OCS Study 

MMS 95-0020 

University Research Initiative 

Bioavailability and Genotoxicity 

of Produced Water Discharges 

Associated with Offshore 

Production Operations 

-, 

LOUISIANA 

c MISSISSIPPI I 

L~_--- 

~ ALA 

TEXAS ( Baton Rouge ~ ~ Mobile 

1 

New Orleans 

- I : ; . . . . 

alveston 

LUMCON Corpus . . 

Christi. ~ 

. . ., . . . . . . . . . 

. . . . . . loom . . , . . . . . . . . . 

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. . . . . . . . .. . . . . . . . . . . . . . . . . . 300m . . . . . 

`. . . . . .,lpppm ~ 

. . . . 

Gulf of Mexico 

U.S. Department of the Interior ,,tom. cooperative Agreement AM Minerals 1management Service Lumton University Research Initiative 

Gulf of Mexico OCS Region Louisiana Universities Marine Consortium 
OCS Study 

MMS 95-0020 

University Research Initiative 

Bioavailability and Genotoxicity 

of Produced Water Discharges 

Associated with Offshore 

Production Operations 

Authors 

Gary W. Winston 

Department of Biochemistry 

322 Choppin Hill 

Louisiana State University 

Baton Rouge, Louisiana 

and 

Jay C . Means 

Department of Veterinary Physiology, 

Toxicology and Pharmacology 

Louisiana State University 

Baton Rouge, Louisiana 

April 1995 

Prepared under MMS Contract 

14-35-0001-30470 

by 

Louisiana Universities Marine Consortium 

8124 Highway 56 

Chauvin, Louisiana 70344 

Published by 

U.S. Department of the Interior Cooperative Agreement 

Minerals Management Service University Research Initiative 

Gulf of Mexico OCS Region Louisiana Universities Marine Consortium 
111 

DISCLAIMER 

This report was prepared under contract between the Minerals Masnagement and the Louisiana 

Universities Marine Consortium (LUMCON). This report has been technically reviewed by the 

MMS and approved for publication. Approval does not signify that the contents necessarily reflect 

the views and policies of the Service, nor does the mention of trade names or commercial products 

constitute endorsement or recommendation for use. It is, however, exempt from review and 

complicance with MMS editorial standards. 

REPORT AVAILABILITY 

Extra copies of the report may be obtained from the Public Information Unit (Mail Stop 5034) at 

the following address: 

U.S . Department of the Interior 

Minerals Management Service 

Gulf of Mexico OCS Region 

Public Information Unit (MS 5034) 

1201 Elmwood Park Boulevard 

New Orleans, Louisiana 70123-2394 

Telephone Number: (504) 736-2519 

1-800-200-GULF 

CITATION 

Winston, G. W. and J. C. Means. 1994. Bioavailability and Genotoxicity of Produced Water 

Discharges Associated with Offshore Drilling Operations . A final report by the 

Louisiana Universities Marine Consortium for the U.S . Department of the Interior, 

Minerals Management Service, Gulf of Mexico OCS Region, New Orleans, Louisiana. 

OCS Study MMS 95-0020. 124 pp. 
1V 

CONTRIBUTORS 

Susan B. Dobias, Department of Biochemistry, Lousiana State University, Baton Rouge, 

Louisiana 70803 

Debra J. McMillin, Department of Veterinary Physiology, Toxicology and Pharmacology, School 

of Veterinary Medicine, Louisiana State University, Baton Rouge, Lousiana 70803 

Jay C. Means, Department of Veterinary Physiology, Toxicology and 

Pharmacology, School of Veterinary Medicine, Louisiana State University, Baton 

Rouge, Lousiana 70803 

Roland Pautz, Department of Biochemistry, Lousiana State University, Baton Rouge, Louisiana 

70803 

Lawrence A. Reily, Department of Veterinary Physiology, Toxicology and 

Pharmacology, School of Veterinary Medicine, Louisiana State University, 

Baton Rouge, Lousiana 70803 

Karen L. Sarradet, Department of Veterinary Physiology, Toxicology and 

Pharmacology, School of Veterinary Medicine, Louisiana State University, Baton 

Rouge, Lousiana 70803 

Gary W. Winston, Department of Biochemistry, Lousiana State University, Baton Rouge, 

Louisiana 70803 

Zuo-Mei Yan, Department of Biochemistry, Lousiana State University, Baton Rouge, Louisiana 

70803 
V 

ABSTRACT 

Discharges from offshore oil and gas production are drilling fluids, drill cuttings, and 

produced water. In coastal regions, effluents are discharged into fresh or brackish marshes or 

wetlands which are spawning sites for many economically important fish and shellfish. The 

assessment of ecological health impact of marine and estuarine pollution require sound technology 

for monitoring and assessing risk of pollutant impacts. Field-exposed organisms yielded much 

lower ethoxyresorufin 0-deethylase activities (EROD) than experimentally induced levels reported 

in microsomes of other fish species. Clear patterns of EROD induction were not evident in fish 

sampled from contaminated areas. EROD was highly variable in liver microsomes of all fish, 

regardless of the area from which they were sampled, and in some cases showed overlap between 

responses in control and impacted areas. Obvious conelations were not evident between 

antioxidant enzyme activity and distance from point source offshore dischage, contaminated coastal 

sites vs. reference sites, species, collection time, depth of habitat, sediment-dwelling fish vs. 

demersal fish and between offshore sites. Microsomal P450 was found to be mostly denatured or 

highly contaminated with other interfering CO-binding chromaphores . EROD, P450 reductase and 

CO-binding data should be reconciled before tenable conclusions of the effects of pollutant outfall 

can be made. Mutagen mobilization into co-solvent extracts was detectable by the Salmonella 

ryphimurium assay, with tester strains varying in sensitivity. Promutagens were detectable in all 

extracts, via activation with Arochlor-induced rat liver S9. Mutagenicity varied among co-solvent 

concentrations, perhaps reflecting mobilization of different arrays of compounds. Mutagens were 

not detected with the umu gene induction assay, which uses S. ryphimurium 1535/psk1002, 

whether assays were conducted in the presence or absence of fish liver S9 or microsomes. Isomer 

specific analyses of bioaccumulated petroleum aromatic hydrocarbons in tissues of aquatic 

organisms indicate the relative bioavailability of these contaminants, which may enter biological 

foodwebs in aquatic systems . This suggests that contaminated sediments remain as bioavailable 

toxic substances even after discharges are suspended . Our data confirm recognized problems in 

biomarker research; fish from large areas display temporal and spatial variability ; biochemical 

responses of fishes are variable as a function of species, sex, gonadal status, season, temperature, 

and nutrition. 

ACKNOWLEDGEMENTS 

The authors gratefully acknowledge the assistance of the Louisiana State University 

Department of Biochemistry office staff for their expert assistance in typing and formatting this 

report. 
TABLE OF CONTENTS 

V11 

Executive Summary. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .1 

Chapter 1 . Induction of EROD in Nereis After Exposure to Contaminated Sediment. 

Four Independent Experiments With Time and Concentration Variables. 

Western Blot Studies of Inducted CYP Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 

Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 

Chapter 2. Induction of Ethoxyresorufin O-Deethylase (EROD) in the Killifish 

(Fundulus Grandis) by Selected Mutagens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 

Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 

Chapter 3. Field Investigations of the Induction of EROD in Fish at St 34 and St 52 

Platforms in the Gulf of Mexico, Pass Fourchon, and Lake Champagne . . 21 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 

Materials and Methods. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 

Chapter 4. Studies of Pro- and Antioxidant Parameters in Liver Cytosol and 

Microsomes of Gulf of Mexico Fish Samples and Other Marine 

Organisms: Evaluation of Superoxide Dismutase, Catalase, Microsomal 

Generation of Oxyradicals, and Oxygen Consumption . . . . . . . . . . . . . . . . . . . . . . . 33 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 

Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 

Chapter 5. Studies of NADH- And NADPH-Dependent Microsomal Electron 

Transport: Relationship of Cytochrome C Reductase and Oxygen 

Consumption to Species and Environmental Conditions of Gulf of 

Mexico Field Sites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 

Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 

Chapter 6: Part A Evaluation of the Bioconcentrarion Potential of Genotoxic Petroleum 

Industry Effluents Using Co-Solvent Mobilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 

Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 

Experimental Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 
viii TABLE OF CONTENTS 

(continued) 

Chapter 6: Part B Studies of Genotoxicity and Potential for Mutagenic Activation in 

Fish Sampled From the Gulf of Mexico as Evaluated by the Umu 

Mutagenicity Assay. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 

Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 

Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 

Chapter 7 . Development and Application of (Parts-Per-Trillion) GUMS Analysis 

Methods for Approximately 60 Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . 77 

Analytical Method Development. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 

Application of Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 

Chapter 8. Investigation of the Bioaccumularion Potential of 60 Individual 

Aromatic Petroleum-Associated Hydrocarbons in the Oyster 

Crassostrea virginica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 

Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 1 

Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .100 

Literature Cited . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .103 

Appendix A Parent and alkylated PAH concentrations (ng/g) in oyster tissue from 

organisms exposed to varying levels of contaminated Pass Fourchon 

sediment for 28 days. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .111 
LIST OF FIGURES ix 

1 Map of samplings sites at Pass Fourchon, LA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 

2 Quantitative analysis of petroleum hydrocarbon contaminants in Pass 

Fourchon sediment collected at PF 400 site, 400 m from produced 

water discharge. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .10 

3 EROD induction in Nereis virens by exposure to petroleumcontaminated sediment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 

4 Western Blot analysis of Nereis intestinal microsomes probed with 

trout anti-CYP lAl . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 

Sa EROD induction in Fundulus grandis larvae in independent replicate 

experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 

Sb EROD induction in Fundulus grandis adults in independent replicate 

experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 

Sc EROD induction in Fundulus grandis embryos in independent replicate 

experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 

6 Induction of EROD in three species of flatfish near offshore 

production facilities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 

7 EROD Activity in Sea Robin as a function of distance from ST 34. . . . . . . . . . 29 

8 Typical EROD standard curve . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 

9 EROD vs. microsomal protein concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 

10 Nitroaromatic-stimulated oxyradical production by sea anemone 

microsomes : effect of cadmium preexposure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36 

11 Detection of purified SOD by SOD-525 assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38 

12 P450 CO-binding spectrum of fish #301 liver microsomes . . . . . . . . . . . . . . . . . . 46 

13 Cytochrome P450 CO-binding spectrum in microsomes of PCBinduced rats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 

14 Polaragraphic tracings of 4NQ0-mediated oxygen consumption 

during microsomal electron transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 

15 Comparison of NADH-dependent oxygen consumption catalyzed by 

LC vs PF catfish: Effect of varying microsomal protein microsomes 

and 4NQ0 concentrations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .50 

16 Relationship of Log P to Log Koc and Log BCF. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 
x LIST OF FIGURES 

(continued) 

17 Effect of mole fraction of three co-solvents on desorption Koc values 

for hexachlorobiphenyl . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .57 

18 Effect of mole fraction acetic acid on Koc for hexachlorobiphenyl . . . . . . . . . . 58 

19 Effect of acetic acid co-solvent concentration upon the Koc of three 

model hydrophobic compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 

20 Co-solvent mobilization of selected pollutants in a contaminated 

sediment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 

21 Dose response of mutagens extracted from sediments . . . . . . . . . . . . . . . . . . . . . . . . 64 

22 Reversion frequencies for direct-acting mutagens extracted at various 

co-solvent fractions from contaminated sediment (TA 102) . . . . . . . . . . . . . . . . . . 65 

23 Reversion frequencies for promutagens extracted at various 

co-solvent fractions from contaminated sediment (TA 97a) . . . . . . . . . . . . . . . . . . 66 

24 Reversion frequencies for promutagens extracted at various 

co-solvent fractions from contaminated sediment (TA 100) . . . . . . . . . . . . . . . . . . 67 

25 Reversion frequencies for promutagens extracted at various 

co-solvent fractions from contaminated sediment (TA 102) . . . . . . . . . . . . . . . . . . 68 

26 2-aminoanthracene activation by liver microsomes of Pass Fourchon 

and Lake Champagne catfish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 

27 Effect of fish liver cytosol on umu gene induction by 

2-aminofluorene and 2-acetyl aminofluorene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 

28 Arylamine activation by fish liver cytosol. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 

29 Concentrations (ppb) of high molecular weight PAH bioaccumulated at 

3 dose levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 

30 Concentrations (ppb) of low molecular PAH bioaccumulated at 4 

dose levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 

31 Concentrations (ppb) of C3-phenanthrenes bioaccumulated at 4 

dose levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 

32 Concentrations (ppb) of C2-phenanthrenes bioaccumulated at 4 

dose levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .101 

33 Concentrations (ppb) of C2-dibenzothiophenes bioaccumulated at 4 

dose levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .102 
X1 

LIST OF TABLES 

TABLE DESCRIPTION PAGE 

3-1 Sediment data for Pass Fourchon and reference site . . . . . . . . . . . . . . . . . . . . . . . . . . . 24 

3-2 Hepatic microsomal EROD levels in specimens from nearshore sites at 

Pass Fourchon and Lake Champagne, LA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 

3-3 Hepatic microsomal EROD levels in fish collected by LDWF near 

offshore platforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 

3-4 Liver microsomal EROD activities of Gulf of Mexico fish samples . . . . . . . . . 30 

3-5 Liver microsomal EROD activities offish species sampled from a 

contaminated (Pass Fourchon) and reference (Lake Champagne) 

coastal site in the Gulf of Mexico . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 

4-1 SOD content of sea anemone (Anthapleura elegantissima) cytosols: 

Effect of 3-methylcholanthrene (3MC) exposure and zooxanthellate 

symbionts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 

4-2 SOD content of Gut Cytosol from Neriei virens exposed to 

contaminated sediment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37 

4-3 Microsomal SOD content of Gulf of Mexico fish samples . . . . . . . . . . . . . . . . . . . . 37 

4-4 Superoxide dimutase activities of liver cytosol from fish sampled from 

various Gulf of Mexico field sites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 

4-5 Specific activities of Catalase in Gulf of Mexico fish liver microsomes. . . . . 40 

4-6 Specific activities of Catalase in Gulf of Mexico fish liver cytosol . . . . . . . . . . 41 

5-1 Microsomal NAD(P)H-dependent cytochrome c reductase activities of 

several fish species sampled from a contaminated (Pass Fourchon) and 

reference (Lake Champagne) site in the Gulf of Mexico . . . . . . . . . . . . . . . . . . . . . 44 

5-2 Microsomal NAD(P)H-dependent cytochrome c reductase activities of 

several fish species sampled from varying distances from offshore oil 

platforms in the Gulf of Mexico . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 

5-3 Cytochrome P450 specific content of sampled Gulf of Mexico fish . . . . . . . . 47 

6A-1 Priority pollutants commonly detected in treated petroleum industry 

effluents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 
xii LIST OF TABLES 

(continued) 

TABLE DESCRIPTION PA GE 

7-1 Selected aromatic hydrocarbon analytes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 

7-2 Summary of regression statistics for two standard curves . . . . . . . . . . . . . . . . . . . . . 84 

7-3 Limit of detection, limit of quanritation, sensitivity values calculated 

from standard deviations of standard curves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 

7-4 Method detection limits for three sample matrices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 

7-5 Percent recovery of analytes spiked at four concentrations . . . . . . . . . . . . . . . . . . . . 87 

7-6 Historical data for sediments from the Pass Fourchon 400m 

(from discharge) sampling site . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 

7-7 Application of analytical methods to aqueous and particulate phases 

of water and sediments in coastal waters near Pass Fourchon . . . . . . . . . . . . . . . . 89 

8-1 Mortality data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 

8-2 Sediment data for Pass Fourchon and reference site . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 
EXECUTIVE SUMMARY 

The major waste streams associated with offshore oil and gas production are drilling 

fluids, drill cuttings, and produced water. The ecological and human health hazards of marine 

and estuarine pollution make it imperative to seek new and improved technology for monitoring 

and risk assessment of pollutant impacts. This report presents results of laboratory and field 

studies directed by the co-principle investigators. 

Laboratory studies of the effects of petroleum-contaminated sediments on 

ethoxyresorufin O-deethylase (EROD) activities the marine worm, Nereis virens, have been 

conducted. Results of initial experiments indicated induction of EROD activity in N. virens, 

exposed to petroleum-contaminated sediment from Pass Fourchon, the site of a major separation 

facility for both the coastal and offshore oil and gas operations in Louisiana . Worms were 

embedded in 20% and 50% (w/w) sediment mixtures for 7 and 14 days. EROD activity was 

detected in microsomal fractions extracted from excised gut tissue. EROD activity in worms 

exposed in 20% sediment was -14-fold higher than controls after 7 days; after 14 days of 

exposure the increase in EROD was -20-fold over controls. When exposed to 50% sediment a 

27-fold induction of EROD was observed after 7 days, and after 14 days induction was -32-

fold. For unexplainable reasons these data could not be reproduced in two subsequent 

experiments. However, EROD activity in invertebrates is traditionally difficult to quantify. N. 

virens exhibited EROD induction in gut tissues, when exposed to benzo(a)pyrene via food (Fries 

and Lee, 1984), thus the data need to be further reconciled. Western blot analysis performed on 

the worm gut microsomes probed with rabbit anti-trout CYP1A1 polyclonal antibody did not 

indicate the presence of proteins with common epitopes of CYP lA proteins. 

Laboratory challenge experiments were conducted with three mutagenic agents, to 

evaluate the responsiveness of EROD induction in adults, larvae, and embryos of the killifish. 

Cyclophosphamide (CP) and ethylmethanesulfonate (EMS) are two benchmark mutagens with 

known genotoxic effects on Fundulus . ; 3-methylcholanthrene (3-MC) is a classical promutagen, MFO inducer and model carcinogen in mammalian studies. EROD activity was 

detected in microsomes from whole embryos, larvae, or adult livers . The mean EROD activity 

in control adult livers was 0 .08 nmol resorufin produced min/mg protein. EROD was not 

detected in control embryos or larvae . 3-MC exposure induced EROD to 2.8, 1 .9, and 2.2 

nmol/min/mg in embryos, larvae, and adults, respectively . EMS induced EROD to 1 .50, 1 .60, 

and 0.57 nmol/min/mg. CP induced to 1 .25, 1 .20, and 0.18 nmol/min/mg. 

Field validation of the EROD assay as a biomarker in teleost fish may provide a sensitive 

index of environmental pollution, to detect exposure history and to qualitatively establish the 

bioavailability of dissolved and sediment-bound aromatic hydrocarbons. Bottom-dwelling 

fishes were periodically collected from offshore petroleum production platforms. S9 and 

microsomal fractions were prepared from pooled or individual livers, and assayed for EROD 

activity . EROD induction was not observed in S9 of bay whiff (Citharichthys spilopterus) ; 

levels in fringed flounder (Etropus crossotus) and shoal flounder (Syacium gunteri) ranged 

between 0.4-5 .2 (mean = 2.9) pmol resorufin produced /min/mg S9 protein and 0.9-9 .1 (mean 

= 3.6), respectively . Field exposed fishes yielded much lower EROD values than in 

microsomes from experimentally induced fish reported in the literature . S9 preparations were 

prone to inhibitory effects and a larger base of field data is needed to validate the use of EROD 

for detecting exposure history and qualitatively demonstrating bioavailability of dissolved and 

sediment-bound chemicals. Collecting trips to offshore and coastal marsh areas yielded fish 

specimens from both contaminated and control (pristine) locations. Pristine locations were 

defined as those which contained less than 50 ppb total aromatic hydrocarbons as measured by 

gas chromatography/mass spectrometry . Generally, microsomes from the livers of both control 

fish and fish from contaminated sites showed very low or no detectable EROD activity. 
2 

Clear patterns of EROD induction are not evident in fish sampled from contaminated 

areas. Liver microsomes of all fish, regardless of the area from which they were sampled 

exhibited highly variable EROD activities, which in some cases showed overlap. It is difficult to 

draw inferences from small samplings, but these data do point to some recognized problems in 

biomarker research . For example, fish from large areas display temporal and spatial variability; 

biochemical responses of fishes are variable as a function of species, sex, gonadal status, 

season, temperature, and nutrition. Thus, field studies can result in a disconcerting array of 

data, even though controlled laboratory studies are very predictable. An objective of biomarker 

research is to establish the efficacy of the use of the sentinel and this is the stage of this research . 

Oxygen radicals (ROS) are produced as a normal part of all aerobic life . Many metals 

and organic chemicals increase ROS production in biological systems with accompanying 

oxidative damage. The toxicities of these chemicals are linked to enhanced generation of ROS. 

The consequences of enhanced ROS production are protein degradation, enzyme inactivation, 

lipid peroxidation, DNA damage and cell death. Superoxide dismutase (SOD) and catalase 

measurements of fish liver cytosol were conducted. SOD activities expressed as units/ mg 

cytosolic protein ranged from less than 1 or below detection limits in Fringed flounder taken 

from 200 m of ST 53a to 530 units/mg in Shoal flounder caught 500 m from ST 34. Fringed 

flounder sampled from 200 m of ST 34 had one of the higher SOD levels . In the few cases 

where antioxidant enzymes were shown to correlate at all with distance from a platform an 

inverse relationship was observed. The significance of this relationship might relate to depletion 

of antioxidant defenses under conditions of high xenobiotic exposure. Obvious correlations 

were not evident between catalase activity and distance from point sources of offshore 

contamination, contaminated coastal sites vs. reference sites, species, collection time, depth of 

habitat, sediment-dwelling fish vs. demersal fish and one offshore site vs another. 

Aside from the heme-containing cytochromes P450 and bs there are also peripheral 

flavoproteins, which serve as reductases for the cytochromes . These reductases are held to the 

membrane by a hydrophobic binding domain and they contain the prosthetic groups FMN and 

FAD. Microsomal flavoprotein reductases are rate-limiting components of NAD(P)H-dependent 

microsomal mixed function oxygenase activities and also extremely active loci of oxyradical 

production . There are relatively few studies of flavoprotein reductases on submammalian 

species. Thus, we have studied both NADH- and NADPH-dependent cytochrome c reduction to 

indicate the infra and inter species range of activities and the effects of various Gulf of Mexico 

field site environments on those activities . In conjunction with reductase activities, attempts 

were made to measure the specific content of cytochromes P450 in an effort to determine the 

efficacy of the microsomal preparations and for use in the evaluation of the MFO studies 

conducted in association with this project. NAD(P)H-dependent cytochrome c reductase 

activities were measured in liver microsomes of fish sampled from various offshore sites. These 

sites were further subdivided as a function of distance from the platform. NADH-dependent 

cytochrome c reductase ranged from below detectable limits in Bay Whiff caught at ST 52 to 

approximately 1000 nmol/min/mg microsomal protein in catfish caught at ST 34 (Table 5 .1) . 

NADH-dependent activities were consistently higher in catfish (870±170 nmol/min/mg, n = 7), 

irrespective of the distance from the platform at which they were caught. Fringed flounder had 

the next highest activities with NADH (83±44 nmol/min/mg, n = 6). Fringed flounder from ST 

36, 37, 52, 53, and 54 had notably lower NADH-cytochrome c reductase activities than those 

from ST 53a or any of those caught from ST 34. NADPH-dependent activities ranged from 0.2 

nmol/min/mg in Bay Whiff caught at ST 53 and 55 to 300 nmol/min/mg in catfish caught at ST 

34. An unexpected observation was the NADH/NADPH ratios of < 1 .0 found in some of the 

Fringed flounder and Bay Whiff microsomes from the most recently provided samples. NADHdependent activites are generally greater than NADPH- dependent activites. The specific content 

of cytochromes P450 were determined in all fish samples for which we were supplied with 

adequate amounts of microsomal protein. In all cases microsomal P450 was found to be mostly 

denatured to cytochrome P420 or highly contaminated with other interfering CO-binding 
3 

chromaphores. EROD, P450 reductase and CO-binding data should be reconciled before tenable 

conclusions of the effects of pollutant outfall can be made. 

The high degrees of variability among the different species, the lack of correlation 

between MFO and field parameters, and the denaturation of MFO catalyst in microsomal 

preparations are all contrary to laboratory conducted exposure studies. Additional major 

problems were the inconsistency of the catches, which were unpredictable with respect to 

species, lack of knowledge of the sex and gonadal status of the fish at the times in which they 

were caught and variable conditions of abiotic factors such as fluctuations in oxygen tension. It 

is not known to what extent Hurricane Andrew perturbed zones of delineation m temperature, 

salinity, partial oxygen pressures, sediment-water interfaces, etc. Hurricane Andrew's impact 

on these abiotic factors could conceivably have resulted in stresses to the organisms which were 

reflected in the biochemical and physiological homeostasis of the sampled organisms. 

Petroleum wastes from produced water separation facilities, marketing terminals and 

production facilities contain a complex array of normal and branch-chained hydrocarbons, 

volatile (benzene, toluene, xylenes and ethyl benzene) and semi-volatile (naphthalenes, 

phenanthrenes, fluorenes, etc .) parent aromatic hydrocarbons, alkylated analogs of these 

aromatic compounds and various heterocyclic aromatic compounds, e.g. dibenzothiophenes and 

aza-arenes. Many of these compounds are lipophilic and thus, tend to bioaccumulate in aquatic 

food chains. To provide a more realistic assessment of genotoxic risk to aquatic organisms 

studies were conducted to evaluate the utility of co-solvent extracts from sediments in a way that 

can be related to bioavailability of contaminants. These studies have yielded the following: 

direct-acting mutagens were mobilized at all co-solvent concentrations but were detected only 

with strain TA 102. Mutagen mobilization into co-solvent extracts was detectable by the 

Salmonella ryphimurium assay, with tester strains varying in sensitivity . Promutagens were 

detectable in all extracts, via activation with Arochlor-induced rat liver S9. Mutagenicity effects 

varied among co-solvent concentrations, perhaps reflecting mobilization of different arrays of 

compounds. Low co-solvent fractions enhanced extraction of water-soluble mutagens but 

suppressed extraction of lipophilic mutagens. In high co-solvent fractions, extraction of 

lipophilic mutagens was significantly increased. Increasing co-solvent fraction appeared to 

suppress the extraction of some water-soluble mutagens. TA 102 was the most responsive tester 

strain with these extracts and yielded very reproducible results . TA 98 showed no response to 

mutagens in these extracts. Attempts to detect mutagens with the umu gene induction assay, 

which uses S . typhimurium 1535/psk1002 did not detect mutagens whether assays were 

conducted in the presence or absence of fish liver S9 or microsomes. Chemical analyses of the 

co-solvent extracts by GC/MS may yield information on the identity of possible mutagens 

present. 

The effects of sediment exposures on the indigenous organism's ability to modulate the 

mutagenicity of the benchmark promutagens 2-aminoanthracene (2AA) and 2-

acetylaminofluorene (2AAF) was investigated by the umu gene induction assay. The ability of 

microsomes from Lake Champagne catfish and Pass Fourchon catfish to activate 100 mM 2AA 

was compared with that of liver microsomes from control rats and from rats pre-treated with 3-

methylcholanthrene (3MC), an aromatic hydrocarbon known to induce enzymes that activate 

these promutagens in the rat . The rat samples were run with 200 mg of microsomal protein and 

the fish with 50, 100 and 200 mg of protein. The results showed a 1 .6-fold and a 2.4-fold 

enhancement of umu gene induction with microsomes from control and 3MC-treated rats, 

respectively, but no increase was observed over background controls with the fish samples. Rat 

liver cytosol contains amine oxidases which are capable of activating certain arylamines to 

mutagens in the Ames mutagenicity assay. Experiments were conducted with cytosolic fractions 

from Atlantic Croaker (Micropogon undulatus) (Pass Fourchon), Threadfin Shad (Dorosoma 

petense) (Lake Champagne) and Pinfish (Lake Champagne) liver to determine if fish liver 

cytosolic fractions catalyzed arylamine activation in the umu mutagenicity assay and if so, was 
4 

such catalysis altered in fish from contaminated sites. Cytosolic protein was tested separately at 

400 mg and 800 mg; no significant increase in gene induction was observed over background . 

An objective of biomonitoring studies is to determine amounts of bioconcentratable 

organics present in petroleum-related waste streams and receiving environments . Several criteria 

for the methods must be met, including: a wide dynamic range (0.01 ppb to 100 ppm)-7 orders 

of magnitude in concentration; compound specificity-discriminaton of individaul compounds 

from biological and geological matrices; compound quantitation- ability to determine 

concentrations of specific compounds; sufficient "fingerprinting" capability to discriminate 

between multiple sources of similar contaminants. Methods should allow for frequent sampling 

and analysis and a minimum of procedural steps to obviate against losses or contamination . 

Sediment bound contaminants associated with chronic produced water discharges which 

accumulate over time are bioavailable to the filter-feeding bivalve Crassostrea virginica. 

Accumulation was observed in three days of exposure and increased toward steady-state for 21 

days or more. Petroleum-associated aromatic hydrocarbons increased in all experiments in a 

dose- and time-dependent fashion . Mortalities observed in higher-dose groups appeared to be 

contaminant-related. Some effects on accumulation of the compromised physiological state of 

toxicant-stressed oysters were observed in the highest dose group at 14 days. Isomer-specific 

analyses of bioaccumulated petroleum aromatic hydrocarbons in tissues of aquatic organisms 

yields information on the relative bioavailability of these contaminants, which rapidly enter 

biological foodwebs in aquatic systems. This suggests that contaminated sediments remain as 

bioavailable toxic substances even after discharges are suspended. Since some of these 

foodwebs potentially lead directly to man, there may be both ecological and human health risks 

associated with in-place, sediment-bound contamination from produced waters. 
5 

CHAPTER 1 

INDUCTION OF EROD IN NEREIS AFTER EXPOSURE TO CONTAMINATED 

SEDIMENT. 

FOUR INDEPENDENT EXPERIMENTS WITH TIME AND CONCENTRATION 

VARIABLES. 

WESTERN BLOT STUDIES OF INDUCED CYP ISOFORMS. 

L.A. Reily, G.W. Winston, J.C. Means, Z.-M. Yan and S.B. Dobias 

INTRODUCTION 

The aquatic environment is continually being challenged by pollutants, with oil spills and 

effluent discharges representing major sources of estuarine and marine pollution . Oil drilling 

operations generate very large volumes of effluent water. During the production of oil and gas, 

water trapped within permeable, petroleum-bearing sedimentary rocks is brought to the surface, 

carrying with it crude oil, drilling fluids, and various geological materials (Boesch and Rabalais, 

1985; Daniels et al., 1990) . This water, called produced water, is composed of a complex 

mixture of sub-surface water (formation water) and solid and liquid geological materials, 

including crude oil, and treatment chemicals, and may contain elevated levels of inorganic (trace 

metals) and organic (petroleum hydrocarbons) substances of a potentially toxic nature. Major 

waste streams of drilling fluids, drill cuttings, and produced water are associated with offshore 

oil and gas production. In Louisisana, produced waters were historically discharged into fresh 

or brackish marshes or wetlands, which also serve as spawning sites for a wide variety of 

species of fish and shellfish . Acute and chronic releases of petroleum hydrocarbons and other 

xenobiotic substances into surface waters affect biota and disrupt entire ecosystems. Moreover, 

bioavailability of hazardous compounds in aquatic environments increasingly presents a direct 

threat to man via seafood consumption . The expanding ecological and human health hazards of 

marine and estuarine pollution make it imperative to to seek new and improved technology for 

monitoring and risk assessment of pollutant impacts . 

Biological monitoring methods assess toxicity of xenobiotic compounds, synergistic 

interactions of multiple compounds, and, moreover, involve components of uptake and 

accumulation phenomena. Biomonitoring regimens yield assessments of toxicant impact which 

are quite tangible. 

Certain polychaete species are the dominant individuals in areas of oil spills, refinery 

effluents, oil field brine effluents, and natural oil seeps (Reish, 1971 ; Baker, 1976; Armstrong et 

al ., 1979 ; Spies et al ., 1979 ; Sanders et al ., 1980). Though reduced in numbers after oil spills, 

species such as Arenicola marina and Nereis diversicolor reestablish in the sediments in spite of 

the continued presence of oil (Prouse and Gordon, 1976; Levell, 1976; Baker, 1976). 

Exposure of vertebrates to benzo(a)pyrene or other natural or synthetic inducers causes 

increases in MFO activity and cytochrome P-450 content, and produces new cytochrome P-450 

(Lu et al ., 1973; Philpot and Bend, 1975 ; Bend et al., 1979; Johnson, 1979; Corbett et al ., 

1982; Stegeman, 1980; Payne et al., 1987). The properties of microsomal enzyme systems in 
6 

marine invertebrates have been extensively reviewed (Stegeman, 1981 ; Lee, 1981 ; Lee et al ., 

1983; James, 1989 ; Livingstone, 1989). The cytochrome P-450 mediated mixed-function 

oxygenase (MFO) system is responsible for the oxidation of a variety of foreign organic 

compounds. Lee (1981) reported that the sandworm, Nereis virens possesses a mixed function 

oxygenase system (MFO) which may provide a useful biological indicator of xenobiotic 

challenge. Evolutionary impetus for the MFO system may be to facilitate the elimination of 

foreign compounds, by converting them to more water-soluble metabolites (Lee, 1981) . This 

apparently enhances the individual's ability to eliminate them. Animals with active hydrocarbon 

metabolism accumulate lower levels of parent PAHs, and the higher rates of metabolism 

associated with P450 induction can enhance the rates of PAH elimination (Lu et al ., 1977) . 

However, the patterns of PAH metabolism in invertebrates can differ from those found in 

vertebrates. 

Molluscan and crustacean microsomal enzymes also transform diverse chemicals, 

including aromatic hydrocarbons (Jewell and Winston, 1989; James, 1989 ; Winston et al., 

1991). Rates of hydrocarbon metabolism detected in vitro, in these invertebrates are 1 to 2 

orders of magnitude lower than those seen in most teleost fish liver microsomes. Levels of 

cytochrome P-450 in mollusks and crustaceans are comparable to those seen in some untreated 

fish (but fish vary markedly, with exposure to inducers) . Polychaete MFO has similarities to 

MFO systems in vertebrates and other invertebrates, involving similar cofactors and being 

comprised of phospholipid, cytochrome P-450 and NADPH-cytochrome P-450 reductase. 

MFO components and cofactors required are the same as those required by insect and vertebrate 

MFO systems (Lee et al ., 1981). Microsomal preparations from the mussel, Mytilus edulis, 

tested in vitro, formed quinone derivatives from benzo(a)pyrene,whereas fish microsomes 

formed epoxides and ultimately yielded hydroxylated derivatives. (Stegeman, 1985 ; Anderson, 

1978; Winston et al ., 1988). Other in vitro metabolites formed by molluscan extracts include 

7,8- and 9, 10- dihydrodiols . Fish rely predominantly on cytochrome P450 acting in epoxide 

formation, while PAH metabolism in mussels has been attributed to radical oxidation . 

(Livingstone et al ., 1989; Winston et al ., 1991). Benzo(a)pyrene hydroxylase, from Nereis 

virens was associated with the lower portion of the intestine, with little or no enzyme activity in 

the pharynx, esophagus, or upper portion of the intestine (Fries and Lee, 1984). Lee (1981) 

reported that full enzyme activity required NADPH, Mg+2, and oxygen. Subcellular 

fractionation of the intestine indicated that the enzyme system was associated with the 

microsomal fraction (Fries and Lee, 1984) Activity was linear with time and protein, was 

unaffected by 0.1% Tween-80, was completely inhibited by 0.1°Io SDS and Triton X-100, and 

incubation with phospholipase C destroyed activity . These findings indicate a phospholipid 

component. Activity was diminished by cytochrome P450 inhibitors, e . g., CO, 7,8-

benzoflavone and SKF 525-A [2(diethylamino) ethyl 2,2-diphenylpentanoate)] . Temperature 

optimum was 30° C; pH optimum was between 7 and 8. Fries and Lee (1984) found that 

benzo(a)pyrene was catalyzed by Nereis virens MFO to the major metabolite, 3-

hydroxybenzo(a)pyrene. Also produced are small amounts of 6-hydroxy- and 9-hydroxyderivatives, and the two diols: trans- 4,5- dihydrodiolbenzo(a)pyrene and trans-7,8-

dihydrobenz(a)pyrene . 

The rates and pathways of xenobiotic metabolism in fish and invertebrates can have a 

direct impact on the organisms themselves, and may affect other organisms via food web and 

commensial associations . Metabolism can activate certain carcinogens and can alter 

bioavailability and bioaccumulation characteristics of toxicant compounds. Thus, uptake and 

metabolic transformation influence the fate of xenobiotic compounds in the environment, and can 

determine the identity and levels of parent compound and metabolite residues which accumulate 

in organisms . Induction of P-450 can influence both of these aspects (Stegeman and Lech, 

1991) . In mammals, certain polycyclic aromatic hydrocarbons are metabolically activated by 

MFO, resulting in the production of unstable epoxides which damage DNA and initiate the 

process of carcinogenesis (Miller and Miller, 1971 ; Ames and Sims, 1972) . Polynuclear 

aromatic hydrocarabons (PAHs) are ecologically significant components of crude oil. Stegeman 
et al . (1988a) have noted the potential for formation of activated carcinogenic derivatives of 

PAHs by fish cytochrome P-450 enzymes. Metabolism of PAH is catalyzed primarily by the 

microsomal mixed function oxygenase (MFO) system, of which cytochrome P-450 is the key 

component. The MFO system generally produces PAH metabolites which are less toxic and 

more easily excreted than the parent compound, although, in some cases the oxidation reactions 

produce electrophilic species capable of DNA binding and mutagenesis (Harvey, 1983). 

In the polychaete Capitella capitata, exposure to crude oil or the PAH, benz(a)anthracene, 

in the sediment resulted in the third generation having higher MFO activity than the first or 

second generation (Lee, 1981). These workers did not detect basal MFO levels in C. capitata, 

but noted the need for dissection of intestinal tissue, which was precluded by the small size of 

this species. 

Molluscs are proven, useful indicators of bioavailable levels of contamination, in 

applications involving direct analysis of pollutant residues, which may be attributable to their 

low metabolic activity . But, based on present knowledge, there is little potential for using 

monooxygenase activity or cytochrome P-450 levels in mollusks, or crustaceans, to indicate 

their exposure to compounds such as the aromatic and chlorinated hydrocarbons (Stegeman and 

Lech, 1991). This is due to the lack of any convincing evidence for induction of specific 

cytochrome P-450 isozymes of monooxygenase activity by any of the hydrocarbon inducers 

known to be active in the vertebrates . The marine polychaete worm, Nereis virens exhibited 

ethoxyresorufin-O-deethylase (EROD) induction in the gut tissues, when exposed to 

benzo(a)pyrene via food (Fries and Lee, 1984). These workers also examined feral worms from 

contaminated and reference sites near Portland, Maine. Field-collected Nereis virens from 

heavily-oiled sediment yielded MFO activities six times greater than that of reference site 

specimens . Cytochrome P450 content was also higher, but not significantly so. 

MFO induction has been suggested to indicate the exposure of organisms to xenobiotic 

challenges in the environment. Stegeman and Lech (1991) noted that P-450 induction would not 

detect direct-acting carcinogens and tumor promotors which are non-inducers, although such 

compounds can be expected to co-occur with pollutants that are inducers. Lee (1981) reported 

that the sandworm, Nereis virens, possesses a mixed function oxygenase system (WO), which 

may provide a useful biological indicator of xenobiotic challenge . Nereis virens is promising as 

a bioindicator species, by virtue of its wide geographic distribution, the availability of specimens 

from commercial sources, and its relative ease to maintain in laboratory culture. Demonstrated 

responsiveness of a quantifiable MFO activity in Nereis virens, such as ethoxyresorufin Odeethylase (EROD), would substantiate the usefulness of this organism assay combination as a 

biological indicator of pollution. This study was conducted to evaluate MFO induction in Nereis 

virens, by exposure to authentic, petroleum-contaminated sediment. 

Oil spills and produced water discharges represent major sources of estuarine and marine 

pollution. Biomarker research is often limited to model pollutant compounds or model 

sediments. However, field use of biomonitoring methods involves not only toxicity of individual 

xenobiotic compounds, but synergistic interactions of multiple compounds and components of 

uptake and accumulation phenomena, as well. It is therefore essential, to include authentic 

environmental samples in the investigation of potential biological monitoring systems. 

Biomonitoring regimens yield tangible assessments of physiological effects and, additionally, 

often suggest food web and ecosystem consequences of pollution. 

MATERIALS AND METHODS 

Authentic field-collected sediments were used to challenge Nereis virens in microcosm 

exposure experiments. Natural petroleum- laden sediments were collected near the producedwater discharge of an oil separation site at Pass Fourchon, in coastal Louisiana (map, Figure 1) . 
8 

The sediments for this study were collected at a site 400 m from the discharge. Clean fluvial 

sand was used as a bedding medium for the test organisms prior to experiments, as a control 

medium for zero- level exposures, and as a diluent for varying sediment concentrations . "Clean" 

sand was collected from a pristine, spring-fed creek in central Louisiana. Concentrated extracts 

of sand samples exhibited no petroleum hydrocarbons detectable by standard GC-MS 

methodology applied in this study. 

A considerable body of information on Pass Fourchon has been assembled from 

laboratory studies (Boesch et al ., 1989; Means et al., 1989; Daniels et al ., 1990; Means et a1 ., 

1990; Rabalais et al ., 1991a,b; Means and McMillin, 1993). Pass Fourchon sediment was 

found to contain a complicated suite of aliphatic and aromatic hydrocarbons (Figure 2). The 

aromatic fraction of the extract was found to contain moderate quantities of normal aromatic 

hydrocarbons associated with combustion sources while very high concentrations of a suite of 

alkylated aromatic hydrocarbons, dominated by the C-2 and C-3 alkylated naphthalenes, with 

lesser amounts of other alkylated naphthalenes, alkylated phenanthrenes and alkylated 

dibenzothiophenes. As reported previously, analyses of the produced waters revealed a complex 

mixture of normal alkyl- and heterocyclic- polynuclear aromatic hydrocarbons (Daniels et al., 

1990). Major volatiles in the produced water were benzene and its C-1 to C-3 homologs. Mean 

benzene concentration was 2,000 ng/1, with various other hydrocarbons present at 

concentrations less than 0.5°Io of the benzene concentration. Benzoic acid was the most 

abundant organic acid, at 3100 ng/ml. Total PAHs were 520 ng/ml, and saturated hydrocarbons 

totalled 16,000 ng/ml. The daily discharge of produced water at the Pass Fourchon facility was 

approximately 18,000 bbl/d (Boesch and Rabalais, 1985) . This discharge caused heavy loading 

of sediments in the area, with petroleum hydrocarbons. The most abundant sediment PAHs at 

this site were the alkyl- substituted phenanthrenes. Daniels and Means (1990) studied genotoxic 

effects of Pass Fourchon produced waters on fish embryos, using Cyprinodon variegates as a 

model. They observed an increase in chromosome aberrations in exposures to decreasing 

dilutions, but noted that the effect was not additive with respect to produced water concentration. 

They further reported that a specific type of aberration was induced, i.e . chromosomal breaks 

and extensive alterations in ploidy . 

ORGANISMS: Specimens of the omnivorous, sediment-dwelling, polychaete worm, Nereis 

virens . were obtained from commercial sources and maintained in laboratory holding tanks 

containing a 7 cm layer of clean sand, with 10 cm overlying water. A commercial salts mixture 

(Fritz Chemical Co., Dallas, TX) was used to prepare the full-strength (32 ppt) seawater used 

throughout the experiment. 

MICROCOSMS: Exposures were conducted in microcosm arrangements, with sediment as 

the route of dosing. Individual and multiple- animal microcosms were both used. Individual 

units were preferred, as casualties observed in multiple- animal arrangements were thought to 

have resulted from territorial aggressiveness among worms. Adequate depth of sediment 

seemed to lend protection from agressiveness and from self- inflicted injuries . Worms were 

exposed to 0%, 20 °Io, and 50°l0 levels of Pass Fourchon sediment. Exposure times were also 

varied in the experiments, with MFO inductions evaluated after 0, 3, 7, and 14 days. All 

treatments were replicated in independent duplicate or triplicate studies. 

EROD ASSAYS : At the conclusion of treatments, worms were euthanized by freezing or icechilling, and dissected. The gut was homogenized in 2 ml TES buffer (0.05 M TRIS-HCI, 1 .0 

mM EDTA, 0.25M sucrose, pH 7 .2) by 4 passes in a 10 ml Potter-Elvehjem tissue grinder, at 

250 rpm. Microsomes were purified by multiple centrifugations of the homogenate, according 

to previously published techniques (Winston et al ., 1989). Gross fragments and cell debris 

were removed at 200 X g for 5 min, and organelles and other elements were sedimented at 

10,000 X g for 15 min. The microsomal fraction was pelleted at 40,000 X g for 180 min. 

Mixed function oxygenase activity (MFO) was evaluated in microsomal fractions, as Odealkylation reactions from substituted alkoxyphenoxazones. Specifically, the ethoxyresorufin 
9 

O-deethylase reaction (EROD) was used to detect the presence of this enzymatic mechanism. 

EROD activity was assayed spectrofluorometrically, by the method of Burke and Mayer (1983) 

and Burke et al . (1985), as modified by Winston et al . (1988), with an excitation wavelength of 

530 nm, and emission read at 585 nm. Reaction product was determined from fluorescence 

response of the instrument, as intensity (I) . A standard curve was prepared, of resorufin diluted 

in the reaction matrix. Microsomal protein was determined by a modified Lowery 

technique,with bovine serum albumin as the standard (Winston et al ., 1988). Specific activity of 

EROD was reported as picomoles or nanomoles of resorufin formed per minute per milligram of 

microsomal protein. 

PF1200N=  1 . .z =' a-_ ^ _:. ;~ ~ 

-67 

=s7. ~ ~ ~ r, 

-PF900N:? - -'=_ 

. 

_-'!-~ ;-CFFEVRON.U S A, INCy=~s~==°- pF800N = =" ^= 

_,-",-':PF1200NE ri 

= w'°= -PF600N r~r =_.r.-x_=c= 

- ' _ =A ~ 

_=~s= ___ ~-A-5- PFZ00_ _ _^~ ~3_s== 

=z- _-+~ CHEVRON 

' _ *_ " ~ 

=PF500Sy-~= ,- - 

__ .- : .:-- -PIPE , LINE 

_: hi`=~ 

=} ~ '= COMPANY 

TATE 

N CE 

'?= - __ - = 3~ _= l I I I I I 

- --'' - ' -- - +Gulf of Mexico 

PASS FOURCHON STUDY AREA 

DISCHARGE _ " PRIMARY STATION 

_ Q SECONDARY STATION 

Figure 1. Map of sampling sites, Pass Fourchon, LA. 
10 

SEDIMENT DATA FOR PASS FOURCHON 400 

NAPHTHALENES Ppb 

2-Methylnaphthalene 3 .0 

1-Methylnaphthalene 1 .5 

2-Ethylnaphthalene 1 .9 

1 -Ethyl naphtha[ one 9 .1 

2,6/2,7-Dimethyl naphthalene 10 

1,3/1,T-Dimethyl naphthalene 48 

1,6-Dimethylnaphthalene 4.7 

1,4/2,3-Dimethyl naphthalene 27 

1,5-Dimethyl naphthalene 25 

1,2-Dimethyl naphthalene 12 

2-Isopropylnaphthalene nd 

1,8-Dimethylnaphthalene 12 

1,6,7-Trimethyinaphthalene 197 

C3-NAPHTHALENES 9,700 

C4-NAPHTHALENES 8,900 

PARENTPAH ppb 

Naphthalene 1 .7 

Acenaphthylane 4,0 

Acenaphthene 11 

Fluorene 1 8 

Dibenzothiophene 5.6 

Phenanthrene 52 

Anthracene 1 6 

Fiuoranthene 73 

Pyrene 5 4 

Benzanthracene 24 

Chrysene 8 4 

Benzo(b)fiuoranthene 29 

Henzo(k)fiuoranthene 8 .4 

Benzo(a)pyrene 1 2 

Indenopyrene 30 

Dibenz(a,h)anthracene 17 

Henzo(g,h,i)perylene 3 8 

DIB EP20THIOPHENES p p b 

4-Methyldibenzothiophone 6 8 

2/3-Methyl dibenzothiophene 55 

1-Methyldibenzothiophene 34 

1,2-Dimethyl dibenzothiophene 5.9 

CZ-QIB ENZOTH10PHENES 8,600 

PHENANTHRENES ppb 

3-Methylphenanthrene 1 83 

2-Methylphenanthrene 8 7 

4/9-Methyl phenanthrene 177 

7-Methyiphenanthrane 102 

3,6-Dimethyl phenanthrene 102 

3,5-Dimethyl phenanthrene nd 

2,6-Dimethyl phenanthrene 9 3 

2,7-Dim ethyl phenanthre no 77 

3,9-Dimethyl phenanthrene 335 

1,612,5/2,9-Dimethyl phenanthrene 228 

1,7-Dimethylphenanthrene 83 

1,9/4,9-Dim ethylphenanthrene 78 

1,5-Dimethyl phenanthrene nd 

1,8-Dimethyl phenanthrene 32 

1,2-Dimethyl phenanthrene 22 

9,10-Dimethyl phenanthrene nd 

1,2,8-Trimethylphenanthrene 84 

C3-PHENANTNRENES 23,000 

TOTALS ppb 

Total Parent PAH 478 

Total Alkylated PAH 52,397 

Total PAH 52,875 

Figure 2. Quantitative analysis of petroleum hydrocarbon contaminants in 

Pass Fourchon sediment collected at PF 400 site, 400 m from 

produced water discharge. 
11 

1 .25 

as 

0 1 .00 

a 

0.75 

E 

c 0.50 

E 

20% 

O 0.251 " -, 

a 

0.00 

0 3 7 14 

Exposure Time (days) 

Figure 3 . EROD induction in Nereis virens by exposure to 

petroleum-contaminated sediment. 

Immunoblot Analyses: CYP1A is an important intermediary in mutagen activation. The 

P450 IA activities are inducible, and are associated with several tissues, but are most often 

studied in liver. In mamalian systems, the CYPIA gene family which is inducible by polycyclic 

aromatic hydrocarbons (PAH), consists of one subfamily of two genes: CYPIAI and CYPIA2. 

Immunoblot (Western blot) comparisons were made of worm gut microsomes to known trout 

cytochrome P450 isozyme lAl (courtesy of Dr. D.R. Buhler, Department of Agricultural 

Chemistry, Oregon State University, Corvallis, OR) and to Arochlor-induced rat liver 

microsomes prepared in house (Dr. G.W. Winston, Department of Biochemistry, Louisiana 

State University). The significance of Western blot analysis is in its ability to indicate the 

presence or absence of a CYP1A ortholog or paralog in this invertebrate's system. The protein 

bands, separated elecrophoretically, were reacted with a purified and highly specific polyclonal 

anti-CYP1A1 antibody. The antibody preparation , also kindly provided to us by Dr. Buhler, 

was produced in rabbit, against trout CYP1A1 . The bands of immune complexes were probed 

with an alkaline phosphatase-linked secondary anti-rabbit antibody. 

RESULTS AND DISCUSSION 

The EROD assay is one of the most robust MFO assays available for biomarker research, 

and is not elevated by normal physiological stressors, such as temperature, high or low salinity, 

high or low oxygen levels . The present study investigated EROD induction in the marine 

polychaete Nereis virens, by exposure to petroleum-contaminated sediments . Nereis virens 

exposed to dilutions of the Pass Fourchon sediment showed good survival rates (greater than 

90%) for periods of up to -20 days. In the first battery of experiments conducted EROD was 

observed to be induced in organisms exposed to authentic petroleum-contaminated sediments for 
12 

at least seven days. Unexposed control organisms had mean EROD activity of 0.03 nmol 

resorufin formed/min/mg microsomal protein. Sediment control organisms exhibited a similar or 

lower level of EROD activity. Nereis virens exhibited ethoxyresorufin-O-deethylase (EROD) 

induction in the gut tissues, when exposed in bedded, petroleum-contaminated sediments. Both 

dose dependent and time dependent responses were observed using this experimental approach. 

The worms in 3 day exposures yielded quantifiable EROD activities only in the 50% 

contaminated sediment dose. This activity was not significantly different from the basal level. 

This may indicate a lag phase in expression of enzyme activity or in the kinetics of xenobiotic 

uptake, or it may relate to a threshold level of tissue residues required for induction. Worms 

exposed in 20°Io Pass Fourchon sediment had EROD activities 14-fold higher than controls after 

7 days and 32-fold higher after 14 days. In 50°Io Pass Fourchon sediment, EROD was 27- fold 

higher than control after 7 d, and 32-fold higher after 14 d. The levels of EROD activity were 

consistently higher in the 50% sediment exposure as compared to the 20% exposure. The 

responses of Nereis in these experiments had suggested sufficient bioavailability of dissolved 

and sediment-bound normal, alkylated and heterocyclic aromatic hydrocarbons to benthic 

invertebrates to elicit an induction of mixed function oxidase activity in the invertebrates . Two 

attempts have been made to confirm the data described in the above experiments; in both cases 

the results were not reproducible . Whether this represents seasonal variation in the response of 

the worm to the contaminants present in the sediments awaits further analysis . The 

irreproducibility of the results is inexplicable . Based upon this fact, no definite conclusions as to 

the efficacy of this marine invertebrate as a sentinel of sediment contamination can be drawn at 

this time. 

There have been no reports of CYP1A induction in marine invertebrates. The present 

studies have demonstrated rime- and concentration- dependent induction of EROD in the nereid 

model. EROD is regarded as the indicator of choice for presence of the PAH-inducible isoforms 

of P450, i e., CYP1A. The immunoblot analysis provides stronger evidence for the presence of 

a CYP1A ortholog in EROD-induced Nereis virens ., and thus, whether it may be a component 

of the observed EROD activity. Western blot analysis was performed on two separate 

occasions with the rabbit antibody raised against trout anti CYP1A1 and compared to a BNFinduced trout liver microsomal fraction . The worm gut microsomes were loaded at 50 4g per 

lane and the trout liver at 10 4g per lane. The data showed strong recognition of a single protein 

band of about 54 kDa in the trout liver microsomal fraction, whereas no recognition of epitopes 

was observed with the worm microsomal fractions (Figure 4). 
13 

KDa 

80 .0 

49 .5 

32 .5 

M - molecular weight markers (x;icdai tcns) 

T-trout liver microsomes 10 ~ig 

W - 50 ~ig worm microsomes 

KDa 

80 .0 

49 .5 

Experiment 1 

ri ri 

--7 

., 

~.i 

Experiment 2 

Figure 4. .Western blot analysis of Nereis intestinal microsomes probed 

with trout anti-CYP lAl. 
15 

CHAPTER 2 

INDUCTION OF ETHOXYRESORUFIN O-DEETHYLASE (EROD) 

IN THE KILLIFISH (FUNDULUS GRANDIS) 

BY SELECTED MUTAGENS 

L.A. Reily, J.C. Means, Z-M. Yan, G.W. Winston 

INTRODUCTION 

Major discharges of drilling fluids, drill cuttings, and produced water are associated with 

offshore oil and gas production . In coastal regions, these wastes are discharged into fresh or 

brackish marshes or wetlands which also serve as the spawning sites for a wide variety of 

economically important species of fish and shellfish. Pollutant impacts on early life stages of 

aquatic organisms and on embryonic and larval fish development are of particular significance in 

coastal estuaries and wetland areas. This has implications for the survival and relative fitness of 

exposed fish at these stages, in contaminated spawning areas which are widespread throughout 

coastal Louisiana. Additionally, bioavailability of hazardous compounds in aquatic 

environments increasingly presents a direct threat to man, via seafood consumption. The 

obvious ecological and human health hazards of marine and estuarine pollution make it 

imperative to to seek new and improved technology for monitoring and risk assessment of 

impacts by such pollution. 

Various studies have demonstrated that exposure to benzo(a)pyrene or polychlorinated 

biphenyls causes increases in mixed-function oxygenase (MFO) activity and cytochrome P-450 

content and produces new cytochrome P-450 (Lu et al ., 1973 ; Philpot and Bend, 1975; Bend, et 

al ., 1979; Johnson, 1979; Stegeman, 1980; Varanasi and Gmur, 1981 ; Corbett et al ., 1982; 

Stegeman et al ., 1986; Varanasi et al ., 1986; Payne et al ., 1987 ; Van Veld et al ., 1988) . 

Similarly, the cytochrome P-450-mediated MFO system exists in invertebrates. MFO is 

responsible for the oxidation of a variety of foreign organic compounds,which facilitates 

elimination of lipophilic xenobiotics by converting them to more water-soluble metabolites (Lee, 

1981) . Animals with active hydrocarbon metabolism accumulate lower levels of parent 

polynuclear aromatic hydrocarbons (PAH), and the higher rates of metabolism associated with 

P-450 induction, can enhance PAH elimination (Lu et al ., 1977). MFO systems in mammals 

and fish are similarly comprised of phospholipid, cytochrome P-450, and NADPH cytochrome 

P-450 reductase (Stegeman, 1989). MFO cofactors required are also the same, although the 

mode of action may differ (Lee, 1981 ; Lee et al ., 1981) . Microsomal preparations from fish 

microsomes formed epoxides and ultimately yielded hydroxylated derivatives (Stegeman, 1985; 

Anderson, 1978; Winston et a1 ., 1988) . The rates and pathways of xenobiotic metabolism in 

fish and invertebrates can have a direct impact on the organisms themselves, and may affect 

other organisms via food web and commensial associations . Metabolism can activate certain 

carcinogens and can alter bioavailability and bioaccumulation characteristics of toxicant 

compounds. In mammals, certain polycyclic aromatic hydrocarbons are metabolically activated 

by MFO, resulting in the production of unstable epoxides which may damage DNA and initiate 

the process of carcinogenesis (Miller and Miller, 1971 ; Ames and Sims, 1972) . Thus, uptake 

and metabolic transformation influences the fate of these compounds in the individual and in the 

environment, and can determine the identity and levels of parent compound and metabolite 
16 

residues which accumulate in organisms. Induction of P-450 can influence all of these aspects 

(Stegeman and Lech, 1991). Stegeman and others have studied the potential for formation of 

activated carcinogenic derivatives of PAHs by fish cytochrome P-450 enzymes (Stegeman and 

Lech, 1991 ; Stegeman et al ., 1988b). 

Polynuclear aromatic hydrocarbons (PAHs) are ecologically significant components of 

crude oil. Metabolism of PAH is catalyzed primarily by the microsomal mixed function 

oxygenase (MFO) system, of which cytochrome P-450 is the key component. The MFO system 

generally produces PAH metabolites which are less toxic and more easily excreted than the 

parent compound, although, in some cases the oxidation reactions produce electrophilic species 

capable of DNA binding and mutagenesis (Harvey, 1983). 

MFO induction has been suggested to indicate the exposure of organisms to xenobiotic 

challenges in the environment although, Stegeman and Lech (1991) noted, P-450 induction in 

sentinel species would not detect direct- acting carcinogens and tumor promotors which are noninducers, but such compounds can be expected to co- occur with pollutants that are inducers. 

The present studies investigate the induction of MFO in the killifish, Fundulus grandis, as a 

biological indicator of pollution . MFO evaluation was focused on detection of O-dealkylation 

reactions from substituted alkoxyphenoxazones, with the ethoxyresorufin O-deethylase reaction 

(EROD) used to detect the presence of this enzymatic mechanism. EROD has been recently 

identified as a sensitive sublethal index of environmental impact on several test organisms, its 

induction indicating : 1) the bioavailability of dissolved and sediment-bound normal, alkylated 

and heterocyclic aromatic hydrocarbons ; 2) the ability of organisms to metabolize these 

compounds; and 3) the genotoxicity of the compounds and their metabolites . Laboratory 

challenge experiments were conducted with model compounds, to evaluate the responsiveness of 

EROD induction in adults, larvae, and embryos of the killifish . Three known mutagenic agents 

were chosen as model compounds for these studies . Cyclophosphamide (CP) and 

ethylmethanesulfonate (EMS) are two benchmark mutagens which have been documented with 

respect to their genotoxic effects on Fundulus. ; 3-methylcholanthrene (3-MC) is classically used 

as a pro-mutagen, MFO inducer and model carcinogen in studies of mammalian systems. 

MATERIALS AND METHODS 

Specimens of the killifish, Fundulus grandis, were obtained from commercial sources 

and maintained in laboratory aquaria, at 7 ppt salinity, 210 C, and 9 hour light photoperiod. 

Spawning was induced by 12 h light, at 250 C. Fertilized embryos were cultivated at 250 C, 7 

ppt salinity, and 12 h light. Hatchling larvae were maintained at 210 C, 7 ppt salinity and12 h 

light, and fed daily with live Artemia. 

Laboratory exposure experiments were conducted with the three major developmental 

stages of Fundulus grandis, i. e. adults, embryos, and larvae . Adults were exposed for 24 h to 

100 nM EMS or CP in water and sacrificed 5 d later. Because of concern over insolubility, 3-

MC was administered to adults by injection (20 mg/kg, in corn oil) on days 0, 4, and 8, and they 

were sacrificed on day 16. Later studies have suggested that water-column exposures of adults 

may also be efficacious. Embryos (11 day post-fertilization) and larvae (11 day post-hatch) 

were exposed to 3-MC, EMS, or CP in the water, at 100 nM concentration and sacrificed 48 

hours later. Stock solutions of the model mutagens were prepared in DMSO, and results were 

compared to DMSO- dosed controls. All experiments were duplicated in independent trials. 

Adult fish were euthanized by decapitation, and dissected. Larvae, whole embryos, and 

adult livers were homogenized in 2 ml buffer (.OS M TRIS-HCl, 1 .0 mM EDTA, 0.25M 

sucrose, pH 7.2) by 5 passes in a 10 ml Potter-Elvehjem tissue grinder, at 250 rpm. 
17 

Microsomes were purified by differential centrifugation of the homogenate, according to 

Winston et al . (1989) . Gross fragments and cell debris were removed at 200 X g for 5 min, 

and organelles and other elements sedimented at 10,000 X g for 15 min. The microsomal 

fraction was pelleted at 105,000 X g for 60 min. Microsomal mixed function oxygenase activity 

(MFO) was evaluated by measuring the rates of ethoxyresorufin O-deethylase (EROD) 

spectrofluorometrically (Burke and Mayer, 1983; Burke et al ., 1985), as modified by Winston et 

al. (1988) . Excitation and emission wavelengths were 530 and 585 nm, respectively . Reaction 

product was determined from a standard curve, prepared of resorufin diluted in the reaction 

matrix. Microsomal protein was determined by the method of Lowry et al . (1951) technique, 

with bovine serum albumin as the standard. Specific activity of EROD was reported as 

nanomoles resorufin formed per minute per milligram of microsomal protein (Figure 5). 

RESULTS AND DISCUSSION 

EROD induction was demonstrated in Fundulus grandis embryos, larvae, and adults by 

the three model MFO inducers investigated in this study. The various developmental stages of 

Fundulus grandis , particularly the embryonic and larval stages, exhibited substantially elevated 

levels of MFO activity following short exposures to these mutagens. The mean EROD activity 

in control adult livers was 0.08 nmol resorufin produced/min/mg microsomal protein. EROD 

activity was not detected in control embryos or larvae . Exposure to 3-MC induced EROD 

activities to 2.80, 1 .90, and 2.21 nmol/min/mg protein in embryos, larvae, and adults, 

respectively . EROD activities induced by EMS treatment were 1 .50, 1 .60, and 0.57 

nmoUmin/mg protein. CP induced activities of 1 .25, 1 .20, and 0.18 nmol/min/mg protein, 

respectively. 

CP and EMS are two benchmark mutagens which have been documented with respect to 

their genotoxic effects on Fundulus . (Baksi and Means) 3-MC is classically used as a promutagen, MFO inducer and model carcinogen in studies in mammalian systems. The various 

developmental stages of Fundulus grandis , and in particular the embryo and larval stages, 

exhibited substantially elevated levels of MFO activity, following exposure short exposures to 

these mutagens, as demonstrated in figure 5). EROD was detected in microsomal fractions from 

whole embryos, larvae, or excised adult livers . The mean EROD activity in control adult livers 

was 0.08 nmol resorufin produced/min/mg microsomal protein. EROD activity was not detected 

in control embryos or larvae . Exposure to 3-MC induced EROD activities to 2.80, 1 .90, and 

2.21 nmol/min/mg in embryos, larvae, and adults, respectively . EROD activities induced by 

EMS treatment were 1 .50, 1 .60, and 0.57 nmol/min/mg protein. CP induced activities of 1 .25, 

1 .20, and 0.18 nmol/min/mg protein, respectively . Significant among these finding are the 

determinations of elevated EROD activities in the early development of the metabolic machinery 

of the embryo and larval fish . This has implications for the survival and relative fitness of 

exposed fish at these stages in contaminated coastal spawning areas so widespread throughout 

Louisiana. Also of interest is the relative activities observed in these early life stages with that 

found in adult livers, a primary organ involved in xenobiodc metabolism . These results suggest 

that there should be more emphasis on examining the metabolic competences of early life stages 

of fish. 

Many chemical carcinogens occur as procarcinogens, and require activation by metabolic 

processes to yield carcinogenic derivatives. Oxidative transformations of these compounds are 

primarily catalyzed by microsomal monooxygenases or mixed function oxidases. The various 

developmental stages of Fundulus grandis , in particular the embryo and larval stages, exhibited 

substantially elevated levels of MFO activity following short exposures to the mutagens. EROD 

activity was not detected in control embryos or larvae. 

A bioindicator system, based upon EROD induction in Fundulus grandis, could prove to 

be a valuable tool for assessment of potential impacts on both ecological and human 
18 

environmental risks associated with increased discharges of petroleum-associated hydrocarbons. 

As the aquatic environment is a major sink for numerous contaminants, the elucidation of 

biochemical responses to contaminants, by aquatic animals, becomes increasingly important. 

The epidemiology of highly elevated rates of idiopathic lesions and neoplasia among some 

populations of aquatic animals inhabiting polluted environments is, in most cases, far clearer 

than is generally the case for human cancers suspected of being related to environmental 

pollution. 

The present study investigated EROD induction in the major life stages of the killifish. 

Experimental subjects showed good survival rates (>90% in all of the treatments used). EROD 

was induced in adults, larvae, and embryos exposed to the model mutagens tested . The 

responsiveness of EROD in the killifish indicates promise for development of this assay/ 

organism system, as a biomarker of contamination in marine and estuarine environments. The 

EROD assay is one of the most robust MFO assays available for biomarker research . It is highly 

selective for CYP1A, the aromatic hydrocarbon-inducible form of cytochrome P450, and is not 

elevated by normal physiological stressors, such as temperature, high or low salinity, high or 

low oxygen levels. Unexposed control organisms in the present study had mean EROD activity 

of 0.08 nmol resorufin formed/min/mg microsomal protein, indicating a low basal level of 

EROD activity . 

Experiment 1 

Experiment 2 

0 6 Experiment 3 

a 

rn 

E 

1 c 4 

c 

G 

w 

0 

3-MC EMS CP CONTROL RAT LIVER 

TREATMENT 

Figure 5a. EROD induction in Fundulis grandis larvae in 

independent replicate experiments. 
19 

8 

o / Experiment 1 duplicate experiments L 

~ Experiment 2 24 h exposures 

6 

c 

£ 4 

c 

0 2 

0 

W 

0 ILL =122M 

3-MC BAS CID CONTROL RAT LIVER 

TREATMENT 

Figure Sb. EROD induction in Fundulis grandis adults in 

independent replicate experiments. 

o / Experiment 1 

dupl. experiments-24 hr exposure 

~ Experiment 2 

g ~-' 

Experiment 3 48 hr exposure >` 

C 

4 -

C 

2- 4:7: 

6 

0 mom 

w 0 ". I 

3-MC EMS CID CONTROL RAT LIVER 

TREATMENT 

Figure Sc. EROD induction in Fundulis grandis embryos 

in independent replicate experiments. 
20 

Since the aquatic environment, particularly sediments, are major sinks for numerous 

contaminants, the elucidation of biochemical responses to contaminants by aquatic animals is an 

increasingly important area of research . The epidemiology of highly elevated rates of idiopathic 

lesions and neoplasia among some populations of aquatic animals inhabiting polluted 

environments is, in most cases, far clearer than is generally the case for human cancers 

suspected of being related to environmental pollution. The data generated in these studies may 

allow the assessment of potential impacts, as well as ecological and human environmental risks 

associated with increased discharges of petroleum-associated hydrocarbons . 

The low background activity, the broad spatial distribution of populations of the species 

and responsiveness of EROD activity in Fundulus grandis indicates the potential usefulness of 

this organism assay combination as a biomarker for environmental contamination . It is 

noteworthy, that induced EROD levels in Fundulus grandis reach a specific activity 

approximately one- third to one- half that observed in Arochlor-induced rat livers . Also of 

significance is the finding that exposures of as little as 24 h will result in a quantifiable and 

significant increase in the activity of EROD in the liver tissues of Fundulus grandis. 
21 

CHAPTER 3 

FIELD INVESTIGATIONS OF THE INDUCTION OF EROD IN 

FISH AT ST 34 AND ST 52 PLATFORMS IN THE 

GULF OF MEXICO, PASS FOURCHON AND LAKE CHAMPAGNE 

L.A. Reily, J.C. Means, S.B. Dobias, G.W. Winston 

INTRODUCTION 

Field studies for validation of MFO induction as a bioindicator in feral fish as a function 

of pollution gradients were conducted by contrasting EROD levels in organisms from control 

(pristine) areas with organisms from known polluted environments . Field validation of the 

EROD assay as a biomarker in teleost fish, should provide a sensitive index of environmental 

pollution, to detect exposure history and to qualitatively establish the bioavailability of dissolved 

and sediment-bound normal, alkylated and heterocyclic aromatic hydrocarbons. This would 

allow environmental managers to assess the potential impacts and both ecological and human 

environmental risks associated with discharges of petroleum-associated hydrocarbons. As the 

aquatic environment is a major sink for numerous contaminants, the elucidation of biochemical 

responses to contaminants, by aquatic animals, becomes increasingly important . The 

epidemiology of highly elevated rates of idiopathic lesions and neoplasia among some 

populations of aquatic animals inhabiting polluted environments is, in most cases, far clearer 

than is generally the case for human cancers suspected of being related to environmental 

pollution . 

MATERIALS AND METHODS 

The present study evaluates biochemical markers recently identified as sensitive sublethal 

indices of environmental impact on several test organisms through detailed studies of: 1) the 

bioavailability of dissolved and sediment-bound normal, alkylated and heterocyclic aromatic 

hydrocarbons to benthic invertebrates and demersal fish, eggs, and larvae; 2) the ability of both 

benthic and pelagic organisms to metabolize these compounds; and 3) the genotoxicity of the 

compounds and their metabolites in benthic organisms and demersal fish . 

Field validation studies of MFO induction as a potential indicator in species of teleost fish 

were undertaken. Collecting trips to offshore and coastal marsh areas (Pass Fourchon) yielded 

fish specimens from both contaminated and control (pristine) locations. The bottom-dwelling 

fishes, Fringed flounder (Etropus crossotus) and Shoal flounder (Syacium gunteri) were trawled 

near offshore petroleum production platforms with known produced water discharges 

(contaminated) and without produced water discharges (control). Collected specimens were 

evaluated for ethoxyresorufin O-deethylase (EROD) induction. Excised livers were pooled 

according to species and collection site, and were homogenized, then centrifuged at 9000 x g. 

The supernates (S9 fractions) were tested for EROD) activity . A scaled-down technique, 

accomodating less tissue, allowed individuals to be tested . S9 fractions were prepared from 

pooled or individual livers, and assayed for EROD activity. Further-purified microsomal 

extracts from the livers of control and contaminated specimens were analyzed for EROD activity . 

Preparations of S9 fractions from contaminated specimens had low levels or zero EROD 

activities . Microsomal fractions were harvested by centrifuging the S9 fractions at 40,000 x g 

for 180 min. Microsomal protein was determined by a modified Lowery (1951) procedure, with 
22 

bovine serum albumin as the standard. Microsomal fractions catalyzed sufficient activity to 

permit quantification and comparison of EROD activities. 

Additional samples of fishes from offshore sites were provided to the laboratory by the 

Louisiana Department of Wildlife and Fisheries. These samples were collected during LDWF 

offshore sampling trips, by trawling at control and impacted stations in the South Terrebonne 

region of coastal Louisiana. Collection dates and hepatic microsomal EROD for these samples 

are presented in Table 3.4 . Our field investigations of EROD inductions were amply enhanced 

by the inclusion of these samples to our database. Sediment cores were collected from the Lake 

Champagne reference site and at distance increments from the Pass Fourchon produced water 

discharge. These samples were analyzed for PAH contents, by methods detailed in chapter 7. 

Data for these analyses are reported in Table 3.1 

RESULTS AND DISCUSSION 

Pass Fourchon sediment was found to contain a complicated suite of aliphatic and 

aromatic hydrocarbons (Table 3 .1). As might be expected, the concentration and makeup, of 

sediment contaminants varies as a function of distance from the discharge. The aromatic fraction 

of the extract was found to contain moderate quantities of normal aromatic hydrocarbons 

associated with combustion sources while very high concentrations of a suite of alkylated 

aromatic hydrocarbons, dominated by the C-2 and C-3 alkylated naphthalenes, with lesser 

amounts of other alkylated naphthalenes, alkylated phenanthrenes and alkylated 

dibenzothiophenes . As reported previously, analyses of the produced waters revealed a complex 

mixture of normal alkyl- and heterocyclic- polynuclear aromatic hydrocarbons (Daniels et al., 

1990) . Total PAHs were 520 ng/ml, and saturated hydrocarbons totalled 16,000 ng/ml . The 

daily discharge of produced water at the Pass Fourchon facility was approximately 18,000 bbl/d 

(Boesch and Rabalais, 1985). This discharge caused heavy loading of sediments in the area, 

with petroleum hydrocarbons. Daniels et al. (1990) reported a concentration of total sediment 

PAHs to be into the parts per thousand range. The most abundant sediment PAHs at this site 

were the alkyl- substituted phenanthrenes. Daniels et al. (1990) studied genotoxic effects of 

Pass Fourchon produced waters on fish embryos, using Cyprinodon variegates as a model. 

They observed an increase in chromosome aberrations in exposures to decreasing dilutions, but 

noted that the effect was not additive with respect to produced water concentration. They further 

reported that a specific type of aberration was induced, i . e. chromosomal breaks and extensive 

alterations in ploidy. Analyses of MFO levels in field-collected organisms were expected to 

show a sharp delineation between pristine and contaminated areas. Specimens taken at various 

distances from pollution sources should exhibit a gradient of MFO induction responses, in 

accordance with decreasing pollutant concentration over distance. 

Sediment concentrations of petroleum hydrocarbons were analyzed at various distances 

from the Pass Fourchon discharge site. These data, presented in Table 3.1, indicated that a 

gradient of pollutant concentrations indeed exists as a function of distance from the discharge. 

Lake Champagne was chosen to represent a control (pristine) area. Analyses of sediment cores 

from this reference site confirmed that it is relatively uncontaminated . In Table 3 .1, these Lake 

Champagne data may be contrasted with sediment data for the pollution gradient samples from 

Pass Fourchon . 

Specimens collected and tested from the Lake Champagne reference area included: 

Pompano (Trachinotus carolinus), Croaker (Micropogonias undulates), and Southern flounder 

(Paralichthys lethostigma) . EROD activity was not detected in these field-collected, control 

specimens (data not shown). This confirms that livers of feral fish in natural clean environments 

do not exhibit elevated EROD activities. . 
23 

In a subsequent field collection, EROD induction was compared for contaminated (Pass 

Fourchon) and control (Lake Champagne) organisms. Harvested specimens were pooled by 

species for analyses, and hepatic EROD data are presented in Table 3.2 . EROD was not detected 

in control Pinfish, Pompano, or Southern flounder. Levels of EROD observed in other control 

organisms may represent uninduced background, levels, but insufficient samples precluded 

substantiation of this . Suprisingly, the contaminated samples of Croaker, Anchovy, and Jack 

had undetectable EROD, whereas measurable levels were found in their Lake Champagne 

counterparts . The reason for this was not evident, although speculations might include 

supressive influences of specific compounds or overall xenobiotic stress in the contaminated 

environment. Influences in the Lake Champagne environment which might be postulated to 

increase EROD expression, might include dietary substances such as phenolic compounds from 

plants . The only well delineated sample contrast was found in the Southern flounder, in which 

single specimens represented the control and contaminated conditions . EROD was not detected 

in the control sample, but hepatic EROD in the contaminated representative was 2,500,000 

pmol/min/mg. It must be emphasized that these data represent only individual organisms. 

Subsequent sampling trips did not produce flounder samples for replicating this comparison. 

Induction of EROD in the flounder, if this was not a chance phenomenon, might relate to its 

tendency to occupy a rather small range, whereas the scattering of EROD levels observed in the 

other organisms may be an artifact of their more nektonic lifestyle. 

Organisms collected from contaminated environments near offshore oil platforms 

included specimens of Fringed flounder (Etropus crossotus) and Sea robin (Prionotus tribulus). 
24 

Table 3 .1 Sediment data for Pass Fourchon and reference site 

NAPHTHALENES ppb 

2-Methylnaphthalene 3 .0 

1-Methylnaphtha(ene 1 .5 

2-Ethylnaphthafene 1 .9 

1-Ethylnaphthalene 9 .1 

2,6/2,7-Dimethyl naphthalene 10 

1,3/1,7-Dimethyl naphthalene 48 

1,6-Dimethyl naphthalene 4.7 

1,4/2,3-Dimethyl naphthalene 27 

1,5-Dimethylnaphthalene 25 

7,2-Dimethyl naphthalene 12 

2-lsopropylnaphthalene rtd 

1,8-Dimethylnaphthalane 12 

1,6,7-Trimethyl naphthalene 197 

C3-NAPHTHALEN ES 9 , 700 

C4-NAP HTHALENES 8,900 

PARENT PAH ppb 

Naphthalene 1 .7 

Acenaphthylene 4.0 

Acenaphthene 1 1 

Fluorene 1 8 

Dibenzothiophene 5.6 

Phenanthrene 52 

Anthracene 1 6 

Fluoranthene 73 

Pyrene 5 4 

Benzanthracene 24 

Chrysene 8 4 

Benzo(b)i(uoranthene 29 

Benzo(k)tluoranthene ' . ~ 8 .4 

Benzo(a)pyrene 1 2 

indenopyrene 30 

Dibenz(a,h)anthracene 17 

eenzo(g,h,i)perylene 38 

DIBENZOTHiOPHENES pp b 

4-Methyldibenzothiophene 6 8 

Z3-Methyldibenzothiophene 55 

1-Methyl dibenzothiophene 3 4 

1,2-Dimethytdibenzothiophene 5.9 

C2-DIBENZOTHIOPHENES 8 ,6 00 

PHENANTHRENES ppb 

3-Methylphenanthrene 183 

2-Methylphenanthrene 87 

419-Methyl phsnanthr en e 177 

1-Methyiphenanthrene 102 

3,6-Dimethylphenanthrene 102 

3,5-Dimethylphenanthrene nd 

2,6-Dimethylphenanthrene 9 3 

2,7-Dimethylphenanthrene 77 

3,9-Dimethylphenanthrene 335 

y,6/2,5/2,9-Dimethylphenanthrerte 228 

1,7-Dimethyiphenanthrene 8 3 

1,9/4,9-Dimethyl phenanthrene 78 

1,5-Dimethyl phenanthrene nd 

1,8-Dimethylphenanthrene 32 

1,2-Dimethylphenanthrene 22 

9,70-Dimethylphenanthrene nd 

1,2,8-Trimethylphenanthrene 8 4 

C3-PHENANTHRENES 23,000 

TOTALS . ppb 

Total Parent PAH 478 

Total Alkytated PAH 52,397 

Total PAH 52,875 
25 

Table 3.2 Hepatic microsomal EROD levels in specimens from nearshore sites at Pass Fourchon 

and Lake Champagne, LA 

EROD, pmol/min/mg 

Organism: Champagne Pass Fourchon 

(Control) (Contaminated) 

# # 

Hardhead catfish 3 80-12,667* 4 204-647 

(Arias fells) 

Croaker 3 450 4 n. d. 

(Micropogonias undulatus) 

Anchovy 26 741 4 n. d. 

(Anchoa mitchilli) 

Pinfish 8 n. d . 16 33 

(Lagodon rhomboides) 

Spot 17 1,075 40 719-2,222 

(Leiostomus xanthurus) 

Pompano 1 n. d. 

(Trachinotus carolinus) 

Southern flounder 1 n. d. 1 2,500,000 

(Paralichthys lethostigma) 

Jack 1 2,000 1 n. d. 

(Caranx hippos) 

* Ranges are given where catch size permitted subsampling; n. d., not detected . 
26 

EROD in Flatfish Near Offshore Structures 

STA55 

STA55 

STA55 

STA55 

STA55 

STA54 

STA54 

STA54 

STA54 

o STA53 

STA53 

STA53 

_, STA53 

STA53 

STA52 

--' STA52 

a' STA52 

STA37 

STA37 

STA37 

STA36 " Bay whiff STA36 

STA36 D Fringed flounder 

STA36 

STA35 m Shoal flounder 

STA35 

STA35 

STA21 

0 0 

o 

0 0 0 0 0 0 0 0 

n o n o ~n 

N N 

EROD, pmol/min/mg microsomal protein 

Figure 6. Induction of EROD in three species of flatfish near offshore 

production facilities . 
27 

Table 3.3 . Hepatic microsomal EROD levels in fish collected by LDWF 

near offshore platforms. 

Site Species Sample Date EROD 

ST 21 Fringed flounder 12%17/93 81 .8 

ST 35 Bay whiff 11/9/93 0.00 

ST 35 Fringed flounder 11/9/93 0.00 

ST 35 Fringed flounder 12/17/93 141 

ST 35 Fringed flounder 12/17/93 331 

ST 35 Fringed flounder 12/17/93 221 

ST 36 Bay whiff 12/17/93 331 

ST 36 Fringed flounder 11/9/93 0.00 

ST 36 Fringed flounder 12/17/93 344 

ST 36 Fringed flounder 11/9/93 0.00 

ST 36 Fringed flounder 12/17/93 283 

ST 36 Fringed flounder 11/9/93 0.00 

ST 36 Fringed flounder 12/17/93 876 

ST 36 Fringed flounder 11/9/93 0.00 

ST 37 Bay whiff 11/9/93 0.00 

ST 37 Fringed flounder 11/9/93 0.00 

ST 37 Fringed flounder 12/17/93 0.00 

ST 37 Fringed flounder 11/9/93 0.00 

ST 37 Fringed flounder 12/17/93 620 

ST 37 Fringed flounder 11/9/93 0.00 

ST 37 Fringed flounder 12/17/93 766 

ST 52 Bay whiff 10/14/93 0.00 

ST 52 Bay whiff 11/3/93 0.00 

ST 52 Bay whiff 12/10/93 0.00 

ST 52 Fringed flounder 5/3/93 0.00 

ST 52 Fringed flounder 12/10/93 0.00 

ST 53 Bay whiff 10/14/93 0.00 

ST 53 Bay whiff 12/10/93 941 

ST 53 Bay whiff 12/10/93 0.00 

ST 53 Bay whiff 11/3/93 0.00 

ST 53 Fringed flounder 3/2/93 0 .00 

ST 53 Fringed flounder 5/3/93 0.00 

ST 53 Fringed flounder 6/14/93 93327 

ST 53 Fringed flounder 8/3/93 821 

ST 53 Fringed flounder 12/10/93 0.00 

ST 53 Shoal flounder 6/14/93 90125 

ST 53 Shoal flounder 8/3/93 1341 

ST 54 Bay whiff 11/3/93 0.00 

ST 54 Bay whiff 10/14/93 0 .00 

ST 54 Fringed flounder 5/3/93 0 .00 

ST 54 Fringed flounder 3/2/93 0 .00 

ST 54 Fringed flounder 12/10/93 0.00 

ST 54 Fringed flounder 12/10/93 81 .8 

ST 54 Fringed flounder 12/10/93 13.4 

ST 55 Bay whiff 11/3/93 0.00 

ST 55 Bay whiff 12/10/93 0.00 

ST 55 Bay whiff 10/14/93 0.00 

ST 55 Fringed flounder 3/2/93 0.00 

ST 55 Fringed flounder 5/3/93 919 

ST 55 Fringed flounder 6/14/93 372 

ST 55 Fringed flounder 8/3/93 5325 

ST 55 Fringed flounder 11/3/93 0.00 

ST 55 Shoal flounder 8/3/93 1332 
28 

Field investigations of EROD inductions were also conducted on feral fish collected by 

LDWF, at control and impacted stations in the coastal region of Louisiana. Data from these 

studies are reported in Table 3.4 and Figure 6. 

Specimens of benthic fish were collected by trawling in control areas, coastal estuaries 

and offshore oil platforms. Initial field studies were performed on fish species from Pass 

Fourchon, a coastal site, and three offshore sites designated STC1 (a control site) and two 

produced water sites, ST34 and ST52. EROD activity in fish liver from these offshore sites 

were studied as a function of distance from the point-source discharge. Although EROD 

readings were obtainable only on two distance points (200 and 2000 m) from the oil platform at 

ST34, and two (500 and 2000 M) from the platform at ST52, an apparent inverse correlation 

was observed in EROD activity of Fringed flounder livers from these sites (data not shown). 

This is not unprecedented and points to the need for immunochemical analysis with antibodies 

prepared against the enzymes (cytochromes P450 IA) catalyzing this activity . We have not been 

successful in obtaining positive western blots with monoclonal anti CYP1A antibodies prepared 

from cod liver or mammalian liver. The array of data does not indicate any statistically 

significant pattern of response to field parameters. Closer scrutiny and reconciliation of data are 

warranted. 

A direct correlation was seen at ST34 in Sea robin (Figure 7), however the sample at the 

closest distance to the site (200 M) was lost during preparation of microsomes. Subsequent 

collections did not produce Sea robins from this distance . The activites are much lower than 

anticipated for the induced state but we note that baseline parameters for this species have not 

been reported. EROD activity (10.5) nmol/min/mg was detected in a Sea robin 1000 M from a 

platform at ST52. Protein analysis was not sensitive enough to obtain values for specific activity 

calculations at the other distance points on these and certain other samples. 

The data in Tables 3.2 - 3.4 indicate that EROD activity varies by as much as four orders 

of magnitude among the various species from the several locations from which they were 

obtained . Because it is known that ethoxyresorufin degrades rapidly even in the freezer, it is 

standard operating procedure to recalibrate standards on a daily basis. From daily recalibrated 

standards new standard curves were prepared for every experiment. Figure 8 below shows a 

typical resorufin standard curve. Note the excellent correlation coefficient of 0.993. 
29 

Figure 7 . EROD Activity in Sea robin as a function 

of distance from ST 34 

120 -

as 

v 

80 -

v 

N 

4) 

L- 0 40 

y = 1 .457 + 114.14x R^2 = 0.993 

0.0 0.5 1 .0 

nmol resorufin 

Figure 8. Typical EROD standard curve 
30 

Table 3.4 Liver microsomal EROD activities of Gulf of Mexico fish samples 

Location/Distance 

Sample# Species from Platform pmoUmin/mg 

301 Hardhead catfish ST 34 100 m 14.8 

302 Hardhead catfish ST 34 50 m BDL 

303 Hardhead catfish ST 34 1000 m 12.5 

304 Hardhead catfish ST 34 50 m 42.2 

305 Hardhead catfish ST 34 50 m 42.2 

306 Hardhead catfish ST 34 1000 m 9 .8 

307 Hardhead catfish ST 34 1000 m BDL 

309 Fringed flounder ST 53a 500 m BDL 

Control rat 10-BDL 

3MC-treated rat 899/775 

308 Fringed flounder ST 53a 200 m 26.2 

310 Shoal flounder ST 53a 500 m BDL 

311 Fringed flounder ST 34 200 m 7 .2 

312 Fringed flounder ST 53a 1000 m BDL 

313 Fringed flounder ST 53a 1000 m BDL 

315 Bay whiff ST 34 1000 m BDL 

316 Bay whiff ST 34 1000 m BDL 

317 Fringed flounder ST 34 1000 m 2.1 

318 Fringed flounder ST 34 1000 m BDL 

319 Sea robin ST 34 1000 m BDL 

320 Bay whiff ST 34 1000 m BDL 

321 Fringed flounder ST 34 500 m 1 .7 

322 Shoal flounder ST 34 500 m 5.4 

37 Fringed flounder ST 34, 500 m BDL 

38 Sea robin ST 34, 500 m 0.01 

39 Fringed flounder ST 34, 200 m BDL 

40 Fringed flounder ST 34, 2000 m 0.04 

41 Gulf menhaden ST 34, 1000 m 0.01 

42 Sea robin ST 34, 1000 m 0 .003 

43 Fringed flounder ST 34, 1000 m BDL 

44 Sea robin ST 34, 2000 m 0 .001 

45 Fringed flounder ST 52, 200 m 0 .02 

46 Fringed flounder ST 34, 200 m 0 .02 

47 Pompano ST 34,200 m BDL 

70 Threadfin shad ST CI 500 0.001 

71 Sea robin ST 52, 500 m BDL 

72 Fringed flounder ST 52, 500 m 0.01 

73 Sea robin ST 52, 1000 m 0.01 

74 Fringed flounder ST 52, 1000 m BDL 

75 Sea robin ST 52, 2000 m BDL 

176 Bay whiff ST 53 32.3 

177 Fringed flounder ST 54 37.0 

178 Fringed flounder ST 52 31 .7 

BDL = below detection limits 

Figure 9 shows a typical microsomal response curve representative of the conditions of 

all of our assays, which were conducted under conditions in which the assay was first order 

with respect to microsomal protein. 
31 

30 -

20-

U 

N 

y 

U 

L. 

1 0-

= 0.952 + 5 .77e-2x R^2 = 0.992 

0' 

0 200 400 

protein (u g) 

Figure 9. EROD vs. microsomal protein concentration 

EROD activities ranged from below detectable limits to values of 42.2 pmol/min/mg in 

offshore catfish. In the coastal sites of Pass Fourchon and Lake Champagne values ranged from 

0.08 to 2.5 pmol/min/mg protein. Rat liver controls from 3MC-induced rats were typically in 

the range of 840 ± 60 pmol/min/mg protein. 

Table 3.5 Liver microsomal EROD acrividtes of fish species sampled from a contaminated 

(Pass Fourchon) and reference (Lake Champagne) coastal site in the Gulf of Mexico. 

Sample# Species Conditions pmol/min/mom 

50 Hardhead catfish LC 0.08 

51 Hardhead catfish LC 0.15 

52 Hardhead catfish LC 1 .07 

53 Hardhead catfish LC 0.27 

55 Croaker LC 0.45 

57 Anchovy LC 0.74 

62 Jack LC 2.00 

65 S pot LC 1 .08 

64 Jack PF 0.21 

66 Spot PF 2.22 

67 Spot PF 0.72 

58 Anchovy PF BDL 

59 Fringed flounder PF 2 .50 

61 Pinfish PF 0.03 

49 Hardhead catfish PF 0 .65 

54 Hardhead catfish PF 0 .21 

48 3-viC-induced Ftcndulcu adult 0 .09 

BDL = below detection limits of 0.001 pmol/min/mg protein 
32 

In environments where carcinogenic PAHs occur, MFO induction does not necessarily 

mean that the parent compound is still at high levels in the fish tissues. Studies have shown that 

animals with active hydrocarbon metabolism accumulate lower levels of parent PAH (Lu et al ., 

1977) . The higher rates of metabolism associated with induction of the hydrocarbon 

metabolizing P-450 can enhance the rates of PAH elimination, via transformation to more 

soluble compounds. The detection of high levels of hydrocarbon metabolites in the bile of fish 

exposed to hydrocarbons is consistent with this reasoning(Statham et al ., 1978; Goddard et al ., 

1987) Highly induced animals can be expected to have lower levels of PAHs in their tissues 

than either animals which are not induced or those that have inherently lower rates of 

hydrocarbon metabolizing activity . Numerous investigations have reported relatively high levels 

of PAH residues in molluscan tissues, intermediate levels in crustaceans and low levels of these 

compounds in fish . This was illustrated in a study by Dunn and Fee (1979), which reported 

levels of BaP in commercial samples of fish and shellfish. BaP content was inversely related to 

the associate rates of in vitro BaP metabolism . The potential risk associated with parent PAH 

carcinogens in seafood would be greater with invertebrates, which are less able to metabolize 

these compounds. There would still be risk associated with consumption of fish from highly 

contaminated sites . While fish may contain a lesser and even undetectable amounts of parent 

compound, carcinogenic metabolic products of these may be present. 

It is difficult and often disconcerting to draw inferences from the small samplings, but 

the data do point to some of the recognized problems in biomarker research. These problems 

are : 1) samplings of fish species from large areas are quite variable from time to time and place to 

place and 2) biochemical responses of fishes are extremely variable as a function of species, sex, 

gonadal status, season, temperature, nutrition etc. Thus, field studies can result in a 

disconcerting array of data, even though controlled laboratory studies may be very predictable. 

This study sought to investigate the differences of hepatic EROD activity between pristine and 

contaminated areas. Significant delineation might ultimately be useful for quantitative or 

qualitative employment in a biomonitoring regimen. The objective of biomarker research is to 

establish the efficacy of the use of the sentinel. 
33 

CHAPTERCh

STUDIES OF PRO- AND ANTIOXIDANT PARAMETERS IN 

LIVER CYTOSOL AND MICROSOMES OF GULF OF MEXICO 

FISH SAMPLES AND OTHER MARINE ORGANISMS: 

EVALUATION OF SUPEROXIDE DISMUTASE, CATALASE, 

MICROSOMAL GENERATION OF OXYRADICALS, AND 

OXYGEN CONSUMPTION 

G.W. Winston, S. B. Dobias and R. Pautz 

INTRODUCTION 

Oxygen radicals are produced as a normal part of all aerobic life . Molecular oxygen is 

reduced by four electrons with the resulting production of two molar equivalents of water for 

each oxygen molecule that is reduced. Thus, partially reduced reactive oxygen species (ROS) 

are produced which can be greatly enhanced under exacerbative conditions of xenobiotic 

exposure. Nicotinamide adenine dinucleotide phosphate, reduced form (NAD(P)H)-dependent 

electron transport by microsomes from virtually all vertebrate and invertebrate species studied 

produces ROS including superoxide anion radical (02'), hydrogen peroxide (H202), and 

hydroxyl radical (-OH) (Jewell and Winston, 1989; Winston et al., 1991). Each of these species 

have been implicated in toxic damage to cells including cell death (Kappus and Seis, 1981). 

Hydroxyl radicals have been shown to participate in the oxidation of certain xenobiotic 

substrates including aliphatic alcohols, benzoate (Winston and Cederbaum, 1982), aniline 

(Ingelman-Sundberg and Ekstrom, 1982), benzene (Ingelman-Sundberg and Hagbjork, 1982) 

and salicylates (Grootveld and Halliwell, 1986). Thus, their production in biological systems is 

an important, often overlooked, consideration in the response of organisms to various 

xenobiotics. The oxidation of a variety of -OH scavenging agents e.g., 2-keto-4-

thiomethylbutyric acid (KMBA) and benzoate have been used to detect the production of potent 

oxidants generated by microsomes of crustaceans, molluscs, fish and mammals (Winston and 

DiGiulio, 1991). The sequence of reactions believed to be involved in the production of H202 

in microsomal systems is outlined below. 

202 + NA.DPH M'crosomes 

202- + NADP+ + H+ (1) 

202 + 2H+ H202 + 02 (2) 

Eq. [2] shows the disproportionation of superoxide resulting in the production of 

hydrogen peroxide. Hydrogen peroxide can serve as the precursor of -OH via the Haber-Weiss 

reaction (Eq. 3) (Haber and Weiss, 1934). 

02 + H202 02 + -OH + OH' (3) 
34 

This reaction is kinetically quite slow, therefore transition metals, especially iron 

generally are invoked as a catalyst for -OH production (Eqs. 4, 5) . 

Fe(III) + 02- 02 + Fe(II) (4) 

Fe(II) + H202 Fe(III) + -OH + OH- (5) 

The sum of Eqs. [4 ] and [5] is the Haber-Weiss reaction (Eq. 3). Eq. [5] is the Fenton 

reaction (Walling, 1975). 

Metal chelates, nitroaromatics, quinones and bipyridyls (Mason, 1979; Kappus and Seis, 

1981 ; Bus and Gibson, 1984) are known to increase ROS production in biological systems with 

accompanying oxidative damage. The toxic sequalae of these various classes has been linked to 

the proliferative generation of oxyradicals that results from these compounds uncoupling 

biological electron transport . The consequence of ROS production is protein degradation, 

enzyme inactivation, lipid peroxidation, DNA damage and cell death (Borg and Schaich, 1984) . 

Two possible loci of 02 production in microsomes are the autooxidation of the oxycytochrome 

P450 complex (Dybing et al., 1976) and/or the autooxidation of the cytochrome P450 reductase 

(Winston and Cederbaum, 1986). Winston and Cederbaum (1986) showed that the oxidation of 

-OH scavenging agents was dependent only upon the amount of P450 reductase in a 

reconstituted MFO system and not the amount of cytochrome P450. In contrast to mammalian 

systems, little work has been done on the mechanism involved in the production of ROS in 

submammalian vertebrates and invertebrates . A characteristic of invertebrate microsomal 

electron transport is the comparatively low specific activity of NAD(P)H-cytochrome c 

reductase, which is about 3-10 % of mammals (James et al., 1979; Jewell and Winston, 1989) . 

The presence of oxyradicals does not preclude physiological damage as they are limited 

by the presence of antioxidant enzymes e.g., superoxide dismutase (SOD) and catalase 

(Diguiseppi and Fridovich, 1984) . Therefore the levels of these antioxidants were assessed as a 

function of produced water discharges in parallel with MFO and genotoxic assessment. The 

hydroxyl radical scavenging agents 2-keto-4-thiomethylbutyric acid (KMBA) and dimethyl 

sulfoxide were oxidized to ethylene and formaldehyde, respectively by microsomes isolated 

from the hepatopancreas of the Red Swamp crawfish (Jewell and Winston, 1989), the common 

marine mussel (Livingstone et al ., 1989 ; Winston et al ., 1990) and the Salmon (Kennish, 1989). 

METHODS 

Studies have been carried out to measure superoxide dismutase (SOD) and catalase levels 

in various species as a function of the conditions described in the previous 3 chapters with the 

newly developed SOD-525 assay from Bioxytech (Cedex, France) . This is the newest system 

for measurement of SOD in biological tissues . The assay exploits the enhanced rate of 

autoxidation of a patented pyrogallol derivative, which offers some distinct advantages over 

previously published assays . One advantage is that confounding substances, which are both 

replete and variable in biological samples and can interfere with detection of SOD, can be diluted 

from the sample without loss of enzyme activity . It is also highly reproducible from one 

laboratory to another which suggests its utility for wide geographical distribution . Preliminary 

studies designed to optimize the concentrations of microsomal and cytosolic fractions of marine 

mussels and sea anemones on the end-point chromaphore (oxidized pyrogallol derivative ; kmax = 

525nm) have been completed. Interferences are noted only in some concentrated biological 

materials but can be eliminated by dilution of samples 1 :10 or greater. 

RESULTS AND DISCUSSION 

Table 4.1 below shows the effects of 3-methylcholanthrene (3MC) exposure on the 

specific activity of SOD in the sea anemone Anthapleura elegantissima . This anemone species is 
35 

known to contain a1ga1 symbionts, which have been shown by others to impose oxidative stress 

on the anemone resulting in elevated SOD levels (Dykens et al ., 1992) . Thus xenobiotic 

exposure might enhance the oxidative stress placed on this organism which may already have its 

antioxidant defenses compromized by the photosynthetically-generated oxygen via 

photosynthesis by the zooxanthellae. It can be seen in Table 4.1 that SOD levels are highest in 

the uv-light-exposed anemones which is consistent with adaptation by the organism to the 

hyperoxic conditions produced by photosynthetically generated oxygen; SOD activities are about 

equivalent and about 30-40% lower in the dark-reared and symbiont-depleted anemones. These 

data corroborate the findings of Dykens et al . (1992) . Depletion of zooxanthellae resulted in 

lower SOD levels in the 3MC-treated anemones, however growing zooxanthellate anemones in 

the dark to prevent photosynthesis did not cause significant changes in SOD activity . This may 

be the result of oxidative stress caused by the 3MC exposure or a response by the zooxanthellae 

to 3MC. The general lack of responsiveness of SOD to 3MC exposure indicates the probable 

ineffectiveness of SOD response for use in biomonitoring, at least with anemone sentinels. 

Table 4.1 SOD content of sea anemone (Anthapleura elegantissima) cytosols: 

Effect of 3-methylcholanthrene (3MC) exposure and zooxanthellate symbionts. 

Conditions SOD (units/ml) 

Controls 

symbionts uv-reared 341 ± 10.7* 

symbiont-depleted 231 ± 11 .4 

symbiont dark-reared 214 ± 14.3 

3MC-exposed 

symbionts uv-reared 258 ± 29.0 

symbiont-depleted 199 ± 13.6 

symbiont dark-reared 226 ± 10.0 

* Mean±SD of 2-3 determinations on pooled cytosol from 5-8 anemones. 

NAD(P)H-dependent microsomal electron transport is a known source of oxyradical 

production in biological systems. The oxidation of a-keto-y-thiomethylbutyric acid (KMBA) to 

ethylene is catalyzed by microsomal electron transport in the presence of iron-EDTA. The latter 

is used as a catalyst of the Fenton reaction to reduce hydrogen peroxide to the hydroxyl radical, 

which is the terminal oxidant in the reaction . Certain nitroaromatic compounds enhance 

oxyradical production by microsomes via a redox cycling reaction, which is linked to univalent 

reduction of the vitro group by microsomal flavoprotein reductases . Group IIB metals, 

especially Cd and Hg are inhibitors of enzymes that contain sulfhydryl groups (-SH) at the active 

site . Flavoprotein reductases, (e.g ., cytochrome P450 reductase) are such enzymes and their 

inhibition by mercurials is classical. Furthermore, To better understand the effects of exposure 

to Group IIB metals on microsomal electron transport in marine invertebrates a pilot study was 

conducted in which sea anemones (Anthapleura elegantissima) were exposed for 5 days to 25 

and 250 gg/liter of CdC12. The temperature was maintained at 25°C and salinity at 32 ppt. 

Following this exposure period microsomes were prepared by differential centrifugation and the 

ability of these microsomes to catalyze NADPH-dependent nitroaromatic reduction assessed . 4-

Nitroquinoline N-oxide (4NQ0) was used as a model redox cycling nitroaromatic because it is 

well characterized in the marine invertebrate literature and is known to stimulate oxygen 

consumption and KMBA oxidation by microsomes from Mytilus edulis digestive gland 

microsomes. 

The data shown in Figure 10 represent preliminary findings of the effect of Cd exposure 

on NADPH-dependent microsomal reduction of 4NQ0 as measured by oxyradical generation, 

i .e . KMBA oxidation . It is noted that microsomes isolated from anemones exposed to either 25 
36 

or 250 ug/liter of CdC12 reduce 4NQ0 at rates significantly less than nonexposed controls . The 

lower rates of 4NQ0 oxidation are reflected in lower rates of KMBA oxidation and hence, 

ethylene production. Microsomal yields from anemones are very low as compared to those 

obtained from organs such as digestive gland or liver. This coupled with small amounts of 

starting material resulted in exhausting the supply of microsomes from the Cd study. The 

preliminary data warrant further study of the effects of metal exposure on microsomal electron 

transport. A tentative working hypothesis for future study of these effects is that Cd binding to 

active site -SH groups results in inefficient electron transport in the microsomal fraction of the 

exposed animals. 

800 

Q 

m 

Y 6000 -~- Control 

--+- 250 ~Lg/L Cd 

---~-- 25 gg/L Cd 

.: 

Y ca 400 

d 

a v 

d 2Q0 

>+ 

t 

W 

0 10 20 

Minutes 

Figure 10. Nitroaromatic-stimulated oxyradical production by sea 

anemone microsomes: Effect of Cadmium preexposure. 

A separate preliminary study to compare NADH-dependent with NADPH-dependent 

KMBA oxidation by sea anemone microsomes in the presence of 4NQ0 was conducted. With 

NADPH as primary electron donor KMBA oxidation was 1 .33±0.14 nmol/min/mg microsomal 

protein and with NADH 0.96±0.11 nmol/min/mg. Based on numerous ethylene standard curves 

1560 area units =1nmo1. 

Exposure of sandworms to sand mixed with petroleum hydrocarbon-contaminated 

sediment (50:50 w/w) for 14 days did not increase SOD levels above the detection limits of the 

assay. This lack of response of SOD in gut tissue cytosolic fractions of the sand worm, Nereis 

virens indicates its ineffectiveness as a marker of sediment contamination. 
37 

Table 4.2 SOD Content of Gut Cytosol from Nereis virens exposed to contaminated sediment. 

Days exposed °Io contamination SOD (units/ml) 

0 0% 0.26±0.16 

14 0% BDL 

14 20% BDL 

14 50% BDL 

BDL = below detection limits 

Results of field sample analysis of SOD in microsomes of several species of fish from 

various stations representing varying degrees of impact in of the Gulf of Mexico are shown in 

Table 4.3 . There it is noted that microsomal SOD activity is below the detection limits or very 

near the detection limits of the assay for all samples tested . These data confirm the 

ineffectiveness of SOD measurements in fish as biomarkers of exposures to the kinds of 

contaminants in these studies in the field. 

Table 4.3 Microsomal SOD content of Gulf of Mexico fish samples 

Location Species SOD-525 units/ml 

ST 34, 200m++ 

ST 34, 200m 

ST 34, SOOm 

ST 34, 1000m 

ST 34, 2000m 

ST 52, SOOm 

ST 52, 1000m 

ST 52, 1000m 

ST 52, 2000m 

ST 52, 2000m 

Lake Champagne 

Lake Champagne 

Pass Fourchon 

3MC* 

Fringed Flounder 

Fringed Flounder 

Sea Robin 

Pogy 

Sea Robin 

Sea Robin 

Sea Robin 

Fringed Flounder 

Fringed Flounder 

Sea Robin 

Hardhead Catfish 

Anchovy 

Anchovy 

Fundulus adult 

BDL 

BDL 

BDL 

4.31±0.47 

0.03±0.01 

BDL 

BDL 

BDL 

BDL 

BDL 

BDL 

BDL 

BDL 

BDL 

BDL = below detection limits ; * Laboratory exposure to 3-methylcholanthrene 

++ numerical values distances in meters from platforms. All trawels were west of 

(down-current from) platforms. 
38 

Figure 11 establishes the efficacy of the SOD-525 assay as confirmed by its robustness 

in detecting with great precision the units of SOD activity of a wide range of concentrations of a 

commercially available preparation of SOD from bovine liver (Sigma Chemical Co., St. Louis, 

MO). 

15 

y 

10 

7 

LA O 

N 

c 5 

O 

N 

y -_ 0.577 + 0.598x R^2 = 0.992 

0 5 10 15 20 25 

Purified SOD units 

Figure 11. Detection of purified SOD by SOD-525 assay. 

SOD determinations of fish liver cytosol were conducted. These data are presented in 

Table 4.4 where it can be seen that SOD activities were markedly higher than in the microsomal 

compartment. The activity expressed as units/ mg cytosolic protein ranged from less than 1 or 

below detection limits in Fringed flounder taken from 200 m from ST 53a platform to 530 

units/mg in Shoal flounder caught 500 m from ST 34. Fringed flounder sampled from 200 m 

from ST 34 had one of the higher SOD levels . In the few cases where antioxidant enzymes were 

shown to correlate at all with distance from a platform an inverse relationship was observed. 

This was noted also for cytosolic catalase (see Table 4.5) . 
39 

Table 4.4 Superoxide dismutase activities of liver cytosol from fish 

sampled from various Gulf of Mexico field sites. 

Sample # Species Location site SOD 

(units/mg protein) 

170 Fringed Flounder ST 54 t 26.1 

172 Fringed Flounder ST 53 17 .6 

173 Bay Whiff ST 55 43 .4 

174 Fringed Flounder ST 54 31 .6 

175 Bay Whiff ST 52 29 .2 

176 Bay Whiff ST 53 21 .0 

177 Fringed Flounder ST 54 26.2 

178 Fringed Flounder ST 52 10.4 

301 Catfish ST 34 100 m * 28 .7 

302 Catfish ST 34 SOm 5 .9 

303 Catfish ST 34 1000m 25 .8 

304 Catfish ST 34 50m 17 .0 

305 Catfish ST 34 50m 41 .5 

306 Catfish ST 34 1000m 20.5 

307 Catfish ST 34 1000m 16.6 

308 Fringed Flounder ST 53a 200m 0 .23 

309 Fringed Flounder ST 53a 200m ND 

311 Fringed Flounder ST 34 200m 437 

312 Fringed Flounder ST 53a 1000m 103 

313 Shoal Flounder ST 53a 1000m 514 

314 Bay Whiff ST 53a 1000m ND 

315 Shoal Flounder ST 34 1000m 33.3 

319 Sea Robin ST 34 1000m 62.3 

320 Bay Whiff ST 34 1000m 334 

322 Shoal Flounder ST 34 SOOm 530 

ND = not detected . t Samples collected by Louisiana Department of Wildlife and Fisheries; 

distances not specified. * Numerical values are distances in meters (m) from platforms. 

Catalase is another important determinant of oxidative stress ; this enzyme along with 

SOD is common to all forms of aerobic life . Together SOD and catalase prevent the 

accumulation of certain partially reduced oxygen species such as superoxide anion radical, 

hydrogen peroxide and thus the formation of potent oxidants such as the hydroxyl radical . 

Catalase activities were measured in all samples for which cytosol and microsomes were 

available. As was noted for the SOD results, catalase measurements are also extremely variable. 

In general catalase activities are higher in cytosols than in microsomes, albeit some overlap in the 

values between the two compartments exists . The range of variability for the microsomes was 

ca. 2 to 850 nmol/min/mg protein and for the cytosol ca. 6 to 22,000 . 
40 

Table 4.5 Specific activities of Catalase in Gulf of Mexico fish liver microsomes. 

Species Location/Distance Cor. Ccef.* Microsomal Protein S. A.** 

(mdml) (nmoUmin/mg) 

LCt Catfish 0.980 2 11 .8 

LC Catfish 0.999 0.6 29 .0 

LC Catfish 0.994 209 6 .8 

LC Croaker 0.970 0.5 20.4 

PF$ Croaker 0.996 1 .7 60.7 

LC Pinfish 0.996 0 .97 16.0 

PF Pinfish 0.934 2 .6 1 .9 

PF Jack 0.999 0 .28 280.0 

PF Jack 0.988 0.42 118.0 

Shad ST CI 500 0.999 1 .62 17.2 

Hardhead Catfish ST 34 SOm 0 .998 42.3 12.7 

Hardhead Catfish ST 34 SOm 0.997 45.2 8.7 

Hardhead Catfish ST 34 SOm 0.994 38.2 4.0 

Hardhead Catfish ST 34 100m 0.998 36 .3 191 .1 

Fringed Flounder ST 34 200m 0.999 28 .0 97.3 

Fringed Flounder ST 34 SOOm 0.997 12 .4 36.6 

Shoal Flounder ST 34 SOOm 0.999 3.2 206.4 

Hardhead Catfish ST 34 1000m 0.982 50.5 18 .8 

Hardhead Catfish ST 34 1000m 0.999 30.3 3 .3 

Hardhead Catfish ST 34 1000m 0.998 19 .4 4 .8 

Shoal Flounder ST 34 1000m 1 .000 12.8 658 .0 

Shoal Flounder ST 34 1000m 1 .000 12.5 422.8 

Fringed Flounder ST 34 1000m 0.999 18 .6 834.6 

Fringed Flounder ST 34 1000m 0.999 17 .1 715.9 

Sea Robin ST 34 1000m 0.996 13.7 98 .8 

Bay Whiff ST 34 1000m 0.997 0.9 504.6 

Fringed Flounder ST 53a 200m 0.999 2.3 110 .0 

Fringed Flounder ST 53a SOOm 0.999 10 .1 506.9 

Shoal Flounder ST 53a SOOm 1 .000 6.4 851 .5 

Fringed Flounder ST 53a 1000m 1 .000 11 .5 416.4 

Shoal Flounder ST 53a 1000m 0.997 2.3 305 .1 

Fringed Flounder ST 36§ 0.998 2 .6 95 .3 

Fringed Flounder ST 37 0.981 2 .62 189 .1 

Bay Whiff ST 52 0.965 1 .77 174.9 

Fringed Flounder ST 52 0.799 3 .8 130.4 

Bay Whiff ST 53 0.989 5 .71 79 .3 

Bay Whiff ST 53 0.940 12 .58 7 .2 

Frin , ized Flounder ST 54 0.989 3.43 216.7 

* Correlation Coefficient. ** S .A. = Specific activity . t LC = Lake Champagne. $ PF = Pass 

Fourchon. § Fish samples from stations in which distance was not specified were collected by 

Louisiana Department of Wildlife and Fisheries. 

For the most part no obvious correlations existed between catalase activity and distance 

from point sources of offshore contamination, contaminated coastal sites vs. reference sites, 

species, collection time, depth of habitat, sediment-dwelling fish vs. demersal fish and one 

offshore site vs another. There was however a noteworthy exception. Fringed flounder caught 

1000 and 500 m from site 53 had low cytosolic catalase activity (< 2000 units/mg), whereas the 
41 

same species from 200 m had a much higher activity (8,800 units/mg). The significance of this 

trend remains to be established. 

Taken as a whole, results from catalase and SOD measurements indicate that oxidative 

stress enzymes may not be reliable markers of exposure to contaminants . Interestingly, no 

pattern of catalase activity was observed between deep-dwelling fish and fish obtained from 

shallow areas, i.e . only a few meters. The enormous variation observed in these activities 

amongst the various species studied indicates highly variable capacities to cope with oxidant 

stress . Of particular importance would be the response of antioxidant enzymes of fish subjected 

to varying periods of anoxia . In such cases injury may occur when oxygen is reintroduced after 

periods of anoxia, i.e . reperfusion injury . Thus, the variability and extreme range of antioxidant 

defenses indicated by the data suggest the importance of studies relating these enzymes to 

various conditions of oxygen concentration as well as conditions that enhance ROS production 

in aquatic organisms. 

Table 4.6 Specific activities of Catalase in Gulf of Mexico fish liver cytosol 

Species Location/Distance Cor. Coef. * Cytosolic Protein S.A. 

(mgJml) (nmoUmin/mom 

Mussel 0.992 3 .18 136 

Anemone 0.997 3 .7 24 

Fringed Flounder ST 34 500 m 0.999 0.01 3750 

Fringed Flounder ST 34 1000 m 0.999 0.12 303 

Sea Robin ST 34 1000 m 0 .995 0.19 180 

Fringed Flounder ST 34 1000 m 0 .999 0.16 284 

Pinfish LCt 0 .984 3 .2 41 

Jack PF$ 1 .000 1 .0 91 

Jack PF 0 .999 1 .7 39 

Shad ST CI 500 0 .993 6.4 27 

Catfish ST 34 50m 0 .997 40.0 206 

Catfish ST 34 50m 0.996 32.4 305 

Catfish ST 34 50m 0.997 16.8 6 

Catfish ST 34 100m 0.997 26 .9 286 

Fringed Flounder ST 34 200m 0.994 0.71 22200 

Shoal Flounder ST 34 SOOm 0.998 0.66 1276 

Catfish ST 34 1000m 0.994 25 .7 199 

Catfish ST 34 1000m 0.991 11 .8 362 

Catfish ST 34 1000m 0.993 11 .2 656 

Shoal Flounder ST 34 1000m 0.999 3 .91 1900 

Sea Robin ST 34 1000m 0.995 6 .26 208 

Bay Whiff ST 34 1000m 0.959 0.18 853 

Fringed Flounder ST 53a 200m 0.999 0.48 8810 

Fringed Flounder ST 53a SOOm 0 .997 3 .71 1410 

Fringed Flounder ST 53a 1000m 0 .997 4.12 1800 

Shoal Flounder ST 53a 1000m 0 .994 0.68 2670 

Bay Whiff ST 53a 1000m 0 .956 --- BDL 

Bay Whiff ST 52§ 0 .999 3.43 936 

Fringed Flounder ST 52 0 .988 15.5 750 

Bay Whiff ST 53 0 .0001 18.07 122 

Bay Whiff ST 53 0.988 15.5 436 

Fringed Flounder ST 54 0.995 9.93 1499 

* Correlation Coefficient. ** S .A. = Specific activity . t LC = Lake Champagne. $ PF = Pass 

Fourchon. § Fish samples from stations in which distance was not specified were collected by 

Louisiana Department of Wildlife and Fisheries. 
43 

Chapter 5 

STUDIES OF NADH- AND NADPH-DEPENDENT 

MICROSOMAL ELECTRON TRANSPORT: 

RELATIONSHIP OF CYTOCHROME C REDUCTASE AND 

OXYGEN CONSUMPTION TO SPECIES 

AND ENVIRONMENTAL CONDITIONS OF GULF OF MEXICO 

FIELD SITES. 

G.W. Winston and S.B. Dobias 

INTRODUCTION 

The microsomal mixed-function oxidase system of fish and marine invertebrates is 

comprised of various components of the microsomal membrane (Winston et al., 1991). Aside 

from the heme-containing cytochromes P450 and b5 there are also periferal flavoproteins, which 

serve as reductases for the cytochromes. These reductases are held to the membrane by a 

hydrophobic binding domain and they contain the prosthetic groups FMN and FAD. 

Cytochrome P450 reductase preferentially accepts electrons from NADPH and cytochrome b5 

from NADH. Typically, microsomal NADPH-dependent reduction of cytochrome c (used as an 

artificial electron acceptor) is taken as a measure of cytochrome P450 reductase and NADHdependent reduction is generally regarded as cytochrome bs reductase (Winston et al., 1991). 

The importance of these enzymes is emphasized by the fact that microsomal flavoprotein 

reductases are rate-limiting components of NAD(P)H-dependent microsomal mixed function 

oxygenase activities and also extremely important loci of oxygen radical production. The 

oxidation of hydroxyl radical scavenging agents was shown to be dependent soley upon the 

concentration of cytochrome P450 reductase in a reconstituted MFO system and not the amount 

of cytochrome P450 (Winston and Cederbaum ; 1986). Furthermore, cytochrome P450 

inhibitors including CO, metyrapone and SKF-525A do not inhibit oxyradical production during 

microsomal electron transport as measured by the production of ethylene from KMBA (Jewell 

and Winston, 1989; Winston et al ., 1991) . 

In mammalian systems, cytochrome P450 reductase is induced by only certain of the 

agents that classically induce cytochrome P450. Phenobarbital and Arochlor 1254 both affect 

increases in cytochrome P450 reductase activity of microsomes (Narayan and Winston, 1988), 

presumably via proliferation of smooth endoplasmic reticulum (Okey, 1990). Thus, 

microsomes prepared from liver of phenobarbital- and Aroclor 1254- pretreated rats have higher 

rates of NADPH-dependent oxygen consumption in the presence of substrates that are 

univalently reduced by flavoprotein reductases, e.g . nitroaromatics, quinones and metal chelates 

(Metosh-Dickey et al ., 1994). A characteristic of invertebrate microsomal electron transport is 

the comparatively low specific activity of NAD(P)H-cytochrome c reductase, which is about 3-

10 % of mammals (James et al ., 1979; Jewell and Winston, 1989). Moreover, relatively few 

investigations of this important rate-limiting enzyme system has been performed with 

submammalian species . Thus, we have studied both NADH- and NADPH-dependent 

cytochrome c reduction to indicate the infra and inter species range of activities and the effects of 

various Gulf of Mexico field site environments on those activities . In conjunction with the 

determination of reductase activities, attempts were made to measure the specific content of 

cytochromes P450 in an effort to determine the efficacy of the microsomal preparations and for 

use in the evaluation of the MFO studies conducted in association with this project. 
44 

METHODS 

Fish liver microsomes were isolated by differential centrifugation according to standard 

techniques in the literature (Winston et al ., 1989). NAD(P)H-cytochrome c (P450) reductase 

was measured by monitoring reduction of ferricytochrome c (Type III from horse heart, Sigma 

Chem. Co., St. Louis, MO) at 550 nm (Phillips and Langdon, 1962) and quantified from the 

millimolar extinction coefficient, 18.5 mM-1 cm-1 for ferrocytochrome c. Cytochrome P450 

specific content was determined according to Omura and Sato (1964) . 

RESULTS AND DISCUSSION 

NAD(P)H-dependent cytochrome c reductase activities were measured in liver 

microsomes of fish sampled from various offshore sites. The offshore sites were further 

subdivided as a function of distance from the platform station. Reminiscent of data obtained on 

these parameters in microsomes from the Lake Champagne and Pass Fourchon fish there was 

extremely wide interspecies and intraspecies variability. NADH-dependent cytochrome c 

reductase ranged from below detectable limits in Bay Whiff caught at ST 52 to approximately 

1000 nmol/min/mg microsomal protein in catfish caught at ST 34 (Table 5 .1). NADHdependent activities were consistently higher in catfish (870±170 nmol/min/mg, n = 7), 

irrespective of the distance from the platform at which they were caught. Fringed flounder had 

the next highest activities with NADH (83±44 nmoUmin/mg, n = 6). When the value for fringed 

flounder caught at ST 53 (9 nmol/min/mg) is eliminated from the data set the mean ± SD was 

98±31 . Fringed flounder from ST36, 37, 52, 53, and 54 had notably lower NADH-cytochrome 

c reductase activities than those from ST 53a or any of those caught from ST 34. NADPHdependent activities ranged from 0.2 nmol/min/mg in Bay Whiff caught at ST 53 and 55 to 300 

nmol/min/mg in catfish caught at ST 34. An unexpected observation was the NADH/NADPH 

ratios of < 1 .0 found in some of the Fringed flounder and Bay Whiff microsomes from the most 

recently provided samples. NADH-dependent activites are generally greater than NADPHdependent acrivites. 

Table 5.1 Microsomal NAD(P)H-dependent cytochrome c reductase activirites 

of several fish species sampled from a contaminated (Pass Fourchon) and 

reference (Lake Champagne) site in the Gulf of Mexico. 

Location NADH NADPH NADH/ 

Species nmol/min/mg NADPH 

Lake Champagne Hardhead Catfish 20 16 1 .3 

Hardhead Catfish 55 145 0.4 

Hardhead Catfish 42 46 0.4 

Pinfish 57 7 10.0 

Spot 95 48 2.0 

Croaker 14 28 0.5 

Pass Fourchon Hardhead Catfish n/a 12 -- 

Croaker 20 5 5 .0 

Pinfish 92 4 26.3 

Jack 54 34 1 .8 

Jack 8 13 0.6 

Spot 48 48 1 .0 

Spot 71 38 1 .9 
45 

Table 5.2 Microsomal NAD(P)H-dependent cytochrome c reductase activirites of 

several fish species sampled from varying distances from offshore oil platforms 

in the Gulf of Mexico. 

Distance from 

platform (meters) Species 

NADH NADPH 

nmol/min/mg 

NADH/ 

NADPH 

ST 53a 200 Fringed flounder 108 104 1 .0 

Fringed flounder 115 44 2 .6 

Shoal flounder 7 14 0 .5 

ST 53a 1000 Fringed flounder 138 30 4.6 

ST 53 Fringed flounder 9 29 0.3 

ST 53 Bay Whiff 11 38 0.2 

ST 34 50 Hardhead catfish 995 300 3 .0 

ST 34 50 Hardhead catfish 852 270 3 .2 

ST 34 50 Hardhead catfish 831 121 6.9 

ST 34 100 Hardhead catfish 1111 203 5 .5 

ST 34 200 Fringed flounder 47 91 0.5 

ST 34 500 Fringed flounder 80 70 1 .1 

ST 34 500 Shoal flounder 163 142 1 .1 

ST 34 1000 Shoal flounder 244 179 1 .4 

ST 34 1000 Bay Whiff 159 ND - 

ST 34 1000 Bay Whiff 94 54 1 .7 

ST 34 1000 Bay Whiff 245 92 2.7 

ST 34 1000 Fringed flounder 321 85 3 .8 

ST 34 1000 Fringed flounder 65 12 5 .4 

ST 34 1000 Sea Robin 16 5 3 .2 

ST 34 1000 Bay Whiff 155 226 0.7 

ST 34 1000 Hardhead catfish 993 177 5.6 

ST 34 1000 Hardhead catfish 757 179 4.2 

ST 34 1000 Hardhead catfish 555 96 5 .8 

ST CI 500 Threadfin shad 20 5 3 .6 

ST 36* Fringed flounder 4 7 0.7 

ST 37* Fringed flounder 11 36 0.3 

ST 52* Fringed flounder 5 17 0.4 

ST 52* Bay Whiff ND 27 -- 

ST 54* Fringed flounder 2 13 1 .8 

ST 55* Bay Whiff 6 38 0.2 

Laboratory-treated 3-MC Fundulus Adult 260 96 2.7 

*Collected by LDWF, distance not specified 

The specific content of cytochromes P450 were determined in all fish samples for which 

we were supplied with adequate amounts of microsomal protein. In all cases microsomal P450 

was found to be mostly denatured to cytochrome P420 (Figure 5. 1) or highly contaminated with 

other in terfering CO-bin ding chromaphores. 
46 

0.2-

0.1 - c 

.0 

0 0.0 

y 

o 

-0.2 

400 420 440 460 480 

manometers 

Figure 12. P450 CO-binding spectrum of fish #301 liver microsomes. 

In some samples in which a positive absorbance change could be measured between 490 

nm (isosbestic point for reduced, CO-liganded and reduced cytochromes P450) and 450 nm an 

attempt to calculate a residual P450 specific content was made. It is recognized that the accuracy 

of the measurements are questionable owing to the possibility of spectral interferences of other 

substances ; the above CO-binding spectrum resembles that of cytochrome oxidase and is 

believed to reflect mitochondrial contamination of the microsomal preparation. Interestingly, the 

spectra resembled more those of S9 preparations than microsomal preparations . Table 5.2 

shows cytochrome P450 specific content estimates of microsomes from fish caught at various 

offshore sites. These fish represent various distances from oil platforms. Table 5.2 indicates 

the cytochrome P450 specific content to vary from 0.027 - 0.39 nmol/mg microsomal protein. 

A microsomal preparation from a rat pretreated with Aroclor 1254, a polychlorinated biphenyls 

(PCB) mixture of coplanar and non-coplanar PCB congeners was used as a positive control; this 

preparation had a specific content of P450 of 1 .62 nmol/mg protein, which is in good agreement 

with the literature (Figure 13). 

A possible explanation for the high P420 (denatured P450) content of the fish 

microsomes is having to freeze liver samples prior to microsomal isolation. This has credence in 

light of a fresh microsomal preparation from a catfish, which showed the expected P450 COspectrum and had a specific content of about 300 pmol/mg protein. 
47 

0.06 -

v 0.04 -

c 

0.02 -

i.. 

0 0.00-

-0.02 -

-0.04 

400 420 440 460 480 

nanometers 

Figure 13 . Cytochrome P450 CO-binding spectrum in microsomes 

of PCB-induced rats . 

Table 5 .3 Cytochrome P450 specific content of sampled Gulf of Mexico fish . 

Species Location/Distance P450 Specific Content 

from platform (nmol/mg protein) 

PCB-treated rat 1 .630 

Hardhead catfish ST 34 100 m .391 

Hardhead catfish ST 34 50 m .031 

Hardhead catfish ST 34 1000 m .050 

Hardhead catfish ST 34 50 m .184 

Hardhead catfish ST 34 50 m .161 

Hardhead catfish ST 34 1000 m .205 

Hardhead catfish ST 34 1000 m .077 

Fringed flounder ST 53a 1000 m .183 

Bay Whiff ST 34 1000 m .024 

Fringed flounder ST 34 1000 m .054 

Fringed flounder ST 34 1000 m --- 

Sea robin ST 34 1000 m --- 

NAD(P)H-dependent oxygen consumption by Gulf of Mexico fish liver microsomes was 

measured by oxygen polaragraphy with a Clark-type oxygen electrode. In conjunction with 

these measurements the ability of the potent nitroarene mutagen, 4-nitroquinoline N-oxide 

(4NQ0), to stimulate oxygen consumption by microsomes was evaluated to assess one-electron 

reduction metabolism by fish microsomes. It was of principle interest to determine if the 

univalent pathway of xenobiotic reductive metabolism was altered in fish microsomes as a 

function of the various field site parameters studied. Various species of fish caught at Pass 

Fourchon and Lake Champagne as well as from offshore sites were compared. The oxygen 
48 

electrode tracings presented in Figure 14 show that all of the fish microsomes studied catalyze 

NAD(P)H-dependent oxygen consumption. NADH-dependent rates tended to be about 2 to 3 

times greater than the corresponding NADPH-dependent rates. Oxygen consumption increases 

with increasing concentrations of microsomal protein. This concentration dependence is variable 

for the different fish species. For the most part, at constant microsomal protein concentration, 

rates of oxygen consumption are proportional to the concentration of 4NQ0. Variation in the 

response to 4NQ0 is noted, i.e . in some cases 4NQ0 is saturating at lower concentrations than 

in others . 

No obvious differences were observed as a function of the field parameters. For 

example, microsomes (200 gg protein) from Hardhead catfish caught at Lake Champagne, a 

relatively pristine reference site, and from Pass Fourchon, a produced water site and 

contaminated shipping channel, catalyze NADH-dependent, 4NQ0-mediated oxygen 

consumption at similar rates and with a similar dependence on toxicant concentration (Figure 

14). Further, the effect of varying the microsomal protein concentration on oxygen consumption 

rates while holding 4NQ0 constant (100 gg) yields similar results with microsomes from Lake 

Champagne and Pass Fourchon Catfish (Figure 15). 

Nitroaromaric compounds are abundant in the marine environment and represent a source 

of oxidative stress owing to the ability of such compounds to undergo redox cycling with 

concomitant production of oxyradicals . Nitroso intermediates are common in reductive 

metabolism of nitroarenes. Attempts to assess C-nitrosoreductase activity in the cytosolic 

fractions of mussels (M. edulis), sea anemones (Aiptasia pallida) and sand worms (Nereis 

virens) were made. p-Nitrosophenol was used as a model C-nitroso (R-C-N=O) compound. 

Nitroso group reduction was measured by monitoring the disappearance of p-nitrosophenol at 

420nm. Based on this activity, cytosol from neither the worms nor the anemones reduced pnitrosophenol to p-aminophenol. M. edulis cytosol catalyzed NADH-dependent p-nitrosophenol 

reduction at a rate of -10 mM/min/mg cytosolic protein and NADPH-dependent reduction at a 

rate of -22 mM/min/mg cytosolic protein. Noteworthy is the observation that this activity is not 

stable and gradually disappears over a period of 3 hours on ice . Approximately 20°l0 of the 

activity was lost in the first hour on ice. By 3 hours approximately 50% of the initial activity 

was lost. Further experiments to determine nitro(so) reductase activities of cytosol should 

account for this time-dependent loss of activity. It is recommended that such activities be 

performed whenever possible on freshly prepared cytosolic fractions. If this is not possible then 

liquid nitrogen freezing of cytosol should be done and activities performed on samples that have 

been freshly thawed. 

As it presently stands, for use in toxicological assessment the data obtained for each of 

the microsomal mixed function oxygenase parameters on field samples are of questionable value. 

The high degrees of variability among the different species, the lack of correlation between MFO 

and field parameters, and the denaturation of MFO catalyst in microsomal preparations are all 

contrary to laboratory conducted exposure studies . Additional major problems were the 

inconsistency of the catches, which were unpredictable with respect to species, lack of 

knowledge of the sex and gonadal status of the fish at the times in which they were caught and 

variable conditions of abiotic factors such as fluctuations in oxygen tension . Variability in 

oxygen tension of the water column possibly contributed to the variability and highly 

unpredicatable nature of field samplings. Correlation of MFO parameters with oxygen tension 

measurements may be of heuristic value. 
Constants 

NADH 

1CAut~~ 4NCa 

r 

0 

25 

50 

1CD 

XD 

400 

LOO 

Mic=oscmal protein concentration varied (ug) 

:51 LC Catfish 

NA,DS I 0 

200ug u5'" " I 

4~~0 Concentration varied (um) 

451 LC Catfish 

NADPH 

'1COurt 4.1QO X10 

I 

:51 LC Catfish 

0 

NADPH 

2GOug uS y,D 

49 

lOJ 

0 

50 

1CC 

~ASx 

'*53 LC Catfish *49 PF Catfish *`61 P° Pinfish 

- I 

0 Q 

lp.3x 

i 

I 

Micosomal protein concentration varied (uq) 

*53 LC Catfish 149 PF Catfish *61 P.° ?inEish 

0 0 

1m M 1QO 

1CD . 3'7D . lp 

4NQO Concentration varied (vM) 

DL\DPH 

200ug u5 

#65 LC Spot #7a Stud *66 PF Soot ;6'7 or Pinfish 

o 

§N 

0 

O 0 

3'D . XD 

- 

2M 

*Each squar e vertical ax--- ------o consumed full scale; 

horizontal axis = 15 min . toll scale; "

* US = mi¢ascmal protein 

Figure 14 . Po(aragraphic tracings of 4NQ0-mediated oxygen consumption 

during microsomal electron transport. 
50 

Constants i53 LC Catfish ;49 PF Catfish 

NIADH 

200ug u5* 

\~ ~ ~ 0 

50 

107 

2M 

5M 

. 4N(p Concentration 

1453 LC Catfish 

NADH ~ 0 0 

100uM 4NQ0 100 100 

230 203 

400 

400 

30 Rzn. fu11 smle 

u s poteici 

Clzrnaltsatiaz . 

. *US = microsomal protein 

Figure 15 . Comparison of NADE-i-dependent oxygen consumption catalyzed by 

LC vs PF catfish : Effect of varying microsomal protein microsomes 

and 4NQ0 concentrations . 
51 

CHAPTER 6 

Part A 

EVALUATION OF THE BIOCONCENTRATION POTENTIAL OF 

GENOTOXIC PETROLEUM INDUSTRY EFFLUENTS 

USING CO-SOLVENT MOBILIZATION . 

J.C. Means, D.J. McMillin and L.A. Reily 

INTRODUCTION 

Characterization of petroleum-related wastes have been part of the scientific literature for 

a very long time. It is well established that environmental petroleum releases from such facilities 

as produced water separation facilities, marketing terminals and production facilities contain a 

very complex mixture of normal and branch chained hydrocarbons and a complex mixture of 

both volatile (benzene, toluene, xylenes and ethyl benzene) and semi-volatile (naphthalenes, 

phenanthrenes, fluorenes, etc.) parent aromatic hydrocarbons but also alkylated analogs of these 

aromatic compounds as well as various heterocyclic aromatic structures such as 

dibenzothiophenes and aza-arenes (Neff et al., 1987). Among these aromatic hydrocarbons are 

several compounds which by virtue of their low aqueous solubilities are likely to bioaccumulate 

in food chains in aquatic systems (Capuzzo et al., 1987 ; Rabalais et al., 1991b). However, 

detailed quantification of the genotoxic potential of many of these constituents has been limited. 

Once these substances are discharged, they are subject to a variety of physical transport 

processes . Further, at the molecular level many of the water insoluble or particle reactive 

substances in petroleum industry effluents may undergo partitioning reactions with suspended 

particulates, colloidal solids and flocculating materials contained in the effluents themselves as 

well as the receiving waters . While many of the constituents of hydrocarbon-related wastes are 

volatile and/or partially biodegradable and therefore present a limited risk to aquatic organisms, 

there is ample evidence to suggest that the vast majority of the semi-volatile components of 

petroleum are relatively refractory and are therefore likely to accumulate in sediments and 

biological tissues. Means et al . (1980 and 1982) showed that PAHs and heterocyclic analogs of 

PAHs were strongly sorbed to sediments and that the relative strength of sorption could be 

predicted from compound aqueous solubilities . Further, we have shown that these same 

compounds may sorb to colloidal organic matter in the water column thus facilitating transport 

and possibly inhibiting bioaccumularion (Means and Wijayaratne, 1982; Wijayaratne and Means, 

1984; Sigleo and Means, 1990 and references therein) . 

Chemical analyses of environmental samples have frequently been directed toward 

meeting some regulatory criteria (eg. NPDES permitting requirements) and therefore much of the 

available literature on hydrocarbon contamination has been limited to studies which report the 

concentrations of only a relative few of the aromatic hydrocarbons potentially present in a 

petroleum industry waste (16 priority pollutant PAHs + 1-methyl and 2-methyl-naphthalene). 

While these compounds may be present as components in the total aromatic fraction of petroleum 

industry waste, recent studies of petroleum-related wastes such as produced waters, suggest that 

these 18 compounds may represent less than 5°Io of the total aromatic materials present. While 

the genotoxicity of a few of these compounds such as Benz[a]pyrene and benzanthracene have 

been extensively studied and reported, the potential for genotoxic impacts of the petrogenic 

compounds is poorly characterized. 
52 

The hydrocarbon load from petroleum industry effluents is normally stated as one 

relatively crude number called oil and grease. Detailed characterizations of what compounds 

comprise this number are not readily available. The EPA conducted two studies in developing 

guidelines for effluent discharge limitations (USEPA 1979 and 1982). There were seventeen 

petroleum industry discharge sources that were sampled and analyzed for priority pollutants. 

Table 1 lists the compounds that were detected in at least one discharge. The majority of these 

compounds were actually detected in less than five discharges . Most other detailed analyses of 

petroleum industry discharges have also focused in on the EPA priority pollutant list . Although 

one study conducted in western Australia investigated petroleum-derived hydrocarbons in a 

petroleum industry effluent and in organisms 250 meters away from the discharge (Alexander et 

al ., 1982a and 1982b). They found both in the discharge and in mullet and mussel tissue : 

naphthalene, methylnaphthalenes, dimethylnaphthalenes, trimethylnaphthalenes, phenanthrene, 

methylphenanthrenes and dimethylphenanthrenes . 

Table 6A.1 Priority pollutants commonly detected in treated 

petroleum industry effluents 

Volatiles Base/Neutral 

Extractables 

Methylene chloride Naphthalene 

trans- l,2-Dichloroethene Dimethyl phthalate 

Chloroform Fluorene 

1,2-Dichloroethane Diethyl phthalate 

Bromodichloromethane Phenanthrene 

Trichloroethene Anthracene 

Benzene Di-n-butyl phthalate 

Tetrachloroethene Fluoranthene 

Toluene Pyrene 

Ethyl benzene Benz(a)anthracene 

Chrysene 

Acid Extractables Bis(2-ethylhexyl) phthalate 

Benzo(a)pyrene 

Phenol Benzo(g,h,i)perylene 

2,4-Dimethylphenol 

2,4-Dichlorophenol 

4-Chloro-3-methylphenol 

4-Nitrophenol 

* EPA (1979, 1982) 

We proposed that a more detailed analysis of the petrogenic components as presented in 

the present study could yield valuable information on the potential for adverse genotoxic effects 

upon aquatic organisms exposed to these complex mixtures. Methodologies developed as pan 

of this study and reported in Chapter 7 were employed to analyze of petroleum-related wastes 

for over 55 specific parent and heterocyclic PAHs and their alkylated homologues. These 

methods were utilized to characterize the exposure concentrations in sediments or water and the 

bioaccumulated residues found in exposed biomonitoring organisms. These data are reported in 

Chapter 3. 
53 

While these data provide an improved understanding of the exposure potential for 

petroleum-related wastes such as produced waters, they do not allow for the assessment of 

genotoxic risks of that exposure. Over the last two decades, investigators have utilized whole 

extracts of soils and sediments to evaluate genotoxic potential of particle-associated toxic 

substances . Typically these methods have employed exhaustive extraction of the sediment or 

soil with an organic solvent followed by concentration of the extract. The final extract was 

typically transferred to another solvent compatible with various bacterial or cell culture 

genotoxicity assays such as dimethylsulfoxide and applied to the assay at different dose levels to 

establish the presence or absence of genotoxic agents in the mixture (Dexter et al ., 1979; Suzuki 

et al., 1982, Osborne et al., 1982 and Daniels et al., 1985). These and numerous other studies 

have reported dose-related responses in a number of genotoxicity assays . Among the problems 

reported in these investigations are the cytotoxicity of many of the extracts, the complexity of the 

mixtures, the high background (positive genotoxicity responses) of extracts from 

uncontaminated soils and sediments. 

A basic conceptual limitation of these experiments has been the inability of these types of 

studies to relate results to actual risk. The limitation is that there has been no way to relate data 

on genotoxicity of sediment extracts to bioavailability and bioaccumulation potential of 

organisms living in contact with contaminated sediments, thus establishing risk. This study 

sought to combine elements of models of bioaccumulation with models of sediment/water 

partitioning theory to evaluate the utility of co-solvent extracts from sediments to evaluate 

genotoxicity in a way that can be related back to the bioavailability of the contaminants and 

therefore provide a more realistic assessment of genotoxic risk to aquatic organisms . 

THEORY 

Sediment Partitioning 

The equilibrium partitioning of organic contaminants in sediment/water systems has been 

the focus of investigation for a long time. Polycyclic aromatic hydrocarbons and their alkylated 

analogs, and various classes of heterocyclic aromatics such as dibenzothiophenes and 

dibenzocarbazoles all have been investigated Means et al., 1980, Hassett et al ., 1980, and 

Banwart et al ., 1982). In all cases, these compounds partition between the organic carbon phase 

of the sediments and water in a predictable way based upon the aqueous solubility of the 

individual compound. Another predictive compound property was found to be the octanol-water 

partition coefficients of the compounds. Wijayaratne and Means (1984) established that similar 

partitioning occurs between organic-rich colloidal phases found in aquatic systems and water. In 

more recent studies, Means and McMillin (1993) showed that this partitioning was completely 

reversible in estuarine sediments and that the kinetics of desorption were sufficiently rapid to 

allow exposure of benthic organisms living in contact with the sediments. 

Figure 16a shows the relationship between octanol-water partition coefficients (log P) 

and the values of sediment-water partition coefficients normalized for the organic carbon content 

of the sediment (log Kim). This relationship has the following equation : 

log Koc =1og P - 0.317 (Means et al ., 1980) 

Bioavailability of Selected Trace Organic Contaminants: Many of the trace metals 

and organic substances which have been found in previous examinations of petroleum 

discharges are known to be toxic both to aquatic organisms and to man NAS, (1985) . In 

particular, many of the aromatic organic compounds are known or suspected mutagens or 

carcinogens EPA (1982) . Therefore, a key aspect of a complete ecological and human risk 
54 

assessment for these discharges must address the bioavailability and bioaccumularion potential of 

the contaminants found in these discharges . 

One approach which has been used in many such assessments has been the collection 

and analysis of indigenous populations of non-migratory species in the area of a discharge site 

and in a spatial gradient away from that site . Bivalve mollusks such as the blue mussel (Mytilus 

edulis) , the oyster (Crassostrea virginica) and the freshwater Asiatic clam (Corbicula fluminea) 

have been used successfully in several large-scale biomonitoring studies in estuaries and coastal 

marine environments (Farrington et al ., 1983, Neff et al ., 1976, NOAA, 1988). These and 

other studies have established the scientific validity of such biomonitoring efforts. However, 

adequate data on the bioavailability and bioaccumulation of petroleum industry-associated 

discharges of aromatic organics remain largely incomplete . 

There remains a certain degree of uncertainty concerning the bioavailability of sedimentassociated contaminants, particularly with respect to suspended particulates . Since many of the 

toxic organics found in petroleum wastes are highly particle-reactive and sorbed to suspended or 

bedded sediments, a more detailed examination of the bioavailability of these contaminants is 

required . 

The vast majority of detailed bioaccumulation studies have been performed using 

aqueous phase exposures of a wide variety of aquatic organisms including fish, crustaceans, 

mollusks, etc . . From these investigations a variety of predictive equations have been derived 

which relate the octanol-water partition coefficient of the compound to the bioconcentration 

potential (BCF) of the compound in tissues . Many of these relationships have further been 

refined and predictive capability improved by normalization of the quantities bioaccumulated in 

the organisms to the lipid content of the tissues examined or the whole organism. Figure 16b 

shows one such relationship . In this case, the slope of the regression equation between log P 

and log BCF is near unity just like the sediment-water partitioning relationship discussed earlier. 

If we assume that exposures of aquatic organisms in sediment/water systems involves a 

sequential process in which the contaminant must first desorb from the sediment and then is 

bioavailable to be accumulated in the organism, then the two predictive relationships shown in 

Figure 16 can be combined to predict sediment exposures. In this figure, log Koc refers to the 

organic carbon-normalized sediment/water partition coefficient, log BCF (lipid) refers to the 

lipid-normalized bioconcentration factor, and log P refers to the octanol/water partition 

coefficient. 

Investigations have been performed by several research groups to establish a predictive 

relationship between sediment concentrations and bioavailable contaminant levels in organisms 

living in association with sediments. These experimental studies have suggested that, for a 

limited number of neutral hydrophobic organic compounds, including aromatic hydrocarbons, 

an exposure level can be estimated (McElroy and Means, 1988 and references therein, Foster, et 

al ., 1987) from the sediment concentration normalized for organic carbon and the lipid content of 

the test organism using the following equation : 

AF- (toxicant sed. conc ./TOCI 

[toxicant tiss . conc ./lipid] 

where AF is an accumulation factor which is an empirically determined constant equal to -2. 
55 

LOG Koc 

LOG P (LOG Kow) 

LOG BCF 

(lipid) 

LOG P (LOG Kow) 

Figure 16. Relationship of Log P to Log Koc and Log BCF. 
56 

Co-solvent Theory Applied to Sediment Partitioning: Co-solvents are defined as 

water miscible organic solvents which when mixed in varying proportions with water result in 

alteration of the solubility of a wide variety of organic chemicals. Typical co-solvents include 

short-chain organic acids, alcohol, ketones, aldehydes, etc. As the fraction of co-solvent added 

to water increases, the solubility in the liquid phase of hydrophobic compounds such as PAH, 

chlorinated hydrocarbons, and even dioxins increases in a log-linear fashion. The slope of the 

relationship is dependent upon the co-solvent and the solubility of the compound. 

Rao et al . (1985) combined co-solvent theory with sediment-water partitioning theory to 

attempt to develop predictive equations for the enhanced transport of organic contaminants in 

sub-surface soil environments. Means et al . (1988) investigated the effects of several cosolvents on the sediment-water partitioning of PCBs and DDT. In that study, they concluded 

that co-solvents predictably reduced the organic carbon normalized sediment-water partition 

coefficients (Koc) for several neutral hydrophobic compounds in a log-linear fashion. They 

further observed that of the organic co-solvents evaluated, the degree to which the hydrophobic 

compounds were solubilized followed the following sequence: propionic acid>acetic 

acid>acetone>ethanol>methanol . 

Figure 17 shows the effect of three co-solvents upon the Koc of the model compound 

2,4,2',4'-tetrachlorobiphenyl . As the mole fraction of each of the co-solvents was increased, 

the log Koc value decreased in a linear fashion. In each case, the regression lines intersect the 

Y-axis at a point corresponding to the log Koc measured or predicted for pure water. Figure 18 

shows the sorption and desorption equilibria for the compound on two different sediments with 

acetic acid as the co-solvent. Importantly, both the Y-axis intercept and the slopes of the 

regressions remain relatively constant from sediment to sediment and regardless of the mode of 

approach to equilibrium (sorption vs. desorption). 

The experimental results may be fit to the equation : 

log (Kacm/Kocm,w) _ -a6f 

where KoCM, w and KoCM are the mole-based, organic carbon-normalized partition coefficient for 

the water and co-solvent systems, f is the mole fraction of the co-solvent, and the product as 

represents the slope of the line . These constants relate solute-soil ((X) and solvent-soil (6) 

interactions . 
6- 

Y1 = 5.6048 - 8.2414x R"2 - 0.990 

Y2 = 5.4207 - 13.245x R"2 = 0.974 

0 

Y3= 5.5614 - 26.731x R"2 a 0.987 

5- 

13 PCB 

v 

0 Y 

0 o Y1aMethand 

J 

Y2=AceUc Add 

o Y3=Proplonic Acid 

3- 

2 -. 

0.0 0 .1 0 .2 0 .3 0.4 

mole traction 

Figure 17. Effect of mole fraction of three co-solvents on desorption Koc values for hexachlorobiphenyl `' 
Y1 = 5.2585 - 11 .979x R"2 a 0.989 

Y2= 5.1719 - 11 .00I x R"2 = 0.979 

e 

s 

Y3 = 5.4344 - 13 .93I x R"2 = 0.987 

Y4= 5.4207 - 13.245x R"2 = 0.974 

v 0 Y 4- 

0 CO 

J 

pS~ o 

o Y1=sed3 sorp 

0 Y2=sed3 desorp 

o Y3.sed4 sorp 

Y4-sed4 damp 

2 

0.0 0.1 0.2 0.3 

mole fraction acetic acid 

00 

Figure 18. Effect of mole fraction acetic acid on Koc for hexachlorobiphenyl 
59 

Figure 19 shows the behavior of three compounds in a single co-solvent systems. With 

a given co-solvent such as acetic acid, the slopes of the regression lines remain constant while 

the Y-axis intercepts change in response to magnitude of the equilibrium partitioning constant 

Koc for the individual compound. 

Co-Solvent Theory And Predictive Assessment Of Bioaccumulatable Mutagenic 

Compounds In Sediments: Since bioaccumulation potential has been observed to be 

strongly correlated with the octanol-water partition coefficient of a compound (log P), especially 

when tissue concentrations at steady-state (log BCF) are expressed on a lipid-normalized basis. 

Similarly, since sediment water partition coefficients normalized to the organic carbon content of 

the sediment (log Koc), which have been inversely related to bioaccumulation potential, are also 

strongly correlated with log P. Bioavailability of contaminants sorbed to sediments have been 

modeled as two-step process involving first, desorption from sediment into interstitial or 

overlying water, followed by bioaccumulation from the aqueous phase. 

Total extracts of sediments/soils using non-polar solvents have been used to determine 

the total concentrations of contaminants in sediments but have been shown to be poor predictors 

of bioaccumulation potential from the sediment, due to differences in the relative bioavailability 

of different pools of the contaminants within the organic and inorganic matrix of the sediment 

particles . Total extracts do not reflect differences in bioavailability due to differences in log P of 

the several contaminants contained in a sediment. Further, the contribution of relatively watersoluble mutagenic compounds may be underestimated when total extracts are used to assess 

sediment mutagenicity or toxicity. 

Co-solvent partitioning behavior of compounds, in which the log Koc for a compound 

decreases in a predictable log-linear fashion with increasing mole fraction of co-solvent, may be 

used to assess the relative desorprion potential of different classes of sediment-bound chemicals. 

The resulting extracts contain amounts of different chemicals, not only in proportion to the total 

amounts present in the sediment but also in relation to their relative desorption potential from 

sediment and therefore in relation to their bioavailability and bioaccumulation potential . In 

addition, relatively water-soluble mutagenic compounds are included in the extracts. A further 

advantage of this approach is that a relatively non-toxic organic solvent such as acetic acid can be 

used which is volatile for concentration purposes and which is not mutagenic in bioassay 

systems like many chlorinated solvents. 

If the co-solvent theory is working as expected in the system, then we would expect to 

see water soluble compounds extracted from the sediment present in the low co-solvent fraction 

extracts and declining in concentration at higher co-solvent fractions. Conversely, we would 

expect neutral hydrophobic compounds extracted from the sediments, such as aromatic 

hydrocarbons, to be very low in low fraction co-solvent extracts and more abundant in higher 

co-solvent extracts . 

Figure 20 shows the concentrations of two model compounds determined in a series of 

acetic acid extracts with increasing mole fraction co-solvent. As expected, bisphenol, a relatively 

polar aromatic compound, was detected in highest amounts in the 0% acetic acid extract and at 

progressively lower concentrations in extracts with higher co-solvent content. Methyl 

naphthalenes, an abundant class of petroleum-associated alkylated aromatic compounds, were 

not detectable in either the 0% or 10°Io acetic acid co-solvent extracts but increased dramatically 

in concentration in 20% to 50°Io acetic acid co-solvent extracts . 
60 

EFFECT OF ACETIC ACID MOLE FRACTION ON KOC OF THREE 

HYDROPHOBIC ORGANICS 

log Koc 

mole fraction acetic acid 

Figure 19 . Effect of acetic acid co-solvent concentration upon the Koc of three 

model hydrophobic compounds . 
35 

30 

~ 25 

Q. 

~ 20 

N 15 

1C 

' " , --~-- BISPHENOL, ppb 

-o- METHYLPHENANTHRENES, ppb 

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., 

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0 , Q 

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1 

COSOLVENT 

50 

Figure 20 . Co-solvent mobilization of selected pollutants in a 

contaminated sediment . 
62 

Based upon the theory described and the basic confirming results from chemical analyses 

of co-solvent extracts of petroleum-contaminated sediments, an experiment was designed to 

evaluate the mutagenicity of co-solvent extracts of Pass Fourchon sediments using a series of 

acetic acid co-solvent extracts assayed in the Ames microbial mutagenicity test. 

EXPERIMENTAL METHODS 

Sediment Extractions 

Approximately 50 g of wet Pass Fourchon sediment were weighed and placed in a 120 

mL glass bottle along with 100 mL of extraction co-solvent . The co-solvents used were 0%, 

10%. 20°Io and 50°Io acetic acid in distilled water. Extractions were performed by rolling the 

bottle containing sediment and solvent together on a roller mill for 24 hrs, separating the phases 

by centrifugation and decanting the solvent. This process was repeated two times and the three 

extracts were combined. The solvents were concentrated to 50 ml by rotary evaporation such 

that 1 ml of extract would contain compounds extracted from approximately 1 g of wet sediment. 

These 1 ml aliquots of extracts were evaporated to near dryness under a stream of nitrogen and 

taken up in 1 ml of dimethylsulfoxide (DMSO). These DMSO solutions were used to apply the 

extracts to petri dishes for the mutagenicity assays . 

Mutageniciry Testing 

Mutagenicity of the co-solvent extracts of Pass Fourchon sediment was assessed using 

the Ames microbial mutagenicity assay, revised as described in Maron and Ames (1983) . 

Several tester strains were used, including strains TA97a, TA98, TA100, and TA102. Each 

extract was assayed both in the presence and absence of a rat S-9 microsomal activating system 

to evaluate the presence of direct-acting and promutagens in the extracts . Each extract was 

evaluated over a range of doses corresponding to 0 to 50 mg of sediment (50 ul of appropriate 

extract dilutions in DMSO, applied to the series of plates). Plates were scored for revenant 

colonies per plate after incubation for 72 hr at 37 C. Results were reported as revenants per plate 

per mg of sediment. 

RESULTS AND DISCUSSION 

The Ames mutagenicity assay employs auxotrophs of Salmonella typhimurium which 

contain errors in the gene which codes for conversion of biotin to the essential amino acid 

histidine (Ames, 1975; Maron and Ames, 1983). The occurance of additional mutations within 

the gene DNA following exposure to a test mutagen, can cause shifts in the genetic reading 

frame which restore cells to a histidine-autotroph genotype. This process is known as reversion. 

Revenant colonies which grow in the absence of histidine may be enumerated as an index of the 

mutagenic potential of mixtures or pure compounds to which they were exposed. The 

mutagenicity of the cosolvent extracts was observed, to assess the various types of mutational 

injury detectable with the array of five tester strains. The experiments were also conducted with 

and without metabolic activation of promutagens by induced rat liver S9. 

Consistent dose-response relationships are prerequisite to the use of Ames test data for a 

particular tester strain/mutagen assay system. Further, in order for the proposed co-solvent 

system to work, the extracts must yield consistent results in the assay system so that results can 

be replicated within reasonable limits of variability . Only then, can the results be used to 

extrapolate to real exposure levels (risks) associated with a particular sediment using the 

proposed casolvent/bioaccumulation model. In order to test the level of replication achieved in 

these co-solvent experiments, assay series which yielded linear dose-response curves were 

assembled and calculated on a revertant/plate/mg basis for a range of doses tested on different 
63 

plates corresponding to 0 to 50 mg of sediment assayed using two co-solvent system extracts 

(0% and 50% acetic acid) in strain TA 102 and a rat S-9 activating system . Adequate replication 

was present in the system, yielding similar values of revenants/plate/mg sediment for all 

sediment doses applied to plates. 

Figure 21 shows the responses obtained in this experiment. It is clear that, except for 

the dose equivalent to 2 .5 mg of sediment from the 50% co-solvent extract, the results are 

reproducible for both water soluble and 50% co-solvent soluble pro-mutagens. This suggests 

that extrapolations using the model can be made with reasonable confidence. 

Both direct acting and promutagenic substances were observed in the extracts from Pass 

Fourchon sediment, however, the direct acting substances were only detected with strain TA 

102. Figure 22 shows the direct acting mutagenic activity of the four co-solvent extracts from 

Pass Fourchon. It should be noted that in this form the data for spontaneous reversions in these 

strains when calculated on an equivalent sediment dose level fall into a range of <1 

revertant/plate/mg. 

Both water soluble and more hydrophobic (eg. 20% co-solvent) direct acting mutagenic 

compound groups were detected with TA 102. A decrease in mutagenicity was observed in the 

10% co-solvent extract relative to all other extracts tested, suggesting that the extraction of water 

soluble mutagens was suppressed in this extract. This is consistent with the results of chemical 

analyses reported before . The highest response was observed at 20°Io co-solvent . The relative 

decrease seen in the 50% co-solvent extract may be due in part to enhanced cytotoxicity of this 

extract to the bacteria. 

Mutagenic compound groups which required metabolic activation with S-9 were detected 

in all three bacterial assay strains utilized . Of the three strains employed, TA 102 was the most 

responsive to the promutagens extracted from this sediment. Figure 23 shows the responses of 

strain TA97a to the four co-solvent extracts. Comparable but low levels of mutagenic activity 

were detected in the 0°Io and 10°Io co-solvent extracts . A three-fold increase in mutagenic activity 

was detected in the 20% co-solvent extract, suggesting that there were more hydrophobic than 

water soluble pro-mutagens which responded in this strain . A great deal of cytotoxicity was 

observed with the 50% co-solvent extract in this strain which suppressed responses in the assay. 

Figure 24 shows the mutagenicity observed in strain TA 100. In contrast to the systems 

reported thus far, the greatest response in this assay was observed with the water soluble extract 

which contained very high levels of promutagens. The extraction of these promutagens was 

completely suppressed in the 10% co-solvent extract. Low levels of more hydrophobic 

promutagens were progressively extracted in the 20°Io and 50°Io co-solvent systems . No 

cytotoxicity was observed in this case with the 50°Io co-solvent extract. This observation 

suggests that the toxicity observed in other strains may be due to very high mutation frequencies 

rather than general cytotoxicity of the extract. This is a fairly common observation in assays of 

complex mixtures. 

Figure 25 shows the responses observed in strain TA 102 with the four extracts. As 

noted earlier, the promutagens in this sediment elicited the greatest response in this assay strain . 

Very high frequencies of mutation were observed in the water soluble extract. Unfortunately, 

the 10% co-solvent extract was lost just prior to this assay and therefore no results for this 

extract were obtained. High levels of mutation were also seen in both the 20% and 50% cosolvent extracts applied to this strain . The slight decrease in mutagenicity was interpreted as 

being to the very high mutation rates leading to reduced survivorship on these plates . 
10, 

Dose Response for Sediment Extracts 

350 << ;~ 

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300 - 

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250 - 

d 

r 

200- 

CL 

a 

" 150 I 50% 

FD 

100 - L 

50 - 0% 

Co-Solvent 

0 0 .5 2 .5 5 25 50 

equivalent mg of sediment 

Figure 21 . Dose response of mutagens extracted from sediments 
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69 

Identification of Problems Associated with Measurement Methods: As with the 

measurement of any complex mixture of chemicals in environmental media, there are potential 

interferences which may influence the results. In the general case, these include but are not 

restricted to matrix effects in the sediments themselves such as the presence of extractable salts, 

nutrients and other dissolved solids (including colloids) or particulates . The presence of 

substances which interfere with the accurate detection and quantification of mutation frequencies 

or which result in matrix effects in the receptor assay organisms (effects on lipids, proteins and 

other biomolecules) may result in detection problems. For example, the presence of the free 

amino acid histidine or metabolizable peptide fragment containing histidine in sediment extracts 

may result in false positives in the Ames assay. In the case of petroleum industry effluents, all 

of these potential interferences are possible . It should be noted that all extracts, regardless of the 

solvent system used have the potential to contain such interferences. Further investigation of 

these potential interferences will need to be undertaken before the co-solvent extract model can 

be applied widely to soil/sediment bioassays. 

Application of the Combined Co-Solvent/Bioaccumulation Model to Results: The 

results of the the mutagenicity assays for a given co-solvent extract reported in units of 

revenants/plate/mg sediment may be converted to an equivalent water exposure frequency by 

application of co-solvent theory and using the equation above and a value for the product of the 

constants as for acetic acid of 12 (Means et al ., 1988). Using an accumulation factor of 2 as 

described above for the bioaccumulation potential (McElroy and Means, 1988), an estimate of 

the bioaccumulatable mutation frequency for the sediment-bound contaminants may be expressed 

but now the units would be revenants/plate/mg lipid in the organisms tissues. 

CONCLUSIONS 

Based upon the results of these experiments several conclusions were developed. They 

were as follows : 

" Direct-acting mutagens were mobilized at all co-solvent concentrations but were detected 

only with swain TA 102 . 

" Mutagen mobilization into co-solvent extracts was detectable by the Salmonella 

typhimurium assay, with tester strains varying in sensitivity. 

" Promutagens were detectable in all extracts, via activation with Arochlor-induced rat liver 

S9. 

" Mutagenicity effects varied among co-solvent concentrations, perhaps reflecting the 

mobilization of different arrays of compounds. 

" Low co-solvent fractions appeared to enhance extraction of water-soluble mutagens but 

suppress the extraction of lipophilic mutagens. In high co-solvent fractions, extraction of 

lipophilic mutagens was significantly increased. 

" Increasing co-solvent fraction appeared to suppress the extraction of some water-soluble 

mutagens. 

" TA 102 was the most responsive tester strain with these extracts and yielded very 

reproducible results. TA 98 showed no response to mutagens in these extracts. 

" Chemical analyses of the co-solvent extracts by GC/MS may yield information on the 

identity of possible mutagens present. 
71 

CHAPTER 6 

Part B 

STUDIES OF GENOTOXICITY AND POTENTIAL FOR MUTAGENIC 

ACTIVATION IN FISH SAMPLED 

FROM THE GULF OF MEXICO AS EVALUATED BY THE UMU 

MUTAGENICITY ASSAY 

G.W. Winston, S.B. Dobias and L. A. Reily 

INTRODUCTION 

A major objective of this project is to identify genotoxic effects in correlation with the 

various biochemical responses from contaminant exposure. Preliminary studies in the Ames 

assay suggest mutagen mobilization into the water column. Previous studies in this laboratory 

(Winston et al ., 1989) indicated that liver microsomes from channel catfish (Ictalurus punctatus) 

caught from a contaminated river basin swamp near Baton Rouge (Devil's Swamp) were more 

effective catalysts of promutagen activation with certain xenobiorics than microsomes prepared 

from the same species obtained from the relatively pristine Ben Hur Reseach Facility at LSU, 

which we used as a reference site . Therefore, the effects of exposures to sediments on the 

indigenous organism's ability to modulate the mutagenicity of the benchmark promutagens 2-

aminoanthracene (2AA) and 2-acetylaminofluorene (2AAF) in the umu gene induction assay 

were tested . 

METHODS 

The ability of microsomes from Lake Champagne catfish and Pass Fourchon catfish to 

activate 100 N.M 2AA was compared with that of liver microsomes from control rats and from 

rats pre-treated with 3-methylcholanthrene (3MC), an aromatic hydrocarbon known to induce 

enzymes that activate these promutagens in the rat. The rat samples were run with 200 gg of 

microsomal protein and the fish with 50, 100 and 200 ~tg of protein. 

The activation of 2AA and 2AAF to mutagenic products was determined by measuring 

the expression of the umu gene in Salmonella typhimurium TA1535/pSK1002 as described in 

Shimada and Nakamura (1987) . Total R-galactosidase activity toward O-nitrophenyl-R-Dgalactoside was taken as a measure of the expression of the chimeric umu DC-lac Z gene product 

and was thus a measure of mutagenicity (Kirchin and Winston, 1992). 

Co-solvent extracts were provided by Dr. Reily, Department of Veterinary Pharmacology 

and Toxicology, Louisiana State University School of Veterinary Medicine. These consisted of 

aqueous extracts in 10%, 20°Io and 50% acetic acid (w/v). Aqueous extracted material was 

concentrated in dichloromethane (DCM), evaporated to dryness under nitrogen and reconstituted 

in DMSO for sue in the mutagenicity assay. 

RESULTS 

Figure 26 shows the results of mutagenicity assays designed to test the ability of 

microsomes from Lake Champagne and Pass Fourchon catfish to activate 100 PM 2AA. These 

data are compared with liver microsomes from untreated control rats and rats pre-treated with 3-

methylcholanthrene (3MC), an aromatic hydrocarbon known to induce enzymes that activate 

various promutagens in mammals and fish . A 1 .6-fold and 2.4-fold enhancement of umu gene 
72 

induction with microsomes from control and 3MC-treated rats was observed, respectively, but 

no increase was observed over background controls with the fish samples. 

We have also shown that the cytosolic fraction of rat liver contains an amine oxidase, 

which is capable of activating certain arylamines to mutagens in the Ames mutagenicity assay 

(Traynor et al, 1992). Therefore experiments were conducted with cytosolic fractions from 

Croaker (Pass Fourchon), Shad (Lake Champagne) and Pinfish (Lake Champagne) liver to 

determine if fish liver cytosolic fractions catalyzed arylamine activation in the umu mutagenicity 

assay and if so, was such catalysis altered in fish from contaminated sites. For these studies 400 

gg and 800 gg of cytosolic protein were tested separately. No significant increase in gene 

induction was observed over background (Figure 27). 

Experiments were repeated in parallel with 200 gg of microsomal protein from control 

and 3MC-treated rats as positive controls . Control and 3MC-treated rat liver microsomes 

catalyzed 3- and 3 .5-fold enhancement of gene expression with 2AA however, as in the previous 

experiments, no enhancement by 2AA of gene induction was seen with fish samples (Figure 

28) . 
ao 

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Figure 28. Arylamine activation by fish liver cytosol. 

Additional experiments were performed to determine if chemicals extracted by the cosolvent method (Methods) from contaminated sediment obtained at Pass Fonrchon were capable 

of umu gene induction in the umu mutageniciry assay. In each of these cases there was no umu 

gene induction observed with any of the samples. 

CONCLUSIONS 

The umu gene induction assay has been widely used to assess the genotoxicity of 

numerous toxicants including complex mixtures (Shimada and Nakamura, 1987). Thus, the 

umu gene induction assay may not be sensitive enough to evaluate mutagens in sediment 

extracts. Typically, promutagens that are activiated by microsomal fractions or S9 fractions in 

the Ames assay are also activated in the umu gene induction assay (Shimada and Nakamura, 

1987). It is also known that polycyclic aromatic hydrocarbon metabolism catalyzed by fish liver 

microsomes is often one to three orders of magnitude lower than that of rat liver microsomes 
76 

(Stegeman, 1981). Presently, it cannot rigorously be established if the relative lack of sensitivity 

of the Salmonella ryphimurium TA1535/pSK1002 strain to extracted mutagens reflects 

susceptibility of the strain to the promutagens in the sediments or a problem of activation 

threshold. Future experiments will address these possibilities. 
77 

ChapterCH

DEVELOPMENT AND APPLICATION OF 

(PARTS-PER-TRILLION) GC/MS ANALYSIS METHODS FOR 

APPROXIMATELY 60 AROMATIC HYDROCARBONS 

D.J. McMillin and J.C . Means 

ANALYTICAL METHOD DEVELOPMENT 

Rationale For Target Analyte Selection 

The compounds of interest in the study of impacts of petroleum production were selected 

based upon knowledge of 1) the composition of petroleum related waste effluents, 2) the 

potential of certain types of compounds to bioaccumulate in aquatic indicator organisms and 3) 

the potential of such compounds to cause adverse effects upon components of ecosystems 

receiving discharges of these compounds on a chronic basis. 

Much of the chemical analysis of environmental samples has been directed toward 

meeting some regulatory criteria and therefore much of the available literature on hydrocarbon 

contamination has been limited to studies which report the concentrations of only a relative few 

of the aromatic hydrocarbons potentially present in petroleum industry wastes (16 priority 

pollutant PAHs + 1-methyl and 2-methyl-naphthalene) (USEPA, 1984, 1986) . While these 

compounds may be present as components in the total aromatic fraction of petroleum refinery 

waste, recent studies of petroleum-related wastes, such as produced waters, suggest that these 

18 compounds may represent less than 5% of the total aromatic materials present. 

The hydrocarbon load from petroleum wastes is normally stated as one relatively crude 

number called oil and grease . Detailed characterizations of what compounds comprise this 

number are not readily available. The EPA conducted two studies in developing guidelines for 

discharge limitations for petroleum refinery effluents (USEPA 1979 and 1982) . Seventeen 

refinery discharge sources were sampled and analyzed for priority pollutants . The majority of 

these pollutants were actually detected in less than five discharges . Most other detailed analyses 

of refinery discharges have also focused in on the EPA priority pollutant list, although one study 

conducted in western Australia investigated petroleum-derived hydrocarbons in a refinery 

effluent and in organisms 250 meters away from the discharge (Alexander et al ., 1982a and 

1982b) . Naphthalene, methylnaphthalenes, dimethylnaphthalenes, trimethylnaphthalenes, 

phenanthrene, methylphenanthrenes and dimethylphenanthrenes were found both in the 

discharge and in mullet and mussel tissue . 

In detailed investigations of the composition and environmental fate of hydrocarbons 

assocated with produced water discharges into fresh, estuarine and marine aquatic ecosystems, 

we have observed that these effluents are very complex mixtures of aliphatic, aromatic, alkylated 

and heterocyclic aromatic hydrocarbons (Rabalais et al ., 1991a, Means et al ., 1990, Means et 

al ., 1989) . In order to conduct a successful biomonitoring program, one necessary precursor 

must be the development and verification of sensitive and compound/element specific analytical 

techniques for application in tissue analysis. We have developed and applied such techniques 

for the analysis of petroleum-related wastes in water, sediments and tissues . To date these 

methods have been utilized in well over 3,000 analyses and the requisite QA/QC practices and 

measures have also been developed and are reported here. 
7s 

The overall data objective of our biomonitoring studies is to determine the presence and 

amounts of bioconcentratable organics present in petroleum-related waste streams and receiving 

environments . In order for this data objective to be accomplished, several criteria for the 

methods must be met, including: 

*a wide dynamic range (-0.01 ppb to 100 ppm)-7 orders of magnitude in concentration 

*compound specificity-the ability to discriminate individaul compounds from both 

biological and geological matrices . 

*compound quantitation- the ability to determine the concentrations of specific 

compounds, as opposed to estimates made by class 

*sufficient "fingerprinting" capability to be able discriminate between possible multiple 

sources of similar contaminants 

*the methods should be designed to allow for frequent sampling and analysis to be 

performed 

*methods should require a minimum of procedural steps to minimize potential for losses 

or contamination. 

Table 7 .1 lists the selected target analytes and the mass fragment used for quantitative 

evaluation ("primary ion") and for confirmation of identity ("confirming ion"). Each of these 

analytes was selected from a more extensive list (over 300 potential analytes) based upon the 

availability of standards, their real or potential presence in effluents and/or previous 

bioaccumulation studies conducted in estuaries in south Louisiana and California. Each of the 

pollutants selected has documented or suspected implications for the ecological health of coastal 

marine ecosystems and also potential human health implications through foodwebs leading to 

man which exist in these ecosystems. In previous and ongoing studies of the fate and transport 

of produced water hydrocarbons under MMS funding, our research group has been able to use 

these techniques to follow both the chemical signature of a discharge for up to 1500 m and longrange pollutant transport in the Gulf of Mexico (Rabalais et al ., 1991a, Means and McMillin, 

1993, Murray, 1994). 

As with the measurement of any complex mixture of chemicals in environmental media, 

there are potential interferences which may influence the results. In the general case these 

include but are not restricted to matrix effects in the discharges themselves such as the presence 

of salts and other dissolved solids (including colloids) or particulates ; the presence of substances 

which interfere with the accurate detection, identification or qualification of other substances (oil 

and grease); matrix effects in the receptor organisms (lipids, proteins and other biomolecules) ; 

and contaminants having sources other than the facility in question which enter the samples 

either at the point of discharge, during sampling or during analysis . In the case of petroleum 

industry effluents, all of these potential interferences are possible and probable. Techniques 

used to overcome or minimize these problems include: 

*the use of Mass Spectrometry as a means of positive identification combined with 

*high-resolution capillary column gas chromotography optimized for resolution of 

individual sample/standard components 

*extensive quality control procedures, including blanks, spikes, duplicates and the 

analysis of standard reference materials 

*target analytes isolation through extraction and sample cleanup steps including solvent 

exchange and Silica gel fractionation. 
79 

Table 7.1 Selected aromatic hydrocarbon analytes 

Analyte Primaa Ion Confirming Ion 

Polvnuclear Aromatic Hydrocarbons 

Naphthalene 128 129 

2-Methylnaphthalene 142 141 

1-Methylnaphthalene 142 141 

2-Ethylnaphthalene 156 141 

1-Ethylnaphthalene 156 141 

2,6/2,7-Dimethylnaphthalene 156 141 

1,3/1,7-Dimethylnaphthalene 156 141 

1,6-Dimethylnaphthalene 156 141 

1,4/2,3-Dimethylnaphthalene 156 141 

1,5-Dimethylnaphthalene 156 141 

Acenaphthylene 152 153 

1,2-Dimethylnaphthalene 156 141 

2-Isopropylnaphthalene 170 155 

1,8-Dimethylnaphthalene 156 141 

1,6,7-Trimethylnaphthalene 170 155 

Acenaphthene 153 154 

Fluorene 166 165 

Dibenzothiophene 184 185 

Phenanthrene 178 179 

Anthracene 178 179 

4-Methyldibenzothiophene 198 197 

2/3-Methyldibenzothiophene 198 197 

1-Methyldibenzothiophene 198 197 

3-Methylphenanthrene 192 191 

2-Methylphenanthrene 192 191 

4/9-Methylphenanthrene 192 191 

1-Methylphenanthrene 192 191 

3,6-Dimethylphenanthrene 206 191 

3,5-Dimethylphenanthrene 206 191 

2,6-Dimethylphenanthrene 206 191 

2,7-Dimethylphenanthrene 206 191 

3,9-Dimethylphenanthrene 206 191 

1,6/2,5/2,9-Dimethylphenanthrene 206 191 

1,7-Dimethylphenanthrene 206 191 

1,9/4,9-Dimethylphenanthrene 206 191 

1,2-Dimethyldibenzothiophene 212 211 

Fluoranthene 202 101 

1,5-Dimethylphenanthrene 206 191 

1,8-Dimethylphenanthrene 206 191 

1,2-Dimethylphenanthrene 206 191 

9,10-Dimethylphenanthrene 206 191 

Pyrene 202 101 

1,2,8-Trimethylphenanthrene 220 205 

Benzo(a)anthracene 228 226 

Chrysene 228 226 

Benzo(b)fluoranthene 252 253 

Benzo(k)fluoranthene 252 253 

Benzo(a)pyrene 252 253 

Indeno(1,2,3-cd)pyrene 276 278 

Dibenz(a,h)anthracene 278 276 

Benzo(g,h,i)perylene 276 278 
80 

Laboratory methodology 

The following is a description of sample preparation methods for various sample 

matrices that have been developed or modified for use in the compound-specific analysis for 

aromatic hydrocarbons of petrogenic origin . The methods were applied in the present study for 

sediment and tissue samples (Chapters 3 and 8, respectively) . 

Tissue Sample Preparation 

Tissue samples for bioaccumulation studies (see Chapter 8) were defrosted overnight in 

a refrigerator. Individual samples were removed from the bags and rinsed with tap water, then 

shelled on a glass cutting board with a Teflon coated knife. The shell-free flesh was rinsed 

with organic free water and then chopped into small pieces. The pieces were placed in a clean jar 

and homogenized using a Tekmar TissumizerO. The homogenized samples were kept frozen at 

-20°C until sample extraction . 

The extraction method is a modification of that described by MacLeod et al . (1985) . 

Approximately 20 g of defrosted homogenized tissue was placed in an 240-m1 amber glass bottle 

with 100 ml nanograde@ dichloromethane (DCM) and four times the tissue weight of cleaned 

sodium sulfate (DCM rinsed) . A surrogate spike of 200ng each of a mixture of deuterated 

standards (US-108, Ultra Sientific) was added to the bottle . The sample was extracted with a 

Tekmar TissumizerO for 2 min at a speed of 24,000 rpm. The probe was rinsed with DCM and 

the rinsings added to the sample bottle . The extract was allowed to settle for 1 min and then 

filtered through a funnel filled with sodium sulfate into a labelled florence flask. The sample 

was extracted as above with two additional 100 ml volumes of DCM. After the third extraction, 

the extract was rinsed with an additional 30 ml of DCM, which was added to the flask. The 

sample extract was reduced in volume to approximately 2 ml with a rotary evaporator . The 

concentrated extract was quantitatively transferred to a 4-ml vial and concentrated to a volume of 

2 ml under a stream of purified nitrogen . Reagent blanks, duplicates and matrix spiked samples 

were extracted to monitor laboratory technique and extraction efficiency. Matrix spikes were 

duplicate tissue samples to which 100 ng of the quantitative calibration standard were added 

instead of the deuterated surrogates mixture. 

Most tissue samples contain a layer of fine white precipitate (extracted proteins) which 

was removed before silica column cleanup . The extracts were "cleaned" by adding 500 g1 of 

hexane, mixing the extract well and then centrifuging for 5 min at 3,000 rpm. The solvent layer 

was transferred to a clean 4-ml vial and the extract was "cleaned" twice more with 1 ml of 

hexane. All solvent layers were combined and reduced in volume to 1 ml under a stream of 

purified nitrogen . 

One half of the extract was separated from the polar lipids on 200 mm x 10 mm columns 

packed with 6.0 g of 100-200 mesh Grade 923 silica, eluted with 100 ml of 60% DCM and 40% 

hexane. The extract was concentrated under nitrogen to 200 g1, after which 2 ~Ll of 2-

fluorobiphenyl (100 ng/g1 in hexane) was added as an internal standard to monitor the GC/MS 

injection performance. 

Sediment Sample Preparation 

Sediment cores (see Chapter 3) and water/filter samples were sealed with electric tape in 

the field and packed on ice for transport to the laboratory. Once in the laboratory, the cores were 

stored at 5°C. until processed, within one week The top 8 cm of sediment from each core was 

placed in a clean glass jar with a teflon-lined cap, mixed, and refrigerated until sampled for 
81 

analyses, then stored at -20°C. The extraction method was a modification of that described by 

MacLeod et al . (1985) . Excess water was decanted from sample jars prior to thorough mixing 

of the sediment sample, and 15 g was weighed into an amber bottle with a teflon-lined cap. The 

sediment was covered with 75 ml of dichloromethane (DCM) and 50 g sodium sulfate 

incorporated until the sediment mixture was free-flowing . A suite of deuterated PAH was added 

(800 ng in hexane of six deuterated PAH) with the first solvent aliquot as a surrogate standard to 

monitor extraction efficiency. The jars were sealed and placed on a roller for 16, 6, and 16 

hours, for a total of three extractions using the modified rock tumbler and rolling rimes specified 

by MacLeod et al. (1985) . The DCM from each jar was decanted into an amber jar and 

refrigerated until 3 extracts were collected. The decanted DCM was filtered through sodium 

sulfate, and concentrated by rotoevaporation and a nitrogen stream to a final volume of 200 g1, 

with solvent exchange to hexane . Activated fine granular copper was added to reduce sulphur 

interferences. 

Water Sample Preparation 

Water samples (20 L) were collected in 20L polypropylene cubitainersTM (Cole-Parmer, 

Inc.), and transferred to a stainless steel compressed nitrogen pressurized vessel to facilitate 

filtration through a 149 mm 0.4 gm membrane filter with glass fiber prefilter. This concentrated 

the particulate phase on the membrane filters, which were stored in small glass jars held at 5°C. 

The filtrate was ultrafiltered using an Amicon Model DC10 equipped with hollow-fiber 

cartridges (MW cutoff 3000 daltons) to separate dissolved and enriched colloidal phases . The 

colloidal fraction was concentrated to 240 ml and stored in amber glass jars at 5°C. Prior to SPE 

extraction 20 .1 of the deuterated surrogate standard at 40 ppm in methanol was added to 4L of 

the ultrafiltrate (dissolved phase) . A two liters aliquot was pumped through an SPE disk 

(EmporeTM, C18) housed in a teflon filter holder attached in line with a MasterFlex pump set for 

a flow rate of 2-3 ml/min. Extraction volume was determined by pumping time. The SPE disk 

was removed from the filter holder and stored in a small glass vial covered with 10 ml of 

methanol. 

Particulate Phase: The membrane and glass fiber prefilters were placed together in taxed teflon 

digestion bombs. The first extraction solvent was 45 ml of 2 :1 hexane acetone. A surrogate 

standard was added (120 ng in methanol of six deuterated PAH and dibromooctafluorobiphenyl) 

at the same time as the first extraction solvent, and the bombs placed in an ice-cooled sonicating 

bath for 6 min. After decanting, two more extractions were performed using 45 ml of hexane 

only. The extracts were combined and acetone removed by adding >75 ml of purified water. 

The top layer of hexane was removed by pipet and filtered through clean sodium sulphate . 

Subsequent concentration steps were the same as for sediment extracts, excluding the addition of 

copper. To obtain the particulate fraction weight, the teflon bombs containing the filter sets were 

dried at 60°C overnight in an oven modified to exchange the atmosphere with charcoal-filtered 

air. The weight of the filters and bombs was recorded and the samples sent to the inorganics 

sample preparation laboratory for acid digestion. 

Colloidal Phase : Samples were transferred to separatory funnels, surrogate standard added (400 

ng in methanol of six deuterated PAH and dibromooctafluorobiphenyl-DBOFBP) and extracted 

with DCM in triplicate, first at ambient (neutral) pH, then at acidic pH (2). The extracts from 

both pH levels were combined and the extract concentrated as for sediment samples, without the 

addition of copper . 

Dissolved Phase: An additional laboratory surrogate standard (120 ng each of DBOFBP, 1,3-

dimethyl-2-nitrobenzene, and triphenylphosphate in methanol) was added to the vials containing 

SPE disks to monitor laboratory performance. The disks were extracted three times with 20m1, 

lOml, and lOml of added DCM and using a sonicating bath for 1 minute per extraction. The 

extracts were combined in a separatory funnel and the DCM separated from methanol (storage 
82 

solvent) and residual water by adding excess water (>25m1) and decanting the DCM layer over 

clean sodium sulphate . Concentration steps were similar to colloidal samples. 

Instrumental Analysis 

Analytical Approach 

The primary analytical tool employed in these studies was a gas chromatograph/mass 

spectrometer (GC/MS) system for organic analyses . This instrument system has the advantage 

that multiple compounds may be quantified simultaneously on a single sample. Repetitive 

analyses are not required. The use of multiple selected ion mass spectrometry on sample extracts 

allow the quantification, at sub-ppb levels, of several classes of pollutants in a single analysis 

without extensive sample fractionation or cleanup techniques. Thus, both pyrogenic and 

petrogenic PAHs may be analyzed simultaneously. 

A Hewlett-Packard 5890 Gas Chromatograph equipped with a 30m by 0.25mm ID, 

0.25~tm DB-5 capillary column (J&W Scientific) is directly interfaced to a Hewlett-Packard 

5970B Mass Selective Detector (MSD). The GC is temperature programmed from 50° to 300°C 

using a series of linear temperature ramps to optimally separate the analytes . The mass 

spectrometer is tuned prior to calibration every 16 hours using Perfluorotributylamine (PFTBA). 

Retention order of the alkylated PAH isomers has been determined in our laboratory by 

analysis of individual "true" components obtained from Chiron Laboratories A>S> (Norway). 

Following initial determination of retention order, the resolution between isomers was optimized 

by adjusting the temperature ramp rates, while considering the effects of band-spreading. Table 

7 .1 presents the analytes in order of elution from the GC column, and also indicates resolved 

versus co-eluting isomers (positional indications separated by a "/" mark). 

Quantitative reference standards were used for daily calibration of the instrument . 2-

fluoro-biphenyl was added as an internal standard to each sample aliquot to monitor performance 

of the GC/MS instrument and injection precision . Standard response curves were prepared 

using 5 to 9 calibration points over several orders of magnitude of concentration for each 

analyte. 

Linearity. Dynamic Range, and Detection Limits 

Linearity and dynamic range of the GC/MS using the SIM method described above was 

demonstrated by two independently acquired standard curves . The first was a 5-point curve 

with concentrations ranging from 0.02 ng to 2.0 ng, or three orders of magnitude, and with each 

point replicated three times. A 9 point curve of concentrations ranging from 0.005 ng to 10 ng, 

with two replications of each point, represented over four orders of magnitude. Table 7.2 

shows the regression statistics and calculated X-intercept (ng) values and response factors for 

both curves . Table 7.3 shows LOD and LOQ values (Keith et al ., 1983) calculated from the 

average standard deviations between replicates for both curves . The 5-point curve is reproduced 

for routine verification of linearity and detection limits in our laboratory, since most sample 

values fall in this range. Also, column overload and detector shut-downs occur when values 

exceed 5-10 ng on column. Thus, the 9-point curve showed lower conelations than did the 5-

point curve. The correlarions for the 9-point curve were improved when a second-order function 

was calculated, and indicates losses in response were occuring at the highest concentration 

points . Detection limits calculated from LOD values for the target analytes are presented in Table 

7.4 showing results for all matrices routinely analyzed. 
83 

Validation Study 

Method detection limits in an environmental matrix were verified using seawater samples 

spiked in triplicate at 5, 10, 50, and 100 parts-per-trillion . Samples (2000 ml) were processed 

using the C18 SPE disks as described above after spiking with the standard solution prepared in 

methanol. Spike concentration corresponded to 0 .1, 0.2, 1 .0, and 2.0 ng on-column, 

respectively, based on concentration of extracts to a final volume of 200 ~.1 and injection of 2g1 

sample onto the GC/MS. Summary statistics are shown in Table 7.5 . All aromatics were 

detected at the lowest spike concentration in all three replicate samples, with the exception of 

Indenopyrene, Dibenzanthracene, and Benzoperylene. Recovery of spiked components 

averaged 47%, or approximately one-half of what was spiked, and indicates that the amount 

detected on-column was in the range of 0.05 ng, which would be nearly equivalent to the mean 

LOD value calculated from the 5-point standard curve. The mean standard deviations between 

replicates averaged 0.093 ng, or 4.7 ppt. 

APPLICATION OF METHODS 

The following tables and discussion show examples of data obtained using the methods 

that have been developed for compound specific analysis of aromatic hydrocarbons of petrogenic 

origin . 

Historical Sediment Data For Pass Fourchon 

Table 7 .6 presents a brief summary of the data accumulated since 1988 for sediments in 

Pass Fourchon. This table shows that, while there is some variablilty over time and depth in 

sediment cores, the overall levels of PAH and saturated hydrocarbons remain high, and FFPI 

values indicate that the source is petroleum contamination. The detailed isomer-specific database 

begins in 1989. The values dated 1994 were obtained for this study. 

Three-Phase Water Data For Coastal Zone Region Near Pass Fourchon 

This data (Table 7.7) is presented as an example of the ability of the analytical methods to 

discriminate specific petroleum contaminant markers at sub-ppb levels, providing additional 

information on transport and availability of these compounds to aquatic organisms. 
84 

Table 7.2 Summary of regression statistics for two standard curves 

5-oolnt curvt 

REGRESSION STATISTICS o.o2-zn 3 neat .sl: 

9-oolnt evrv~ 

REGRESSION STATISTICS 0.005 .10n 2 npilcates 

SLOPE INT COfiFt X-INT Main Std. FF SLOPE INT COFIFL X-INT Main Std . RF 

ANALYTE n Dav, n n Dav. (n 

d8 NAPH 2.13E+06 2.34fi+04 0.999 -0 .011 0.0037 4.65E-07 3.11 E.06 -236E+05 0.999 0.076 0.0027 9.47E-07 

Naphthalene 2.74E+06 1 .85E+04 1 .000 -0 .007 0.0062 7.62E-07 4.01E+06 -3 .03E .05 0.999 0.076 0.012 2.70E-07 

2-MN 2.14E+06 1 .58Ei04 0.999 -0 .007 0.0024 4.54E-07 327E+06 -2.98E+05 0.998 0.091 0.00.11 .77E-07 

1-MN 2.03Ei06 8.10E+03 1 .000 -0 .004 0.0035 4.90E-07 3.04E+06 -2.61E+05 0.958 0.086 0.007 .60E-07 

2-Fl'! 120E+06 4.96E+03 0.999 -0 .004 0.0048 8.28E-07 1.83Er06 -2.42E+05 0.998 0.13 0.0064 628E-07 

1-EN 1.14E+06 1.04Ei04 0.999 -0 .009 0.0091 8.70E-07 1.72E+06 -2 .OOEi05 0.998 0,12 0.0075 6.60E-07 

2,62,7-DMN 4.17E+06 1.11E+02 1 .000 0.000 0.014 4.80E-07 627E+06 -5 .98E+05 0.998 0.19 0.0059 3.53E-07 

1,9J1,7-DMN 5.09E+06 -821E+03 1 .000 0 .003 0 .016 3.94E-07 7S1E+o6 -6 .4bE+a5 0.998 0.17 0 .00.59 2.92E-07 

1,6-DMN 2.40E+06 1 .36E+04 0.999 -0 .006 0.0077 4.15E-07 3.59E+06 -3 .15E+05 0.998 0.088 0.0049 3.05E-07 

1,4/2,3-DMN 3.94Ea06 -8.34E+02 1.000 0.000 0.013 5.08E-07 5.70E+06 -424E+05 0.998 0.15 0.0090 3.79E-07 

1,5-DMN 2.05E+d6 1 .21 E+04 0.999 -0 .006 0.011 4.86E-07 3.06E+06 -2.86E+05 0.998 0.093 0.0064 .60E-07 

Aeorsaphthybrw 3.14E+06 -2 .35Ea04 1 .000 0.007 0.0065 321E-07 4.97Et06 -7 .72E+05 0.997 0.16 0.0033 2.38E-07 

1Z-DMN 2.9E+06 1.64E+03 0.999 -0 .001 0.0089 3.41E-07 4.36fir06 3.89E+05 0.998 0.089 0.0050 2.52E-07 

1,&OAAN 1 .80E+06 5.63E+03 0.999 -0 .003 0.0041 5.54E-07 2.74E+06 1.3E+d5 0.998 0.12 0.0089 4.16E-07 

d10 ACE 1 .73E+06 4.65E+03 1.000 -0 .003 0.0074 5.75E-07 2.63E+06 .46E+05 0.998 0.13 0.0044 4.37E-07 

Aoenaphthone 2.41E+06 9.32E+03 1.000 -0 .004 0.0069 4.14E-07 3.33E+06 -128E+05 0.986 0.038 0.0086 3.1E-07 

1,6,7-TMN 2.62E+06 1 .63Ef04 1,000 -0 .006 .O .oQ62 3.80E-07 4,00E+06 -5 .18E+05 0.997 0.129 0,0067 2.87E-07 

Fluorena 2.79+06 1 .77Ei04 1 .000 '-0.006 0.0067 3.56E-07 4.3E+06 27E+05 0.997 0.14 0.0070 2.70E-07 

Dbanzothicphana 2.47E+06 6.4SE+03 0.999 -0 .003 0.0081 4.04E-07 3.95E+06 -8 .37Ei05 0.996 0.16 0.012 3.02E-07 

d10 PHEN 3.56E+06 6.43E+02 1 .000 0.000 0.005 2.81E-07 5.98E+06 -9.04Er05 0.996 0.15 0.0052 1.97E-07 

PhanantFxona 420E+06 4.04E+03 1 .000 -0 .001 0 .0073 2.38E-07 6 .87E+06 -9.78E+05 0.996 0 .14 0.00.39 1 .70E-07 

Anthracene 4,17E+06 -1 .91 E+04 1 .000 0.005 0.010 2.41E-07 6.95E+06 -1 .12E+06 0.995 0.16 0.0070 1 .71E-07 

4MDBT 1 .43E+08 1 .04Et03 0.949 0.000 0.0044 3.49E-07 2.43E+06 -4.11Et05 0.995 0.085 0.0043 248E-07 

2/3-MOST 2.36E+06 5.41E+03 0.99'9 -0.002 0.0088 3.17E-07 4.OOE+06 -6 .37E+05 0.956 0.12 0.0044 223E-07 

3-MP 1 .32E+06 1 .54E+05 0.931 -0 .117 0.13 6.79E-07 5.45E+06 -8,76E+05 0.995 0.16 0.0049 2.18E-07 

1-MOBT 1 .82E+06 1 .13E+04 1.000 -0 .00d 0.0029 2.73E-07 3.10E+06 -4 .63Et05 0.996 0.075 0.0045 1.89E-07 

2-MP 6.05E+06 -729E+04 0.999 0.018 0.012 2.51E-07 9.78Er06 -1 .54E+06 0.995 024 0.0082 1.82E-07 

4/9-MP 4.87E+06 8.95E+03 0.999 -0 .003 0.012 3.08E-07 7.97E+06 -126E+06 0.995 024 0.0096 224E-07 

1-MP 3 .92E+06 9.72E+d3 0.999 -0 .002 0.0078 2.54-07 6.45E+06 -1 .04E*O6 0.996 0.16 0.0086 1.85E-07 

3,6-OMP 1 .45E+06 -1 .12E+04 0.999 0.004 0.0056 3.47E-07 2.48E+06 5.45Er05 0.994 0.11 0.0051 258E-07 

3,5-OMP 121E+06 -921E+03 0.998 0.004 0.011 4.16E-07 2.02E+06 -4.39E+05 0.994 0.1 1 0.000.1 3.16E-07 

2,6-OMP 243E+08 -1 .36E+04 1.000 0.003 0.0036 2.07E-07 4.18E+06 -920Er05 0.994 0.11 0.0040 159E-07 

2,7-OMP 1 .10Ei08 420E+03 0.999 -0 .004 0.013 9.08E-07 1.87Er06 3.81E+05 0.994 020 0.013 8.71E-07 

9,9-OMP 2.00E+06 9.17E+01 1.000 0.000 0.0046 2.49E-07 3.41E+06 -6 .74E+05 0.994 0.099 0.0044 1 .83E-07 

1,62,Sf2,9-OMP 5.32E+06 -2.44Ei04 1 .000 0.006 0.011 2.96E-07 8.86Ei06 -1 .49E+06 0.995 021 0.0088 1 .70E-07 

1,7-DMP 2.52E+06 -2.83E+03 1 .000 0.001 0.011 4.32E-07 4.07Et06 -8 .89Ey05 0.994 0?2 0.0076 3.14E-07 

1,914,9-DMP 2.55E+06 -1 .SSE+04 1 .000 0.005 0.0082 2.96E-07 5.02Er06 -8 .72E+05 0.997 0.13 0.0067 1.81E-07 

Fiuaanthana 4.94E+06 2.72E+04 0.999 -0 .006 0.013 2.01E-07 1.06E+07 -124E+06 0.997 0.12 0.007 1.07E-07 

1,2-DMDBT 1 .65E+06 1 .65Er04 0.999 -0 .008 0.0084 4.80E-07 3SOE+06 -4.44E+o5 0 .998 0.10 0.0081 2.62E-07 

1,-5-DMP 1 .15E+06 210E+03 1,000 -0 .002 0.0096 8.71E-07 1 .99E+06 -3 .57Ei05 0.995 0.18 0.014 6.13E-07 

1,&DMP 1 .78Et06 5.61E+09 1 .000 -0.002 0.0038 280E-07 321E+06 .8.52E+o5 0.993 0 .10 0.0056 1 .95E-07 

12-OMP 156E+06 3.47Et03 0.999 -0 .001 0.0063 320E-07 2.78E+06 -6.11 E+05 0.993 0.11 0.0084 2.30E-07 

9,10-DMP 1.54Ei06 8.68E+03 0.999 -0 .003 0.0078 22E-07 2.65E+06 5.40E+05 0.994 4.10 0.0041 2.37E-07 

Pyrorw 5.89Et06 -249E+04 0.999 0.004 0.012 1.70E-07 1.09E+07 -1 .61E+06 0.993 0.16 0.0045 1 .15E-07 

1,2,8-TMP 3.77E+06 -128E+04 0.999 0.009 0.013 2.66E-07 7.16Ei06 -1 .42E+06 0.992 020 0.011 1 .74E-07 

Banzanthracena 5.48E+06 -1 .49E+05 0.999 0.027 0.017 1 .88E-07 9.66Es06 -1 .05E+06 0.992 0,11 0.0087 1 .16E-07 

d12CNRYS 4.60E+06 -8 .54E+04 0.999 0.019 0.019 221E-07 8.30E+06 -8 .31 E+05 0.994 0.10 0.012 1 .34E-07 

Cheysone, 5.33E+06 -7 .87E+04 0.999 0.015 0.015 1 .91E-07 9.40E+06 -8 .75E+05 0.993 0.093 0.014 1 .17E-07 

Banzo (b) fiuor 3.35E+06 -2 .39E+05 0.991 0.071 0.059 321E-07 1 .06E+07 -5 .71E+06 0.96b 0.54 0.015 2.06E-07 

8anzo(k),Ouor 4.49E+06 -2-15Ef05 0.998 0.048 0.015 2.14E-07 1 .15E+07 -421E+06 0.975 0.37 0.018 1.38E-07 

Banzo(a)pyrana 2.59E+06 -1 .38E+05 0.994 0.053 0.046 4.07E-07 8.36Ero6 S.37E*06 0.966 0.64 0.021 .35E-07 

412 PEAYL 2.03E+06 -7.42E+04 0.996 0.036 0.042 5.10E-07 6.12E+06 -2.96E+06 0.972 0.47 0.027 298E-07 

Indancpyrarw 8.86Eid5 -1 .37E+05 0.981 0.154 0.058 1.31E-06 3.42E+06 a.08E+06 0.969 0.90 023 2.95E-06 

Dbonzanthracana 6.06E+05 -8.46Et04 0.988 0.140 0.063 1.92E-06 2.61E+06 -2SSE+06 0.961 0.97 0.65 1.48E-05 

Banzoparyiana , 7.18Ei05 -822E+04 0.992 0.087 0.026 153E-06 251E+06 -1 .89E .o6 0.978 0.76 0.029 1.6:1E-08 

MEANS 2.79E+06 -1 .44E+04 0.997 0.009 0.015 4.57E-07 5.07E+06 -1 .06E+06 0.992 0200 0.024 6.14E-07 

MIN 6.06E+a5 -2 .99E+a5 0.931 -0 .117 0.002 1.70E-07 1 .72Et06 -5 .71E .06 0.965 0.038 0.00.1 1 .07E-0T 

MAX 6.05Er06 1 .54Et05 1 .000 0.154 0.132 1 .92E-06 1 .15E+07 -128E+05 0.999 0.974 0.652 1 .48E-05 
85 

Table 7.3 Limit of detection, limit of quantitation, and sensitivity values 

calculated from standard deviations of standard curves 

S 

ANALYTE 

-PT CURVE 

LOO LOG 

(35.D.np) (10'S.O.nq) 

9 

93% C1 

(n0) 

-FT CURVE 

LOO 

(3'S .Onp) 

LOO 

(10'S.D .nq) 

25% C1 

(nq) 

d8 NAPH 0.011 0.037 0.010 0.008 0.027 0.008 

Naphthalene 0.019 0.062 0.017 0.035 0.118 0.033 

2-MN 0.0073 0.024 0.007 0.009 0.031 0.009 

1-MN 0.010 0.035 0.010 0.011 0.037 0.011 

2-EN 0.014 0.048 0.013 0.019 0.064 0.018 

0.027 0.091 0.028 0.022 0.075 0.021 

2,612,7-OMN 0.041 0.14 0.039 0.018 0.058 0.017 

1311,7 .ptiW 0.049 0.18 0.047 0.018 0.059 0.017 

1,8-OMN 0.023 0 .077 0 .022 0 .015 0 .049 0 .014 

1,4/2,-OMN 0.038 0.13 0.038 0.027 0.090 0.028 

0.034 0.11 0.032 0.018 0.054 0.415 

AcanaQhthylene 0.020 0.065 0.018 0.010 0.034 0.010 

0 .027 0 .089 0 .025 0 .015 0 .050 0 .014 

1,8-DMN 0.012 0.041 0.072 0.027 0.089 0.025 

d10 ACE 0.022 0.074 0.021 

. 

0.013 0.044 0.013 

Acenephinene 0.021 0.009 0.020 0.028 . 0.086 0.024 

1,8,7.TMP1 0.019 0.082 0.017 0.020 0.067 0.019 

Ruaene 0.024 0.067 0.019 0.021 0.070 0.020 

DibenZOthbphene 0.024 0.081 0.023 0.0.38 0.121 0.034 

d10 PHEN 0 .011 0 .035 0 .010 0.018 0.052 0 .015 

Phenenthrene 0.022 0.073 0.021 0.012 0.039 0.011 

Anthracene 0.030 0.10 0.028 0.021 0.070 0.020 

4-MOST 0.013 0 .044 0 .013 0 .013 0 .043 0 .012 

?13-M08T 0.028 0.088 0.025 0.013 0.044 0.013 

3-MP 0.40 1 .3 0.373 0.015 0.049 0.014 

1-MOST 0.0086 0.03 0.008 0.013 0.045 0.013 

2-MP 0.035 0.12 0.033 0.025 0.082 0.023 

p 0.037 0.12 0.035 0.029 0.096 0.027 

p 0.023 0.078 0.022 0.028 0.086 0.024 

3,8-OMP 0.017 0.058 0.018 0.015 0.051 0.014 

3,5-DMP 0.033 0.11 0.032 0.013 0.043 0.012 

2,6-DMP 0.011 0.038 0.010 0.012 0.040 0.011 

2,7-OMP 0.039 0.13 0.037 0.038 0.127 0.038 

3,9-DMP 0 .014 0 .046 0 .013 0 .01 0 .044 0.013 

1,BI2,52,9-DMP 0.033 4.11 0.031 0.028 0.088 0.025 

1,7-OMP 0.032 0.11 0.031 0.023 0.078 0.021 

1,9/4,9-OMP 0.025 0.082 0.023 0.020 0.067 0.019 

Fluoranthene 0.038 0.13 0.038 0.022 0.079 0.021 

1 Z-OMDBT 0 .025 0.084 0.024 0 .024 0.081 0 .023 

1,5-OMP 0.029 0.098 0.027 0.042 0,141 0.040 

1,8-OMP 0.011 0.038 0.011 0.017 0.058 0.016 

1,2-DMP 0.019 0 .063 0 .018 0 .025 0.084 0.024 

9,10-DMP 0.023 0.079 0.022 0.012 0.041 0.012 

ppr9ne 0,037 0.12 0.035 0.013 0.045 0.013 

1,2,8-TINP 0.039 0.13 0.037 0.034 0.112 0.032 

BanZantfiracene 0.05 0.17 0.049 0.026 0.087 0.025 

d12 CNFiYS 0.06 0.19 0.054 0.036 0.120 0.034 

Cnrysene 0.05 0 .15 0 .044 0.041 0 .18 0 .08 

0.18 0.59 0.188 0.045 0.750 0.042 

Benzo(k),Auor 0.045 0.15 0.042 0.05 0,178 0.050 

Benzo(a)pyrene 0.14 0 .46 0 .131 0 .064 0212 0 .060 

d12 PEAYL 0.13 0.42 0.118 0.080 0.257 0.078 

Indenopyrena 0.17 0.58 0.164 0.692 2307 0.652 

Dibenzanthracene 0.19 0.63 0.179 1958 A.519 1 .843 

Benzoperyiene 0.078 028 0.074 0.087 0.290 0.082 

MEANS 0.046 0.155 0.044 0.072 0.241 0.068 

0 .007 0 .024 0 .007 0 .008 0 .427 0.008 

MqX 0 .398 1 .320 0 .373 1 .958 9.519 1 .841 
86 

Table 7.4 Method detection limits for three sample matrices 

LOD MATRIX 

S.D .ng) TISSUES SEDIMENTS WATERS 

AVE SAMPLE AMT. 20.459 7.429 " ' 2000m1 

Naphthalene 0.019 0.091 0.25 0.00093 

2-MN 0.0073 0.05 0.098 0.00036 

1-MN 0.010 0.051 0.14 0.00052 

2-EN 0.014 0.070 0.19 0.00071 

1-EN 0.027 0.13 0.37 0.0014 

2,62,7-DMN 0.041 0.20 0.56 0.0021 

1,3/1,7-DMN 0,049 0.24 0.67 0.0025 

1,6-DMN 0.023 0.11 0.31 0.0012 

1,42,3-DMN 0.038 0.18 0.51 0.0019 

1,5-DMN 0.034 0.17 0.46 0.0017 

Acenaphthyfene 0.020 0.095 0.26 0.00098 

1,2-DMN 0.027 0.13 0.36 0.0013 

1,8-DMN 0.012 0.060 0.17 0.00061 

Acenaphdiana 0.021 0.10 0.28 0 .0010 

1,6,7-TINN 0.019 0.091 0.25 0.00093 

Fluotene 0.020 0.098 027 0.0010 

Dibenzottuaphene 0.024 0.12 0.33 0.0012 

Phenanthrens 0.022 0.11 0.29 0.0011 

Anthracene 0.030 0.15 0.41 0.0015 

4-MDBT 0.013 0.065 0.18 0.00066 

2/3-M DBT 0.026 0.13 0.36 0.0013 

3-MP 0.40 1 .9 5.3 0.0198 

1-MOST 0.0086 0.042 0.12 0.00043 

2-MP 0.035 0.17 0.48 0.0018 

4J9-MP 0.037 0.18 0 .50 0.0018 

1-MP 0.023 0.11 0.31 0 .0012 

3,r,-DMP 0.017 0.082 0.23 0.00084 

3,5-OMP 0.033 0.16 0.45 0 .0017 

2,6-DMP 0.011 0.053 0.15 0.00054 

2,7-DMP 0.039 0.19 0.52 0.0019 

3,9-DMP 0.014 0.067 0.18 0.00068 

1,6l2,5r2,9-OMP 0.033 0.16 0.45 0.0017 

1 .7-DM P 0.032 0.16 0.44 0.0016 

1,9/4,9-OMP 0.025 . 0.12 0.33 0.0012 

Fiuorantfiene 0.038 0.19 0.51 0.0019 

1,2-DMDB'T 0.025 0.12 0.34 0.0013 

1,5-DMP 0.029 0.14 0.39 0.0014 

i,&OMP 0.011 0.055 0.15 0.00057 

1,2-DMP 0.019 0.092 0.25 0.00095 

9,10-DMP 0.023 0.11 0.32 0.0012 

Pyrene 0.037 0.18 0.50 0 .0019 

1,2,8-TEA P 0.034 0.19 0.53 0.0020 

Benzanthracene 0.052 0.25 0.70 0 .0026 

Chrysene 0.046 023 0.62 0.0023 

Benzo (b) fluor 0.18 0.87 2.4 0.0089 

Benzo(k),fluor 0.045 022 0.60 0.0022 

Benzo(a)pyrene 0.14 0.68 1 .9 0.0070 

Indenopyrene 0.17 0.85 23 0 .0087 

Dibenzanduaeene 0.19 0.93 2.6 0.0095 

Benzoperylene 0.078 0.38 1 .1 0.0039 

MEANS 0.050 0.24 0.67 0.0025 

MIN 0.0072 0.035 0.097 0.00036 

MAX 0 .40 1 .9 5 .3 0.020 

'MDL (ppb)_lOD'200ui final vol . / (sample amt' 2ul inj d) 

"'typical dry wt of 15q wet se6ment 
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88 

Table 7 .6 Historical data for sediments from the Pass Fourchon 400m (from discharge) 

sampling site 

DATE DEPTH TOTAL PAH SATURATED HC FFPI Reference 

(CM) ppb ppb 

Oct-87 0-5 7,700 260,000 0 .91 1 

5-10 4,200 184,000 0 .83 

10-15 6,800 210,000 0 .89 

15-20 1,000 250,000 0 .76 

Jan-88 0-5 43,000 650,000 0 .93 1 

Feb-89 0-2 21,000 490,000 0 .92 2 

2-5 22,000 570,000 0 .91 

5-10 20,000 490,000 0 .92 

10-15 21,000 630,000 0 .91 

15-20 7,900 140,000 0 .92 

20-27 ' '31,000 580,000 0 .93 

May-89 0-2 35,000 680,000 0 .90 2 

Oct-89 0-2 22,000 760,000 0 .91 2 

Feb-90 0-2 91,000 1,700,000 0 .94 2 

2-5 36,000 930,000 0 .93 

5-10 61,000 1,000,000 . 0 .95 

10-15 20,000 750,000 0 .88 

15-20 14,000 660,000 0 .87 

20-25 12,000 510,000 0 .87 

25-30 17,000 471,000 0 .93 

30-35 12,000 310,000 0 .80 

35-40 22,000 490,000 0 .96 

1994 53,000 NA 0 .98 3 

1 Boesch & Rabalais, 1989 

2 Rabalais et al, 1991 . Vol . III 

3 this report 
89 

Table 7.7 Application of analytical methods to aqueous and particulate phases 

of water and sediments in coastal waters near Pass Fourchon 

ANuM assavm afasE 

SUFFpCE 

ml 

BOTTOM 

ml 

PARTICULATE awse 

SIFFOCFE: BOTTDM 

n n 

sEDRAExr 

(BEDOED) 

n 

Naphthalene 134 96 2673 2420 2.2 

2-MN 223 141 132 96 2.5 

1-MN 125 e0 S3 40 2.2 

2-EN 26 12 27 14 1 .0 

1-EN 5.1 TA TA 7R 9 .2 

2,D-/2,7-OMN 62 92 65 36 nd 

1,9/1,7-OMN 43 26 38 27 2.2 

1,8-DMN 28 17 3.7 22 1 .8 

1,4/2,9-OMN 79 9.3 24 12 2.3 

1,SOMN 32 18 9.3 7.6 nd 

Acanaphtfiylene TR TR nd TR 0.63 

1,2-OMN 7.4 4.4 9.0 TA 0.39 

2-IPN 13 8.9 nd nd NA 

1,8-DMN nd nd nd nd NA 

ACenaphthens 19 15 13 6.2 0.51 

1,8,7-TMN ~9.2 nd 24 19 t .0 

Fiuorene 13 11 12 72 nd 

Dibanzothiophena 6.3 7.6 nd 7.6 0.71 

Pfienenduene 17 13 11 38 5 .6 

Anduacene nd TFl nd 0.28 0.92 

4-M08T TR TR nd 15 1 .3 

2/3-MOST nd nd nd 6.0 nd 

3-MP 1,5 nd nd 13 3 .0 

1-MOST nd nd nd 0.12 NA 

2-MP 1.0 nd nd 17 1 .8 

4/9-MP 27 3.0 nd nd 1 .9 

1-MP nd nd nd 3.4 0.98 

3,8-DMP nd nd nd 6.3 1 .1 

9,SDMP nd nd nd nd 

2,8-OMP nd nd nd 7.3 0.77 

2,7-OMP nd nd nd 20 0.83 

9A-OMP nd nd nd 2.9 2.8 

1,8/2,5/2,9-OMP nd nd nd 21 1 .7 

1,7-OMP nd nd nd 1 .9 2.1 

1,9/4,9-OMP nd nd nd nd 1 .8 

1,2-DMDBT nd nd nd nd nd 

Fluoranthens 1.4 TA 55 128 4.9 

1,SOMP nd nd nd nd nd 

1,8-OMP nd nd nd 2,4 0 .60 

.1,2-DMP nd nd nd nd 0 .78 

9,10DMP nd nd nd nd nd 

Pytene 1 .0 t .t 57 115 8.6 

1,2,&TMP nd nd nd nd nd 

Benzenthracene 4.3 nd 47 T7 3.4 

Chry3ene 23 nd 62 118 4 .3 

Bsnzo(b)fluor nd nd 333 1297 9 .9 

Ben=o(k)fluor nd nd nd nd nd 

Benzo(a)pyrene nd nd nd 381 3 .7 

Indenopyrene nd nd TR 418 4.7 

Dibenzanthracene nd nd nd TA TA 

Benzoperylene nd nd 77 113 4.9 

Estimated Totals 

Cl-Naphifialonas 347 221 185 136 4.7 

C2-Naphthalanas 222 119 206 120 11 

C1-Naphlhaknes 96 65 124 1?3 NA 

CA-Naphthalorws nd 5.9 Za 97 NA 

CI-Dibonzothiophonas TR TF1 ^d 21 1 .30 

a-Dibaruolhiaphonos 8.5 8.6 nd 64 NA 

Ct-Plwnanthrones 5.1 9.0 nd 94 7.7 

C2-Phonanthranes nd nd nd 43 17 

G1-Plwnanihtvrws 6.7 nd nd 17 NA 
91 

CHA TER 8 

INVESTIGATION OF THE BIOACCUMULATION 

POTENTIAL OF 60 INDIVIDUAL 

AROMATIC PETROLEUM-ASSOCIATED 

HYDROCARBONS IN THE 

OYSTER CRASSOSTREA VIRGINICA 

J. C. Means, K.L. Sarradet, L.A. Reily and D.J. McMillin 

INTRODUCTION 

Owing to their chemical structures and inherent physiochemical properties, a number of 

anthropogenically generated organic chemicals are bioaccumulated by marine organisms via 

various routes of uptake . Crassostrea virginica are bottom dwelling-filter feeders and tend to 

burrow slightly into the sediment along the benthic zone. Petroleum hydrocarbons, which are 

considered toxicologically relevant, are being released into the water of coastal Louisiana and the 

Gulf of Mexico due to the ongoing extraction of petroleum. Once released, the chemicals are 

sorbed onto the sediment and as our study shows, become bioavailable to benthic bottom 

feeders. Concentrations of contaminants will vary from site to site but where uptake does not 

cause a physiologically compromised state or death, introduction to aquatic food webs is 

probable. 

MATERIALS AND METHODS 

Microcosm Exposures 

Live oysters (Crassostrea virginica), with an average shell length of 5 cm, from Black 

Bay, LA were purchased from a local seafood store (Baton Rouge, LA.) The oysters were then 

brought to our aquatic laboratory where they were placed in a large holding tank for acclimation. 

In the holding tank, oysters were submersed in sea water (pre-dissolved FRITZ Super Salt - a 

synthetic sea water salt mixture plus carbon filtered/ dechlorinated water 11 ppt salinity ; 21-

23°C) for a period of 4.5 days. Oysters were acclimated to a 12-hr. day and night. Spawning 

began on day 1 and ceased around day 4. During the daylight hours of this period, 1 day old 

prepared sea water was constantly flushed through the holding tank to discourage spawning and 

to keep ammonia levels to a minimum. Holding tank was aerated with 2 aquaria air pumps of 

appropriate pumping capacity with air stones attached during the duration of the acclimatization . 

A liquid diet for invertebrate marine animals by Hawaiian Marine Imports, Inc. was chosen as 

the diet and their first feeding was on day 3 in the holding tank. 

Twenty, 10-gallon aquarium tanks (glass sides and bottoms) were set up as follows. 

First, sediment obtained from the contaminated area as well as sediment from the control area 

was hand pressed through a 2 mm sieve to remove stones, shell hash and debris . Four separate 

dilutions were then prepared using a mixture of the contaminated and control sediments in 

varying amounts to yield a 0% contaminated batch, comprised solely of sediment from Lake 

Champagne; a 12.5% contaminated batch; 25°Io; and 50% with the latter percentages indicating 

of contaminated sediment from Pass Fourchon accordingly. 
92 

Oysters were exposed to dilutions of sediment contaminated with 52 ppm total petroleum 

aromatic hydrocarbons to study the rates and magnitude of bioaccumulation. A period of 28 

days was chosen for the exposure study based upon previous experiments which indicate that 

the organism can achieve steady-state within that period . (McElroy and Means, 1988) . Four 

concentrations of contaminated sediment from Pass Fourchon (52 ppm total petroleum aromatic 

hydrocarbons) were prepared by mixing with "clean" sediment from Lake Champagne (< 0.015 

ppm total petroleum aromatic hydrocarbons) and rolling in jars for 24 hr. to ensure 

homogeneity. The resulting mixtures, -3000 kg total (wet wt.), were comprised of 0°Io, 12.5%, 

25%, and 50% of contaminated sediment, respectively . Oysters (4-5 per chamber) were held in 

separate aquaria and samples were taken on days 0, 3, 7, 14, 21, and 28 to assess uptake of the 

aromatic hydrocarbons from the sediments by the oysters. Overlying water in the tanks was 

exchanged every two days to prevent the accumulation of metabolic wastes . Feeding occurred 

immediately after each water exchange. Mortality was noted every two days and recorded. 

Oysters were recovered from the tank, rinsed and frozen at -40°C until analyses were 

performed. Tissues were prepared for hydrocarbon analysis by thawing, followed by removal 

of the soft tissues from the shell and rinsing with distilled water to remove any residual 

sediment. 

Chemical Analysis 

Tissue Sample Preparation 

Oysters samples were defrosted overnight in a refrigerator . Individual samples were 

removed from the bags and rinsed with tap water, then shelled on a glass cutting board with a 

TeflonO coated knife. The shell-free flesh was rinsed with organic free water and then chopped 

into small pieces . The pieces were placed in a clean jar and homogenized using a Tekmar 

Tissuemizer@. The homogenized samples were kept frozen at -20°C until sample extraction . 

The extraction method is a modification of that described by MacLeod et al. (1985) . 

Approximately 20 g of defrosted homogenized tissue was placed in an 240-m1 amber glass bottle 

with 100 ml nanograde@ dichloromethane (DCM) and four times the tissue weight of cleaned 

sodium sulfate (DCM rinsed) . A surrogate spike of 200 ng each of a mixture of deuterated 

standards (US-108, Ultra Scientific) was added to the bottle . The sample was extracted with a 

Tekmar TissuemizerO for 2 min. at a speed of 24,000 rpm. The probe was rinsed with DCM 

and the rinsings added to the sample bottle . The extract was allowed to settle for 1 min. and then 

filtered through a funnel filled with sodium sulfate into a labeled florence flask. The sample was 

extracted as above with two additional 100 ml volumes of DCM. After the third extraction, the 

extract was rinsed with an additional 30 ml of DCM, which was added to the flask. The sample 

extract was reduced in volume to approximately 2 ml with a rotary evaporator. The concentrated 

extract was quantitatively transferred to a 4-ml vial and concentrated to a volume of 2 ml under a 

stream of purified nitrogen. Reagent blanks, duplicates and matrix-spiked samples were 

extracted to monitor laboratory technique and extraction efficiency . Matrix spikes were duplicate 

tissue samples to which 100 ng of the quantitative calibration standard was added instead of the 

deuterated surrogates mixture. 

Most tissue samples contained a layer of fine white precipitate (extracted proteins) which 

was removed before silica column cleanup. The extracts were "cleaned" by adding 500 ~l of 

hexane, mixing the extract well and then centrifuging for 5 min. at 3,000 rpm. The solvent layer 

was transferred to a clean 4-ml vial and the extract was "cleaned" twice more with 1 ml of 

hexane. All solvent layers were combined and reduced in volume to 1 ml under a stream of 

purified nitrogen . 
93 

One half of the extract was separated from the polar lipids on 200 mm x 10 mm columns 

packed with 6.0 g of 100-200 mesh Grade 923 silica, eluted with 100 ml of 60% DCM and 40°Io 

hexane. The extract was concentrated under nitrogen to 200 ~t1, after which 2 g1 of 2-

fluorobiphenyl (100 ng/g1 in hexane) was added as an internal standard to monitor the GC/MS 

injection performance. Instrumental analysis (GC/MS) was performed as described in Chapter 

7 . 

Sediment Sample Preparation 

Sediment samples from the microcosms were sealed in glass jars with a Teflon-lined 

cap, mixed, and refrigerated at -20°C until processed for analyses. The extraction method was a 

modification of that described by MacLeod et al . (1985) . Excess water was decanted from 

sample jars prior to thorough mixing of the sediment sample, and 15 g was weighed into an 

amber bottle with a Teflon-lined cap. The sediment was covered with 75 ml of dichloromethane 

(DCM) and 50 g sodium sulfate incorporated until the sediment mixture was free-flowing . A 

suite of deuterated PAH was added (800 ng in hexane of six deuterated PAH) with the first 

solvent aliquot as a surrogate standard to monitor extraction efficiency . The jars were sealed and 

placed on a roller for 16, 6, and 16 hours, for a total of three extractions using the modified rock 

tumbler and rolling times specified by MacLeod et al. (1985) . The DCM from each jar was 

decanted into an amber jar and refrigerated until 3 extracts were collected. The decanted DCM 

was filtered through sodium sulfate, and concentrated by rotoevaporation and a nitrogen stream 

to a final volume of 200 g1, with solvent exchange to hexane. Activated fine granular copper 

was added to reduce sulfur interferences. Instrumental analysis (GC/MS) was performed as 

described in Chapter 7. 

RESULTS AND DISCUSSION 

Mortality 

Significant mortality was observed in the two highest dose rates. Table 8 .1 shows the 

mortality data for all exposures. After two separate experimental attempts at sustaining viability 

in the 25% and 50°Io exposure chambers for longer than 14 days, we were unable to maintain 

enough live organisms in order to assess the rate of uptake and bioaccumularion of the petroleum 

hydrocarbons for those periods due to the mortality. Survivorship in the control sediments and 

12.5% contaminated dose tanks was satisfactory to obtain bioaccumulation data. Our conclusion 

in this case is that the concentrations of petroleum HC's were too great to maintain viability at the 

high dose levels for more than 14 days. Maintenance of the laboratory experimental conditions 

proved adequate to sustain life in all other chambers. 
94 

Table 8.1 Mortality data 

°Io of oysters surviving at collection date 

Dose 

0% 12 .50% 25% 50% 

DAY 0 100% 100% 100% 100% 

DAY 3 100% 75% 75% 100% 

DAY 7 100% 75% 25% 25% 

DAY 14 75% 50% 25% 25% 

DAY 21 75% 25% 0% 0% 

DAY 28 25% 50% 0% 0% 

Sediment Contaminant Chemistry 

Both of the sediments used in these experiments are typical of fine-grained sediments 

found in coastal regions of Louisiana. The contaminated sediment from Pass Fourchon is a 

sandy silt, -70°Io sand, 25°Io silt and < 5% clay. The Lake Champagne sediment is also a sandy 

silt, 38 .5% sand, 56.6°Io silt and <5°Io clay . The total organic carbon content of the two 

sediments were 2.0% for the Pass Fourchon and 4.0% for the Lake Champagne sediments, 

respectively. 

Total aromatic hydrocarbon content was measured in both sediments. The undiluted 

Pass Fourchon sediment contained 52.8 ppm of total aromatics of which only 0.5 ppm were 

parent aromatic compounds. The vast majority of the aromatic fraction was composed of 

alkylated analogs of parent, and heterocyclic aromatics typically associate with petroleum 

sources, including produced water discharges (Means and McMillin, 1993). In contrast, the 

Lake Champagne reference sediment contained only 0.015 total aromatic hydrocarbons of which 

<0.001 ppm were petroleum-associated materials. 

Table 8.2 shows the concentrations of each individual compound determined in the 

sediments and reported on a dry-weight basis. The values are consistent with data obtained at 

this site in previous investigations (Rabalais et al ., 1991a; Means and McMillin, 1993). 

Bioaccumulation 

The filter-feeding benthic organism [oyster (Crassostrea virginica)] was exposed to four 

dilutions of a contaminated estuarine sediment [52 ppm total petroleum aromatic hydrocarbons] . 

An assessment of the amount of bioaccumulation of 60 individual aromatic hydrocarbons, 

measured by gas chromatography/mass spectrometry (GC/MS) revealed that accumulation was 

both time and dose dependent. A summary of the concentrations of each individual compound 

detected in each set of oysters at each time and dose level are presented in Appendix A. Selected 

data, showing major trends, are presented here. 

Figure 29a shows the accumulation of high molecular weight parent PAH from the 0°Io 

contaminated dose sediment (pure reference sediment) . Chrysene and benzo(b and k) 

fluoranthene were not detected in any samples. Fluoranthene and benzanthracene were found at 

low levels in the tissues of acclimated but unexposed organisms. The levels of these two 

compounds appeared to decrease through day 14 of the experiment but were elevated after 21 

days of exposure. Pyrene, absent at the early time points, was observed at day 21 in the tissues 

at low levels . 
95 

Table 8.2 Sediment data for Pass Fourchon and reference site 

SAMPLE LCHAMP PF400 

Units, dry wt, basis ppb' ppb 

Naphthalene nd 1 .7 

2-Methylnaphthalene nd 3 .0 

1-Methylnaphthalene nd 1 .5 

2-Ethylnaphthalene nd 1 .9 

1-Ethylnaphthalene nd 9 .1 

2,6/2,7-Dimethylnaphthalene nd 10 

1,3!1,7-Dimethylnaphthalene nd 48 

1,6-Dimethylnaphthalene nd 4.7 

1,412,3-Dimethylnaphthalene nd 27 

1,5-Dimethylnaphthalene nd 25 

1,2-Dimethylnaphthalene nd 12 

2-Isopropylnaphthalene nd nd 

1,8-Dimethyl naphthalene nd 12 

1,6,7-Trimethylnaphthalene nd 197 

Acenaphthylene nd 4.0 

Acenaphthene nd 11 

Ruorene nd 18 

Dibenzothiophene nd 5 .6 

Phenanthrene 1 .0 52 

Anthracene nd 16 

4-Methyldibenzothiophene nd 68 

2/3-Methyl dibenzothiophene nd 55 

1-Methyldibenzothiophene nd 34 

3-Methylphenanthrene nd 183 

2-Methylphenanthrene 1 .0 87 

4/9-Methylphenanthrene nd 177 

1-Methylphenanthrene nd 102 

3,6-Dimethylphenanthrene nd 102 

3,5-Dimethylphenanthrene nd nd 

2,6-Dimethylphenanthrene nd 93 

2,7-Dimethylphenanthrene nd 77 

3,9-Dimethylphenanthrene nd 335 

1,6/2,5/2,9-Dimethylphenanthrene nd 228 

1,7-Dimethylphenanthrene nd 83 

1,9/4,9-Dimethylphenanthrene nd 78 

1,5-Dimethylphenanthrene nd nd 

1,8-Dimethylphenanthrene nd 32 

1,2-Dimethylphenanthrene nd 22 

9,10-Dimethylphenanthrene nd nd 

Fiuoranthene 4 .5 73 

1,2-Dimethyldibenzothiophene nd 5 .9 

Pyrene 2.7 54 

1,2,8-Trimethylphenanthrene nd 84 

Benzanthracene 3.0 24 

Chrysene 1 .9 8 4 

Benzo(b)tluoranthene nd 29 

Benzo(k)fluoranthene nd 8.4 

Benzo(a)pyrene nd 12 

Indenopyrene nd 30 

Dibenz(a,h)anthracene nd 17 

Benzo(g,h,i)perylene nd 38 

C3-NAPHTHALENES nd 9,700 

C4-NAPHTHALENES NA 8,900 

C2 DIBEIVZOTHIOPHENES nd 8,600 

q-PHEIVArfTHRENES nd 23,000 

Total Parent PAH 12 478 

Total Alkylated PAH 0.50 52,397 

Total PAH 13 52,875 

FFPI (missing C3-DBTs) 0 .06 0 .98 

'mean of lab duplicates 
96 

Figure 29b shows the same group of parent PAH accumulating from the 25% 

contaminated sediment. In this exposure, each of the compounds detected was observed to 

increase with time through day 14, except for benzanthracene which decreased and then 

increased at day 14. Chrysene was never observed in the 0% dose, but was found at significant 

levels in tissues exposed to 25% contaminated sediment. 

Figure 29c shows the accumulation of high molecular weight PAH in oysters exposed at 

the 50°Io dose. Each of the compounds, including the benzofluoranthene isomers, were 

observed in these tissues after 7 to 14 days. The levels of PAH accumulated were much higher 

than any of the lower exposure doses and each of the compounds was observed to increase with 

time except benzanthracene, which decreased and then increased as reported at the other dose 

levels . At the 50°lo dose level, tissue concentrations at 7 days were approximately 5 times higher 

than seen at the lower doses at that time point. It is interesting to note that between day 7 and 

day 14 of this experiment, the concentrations of PAH declined in the oyster tissues. We 

interpret this as being a reflection of the physiologically compromised nature of the organisms, 

since mortality was so high at this exposure level. 

Figure 30(a-d) shows the bioaccumulation pattern of low molecular weight PAH in 

oysters at four sediment doses. In the 0% dose system, a pattern similar to that observed with 

the high molecular weight PAH of decreasing followed by increasing PAH concentrations . At 

the 12.5°Io dose, a general pattern of declining PAH concentrations with time was observed. At 

the 25 and 50°Io dose levels, these compounds appear to increase in tissues with time but there is 

no clear dose dependent pattern of increasing tissue levels . 

The bioaccumulation behavior of the more abundant alkylated aromatic hydrocarbons 

present in the sediments give a more consistent pattern tissue distributions with both time and 

sediment dose. Figure 31(a-d) show the bioaccumulation of C3-phenanthrenes in oyster tissues 

as a function of time and dose. In the 0% dose, where sediment C3-phenanthrenes were not 

detected, these compounds were not detected in the day 0 oysters, but were detected at very low 

but consistent levels (-12 ppb) in tissues samples after day three. At the 12 .5% and 25°Io dose 

levels, there was a rapid, dose-dependent increase in C3-phenanthrenes in the oyster tissues to 

14 days. Only oysters at the 12.5°Io dosage level survived to 28 days, and these showed 

declining PAH levels, again probably due to compromised physiology. At the 50°Io dose level, 

tissue concentrations increased even more rapidly to high levels . After only 7 days, the tissue 

concentrations were comparable to the levels observed in the 25% dose level after 14 days. The 

highest concentration reached for this compound group was 670 ppb, wet weight, which is 

equivalent to 33,000 ppb on an average lipid-weight basis. 

Figure 32(a-d), shows the bioaccumulation pattern of C2-phenanthrenes, another group 

of alkylated PAH which were absent from the reference sediment but very concentrated in the 

contaminated sediment. A dose-dependent increase and time-dependent increase in tisssue 

residues of each of the individual dimethyl phenanthrenes was observed in the oyster tissues. 

As with several of the contaminant groups at the high level dose, tissues residues declined 

between day 7 and day 14, and after 14 days in the 12.5% dosage level. This was attributed to 

the toxicity of the sediments to the oysters with time. 

Another compound group, which is unique to petroleum-related contaminated sediments 

are the alkyl dibenzothiophenes. This group of heterocyclic aromatic hydrocarbons were absent 

from the reference sediment and relatively abundent in the contaminated sediment. Figure 33(ad) show the bioaccumulation of the C2-dibenzothiophenes in oyster tissues. Total dimethyl 

dibenzothiophenes showed a dose and time dependent increase in tissue residues, similar to the 

trend seen for C2- and C3-phenanthrenes . 
97 

60 0% PF400: HIGH MW PAH 

0 Fluoranthene 

..: 50 ~ pyrene 

3 40 ~ Benzanthracene 

30 ~ Chrysene 

~ Benzo(b)fluoranthene 

a 20 ~ Benzo(k)fluoranthene 

10 

0 

0 3 7 14 

Days of Exposure 

VO 

50 

3 40 

N 

3 30 

a 

a 20 

10 

0 

60 

50 

r 3 40 

as 

3 30 

.a 

a 20 

10 

0 

21 

a. 

Figure 29. Concentrations (ppb) of high molecular weight PAH bioaccumulated at 3 dose levels . 

0 3 7 14 

Days of exposure 

0 3 7 14 

Days of Expos ure 
20 0% PF400 : LOW MW PAH a_ zo 12 .5% PF400 : LOW MW PAH 

D Acenaphthylene o Dibenzothlophene 

~s i5 o Acenaphthene D Phenanthrene 

D Fluorene e Anthracene 

r+ 

~0 

++ 

010 

a a 

a s 

o " o 

0 3 7 11 21 0 3 7 14 21 2B 

Days of Exposure Days of Exposure 

20 25% PF400 : LOW MW PAH C . zo 50% PF400: LOW MW PAH 

d. 

15 15 

5 

r 

10 

r 

310 

.a 

a a 

a a 

b 5 

0 ~ 0 

0 3 7 11 0 S 7 14 

Days of Exposure Days of Exposure 

03 

Figure 30. Concentrations (ppb) of low molecular PAH bioaccumulated at 4 dose levels . 
ISO 0% PF400: C3-PHENANTHRENES 7SO 12.5% PF400: C3-PHENANTHRENES 

a. b . D "1,2,8-Trimethylphenanthrene" 600 

a C3-Phenanthrenes 

goo 

rs° 

3 3 

a ~a3°° 

CL so (SCALEzSX) a 

ISO 

0 

0 3 7 14 21 

0 

0 9 7 14 21 28 

Days of Exposure Days of Exposure 

750 25% PF400: C3-PHENANTHRENES C . 750 50°/. PF400 : C3-PHENANTHRENES 

d . 

600 

450 450 

m 

3 

m 

; 

300 -0300 CL a Q, 

ISO ISO 

o 

0 9 7 11 

a 

0 y 7 14 

Days of Exposure Days of Exposure 

Figure 31 . Concentrations (ppb) of C3-phenanthrenes bioaccumulated at 4 dose levels. 
100 

Other compound groups were also examined and followed trends observed with the 

preceeding groups discussed. They include: methyl dibenzothiophenes, C-2 naphthalenes, C3-

naphthalenes and C4-naphthalenes. These data are contained in Appendix A. 

CONCLUSIONS 

The sediment-bound contaminants associated with chronic produced water discharges 

which accumulate over time were found to be bioavailable to the filter-feeding bivalve 

Crassostrea virginica. Accumulation could be observed in as little as three days of exposure. 

Petroleum- associated aromatic hydrocarbons increased in a dose- and time-dependent fashion. 

Mortalities observed in the higher dose groups appeared to be contaminant related and effects on 

accumulation due to the compromised physiological state of intoxicated oysters was observed 

after 14 days in the highest dose group, and after 21 days at the 12.5°Io dosage level. 

Isomer-specific analyses of bioaccumulated petroleum aromatic hydrocarbons in tissues 

of aquatic organisms yield information on the relative bioavailability of this important group of 

contaminants. These chemicals have been shown to rapidly enter biological foodwebs in aquatic 

systems as the result of exposure to contaminated sediments even in the absence of a continuous 

source of fresh contaminants. This suggests that these contaminated sediments will remain a 

source of bioavailable toxic substances even after discharges are suspended at a site . Since some 

of these foodwebs potentially lead directly to man, there are both ecological and human health 

risks associated with in-place, sediment-bound contamination from produced waters . 
,z0 0°/. PF400: C2-PHENANTHRENES 

a. 

p 9,s-Dlmethylphenanthrene 

pp p 2,6-Dlmoth ylphananthnne 

2,7-Dimefhylphsnanthrene 

so 0 3,9-Dlmothylphenanthrene 

1,8/2,b/2,9-Dimethylphe nanthren 

1,7-Dlmethylphenanthrans 

p 1,9/4,9-Dimsihylphenanthrene 

d D 1,8-Dimathylphensnthnne 

a ,o 

201 E3 1,2-Dimsthylphenanthrem 

p 9,70-Dlmethylphenanthrane 

0 3 7 14 Yl 

Days of Exposure 

rr 

m 

3 

.a 

a 

a 

m 

3 

a 

a 

m 

3 

a 

a 

Figure 32. Concentrations (ppb) of C2-phenanthrenes bioaccumulated at 4 dose levels . 

F~ 

0 

r 

Days of Exposure 

Days of Exposure Days of Exposure 
no 0°ti PF400: C2-DIBENZOTHIOPHENES 270 12.5"/. PF400: C2-DIBENZOTHIOPHENES b. 

240 240 

210 13 1,2-Dimethyidibenzothlop hone 210 

0 C2-Dibenzothiophenes Leo ISO 

ISO ISO 

m 

120 120 

a 90 

a °90 

so `° 

30 3° 

o 

o s > >4 s~ 0 3 ~ u 21 xe 

Days of Exposure Days of Exposure 

:70 2594 PF400: C2-DIBENZOTHIOPHENES =70 50% PF400: C2-DIBENZOTHIOPHENES 

c. d. 

240 240 

210 210 

180 180 

160 ,., 1b0 

3 3 

-1so -1so 

a CL 90 

I 

a 

a90 

w so 

30 90 

0 

0 S 7 14 

0 

0 3 7 

IL 

11 

Days of Exposure Days of Exposure 

O 

N 

Figure 33 . Concentrations (ppb) of C2-dibenzothiophenes bioaccumulated at 4 dose levels . 
103 

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Appendix A: Parent and alkylated PAH concentrations (ng/g) in oyster tissue from organisms exposed to varying 

levels of contaminated Pass Fourchon sediment for 28 days . 

EXPOSURE LEVEL 

SAMPLING DAY 

ANALYTE 

0% LEVEL (CONTROL) 

0 0-dup. 3 3-dup. 7 

n n n n n 

14 

ng/g 

21 

n 

12.5% LEVEL 

3 7 14 

ng/g n n 

21 

ng/ 

28 

n 

25% LEVEL 

3 7 14 

n n ng/ 

50% LEVEL 

3 7 14 

n n n 

Naphthalene nd nd 16 20 23 13 261 69 16 9.6 14 21 30 82 90 37 42 21 

2-Methy1naphdialene nd nd 1 .5 3.0 1 .5 5.1 139 24 2.8 1 .4 4.1 4.9 7.5 29 8.3 2.4 12 5.7 

1-Methylnnphthalcne nd nd 0.81 1 .7 0.78 1 .9 65 15 1 .6 0.96 3 .3 4.1 4.7 19 5.1 1 .4 6.0 2.6 

2-Elltylnaplilhalcne nd nd 0?4 nd 0.13 nd nd nd 0.47 nd 0.91 1 .0 1 .2 3.9 nd 0.29 1,9 nd 

1-Elhylnaphthalene nd nd nd nd nd nd nd nd nd nd 0.59 0.72 0.48 nd nd 0.16 nd nd 

2,6/2,7-Dimelhylnaphdialene nd nd 0.91 0.93 0.70 nd nd nd 1 .1 nd 1 .6 1 .7 1 .9 14 4.6 1 .8 ~ 4.3 nd 

1,3/1,7-Dimethy1naphthalene nd nd 0.53 0.93 0.34 1.4 26 7.1 0.74 0.55 1.5 1 .6 2.7 8.0 2.6 2.4 4.5 1 .7 

1,6-Dimediylnaphtlialene nd nd 0.39 0.68 0.24 1 .1 20 5.7 0.51 0.20 0.99 1 .1 1 .3 5.4 2.0 0.65 2,3 1 .5 

1,4/2,3-Dimethylnaphthalene nd nd 0.32 nd 0.18 nd nd nd 0.57 0.60 1.0 1 .0 1 .6 5.3 2.6 2.5 3.7 nd 

1,5-Dimethylnaph(halene nd nd 0.16 nd nd nd nd nd 0.28 0.35 0.36 0.44 1 .5 3.0 0.55 1 .8 2.2 nd 

1,2-Dimethylnaphthalene nd nd 0.16 nd nd nd nd nd 0.26 nd 0.62 0.70 0.73 2.8 nd 0.79 0.97 nd 

2-Isopropylnuphthalene nd nd nd nd nd nd nd nd nd nd 0.09 nd nd nd nd nd nd nd 

1,8-Dimcthylnaplithulene nd nd nd nd nd nd nd nd nd nd 0.14 nJ nd nJ nd nd nd nd 

1,6,7-Trimethylnaphthalene nd nd 0.59 0.59 0.25 nd nd 28 1 .8 9.5 2,8 1 .8 11 28 19 21 49 6.2 

C3-NAPHTHALENES, est . nd nd 3.5 9.9 1 .8 nd nd 112 9.2 36 13 8.5 49 138 80 77 203 39 

C4NAPHTfiALENES, est . nd nd 3.1 nd 1 .5 nd nd 193 20 95 33 17 50 262 184 81 320 57 

Accnaphlhylcne nd nd nd nd nd nd nd nd nd nd 0.48 0.38 nd nd nd 0.24 . nd nd 

Accnaphthcne nd 3.9 0.76 1,9 0.74 nd 9.2 2.4 2.0 1 .5 1 .6 1 .7 4.0 7.5 5.0 6.2 6.7 1 .4 

Fluorene nd 6.0 0.94 2.7 0.86 2.0 14 8 .2 2.9 1 .7 2.1 2.6 5.6 11 10 6.5 8.0 2.5 

DibenzoUuoplune nd nd 1 .1 0.96 0.75 nd nd 0.68 0.45 nd 0.58 0.53 1 .5 0.86 2.1 6.5 2.1 nd 

Phenanthrene 13.2 6.9 3 .2 6.1 3.1 5 .3 18 7.6 6.0 4.0 7.6 8 .2 9.2 14 14 11 13 6.2 

Anthrucene nd 1 .2 0.36 0.79 0.55 0.76 nd 0.85 0.85 1 .2 0.87 0.90 2.8 2.8 3 .6 3.4 5.0 2.1 

4-Mcthyldibenzothiophene 2.7 1 .9 3.9 2.2 1 .8 nd nd 2.5 1 .2 5.3 2.7 1 .8 5.7 4.0 14 18 20 2.0 

2/3-Methyldibenzothiophena nd nd 0.87 nd nd nd nd nd 0.36 2.2 0.75 0.55 2.1 1 .3 3.6 8 .3 6.9 nd 

1-Methyldibenzothiophene nd nd 1 .0 nd nd nd nd nd 0.44 2.6 0.93 0.40 2.8 0.98 5.0 9.1 11 nd 

3-Melhylphenanthrene nd nd 0.78 0.67 nd nd nd 3.3 1 .2 7.6 2.6 1 .7 8.0 5.3 10 15 30 nd 

2-Melhylphenanthrene nd nd 0.36 0.38 nd nd nd 0.79 0.58 1 .6 1 .5 1 .4 1 .5 1 .1 3 .0 3 .7 4.7 nd 

4/9-Melhylphenanllircne nd nd 0.50 nd nd nd 239 1 .9 nd 4.9 3.0 1 .9 7.1 3.8 12 IS 36 nd 

1-Methylphenanthrene nd nd 0.60 0.50 nd nd nd 1 .8 nd 4.1 1 .6 1 .1 4.1 2.3 6.4 9.4 17 327 

3,6-DimeUiylphenanthrene nd nd 0.35 nd nd nd nd 3 .8 4.6 IS 9.9 6.8 8.1 13 30 13 43 3 .3 

3,5-DimcUiylphenvnthrene nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd 

2,6-DimethylphenunQuene nd nd 0.43 nd nd nd nd 2.3 3.1 7.9 5.4 3.7 4.0 6.6 16 6.8 23 nd 

F-~ 

hr 
!-+ Appendix A: Parent and alkylated PAH concentrations (ng/g) in oyster tissue from organisuis exposed to varying levels of contaminated pass Fourchon sediment for 28 days . 

N 

EXPOSURE LEVEL 

SAMPLING DAY 

ANALYTE 

2,7-Dimelhylphenant}uene 

3,9-Dimethylphenanthrcne 

1,6/2,5/2,9-Dimethylphenanthrene 

I ,7-Dimethylphenanlhrene 

1,9/4,9-Dimethylphenanlhrene 

1,S-Dimethylphenanthrene 

1,8-Dimethylphenanthrene 

I ,2-Dimethylphenanthrene 

9, l0-Dimelhylphenanthrene 

Fluoranlhcne 

1,2-Dimelhyldibenzothiophcne 

C2-DIBENZOTHIOPHENES, est . 

Pyrene 

1,2,8-Trimethylphenanlhrene 

C3-PHENANTHRENES, est. 

Benzanthracene 

Chrysene 

Benzo(b)fluoranthene 

Benzo(k)fluoranlhene 

Benzo(a)pyrene 

indenopyrene 

Dibenz(a,li)anthracene 

Henzo(S)i,i)perylene 

0% LEVEL (CONTROL) 112.5% LEVEL I25% LEVEL I50% LEVEL 

0 0-dup. 3 3-dup. 7 14 21 3 7 14 21 28 3 7 14 3 7 14 

nd nd nd nd nd nd nd nd 0.51 4.8 1 .1 0.78 0.99 7.7 3 .9 2 .2 13 nd 

nd nd 1.2 1.3 0.96 0.83 nd 9.6 7.0 37 19 9.4 19 23 72 29 112 26 

nd nd 0.62 nd nd nd nd 6.5 5.0 28 7.8 3 .2 13 15 39 22 89 14 

nd nd 0.37 nd nd nd nd 2.1 1 .9 83 4.7 2.2 4.2 5.7 15 7.0 26 4.7 

nd nd nd nd nd nd nd 2.1 1 .4 9.9 5.1 2.4 5.6 6.1 15 9.6 35 6.3 

nd nd nd nd nd nd nd nd nd nd 5.6 1 .7 nd nd nd nd nd nd 

nd nd nd nd nd nd nd 2.1 2.5 6.0 4.5 4.0 3.3 7.8 20 4 .9 19 6.0 

nd nd nd nd nd nd nd 0.95 1,0 3.5 1 .7 1 .1 1 .3 2.7 6 .7 2.7 11 3.3 

nd nd nd nd nd nd nd nd nd nd 0.36 nd nd nd nd 1 .14 nd nd 

2.6 1 .6 0.48 0.86 0.44 0.87 4.0 4.2 2.8 12 6.0 0.27 7.2 8 .9 19 15 53 7.9 

nd nd nd nd nd nd nd nd nd 0,64 0.67 nd nd nd 1 .2 0.61 1 .8 nd 

nd nd 19 15 16 13 nd 30 24 94 78 50 63 71 177 99 240 77 

nd nd 0.22 0.33 0.14 nd 2.7 1 .7 1 .5 7.2 3.8 0.21 3.8 2.8 11 8.1 27 1l 

nd nd 0.21 nd 0.12 nd nd 1 .4 4.4 10 6.0 5.3 2.4 4.0 26 5 .8 20 6.7 

nd nd 11 19 12 13 nd 56 117 363 200 150 90 176 723 194 684 264 

7.1 4.6 nd nd nd nd 18 nd 2.0 3.6 2.0 1 .5 1 .9 1 .6 6.2 1 .9 9.8 13 

nd nd nd nd nd nd nd 2.3 3.0 12 9.0 4.9 7.1 8.4 17 8.4 31 30 

nd nd nd nd nd nd nd nd nd nd 1 .6 1 .1 nd 0.59 nd 0.91 3 .0 4.3 

nd nd nd nd nd nd nd nd nd nd 0.58 0.19 nd 0.52 nd 0.40 0.78 1 .2 

nd nd nd nd nd nd nd nd nd nd 0.21 nd nd nd nd nd nJ nd 

nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd 

nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd nd 

nd nd nd nJ nd nd nd nd nd nd nd nd nJ nd nd nd nd nd 
N, Nt QPP OaTy ~'y The Department of the Interior Mission 

a`' ~ p As the Nation's principal conservation agency, the Department of the Interior has responsibility for 

most of our nationally owned public lands and natural resources. This includes fostering sound 

use of our land and water resources; protecting our fish, wildlife, and biological diversity; 

M 

preserving the environmental and cultural values of our national parks and historical places; 

ACH s rt t ~ and providing for the enjoyment of life through outdoor recreation. The Depa men assesses our 

energy and mineral resources and works to ensure that their development is in the best interests 

of all our people by encouraging stewardship and citizen participation in their care. The 

Department also has a major responsibility for American Indian reservation communities and for 

people who live in island territories under U.S . administration . 

The Minerals Management Service Mission 

As a bureau of the Department of the Interior, the Minerals Management Service's (MMS) primary 

responsibilities are to manage the mineral resources located on the Nation ' s Outer Continental Shelf 

(OCS), collect revenue from the Federal OCS and onshore Federal and Indian lands, and distribute 

those revenues. 

+ . GIL w. ~ Moreover, in working to meet its responsibilities the Offshore Minerals Management Program 

administers the OCS competitive leasing program and oversees the safe and environmentally sound 

exploration and production of our Nation's offshore natural gas, oil and other mineral resources. 

The MMS Royalty Management Program meets its responsibilities by ensuring the efficient, timely 

and accurate collection and disbursement of revenue from mineral leasing and production due to 

Indian tribes and allottees, States and the U.S. Treasury . 

The MMS strives to fulfill its responsibilities through the general guiding principles of: (1) being 

responsive to the public's concerns and interests by maintaining a dialogue with all potentially 

affected parties and (2) carrying out its programs with an emphasis on working to enhance the 

quality of life for all Americans by lending MMS assistance and expertise to economic development 

and environmental protection.