds in both, the polycyclic hydrocarbon and DNA, a reasonable assumption is that the chemical change and, thereby, the in vivo interactions of the hydrocarbon with DNA, is enzymatic in nature. In 1969 Gelboin (24) explored this possibility with regard to AHH. He found that isotopically-labeled BaP, incubated in the presence of rat liver AHH and calf thymus DNA, binds covalently to the DNA. The

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binding required NADPH and was greater when the microsomal enzyme was derived from rats pretreated with the inducing substance, MC. The extent of binding with the induced system was of the order of 1 BaP covalently bound per 50,000 nucleotide bases. Later studies (98) revealed that microsomes derived from rat lung also are able to catalyze covalent binding between BaP and DNA. Furthermore, the DNA-binding product in such reactions may be a precursor of 3-mono-hydroxy BaP, the major product of microsomal conversion of BaP, since both DNA-binding and 3-monohydroxy BaP production are similarly suppressed by hematin, an inhibitor of microsomal enzymes.

Nevertheless, these findings suggest that the microsomal enzyme system responsible for converting BaP in vivo in a number of tissues in various species, including man, is capable of covalently binding BaP to DNA. Because the involvement of carcinogen metalolizing enzymes in chemical reactions with DNA is of fundamental importance to the mechanism of chemical carcinogenesis, further objectives of the present research are to study the interactions of biologically transformed products of BaP with DNA and other polynucleotides and, on the basis of studies with polynucleotides, enzymatically modified with BaP, attempt to evaluate the biological significance of such interactions.

16 - FIGURE 1. CHEMICAL STRUCTURES OF BENZO(A)PYRENE AND BENZO(E)PYRENE. edit

17 - CLICK HERE FOR PAGE-017-FIGURE-01

PAGE-017-FIGURE-01

18 - FIGURE 2. ELECTRON FLOW PATHWAY IN THE MICROSOMAL MIXED-FUNCTION OXIDATION SYSTEM. edit

“Fp” is a flavoprotein and P-450 is a cytochrome oxidase. In its reduced form, P-450 combines with carbon monoxide to produce a complex able to absorb light at 450 mμ.

19 - CLICK HERE FOR PAGE-019-FIGURE-02

PAGE-019-FIGURE-02

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MATERIALS edit

All rats used were purchased from Charles Rivers Breeding Laboratories, Wilmington, Massachusetts. Human tissues were obtained from the Obstetrics Department of the Children’s Hospital at Buffalo. E. Coli K-12 bacterial cells (grown to three-fourths log phase) were supplied by Grain Processing Corporation, Muscatine, Iowa. Calf thymus DNA (highly polymerized), yeast RNA, and bovine pancreas deoxyribonuclease I (electrophoretically pure) were obtained from Worthington Biochemical Corporation, Freehold, New Jersey. Synthetic ribopolynucleotides (Poly G, Poly A, Poly U, Poly C, Poly I, Poly (I·c), Poly I· Poly C) were procured form P-L Biochemicals, Milwaukee, Wisconsin. Tritium-labeled benzo(a)pyrene (3,4 – ³H) and adenosine ribonucleoside-5’-triphosphate (8-³H) were purchased from Schwartz-Mann, Orangeburg, New York. Benzo(a)pyrene (nonradioactive), 3-methylcholanthrene, 8-quinolinol and naphthaline (recrystallized) were obtained from Eastman Organic Chemicals, Rochester, New York; benzo(e)pyrene from Pfaltz and Bauer, Flushing, New York; and bovine serum albumin (crystallized) from Armour Pharmaceutical Company, Kankakee, Illinois. Deoxyriboncleoside-5’-monophosphates (dGMP, dAMP, dUMP, dCMP), ribonucleoside-5’-monophosphates (GMP, AMP, UMP, CMP), ribonucleoside-5’-triphosphates (GTP, ATP, UTP, GTP), dithiothreitol (Cleland’s reagent), reduced nicotinaminde adenine dinucleotide (NADPH), Bio-Gel A-5m (100-200 mesh) and Bio-Gel A-1.5 m 9100-200 mesh) were

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obtained from Calbiochem, San Diego, California. Reduced nicotinamide adenine dinucleotide (NADH), flavin adenine dinucleotide (PAD), riboflavin-5-phosphate (FMN), reduced glutathions (GSH), iodacetamide, p-chloromercuribenzoic acid (PCMB), estrone, androstenedione, testosterone, dehydroepiandrosterone, dehydroepiandrosterone sulfate (sodium salt), pregnenolone, cholesterol, cortisone and tris-(hydroxymethyl)aminomethane (Trizma Base) were all obtained from Sigma Chemical Company, St. Louis, Missouri; β-estradiol and progesterone from General Biochemicals, Chagrin Falls, Ohio; and ammonium sulfate (enzyme grade) from Nutritional Biochemicals, Cleveland, Ohio. 2,5-diphenyloxazole (PPO) and 1,4-di[2-(5-phenyloxazolyl)]-benzene (POPOP) were purchased from Amersham-Serle, Des Plaines, Illinois, dimethyl POPOP and Triton X-100 from Packard Instruments, Downers Grove, Illinois; Superbrite (type 100-5005) glass beads from 3M Company, St. Paul, Minnesota; Whatman microgranular DEAE-cellulose (DE-52) (1.0 meq per g, dry weight) from Reeve Angel Company, New York; and carbon monoxide, oxygen and nitrogen from the Matheson Company, East Rutherford, New Jersey. Other reagents and solvents were obtained from Fisher Scientific Company, Pittsburgh, Pennsylvania and J.T. Baker Chemicals, Phillipsburg, New Jersey.

Unless otherwise specified, equipment used included an Aminco-Bowman spectrophotofluorometer, Gilford Model 2400-S spectrophotometer, Packard Tri-Carb Model 3375

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liquid scintillation spectrometer, Nuclear Chicago Actigraph III paper strip counter, Beckman Model L-2 refrigerated ultracentrifuge with associated 40, 30 and SW 39 rotors, Servall Model RC2-B automatic superspeed refrigerated centrifuge with associated SS-34 and GSA rotors, Isco Model D density gradient fractionator equipped with an ultraviolet analyzer and teflon flow cell (2 mm optical path; 0.1 ml illuminated volume) designed for use with ½” X 2” ultra-centrifuge tubes, Isco Model 820 fraction collector, Radiometer Model 26 and Photovolt digital readout pH meters, Radiometer Type CDM 2d conductivity meter, Mettler Type H15 analytical balance, Heller Model GT-21 electronic laboratory stirrer, American optical Model 03156 isothermic water bath shaker, Burrel wrist-action shaker with 12 place capacity, Buchler sucrose density maker with associated peristaltic pump and other standard laboratory equipment.

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METHODS edit

PREPARATION OF HOMOGENATES IN FETAL-PLACENTAL UNIT STUDIES edit

Human placental tissues were obtained at term following normal vaginal deliveries of cesarean sections. Immature human placental and fetal tissues were obtained from healthy patients at operation (dilatation and curettage or hysterotomy for therapeutic abortion). Gestational ages were based on measurements of crown-rump length and foot length and from the last menstrual period. Tissues were immediately refrigerated at approximately 4°C. Fifteen-day pregnant rats (CD strain, 200-300 grams each) were injected intraperitoneally with either 20 mg/kg MC in corn oil (Mazola brand) or corn oil alone (injected volume was approximately 0.5 ml). Forty-eight hours later the rats were sacrificed by cervical dislocation and the placentas and maternal and fetal livers were dissected from the carcasses. (Preliminary experiments showed the dose of MC and time of sacrifice of animals to be optimal for stimulation of AHH activity in all tissue systems; i.e., higher doses did not significantly increase specific activities).

Tissues, including those of human and, unless otherwise stated, those pooled from two rats, were extensively washed with ice-cold 0.25 M sucrose (until essentially free of blood), blotted, weighed, diluted with two volumes ice-cold 0.25 M sucrose and minced with scissors. This and the following steps were performed essentially according to procedures by Juchau (90). Human placental tissues were homogenized in a Waring blender

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at high speed for 30 seconds. Other tissues were homogenized with a Potter homogenizer fitted with a teflon pestle (ten strokes). In some experiments requiring greater tissue disruption, rat placental tissue, after mincing, was forced through a tissue press, diluted with two volumes of 0.25 M sucrose, homogenized in a Virtis ‘45’ homogenizer (fitted with turboshear blades) at maximum speed for one minute, and then homogenized with a Potter homogenizer.

Placental homogenates were centrifuged at 1000 x g for 10 minutes; the resulting pellets were resuspended in the original volume of 0.25 M sucrose and frozen (approximately -20°C) until needed (within two weeks). Liver homogenates were centrifuged at 9000 x g for 20 minutes and the resulting supernatants were then frozen. It should be noted that hydroxylase actvities were not significantly affected when homogenates were frozen (with dry ice and methanol) and thawed 25 times within six hours or stored frozen for three weeks. Before assaying, rat placental homogenates were further diluted 1:1.5 with 0.25 M sucrose solution (human placental tissue homogenates were not further diluted). In the case of maternal liver homogenates, 0.25 M sucrose was added to further dilute the fraction 1:79 prior to incubation. All rat tissue homogenates were derived from MC pretreated animals unless noted otherwise.

In dialysis experiments, homogenates (10-15 ml) were dialyzed in an Oxford Laboratory Multiple dialyzer for 24 hours at 4°C with three equally spaced changes of 700 ml

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dialysate. In boiling experiments, thin-walled test tubes containing the homogenate subfractions were placed in boiling water for one minute and subsequently reconstituted to the original volume.

PREPARATION OF RAT LIVER MICROSOMES edit

Female rats (CD strain, 180-230 grams each) were injected intraperitoneally with either 20 mg/kg MC in corn oil or corn oil alone (approximately one ml injected volume). After 48 hours rats were sacrificed by cervical dislocation and the dissected livers (pooled from at least six animals) were immediately placed in ice-cold 0.25 M sucrose. Blood was essentially removed after tissues were extensively washed in ice-cold 0.25 M sucrose. Microsomes were prepared according to slight modifications of a previous method described by Gelboin (24). Diced livers were homogenized with a Potter homogenizer fitted with a teflon pestle for 10-13 strokes at 5000 rpm. Homogenates were centrifuged at 100 x g for 20 minutes; resulting supernatants at 12,000 x g for 20 minutes (twice); and, finally, at 105,000 x g for one hour. The 105,000 x g pellets (crude microsomes) were washed by suspension in the original volume of sucrose solution and recentrifuged at 105,000 x g for one hour (repeated twice). The final microsomal pellets were resuspended in 0.25 M sucrose equivalent to 1 gram wet weight tissue per 1.4 ml sucrose solution (approximately 5 mg microsomal protein per ml), divided into small test tubes containing approximately one ml each

26

and frozen at -70 to -100°C until needed (hydroxylase activities remained stable for at least four months). Microsomal preparations, except in fetal-placental unit studies, served as the hydroxylase enzyme source in all experiments involving BaP biotransformation and, unless otherwise noted, were derived from MC pretreated animals.

ENYZME ASSAYS IN FETAL-PLACENTAL UNIT STUDIES edit

The rate of BaP hydroxylation in tissue homogenates was determined by measuring the appearance of the fluorescent hydroxylated products according to slight modification of a previous method (88). Typical reaction mixtures consisted of 0.5 ml tissue homogenate as the enzyme source, 0.1 ml BaP in acetone (0.5 g/ml) as substrate, 0.2 ml NADPH (2 mg/ml for assays of rat maternal liver homogenates and 10 mg/ml for all other tissue homogenates) as the electron donor and 0.7 ml 0.1 M phosphate buffer at pH 7.35. Phosphate buffer alone or with other additions (as noted) were added to give a final volume of 2.0 ml. This mixture was incubated with shaking in a Dubnoff metabolic incubator (50-60 rpm) under saturated oxygen, nitrogen or carbon monoxide gas phases at 37°C for 15 minutes. Triplicate flasks were used in each experiment. Appropriate tissue and substrate blanks were employed. Under these conditions enzyme activity was of first-order kinetics with respect to enzyme concentration and incubation time and of zero-order kinetics with respect to substrate and NADPH concentrations.

The room was darkened for the reminder of the assay.

27

After incubation, 2.0 ml of ice-cold acetone were added to reaction vessels to terminate the reaction. 8.0 ml of hexane were then added to the mixture. The flasks were stoppered and incubated at 37°C with shaking for 10 minutes. A 5.0 ml aliquot of the hexane layer was then added to 8.0 ml of 1.0 N NaOH in separate 50 ml centrifuge tubes. The tubes were stoppered, shaken for five minutes and centrifuged for one minute. A 4.5 ml sample of the NaOH layer transferred to fluorometric cuvettes. The amount of extracted products were measured as fluorescence units in a Model III Turner fluorometer (sensitivity 30) fitted with a primary activating filter (405 mμ) and a secondary narrow pass filter (525 mμ). Pure 1.0 N NaOH was utilized as a blank. One fluorescent unit of extracted hydroxylated BaP in 1.0 N NaOH was equivalent to 100 ng/ml quinine sulfate in 0.1 N sulfuric acid.

NADPH oxidase activity was determined by following the disappearance of NADPH at 340 mμ on a Beckman DU spectrophotometer equipped with a Gilford optical density converter and recorder. The reaction mixtures contained 0.2 ml of 33% homogenate suspensions (approximately 4.7 mg and 2.8 mg of liver and placental protein respectively), 0.3 ml NADPH (1 mg/ml) and 2.5 ml of 0.1 M phosphate buffer at pH 7.35. NADPH, homogenate and NADPH/homogenate blanks were run concurrently, and NADPH oxidase activity was calculated by determination of concentrations of NADPH in reaction cuvettes from a standard curve.

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ASSAY FOR ARYL HYDROCARBON HYDROXYLASE edit

Except for fetal-placental unit studies, the following assay for the appearance of fluorescent hydroxylated BaP products was used in the present research. The reaction and incubation conditions were similar to those used by Gelboin (24). Typical reaction mixtures consisted of approximately 0.1 ml rat liver microsomal preparation (0.5 mg microsomal protein) as the enzyme source, 0.2 ml NADPH (10 mg/ml in 0.25 M sucrose) as the electron donor, 0.6 ml 0.25 M sucrose (adjusted in relation to the exact volume of microsomal suspension added) and 2.0 ml of solution A containing 100 μmoles EDTA (to inhibit nuclease activity) and 50 μmoles sodium phosphate buffer at pH 7.40. In some experiments microsomes were heated in a boiling water bath before adding to reaction mixtures. Unless otherwise noted, only microsomes derived from MC pretreated animals were used.

Various polynucleotides and mononucleotides were added to incubation mixtures after dissolution in solution A (usually 1.0 mg/ml; DNA solutions contained 0.75 mg/ml). In the case of Poly I, dissolution in solution A required a basic pH (pH 9.00) which was later adjusted to pH 7.40 with phosphoric acid before addition to mixtures. To assure the double-stranded structure, Poly I·Poly C was dissolved in solution A containing 1.27% NaCl (NaCl solutions, but without Poly I·Poly C, served as controls in appropriate studies). In studies requiring denatured DNA, native DNA dissolved in solution A was heated for

29

15 minutes in a boiling water bath followed by immediate chilling in an ice-water bath (denaturation of DNA was verified by hyperchromicity studies).

Mixtures, with and without polynucleotide additions, were preincubated at 37°C with shaking (50-60 rpm) for one minute after which 0.1 ml BaP in acetone (0.8 mg/ml) as substrate was added to initiate the enzyme reactions. In some experiments BaP was replaced by an equivalent amount of BeP; in other experiments 0.1 ml of fluorescent BaP products in a concentrated hexane solution substituted (see below). After incubation for 14.0 minutes, reactions were terminated by adding one volume (3 ml) ice-cold acetone to mixtures and immediately chilling in ice for at least five minutes. Under the above conditions, enzyme activities were of first-order kinetics with respect to enzyme concentrations and of zero-order kinetics with respect to substrate and NADPH concentrations.

Terminated reaction mixtures were transferred to low actinic light test tubes and extracted with 9.0 ml hexane with shaking for 20 minutes on a Burrell wrist-action shaker adjusted for maximum speed. A 5.0 ml aliquot of the hexane layer was next extracted with 5.0 ml 1.0 N NaOH in separate test tubes for one minute. Exactly 10 minutes after addition of hexane aliquots to NaOH solutions, extracted fluorescence present in the NaOH layer was measured on an Aminco-Bowman spectrophotofluorometer (initially adjusted with a standard quinine sulfate solution, 3000 ng/ml in

30

0.1 N H₂SO₄, to read 880 units at 340 mμ - excitation wavelength and 445 mμ - emission wavelength) at 390 mμ - excitation maxima and 515 mμ - emission maxima (under these maxima, the quinine sulfate solution read 34 units). Under similar fluorometric conditions, Kuntzman (87) found that a solution of 3-hydroxy BaP (10 ng/ml), the major product of BaP biotransformation, read 260 units (readings were linear up to at least 100 ng/ml concentration). In the present research, some experiments studied fluorospectal characteristics of extracted BaP and BeP products by usual wavelength scanning procedures.

PREPARATION OF FLUORESCENT BaP HYDROXYLATED PRODUCTS edit

Some studies involving various polynucleotides required the inclusion of fluorescent BaP products, substituted for the parent BaP, in reaction mixtures used in the AHH assay. A solution of such products was prepared enzymatically according to methods described earlier with regard to the AHH assay. A reaction mixture containing 2.0 ml microsomal suspension, 1.0 ml BaP in acetone (0.8 mg), 2.0 ml solution A (0.05 M EDTA and 0.025 M sodium phosphate buffer at pH 7.40) was incubated for 15 minutes at 37°C. After terminating the reaction with 30 ml ice-cold acetone, the mixture was extracted with 90 ml hexane by shaking in a 250 ml separatory funnel for 20 minutes. Immediately following extraction of the hexane layer with 45 ml 1.0 N NaOH for one minute, the NaOH layer (containing hydroxylated BaP products) was acidified to pH 5.00 with 3 N HCl and

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extracted with 45 ml hexane for 10 minutes. The resulting hexane extract, evaporated sub vacuo to one ml, served as the source of fluorescent BaP hydroxylated products in appropriate studies (approximately 54,000 fluorescent units/ml).

SUCROSE GRADIENT SEDIMENTATION STUDIES edit

Reaction mixtures contained 0.2 ml microsomal suspension, 0.2 ml NADPH (2 mg), 0.5 ml 0.25 M sucrose, and 2.0 ml solution A (0.05 M EDTA and 0.025 M sodium phosphate buffer at PH 7.40) containing 2 mg of the polynucleotide under study. After preincubation for one minute with shaking (50-60 rpm) at 37°C, 0.1 ml of ³H-BaP in acetone (0.8 mg/ml; 5.4 mC/ml) was added and mixtures further incubated for 14 minutes. Reactions were terminated by chilling in ice for five minutes. Mixtures were then centrifuged at 105,000 x g for one hour and the resulting supernatants extracted with three volumes of hexane for 20 minutes. Aliquots (0.075 or 0.050 ml) of the aqueous layer were carefully layered on the top of 5-20% sucrose gradients (5.0 ml) prepared in ½” x 2” cellulose nitrate centrifuge tubes by combining a 5% sucrose solution (containing 0.15 M NaCl and 0.01 M Tris-HCl buffer at pH 7.50) with a similar solution containing 20% sucrose in a conical sucrose density maker. Gradients were centrifuged for various times (12, 18 and 24 hours) in an SW 39 ultracentrifuge rotor at 35,000 rpm (completed runs were decelerated freely). In an Isco Model D fractionator, centrifuged gradients were gently

32

forced (by means of a 30% sucrose solution cushion entering at the bottom of pierced gradient tubes) through an ultraviolet analyzer unit set at 254 mμ at a rate of 0.5 ml per minute. UV readings were chart recorded and fractions (approximately 0.20 ml each) of the gradients were collected directly in scintillation vials. After water (0.8 ml) was added to vials, 10.0 ml of a scintillation fluid (containing O.1 g POPOP, 5·5 g PPO and 333 ml Triton x-100 per liter toluene solution) was added and the final solutions mixed well. Fractions were then counted in a refrigerated liquid scintillation counter adjusted for tritium counting (approximately 25% counting efficiency).

PREPARATION OF DNA MODIFIED WITH NITROGEN MUSTARD edit

The preparation of DNA modified with nitrogen mustard was based on the method of Chmielewicz et al. (100). Highly polymerized calf thymus DNA (0.905 mg/ml) in 0.10 M sodium phosphate buffer at pH 7.00 was incubated with shaking (50-60 rpm) in the presence of nitrogen mustard (HCl salt) (6 x 10-4 M) at 37°C. After incubation for 18 hours, DNA was precipitated with 2 volumes cold pure ethanol, spooled onto a glass rod, washed extensively in 70% ethanol, 90% ethanol and pure ethanol, air dried and finally dissolved in solution A (0.050 M EDTA and 0.025 M sodium phosphate buffer at pH 7.40) with slow stirring at 4°C for 48 hours. Under these conditions template activity of modified DNA in a bacterial DNA-dependent RNA polymerase system was nearly abolished (100). Several unmodified DNA controls

33

were required in studies involving nitrogen mustard modified DNA. These DNA controls were prepared at the same time as nitrogen mustard-modified DNA but differed from the modification procedure in one of three ways. DNA was incubated for 18 hours but in the absence of nitrogen mustard; DNA was prepared initially in the presence of nitrogen mustard but was not incubated; and DNA was prepared initially in the absence of nitrogen mustard and was not incubated.

PREPARATION OF DNA FOR TEMPLATE STUDIES edit

DNAs treated with either BaP, BeP, or vehicle alone in microsomal incubation mixtures were prepared for use in template studies employing an E. coli DNA-dependent RNA polymerase system. Three reaction mixtures, each containing 0.3 ml rat liver microsomal preparation (1.5 mg microsomal protein), 0.6 ml NADPH (10 mg/ml in 0.25 M sucrose), 1.8 ml 0.25 M sucrose, and 6.0 ml of solution B (5.4 mg calf thymus DNA in 0.050 M EDTA and 0.025 M sodium phosphate buffer at pH 7.40), were preincubated with shaking (50-60 rpm) at 37°C for one minute. Either 0.3 ml BaP in acetone (0.8 mg/ml), 0.3 ml BeP in acetone (0.8 mg/ml) or 0.3 ml acetone alone was then added and mixtures further incubated for 14 minutes. After incubation, mixtures were chilled in ice five minutes and then centrifuged at 104,000 x g for one hour (4°C). Supernatants were extracted with 27 ml hexane for 10 minutes. After centrifugation at 10,000 rpm in a Sorvall SS-34 rotor for five minutes, hexane was separated from the

34

aqueous layers by aspiration and, finally, by evaporation under vacuum at 30°C. The extracted DNA solutions were then dialyzed in Bio-Rad hollow fiber mini-beaker dialyzers against solution A (0.050 M EDTA and 0.025 M sodium phosphate buffer at pH 7.40) at a flow rate of 40 ml/hour for 15 hours at 4°C. This step removed NADPH from DNA solutions as indicated by spectrophotometric measurements at 340 mμ. By applying a vacuum (580 mm Hg) to the mini-beaker dialyzers, DNA solutions were concentrated to 6 ml. These solutions, essentially the same as solution B, were reintroduced into incubation mixtures as described above and the procedure up to this point repeated. After one further repetition, DNA solutions were dialyzed against 0.10 M Tris-HCl buffer at pH 7.50 in mini-beaker dialyzers (40 ml/hour for 15 hours) and concentrated to 5 ml.

DNA solutions were then treated with one volume of a freshly prepared phenol solution (made by mixing 317 ml distilled phenol with 37.5 ml distilled m-cresol and 0.25 g 8-quinolinol, saturating with 0.1 M Tris-HCl buffer at pH 7.50 and adjusting the pH to 7.50 with 1 N NaOH). After gently shaking for three minutes, the phenol phases were separated from the DNA solutions by centrifuging at 4300 x g for 10 minutes at room temperature. Without disturbing the deproteinized interfaces, clear supernatants were collected and the phenol extraction procedure repeated twice more. (Phenol solutions wee back titrated with 1 ml 0.10 M Tris-NCl buffer at pH 7.50 and the Tris solutions

35

combined with the respective DNA solutions before each further extraction.) The phenol extracted solutions were then extensively extracted with diethyl ether. After reserving residual ether under vacuum, solutions were chemically analyzed for DNA content. (In all cases, approximately 55% of the original DNA was recovered.) These solutions served as sources for DNA in template studies using a DNA-dependent RNA polymerase system.

ASSAY FOR DNA TEMPLATE ACTIVITY edit

The ability of various DNAs to serve as templates in a DNA-dependent RNA polymerase system was determined essentially according to an assay procedure reported by Burgess (101). Reaction mixtures (0.5 ml final volume) contained 0.13 ml solution N (freshly prepared with 3.84 ml 1.0 M Tris-HCl buffer at pH 7.9, 48.0 mg bovine serum albumin, 1.075 g KCl, 0.384 ml 0.1 M potassium phosphate buffer at pH 7.5, 0.096 ml 0.1 M EDTA at pH 7.0, 0.096 ml 0.1 M dithiothreitol, 0.96 ml 1.0 M MgCl₂ and enough deionized distilled water to produce 25.0 ml final volume); 0.075 μmoles each of UTP, CTP, GTP and ³H-ATP (1.0 μC); 0.025 ml RNA polymerase preparation (260 μg protein); and various amounts of DNA (0-55 μg) in 0.1 M Tris-HCl buffer at pH 7.5. Assays were incubated 30 minutes at 37°C with shaking (50-60 rpm), chilled in ice and precipitated with 0.1 ml 50% trichloroacetic acid (TCA). After 15 minutes, TCA-insoluble products were collected on Millipore filters (no. HAWP-0220; 0.45 μ pore size) mounted in a Tracerlab Precipitation Apparatus (type E8B)

36

and washed 10 times with 1 ml of 5% TCA containing 0.01 M sodium pyrophosphate. Filters were dissolved in 10.0 ml of a scintillation fluid (prepared by dissolving 300 g naphthalene, 21 g PPO and 0.9 g dimethyl POPOP in 3 liters p-diozane) and the amount of radioactivity determined in a liquid scintillation counter. Under these conditions, enzyme activity, represented by the amount of precipitated radioactivity, was of zero-order kinetics with respect to enzyme concentration, substrate concentrations (nucleoside triphosphates) and incubation time.

The enzyme assay used during the preparation of RNA polymerase (see below) was essentially the same as the assay described above except that excessive DNA (100 μg) was included in the reaction, and mixtures were incubated for 10 minutes. Under these conditions, one activity (Burgess) unit of enzyme incorporates ne mμmole AMP into a TCA-insoluble product.

PREPARATION OF RNA POLYMERASE edit

E. Coli DNA-dependent RNA polymerase was prepared essentially according to the method of Burgess (101). Unless noted otherwise, all steps were preformed in the cold (4°C). 200 g of frozen E. coli K-12 bacterial cells, broken into small places with a mallet, were placed in a 1-liter Waring Blendor. Prechilled Superbrite 100 glass beads (500 g; prepared by soaking in concentrated HCl overnight, 0.5 N NaOH, and drying overnight in shallow dishes at 120°C) and buffer G (200ml; containing 0.05 M Tris-HCl buffer at pH 7.5, 0.01 M MgCl₂,

37

0.20 M KCl, 0.10 mM EDTA at pH 7.0, 0.10 mM dithiothreitol and 5% glycerol, v/v) were added and the cells homogenized at low speed for five minutes and at high speed for ten minutes. The extract was poured into a 1-liter beaker and chilled in a dry ice-isopropanol bath. 2 ml of a freshly prepared solution of deoxyribonuclease I in Buffer G (1.0 mg/ml) was added with mixing, and the beads were allowed to settle for 30 minutes. The supernatant was decanted into a 600 ml beaker. Beads were poured into a large funnel loosely plugged with glass wool and residual supernatant gently drawn off under vacuum. Beads were rinsed with 100 ml Buffer G and the rinsing drawn through by gentle suction. The filtrates were combined with the supernatant (Fraction 1; 400 ml).

Fraction 1 was centrifuged at 30,000 rpm for two hours in a Spinco No. 30 rotor. The resulting clear amber supernatant was Fraction 2 (305 ml). 70.0 g solid ammonium sulfate (AS) and 0.35 ml 1 N NaOH were added to Fraction 2 with stirring and the solution further stirred for 30 minutes. After centrifugation at 10,500 rpm in a Sorvall SS-34 rotor for 30 minutes, 30.0 g solid AS was added with stirring to the supernatant and the solution further stirred for 30 minutes. The pellet resulting from centrifugation at 10,500 rpm in a Sorvall SS-34 rotor for 30 minutes was resuspended in 260 ml 42% AS in Buffer A (containing 0.01 M Tris-MCI at pH 7.9; 0.01 M MgCl₂; 0.01 mM EDTA at pH 7.0; 0.10 mM dithiothreitol;

38

and 5% glycerol, v/v) and stirred 45 minutes. After centrifugation at 10,500 rpm in a Sorvall SS-34 rotor for 60 minutes, the resulting pellet was dissolved in 100 ml Buffer A and further diluted with Buffer A until the specific conductivity of the solution was equivalent to that of a solution of Buffer A containing 0.13 M KCl (6.6 x 1023 umhos/cm). The resulting solution (approximately 300 ml) was Fraction 3.

Fraction 3 was applied to a DEAE-cellulose column recently equilibrated with Buffer A. (This chromatographic column, 3 cm diameter, was prepared from Whatman microgranular DE-52 such that a 100 ml bed volume resulted. Before pouring the column, DE-52 was treated with 5 volumes 0.5 N HCl, rinsed with water to pH 4.0, treated with 5 volumes 0.5 N NaOH, rinsed to pH 8.0, and, finally, resuspended in 2 volumes 0.05 M Tris-HCl buffer at pH 7.9.) Fraction 3 was washed into the column with 50 ml Buffer A and then with 400 ml Buffer A containing 0.13 M KCl. After elution with 300 ml Buffer C (0.25 M Tris-HCl at pH 7.9; 0.10 mM EDTA at pH 7.0; 0.10 mM dithiothreitol; and 5% glycerol, v/v) containing 0.23 M KCl, fractions (10 ml each) containing peak polymerase activities were pooled. (Fraction 4; see above for polymerase assay procedure).

Protein in Fraction 4 was precipitated with 1.5 volumes saturated AS solution. The pellet resulting from centrifugation at 10,500 rpm in a Sorvall SS-34 rotor for 60 minutes was dissolved in a minimum volume of Buffer C

39

(approximately 5 ml) and dialyzed for two hours for each of two changes of the dialysate (100 volumes Buffer C). The dialyzed enzyme solution was diluted to 40 mg protein/ml with Buffer C and applied (3.2 ml aliquots per separation) to a Bio-Gel A-5m column recently equilibrated with Buffer C. (This column, 250 ml bed volume, was prepared by packing a slurry containing Bio-Gel A-5m beads, extensively washed with Buffer C, in a 2.5 cm diameter chromatographic column.) After elution with Buffer C at a flow rate of 50 ml/hour, fractions (4.2 ml each) containing peak polymerase activities were pooled (Fraction 5).

Fraction 5, like Fraction 4, was precipitated with 1.5 volumes of saturated AS and centrifuged at 10,500 rpm for 60 minutes. The resulting pellet was dissolved in a minimum amount of buffer A and dialyzed for two hours for each of two changes of the dialysate (100 volumes Buffer A). The dialyzed enzyme solution was diluted with Buffer A to a protein concentration of 16 mg/ml and applied (4.0 l aliquots per separation) to a Bio-Gel A-1.5 m column recently equilibrated with Buffer A containing 1.0 M KCl. (This column, 250 ml bed volume, was prepared by packing a slurry containing Bio-Gel A-1.5m beads, extensively washed with Buffer A containing 1.0 M KCl, in a 2.5 cm diameter chromatographic column.) After elution with Buffer A containing 1.0 M KCl at a flow rate of 40 ml/hour, fractions (3.3 ml each) containing a constant specific polymerase activities were pooled (Fraction 6).

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Fraction 6 was precipitated with 1.5 volumes saturated AS solution. The pellet, after centrifugation at 10,500 rpm in a Sorvall SS-34 rotor for 60 minutes, was dissolved in 6 ml Buffer E (0.01 M Tris-HCl at pH 7.9; 0.01 M MgCl₂; 0.10 M KCl; 0.10 mM EDTA at Ph 7.0; 0.10 mM dithiothreitol; and 50% glycerol, v/v) resulting in a solution with a 280 mμ/260 mμ ratio of 1.76 and containing 10.4 mg protein/ml. This final enzyme solution (450 Burgess units per mg protein) was stored at -20°C and served as the source of highly purified DNA-dependent RNA polymerase used in template studies (within five weeks).

DETERMINATION OF DNA edit

DNA was analyzed according to the diphenylamine method of Burton (102). Diphenylamine reagent was prepared by dissolving 1.5 g recrystallized diphenylamine in 100 ml glacial acetic acid; 1.5 ml of concentrated H₂SO₄ was added and immediately before use, 0.50 ml cold acetaldehyde (16 mg/ml) was added. The assay mixture contained 0.5 ml sample (30-100 μg DNA) in 0.10 M Tris-HCl buffer at pH 7.5, 0.5 ml 0.005 M NaOH and 1.0 ml 1 N HCl04. After heating at 70°C for 15 minutes, 4 ml diphenylamine reagent was added and the mixture left undisturbed for 16 hours at 30°C in the dark. Absorbancies were read against an appropriate blank at 600 mμ. DNA content was calculated from a standard curve using calf thymus DNA as a standard. The standard curve was linear throughout the range studied.

DETERMINATION OF RNA edit

RNA was analyzed according to the

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orcinol method of Fleck et al (103); Orcinol reagent was prepared by dissolving 100 mg recrystallized orcinol in 10 ml of 0.1% FeCl₃ in concentrated HCl. The assay mixture contained 0.5 ml sample (100-200 μg RNA) in 0.1 M Tris-NCl buffer at pH 7.5, 0.5 ml 10% TCA and 1.0 ml orcinol reagent. The mixture was heated for one hour in a boiling water bath. After cooling and adding 2 ml water, absorbancies were read against an appropriate blank at 660 mμ. Yeast RNA served as a standard. The standard curve was linear throughout the range studied.

DETERMINATION OF PROTEIN edit

Protein was analyzed according to the method of Lowry et al. (104). Solution D was prepared by mixing 0.5 ml 0.5% CuSO₄ and 0.5 ml 1.0% Na-K tartrate per 50 ml 2.0% Na₂CO₃ in 0.1 N NaOH. The assay mixture contained 1.0 ml sample (30-130 μg protein) and 5.0 ml solution D. After standing 10 minutes at room temperature, 0.5 ml 1 N phenol was added. After development for 30 minutes, absorbancies were read against an appropriate blank at 750 mμ. Bovine serum albumin was used as a standard. The standard curve was linear throughout the range studied.

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RESULTS edit

BIOTRANSFORMATION OF BaP IN VARIOUS TISSUES OF THE HUMAN AND RAT FETAL-PLACENTAL UNIT edit

For reasons discussed in the Introduction, one objective of the present research was to study the biological transformation of BaP in various tissues of the human and rat fetal-placental unit. A comparison of characteristics of BaP biotransformation processes in the various tissues therefore was attempted and was based on known features of the aryl hydrocarbon hydroxylase system including inducibility, subcellular localization, requirements for optimal activity and the effect of various agents.

Initial studies revealed no detectible AHH activity in boiled human and rat fetal liver and placental homogenates and rat maternal liver homogenates. No difference in specific activities were observed between dialyzed and non-dialyzed homogenates. Furthermore, EDTA, a divalent cation chelating agent (at a final concentration of 10^-3 M) had no effect on placental hydroxylase activity. However, an equivalent concentration of EDTA increased the activity in the rat maternal liver system by 20% over control values.

As previously established in the rat (64) and human (87), hydroxylase activity in liver homogenates was localized in microsomal subfractions. However, as shown in Figure 3, assignment of activity to a single subfraction of placental homogenates was more difficult. Each fraction, except the 500 x g pellet, was obtained by centrifuging the supernatant of the previous fractions. For example, centrifuging

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the 500 x g (10 minutes) supernatant at 1000 x g for 10 minutes produces the 1000 x g pellet. The 1000 x g supernatant was centrifuged at 5000 x g for 10 minutes, the 5,000 x g supernatant at 10,000 x g for 30 minutes, and the 10,000 x g supernatant at 104,000 x g for 1 hour. On a mg protein basis, placental activity did not appear to be localized in any one subfraction although later studies, using procedures to insure greater tissue disruption (see Methods), demonstrated greatest activity in microsomal subfractions. Nevertheless, in all such studies the bulk of activity on a gm wet weight tissue basis was localized in the 500 x g pellet. Insufficient disruption of essential cellular components apparently accounted for the relativity high activity observed in this subfraction. This is hardly surprising when one considers the difficulty in homogenizing highly vascular tissues by usual techniques.

Table 1 compares AHH activity in the various tissue homogenates studied. In humans, only homogenates of term placentas from cigarette smokers contained significant enzymatic activity. In contrast, activities in immature human placental homogenates were barely detectible regardless of maternal smoking history. Similarly, placental homogenates from 17 day pregnant rats exhibited substantial activity only when animals were induced by pretreating with MC prior to sacrifice. Separate experiments with rats indicated that the optimal activity in pretreated rat placentas could be observed on days 17-19

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and that activities at term or at 15 days were relatively less (approximately 40% less at 15 days and 20% less at term). Determinations on rat fetal or placental tissues from earlier than 15 days gestation were not made due to insufficient tissue.

Only minimal activity was detected in human and rat fetal livers and livers from several human neonates succumbing shortly after birth. Although rat fetal liver activity was increased when maternal animals were pretreated with MC, no similar induction in activity was observed in human fetal livers derived from smoking mothers. Based on results using human placentas, this may be due, at least in part, to the immature status of the human fetuses studied. (It should be noted that Juchau (113), using longer incubation times and higher reactant concentrations than those used in the present assay procedure, recently reported results establishing the presence of the AHH system in human fetal liver homogenates.) Since appropriate human neonates were limited for study, determinations of neonate liver activity from smoking and nonsmoking mothers could not be made.

Even under optimal conditions of smoking history or inducer pretreatment, human and rat fetal liver and placental homogenate activities were substantially less than the activity obtained from noninduced rat maternal liver. Activity in maternal liver was increased over 30-fold when animals were pretreated with MC. It should be noted that

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the maternal liver system appeared to be nearly three times more sensitive to MC induction than the placental enzyme system when an optimal dose of MC was employed.

Table 2 compares apparent kinetic constants among the various tissue homogenates studied. Subfractions of human and rat placental and rat maternal liver homogenates were compared with respect to apparent km and Vmax values for BaP as well as NADPH. Although the data should be interpreted with cautions, since they were obtained with crude enzyme preparations, the results may provide important clues to the reasons for differences in enzymatic activity among the various tissue homogenates.

Rat maternal livers exhibited the lowest km and highest Vmax values with regard to both BaP and NADPH. In placental preparations, the human system had higher Vmax values than the rat system suggesting that the human enzymes are able to transform larger amounts of BaP in the presence of very high NADPH and/or BaP concentrations. Concentrations of NADPH in vivo, however, might be expected to limit the actual turnover rate.

Figure 4 typifies the reciprocal activity vs reciprocal concentration plots used to obtain kinetic parameters. In this instance the liver enzyme system appeared to have over 10 times the affinity for NADPH than the placental system. The difference in Km values may be greater if one takes into consideration the NADPH oxidizing capacity in placental homogenates (practically negligible) and liver homogenates

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(considerable) (Figure 5).

All systems required NADPH and oxygen for maximal activity. NADH at 10^-3 M (final concentration) could substitute for NADPH as an electron donor in placental and liver systems but was only about one-fifth as active (Figure 6). Also similar were the effects of incubating maternal liver and placental homogenates under various saturated gas phases (Figure 7). Maximal activity was observed under oxygen.

Saturation of the gas phase with CO produced a 70% decrease in homogenate activities. At least part of this effect was due to anaerobiasis, since activity was 30% less in 100% nitrogen than in 100% oxygen. The actual inhibitory effect of CO on the systems (approximately 40% after correction for lack of oxygen) could be explained by inhibition of essential CO-sensitive enzyme components. It should be pointed out that cytochrome P-450 is an established CO-sensitive component of liver AHH preparations (56) and, more recently, has been identified in placental homogenates (105).

The 25-30% inhibition under nitrogen might suggest to some that H₂0 rather than O₂ is the source of the OH group for BaP hydroxylation or that the nitrogen gas was contaminated with O₂. Bubbling of nitrogen gas through a concentrated dithionite solution (1.4 M) and through solid KOH crystals, however, did not alter the activity of placental nitroreductase as assayed by previously described

47

methods (115). (The reductase system is very sensitive to oxygen.) Also, if H₂O were the source of the OH group, no inhibition would be expected under a nitrogen atmosphere.

The optimal pH observed for liver and placental enzymes was approximately 7.6. The pH range studied was between 5.0 and 8.5 and was adjusted by varying the ratio of monobasic to dibasic potassium phosphate in the 0.1 M buffer system. Reactions were run at 37°C for 15 minutes with the usual substrate and cofactor concentrations.

Figure 8 represents the effects of FMN and FAD on maternal liver and placental homogenate activities. With maternal liver homogenate FMN (10^-3 M final concentration) inhibited by 20%, whereas equimolar concentrations of FAD nearly abolished activity. However, either flavin at 10^-3 M inhibited the placental activity to the same extent (approximately 60%). These results suggest that a difference in electron transport characteristics may exist between the liver and placental systems. It should be noted that levels of activity in maternal liver homogenates were comparatively high in these experiments, but this was not surprising in view of the wide variation in hydroxylase activity observed in maternal livers of MC-pretreated animals. Comparisons of several pools (2 livers/pool) of maternal liver homogenates from MC-pretreated animals indicated that specific activities varied from as low as 1220 units/mg protein/hr to as high as 3670 units/mg protein/hr.

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The addition of NaCN (10^-3 M final concentration) to incubation mixtures had no significant effect on liver or placental activities, suggesting that electron transport in either system did not depend on cytochrome oxidase. Reduced glutathione at 10^-3 M (final concentration) had no discernible effect on enzymic activity in placental homogenates.

Figure 9 shows the effects of PCMB addition to incubation mixtures. PCMB at 10^-3 M (final concentration) inhibited both systems almost completely. However, at 10^-5 M the placental system was less sensitive to PCMB inhibition than the liver system. At first it was assumed that PCMB acted as a sulfhydryl inhibitor in these preparations. However, iodoacetamide (perhaps a more specific sulfhydryl inhibitor than PCMB) at 10^-5 M or 10^-3 M had no observable effect on the hydroxylation reaction in either system.

Increasing evidence suggests that certain hepatic mixed-function oxidases able to enzymatically alter drug substrates are involved in the biotranformation of endogenous steroids (106,107). In the human placenta, mixed-function oxidases are known to be involved in the side chain oxidation of cholesterol to pregnenolone and progesterone and in the aromatization of androgens to estrogens (108). The rat placenta, however, does not catalyze synthesis of estrogens or progesterone (109). For these reasons and in order to further compare liver

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and placental systems, a series of steroids were studied for their effects on the biotransformation of BaP in homogenate preparations. The results of these studies are shown in Table 3.

In all tissues studied, estrogens and progesterone were the most effective inhibitors. With regard to placental homogenates, the rat system was more sensitive to estrogens than the human system whereas the human system appeared to be the more sensitive to inhibition by androgens. Dehydroepiandosterone sulfate and pregnenolone both produced slight activation in liver homogenate activity but did not alter activity in placental homogenates. This experiment was repeated twice (in triplicate) with similar results. None of the systems were appreciably affected by 10^-3 M cholesterol or pregnenolone or 10^-5 M concentrations of progesterone or androgens. In general, difference between the various systems were more quantitative than qualitative and were not great. Moreover, tissue AHH appeared to be unrelated to mixed-function oxidases utilizing cholesterol or androgens as substrates. (It should be noted that the aromatization reaction is inhibited by cyanide but not by carbon monoxide (108), characteristics opposite to those of the AHH system.)

Although differences were observed between the various systems studied (apparent specific activities and certain kinetic parameters, effect of EDTA and flavins), the similarities appeared to be more striking. All systems

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were inducible mixed-function oxidases requiring NADPH and oxygen (but not dialyzable cofactors) for maximal activity, had similar pH optima and, on the basis of preliminary studies, appeared to be localized in microsomal subfractions. Furthermore, the effects of various inhibitors (carbon monoxide, nitrogen, PCMB, iodoacetamide and steroids) were similar. Other investigators have reported similar characteristics (notably those related to cofactor requirements and subcellular localization) with regard to AHH systems found in human adult liver tissue (87), neonate foreskin tissue (114) and fetal kidney and adrenal tissue (113) as well as in numerous rat tissues (88,89). On this basis then, and because of established characteristics, high enzyme content, availability and ease of preparation by usual techniques, the adult rat liver system was chosen as a model for further studies dealing with the biotransformation of BaP as related to further objectives of the present research.

EFFECT OF CALF THYMUS DNA ON THE APPEARANCE OF FLUORESCENT PRODUCTS OF BaP BIOTRANSFORMATION edit

In the Introduction, studies were cited which demonstrated covalent bond formation between BaP and DNA as a result of microsome-dependent reactions. Since other evidence cited suggested the DNA binding product to be a precursor of 3-hydroxy BaP, a major product of BaP microsomal transformation, the possibility was considered that DNA binding could be demonstrated in studies involving the microsomal appearance of hydroxylated BaP products.

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On this basis and in an effort to provide a useful approach to the mechanism of enzyme-dependent interactions between DNA and BaP, work was undertaken utilizing a highly sensitive assay method, similar to that adopted in previous studies, which measures the enzymatic appearance of fluorescent hydroxylated BaP products.

Using reaction mixtures (containing adult rat liver microsomes) and incubation conditions similar to those used by Gelboin (24) in earlier studies on bindng of BaP to calf thymus DNA, initial experiments showed that the amount of fluorescent BaP products extracted (first with hexane and then with 1 N NaOH – see Methods) from mixtures incubated in the presence of DNA was substantially less than the amount extracted from mixtures without added DNA (Table 4). This decrease was evident when microsomes used were derived either from animals pretreated with MC in corn oil prior to sacrifice or from non-induced animals pretreated with vehicle alone.

Terminating incubation mixtures by chilling in ice, rather than by precipitation with cold acetone, did not overcome the DNA effect (Table 5). This eliminated the possibility that the decrease in extractible fluorescence by DNA was due to coprecipitation of BaP products with DNA when acetone was added at the end of the incubation period.

Figure 10 shows the amount of fluorescence extracted from incubation mixtures as a function of DNA concentrations.

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At higher DNA levels, the loss of extractible fluorescence varied directly with the DNA concentration. However, at lower levels, no loss of extractible fluorescence was observed, suggesting that some reaction component(s) has the capacity to titrate DNA in a manner which is not directly related to the loss of extractible fluorescence apparent at higher DNA concentrations. It should be noted that apparent titration of DNA from similar mixtures was also reported by Gelboin (24) on the basis of studies involving isotopically-labeled BaP.

In the present research, the amount of fluorescent products extracted from reaction mixtures in the absence or presence of DNA as a function of incubation time was also studied (Figure 11). The difference in the levels of extracted fluorescence between the curves represents the loss in fluorescence due to the presence of DNA in incubation mixtures. This loss is seen to vary directly with the incubation period. Evidently then, the fluorescence loss produced by DNA is incubation-dependent.

In a related study (Figure 12) the DNA effect was found to be maximal when DNA was present initially in mixtures for the complete 14 minute incubation interval but was lessened when DNA was added at various times after the start of the incubation period. This further supports the dependency of the DNA action on the incubation period.

Other studies measured the amount of fluorescent products extracted from reaction mixtures, incubated with

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and without DNA, as a function of BaP concentrations (Figure 13) and NADPH concentration (Figure 14). For reasons which will become clear below, these experiments were performed under conditions in which the enzyme concentration and the incubation time were of first-order kinetics with respect to the appearance of fluorescent products. If titration of the substrates by DNA were responsible for losses in fluorescence, then the only difference between curves involving DNA and those not involving DNA would be one of displacement along the abscissas. No such displacements were observed with regard to either reactant. These graphs also demonstrate that DNA does not act by competitively interfering with either reactant for an enzymatic site. However, to indicate this in a more rigorous fashion, the above results were expressed in the form of reciprocal activity vs. reciprocal reactant concentration plots (Figures 15 and 16). If DNA competed with either BaP or NADPH for an enzymatic site, then the curves, one involving DNA and the other not involving DNA, should intersect on the ordinate of such a plot. No competitive interactions of this sort were found between DNA and either reactant.

Also considered was the possibility that DNA acted by complexing with the fluorescent hydroxylated product itself. In order to investigate this possibility, the hydroxylated products were isolated in the form of a concentrated hexane solution (see Methods) and incubated, substituting for the parent BaP, in reaction mixtures with

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and without added DNA. If DNA acted primarily by complexing with these products, then less of the green fluorescence due to these substances would be expected to be extracted from reaction mixtures containing DNA compared to those incubated in the absence of DNA. The opposite proved to be the case (Figure 17). The inclusion of DNA in reaction mixtures actually increased the amount recoverable by hexane extraction when compared to control mixtures not containing DNA. As demonstrated in the graph, DNA is seen to decrease an incubation-dependent disappearance of hydroxylated products. This further reaction of hydroxylated products to a form not detectible by our fluorometric procedure was found to be enzymatic in nature since boiled microsomes were ineffective in producing incubation-dependent changes (Figure 18). Furthermore, the reaction was found to be NADPH-dependent and inducible.

Nevertheless, these studies indicate that DNA decreases both the enzymatic appearance and disappearance of fluorescent products. Since limiting enzyme amounts were used in all studies described earlier, the possibility was considered that DNA acts by reducing the effective concentration of the microsomal enzyme system itself. Such a possibility was dramatically supported when reaction mixtures, with and without DNA, were studied at various preincubation times before the addition of BaP to initiate the enzymatic appearance of fluorescent BaP products (Figure 19). The amount of fluorescence extracted from reaction mixtures

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varied inversely with the preincubation time when DNA was included in mixtures but was unaffected by the preincubation time when DNA was omitted from mixtures.

In further studies the effect of DNA on the appearance of fluorescent BaP products was studied as a function of microsomal protein concentration (Figure 20). At high protein concentrations, no decrease in fluorescence was observed in mixtures containing DNA compared to control mixtures without DNA. Therefore, decreases in the appearance and disappearance of fluorescent BaP products observed in the presence of DNA under conditions of limiting amounts of enzyme appear to be due, at least in part, to interactions between DNA and microsomal protein resulting in the reduction of effective enzyme concentrations present in incubation mixtures.

Further analysis of previously described experiments are consistent with this conclusion. For example, in studies with various BaP concentrations (Figure 13), maximal enzyme activity (represented by the maximal amount of fluorescence extracted from incubation mixtures at high BaP concentrations) was substantially less in the presence of DNA than in the absence of DNA. Furthermore, at lower BaP concentrations, where enzyme is less limiting, no or smaller decreases were observed in extracted fluorescence due to DNA compared to controls without DNA. Similar results were observed in the case of various NADPH concentrations (Figure 14).

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Preliminary studies demonstrated that DNA forms a precipitable complex with microsomes. In these experiments reaction mixtures containing DNA and microsomes, but no substrates, were incubated for 15 minutes as before. Afterwards, mixtures were centrifuged at 104,000 x g for 1 hour. The resulting supernatants, devoid of particulate microsomes, were chemically analyzed for DNA content and compared to control mixtures containing DNA but no microsomes. It was found that the presence of microsomes increases the loss of DNA remaining in analyzed supernatants (Table 6). (Not unexpectedly, DNA, centrifuged in the absence of particulate reaction components, separated from the upper portions of the centrifuged supernatants sampled in these studies.) Increases in the DNA loss also were found in similar studies involving BaP. When mixtures containing the complete enzyme system (microsomes and substrates) were compared, substantial increases in the DNA loss were observed and appeared to be greater than the sum produced by individual reaction components (Related studies involving UV spectral analysis of supernatants revealed little difference between absorbance values at 260 mμ in supernatants containing both DNA and NADPH and the summation of absorbance values observed in supernatants containing DNA alone and NADPH alone).

Nevertheless, precipitable complexes responsible for the DNA losses need not be related to the enzymatic studies described earlier since, as demonstrated with regard to the

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effect of various DNA concentrations on the fluorometric appearance of BaP products (Figure 10), reaction components are able to titrate DNA in a manner which appears to be unrelated to the enzymatic conversion of BaP to fluorescent products.

EFFECT OF VARIOUS MONONUCLEOTIDES AND SYNTHETIC POLYNUCLEOTIDES ON THE APPEARANCE OF FLUORESCENT PRODUCTS OF BaP BIOTRANSFORMATION edit

In a further effort to provide insight into the intricacies of microsome-dependent BaP-DNA interactions, the basic approach adopted in the preceding studies was extended to include various mononucleotides and synthetic polyribonucleotides.

Initial studies revealed no difference from controls in the amounts of fluorescent BaP products extracted from incubation mixtures under usual conditions of excessive substrate and limiting enzyme concentrations when various deoyribomononucleotides (Table 7) or ribomononucleotides (Table 8) were studied. However, when a series of synthetic ribopolynucleotides were studied, substantial decreases in the extractible fluorescence were observed in the case of Poly G and, to lesser extents, in the cases of Poly I and the alternating copolymer Poly (I·C) (Table 9). In contrast, Poly A, Poly C, Poly U or Poly I·Poly C (double-stranded structure was maintained in these experiments as indicated by a 41% increase in absorbancy at 248 mμ at 100°C compared to readings at room temperature) did not significantly alter the amounts of fluorescence recovered

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from incubation mixtures.

In order to determine whether a relationship exists between the ability of polynucleotides to decrease the fluorometric appearance of enzymic BaP products and their ability to enzymatically bind BaP, Poly G, representative of a polynucleotide which decreases recoverable fluorescence, and Poly A, representative of a polynucleotide which does not alter recoverable fluorescence, were incubated in mixtures containing isotopically-labeled BaP. After centrifugation to remove the particulate microsomes and extraction with hexane, reaction mixtures were carefully layered on top of a sucrose gradient and centrifuged for various times. In the case of Poly A, no peak of radioactivity was associated with the location of the polynucleotide in the gradient whether developed by a 12 hour run (Figure 31) or by a 24 hour run (Figure 22). However, in the case of Poly G, a peak of radioactivity was associated with the polynucleotide after centrifuging the gradient for 12 hours (Figure 23) and was seen to migrate with continued centrifugation (Figure 24). (Maximal levels of peak radioactivity were obtained only when NADPH was included in incubation mixtures). Similar results, but involving smaller amounts of associated radioactivity, were obtained when Poly I, also able to decrease recoverable fluorescence as Poly G but to a lesser extent, was studied (Figure 25). However, Poly I·poly C, unable to alter recoverable fluorescence, was not associated with

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radioactivity in gradient studies (Figure 26). (Poly I·Poly C, sedimented in gradient tubes, maintained the double-stranded structure as revealed in hyperchromicity studies described earlier.) These findings suggest, at least in the present polymer series, that the ability of polynucleotides to decrease the extraction of enzymic BaP products is directly related to their ability to form a chemical bond with BaP.

In other studies the decreases resulting from the inclusion of Poly G in incubation mixtures were not overcome when the excessive substrate (BaP and NADPH) concentrations normally used in these studies were doubled. However, the fluorometric decreases due to Poly G, unlike those due to DNA (Figure 19), were unaffected when various preincubation times were studied (Figure 27). Also unlike results with DNA (Figure 17), Poly G, as well as Poly I, did not appreciably alter the enzymatic disappearance of hydroxylated fluorescent BaP products (Figure 28). These results demonstrate that Poly G acts to produce decreases in the fluorometric appearance of BaP products in a way basically different from the manner observed with DNA. Furthermore, on the basis of the aforementioned studies, it seems likely that Poly G decreases fluorometric appearance of BaP products by strongly binding with enzymic BaP products under usual conditions of limiting enzyme concentrations. (At excessive enzyme levels, 3.115 mg microsomal protein, increases amounting to more

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than 20% in recoverable fluorescence were observed in mixtures containing Poly G compared to control mixtures.)

EFFECT OF CALF THYMUS DNA MODIFIED BY NITROGEN MUSTARD ON THE APPEARANCE OF FLUORESCENT PRODUCTS OF BaP BIOTRANSFORMATION edit

The possibility was suggested by the preceding studies that enzymic binding between DNA and BaP might involve preferential reaction with guanine residues on the DNA molecule. Indeed, studies reported by others (95), using nonenzymatic model BaP hydroxylating systems, have established guanine specificity with regard to chemical binding between DNA and BaP. In order to test such a supposition in the present research, DNA was alkylated with nitrogen mustard (HN₂) according to procedures published previously (100). Modifications of DNA in this manner has been shown to specifically involve chemical binding between HN₂ and the N-7 position of guanine in the DNA molecule (110). As shown in Table 10, no substantial difference was observed in the fluorometric appearance of BaP products under usual conditions when the effect of including HN₂-modified DNA in incubation mixtures was compared with control mixtures without modified DNA. In contrast, control unmodified DNAs decreased extractible fluorescence as before. These findings, although consistent with the possibility that guanine is involved in the enzyme catalyzed reaction between DNA and BaP, should be interpreted with caution since interactions between DNA and other reaction components, notably enzyme, are known

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to occur under the usual experimental conditions employed in these studies (see above).

EFFECT OF YEAST RNA AND DENATURED CALF THYMUS DNA ON THE APPEARANCE OF FLUORESCENT PRODUCTS OF BaP BIOTRANSFORMATION edit

Described results demonstrated a direct relationship between covalent BaP binding and a decrease in the microsome-dependent appearance of fluorescent BaP products for a series of synthetic polyribonucleotides. On the basis of these experiments as well as studies involving HN₂-modified DNA, the possibility was supported that BaP binds preferentially to guanine sites on DNA as a result of microsome catalyzed reactions. For these reasons and in order to further test the usefulness of the adopted fluorometric approach, it seemed worthwhile to extend these studies to include denatured DNA and yeast RNA.

The inclusion of denatured DNA and yeast RNA in incubation mixtures increased, in contrast to results with native DNA and Poly G, the amount of extractible fluorescence compared to control mixtures (Table 11). Preliminary studies indicated that increases due to RNA (approximately 10%) were unaffected when incubation mixtures contained either limiting enzyme (0.534 mg microsomal protein) or excessive enzyme (3.115 mg microsomal protein). Increases due to denatured DNA, on the other hand, were greater at excessive enzyme levels (approximately 40% over controls) than at limiting levels (approximately 25%). Furthermore, the effect of various preincubation times (Figure 29) did

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not alter the increase in extractible fluorescence due to RNA although slightly less increase due to denatured DNA was observed at longer preincubation periods.

RNA and denatured DNA were also studied with regard to the microsome-dependent disappearance of hydroxylated fluorescent BaP products (Figure 30). Substantial increases in the disappearance were noted in both cases and the extent of these increases were roughly equivalent to the increases in the enzymatic appearance of fluorescent BaP products in each case respectively (RNA increased the appearance and disappearance reactions approximately 10-15% after incubation for 14 minutes whereas denatured DNA increased both processes approximately 25%).

When RNA was prepared as before by incubating in the presence of isotopically-labeled BaP and sedimented in a sucrose gradient, the crude yeast RNA separated into two RNA fractions: one sedimenting near the bottom of the gradient and the other remaining at the top of the gradient (Figure 31). Only the RNA fraction remaining at the top of gradients was associated with a peak of radioactivity. Although not analyzed it seems likely that the sedimenting fraction contains the heavier ribosomal RNA subunits whereas the unsedimenting fraction contains the lighter ribosomal subunits as well as the transfer RNA.

Nevertheless, single-stranded DNA and RNA appear to be associated with a process at limiting enzyme concentrations which serves to increase the appearance and

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disappearance of fluorometric products of BaP biotransformation and, at least in the case of an RNA fraction which sediments easily in a sucrose gradient, does not appear to be associated with chemical bond formation involving BaP.

BIOLOGICAL ACTIVITY OF CALF THYMUS DNA CHEMICALLY MODIFIED BY BaP BIOTRANSFORMATION edit

A final objective of the present research was to determine the biological significance of DNA chemically altered by microsome-dependent transformation of BaP. The approach adopted in these experiments involved a comparison between the abilities of modified DNA and various unmodified DNA controls to serve as templates in a DNA-dependent RNA polymerase system. In order to make the results even more meaningful, DNA treated with BeP, substituting for BaP in modification procedures, was also studied. BeP, a noncarcinogenic structural isomer of BaP, has been shown by others (111, 112) to be transformed by rat liver microsomes to hydroxylated products in much the same manner at BaP. In the present research BeP, like BaP, yielded fluorescent products when usual assay procedures were employed. However, the fluorospectral characteristics of BeP products (Figure 32) differed markedly from those of BaP products (Figure 33). Furthermore, maximal fluorescent intensity of BeP products was less than one-tenth that of the BaP products at their respective fluorospectral maxima (BeP: 365 mμ excitation, 485 mμ emission: BaP: 390 mμ excitation, 515 mμ emission). As shown in Table 12, the appearance of fluorescent BeP

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products was microsome- and incubation-dependent and required NADPH and microsomes derived from MC-pretreated animals for maximal yield. When DNA was added to incubation mixtures, decreases in the appearance of extractible fluorometric BeP products were observed and were proportionately similar to decreases observed in the fluorometric appearance of BaP products previously described.

With regard to template studies, DNA was incubated in mixtures containing either BaP, BeP or vehicle alone. After the incubation period mixtures were centrifuged at high speed to remove particulate reaction components, extracted with hexane, dialyzed to remove NADPH and, finally, concentrated to an appropriate volume. The resulting DNA solutions were reintroduced into their respective incubation mixtures and the isolation procedure was repeated. After a further reintroduction into mixtures and isolation, DNA solutions (which, in the case of BaP-treated DNA, was unable to decrease the enzymatic appearance of fluorescent BaP products) were dialyzed against 0.1 M Tris-HCl buffer (pH 7.5) and extracted with a fresh phenol solution. The final DNA solutions, representing approximately 55% of the original starting DNA, were assayed for template abilities in a DNA-dependent RNA polymerase system purified from E. coli bacterial cells. A nonlinear incubation period of 30 minutes was employed in all studies. The results are summarized in Figure 34. Each point on the graph represents the mean of three experiments. Control DNA, not treated

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with either BaP or BeP, was as efficient a template (represented by incorporation of tritiated-ATP into an acid-insoluble product) as DNA not undergoing the modification procedure. By comparison, DNA modified with BaP was substantially less effective a template and was indistinguishable in template ability from DNA treated with BeP. These findings suggest that DNA, treated enzymatically with benzopyrenes, is biologically modified on the basis of studies involving a model transcription system, but in a manner which does not appear to be carcinogen-specific.

66 - FIGURE 3. SUBFRACTIONAL DISTRIBUTION OF ARYL HYDROCARBON HYDROXYLASE ACTIVITY IN RAT PLACENTAL HOMOGENATES. edit

67 - CLICK HERE FOR PAGE-067-FIGURE-03

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68 - TABLE 1. SUMMARY OF ARYL HYDROCARBON HYDROXYLASE ACTIVITY IN HUMAN AND RAT PLACENTAL AND HEPATIC HOMOGENATES. edit

69 - CLICK HERE FOR PAGE-069-TABLE-01

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70 - TABLE 2. SUMMARY OF APPARENT KINETIC CONSTANTS+ OF ARYL HYDROCARBON HYDROXYLASE IN HUMAN TERM AND RAT++ PLACENTAL AND RAT MATERNAL HEPATIC HOMOGENATES. edit

+Determined by the Lineweaver-Burke graphical method of analysis. Analyses were performed on placental and hepatic homogenates of MC-pretreated rats (pool from 12 animals) and women who smoked cigarettes (pool from 4 women at term).

++Rat tissues were analyzed at 17 days gestation.

71 - CLICK HERE FOR PAGE-071-TABLE-02

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72 - FIGURE 4. RECIPROCAL ACTIVITY vs. RECIPROCAL NADPH CONCENTRATION PLOTS FOR RAT MATERNAL LIVER AND PLACENTAL PREPARATIONS. edit

These plots are uncorrected for NADPH oxidase activity.

73 - CLICK HERE FOR PAGE-073-FIGURE-04

PAGE-073-FIGURE-04

74 - FIGURE 5. DISAPPEARANCE OF NADPH IN RAT MATERNAL LIVER (9000 x g supernatant) AND PLACENTAL (1000 x g pellet) HOMOGENATE SUBFRACTIONS. edit

Protein concentrations were 1.6 and 0.9 mg protein/ml for liver and placenta respectively.

75 - CLICK HERE FOR PAGE-075-FIGURE-05

PAGE-075-FIGURE-05

76 - FIGURE 6. COMPARISON OF THE EFFECTS OF NADPH AND NADH ON THE ARYL HYDROCARBON ACTIVITY FOUND IN RAT MATERNAL LIVER AND PLACENTAL HOMOGENATES. edit

A relative specific activity of 100 represents 2440 fluorescent units/mg protein/hr for the liver preparation and 25.8 fluorescent units/mg protein/hr for the placental preparation.

77 - CLICK HERE FOR PAGE-077-FIGURE-06

PAGE-077-FIGURE-06

78 - FIGURE 7. EFFECT OF INCUBATING RAT MATERNAL LIVER AND PLACENTAL HOMOGENATE UNDER VARIOUS SATURATED GAS PHASES. edit

The specific activity of the liver enzyme at 0% inhibition was 3000 fluorescent units/mg protein/hr and 26.3 units/mg protein/hr for the placenta enzyme.

79 - CLICK HERE FOR PAGE-079-FIGURE-07

PAGE-079-FIGURE-07

80 - FIGURE 8. EFFECT OF FMN AND FAD ON THE ARYL HYDROXYLASE ACTIVITY IN RAT MATERNAL LIVER AND PLACENTAL HOMOGENATES. edit

A relative specific activity of 100 represents 3470 fluorescent units/mg protein/hr for the liver homogenate and 19.8 fluorescent units/mg protein/hr for the placental homogenate.

81 - CLICK HERE FOR PAGE-081-FIGURE-08

PAGE-081-FIGURE-08

82 - FIGURE 9. EFFECT OF PCMB ON RAT MATERNAL LIVER AND PLACENTAL ARYL HYDROCARBON HYDROXYLASE ACTIVITY. edit

A relative specific activity of 100 represents 2920 fluorescent units/mg protein/hr for the liver preparations and 24.3 fluorescent units/mg protein/hr for the placental preparations.

83 - CLICK HERE FOR PAGE-083-FIGURE-09

PAGE-083-FIGURE-09

84 - TABLE 3. EFFECT OF VARIOUS STEROIDS ON THE RELATIVE SPECIFIC ACTIVITY OF ARYL HYDROCARBON HYDROXYLASE IN HUMAN AND RAT PLACENTAL HOMOGENATES AND RAT LIVER HOMOGENATES. edit

A relative specific activity when no steroid was added to incubation mixtures was 100 for liver and placental preparations. A relative specific activity of 100 represents a mean specific activity of 1610 fluorescent units/mg protein/hr, 46.0 fluorescent units/mg protein/hr and 64.0 fluorescent units/mg protein/hr for rat liver, rat placental and human (term) placental homogenates respectively. Rat homogenates were prepared from the pooled tissues of 12 animals. All experiments were in triplicate and verified by repeating at least once with the same homogenate. Results as expressed represent the percentage of the mean specific activity of 6 incubation flasks.

85 - CLICK HERE FOR PAGE-085-TABLE-03

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86 - TABLE 4. EFFECT OF DNA ON EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF ANIMAL PRETREATMENT. edit

Female rats (180-230 grams each) were pretreated with MC (20 mg/kg, i.p.) in corn oil or corn oil alone 48 hours before sacrifice. 1.50 mg calf thymus DNA was added to indicated reaction mixtures before incubation. Results as expressed represent mean values (per 14 minutes incubation) of four determinations with standard error.

87 - CLICK HERE FOR PAGE-087-TABLE-04

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88 - TABLE 5. EFFECT OF DNA ON EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF THE MANNER INCUBATION MIXTURES WERE TERMINATED. edit

(See legend for Table 4.)

89 - CLICK HERE FOR PAGE-089-TABLE-05

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90 - FIGURE 10. EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF DNA CONCENTRATION. edit

Results as expressed represent total extracted fluorescence per 14 minutes incubation.

91 - CLICK HERE FOR PAGE-091-FIGURE-10

PAGE-091-FIGURE-10

92 - FIGURE 11. EFFECT OF DNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF INCUBATION TIME. edit

1.5 mg calf thymus DNA was added to appropriate mixtures.

93 - CLICK HERE FOR PAGE-093-FIGURE-11

PAGE-093-FIGURE-11

94 - FIGURE 12. EFFECT OF ADDING DNA AT VARIOUS TIMES DURING THE INCUBATION PERIOD ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE. edit

1.5 mg calf thymus DNA was added to appropriate mixtures. A relative value of 100 represents approximately 4950 total extracted fluorescent units/incubate/14 minutes.

95 - CLICK HERE FOR PAGE-095-FIGURE-12

PAGE-095-FIGURE-12

96 - FIGURE 13. EFFECT OF DNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF BaP CONCENTRATIONS. edit

1.5 mg calf thymus DNA was added to appropriate mixtures. Appearance of fluorescent products was linear during the 5 minute incubation period.

97 - CLICK HERE FOR PAGE-097-FIGURE-13

PAGE-097-FIGURE-13

98 - FIGURE 14. EFFECT OF DNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF NADPH CONCENTRATIONS. edit

(See legend for figure 13.)

99 - CLICK HERE FOR PAGE-099-FIGURE-14

PAGE-099-FIGURE-14

100 - FIGURE 15. RECIPROCAL ACTIVITY vs. RECIPROCAL BaP CONCENTRATION PLOTS FOR ARYL HYDROCARBON HYDROXYLASE ACTIVITIES IN THE PRESENCE AND ABSENCE OF DNA. edit

These plots are derived from data presented in Figure 13.

101 - CLICK HERE FOR PAGE-101-FIGURE-15

PAGE-101-FIGURE-15

102 - FIGURE 16. RECIPROCAL ACTIVITY vs. RECIPROCAL NADPH CONCENTRATION PLOTS FOR ARYL HYDROCARBON HYDROXYLASE ACTIVITIES IN THE PRESENCE AND ABSENCE OF DNA. edit

These plots are derived from data presented in Figure 14.

103 - CLICK HERE FOR PAGE-103-FIGURE-16

PAGE-103-FIGURE-16

104 - TABLE 6. EFFECT OF HIGH SPEED CENTRIFUGATION ON THE SEDIMENTATION OF CALF THYMUS DNA IN INCUBATION MIXTURES CONTAINING VARIOUS REACTION COMPONENTS OF THE ARYL HYDROCARBON HYDROXYLASE ASSAY SYSTEM. edit

Usual substrate, cofactor and microsomal protein concentrations were employed in these studies. 0.25 M sucrose solution replaced microsomal preparation and NADPH solution in appropriate mixtures; acetone replaced BaP solutions. The final volume in all mixtures was 3.0 ml. Only the top 0.5 ml of supernatants were chemically analyzed for DNA. Results as expressed represent the mean of two determinations. See the text for further experimental details.

105 - CLICK HERE FOR PAGE-105-TABLE-06

PAGE-105-TABLE-06

106 - FIGURE 17. EFFECT OF DNA ON THE DISAPPEARANCE OF FLUORESCENT BaP PRODUCTS FROM MICROSOMAL MIXTURES AS A FUNCTION OF INCUBATION TIME. edit

Only microsomes derived from MC-pretreated animals were used. 1.5 mg calf thymus DNA was added to appropriate mixtures. A relative value of 100 represents 7200 extractible fluorescent units of BaP products originally added to incubation mixtures.

107 - CLICK HERE FOR PAGE-107-FIGURE-17

PAGE-107-FIGURE-17

108 - FIGURE 18. EFFECT OF VARIOUS REACTION CONDITIONS ON DISAPPEARANCE OF FLUORESCENT BaP PRODUCTS FROM MICROSOMAL MIXTURES AS A FUNCTION OF INCUBATION TIME. edit

Microsomes were derived from corn oil (CO)-pretreated animals only where indicated; otherwise, microsomes were derived from MC-pretreated animals. A relative value of 100 represents 7200 extractible fluorescent units of BaP products originally added to incubation mixtures.

109 - CLICK HERE FOR PAGE-109-FIGURE-18

PAGE-109-FIGURE-18

110 - FIGURE 19. EFFECT OF DNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF PREINCUBATION TIME. edit

1.5 mg calf thymus DNA was added to appropriate mixtures. After preincubating mixtures (minus substrate) at 37°C, BaP was added and mixtures further incubated for 14 minutes.

111 - CLICK HERE FOR PAGE-111-FIGURE-19

PAGE-111-FIGURE-19

112 - FIGURE 20. EFFECT OF DNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF MICROSOMAL PROTEIN CONCENTRATION. edit

1.5 mg calf thymus DNA was added to appropriate mixtures.

113 - CLICK HERE FOR PAGE-113-FIGURE-20

PAGE-113-FIGURE-20

114 - TABLE 7. EFFECT OF VARIOUS 5’-DEOXYRIBOMONONUCLEOTIDES ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE. edit

A relative value of 100 represents 8210 total extracted fluorescent units/incubate/14 minutes.

115 - CLICK HERE FOR PAGE-115-TABLE-07

PAGE-115-TABLE-07

116 - TABLE 8. EFFECT OF VARIOUS 5’-RIBOMONONUCLEOTIDES ON THIS EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE. edit

A relative value of 100 represents 8496 total extracted fluorescent units/incubate/14 minutes.

117 CLICK HERE FOR PAGE-117-TABLE-08

PAGE-117-TABLE-08

118 - TABLE 9. EFFECT OF VARIOUS SYNTHETIC RIBOPOLYNUCLEOTIDES ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE. edit

A relative value of 100 represents approximately 7000 total extracted fluorescence units/incubate/14 minutes. Each experimental result represents the mean of at least two determinations.

119 - CLICK HERE FOR PAGE-119-TABLE-09

PAGE-119-TABLE-09

120 - FIGURE 21. SUCROSE GRADIENT SEDIMENTATION OF POLY A TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 12 HOUR RUN. edit

A 0.075 ml aliquot of incubation mixture supernatant (see Methods) was layered on top of a 5.0 ml sucrose gradient (5-20%) and centrifuged for 12 hours. Fraction 1 represents the top of the gradient.

121 - CLICK HERE FOR PAGE-121-FIGURE-21

PAGE-121-FIGURE-21

122 - FIGURE 22. SUCROSE GRADIENT SEDIMENTATION OF POLY A TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 24 HOUR RUN. edit

A 0.050 ml aliquot of incubation mixture supernatant (see Methods) was layered on top of a 5.0 ml sucrose gradient (5-20%) and centrifuged for 24 hours. Fraction 1 represents the top of the gradient.

123 - CLICK HERE FOR PAGE-123-FIGURE-22

PAGE-123-FIGURE-22

124 - FIGURE 23. SUCROSE GRADIENT SEDIMENTATION OF POLY G TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 12 HOUR RUN. edit

(See legend for Figure 21.)

125 - CLICK HERE FOR PAGE-125-FIGURE-23

PAGE-125-FIGURE-23

126 - FIGURE 24. SUCROSE GRADIENT SEDIMENTATION OF POLY G TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 24 HOUR RUN. edit

(See legend for Figure 22.)

127 - CLICK HERE FOR PAGE-127-FIGURE-24

PAGE-127-FIGURE-24

128 - FIGURE 25. SUCROSE GRADIENT SEDIMENTATION OF POLY I TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 18 HOUR RUN. edit

A 0.075 ml aliquot of incubation mixture supernatant (see Methods) was layered on top of a 5.0 ml sucrose gradient (5-20%) and centrifuged for 18 hours. Fraction 1 represents the top of the gradient.

129 - CLICK HERE FOR PAGE-129-FIGURE-25

PAGE-129-FIGURE-25

130 - FIGURE 26. SUCROSE GRADIENT SEDIMENTATION OF POLY I·POLY C TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 18 HOUR RUN. edit

(See legend for Figure 25.)

131 - CLICK HERE FOR PAGE-131-FIGURE-26

PAGE-131-FIGURE-26

132 - FIGURE 27. EFFECT OF POLY G ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF PREINCUBATION TIME. edit

2.0 mg Poly G was added to appropriate mixtures. After preincubating mixtures (minus substrate) at 37°C, BaP was added and mixtures further incubated for 14 minutes.

133 - CLICK HERE FOR PAGE-133-FIGURE-27

PAGE-133-FIGURE-27

134 - FIGURE 28. EFFECTS OF POLY G AND POLY I ON THE DISAPPEARANCE OF FLUORESCENT BaP PRODUCTS FROM MICROSOMAL MIXTURES AS A FUNCTION OF INCUBATION TIME. edit

Only microsomes derived from MC-pretreated animals were used. 2.0 mg of either Poly G or Poly I was added to appropriate mixtures. A relative value of 100 represents 7200 extractible fluorescent units of BaP products originally added to incubation mixtures.

135 - CLICK HERE FOR PAGE-135-FIGURE-28

PAGE-135-FIGURE-28

136 - TABLE 10. EFFECT OF DNA, CHEMICALLY MODIFIED WITH NITROGEN MUSTARD (HN₂), ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE. edit

1.5 mg calf thymus DNA, treated in various ways with and without HN₂ (see Methods), was added to appropriate mixtures. A relative value of 100 represents 4680 total extracted fluorescent units/incubate/14 minutes. Each experimental result represents the relative mean of two determinations.

137 - CLICK HERE FOR PAGE-137-TABLE-10

PAGE-137-TABLE-10

138 - TABLE 11. EFFECT OF CALF THYMUS DNA AND YEAST RNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE. edit

A relative value of 100 represents approximately 8100 units/incubate/14 minutes. Experimental results as expressed represent the relative means of at least two determinations.

139 - CLICK HERE FOR PAGE-139-TABLE-11

PAGE-139-TABLE-11

140 - FIGURE 29. EFFECT OF YEAST RNA AND DENATURED CALF THYMUS DNA ON THE EXTRACTIBLE FLUORESCENCE PRODUCED BY ARYL HYDROCARBON HYDROXYLASE AS A FUNCTION OF PREINCUBATION TIME. edit

2.0 mg RNA and 1.5 mg denatured DNA was added to appropriate mixtures. After preincubating mixtures (minus substrate) at 37°C, BaP was added and mixtures further incubated for 14 minutes.

141 - CLICK HERE FOR PAGE-141-FIGURE-29

PAGE-141-FIGURE-29

142 - FIGURE 30. EFFECT OF YEAST RNA AND DENATURED CALF THYMUS DNA ON THE DISAPPEARANCE OF FLUORESCENT BaP PRODUCTS FROM MICROSOMAL MIXTURES AS A FUNCTION OF INCUBATION TIME. edit

Only microsomes derived from MC-pretreated animals were used. 2.0 mg RNA and 1.5 mg denatured DNA was added to appropriate mixtures. A relative value of 100 represents 7200 extractible fluorescent units of BaP products originally added to incubation mixtures.

143 - CLICK HERE FOR PAGE-143-FIGURE-30

PAGE-143-FIGURE-30

144 - FIGURE 31. SUCROSE GRADIENT SEDIMENTATION OF YEAST RNA TREATED WITH ³H-BaP IN A MICROSOMAL INCUBATION MIXTURE – 18 HOUR RUN. edit

A 0.075 ml aliquot of incubation mixture supernatant (see Methods) was layered on top of a 5.0 ml sucrose gradient (5-20%) and centrifuged for 18 hours. Fraction 1 represents the top of the gradient. To clearly illustrate the association of RNA remaining at the top of centrifuged gradients and radioactivity, graphs were corrected for absorbance due to NADPH and radioactivity on the basis of similar studies not employing RNA.

145 - CLICK HERE FOR PAGE-145-FIGURE-31

PAGE-145-FIGURE-31

146 - FIGURE 32. EXCITATION-EMISSION FLUOROSPECTROGRAPH OF PRODUCTS OF BeP MICROSOMAL TRANSFORMATION. edit

The excitation spectra was the result of a fluorometric wavelength scan at the emission maximum,485 mμ; the emission spectra at the excitation maximum, 365 mμ. See the Methods for further experimental details.

147 - CLICK HERE FOR PAGE-147-FIGURE-32

PAGE-147-FIGURE-32

148 - FIGURE 33. EXCITATION-EMISSION FLUOROSPECTROGRAPH OF PRODUCTS OF BaP MICROSOMAL TRANSFORMATION. edit

The excitation spectra was the result of a fluorometric wavelength scan at the emission maximum, 515 mμ; the emission spectra at the excitation maximum, 390 mμ. See the Methods for further experimental details. It should be noted that identical excitation and emission spectra resulted when authentic 3-hydroxy BaP (kindly supplied by Dr. L.W. Wattenberg) was studied.

149 - CLICK HERE FOR PAGE-149-FIGURE-33

PAGE-149-FIGURE-33

150 - TABLE 12. EXTRACTIBLE FLUORESCENCE DUE TO BeP MICROSOMAL TRANSFORMATION UNDER VARIOUS EXPERIMENTAL CONDITIONS. edit

Reaction and incubation conditions were the same as those usually employed in studies of BaP microsome transformation (see Methods). Fluorometric determinations were performed at 365 mμ (excitation maximum) and 485 mμ (emission maximum) (see Figure 32). Microsomes were derived either from MC- or CO- pretreated animals as indicated. 1.5 mg calf thymus DNA was added to appropriate mixtures.

151 - CLICK HERE FOR PAGE-151-TABLE-12

PAGE-151-TABLE-12

152 - FIGURE 34. TEMPLATE ACTIVITY OF CALF THYMUS DNA, TREATED WITH BENZOPYRENE IN MICROSOMAL INCUBATION MIXTURES, IN AN E. COLI DNA-DEPENDANT RNA POLYMERASE SYSTEM AS A FUNCTION OF DNA CONCENTRATION. edit

As described in Methods, DNA was chemically analyzed according to a published procedure (102). Each point on the graph represents the mean of 3 determinations.

153 - CLICK HERE FOR PAGE-153-FIGURE-34

PAGE-153-FIGURE-34

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DISCUSSION edit

One objective of the present research was to study the biological transformation of BaP in various tissues of the human and rat fetal-placental unit. A comparison of characteristics of the BaP biotransformation processes in the various tissues was attempted and was based on reported features of the aryl hydrocarbon hydroxylase system including inducibility, subcellular localization, requirements for optimal activity and the effect of various agents.

Biotransformation of BaP in all tissues studied was found to be enzymatic in nature and required NADPH and oxygen for maximal activity. Furthermore, the systems had similar pH optima and, on the basis of preliminary studies, appeared to be localized in microsomal subfractions. The effects of various inhibitors including carbon monoxide, nitrogen, PCMB, iodacetamide and a series of steroids were similar. Generally, tissue homogenates derived from inducer pretreated animals and smoking human subjects (especially with regard to term placentas) were associated with substantially increased activities compared to control pretreated animals and nonsmoking human subjects respectively. Although differences were observed between the various systems studied (apparent specific activities and certain kinetic parameters, effect of EDTA and flavins), the similarities appeared to be more striking. On this basis then, and because of established characteristics, high enzyme content, availability and ease of preparation by usual techniques, the adult rat liver system was chosen

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as a model for further studies dealing with the biotransformation of BaP as related to further objectives in the present research.

Fetal placental unit studies involving various steroids demonstrated substantial inhibition by estrogens (i.e., estrone and β-estradiol) of the recovery of fluorescent BaP products in all tissues studied. Although the exact nature of the inhibition was not determined in the present research, one likely possibility is that estrogens are biologically transformed themselves by the AHH system and, in this manner, interfere with the enzymatic appearance of fluorescent BaP products. Further studies of the relationship between the AHH system and estrogens may be useful in providing insight into the currently outstanding problem of estrogen catabolism in vivo especially with regard to the little known role played by the placenta during pregnancy in regulating plasma estrogen levels.

As related to carcinogenesis, the presence of the BaP biotransformation processes in developing fetal tissue may be of considerable significance in the initiation of carcinogenesis arising during early development. This is especially relevant when it is considered that fetal tissue, due to the greater proliferating rate of cells during early development stages, may be associated with an increased carcinogenic risk as a result of exposure to biologically activated carcinogens than more mature, but slower dividing, tissue.

Nevertheless, the biological significance of fetal

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placental unit studies to the potential problem of polycyclic hydrocarbon carcinogenesis arising early in development requires that the ability of hydrocarbons to penetrate membrane barriers, including the placental barrier, be considered. Polycyclic hydrocarbons are highly lipophilic chemically and, as such, would be expected to easily penetrate such barriers. Indeed, studies demonstrate that not only are polycyclic hydrocarbons able to transgress the placental barrier but, more importantly, substantial amounts of tritium-labeled hydrocarbons are found in fetal liver tissue after intraperitoneal pretreatment of maternal rats (116). In the present studies, significant induction of AHH activity in fetal livers resulted after pretreating maternal animals with polycyclic hydrocarbons. This suggests that hydrocarbons penetrate the placental barrier as well as hepatocellular membranes and, presumably, subcellular barriers in order to interact with receptor material essential to the induction response. Recent studies by Carlassare et al. (117) demonstrate covalent binding between BaP and DNA from liver, spleen and skin after a single does of tritium-labeled BaP administered orally to rodents. Apparently then, biological membrane barriers are ineffectual in preventing the penetration of BaP and related hydrocarbons into a variety of tissues including those important to fetal development. The presence in fetal tissues of BaP as well as an enzymatic system similar to one which has been demonstrated to covalently attach BaP to DNA in a manner, as shown in the present

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research, which impairs DNA template function, may be related to the carcinogenic effects observed in utero after pretreating maternal animals with BaP (94) (see Introduction). This observation, however, need not be limited to carcinogenic events initiated during early stages of development and may be equally applied to carcinogenicity occurring at maturity and involving developed liver, skin and intestinal tissue and, of particular relevance to carcinogenesis associated with cigarette smoking, lung tissue.

Because the involvement of carcinogen metabolizing enzymes in chemical reaction with DNA is of fundamental importance to the mechanism of chemical carcinogenesis, other studies in the present research considered the interaction between BaP and various polynucleotides in the presence of the model AHH system. Inclusion of DNA in incubation mixtures containing BaP and the AHH system substantially reduced the recovery of BaP products as compared to controls. Initial studies showed the reduction to be incubation-dependent and, at higher DNA levels, to vary directly with the DNA concentration. Furthermore, the reduction by DNA did not appear to be due to interference with either BaP or NADPH for an enzymatic site nor to involve complexation with recoverable hydroxylated products. When various preincubation times were studied, the amount of recoverable products was found to vary inversely with the preincubation time when DNA was included in mixtures but was unaffected when DNA was omitted. Further, the DNA action was overcome when the usual enzyme concentration

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was increased. These findings strongly suggest that DNA acts to decrease recoverable products, at least in part, by interacting with microsomal protein in a manner which reduces the effective enzyme concentrations present in incubation mixtures. Such interactions are hardly surprising in view of the many examples of nucleic acid-protein interactions observed in biological system including the close association of nucleoprotein and nucleic acids in chromatin material derived from nucleated cells. It should be noted that, due to subcellular compartmentalization, DNA and microsomal enzymes would not be expected to interact in vivo and, for this reason, such interactions may be unrelated to chemical carcinogenesis generally and, specifically, to covalent binding between BaP and DNA either in vivo or, more related to the present research, in vitro. However, in view of reported findings that BaP covalently binds to DNA as demonstrated by sedimentation studies (24), the decreases in appearance of fluorescent BaP products by DNA need not necessarily be accounted for by titration of the enzyme only. On this basis, then, studies involving model systems were initiated.

When various component nucleotides were studied, no differences from controls in the amount of fluorescent BaP products recovered from incubation mixtures were observed. However, when a series of synthetic polynucleotides were studied, substantial decreases in extractible fluorescence were observed in the case of Poly G and, to lesser extents, in the cases of Poly I and the alternating copolymer Poly (I·C).

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In contrast, Poly A, Poly C, Poly U and the double-stranded Poly I·Poly C did not significantly alter the amounts of fluorescence recovered from incubation mixtures. Decreases due to Poly G, unlike those due to DNA, were unaffected when various preincubation times were studied. When Poly G was incubated in mixtures containing labeled BaP, isolated and sedimented in a sucrose gradient, a peak of radioactivity was associated with the polynucleotide. Similar results, but involving smaller amounts of associated radioactivity were obtained with Poly I. In contrast, related studies involving Poly A and the double stranded Poly I·Poly C, demonstrated no detectible peak of radioactivity associated with the location of polynucleotides in centrifuged gradients. These findings, when compared with the decreases in fluorescent BaP products produced by polynucleotides, demonstrate a direct correlation with the ability to covalently bind BaP as a result of microsomal reactions.

Results indicated that Poly G substantially decreased the enzymatic appearance of fluorescent BaP products while equivalent amounts of monomeric guanine were without observable effect. Apparently, the polymerized form provides suitable positioning of the BaP molecule in relation to crucial sites of reaction. Since Poly G molecules tend to aggregate in multistranded complexes in aqueous solution (120), a likely possibility is that BaP intercalates between bases within the Poly G molecule such that the resulting conformational fitting favors chemical reaction with sites on adjoining

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Poly G molecules. Presumably, Poly I, being similar to Poly G in chemical structure and in ability to form multistranded aggregates in solution (121), interacts with BaP in much the same manner as Poly G. With regard to the double stranded Poly I·Poly C, no covalent binding with BaP was observed nor was Poly I·Poly C able to decrease the enzyme-dependent appearance of fluorescent BaP products. One possible explanation is that the BaP molecule can intercalate between base stacks in the Poly I strand but cannot chemically react with cytosine on the adjoining strand. It should be noted that, due to base pairing interactions between inosine and cytosine in the double-stranded structure, a likely site of chemical reaction on the cytosine molecule, position N-3, may not be available for reaction with BaP. However, one might expect the alternating copolymer, Poly (I·C), to be suitable for both physical and chemical reaction with BaP since one strand may serve as a structure permitting suitable intercalation of the BaP molecule while an adjoining strand containing inosine residues may serve as a structure proving suitable sites of chemical reaction. This may explain the observed finding that Poly (I·C), unlike Poly I·Poly C, is able to decrease the enzymatic appearance of recoverable BaP products.

In the case of RNA derived from yeast, results demonstrated no covalent binding between BaP and RNA sedimenting in a sucrose gradient whereas BaP was associated strongly with an RNA peak not sedimenting in the gradient. In this

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regard it appears that the low molecular weight transfer RNA might be expected to be the least likely of the various RNA species to sediment and, due to extensive base pairing in the native state (122), may represent a structure suitable for physical and chemical interaction with BaP.

On the basis of the above considerations, a suitable conformation of the BaP molecule within the native DNA structure may be required for chemical reaction. According to studies reported by Lesko et al. (95, 118) using a nonenzymatic model hydroxylating system, procedures designed to optimize physical binding between BaP and DNA proved favorable to increased reaction yield of chemical complex. Nevertheless, bifunctional alkylating agents, like HN₂, which are able to covalently cross link guanine residues on the DNA polymer (119), might be expected to physically prevent the suitable fitting of BaP within the DNA molecule and, in this manner, may explain, at least in part, the inability of DNA modified with HN₂ to substantially decrease the enzymatic appearance of BaP products observed in the present studies.

The above discussion stresses the apparent importance of considering physical interactions and secondary structure of nucleic acids in the mechanism of chemical reactions between BaP and polynucleotides and may be of fundamental significance to the observed specificity of such reactions.

As stated previously, various polynucleotides, notably native DNA, Poly G, Poly I and the alternating copolymer

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Poly (I·C), were found to decrease the enzymatic appearance of recoverable BaP products. In contrast, inclusion of denatured DNA and RNA in incubation mixtures enhanced the amount of recovered products. Poly G also was able to significantly increase the yield of product but, unlike denatured DNA and RNA, only under conditions of excessive enzyme concentration. According to Hayakawa and Udenfriend (123), observing similar activating effects of polynucleotides on the enzymatic appearance of BaP products as determined by a radioassay procedure, nucleic acids may act to promote product recovery by protecting the enzyme from inactivation by highly reactive BaP intermediates. In the present studies, denatured DNA and RNA may interact with enzyme and, in this manner, protect against enzyme inactivation by reactive BaP products. Although Poly G may act likewise, it should be noted that studies suggest covalent binding between Poly G and intermediate BaP products. An alternative possibility is that such polynucleotides stabilize the biotransformation process by protecting the AHH system from degradation by proteolytic enzymes present in microsomal preparations.

In other studies results revealed that polynucleotides (specifically native DNA, Poly G and Poly I), known to covalently bind BaP as a result of microsomal reactions, were unable to increase the enzyme-dependent disappearance of isolated fluorescent BaP products. These findings demonstrate that the chemical binding of the various polynucleotides with BaP is not the result of the interaction

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with fluorescent biotransformed BaP products and suggests that binding occurs as a result of interaction with a reactive BaP intermediate product.

Studies cited earlier (see Introduction) reported 3-hydroxy BaP to be the most significant product formed as a result of the BaP oxidation in vivo in the rat and in vitro by microsomal systems derived from rat tissues. This product apparently was present in substantial amounts in the green fluorescent BaP products isolated in the present research since activation and fluorescence spectra of isolated products and authentic 3-hydroxy BaP were found to be identical in NaOH solution (see legend to Figure 33), supporting similar findings reported earlier by Kuntzman (87). However, other oxidation products, notably 6-hydroxy BaP, 1,6-dihydroxy BaP and 3,6-dihydroxy BaP as well as various dihydrodiols ad quinones, result from microsome-dependent oxidation of BaP (64, 66, 124). As demonstrated in the present studies, the green fluorescence of isolated BaP products was found to disappear in appropriate incubation mixtures in a manner which suggests further conversion of products by the AHH system. It should be noted that of the various BaP products identified in microsomal oxidation reactions, only monohydroxylated products exhibit green fluorescence in alkaline solution after ultraviolet illumination (124). Possibly, monohydroxylated products can act as substrates in the enzymatic generation of other oxidized products. This may explain, on a more established basis than before, the origin of secondary oxidation products observed in the oxidation

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of BaP in vivo (57).

As early as 1950, Boyland proposed that reactive epoxide intermediates resulted from the metabolism of polycyclic aromatic compounds (125). Recently, studies have provided evidence that epoxides are oxidation products in the microsome-dependent oxidation of such polycyclic hydrocarbons as phenanthrene (128), benz(a)anthracene (BA) (128), dibenz(a,h)anthracene (DBA) (127,128), 7,12-dimethylbenz(a)anthracene (DMBA) (130) and, most recently, pyrene and BaP (129). Typically, polycyclic aromatic epoxides rearrange to phenols, give rise to related dihydrodiols in the presence of microsomal epoxide hydrase and conjugate glutathione in the presence of soluble epoxy glutathione transferase (131). The latter enzymes are found in rat liver tissue (132,133). Further, epoxides, such as those derived from phenanthrene, 7-methylbenz(a)anthracene (MBA), BA and DBA, are reported to chemically react with Poly G, less so with Poly A and apparently not at all with Poly U and Poly C (130). Generally, polycyclic aromatic epoxides appear capable of covalently binding DNA both chemically (in the absence of microsomes) (137) and in cell culture (138, 139). In the case of BaP, the 4,5-oxide reportedly has been detected as the result of microsomal reaction (129). The epoxide was reported to rearrange to acid to 4-hydroxy baP, react in the presence of epoxide hydrase to form BaP 4,5-dihydrodiol and react with glutathione to yield conjugates. Moreover, BaP 4,5-oxide reportedly was reactive toward Poly G (129) and, very recently, has been suggested as being involved

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in chemical reactions with DNA as a results of microsome-dependent conversion of BaP (135).

Besides BaP 4,5-dihydrodiol, such vicinal dihydrodiols as BaP 1,2-diol, BaP 9,10-diol and BaP 11,12-diol have been reported as products of rat liver microsomal oxidation and used as evidence for the intermediacy of the corresponding epoxides (111,112). Reported detection of various phenols, including 1-hydroxy BaP, 5-hydroxy BaP and 12-hydroxy BaP, as a result of microsomal BaP oxidation, might be used to further support the formation of epoxide intermediates in these reactions. However, substantive evidence supporting the involvement of an epoxide intermediate in the formation of the major oxidation product, 3-hydroxy BaP (i.e., detection of BaP 2,3 oxide or BaP 2,3-dihydrodiol) remain unreported. Based on the ability of BaP to form radical cations in nonenzymatic model hydroxylating systems (118) and evidence apparently suggesting the generation of electrophilic center at position 3 of the BaP molecule as a result of electrophilic attack of position 6 by a positive oxygen atom produced by the AHH system (134), an intermediate of 3-hydroxy BaP formed enzymatically could be a positively charged species directly generated by the AHH system. Such an intermediate would be expected to react with water to form the 3-hydroxylated product but would not be expected to yield 2,3-dihydrodiols. Nevertheless, Sims et al. (129) maintains as doubtful that such products as 3-hydroxy BaP and 6-hydroxy BaP “arise by a mechanism that does not involve the initial formation of an epoxide)” (the

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exact nature of an epoxide at position 6 is left unexplained) and explains the absence of related dihydrodiols to be the result of a more rapid rate of epoxide rearrangement to phenols than the rate of microsome-dependent hydration to dihydodiols. In any case, the covalent binding between BaP and polynucleotides demonstrated in the present studies might be related to a reaction involving either a BaP epoxide or a BaP radical cation. It should be recalled that results derived from the present research suggest the involvement of a BaP intermediate in polynucleiotide binding reactions.

Other studies in the present research attempted to determine the biological significance of DNA treated with BaP in the presence of the AHH system. This involved a comparison of the abilities of DNA treated with BaP in the presence of the AHH system and various DNA controls to serve as templates in a DNA-dependent RNA polymerase system. The template ability of DNA treated with the noncarcinogenic BeP also was compared to these studies. It should be noted that no physical, chemical or biological properties of BaP-treated DNA derived from microsomal reactions have been reported earlier and may be related to the relatively low extent of binding so far achieved (approximately one BaP molecule per 50,000 nucleotide bases) (24). Use of the template approach might be expected to provide substantive evidence for chemical alteration of DNA by BaP since changes in the template ability of DNA has been shown to be a very sensitive indicator of chemical modification (136).

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The results of template studies demonstrate that template activities of DNA treated with either BaP or BeP are decreased (and to a similar extent) when compared to template activities of DNA controls. These findings provide evidence that DNA is chemically altered by BaP and BeP as the result of microsomal reactions. More importantly, these findings suggest that DNA, treated enzymatically with benzopyrenes, is biologically modified as determined by studies involving a model transcription system, but in a manner which does not appear to be carcinogen-specific.

Summarizing the major findings of the present research, BaP apparently can biologically and chemically modify mammalian nucleic acids in the presence of a mixed-function oxidase system found in the rat and human tissues in a reaction which appears to involve an enzyme generated BaP intermediate product, to be dependent on the secondary structure of nucleic acids and to be base-specific. Overall, then the in vitro studies described herein appear to be consistent with the general hypothesis that initiation of carcinogenic processes in mammalian systems in vivo can be a result of chemical reactions between biologically activated chemical carcinogens and critical sites on cellular macromolecules.

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"INTERACTION OF BENZO (α) PYRENE AND NUCLEIC ACIDS IN THE PRESENCE OF A MIXED-FUNCTION OXIDASE SYSTEM"
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USA – STATE UNIVERSITY OF NEW YORK AT BUFFALO (SUNYAB) – 1973 (ProQuest 302781013; ISBN 979-8661021359)

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