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dissertation entitled

Instrumental and Bioanalytical Measures of
' Endocrine Disruptors in Water

presented by
Shane Allen Snyder

has been accepted towards fulfillment
of the requirements for

Doctor of Philom degree in ZOOlOQ‘V and
Environmental Toxicology

 

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Instrumental and Bioanalytical Measures of Endocrine Disruptors in Water

By

Shane Allen Snyder

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Zoology and Institute for Environmental Toxicology

2000

 

ABSTRACT

Instrumental and Bioanalytical Measures of Endocrine Disruptors in Water

By

Shane Allen Snyder

A number of compounds released into the environment by human
activities can modulate endogenous hormone activities and have been termed
endocrine-disrupting compounds (EDCs). It has been hypothesized that such
compounds may elicit a variety of adverse effects in both humans and wildlife,
including promotion of hormone-dependent cancers and reduction in
reproductive fitness. Concern over EDCs has created a need to monitor for
these compounds in the environment. This need is underscored by recent
legislation mandating that chemicals and formulations be screened for potential
estrogenic activity before they are manufactured or used in certain processes
(Safe Drinking Water Act Amendments of 1995 - Bill Number S1316; Food
Quality Protection Act of 1996 - Bill Number PL. 104-170). Several methods for
the sensitive detection and quantitation of EDCs from water and tissue samples
have been developed.

Nonylphenol (NP), a minor transient degradation product of the widely
used nonylphenol polyethoxylate (NPE) non-ionic surfactants, was tested for
bioaccumulation in fathead minnows. After a 42 d continuous exposure, fish

tissues were analyzed. The resulting bioconcentration factors (BCFs) ranged

from 203 — 268. While this method was appropriate for fathead minnows
weighing less than 2 9, it was not applicable for larger tissue sample sizes.
Therefore, a more reliable method was desired. Extractive steam-distillation was
employed to extract NP and lower oligomer NPEs from fish tissue samples.
Optimization of this method resulted in consistent recoveries greater than 70%
and detection limits less than 20 ng/g. The method was tested using carp
captured in the Las Vegas Bay of Lake Mead, Nevada. The average
concentration of NP detected in the carp tissue was 184 ng/g wet weight.

Wastewater effluents and surface waters from Michigan and Nevada were
screened for estrogenic and anti-estrogenic compounds using a novel method
which combines analytical chemistry and in vitm bioassays. Bioassay-directed
chemical fractionation and instrumental analyses indicated that all observed
estrogenicity was caused by the fraction of greatest polarity. Results indicate
that the natural steroid 17B-estradiol and the synthetic steroid ethynylestradiol
were most likely responsible for observed bioactivity.

A follow-up study was initiated at Lake Mead, Nevada. Water samples of
100 L volume were extracted and analyzed using various instrumental and
bioanalytical methods. Concentrations of priority pollutants were found to be low
relative to other sites in the US. However. several new contaminants were
identified. These included pharmaceuticals, flame-retardants, and personal care
products. Many of these compounds have not been reported previously in the

surface waters of North America.

Copyright by
Shane Allen Snyder
2000

 

ACKNOWLEDGMENTS

The reseach contained herein would not have been possible without the
guidance and support of many people and organizations. I would like to thank
Dr. John Giesy for serving as my advisor and for his years of support and
friendship. I would also like to thank Dr. Frank D’ltri, Dr. Robert Huggett, and Dr.
Matthew Zabik for serving as members to my graduate committee and for their
time and efforts in helping me achieve my research and professional goals.

I would like to express my sincere appreciation to my friends and
colleagues, without their support this research would not have been possible.
Acknowledgments for specific components of my research are made in the
following chapters. The MSU Aquatic Toxicology Laboratory has provided me
with a breadth of experience. This was in no small part due to interactions with
peers of various expertise and I would like to express my appreciation for their
help and support. I would like to specifically thank Erin Snyder, Dan Villenueve,
Tim Keith, Kevin Henry, Dave Verbrugge, K. Kannan, and Pete Horvath.

The organizations which provided funding and material and technical
assistance are acknowledged in the appropriate chapters of this dissertation.
However, I would like to specifically thank those who have inspired and guided
my research, namely: Dr. Carter Naylor of Huntsman Corporation; Dr. Kevin
Kelly, Margaret Lake, and Dr. James LaBounty of US. Department of the Interior
- Bureau of Reclamation; Peggy Roefer, Kim Zikmund, and Kay Brothers of the

Southern Nevada Water Authority; Doug Karafa and Bill Shepherd of the Clark

 

County Sanitation District; and Bill Burke, Fred DeSiIva, and Roy Irwin of the

National Park Service.

vi

TABLE OF CONTENTS
LIST OF TABLES ............................................................................................. . ..... x
LIST OF FIGURES ............................................................................................... xii
INTRODUCTION ................................................................................................... 1

CHAPTER 1. OVERVIEW OF INSTRUMENTAL AND BIOANALYTICAL

MEASURES OF ENDOCRINE DISRUPTERS IN WATER ................................... 6
INTRODUCTION ......................................................................................... 6
IN VITRO BIOASSAYS ............................................................................. 11
IN VIVO VERSUS IN VITRO BIOASSAYS FOR TESTING EDCs ........... 16
USE OF FISH TO TEST FOR EDCs ........................................................ 16

Selection of test species ................................................................. 18
End points and biomarkers used to assess effects of EDCs .......... 18
Field Studies ................................................................................... 19
Laboratory Studies ......................................................................... 20
MONITORING OF BUTYL TIN COMPOUNDS IN WATER ............... . ...... 26
Ecological impacts .......................................................................... 28
Instrumental analysis and biomoniton'ng ........................................ 29
BlOASSAY-DIRECTED CHEMICAL FRACTIONATION .......................... 32
CONCLUSIONS ........................................................................................ 35
REFERENCES .......................................................................................... 38

CHAPTER 2. BIOCONCENTRATION OF NONYLPHENOL IN FATHEAD

MINNOWS (PIMEPHALES PROMELAS) ............................................................ 50
INTRODUCTION .................................... ' ...... . ............................................ 5 0
EXPERIMENTAL SECTION ..................................................................... 53

Standards and reagents ............ . .................................................... 53
Fish exposure .............................. . .................................................. 54
Water extraction ................................ - ............................................ 55
Fish extraction and clean-up .......................................................... 56
Quantitation of compounds ............................................................ 57
Recovery and precision ............... . .................................................. 57
RESULTS .................................................................................................. 58
DISCUSSION ............................................................................................ 59
ACKNOWLEDGMENTS ............................................................................ 60
REFERENCES .......................................................................................... 60

vii

CHAPTER 3. A METHOD FOR DETERMINING CONCENTRATIONS OF
NONYLPHENOL AND LOWER OLIGOMER NONYLPHENOL ETHOXYLATES

IN FISH TISSUES ................................................................................................ 69
ABSTRACT ............................................ . .................................................. 69
INTRODUCTION ....................................................................................... 70
EXPERIMENTAL SECTION ..................................................................... 72

Standards and reagents ................................................................. 72
Sample collection and preservation ............................................... 72
Extraction ....................................................................................... 73
NonnaI-phase HPLC fractionation .................................................. 74
Identification and Quantitation ........................................................ 75
Recovery and Detection Limits ....................................................... 75
RESULTS AND DISCUSSION .................................................................. 76
ACKNOWLEDGMENTS ................................................................ . ........... 77
REFERENCES .......................................................................................... 77

CHAPTER 4. ANALYTICAL METHODS FOR DETECTION OF SELECTED

ESTROGENIC COMPOUNDS IN AQUEOUS MIXTURES ................................. 84
ABSTRACT .................................................. . ........................................... 84
INTRODUCTION ....................................................................................... 85
EXPERIMENTAL SECTION ..................................................................... 88

Standards and reagents ................................................................. 88
Sample collection and preservation ............................................... 90
Extraction ....................................................................................... 92
Normal-phase HPLC fractionation .................................................. 92
Quantitation of compounds ............................................................ 93
Radioimmunoassay (RIA) .............................................................. 94
Recovery and Precision ................................................................. 95
Detection Limits ................................................. .......................... 97
RESULTS .................................................................................................. 97
DISCUSSION ............................................................................................ 99
ACKNOWLEDGMENTS .......................................................................... 1 03
REFERENCES ........................................................................................ 104

CHAPTER 5. TOXICITY IDENTIFICATION AND EVALUATION OF
ESTROGENIC AND DIOXIN-LIKE COMPOUNDS IN WASTEWATER

EFFLUENTS ...................................................................................................... 118
ABSTRACT ............................................ .. ................................................ 118
INTRODUCTION ..................................................................................... 119
MATERIALS AND METHODS ................................................................ 122

Sample collection and fractionation .............................................. 122

viii

Cell culture and bioassay ............................................................. 122

RESULTS AND DISCUSSION ................................................................ 126
Dioxin-Iike activity .............................. . .......................................... 126
Estrogen-like activity .................................................................... 127

SUMMARY .............................................................................................. 1 32

REFERENCES ........................................................................................ 133

CHAPTER 6. BIOACTIVE ORGANIC COMPOUNDS IN THE WATERS OF

LAKE MEAD, NEVADA, USA ............................................................................ 154
ABSTRACT ............................................................................................. 154
INTRODUCTION ..................................................................................... 155
EXPERIMENTAL SECTION ................................................................... 158

Standards and reagents ............................................................... 158
Sample collection ........................................................................ 159
Extraction .................................... , ................................................ 160
Silica gel fractionation ............................................... . .................. 162
Identification and quantitation ....................................................... 163
Recovery and precision ................................................................ 165
Detection Limits ............................................................................ 165
RESULTS AND DISCUSSION ................................................................ 166
Analytical considerations .............................................................. 166
Organochlon'ne pesticides and PCBs ........................................... 167
Polycyclic aromatic hydrocarbons ................................................ 168
Alkylphenols and EPA .................................................................. 169
Pharmaceuticals .......................... . ................................................ 1 70
Other compounds ......................................................................... 1 71
High-resolution mass spectrometry .............................................. 172
SUMMARY .............................................................................................. 172
ACKNOWLEDGMENTS .......................................................................... 172
REFERENCES ........................................................................................ 173

LIST OF TABLES

CHAPTER 1

Table 1. End points used to assess reproductive impacts of EDCs in

fish ............................................................................................................. 37
CHAPTER 2

Table 1. Detection Limits for Compounds of Interest ............................... 65

Table 2. Bioconcentration of NP in Fathead Minnows (mean

concentrations) .......................................................................................... 66
CHAPTER 3

Table 1. Ions Monitored and Recovery Data ........................................... 80

Table 2. Detection Limits for Compounds of Interest ............................... 81

Table 3. Average Concentrations of NP and NPEs in Lake Mead Carp..82

CHAPTER 4
Table 1. Recoveries through the analytical procedure (n=3) (%
Recovered i SD) ..................................................................................... 108
Table 2. Dection Limits for Selected Xenoestrogens ............................. 109
Table 3. Concentrations of Selected Xenoestrogens in the Trenton
Channel ................................................................................................... 110
Table 4. Concentrations of Selected Xenoestrogens at
WWTPs ................................................................................................... 111
Table 5. Concentrations of Selected Xenoestrogens in Lake Mead ...... 112
CHAPTER 5
Table 1. Extract Concentrations ............................................................ 138
Table 2. E2-EQs and Predicted Responses .......................................... 139
CHAPTER 6

Table 1. F1 and F2 Compounds Detected, Estimated Method Detection

Limits (nglL), and Ions Monitored ............................................................ 176
Table 2. Pharmaceuticals Compounds (F3) Detected, Estimated Method
Detection Limits (nglL), and Ions Monitored ........................................... 177
Table 3. F3 Compounds Detected, Estimated Method Detection Limits
(nglL), and Ions Monitored — Part I ......................................................... 178
Table 4. F3 Compounds Detected, Estimated Method Detection Limits
(nglL), and Ions Monitored — Part II ........................................................ 179
Table 5. Concentrations of HCHs in Lake Mead (ng/L) ......................... 180

Table 6. Concentrations of DDTs Detected in Lake Mead (nglL) .......... 181

Table 7. Concentrations of Alkylphenols and Bisphenol A in Lake Mead
(ng/L) ....................................................................................................... 182

Table 8. Concentrations of Pharmaceuticals in Lake Mead (ng/L),
Part I ........................................................................................................ 183

Table 9. Concentrations of Pharmaceuticals in Lake Mead (ng/L),
Part II ....................................................................................................... 184

Table 10. Concentrations of Pharmaceuticals in Lake Mead (ng/L),
Part III ...................................................................................................... 185

Table 11. Concentrations of Other Compounds in Lake Mead (ng/L)...186
Table 12. Compounds Identified in Las Vegas Bay by HRMS .............. 187

xi

LIST OF FIGURES

CHAPTER 1
Figure 1. Mechanism for activation of the estrogen receptor (ER) in
MVLN cells ................................................................................................ 49
CHAPTER 2
Figure 1. Structures of target compounds ............................................... 67

Figure 2. Chromatograms of target compounds in fish tissues: (A)
standards (B) fish tissue extract from 3.4 pg NP/L exposure (C) fish

tissue extract from 0.4 pg NP/L exposure ................................................. 68
CHAPTER 3

Figure 1. Structures of compounds of interest ........................................ 83
CHAPTER 4

Figure 1. Structures of target compounds ............................................. 113

Figure 2. Flow diagram of analytical method ......................................... 114

Figure 3. Reverse-phase HPLC of F2: (A) spiked standards; (B) Trenton
Channel - Black Lagoon 8/30/97; (C) WWI'P - BV effluent 10l8l97 ....... 115

Figure 4. Reverse-phase HPLC of F3: (A) spiked standards; (B) Trenton-
Channel — Black Lagoon 8/30/97; (C) WWTP — BV effluent 10l8l97 ...... 116

Figure 5. Normal-phase HPLC of NPE oligomer distribution: (A) NPE

standard; (B) extracted NPE ................................................................... 117
CHAPTER 5

Figure 1. Flow diagram of analytical method ......................................... 140

Figure 2. Reverse-phase HPLC fine fractionation ................................. 141

Figure 3. Dioxin-Iike bioactivity (various sites): Significance bars
represent 3x SD of the background sign ................................................. 142

Figure 4. Estrogen-like bioactivity — Michigan WWTPs (October 1997):
Significance bars represent 3x SD of the background signal .................. 143

xii

Figure 5. Estrogen-like bioactivity — Trenton Channel (August 1997):
Significance bars represent 3x SD of the background signal .................. 144

Figure 6. Estrogen-like bioactivity - Lake Mead (April 1997): Significance
bars represent 3x SD of the background signal ...................................... 145

Figure 7. Estrogen-like bioactivity — Lake Mead (September 1997):
Significance bars represent 3x SD of the background signal .................. 146

Figure 8. Fine fractionation and corresponding estrogen-like bioactivity of

WWTP BV-effluent F2 extract ................................................................. 147
Figure 9. Fine fractionation and corresponding estrogen-like bioactivity of
Black Lagoon, Trenton Channel, F2 extract ............................................ 148
Figure 10. Fine fractionation and corresponding estrogen-like bioactivity
of LV Wash, Lake Mead (April 1997), F3 extract .................................... 149
Figure 11. Fine fractionation and corresponding estrogen-like bioactivity
of LV Bay, Lake Mead (April 1997), F3 extract ....................................... 150
Figure 12. Further fractionation and corresponding estrogen-like
bioactivity of LV Wash, Lake Mead (April 1997), F3 extract ................... 151
Figure 13. Fine fractionation and corresponding estrogen-like bioactivity
of WWTP BV-effluent, F3 extract ............................................................ 152
Figure 14. Fine fractionation and corresponding estrogen-like bioactivity
of Black Lagoon, Trenton Channel, F3 extract ........................................ 153
CHAPTER 6
Figure 1. Map of the Boulder Basin study area of Lake Mead,
Nevada .................................................................................................... 188
Figure 2. Flow diagram of analytical method ........................ . ................. 189

Figure 3. Mass spectra of phenytoin (Dilantin): (A) authentic standard;
(B). LV Bay water extract ........................................................................ 190

Figure 4. Mass spectral scan of phenobarbital: (A) authentic standard;
(B) LV Bay water extract ........................ . ................................................ 191

Figure 5. Mass spectral SIM Chromatograms I: (A) authentic standards;
(B) LV Bay water extract ......................................................................... 192

xiii

Figure 6. Mass spectral SIM Chromatograms II: (A) standards; (B) LV
Bay water extract. Peak # 1. DEET; 2. Guafenesin; 3. Acetaminophen; 4.
Meprobamate; 5. Caffeine; 6. Oxybenzone; 7. Bisphenol A; 8. A8 THC; 9.
A9 THC; 10. Primidone; 11. Carbamazepine; 12. Hydrocodone ............. 193

Figure 7. Mass spectral scan of DEET: (A) authentic standard; (B) LV
Bay water extract .................................................................................... 194

Figure 8. Mass spectral scan of tris(1 ,3-dichIoroisopropyl)phosphate: (A)
NIST database spectra; (B) LV Bay water extract .................................. 195

xiv

INTRODUCTION

The threat of modulation of animal endocrine systems by xenobiotic
compounds has become a major issue facing society today. These compounds
are varied in form and mechanism of action. This poses unique challenges in the
identification and evaluation of these compounds from environmental matrixes.
The aquatic environment is exceptionally susceptible to xenobiotic insult. Water
can be polluted by a multitude of sources and acts as a sink for many types of
pollution. This chapter outlines several methods for the detection and
quantitation of endocrine disrupting compounds (EDCs) in the aquatic
environment. However, no single method alone can predict or detect all EDCs
present in an environmental sample, nor can all the biological mechanisms of
action be accounted for in one simple test. Therefore, comprehensive screening
for EDCs must combine several types of analyses including in vivo and in vitro
bioassays, and analytical chemistry. This dissertation will discuss several of
these types of analyses and demonstrate advantages and disadvantages of
each. Each chapter of this dissertation is in journal format and publication status
is provided.

Chapter one of this dissertation is an overview of the research conducted
at the Aquatic Toxicology Laboratory (ATL) at Michigan State University (MSU)
related to endocrine disruption in the aqautic environment. The role of various
types of in vitro and in vivo bioassays for the detection of enodcrine disrupting
chemicals will be discussed. Specific examples are provided as well as

advantages and disadvantages of each type of analysis. A discussion of

butyltins is provided as an example of a unique type of endocrine disrupting
chemical (EDC) which has been shown to cause adverse effects in the
environment. The final section in this chapter deals with toxicity identification and
evaluation of EDCs. Here we integrate results from in vitro bioassays with
instrumental analyses. By using bioassay-directed fractionation and subsequent
instrumental analyses, we are able to determine the compounds of greatest
potency from complex mixtures. In conclusion, we discuss the need to use an
integrated approach combining in vitro and in vivo bioassays with advanced
instrumental techniques to determine what compounds in the aquatic
environment may have the potential to impact the endocrine systems of aquatic
organisms.

Chapter two discusses a controlled laboratory study to determine the
bioconcentration of p-nonylphenol (NP) in fathead minnows. NP is one of
several transient intermediate degradation products of the widely used
nonylphenol polyethoxylate (NPE) nonionic surfactants. NP is a ubiquitous
contaminant of wastewater treatment plant (WWT P) effluents. In both in vitro
and in vivo bioassays, NP has been shown to be estrogenic. NP is fairly
lipophilic and persistent, therefore, it was desirable to determine
bioconcentrations factors (BCFs) in an aquatic organism. Methods for
determining NP concentrations in water and tissue were developed. BCFs
ranging from 203 to 268 were determined.

Chapter three expands upon the methodology developed in chapter two.

A monitoring study for NP in the tissues of fish was desired. However, no

reliable and sensitive methods for the identification and quantitation of NP in
tissue samples were available. The methodology presented in chapter two did
not provide the necessary sensitivity for a monitoring study. Several methods
were investigated. Also, the methodology was to be readily transferable to other
laboratories by using common and/or inexpensive equipment. Extract steam-
distillation followed by gas chromatography with mass selective detection
(GC/MS) was developed. This method provided method detection limits (MDLs)
ranging from 2 — 5 ng/g, wet weight, for NP and nonylphenol mono— and di-
ethoxylate. Themethod was also highly reproducible with coefficients of varation
(CVs) generally less than 10% among subsamples. The method was tested
using carp captured in the Las Vegas Bay of Lake Mead, Nevada. Average
tissue concentrations of NP and nonylphenol monoethoxylate (NPE1) were
determined to be 184 and 242 ng/g, respectively.

The fourth chapter presents a novel method to determine concentrations
of certain estrogenic compounds in aqueous mixtures. It provides the first report
in North America of natural and sythetic estrogens in surface waters.
Octylphenol (OP), NP, and NPE were also identified and quantified. These
compounds were extracted in situ from 5 L water samples using solid-phase
extraction (SPE) disks. The resulting extracts were fractionated based on
polarity. Various instrumental techniques were used to identify and quantify
compounds of interest. The study locations include four WWl'Ps in central
Michigan; the Trenton Channel of the Detront River, Michigan; and Lake Mead,

Nevada. Concentrations of NP and OP ranged from less than the MDL to 37 and

0.7 pg/L, respectively. NPE concentrations ranged from less than the MDL to
332 pg/L. Concentrations of 17B-estradiol (endogenous estrogen) and
ethynylestradiol (synthetic estrogen) ranged from less than the MDLs to 3.7 ng/L
and 0.8 ng/L, respectively.

The fifth chapter describes a toxicity identification and evaluation (TIE)
method for estrogen- and dioxin-like compounds in complex aqueous mixtures.
Extracts from the method presented in chapter three were tested using in vitro
bioassays. Whole extracts and corresponding fractions were tested for estrogen-
and dioxin-like activity using MVLN and H4IIE-Iuc in vitro bioassays, respectively.
No significant dioxin-like activity was observed. Estrogenic activity was
quantitated by comparing the magnitude of induction elicited by the extract or
fraction to the maximum induction caused by the endogenous estrogen, 17B-
estradiol (E2). Several extracts and fractions displayed significant estrogenic
activity. In each case, the most polar fraction (F3) was associated with the
greatest estrogenicity. Instrumental analyses and further fractionation were used
to identify compounds associated with estrogenic responses. Mass balance
calculations suggested that E2 and the synthetic estrogen, ethynylestradiol
(EE2), were the most likely cause of the responses observed. Alkylphenols
(APs) did not appear to contribute significantly to the observed estrogenicity.

The final chapter of this dissertation (chapter 6) expands upon the
methods developed in the previous chapters to determine the concentrations of
several classes of bioactive compounds in the waters of Lake Mead, Nevada.

Several recent reports have indicated a flux of organic compounds flowing into

the Boulder Basin of Lake Mead, Nevada from the Las Vegas Wash. The
primary objective of this study was to screen for a wide range of organic
contaminants in Lake Mead. Mass spectrometry was utilized to screen for
previously unreported compounds. Samples of 100 L of filtered water were
extracted using large-scale SPE. Following a series of elution and extraction
procedures, resulting extracts were fractionated into 3 polarity classes. Each
fraction was analyzed by gas chromatography with mass selective detection
(GC/MS). DDT, DDE, and DDD (collectively, DDTs) and four isomers of
hexachlorocyclohexane (HCH) were detected at some sites, with concentrations
ranging from less than detection to 28 nglL. Octylphenol, nonylphenol,
nonylphenol monoethoxylate, and nonylphenol diethoxylate were detected in
some samples, with concentrations ranging from less than detection to 1500
nglL. Caffeine, nicotine, oxybenzone, N,N-diethyl-meta-toluamide (DEET),
phosphate-based flame-retardants, and several pharmaceuticals were detected
in some samples with concentrations ranging from less than detection to 360
nglL. The identities of several of these previously unreported compounds were
verified using high-resolution mass spectrometry. PAHs were detected at each
site and did not vary significantly among the sites in concentration or in pattern.
Total PAH concentrations ranged from 5 to 78 nglL. Polychlorinated biphenyls,

triazine herbicides, and steroids were not detected in any of the samples.

Chapter 1

OVERVIEW OF INSTRUMENTAL AND BIOANALYTICAL MEASURES OF
ENDOCRINE DISRUPTERS IN WATER

(Published as a chapter in the American Chemical Society Book: “Analysis of
Environmental Endocrine Disruptors” 2000)

A number of compounds released into the environment by human
activities can modulate endogenous hormone activities and have been termed
endocrine-disrupting compounds (EDCs) (1, 2). Environmental EDCs have been
defined as exogenous agents which interfere with the "synthesis, secretion,
transport, binding, action, or elimination of natural hormones in the body that are
responsible for the maintenance of homeostasis, reproduction, development,
and/or behavior" (3). It has been hypothesized that such compounds may elicit a
variety of adverse effects in both humans and wildlife, including promotion of
hormone-dependent cancers, reproductive tract disorders, and reduction in
reproductive fitness (4-10). Endocrine "disruption" by environmental
contaminants has become a cause for concern (11) and as a result there is an
urgent need to develop methods for monitoring EDCs in the environment. This
need is underscored by recent legislation mandating that chemicals and
formulations be screened for potential estrogenic activity before they are
manufactured or used in certain processes (Safe Drinking Water Act
Amendments of 1995 - Bill Number 8.1316; Food Quality Protection Act of 1996 -
Bill Number PL. 104-170).

EDCs act through a number of mechanisms including receptor-mediated
hormonal responses such as hormone mimics (1). These include xenobiotic
effects through thyroid hormone receptor, epidermal growth factor (EGF)
receptor, aryl hydrocarbon receptor (AhR), estrogen receptor (ER) and androgen
receptor (AR) (12, 13). Of all the EDCs, those that are direct-acting estrogen
receptor (ER) agonists (xenoestrogens or estrogen mimics), direct acting ER
antagonists (antiestrogens), or androgen antagonists have received the greatest
attention, due in part to their importance in embryonic development. The
“estrogen hypothesis” stemmed from the observation that some of the effects
observed in oviparous wildlife exposed to persistent and bioaccumulative
chemicals were similar to those that could be caused by injecting estrogen into
eggs (2). This hypothesis was supported by the fact that naturally occurring
exoestrogens, such as phytoestrogens, could cause reproductive dysfunction in
animals. Further support came from the observation that some xenobiotics that
can bind the ER are weak estrogen agonists or antagonists in in vitro expression
assays (12). Because the major initial concern over endocrine modulating
compounds has been the estrogen mimics (xenoestrogens), we will restrict our
discussion to “estrogenic” and “antiestrogenic” compounds, but the techniques
presented can be applied to any receptor-mediated process for which a suitable
in vitro cell system can be developed (12).

There are a wide variety of ER-active compounds in the environment (14).
These include both natural products (11) and synthetic compounds (10). The

synthetic compounds include both chlorinated and non-chlorinated compounds

that can either mimic or antagonize the effects of endogenous estrogen (12).
Some of the compounds found in the surface waters that have been reported to
be estrogenic include nonylphenol (NP), octylphenol (OP), 17B-estradiol (E2)
(produced in the body) and the synthetic estrogen ethynylestradiol (EE2) (used in
oral birth control medication) (15).

Several types of in vitro assays are available for measuring the estrogenic
activity of single compounds or complex mixtures. The range of responses
includes everything from simple receptor binding, to expression of endogenous
or exogenous genes, to cell proliferation and differentiation. In vitro systems are
attractive as screening tools because they are rapid, inexpensive, and fairly
reproducible. For these reasons, precise estimates of the relative potency of a
great many samples or compounds can be obtained in a rather short period of
time. Expression assays examine induction or suppression of proteins encoded
by genes whose transcription is thought to be modulated through an ER-
mediated mechanism. Increases or decreases in the activity of the protein of
interest upon exposure to a single compound or complex mixture, such as an
environmental extract, suggest the presence of one or more ligands with the
potential to modulate a broad range of genomically controlled estrogenic
responses.

Although in vitro assays are an attractive option for screening and
mechanistic studies, they may miss effects that would take place only in whole
organisms. Some toxicants require metabolic activation through one or more

pathways that occur in vivo, but not in vitro. Homeostatic controls and

bioaccumulation generally are not simulated by in vitro testing systems. Toxicant
effects can occur by different mechanisms in multiple tissues simultaneously. In
addition, the same chemical exposure can result in very different responses in an
animal depending upon its life stage, sex, and reproductive state. Fish are useful
in vivo models for testing for effects of endocrine disrupting chemicals in water.
Some species are extremely sensitive to such effects, and fish can act as
integrators of responses to mixtures of toxicants that occur in the environment.
In addition, effects related to growth and reproduction in fish are more easily
related to population-level and ecological effects than are effects in in vitro
systems. Fish can also be used in conjunction with in vitro testing systems to
investigate mechanisms of toxicant action and to calibrate in vitro to in vivo
responses.

The aquatic environment is especially susceptible to the effects of
contaminants. Effluents from municipal and industrial wastewater treatment
facilities, storm and agricultural run-off, and boating add many exogenous
compounds to the aquatic ecosystem. Of the most frequently studied EDCs,
butyltin compounds are next in abundance to alkylphenols in wastewater
effluents. These compounds are also present in surface waters from many
locations. The impact of tributyltin on the marine environment has received
considerable attention during recent years because of its high toxicity to aquatic
organisms (16-20). The use of TBT as an antifouling agent in paints applied on

ships results in direct contact with aquatic environments. However, studies on the

contamination of surface water (lakes and rivers) by butyltin compounds
originating from the disposal of municipal wastewater are limited.

In the aquatic environment, instrumental analytical techniques are
used to identify and quantify EDCs in sediments, tissues, and water. In some
cases the toxicant is already known. In other cases, screening can be used to
identify new toxicants based on bioassay-directed fractionation coupled with
subsequent instrumental analyses. This is part of a toxicity identification and
evaluation (TIE). By using bioassay-directed fractionation, compounds with
certain mechanisms of action can be isolated and identified. A mass-balance
can then be performed to see if all observed bioactivity can be accounted for by
the compounds identified (21). This comprehensive approach allows for
screening of various types of EDCs and may identify yet unknown compounds
with significant bioactivity. The issue of environmental endocrine disruption is
extremely complex. No single method will identify or explain all types of EDCs in
the aquatic environment. Integrated approaches that combine in vitro and in vivo
bioassays with analytical chemistry techniques are needed to screen for and
assess the effects of EDCs in the environment. This introduction should serve as
an overview of work completed at the Aquatic Toxicology Laboratory (ATL) at
Michigan State University relative to instrumental and bioanalytical techniques

used for the determination of endocrine disrupters in water.

10

IN VITRO BIOASSAYS

In vitro bioassays have an important role to play in an integrated approach
to the study of endocrine disrupting compounds in the environment (12). In vivo
studies in both the field and laboratory are critical for linking exposure to
biologically relevant effects. They are, however, impractical for routine, high
throughput screening, and characterization of individual compounds or
environmental samples. Procedurally, in vitro bioassays are typically performed
more quickly and at significantly less cost than in vivo studies. Use of simplified
biological systems circumvents much of the inter-individual, seasonal, and
temporal variability, which can confound interpretation of in vivo responses.
Additionally, in vitro bioassays avoid many of the complex socio—political and
ethical issues associated with whole animal studies. Thus, in vitro bioassays are
more amenable systems for routine use than in vivo assays. In vitro bioassays
use simplified biological or biochemical systems to measure responses elicited
by exposure to individual compounds or environmental samples. The responses
measured are generally highly sensitive, integrated, and mechanistically-based.
These properties allow in vitro bioassays to overcome many of the limitations of
in vivo studies and instrumental analyses.

While instrumental analyses are essential for the identification and
quantitation of compounds, they provide no information regarding the efficacy
and potency of these compounds to modulate endocrine function. In vitro

bioassays measure mechanistically-based biological responses. In vitro

11

bioassays provide an integrated measure of the combined potency of all
compounds in a sample. This is a practical advantage, because instrumental
analysis of complex mixtures can be very expensive, difficult, and time
consuming. Additionally, in vitro bioassays can detect compounds for which
there are no analytical methods available and can be useful tools to measure
potential additive and non-additive interactions between compounds. When the
biological response being measured is highly sensitive, in vitro bioassays can
also account for compounds that can exert a biological effect at concentrations
less than analytical detection limits. 17B-estradiol and ethynylestradiol, for
instance, can elicit significant responses in MVLN cells at concentrations less
than 50 pM (a 15 ppt) (12), while sensitive instrumental analyses, such as HPLC
fluorescence which has a detection limit of approximately 100 ng/mL (15) (z 350
nM) for these compounds. As a result, in vitro bioassays provide information
which can complement instrumental analyses.

In vitm bioassays have been used widely for the study of potentially
estrogenic compounds (12). Competitive receptor binding assays (12, 22-25),
cell proliferation assays (26-30) and in vitro gene expression assays (12, 24, 29,
31-35) are among the most common in vitro approaches employed in the study
of estrogenic and other endocrine disrupting compounds. The mechanistic
foundation, advantages, and disadvantages of these approaches have been
described elsewhere (12, 22, 36).

Work in our laboratory has primarily involved the use of the MVLN (MCF-

7-Iuc) in vitro gene expression assay. MVLN cells are human breast carcinoma

12

cells stably transfected with a Iuciferase reporter gene under control of estrogen
responsive elements (EREs) of the Xenopus vitellogenin A2 gene (37, 38).
When cells are exposed to a single compound or environmental sample, ER
ligands can enter cells and bind the ER (Fig. 1). In MVLN cells, binding also
upregulates expression of an exogenous Iuciferase reporter gene (Fig. 1). Upon
addition of the appropriate substrate, Iuciferase catalyzes a light producing
reaction. As a result, luciferase activity can be measured conveniently and with
great sensitivity using a 96 well plate-reading Iuminometer. The measured
Iuciferase activity is proportional to the sample’s ability to modulate ERE-
mediated gene expression.

Although gene expression in MVLN and other estrogen responsive cells is
generally regarded to be ligand induced, there are also ligand-independent
pathways that can modulate the transcriptional activity of the ER through
activation of protein kinases (12). For example, treatment of cells with growth
factors or agents that activate protein kinases in general, can cause the ER to be
activated (without ligand binding) to bind to EREs and activate transcription.
Complex interactions within mixtures are also possible. Investigations have
included mixtures of estrogenic and anti-estrogenic compounds (39) and
mixtures of weak estrogens (12).

Use of the MVLN assay by our laboratory provides examples of useful
applications of in vitro bioassays as part of an integrated approach to the study of
endocrine disrupters. Screening with the MVLN bioassay has identified both

individual compounds and complex mixtures extracted from environmental

13

samples that were able to induce ERE-mediated gene expression (12, 15, 40-
42). Active samples have been compared in terms of either their relative efficacy
(the magnitude of response elicited; (15, 40, 41)) or relative potency (the mass,
volume, or concentration of sample or compound needed to elicit some defined
level of response; 1). Environmental samples from a diverse array of sources
including surface waters (15, 43), wastewaters (15), sediments (40), and
commercial fish diets (42) have been evaluated successfully. MVLN—based
screening and relative potency or efficacy comparison has been used to focus
analytical efforts on samples that show the greatest potential to exert estrogenic
effects, identify field sites that warrant more extensive study, and profile the
distribution of potentially estrogenic agents in relation to point sources, land use,
and wastewater treatment processes. In cases where chemical fractionation has
been coupled with MVLN screening, the results have facilitated the formulation
and testing of hypotheses regarding which specific compounds or groups of
compounds in an environmental sample which may be responsible for the in vitro
responses observed (40). MVLN responses to individual compounds have been
used to characterize the relative “estrogenic" potencies of compounds (12).
Comparison of responses elicited by a compound in several in vitro bioassays
based on different mechanistic assumptions can provide lnfonnation regarding
that compound’s mechanism of action. Thus, in vitro bioassays can provide a
great deal of useful information to aid in the assessment of endocrine disrupting

compounds.

14

As with other approaches, in vitro bioassay responses must be interpreted
with a clear understanding of their limitations. They can detect and provide an
integrated measure of the ability of a compound or sample to elicit a mechanism-
specific biological response. The relevance of that response for modeling or
predicting effects in vivo is dependent on the relevance of the mechanistic
principles and assumptions upon which the assay is based. The MVLN assay,
for example, would not detect compounds that exert estrogenic responses in vivo
by acting as an anti-androgen or by altering endogenous estradiol levels through
effects on the hypothalamus or pituitary. Thus, unless a general and fairly
universal mechanism of action for a specific type of in vivo effect can be defined,
in vitro bioassays provide a rather narrow field of view for detecting compounds
which may alter endocrine function on a whole-organism level. Furthermore,
because they use simplified systems, in vitro assays fail to account for a number
of important factors which may modulate responses via a given mechanism of
action in vivo. Such factors include toxicokinetics, cross-talk between biological
pathways, and environmental factors (12, 22). In order to be useful for predicting
effects at the whole-animal or population level, in vitro bioassays must be
rigorously calibrated to in vivo responses (44, 45). The greatest limitation of in
vitro bioassays for either screening or monitoring is the need for a priori
knowledge about the mechanisms of action of a compound or mixture for which
the assay is designed to screen. Thus, although in vitro bioassays have a role to
play in the evaluation of endocrine disrupting compounds, they are just one of the

tools needed to address the issue.

15

IN VIVO VERSUS IN VITRO TESTING FOR EDCs

Despite public pressure to use in vitro testing whenever possible to spare
animal lives and minimize the cost and time involved, whole animal studies are
crucial for the development of biologically relevant methods to screen for and
characterize EDCs in water. In vitro bioassays are mechanism specific and
cannot account for the broad spectrum of potential mechanisms of effects that
may be relevant in vivo (22). Whole animals can integrate and account for
effects of chemicals in a mixture that may act through different mechanisms, at
different tissues, either directly or indirectly to modulate overall endocrine
function as well as specific endpoints. Additionally, effects of some compounds
may be affected by metabolic transformation. Such transformations are
generally not accounted for by in vitro bioassays (22). Furthermore, whole
animal studies account for bioaccumulation and homeostatic controls that affect
the potency or efficacy of compounds in vivo. Although not necessarily
amenable for routine monitoring and screening, in vivo studies are critical for
relating exposure to population level-effects and ecologically relevant responses
as well as the development and validation of relevant in vitro screening assays

and biomarkers.

16

USE OF FISH TO TEST FOR EDCs.

Fish have long been used as biosentinel models of water quality,
particularly because of the concerns associated with the increased production,
use, and disposal of chemicals in the environment (13, 46). Among these
chemicals are the EDCs, which are thought to interfere (directly or indirectly) with
the processes governing reproduction, behavior, and other aspects used to
evaluate population health.

Many factors must be taken into account when using fish to test for effects
of EDCs. It is well known that environmental factors play an important role in fish
reproduction. Temperature, photoperiod, diet, availability of spawning
substrates, proximity and reproductive readiness of potential mates (47), and
water quality all affect reproductive status, ovulation, and spawning in fish (48).
Stress, including that from capture, handling, and confinement, can have a rapid
and marked effect on plasma sex hormone levels (49, 50). Seasonal and dailly
changes in sex hormone levels must be considered in sampling schemes (49). If
fish are fed during a study, diet must be selected carefully since some animal
diets can contain hormonally active components or contaminants (30, 51, 52).

Fish exhibit a variety of reproductive strategies. Some are ovovivlparous
and some are oviparous. Some, like the fathead minnow (Pimephales
promelas), demonstrate parental care of eggs or offspring, while others, like the
goldfish (Carassius auratus), ignore or even consume their eggs. Some spawn

only once per breeding season, while others spawn repeatedly. This last point is

17

particularly important to consider, since hormone levels and other end points can

change rapidly and dramatically around the time of spawning.

SELECTION OF TEST SPECIES

Different species of fish have been used to test for effects of EDCs, and
each has its advantages and disadvantages. Desirable species should be easy
to culture and handle in the laboratory, readily available year-round, widespread,
and economically or aesthetically important. It is also important to choose a
species for which there is a great deal of knowledge about its reproductive
biology. It is difficult to determine whether an EDC has caused a departure from
the normal condition if the normal condition is not known. Easily recognized
sexual dimorphism is important for sexing fish, and in some cases, reduction of
prominent secondary sex characteristics is used as evidence of endocrine
disruption (44, 45). Some species that are currently used include trout and
salmon, particularly rainbow trout (Oncorhynchus mykiss), carp (Cyprinus
carpio), goldfish, white sucker (Catostomus commersonr), Iargemouth bass
(Micmpterus salmoides), fathead minnow, Japanese medaka (Oryzias Iatipes),

mummichog (Fundulus heteroclitus), and mosquitofish (Gambusia affinis).

18

END POINTS AND BIOMARKERS USED TO ASSESS EFFECTS OF EDCs

Fish have been used in both laboratory and field studies to test individual
chemicals or mixtures, effluents, and natural waters impacted by human activity
for potential to alter reproductive structure and function. Some end points used
to assess the effects of EDCs on fish and some studies that employed these end
points are listed (Table 1). This table does not list all end points that could be
used or all of the studies in which they have been used. For a review of methods
to assess effects of EDCs on fish, see (53).

Plasma vitellogenin (Vtg) induction is a popular marker of exposure of fish
to chemicals that bind to the estrogen receptor. Vtg is an egg-yolk precursor
synthesized in the liver of oviparous animals in response to estrogen (54). Vtg is
released from the liver to the bloodstream where it is carried to the ovary and
sequestered into developing oocytes. Vtg usually is not detected in significant
amounts in the plasma of male or immature fish, but they can be stimulated by
exposure to natural or synthetic estrogenic chemicals to produce Vtg (55). Thus,
Vtg is a useful tool to detect estrogenic activity of test chemicals or environmental
waters potentially impacted by estrogenic EDCs. However, it is important to
keep in mind that Vtg induction has not been shown to have any adverse effects
in fish and is a functional measure of exposure useful only for monitoring for

estrogenic compounds (55).

19

FIELD STUDIES

Other researchers in the United States, the United Kingdom, and Canada
have conducted field studies to determine whether EDCs in the environment
have the ability to alter the reproductive structure and function of feral or wild (44,
56—60) and caged fish (61-63). Representative examples of these studies are
those conducted with adult fish at the Aquatic Toxicology Laboratory at Michigan
State University.

In situ exposure of caged fathead minnows to municipal wastewater
treatment plant (VVWT P) effluents was conducted. Fathead minnows were caged
in rivers directly in the outfalls of municipal WWI'Ps in mid-Michigan (64).
Plasma Vtg, E2, testosterone (T), gonad histology, and male secondary sex
characteristics were examined. Adult male and female minnows were caged for
3 weeks. No changes in any of these end points could be attributed to EDCs in
WWTP effluents.

In situ exposure of caged common goldfish to municipal waste water
treatment plant effluents was conducted. This study was similar to the fathead
minnow study previously described. Adult male and female goldfish were caged
for 6 weeks at the same sites with the fathead minnows (65). Plasma Vtg, E2,
and T were measured by enzyme immunoassay. Gonad histology was
examined, and gonad tissue was removed and incubated in vitro to evaluate

gonadal steroidogenic activity. Interpretation of the data is in progress.

20

 

 

LABORATORY STUDIES

The Aquatic Toxicology Laboratory at Michigan State University has
conducted both field and controlled laboratory studies. All of the laboratory
studies were conducted with adult fathead minnows exposed to test compounds
in a proportional flow-through diluter system.

Fathead minnows were exposed for 19 days to water borne E2 at
measured concentrations ranging from 3.5 to 15 ng/L in a proportional flow-
through diluter (66). Fish were exposed in duplicate groups of six, with three
males and three females per exposure tank. Interestingly, E2 was detected even
in control and solvent control tanks, indicating that the fish were a source for
some of the estrogen in the exposure tanks. Egg production, plasma Vtg, plasma
E2, and gonad histology were examined. The E050 (concentration expected to
cause 50% effect) for inhibition of egg production was 120 nglL. and the E050 for
Vtg induction in males was 251 ng/L. Vtg was measured indirectly as alkaline
labile phosphorus. In male fish, plasma E2 increased in a dose-dependent
manner with concentrations of water borne E2. Plasma E2 levels in females
were not affected by water borne E2 exposure. This difference in response may
be due to sex-related differences in homeostatic control or metabolism of
estrogen. Histopathological changes of the gonad were noted in a companion
study (45) (see next section).

The fathead minnow was used as a test subject in part because of its
prominent sexual dimorphism. Males have a fatpad (for which the fish is named)

on the dorsal surface of the head. Males also develop nuptial tubercles, rows of

21

 

small, hardened projections on the rostrum. The fatpad and nuptial tubercles are
easily seen with the naked eye, and are absent in females. The female is
generally lighter in color and smaller than the male, which usually develops a
darker color pattern when sexually mature. In addition, the female has a more
tapered head shape.

Sexually mature fathead minnows were exposed to water borne E2 as
described in the previous section (66). In males, E2 exposure caused atrophy of
secondary sex characteristics and histologic lesions of the testes. Lesions of the
testes included proliferation of Sertoli cells, degenerative spermatozoa, and
enlarged phagolysosomes containing necrotic spermatozoa and other cellular
debris. Ovaries of females exposed to E2 contained more primary follicles and
fewer secondary and Graafian follicles than ovaries of control fish (45), indicating
that ovarian development in preparation for spawning had been retarded.

Adult fathead minnows were exposed to water borne 4-nonylphenol (NP)
in a proportional flow-through diluter at concentrations ranging from <0.01 to 3.4
[lg/I. for 42 days (67). The study was conducted first early in the natural breeding
season (Experiment I, or EX l) and again later in the breeding season
(Experiment II, or EX ll). Measured exposure concentrations for the two
experiments were similar (EX I: 0.05, 0.16, 0.4, 1.1, and 3.4 pg NPIL ; EX II:
<0.01, 0.10, 0.33, 0.93, and 2.4 pg NPIL ). Examination of dose-response
relationships between NP and egg production per female and between NP and
plasma E2 in females revealed curves indicative of "Inverted-U" type responses,

with lesser doses resulting in an increased response and greater doses resulting

22

in lesser responses. For males and females in EX l, plasma E2 levels were
greater than controls for all of the NP treatment groups except the greatest (3.4
pg NPIL), which was not different from the controls. Plasma E2 levels for both
sexes ranged roughly from 2 to 25 ng/mL. For females in EX ll, two intermediate
exposure concentrations (0.1 and 0.33 pg NP/L) produced plasma E2
concentrations statistically greater than those in both controls groups. The
plasma E2 concentrations for females ranged from about 2 to 8 ng/mL. In EX II
males, none of the treatment levels produced plasma E2 levels statistically
greater than both controls, but the sample sizes were small, and the shape of the
curve indicated a trend similar to that seen in the females. For females in EX l,
the dose-response curve for NP versus plasma Vtg suggests that NP caused a
decrease in plasma Vtg levels, with the two greatest NP concentrations resulting
in the least plasma Vtg levels. For females in EX II, the dose-response curve for
NP versus plasma Vtg suggests an Inverted-U response, with the least NP
exposure level producing a statistically significant increase over the controls, and
greater exposure levels producing statistically lesser Vtg levels that are still
significantly greater than the control Vtg levels. For males in both EX | and EX II,
there was no significant increase in plasma Vtg in any of the NP exposure
concentrations. In EX I, NP caused no statistically significant effect on egg
production per female. However, the sample sizes used were small, and the
shape of the dose-response curve suggests the same inverted-U type response,
with increased egg production at the least treatment level (0.05 pg NPIL) and a

decrease in greater treatment levels. In EX ll, there was an increase in egg

23

production per female for the greatest treatment level (2.4 pg NPIL) and for one
of the intermediate treatment levels (0.01 pg NPIL), with two treatment levels
between exhibiting no difference from the controls.

The mechanism of NP action in the alteration of fathead minnow
reproductive end points is unclear, but may be related to positive and negative
feedback on endogenous estrogen production. The differences in response over
the two different experiments may have resulted from differences in the
reproductive state of the fish. This underscores the importance of timing of the
study relative to the reproductive state in fish. Furthermore, while NP has been
reported to be a weak estrogen agonist, it is likely that the mechanism of action is
not due to the binding of NP to the ER.

Adult fathead minnows were exposed to an industrial mixture of NPEO for
42 days in a proportional flow-through diluter. Three males and three females
were placed in each exposure tank. NPEO exposure concentrations ranged from
0 to 10 pg/L. Plasma Vtg, E2, and T were measured by enzyme immunoassay.
Egg production per female (fecundity) was determined. No statistically significant
concentration-dependent relationship was demonstrated between fecundity and
NPEO exposure. However, the shape of the curve indicated a possible inverted-
U response. NPEO was not observed to have any statistically significant effect
on any of the other end points examined.

The mechanisms playing a role in the disruption of endocrine, and related
systems, in fish need a better understanding (68). Considering that testing of

EDCs in feral populations, or even representative captive adults in laboratory

24

studies represent an economical, practical, and even technological obstacle,
attention has been shifted to possible alternatives, not just of surrogate species,
but also of exposure methodologies. The medaka (Oryzias Iatipes), a small fish
frequently used as adults and/or early life stages (embryos and/or larvae) in
aquatic toxicity studies (69-71) has proven useful as a tool in those alternative
studies. The attributes that make this species attractive for in vivo
experimentation have been discussed elsewhere (46); however, only until
recently has micromanipulation technology started to take advantage of a more
novel exposure approach: in ovo nanoinjection of chemicals into fertilized
medaka eggs (72, 73).

While egg microinjecting techniques have been developed in several fish
(mainly salmonids) over ten years ago (74-76), few studies have been attempted
in species with small eggs. Recently, Villalobos et al. (73) reported success in
using this medaka assay to test the toxicity of several technical mixtures of
polychlorinated napthalenes (Halowax). The authors injected 0.5 nl of Halowax
1014, Halowax 1013, or Halowax 1051 in triolein as a carrier solvent. Mortality
for the injected triolein controls was below 10%, and the Halowax-induced
responses suggested involvement of the arylhydrocarbon- (Ah-) receptor. The
morphological effects of each of the Halowaxes was easily observed in the
embryos and could be associated with structure-activity dependent effects
(responses varied according to the chloronaphthalene speciation, chlorine
localization within the molecule, and total mass contribution). Furthermore, the

effects were consistently reproduced after repeated tests. However, these

25

compounds did not appear to have elicited a disrupting effect in the medaka
endocrine system. These studies, done in collaboration with the US. Geological
Survey (USGS) Columbia Environmental Research Center (Columbia, MO),
demonstrate the feasibility of nanoinjection of compounds into the small eggs of

medaka.

MONITORING OF BUTYLTIN COMPOUNDS IN WATER

Recently, widespread industrial and agricultural applications of organotin
compounds, particularly butyltins, have led to increasing concerns about
environmental contamination and biological effects caused by these compounds.
Di- (DBT) and mono-butyltins (MBT) have been widely used as heat and light
stabilizers for polyvinyl chloride (PVC) plastics and catalysts in the production of
polyurethane and silicone. TBT has been used as antifouling agents, biocides,
fungicides, molluscicides, insecticides and wood preservatives (77). The use of
butyltin compounds in a variety of consumer products and household items
(including sanitary napkins, diaper, cellophane wrap, sponges used in washing
dishes, butter parchments, and gloves) (78, 79) and the leachates from PVC
pipes have led to their occurrence in municipal wastewater and sewage sludge
(80, 81). In addition, TBT is used as a disinfectant in waxes, polishes, sprays
and laundry washes, which may allow contamination of municipal wastewater

(82). Industrial discharges of TBT used as a slimicide in the paper industry and

26

for textile and lumber treatment, and in cooling water treatment are further
sources of contamination.

Input of DBT and TBT into surface waters from applications other than
antifouling paint on vessels (i.e., use of TBT as a slimicide in the cooling water of
a thermoelectric power plant in Italy) was first reported in the late 19805 (83).
Concentrations of butyltins in raw wastewater in Zurich, Switzerland, on six
sampling days ranged from 136 to 564, 127 to 1026 and 64 to 217 ngIL,
respectively (84). Approximately 90% of each butyltin species was associated
with suspended particles, and butyltins are removed at a rate of 62, 80 and 66%
for MBT, DBT and TBT, respectively, by sedimentation (clarifier) in the sewage
treatment plant. After secondary clarification, which usually represents the
outflow from treatment plants, MBT, DBT and TBT ranged from 7 to 47 ng/L.
MBT, DBT and TBT concentrations in the range of 01-097, 0.41-1.24 and 0.28-
1.51 png (dry wt) were found in sewage sludge. Similarly, MBT was found in all
the 36 influent and effluent samples from VWVTPs in five Canadian cities.

Concentrations of MBT were in the range of 28-31 pglL for influent and 1-22

ngL for effluent. DBT and TBT were found only infrequently at a few pglL (85).
Concentrations of MBT, DBT and TBT in sewage sludge were as great as 650,
600, and 675 pg/g, dry wt, respectively (85). In Bordeaux, France,
concentrations of MBT and DBT in influents from a WWTP were 18 and 12 ng/L,
respectively (86), whereas those in effluent samples were 15 and 8 ng/L,
respectively. TBT was not detected in these samples. Effluents from a WWTP in

Mainz, Germany, contained MBT and DBT concentrations in the range of 13-28

27

and 6-96 ng/L, respectively (87). Total butyltin concentration in sewage sludge
from Patna, India, was 340 ng/g, dry wt (88).

Studies describing the occurrence of butyltins in wastewater
effluents in the US. are not available. Nevertheless, results from the above
studies suggest that butyltins are widespread contaminants in wastewater. Even
in those WWI'Ps with exceptionally modern systems (84), the concentrations of
TBT remaining in effluents were greater than the environmental quality standard
of 2 ngIL (89). Occurrence of butyltin compounds in unnavigable areas in certain
rivers (90), as well as in aquatic organisms of certain unnavigable rivers, suggest

the release of these compounds from wastewater (91).

ECOLOGICAL IMPACTS OF BUTYLTINS

Butyltin compounds, particularly TBT has been implicated in the
development of “imposex” (development of male sex characteristics in females)
in about 50 species of gastropods worldwide (92). This effect was attributed to
inhibition by TBT of cytochrome P450-dependent aromatase, the enzyme
responsible for the conversion of testosterone to 178-estradiol (17). Because
cytochrome P450 systems control the conversion of cholesterol into a variety of
hormones including testosterone and estradiol, effects of TBT on this enzyme
system can result in hormonal imbalance in animals. Although the
concentrations of TBT in wastewater effluents were greater than those known to

cause imposex in several sensitive gastropods, discharge of effluents into

28

surface waters could result in dilution. Nevertheless, aquatic animals
concentrate TBT in tissues, particularly in liver or hepatopancreas (20, 93, 94).
Because lipovitellin (provides protein and lipid for the developing embryo) is
produced in the hepatopancreas or liver, accumulation of butyltins in these
organs can interfere with vitellogenesis (95). TBT can also interact with
cytochrome P450 and modulate the toxic effects of contaminants such as
polychlorinated biphenyls (96). Although MBT and DBT were not shown to
induce imposex in gastropods, they have been reported to be toxic to rainbow
trout yolk sac fry (97). No observed effect concentrations (NOEC) for DBT and
TBT in water were 40 ng/mL and 40 ngmL, respectively, for the mortality of
rainbow trout yolk sac fry (97). Similarly, DBT has been reported to produce a

wide variety of toxic effects such as immunosuppression in exposed fish (97, 98).

INSTRUMENTAL ANALYSIS AND BIOMONITORING OF BUTYLTINS

The need to identify and determine concentrations of different butyltin
species at trace levels was driven by their great toxicity. There are now a variety
of instrumental methods available for the routine determination of butyltins in
different matrices, but it should be noted that these analytical techniques are still
somewhat laborious. The modern techniques are generally sensitive enough to
measure butyltin species at sub nglL levels, but quality assurance and quality
control efforts should be implemented on a regular basis to assess the accuracy

and precision of analytical results. Some of the recent instrumental analytical

29

 

techniques for butyltins are discussed elsewhere (18). The most sensitive
methods for analysis of butyltins involve the conversion to either alkyl derivatives
or volatile hydrides followed by chromatographic separation and determination
with specific detectors. The gas chromatography (GC) techniques used in
butyltin analyses include: GC-MS (mass spectrometery), GC-GFAA (graphite
furnace atomic spectrometry), and GC-FPD (flame photometric detector).
Methods based on G0 are widely used due to its high resolution and detector
versatility.

Due to the potential for TBT to Induce imposex in gastropods at
water concentrations of a few ng/L, studies have suggested the application of
field bioassays using transplanted bivalves to assess TBT contamination in
aquatic systems (17, 92). The intensity of expression of imposex in Nucella
IapiIIus can generally be related to the water concentration of TBT. The
appearance of a small penis and the partial development of a vas deferens first
occurs at TBT concentrations below 0.5 ng/L (as tin), although reproduction is
unaffected at this level (89). At 1-2 nglL (as tin) penis size is markedly increased
and in some females proliferation of vas deferens tissue overgrows the genital
papilla, thus sterilizing the animal. At slightly greater concentrations all females
become sterile and a concentration of 10 ng tin/L caused suppression of
oogenesis and initiated sperrnatogenesis (89). For monitoring purposes, the
intensity of imposex is most simply described by the relative penis size (RPS)
index in which the bulk of the female penis [(female length)3/(male length)3 x 100]

is expressed as a percentage of that of the male. Measurement of the length of

30

 

a penis in species such as Nugefla is a simple procedure since the penis can be
expressed without dissection by placing the animal (minus shell) dorsal side up
and drawing back the flap of tissue forming the rook of the mantle cavity. Under
a binocular microscope and using 1 mm graduated graph paper, the length of the
penis can be measured to the nearest 0.1 mm from its tip to its base. Bifurcate
or trifurcate penises are occasionally found, particularly on females, and in these
cases the longer or the longest structure is measured. A female lacking any
measurable penile outgrowth is registered as zero and this value is included in
the calculation of the mean. Whilst length is the most convenient parameter of
penis size to measure, it does not convey a true impression of the difference in
mass between, for example, a penis of 1 mm and another of 3 mm. However,
the weight of the penis is related to the cube of its length (89) and thus the RPS
calculation involves the cube of the length. An RPS of 50% indicates that the
mean penis size of the female is half the bulk of that of the male. RPS is a
reliable measure in all areas except those where contamination is severe. In
these areas the male penis is often deformed and consequently the RPS value is
lowered and imposex thereby underestimated. Because RPS does not positively
identify the presence of sterilized females and thus a second index, the vas
deferens sequence (VDS) was devised (89). This index is based on the division
of imposex into six stages ranging from early development to the final stage of
sterilization. Calculation of the mean VDS stage provides an index by which to
compare imposex in different populations. Any population with an index above 4

contains sterilized females and thus has a reduced reproductive capacity.

31

Further details on this biomonitoring technique are given in Bryan et al., 1988. A
positive relationship between the incidence of imposex and body residues of TBT
has been shown (99). Although the induction of imposex tends to be regarded
as a specific response to TBT, field bioassays can be confounded by several
environmental factors and, therefore, their applicability to quantitatively monitor
butyltin contamination is in question. In vitro bioassays have been used to study
the effects of butyltin compounds on EROD activity. Due to high cytotoxic
potential of butyltin compounds, use of in vitro studies to elucidate the
mechanism of toxic effects of butyltin compounds has been hindered. This
indicates the need for a combination of several techniques to identify the sources

and effects of butyltins in the environment.

BIOASSAY-DIRECTED CHEMICAL FRACTIONATION

Unique challenges face the analytical chemist when trying to develop
methods for EDCs. The primary difficulty is determining what compounds are
EDCs. While various groups and agencies have published lists of EDCs, the
question of new or unknown EDCs in the environment remains. By the use of in
vitro bioassays like those previously discussed as an analytical tool, the chemist
may begin to develop a more comprehensive screening program. When a
bioactive sample is discovered, subsequent chemical fractionation and in vitro
bioassays may lead to compounds or classes of compounds likely responsible

for the observed effect. Once candidate compounds have been identified, a

32

predicted activity can be estimated by applying relative potency factors derived
from pure compounds. This predicted activity may be compared to the actual
activity measured in the bioassay to determine if a mass-balance exists.

A reliable method for sensitive detection of certain EDCs in complex
aqueous mixtures has been developed using solid-phase extraction (SPE) and in
vitro bioassays (15). Dissolved and particulate phase organics were
simultaneously extracted from 5 L water samples at the field site by pumping the
samples sequentially through a 90 mm glass fiber filter and a 90 mm Empore
styrenedivinylbenzene (SDB) SPE disk. The SDB matrix permits the extraction
of both non-polar and semi-polar organics from water. Flow rates and volumes
were measured with an in-Iine electronic flow meter/totalizer. Vacuum filtration
was used at the laboratory to extract the SPE disks sequentially with acetone,
dichloromethane and hexane. All extracts were dried over sodium sulfate and
concentrated to near dryness. HPLC with a preparative silica column was used
to generate three fractions of the extract based on polarity. The least polar
fraction contained halogenated aromatic hydrocarbons (HAHs) and polycyclic
aromatic hydrocarbons (PAHs), the moderately polar fraction contained
alkylphenols and phthalates, and the fraction of greatest polarity contained
steroids, bisphenol A, and alkylphenol polyethoxylate surfactants. Both total and
fractionated extracts were assayed for estrogenic/antiestrogenic (ERE-mediated)
activity or dioxin-like (AhR-mediated) activity using cell lines transfected with
Iuciferase reporter genes under control of estrogen- or dioxin- responsive

enhancer sequences, respectively. Additional fractionation by reverse phase

33

HPLC and in vitro testing were sometimes used to more narrowly isolate
bioactive compounds. Compounds identified using this strategy included:
alkyphenols, steroids, PAHs, and HAHs. Instrumental analyses performed
included: HPLC/fluorescence, GCIMS and GCIECD. ELISA was used to quantify
synthetic and endogenous estrogen in some bioactive extracts. Sites screened
included several wastewater treatment plants in south central Michigan, effluents
entering the Trenton Channel of the Detroit River, and areas of Lake Mead in
Nevada.

The degree of induction over background was used as a measure of the
activity of unknown samples to direct further fractionation and instrumental
analyses. Most AhR-mediated activity occurred in the most non-polar fraction,
while the greatest ERE-mediated activity occurred in the most polar of the three
fractions. These polar fractions were further separated by reverse-phase HPLC
into 9 fractions. In every case, the estrogenic activity occurred in the fractions
containing E2 and EE2. These compounds were determined to be the most
likely causes of observed in vitro estrogenicity. Further confirmation was
completed using radioimmunoassays (RIAs) for E2 and EE2. Water
concentrations of E2 and EE2 ranged from less than detection to 3.7 ng/L and
0.8 nglL, respectively.

AhR-active compounds were detected by bioassay in only one sample,
indicating that the levels of “dioxin-like” compounds were less than the method
detection limits for most of the water samples. This is not suprising as only 5 L of

water was extracted.

34

 

Although NP and OP, known environmental estrogens, were present in
many water samples, there was no correlation between these alkylphenols and
observed estrogenicity. Not only did the fraction containing these compounds
show no bioactivity, the mass balance also would not predict a significant
response based on the concentrations in the resulting extracts. Fine
fractionation also confirmed that the lack of estrogenicity in the alkylphenol
fraction was not due to antagonistic effects. The concentrations of NP and OP
ranged from less than detection to 37 and 0.7 pglL, respectively.

The in vitro bioassay systems were found to be effective tools for the
screening of estrogen and dioxin-like activity in extracts of surface waters and for
directing further fractionation and instrumental analyses. The results of the
assay can also be compared to the predicted activity calculated from an additive
model of estrogenic activity. In the second method, the concentrations of all of
the putative estrogenic compounds in a mixture are multiplied by their relative
potency factors (determined for individual compounds) and the total activity is
predicted by summing the products of these two values. This method allows a
mass balance of activity to be calculated. By comparing the response of the cells
with whole extracts and various fractions, it is possible to determine whether all
of the likely estrogenic compounds have been identified and to investigate

nonadditive interactions among compounds.

35

 

CONLUSIONS

Instrumental and bioanalytical methods for the determination and
measurement of EDCs in the aquatic environment have been presented. There
are many tools available to aid in the identification and characterization of EDCs.
In vitro bioassays have great potential for routine use in characterizing EDCs.
They become particularly useful tools when used in conjunction with instrumental
analyses. They are, however, only relevant as the mechanisms the are based
on. Calibration to in vivo responses are crucial for proper application and
interpretation of in vitro bioassay results. In vivo bioassays offer an integrated
measure of endocrine disruption as the whole animal is taken into account. In
vivo testing suffers from the complexity of sorting out mechanisms of toxicity.
Analytical chemistry techniques are important in the identification and
quantitation of EDCs in the environment and in controlled laboratory studies.

The issue of endocrine disruption is extremely complex. Only by
combining various investigative techniques can the risks, if any, of EDCs be
better understood. EDCs have a wide variety of toxicity mechanisms, while also
having a great range of physical properties. An integrated approach is required

to better detect, quantify, and understand EDCs in the environment.

36

TABLE 1: End points used to assess reproductive impacts of EDCs in Fish

 

End point

vitellogenin (alkaline labile phosphorus,
immunoassay, liver mRNA)

gonadosomatic index (GSI)

plasma sex hormone levels (T, 11-KT, E2 and other
estrogens)

steroidogenesis enzyme activities and intermediates

production of hormones during in vitro organ
incubation (gonad, pituitary)

gonadotropins and gonadotropin releasing hormone

gonad structure and histology (intersex, gonad
duct development)

activity of liver enzymes involved in steroid
metabolism (mixed function oxygenase, or MFO,
activity)

fertility and fecundity

secondary sex characteristics

reproductive behavior

37

Selected references

(56, 62)

(100-103)

(104,105)

(105)

(103,105)

(103, 104)

(107-109)

(57,110)

(57,102)

(44, 45)

(44,111)

(1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

(11)

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48

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:04

l
‘-—---0’

   

ERE-Luc . ,
\ gene \LUleel‘ase III
\ \ m NA ’I’I

’ ’
‘

  
      
 

 

- o luiferin . . '
xy lucrferrn

Luciferase

Figure 1. Mechanism for activation of the estrogen receptor (ER) in
MVLN cells

49

Chapter 2

BIOCONCENTRATION OF NONYLPHENOL IN FATHEAD MINNOWS
(PIMEPHALES PROMELAS)

(Submitted to Chemosphere 2000)

ABSTRACT

Bioaccumulation of p-nonylphenol (NP) by fathead minnows was
determined under laboratory conditions. Fish were exposed continuously for 5
wk to 3.4, 1.6, 0.4, 0.16 and 0.05 pg NPIL in a flow-through system. NP was
Soxhlet extracted from whole fish homogenates with dichloromethane (DCM).
The resulting extract was concentrated and bulk lipids removed by gel
permeation and silica gel chromatography. Compounds were identified and
quantified by reverse-phase high-pressure liquid chromatography (RP-HPLC)
with fluorescence detection. Mass spectrometry was used for verification of peak

assignments. Bioconcentration factors ranged from 203 to 268.

INTRODUCTION

Alkylphenol polyethoxylates (APEs) are non-ionic surfactants that are
used in various industrial, institutional, agricultural, and household applications
(T almage, 1994). APEs are among the most widely used non-ionic surfactants,

with approximately 250 million kg per year used in the United States (US)

50

(Chemical Market Reporter, Oct. 18, 1999). The molecular structure of APEs
consist of two moieties, an alkyl-substituted phenol and an ethoxylate oligomer
chain. The alkylphenol (AP) portion of the surfactant is fairly non-polar and
allows the APEs to dissolve grease and other materials that have small water
solubilities. The ethoxylate portion of the surfactant is water-soluble and aids in
the transfer of materials to the aqueous phase. APEs are produced
commercially by the base-catalyzed ethoxylation of alkylphenols (Cahn & Lynn
Jr., 1978). Greater than 90% of the APs used in APE production are of the para
substituted isomer. Nonylphenol polyethoxylates (NPEs) that contain a 9 carbon
branched isomeric alkyl group (CgI‘I1g) (Figure 1) represent approximately 80% of
production (Lye et al., 1999; Naylor et al., 1992; Talmage, 1994). The remaining
production is comprised almost entirely of octylphenol polyethoxylates (OPEs).
NPEs and the less widely used OPEs degrade during wastewater
treatment or in the environment to form transient intermediates such as
alkylphenol ethoxycarboxylates (APECs), shorter-chain oligomers of APEs, and
alkylphenols (Bennie et al., 1997; Field & Reed, 1996; Naylor et al., 1992; Rudel
et al., 1998; Tanghe, Dhooge & Verstraete, 1999). The type and efficiency of
wastewater treatment affects the total removal rate of NPEs and the type and
proportion of the transient intermediates. Primary treatment removes only 25 -
78% of the total NPE entering wastewater treatment plants (WWTPs). With
primary treatment alone, the hydraulic retention time is short and degradation is
minimal. Primary effluents contain approximately 82% NPE3 - 20, 12% NPEm,

3% nonylphenol monoethoxycarboxylate and nonylphenol diethoxycarboxylate

51

(NPECm), and 3% nonylphenol (NP) from the initial NPE loading (Ahel, Giger &
Koch, 1994). In the United States (US), WWTPs with secondary or tertiary
treatment achieve average removal rates in excess of 97% for NPE. Effluents
from these types of WWTPs contain approximately 28% NPE3 .. 20, 22% NPEm,
46% NPECm, and 4% NP from the initial NPE loading (Ahel et al., 1994).

Although NP is a minor degradation component of NPE, significant
amounts of this slightly lipophilic, moderately toxic compound enter the
environment. NP has a log K0W of 4.84 (Ahel & Giger, 1993b), water solubility of
5.4 mg/L (Ahel & Giger, 1993a), and a log Koc of 3.58 (Naylor et al., 1992). This
physico-chemical profile indicates that NP may bioaccumulate in aquatic
organisms. This has been documented in several species of fish from natural
waters and from controlled laboratory exposures (T almage, 1994). NP
bioconcentration factors (BCFs) for the fathead minnow (Pimephales promelas)
exposed to 5 and 25 pg NP/L for 20-d was determined to be 270 and 350,
respectively (Naylor, 1995). Half-times for depuration have been determined to
be 1.4 and 1.2 (1, respectively, following exposure to 5 and 25 pg NPIL (Naylor,
1995).

Nonylphenol has been shown to be weakly estrogenic in a variety of both
in vitro (Desbrow et al., 1998; Jobling & Sumpter, 1993; Mueller & Kim, 1978;
Soto et al., 1992; White et al., 1994) and in vivo bioassays (Giesy et al., 2000;
Jobling et al., 1996; Nimrod & Benson, 1996). L050 values for acute toxicity to
fish range from 0.135 to 0.3 mg NPIL for fathead minnows (Pimephales

promelas) (Dwyer et al., 1995) and from 0.167 to 0.221 mg NPIL for three

52

species of trout (Naylor, 1995). Reports of APEs and their degradation products
in the environment in the US. are scarce. The most comprehensive survey
reports concentrations from 30 US. rivers that are influenced by municipal or
industrial wastewater effluents (Naylor et al., 1992). That study found that 60-
75% of samples had no detectable levels of NP, NPE1, and NPEZ. NP
concentrations observed in that study ranged from <0.11 to 0.64 pglL (Naylor et
al., 1992). Concentrations of NP in effluents and surface waters ranging from not
detectable to 37 pglL have also been reported (Snyder et al., 1999).

The objectives of this study were to develop a suitable method for
extraction, clean-up, and quantification of NP in fish tissues and to use this
method to determine the BCF for partitioning of NP into fathead minnows during

a controlled laboratory exposure.

EXPERIMENTAL SECTION

Standards and Reagents: All standards and reagents used were of the highest
purity commercially available. High purity standards of p-nonylphenol (NP), p-tert
octylphenol (OP), and p-tert butylphenol (BP) were obtained from Schenectady
International (Freeport, TX). Pesticide residue grade acetonitrile (ACN), hexane,
dichloromethane (DCM), and acetone were obtained from Burdick and Jackson
(Muskegon, MI). Reagent water was first purified by reverse osmosis (RO)
followed by NanopureTM (Barnstead, Dubuque, IO) treatment. Glass wool (8 Elm,

Corning Glass Works, Corning, NY) was Soxhlet extracted overnight with DCM,

53

then dried and stored in glass containers until needed. Glass fiber filters, fine
grade (GF/F) of 0.7 pm nominal pore size were heated at 450 °C for 4 h in
aluminum foil prior to use. Anhydrous granular sodium sulfate (EM Science,
Gibbstown, NJ) was heated at 450 °C overnight then stored at 125 °C until day of

use.

Fish Exposure. Fathead minnows (Pimephales promelas) were cultured in the
Michigan State University Aquatic Toxicology Laboratory (MSU-ATL) under a
16:8 lightzdark cycle in continuously flowing, well water at 17-19 °C. Fish were
fed a 1:1 (v:v) mixture of Tetramin® (Tetrawerke, Melle Germany) and ground
Sterling Silver Cup (2-4 mm pellet size) (Murray, UT) once daily at a rate of
approximately 0.5% of fresh body weight. The dry food was supplemented with
Artemia (brine shrimp) nauplii. Freeze-dried Spirulina sp algae was fed to fry up
to 2.5 cm length. Reproductively mature fish, 12 to 18 mo of age, were
transferred to randomly assigned 19 L glass aquaria in a circulating water-
jacketed bath (Frigid Units Living Stream, Toledo, OH). Each tank received 3 Uh
treated well water through a solenoid-controlled proportional-flow diluter
apparatus (Ace Glass, Vineland, NJ). Each tank received vigorous aeration, and
glass lids prevented escape of aerosols from the tanks. The temperature of the
water jacket was maintained at 26 °C.

Fish were randomly assigned to 20 groups, each containing 2
males and 2 females and placed into 19L aquaria. Treatments were in triplicate

with the exception of the solvent control group, which due to the restriction of

54

space in the water bath, contained only two aquaria. Fish were allowed to
acclimate in control water for 7 d then exposed to NP for a total of 42 d. The
aquaria were randomly assigned to receive treatment concentrations of NP or
control solutions delivered by a proportional-flow diluter. Treatments included
control, which received only water, and a solvent control, which received the
same concentration of solvent (0.0001% ethanol) as the NP treatments. Fathead
minnows were exposed to target (nominal) concentrations of 0.1, 0.3, 1.0, 3.0

and 10 pg NPIL. The actual, measured concentrations were 0.05, 0.16, 0.4, 1.1

and 3.4 pg NPIL (Table 2). Three fish from each exposure level were analyzed

for NP.

Water Extraction. Water samples were collected 3 times during the exposure to
determine actual NP concentrations. Analytical methods were adapted from
those published previously (Snyder et al., 1999). One L samples were spiked
with 1.0 pg BP in methanol as a surrogate standard. The fortified samples were
vacuum filtered and extracted using a 47 mm glass fiber filters (GF/F) directly
above a 47 mm styrenedivinylbenzene (SDB) EmporeTM solid-phase extraction
(SPE) disk. The SPE disk was frozen until extraction. The SPE disk was solvent
extracted with 15 mL of acetone followed by 25 mL DCM. Moisture was removed
by passing the extract through 30 g of anhydrous sodium sulfate. The dried
extract was concentrated to 1 mL in iso-octane. A 2 g gravimetric silica column
(60 - 100 mesh, Aldrich Chemical Company, Milwaukee, WI) column was

washed with 100 mL of 20% DCM in hexane and the extract loaded.

55

Alkylphenols were eluted with 100 mL of DCM. This fraction was concentrated to

1 mL in acetonitrile (ACN) for RP-HPLC analysis.

Fish Extraction and Clean-up. Individual, whole fathead minnows were
weighed then homogenized with 30 g of anhydrous sodium sulfate. The
homogenate was spiked with 0.50 pg of OP as a surrogate standard and Soxhlet
extracted for 12 h using 300 mL DCM. The resulting extract was concentrated at
30 °C by rotary evaporation followed by a gentle stream of nitrogen to a final
volume of 2 mL.

Gel permeation chromatography (GPC) was used to remove bulk
lipids from the extracts. The GPC system consisted of a Rheodyne 7725i injector
(Cotati, CA) with a 5 mL sample loop, a quaternary high-pressure liquid
chromatography (HPLC) pump (Perkin Elmer, Series 410, Norwalk, CT), and an
electronic fraction collector (ISCO Foxy 200, Lincoln, NE). Two Phenomenex
(Torrance, CA) GPC columns were connected in tandem; a PhenogelTM 5 pm
500 A (300 x 21.20 mm) followed by an Envirosep-ABC (350 x 21 .20 mm). Both
GPC columns were preceded by an Envirosep-ABC (60 x 21.20 mm) guard
column (Phenomenex, Torrance, CA). The pump was operated isocratically at 3
mL/min with DCM as the mobile-phase solvent. A fraction containing OP and NP
was collected from 20-35 min. This fraction was concentrated and cleaned-up in

the same manner as the water extracts.

56

Quantitation of Compounds. Extracts were analyzed by RP-HPLC as
described previously (Snyder et al., 1999). The reverse-phase HPLC system
included a Perkin Elmer (PE) (Norwalk, CT) series 200 autosampler, a PE series
200 binary pump, a Hewlett Packard (HP) 1046A fluorescence detector, and PE
TurboChrome 4.0 data software package. A Phenomenex ProdigyTM 5 pm
octadecylsilica (ODS) 100 A (250 mm x 4.6 mm, Torrance, CA) preceded by a 30
mm x 4.6 mm guard column of the same packing material was used. For all
compounds of interest, the fluorescence detector settings were: 229 nm
excitation, 310 nm emission, PMT gain of 12, lamp time of -1, response time of 2
sec, stop time at 27 min, gate and delay at zero.

The elution profile was a 20 min gradient (curve = -2) from 50%
water in ACN to 2% water in ACN followed by a 10 min ACN purge. Each
sequence began with one blank and one standard. The injection volume of
standards and samples was 25 pL. Peak areas were determined by electronic
integration, and sample concentrations were determined using Microsoft Excel
version 7.0. Chromatography and retention times for the compounds of interest
are given (Figure 2). All calibration curves were linear (r‘2 > 0.99). An HP 5890
series ll plus GC with a 30 m DB5-MS capillary column (J&W Scientific, Folsom,
CA) and a HP 5972 series mass selective detector (MSD) were used for

confirmation of peak assignments.

Recovery and Precision. Instrumental detection limits (lDLs) and limits of

quantitation (LOQs) were determined by analyzing dilute standards (near the

57

estimated IDL) and calculating the signal-to-noise ratio [peak height (mV) divided
by baseline noise] for each standard concentration. Linear regression of the
signal to noise ratios against concentrations was then used to determine the IDL
(signal to noise ratio = 3) and LOQ (signal to noise ratio = 10). The method
detection limit (MDL) was determined by spiking seven 1.5 g homogenized
fathead minnow tissue samples with dilute concentrations of NP and OP (at the
estimated MDL) and analyzing as described previously. The MDL was
calculated by multiplying the standard deviation (SD) of the resulting

concentrations by a t—value of 3.1 (for n=7).

RESULTS

Recoveries for each of the compounds of interest were in excess of 70%.

For both water and tissue analyses, data were adjusted for surrogate recovery.

The lDLs, LOQs, and MDLs for these compounds in water and tissue are shown

(Table 1). The clean-up steps for this method did not remove all lipids present in

the fish tissue extracts, resulting in an increased baseline and elevated operating

pressures. Therefore, after approximately 5 injections of fish tissue extracts, the

analytical column was purged with successively less polar solvents to remove
residual lipids.

With the exception of the controls, NP was detected in the water in

each of the exposure tanks (Table 2). For each exposure level, measured

concentrations were approximately 1/3 of nominal concentrations.

58

Concentrations were fairly constant throughout the exposure period.
Nonylphenol was not detectable in fish tissues from the lowest (measured)
exposure concentrations, 0.16 and 0.05 pg NPIL (Table 2). Therefore,
bioconcentration factors (BCFs) were calculated only for the 0.4, 1.6, and 3.4 pg
NPIL (measured) exposure levels (Table 2). The coefficient of variation (CV) for
triplicate water samples was less than 26%. These fish were homogenized, then
1 g aliquots were removed for analysis. The CV for triplicate measurements of
NP in fish tissue was less than 29%. BCFs were calculated by dividing the
average tissue concentration by the average measured exposure concentration

(Table 2).

DISCUSSION

The BCFs determined here are similar to previously reported values
ranging from 100 -300 (Liber et al., 1999; Naylor, 1995; Staples et al., 1998).
The analytical method described here works well for tissue samples with a mass
of less than 1.5 9. For environmentally exposed fish, it would be necessary to
assure that other residues that can fluoresce (for example, polycyclic aromatic
hydrocarbons) present in the tissue do not interfere with the HPLC analysis.
Concentrations of NP in surface waters are often less than 0.16 pg NPIL (Naylor

et al., 1992). Fish exposed to 0.16 pg NPIL would, assuming a BCF of 300,

contain a concentration of 48 pg NPlkg. Thus, the MDL for tissues presented

59

here may be insufficient to assess accurately the degree of accumulation of NP

in fish from surface waters containing small concentrations of NP.

ACKNOWLEDGMENTS

This work was supported by the Chlorine Chemical Council of the
Chemical Manufacturer’s Association, the National Institute of Environmental
Health Sciences (NIEHS) Superfund Basic Research Program (E80491), and the
US. Environmental Protection Agency (EPA) Office of Water Quality (CR
822983-01-0).

REFERENCES

Ahel, M. & Giger, W. (1993a). Aqueous solubility of alkylphenols and

alkylphenol polyethoxylates. Chemosphere 26, 1461-1470.

Ahel, M. & Giger, W. (1993b). Partitioning of alkylphenols and alkylphenol
polyethoxylates between water and organic solvents. Chemosphere 26, 1471-

1478.

Ahel, M., Giger, W. & Koch, M. (1994). Behaviour of alkylphenol

polyethoxylate surfactants in the aquatic environment - ll. Occurence and

transformation in sewage treatment. Water Research.28, 1131 -1 142.

60

Bennie, D. T., Sullivan, C. A., Lee, H. B., Peart, T. E. & Maguire, R. J.
(1997). Occurence of alkylphenols and alkylphenol mono- and diethoxylates in
natural waters of the Laurentian Great Lakes basin and the upper St. Lawrence

River. The Science of the Total Environment 193, 263-275.

Cahn, A. & Lynn Jr., J. L. (1978). Surfactants and Detersive Systems. In
“Kirk-Othmer Encyclopedia of Chemical Technology”, vol. 22, pp. 365. J. Wiley &

Sons, Inc., New York.

Desbrow, C., Routledge, E. J., Brighty, G. C., Sumpter, J. P. & Waldock,
M. (1998). Identification of Estrogenic Chemicals in STW Eflleunt. 1. Chemical
Fractionation and in vitro Biological Screening. Environmental Science &

Technology 32, 1549-1558.

Dwyer, F. J., Sappington, L. C., Buckler, D. R. & Jones, S. B. (1995). “Use
of surrogate species in assessing contaminant risk to endangered and

threatened species”, pp. 71. National Technical lnforrnation Service, Springfield,

VA.

Field, J. A. & Reed, R. L. (1996). Nonylphenol Polyethoxy Carboxylate
Metabolites of Nonionic Surfactants in US. Paper Mill Effluents, Municipal
Sewage Treatment Plant Effluents, and River Waters. Environmental Science &

Technology 30, 3544-3550.

61

Giesy, J. P., Pierens, S. L., Snyder, E. M., Miles-Richardson, S., Kramer,
V. J., Snyder, S. A., Nichols, K. M. & Villeneuve, D. A. (2000). Effects of 4-Nonyl
Phenol on Fecundity and Biomarkers of Estrogenicity in Fathead Minnows

(Pimephales promelas). Environmental Toxicology and Chemistry (in press).

Jobling, S., Sheahan, D., Osborne, J. A., Matthiessen, P. & Sumpter, J. P.
(1996). Inhibition of testicular growth in rainbow trout (Oncorhynchus mykiss)

exposed to estrogenic alkylphenolic chemicals. Environmental Toxicology and

Chemistry 1 5, 194-202.

Jobling, S. & Sumpter, J. P. (1993). Detergent components in sewage
effluent are weakly oestrogenic to fish: an in vitro study using rainbow trout

(Oncorhynchus mykiss) hepatocytes. Aquatic Toxicology 27, 361 -372.

Liber, K., Gangl, J. A., Corry, T. D., Heinis, L. J. & Stay, F. S. (1999).
Lethality and Bioaccumulation of 4-Nonylphenol in Bluegill Sunfish in Littoral

Enclosures. Environmental Toxicology and Chemistry 18, 394-400.

Lye, C. M., Frid, C. L. J., Gill, M. E., Cooper, D. W. & Jones, D. M. (1999).

Estrogenic Alklphenols in Fish Tissues, Sediments, and Waters from the UK.

Tyne and Tees Estuaries. Environmental Science & Technology 33, 1009-1014.

62

Mueller, G. C. & Kim, U. H. (1978). Displacement of estradiol from

estrogen receptors by simple alkylphenols. Endocrinology 102, 1429-1435.

Naylor, C. G. (1995). Environmental fate and safety of nonylphenol

ethoxylates. Text. Chem. Color. 27, 29-33.

Naylor, C. G., Mieure, J. R, Adams, W. J., Weeks, J. A., Castaldi, F. J.,
Ogle, L. D. & Romano, R. R. (1992). Alkylphenol ethoxylates in the environment.
JAOCS 69, 695-703.

Nimrod, A. C. & Benson, W. H. (1996). Environmental Estrogenic Effects

of Alkylphenol Ethoxylates. Critical Reviews in Toxicology 26, 335-364.

Rudel, R. A., Melly, S. J., Geno, P. W., Sun, G. & Brody, J. G. (1998).
Identification of Alkylphenols and Other Estrogenic Phenolic Compounds in
Wastewater, Septage, and Groundwater on Cape Cod, Massachusetts.

Environmental Science & Technology 32, 861-869.

Snyder, S. A., Keith, T. L., Verbrugge, D. A., Snyder, E. M., Gross, T. S.,
Kannan, K. & Giesy, J. P. (1999). analytical methods for detection of selected
estrogenic compounds in aqueous mixtures. Environmental Science &

Technology 33, 2814-2820.

63

Soto, A. M., Tien-Min, L., Honorato, J., Silvia, R. M. & Sonnenschein, C.
(1992). An "in culture" bioassay to assess the estrogencity of xenobiotics (e-
screen). In “Chemically-induced alterations in sexual and functional development:
The wildlife/human connection”, vol. 21 (ed. M. A. Mehlman). Princeton Scientific

Publishing Co.,lnc, Princeton,NJ.

Staples, C. A., Weeks, J., Hall, J. & Naylor, C. (1998). Evaluation of
Aquatic Toxicity and Bioaccumulation of C8- and 09- Alkylphenol Ethoxylates.

Environmental Toxicology and Chemistry 17, 2470-2480.

Talmage, S. S. (1994). “Environmental and Human Safety of Major
Surfactants: Alcohol Ethoxylates and Alklphenol Ethoxylates”. Lewis Publishers,

Boca Raton.

Tanghe, T., Dhooge, W. & Verstraete, W. (1999). Isolation of a bacterial
strain able to degrade branched nonylphenol. Applied and Environmental

Microbiology 65, 746-751.

White, R., Jobling, S., Hoare, S. A., Sumpter, J. P. & Parker, M. G. (1994).

Environmentally Persistent Alkylphenolic Compounds Are Estrogenic.

Endocrinology 135, 175-1 82.

64

TABLE 1. Detection Limits for Compounds of Interest

 

 

Detection Limit BP OP NP
IDL (nglml) 2 2 7
L00 (nglml) 7 8 23
MDL (ng/L Water) 23 16 45
MDL (nglg Tissue) NA 39 67

 

NA = not applicable

BP = butylphenol

OP = octylphenol

NP = nonylphenol

IDL = instrumental detection limit
LOQ = limit of quantitation

MDL = method detection limit

65

TABLE 2. Bioconcentration of NP in fathead minnows (mean

 

 

concentrations)
Nominal Measured Tissue BCF
Concentration Concentration Concentration
(HQ/LI (nglL) (91kg)
0.1 0.05 <MDL NA
0.3 0.16 <MDL NA
1 .0 0.40 81 203
3.0 1 .6 287 252
10 3.4 912 268

 

MDL = method detection limit (67 g/kg)
NA = not applicable

NP = nonylphenol

66

How H0 (CQH19)

p-t-Octylphenol p-Nonylphenol

H— (o— c— (3)1-230—9—(09H19)

Nonylphenol polyethoxylate

Figure 1. Structures of target compounds

67

 

OP

 

 

 

NP
A.
B.

 

 

Figure 2. Chromatograms of target compounds in fish tissues: (A)
standards (B) fish tissue extract from 3.4 pg NPIL exposure (C) fish

tissue extract from 0.4 pg NPIL exposure

68

Chapter 3

A METHOD FOR DETERMINING CONCENTRATIONS OF NONYLPHENOL
AND LOWER OLIGOMER NONYLPHENOL ETHOXYLATES IN FISH TISSUES

(To be submitted to Environmental Science and Technology)

ABSTRACT

A great deal of research is currently focused on the toxicological effects of
alkylphenol ethoxylates (APEs) and alkylphenols (APs) on aquatic animals.
Considerable data are available on the concentrations of APEs and APs in river
systems in the United States; however, few if any data are available on the tissue
concentrations of fish living in these rivers. A reliable method for the analysis of
nonylphenol (NP) and lower oligomer nonylphenol ethoxylates (NPE1-3) in fish
tissues has been developed. NP and NPE1.3 were extracted from fish tissues
using extractive steam distillation. Initially nonnal-phase HPLC with fluorescence
detection was used for the identification and quantification of these alkylphenols.
The method was developed and tested using laboratory-raised goldfish spiked
with the compounds of interest. However, when these environmental samples
were tested using this method, interferences prevented accurate identification
and quantitation. Therefore, normal phase HPLC was used as a cleanup step
prior to analysis by GC/MS using selected ion monitoring (SIM). Optimization of
this technique resulted in consistent recoveries in excess of 80%. Method

detection limits (MDLs) using the technique range from 2 - 5 ng/g, wet weight.

69

As a test for the methods ability to analyze environmental samples, common carp
(Cyprinus carpio) from the Las Vegas Bay of Lake Mead, Nevada were obtained
and processed. NP and NPE1 were detected in the Lake Mead carp at average
concentrations of 184 i 4 ng/g and 242 i 9, wet weight, respectively. NPE2&3

were not detected in any carp collected at Lake Mead.

INTRODUCTION

Alkylphenol polyethoxylates (APEs) are nonionic surfactants widely used
in various industrial (55% of total demand), institutional (30% of total demand)
and household applications (15% of total demand) (1). APEs are among the
most widely used nonionic surfactants, with 1998 US. use of approximately 250
million kg per year (Chemical Market Reporter, Oct. 18, 1999). APEs are
primarily used as surfactants, which can function as detergents, wetting agents,
dispersants, emulsifiers, solubilizers and anti-foaming agents. APEs are used in
many industrial applications, including pulp and paper, textiles, coatings,
agricultural pesticides, lube oils and fuels, metals and plastics. Most of the APEs
used are of the nonylphenol polyethoxylate (NPE) types that contain a 9 carbon
branched isomeric alkyl group (1, 2). NPEs and the less widely used octylphenol
polyethoxylates (OPEs) degrade during wastewater treatment or in the
environment through transient intermediates including alkylphenol
ethoxycarboxylates (APECs), lower oligomer APEs such as NPE(1-3), and

alkylphenols such as octylphenol (OP) and nonylphenol (NP) (Figure 1) (2-5).

70

The type and efficiency of wastewater treatment affects the degree of APE
removal during treatment. In the US. where most wastewater treatment plants
(VWVTPs) use secondary or tertiary treatment, removal rates generally exceed
95% (6).

Various reports have shown that APs and certain APEs can have adverse
affects on aquatic organisms when concentrations are great (1, 7-9). NP and OP
have been shown to be estrogenic in a variety of both in vitro (9-13) and in vivo
bioassays (14, 15). Some oligomers of NPE have also been found to be
estrogenic in vitro (9, 14, 16, 17) and in vivo (14).

The most comprehensive survey from the US. reported concentrations of
NP and NPEs from 30 rivers that are influenced by municipal or industrial
wastewater effluents (2). That study found that 60-75% of samples had no
detectable levels of NP, NPE1, or NPE; (2). However, data are scarce on the
concentrations of APs and APEs in fish in US. waters. Recently, NP and NPE
have been reported to occur in the Las Vegas Bay of Lake Mead, Nevada (18).
Because of this known contamination, it was determined a suitable site to
capture fish likely to have detectable NP and NPE tissue concentrations.

The objective of this study was to develop a reliable, cost-effective,
and simple method to sensitively detect and quantify NP and NPEM, in the
tissues of fish. Extractive steam-distillation has been used previously for the
extraction of alkylphenols from sediment, water, and fish tissue (19-22).
Extractive steam-distillation minimizes the coextraction of large molecular weight

interferences (such as lipids) while efficiently extracting compounds with

71

sufficient volatility (23). This method also uses far less solvent then would be
required for Soxhlet extraction. Therefore, extractive steam-distillation coupled
with norrnaI-phase high-pressure liquid chromatography (NP-HPLC) clean-up
and gas chromatography with mass selective detection (GC/MSD) was
developed as a reliable and sensitive method for the identification and
quantitation of NP and NPE(1-3). Goldfish spiked with various concentrations of
the compounds of interest were used for method development. Carp captured

from the Las Vegas Bay of Lake Mead, Nevada were used for method validation.

EXPERIMENTAL SECTION

Standards and Reagents. All standards and reagents used were of the highest
purity commercially available. High purity standards (approximately 96% or
greater purity) of p-nonylphenol (NP), p-cumylphenol (CP), and 4-tert-butyl
orthocresol (TBC) (Figure 1) were obtained from Schenectady International
(Freeport, TX). Standards of NPE(1-3) were obtained from Huntsman Corporation
(Austin, TX). Pesticide residue grade hexane, dichloromethane (DCM), and iso-
octane were obtained from Burdick and Jackson (Muskegon, MI). Reagent water
was first purified by reverse osmosis (R0) followed by NanopureTM (Barnstead,

Dubuque, IO) treatment.

Sample Collection and Preservation. Laboratory-raised goldfish were

euthanized using MS-222 then frozen until analysis. Carp from Las Vegas Bay

72

of Lake Mead, Nevada, were captured in June of 1999 via gill nets, euthanized
with MS-222, then frozen until analysis. An approximately 150 g representative
cross-section from a large carp (approximately 1 kg) captured from the Las
Vegas Bay of Lake Mead, Nevada, was homogenized. Twenty g aliquants of this

homogenate were processed as described previously.

Extraction. A 20 g representative cross-section was cut from the whole fish and
added to a homogenizer (Blender 700, Waring Corporation, New Hartford, CN)
with 350 mL of laboratory water. This mixture was homogenized for 2 min. The
homogenate was added to a 2 L boiling flask with 20 g sodium chloride (ACS
reagent grade, JT Baker, Phillipsburg, NJ), 3 mL concentrated sulfuric acid (GR
grade, EM Science, Gibbstown, NJ), several glass boiling chips (3 mm diameter,
Pyrex, Big Flats, NY), and a TeflonTM stir-bar. An additional 650 mL of laboratory
water is added to the homogenizer in small portions to rinse and transfer any
remaining homogenate into the 2 L boiling flask. Five hundred ng of CP was
added to the boiling flask mixture as a surrogate standard.

The mixture described above was placed onto a heating mantle
with a magnetic stir plate below the mantle. A Nielsen-Kryger improved version
steam-distillation column (Ace Glass, Vineland, NJ) was attached to the round
bottom flask containing the sample. Three mL of laboratory water and 10 mL of
iso-octane were added to the steam-distillation column. Tap water was
circulated through the condenser at a fast flow rate. The heating mantle was

operated on the maximum setting and the stir plate speed was set to

73

approximately 50%. The mixture was boiled for 1.5 h and the water layer in the
distillation column discarded. The iso-octane layer was collected in a 15 mL
centrifuge tube. An additional 3 mL of laboratory water and 10 mL of iso-octane
were added to the distillation column. The mixture was then boiled again for 1.5
h and the iso-octane layer combined with the initial extract. The combined

extract was concentrated to 1 mL at 30 °C under a gentle stream of purified

nitrogen and stored at -20 °C until analysis.

Normal-Phase HPLC Fractionation. The NP-HPLC fractionation system
consisted of a Perkin Elmer (PE) (Nonrvalk, CT) series 200 autosampler, a PE
series 200 binary pump, a Hewlett Packard (HP) 1046A flourescence detector,
and PE TurboChrome 4.0 data software package. An 800 pL injection volume of
the iso—octane extract was separated using a Phenomenex Luna 5 pm silica
column (250 mm x 4.6 mm, Torrance, CA) with isocratic elution of 88% hexane
and 12% 1:4 MeOHzDCM and a flow rate of 0.65 mUmin. Fluorescence
detection was used to determine surrogate recovery during this fractionation at
an excitation of 229 nm and emission of 310 nm. The HPLC effluent containing
the analytes of interest was collected during the time period of 7 to 16 min. To
this fraction, 3.0 pg of TBC was added as an internal standard. This fraction was
then concentrated using a gentle stream of purified nitrogen to 100 pL iso-

octane.

74

Identification and Quantitation. Compounds of interest were identified and
quantified using a HP 5890 series ll plus GC and a HP 5972 MSD. Separation
was accomplished using a 30 m DB-17MS capillary column (0.25 mm ID, 0.15
pm film, J&W Scientific, Folsom, CA) and a 4 pL injection volume. The G0 was
programmed with a starting temperature of 100 °C for 2 min, followed by a 4
°C/min temperature ramp to a final temperature of 300 °C that was held for 10
min. The MSD was operated in selected ion monitoring (SIM) mode with 3 ions
monitored for each compound of interest (Table 1). Each sequence began with
one blank and one standard mixture. Identification was made by proper retention
time and ion ratios. Peak areas were determined by electronic integration and
concentrations were determined using external calibration and Microsoft Excel
version 7.0 spreadsheets (Seattle, WA). Standard calibration curves were linear

(r2 > 0.9) across the entire calibration range.

Recovery and Detection Limits. Instrument detection limits (lLDs) were
defined as the minimum mass of analyte required to yield a signal to noise ratio
of 3. Seven replicates of homogenized goldfish tissues were spiked with NP and
NPE(1-3) at the estimated method detection limits (MDLs) to determine recovery
and precision (Table 1). MDLs were calculated by multiplying the standard
deviation (SD) of the recovered concentrations by a t-value of 3.1427 (for n=7
replicates). The limits of quantitation (LOQs) were calculated by adding 5 80s to
the MDL.

75

RESULTS AND DISCUSSION

It was determined that two successive distillations with fresh iso-octane
were required to achieve acceptable analyte recovery. It was also observed that
two successive extractions of 1.5 h each were more efficient than one 3 h
distillation. Recoveries of the compounds of interest using two distillations were
consistently greater than 70% with the exception of NPE3 (Table 2). The
coefficient of variation (CV) in recoveries were less than 20% (T able 2). The
lesser recovery of NPE3 is a function of its lesser volatility and lipophillicity. Initial
method development with steam-distillation resulted in extreme foaming that
resulted in contamination of the iso-octane extract. While silicon-based anti-
foaming agents successfully reduced foaming, recoveries were reduced and
inconsistent. The addition of concentrated sulfuric acid reduced foaming and did
not negatively effect recovery or precision. Although cooling the distillation
condensers resulted in slightly greater recovery, the improvement was small.

NP and NPE1 were detected in the tissue from the carp captured in Las
Vegas Bay (Table 3). The variability in concentrations of NP and NPE1 among
the subsamples was small (Table 3). These concentrations observed are
reasonable, based upon reported water concentrations (18) and published
bioconcentration factors (BCFs) of 100 to 300 (8, 24, 25). Although
concentrations of NP and NPE in sediments of Las Vegas Bay are unknown, it is
likely that sediment bound NP and NPE effected the carp tissue concentrations,

as carp are bottom-feeders.

76

The method presented here was successful for sensitive and
reproducible measurements of concentrations of NP and NPE“ a. 2) in fish tissues.
The resulting detection limits are sufficient to determine toxicologicalIy-relevant
concentrations of the compounds of interest in fish tissues. This method also
utilizes laboratory equipment that is common or inexpensive, which makes the

method ammenable for use in most environmental laboratories.

ACKNOWLEDGMENTS

Funding for this project was provided by the Alkylphenol Exthoxylate
Research Coucil (APEC). Carp were provided by the Nevada Division of Wildlife,
Las Vegas, Nevada. The authors would like to thank Erin M. Snyder for critically
reviewing this manuscript. We would also like to thank Bob Fensterhelm of the

APERC for technical guidance of this project.

REFERENCES

(1) Talmage, S. S. Environmental and Human Safety of Major Surfactants:
Alcohol Ethoxylates and Alklphenol Ethoxylates; Lewis Publishers: Boca
Raton. 1994.

(2) Naylor, C. G.; Mieure, J. R; Adams, W. J.; Weeks, J. A.; Castaldi, F. J.;
Ogle, L. D.; Romano, R. R. JAOCS1992, 69, 695-703.

(3) Field, J. A.; Reed, R. L. Environ. Sci. Technol. 1996, 30, 3544-3550.

77

(4)

(5)

(6)
(7)

(8)

(9)

(10)
(11)

(12)

(13)

(14)

Rudel, R. A.; Melly, S. J.; Geno, P. W.; Sun, 6.; Brody, J. G. Environ. Sci.
Technol. 1998, 32, 861-869.

Bennie, D. T.; Sullivan, C. A.; Lee, H. B.; Peart, T. E.; Maguire, R. J. Sci.
Total Environ. 1997, 193, 263-275.

Naylor, C. G. Text. Chem. Color. 1995, 27, 29-33.

Weeks, J. A.; Adams, W. J.; Guiney, P. D.; Hall, J. F.; Naylor, C. G. The
Alkyphenols and Alkylphenol Ethoxylates Review 1998, 1, 64-74.

Staples, C. A.; Weeks, J.; Hall, J.; Naylor, C. Environ. Toxicol. Chem.
1998, 17, 2470-2480.

White, R.; Jobling, S.; Hoare, S. A.; Sumpter, J. R; Parker, M. G. Endo.
1994, 135, 175-182.

Mueller, G. C.; Kim, U. H. Endo. 1978, 102, 1429-1435.

Soto, A. M.; Tien-Min, L.; Honorato, J.; Silvia, R. M.; Sonnenschein, C. In
Chemically-induced alterations in sexual and functional development: The
wildlife/human connection; Mehlman, M. A., Ed.; Princeton Scientific
Publishing Co.,lnc: Princeton,NJ, 1992; Vol. 21.

Desbrow, C.; Routledge, E. J.; Brighty, G. C.; Sumpter, J. P.; Waldock, M.
Environ. Sci. Technol. 1998, 32, 1549-1558.

Jobling, S.; Noylan, M.; Tyler, C. R.; Brighty, G.; Sumpter, J. P. Environ.
Sci. Technol. 1998, 32, 2498-2506.

Jobling, S.; Sheahan, D.; Osborne, J. A.; Matthiessen, P.; Sumpter, J. P.

Environ. Toxicol. Chem. 1996, 15, 194-202.

78

(15)

(15)

(17)

(13)

(19)

(20)

(21)

(22)

(23)

(24)

(25)

Nimrod, A. C.; Benson, W. H. Critical Reviews in Toxicology 1996, 26,
335-364.

Jobling, S.; Sumpter, J. P. Aquat. Toxicol. 1993, 27, 361-372.

Routledge, E. J.; Sumpter, J. P. Joumal of Biological Chemistry 1997,
272, 3280-3288.

Snyder, S. A.; Keith, T. L.; Verbrugge, D. A.; Snyder, E. M.; Gross, T. S.;
Kannan, K.; Giesy, J. P. Environ. Sci. Technol. 1999, 33, 2814-2820.

Lye, C. M.; Frid, C. L. J.; Gill, M. E.; Cooper, D. W.; Jones, D. M. Environ.
Sci. Technol. 1999, 33, 1009-1014.

Kubeck, E.; Naylor, C. G. JAOCS 1990, 67, 400-405.

Fowler, B. R.; Haviland, L.; Kennedy, S. SETAC Presentation 1998.

Ahel, M.; McEvoy, J.; Giger, W. Environmental Pollution 1993, 79, 243-
248.

Veith, G. D.; Kiwus, L. M. Bull. Environ. Contam. Toxicol. 1977, 17, 631-
636.

Liber, K.; Gangl, J.; Corry, T.; Heinis, L.; Stay, F. Environ. Toxicol. Chem.
1999, 18, 394-400.

Snyder, S. A.; Keith, T. L.; Pierens, S. L.; Snyder, E. M; Giesy, J. P.
Chemosphere, 2000 (submitted).

79

TABLE 1. lens Monitored and Recovery Data

 

 

Compound Ions (mlz) Spike (nglg) % Recovered
NP 107, 135, 149 15.4 78.1 x 9.21
NPE1 135, 179, 193 74.8 76.1 i 9.84
NPE; 135, 223, 237 67.2 69.4 i 13.0
NPE3 135, 267, 281 40.0 17.0 i 20.1
TBC 149, 164, 121 NA NA

 

NA = not applicable

NP = nonylphenol

NPE1 = nonylphenol monoethoxylate
NPE2 = nonylphenol diethoxylate
NPE3 = nonylphenol triethoxylate

TBC = tert-butyl orthocresol

80

TABLE 2. Detection Limits for Compounds of Interest

 

Compound IDL(ng) MDL(ngIg) LOQ(ngIg)

 

NP 5.12 3.30 4.82
NPE1 15.5 16.8 18.5
NPE; 17.3 18.2 20.6
NPE3 112 20.6 28.9

 

NP = nonylphenol
NPE1 = nonylphenol monoethoxylate
NPE; = nonylphenol diethoxylate

NPE3 = nonylphenol triethoxylate

81

TABLE 3. Average Concentrations of NP and NPEs in Lake Mead Carp

 

 

Compound nglg wet wt. %CV
NP 184 4.4
NPE1 242 9.3
NPEz ND NA
NPE3 ND NA

 

ND = not detectable at MDL

NA = not applicable

NP = nonylphenol

NPE1 = nonylphenol monoethoxylate
NPE; = nonylphenol diethoxylate

NPE3 = nonylphenol triethoxylate

82

Ho. Ho.

p-tert-butyl orthocresol p-Cumylphenol

H0 (C9H19)

p-Nonylphenol

H— (o— c—cy; o—.—(c,H,,)

Nonylphenol ethoxylate1 4

Figure 1. Structures of Compounds of Interest.

83

Chapter 4

ANALYTICAL METHODS FOR DETECTION OF SELECTED ESTROGENIC
COMPOUNDS IN AQUEOUS MIXTURES

(Published in Environmental Science and Technology, 1999)

ABSTRACT

Both natural estrogens and synthetic compounds that mimic estrogen can
reach the aquatic environment through wastewater discharges. Because
nonylphenol (NP), octylphenol (OP), nonylphenol polyethoxylates (NPE), 17B-
estradiol (E2), and ethynylestradiol (EE2) have previously been found to be
estrogenic and occur in wastewater effluents, they were the primary analytes for
which the method was developed. Water samples were extracted in situ with
solid-phase extraction disks. Analytes were separated by high-pressure liquid
chromatography (HPLC) and detected by fluorescence or competitive
radioimmunoassay (RIA). Method detection limits (MDLs) using HPLC with
fluorescence detection were 11, 2, and 52 nglL of water for NP, DP, and NPE,
respectively. The RIA MDLs for E2 and EE2 were 107 and 53 pglL, respectively.
Samples were collected from 4 municipal wastewater treatment plants (WWTPs)
in south central Michigan, 8 locations on the Trenton Channel of the Detroit
River, Michigan, and 5 locations in Lake Mead, Nevada. Concentrations of NP

and OP ranged from less than the MDL to 37 and 0.7 pglL, respectively. NPE

84

concentrations ranged from less than the MDL to 332 pglL. Concentrations of E2

and EE2 ranged from less than the MDLs to 3.7 nglL and 0.8 nglL, respectively.

INTRODUCTION

The potential effects of chemicals that alter the normal endocrine function
and physiological status of animals have been an increasing concern in recent
years (1). Reports of apparent increases in hormone-dependent cancers and
decreases in sperm quantity and quality in humans have raised questions about
the role of natural and synthetic chemicals in these trends (2-7). A number of
pollutants including pesticides, certain polychlorinated biphenyls (PCBs), dioxins,
furans, alkylphenols, synthetic steroids and natural products such as
phytoestrogens have been reported to disrupt normal hormonal pathways in
animals and collectively have been referred to as endocrine disrupters or
endocrine disrupting chemicals (EDCs) (2, 8). While EDCs can operate through
a number of both direct and indirect mechanisms of action, of particular concern
are those compounds that mimic endogenous estrogens. The Safe Drinking
Water Act Amendments of 1995 (bill number S1316) and the Food Quality
Protection Act of 1996 (bill number PL. 104-170), which mandate
comprehensive screening for estrogenic and anti-estrogenic chemicals, are
examples of the increasing public concern regarding endocrine disruption. While
it is known that many natural and synthetic chemicals are estrogenic, it is unclear

whether or not the concentrations of estrogenic agents present in the

85

environment are sufficient to cause adverse physiological effects. One aspect of
conducting human or wildlife risk assessments is an exposure assessment. This
suggests the need for assays and techniques to monitor the quantity and effects
of endocrine disrupters.

WWTPs receive natural and synthetic EDCs from urban and
industrial dischargers. WWTPs use a variety of treatment processes of varying
efficiency such that in some cases compounds are not completely removed by
the treatment processes and are ultimately discharged into surface waters. Fish
caged below WWTP outfalls have been reported to have abnormal ratios of sex
steroid hormones as well as to exhibit histological changes in their reproductive
organs (9-11). Fish living in waters influenced by WWTP effluents have been
observed to express similar effects attributed to unknown or unspecified EDCs
(12-14). Extracts of some WWTP effluents in the United Kingdom have been
shown by use of in vitro assays, to be estrogenic due to the presence of
alkylphenols and natural and synthetic estrogens (15).

Alkylphenol polyethoxylates (APEs) are non-ionic surfactants
widely used in various industrial (55% of total demand), institutional (30% of total
demand) and household applications (15% of total demand) (16). APEs are
among the most widely used non-ionic surfactants, with 1998 US. use of
approximately 204 million kg per year (16). Most of the APEs used are of the
nonylphenol polyethoxylate (NPE) types that contain a 9 carbon branched
isomeric alkyl group (16, 17). NPEs and the less widely used octylphenol

polyethoxylates (OPEs) degrade during wastewater treatment or in the

86

environment to alkylphenol ethoxycarboxylates (APECs), lower oligomer APEs
such as NPE1-3, and alkylphenols such as octylphenol (OP) and nonylphenol
(NP) (Figure 1) (17-20). NP and OP have been shown to be estrogenic in a
variety of both in vitro (9, 15, 21-23) and in vivo bioassays (24-26). Some
oligomers of NPE have also been found to be estrogenic in vitro (9, 23, 24, 27)
and in vivo (24). Reports of APEs and their corresponding degradation products
from the US. are scarce. The most comprehensive survey reports
concentrations from 30 US. rivers that are influenced by municipal or industrial
wastewater effluents (17). That study found that 60-75% of samples had no
detectable levels of NP, NPE1, and NPEz (17).

The endogenous hormone 178-estradiol (E2) and the synthetic
hormone ethynylestradiol (EE2) have been detected in several WWTP effluents
(Figure 1) (15, 28-31). Synthetic hormones are generally more stable in water
than are natural hormones and have greater potency (31, 32). EE2 was found to
be unchanged after 120 h in activated sludge treatment, indicating possible
release into surface waters after waste water treatment (29). EE2 is a widely
used pharmaceutical with daily oral doses ranging from 30 - 50 pg (33).
Estrogens are excreted by humans in concentrations as great as 2.7 mg/L on a
daily basis (34). Conjugated estrogens used in the treatment of cancer, hormonal
imbalance, osteoporosis, and other ailments are the second most prescribed
drug in the US. and are administered orally in doses ranging from 0.3 — 2.5 mg
(33). Estrogens are excreted by mammals as water-soluble conjugates in urine

or as “free” estrogens in feces (30, 35, 36). Conjugated forms of these estrogens

87

can be deconjugated during wastewater treatment and In the environment to
generate the more potent “free” estrogen (15, 31). Environmental concentrations
of E2 and EE2 range from less than detection to greater than 140 ng/L and less
than detection to 15 nglL, respectively (15, 28, 29, 32). Fish exposed to E2 and
EE2 exhibit changes in biomarkers for estrogenicity at concentrations as low as
0.5 nglL (10, 15, 37, 38).

In this report, we describe a rapid and sensitive method for
detecting and quantifying EDCs in surface waters. The method was designed to
isolate and concentrate a broad range of organic compounds from surface
waters so that it could be used as an analytical tool for toxicity identification and
evaluation (TIE). An in situ extraction technique using 90 mm
styrenedivinylbenzene (SDB) EmporeTM solid-phase extraction disks was
developed. This technique permits the extraction of greater volumes of water
more rapidly and efficiently than the typical vacuum filtration methods (39). This
method also provides detection limits less than the reported effects
concentrations for these compounds in the aquatic environment. The SDM
matrix is effective for extracting polar organics, which is necessary has the
compounds of interest all have polar hydroxyl functional groups. Previously
EmporeTM disks have been used for the extraction of natural water sucessfully
(18, 39-41). However, these compounds of interest were not investigated. APs
and certain steroids are extracted by this method without requiring the transport
of large volumes of water, thus reducing losses due to adsorption to transport

containers and due to microbial degradation (Figure 2). To demonstrate the

88

 

 

 

utility of this method, samples of effluents and surface waters were collected from
several locations in the Trenton Channel of the Detroit River, Michigan; below the
outfalls of municipal WWTPs in south central Michigan; and in the Las Vegas

(LV) Wash and in Lake Mead, Nevada.

EXPERIMENTAL SECTION

Standards and Reagents. All standards and reagents used were of the highest
purity commercially available. High purity standards (98% pure) of p-nonylphenol
(NP), p-tert butylphenol (BP), and p-tert octylphenol (OP) were obtained from
Schenectady International (Freeport, TX). Standards of nonylphenol
monoethoxylate (NPE1) and nonylphenol polyethoxylates (SurfonicTM N-95) were
obtained from Huntsman Corporation (Austin, TX). Standards of E2 and EE2 of
98% purity were obtained from Sigma Chemical Company (St. Louis, MO). High
purity, pesticide residue grade ACN, methanol (MeOH), hexane,
dichloromethane (MeClz), iso-octane, 2-propanol, and acetone were obtained
from Burdick and Jackson (Muskegon, MI) or Sigma-Aldrich (Milwaukee, WI).
Solvents were also tested for interferences by examining 150-fold concentrated
solvents using high-pressure liquid chromatography (HPLC) with fluorescence
detection and gas chromatography/mass spectrometery (GCIMS). Reagent
water was first purified by reverse osmosis (RO) followed by NanopureTM
(Barnstead, Dubuque, IA) treatment. Glass fiber filters, fine grade (GF/F) of 0.7

pm nominal pore size and coarse grade (GF/C) of 1.2 pm nominal pore size

89

(\Nhatman, Maidstone, England), were heated at 450 °C for 4 h in aluminum foil
prior to use. Anhydrous granular sodium sulfate (EM Science, Gibbstown, NJ)

was heated at 550 °C overnight then stored at 125 °C until day of use.

Sample Collection and Preservation. Water samples were collected from 8
sites in the Trenton Channel of the Detroit River, MI; 4 WWTPs in south central
Michigan, and 5 sites in Lake Mead, NV. To minimize effects due to dilution,
effluent samples were collected as close to the source as possible. Samples
from the Trenton Channel and Lake Mead were collected at 50% of the
maximum depth or 3 m, whichever was less. Samples were collected between
April and October 1997.

All samples were filtered and extracted at the sample location
(Figure 2). Water samples were pumped through fluoropolymer (PFA) tubing
(3l8” o.d.) by a Fluid Metering (Oyster Bay, NY) DC pump (model QB) fitted with
a ceramic lined stainless steel pump head (model Q1-CSC) to a 90 mm stainless
steel pressure filtration holder (Millipore, Bedford, MA). Flow rate was
maintained at 100 mL/min until 5 L total volume had been extracted or until the
pressure exceeded 100 psi. Stainless steel screen (250 pm mesh, Aquatic
Ecosystems, Apopka, FL) was used to prevent large particles from entering the
inlet tubing. A glass fiber filter was placed on each side of a 90 mm poly
(styrenedivinylbenzene) (SDB-XC) EmporeTM (3M Corporation, St. Paul, MN).
The bottom filter (90 mm GF/C filter) was used to improve flow characteristics,

and the top filter (90 mm GF/F) was used to filter particulates from the sample.

90

 

Fifteen mL each of acetone, MeClz, and MeOH, and 20 mL reagent water were
injected sequentially through a custom Luer lock fitting on the relief valve of the
filtration holder with a 2 min residence period to condition the EmporeTM disk. A
McMillan (Georgetown, TX) electronic flow meter (model 102-6TP), calibrated to
i 1% accuracy with National Institute of Standards and Technology (NIST)
traceable standards, was installed downstream of the filtration holder. This
digital rate meter and totalizer was used to monitor flow rate and to record the
total volume. Once the desired sample volume had been reached, a volume of
20 mL of air was pushed through the filter holder to move headspace water
through the holder. The filter holder then was disassembled and the GF/F and
EmporeTM disk were stored separately in clean aluminum foil and kept on ice for
transport to the laboratory. The GF/C filter was discarded.

At each sampling site, prior to collecting the samples, the apparatus
was purged with sample for 10 min by bypassing the filter holder. At sites with
concentration gradients, samples were taken sequentially from the least to
greatest predicted concentrations. Flow meter calibration was verified prior to
each sampling trip by comparing the reported totalizer output to the measured
volume of water exiting the meter. The sampling apparatus was cleaned with
water and solvents before and after each sample.

Field blanks were taken daily by passing 5 L of NanopureTM water through
the system as previously described. Laboratory blanks were performed on each
day of extractions and instrument blanks were conducted daily. No blank had

detectable concentrations of any of the compounds of interest. None of the

91

 

compounds of interest were detected in the blanks. Breakthrough tests were
performed by placing two EmporeTM disks in series and analyzing each disk
separately. Breakthrough was detected only at the greatest concentrations

spiked and the breakthrough was determined to be less than 11%.

Extraction. EmporeTM disks were eluted on a 90-mm vacuum manifold (3M
Corporation, St. Paul, MN). Fifteen mL of acetone, 25 mL of MeClz, and 10 mL
of hexane were added sequentially to the reservoir and collected in a 50-mL
centrifuge tube. Slight vacuum was applied to initiate solvent transfer and the
remainder was permitted to flow without vacuum. Vacuum was applied once all
solvents had passed through the disk to remove residual solvents from the disk.
Moisture was removed from the solvent extract by passing it through 30 g
anhydrous sodium sulfate placed in a custom glass funnel containing a stopcock
at 1 drop/sec. The eluate was collected in a flat bottom flask and rotary
evaporated at 30 °C to approximately 5 mL. The sample was concentrated and
solvent-exchanged to 1 mL iso-octane under a gentle stream of nitrogen at 30
°C. One mL of 40% hexane in MeClz was added to the test tube and vortexed
for 5 sec. The sample was collected in a 2.5 mL Hamilton syringe (Reno, NV)

and injected into a normal phase HPLC system for fractionation.
Normal-Phase HPLC Fractionation. The HPLC fractionation system consisted

of a Rheodyne 7725i injector (Cotati, CA) with a 5 mL sample loop, a quaternary

pump (Perkin Elmer, Series 410, Nonrvalk, CT), and an electronic fraction

92

collector (ISCO Foxy 200, Lincoln, NE). A Phenomenex Luna 5 pm silica column
(250 mm x 4.6 mm, Torrance, CA) was used for nonnal-phase separations with
3-step isocratic elution. The mobile-phase solvent profile was 30% MeClz in
hexane for 15 min, MeCIz for 20 min, and MeOH for 20 min, each at 1 mUmin
with no gradient curves. The column was returned to initial conditions by passing
MeClz for 10 min followed by 30% MeClz in hexane for 35 min at 1mUmin.
Fractions were collected from 0-20 min (F 1), 20-45 min (F2), and 45-70 min (F3)
with no delay. F1 contained the most non-polar compounds such as PAHs,
PCBs, organochlorine (OC) pesticides, and fluorescent compounds leached from
the EmporeTM SDB-XC disk. F2 contained BP, NP, and OP. F3 contained the
most polar compounds, including E2, EE2, and NPEs. Each fraction was
concentrated by rotary evaporation at 30 °C to approximately 5 mL. The sample
was evaporated and solvent-exhanged to 1 mL ACN under a gentle stream of
nitrogen. No internal or surrogate standards were added to the samples since
these could interfere with the in vitro bioassays used to screen for total

estrogenic and dioxin-like activity.

Quantitation of Compounds. F2 and F3 were analyzed by reverse-phase
HPLC (Figures 3 and 4). The reverse-phase HPLC system included a Perkin
Elmer (PE) (Nomalk, CT) series 200 autosampler, a PE series 200 binary pump,
a Hewlett Packard (HP) 1046A fluorescence detector, and PE TurboChrome 4.0
data software package. The column used was a Phenomenex Prodigy 5 pm

octadecylsilica (ODS) 100 A (250 mm x 4.6 mm, Torrance, CA) preceded by a 30

93

 

mm x 4.6 mm guard column of the same packing material. For all compounds of
interest, thefluorescence detector settings were: 229 nm excitation, 310 nm
emission, PMT gain of 12, lamp time of -1, response time of 2 sec, stop time at
27 min, gate and delay at zero.

Elution solvents for gradient elution were reagent water and ACN
delivered at a constant flow rate of 1 mL/min. The elution profile was a 20 min
gradient (curve = -2) from 50% water and 50% ACN to 2% water and 98% ACN
followed by a 10 min isocratic ACN purge. Each sequence began with one blank
and one standard. The injection volume of standards and samples was 10 pL.
Peak areas were determined by electronic integration, and sample
concentrations were determined using Microsoft Excel version 7.0.
Chromatography and retention times for the compounds of interest are shown in
Figures 3 and 4. All calibration curves were linear (r2 > 0.99) across the entire
calibration range. An HP 5890 series ll plus GC with a 30 m DB5-MS capillary
column (J&W Scientific, Folsom, CA) and a HP 5972 series mass selective

detector (MSD) were used for confirmation of peak assignments.

Radioimmunoassay (RIA). Competitive radioimmunoassay (RIA) was used for
detection of E2 and EE2 following a procedure previously described for
measurement of these compounds in plasma and adapted for environmental
samples (14). The most polar fraction (F3) from each of the HPLC fractionation
procedures was analyzed in duplicate for both E2 and EE2. A 20 pL aliquot of

sample was evaporated to dryness in a 5 mL glass test tube by use of a vortex

94

evaporator (Labconco, Kansas City, MO) and nitrogen gas. Each sample was
reconstituted in 0.5 M sodium phosphate buffered saline buffer (PBSGA).
Standard curves were prepared in PBSGA.

Accuracy and precision of the assay were determined in several ways.
Cross-reactivities of the E2 antiserum with other steroids are 11.2 % for estrone;
1.7% for estriol; <1.0 % for 178-estradiol and androstenedione; and <0.1% for
EE2, DES, and all other steroids examined. Cross-reactivities of the EE2
antiserum with other steroids were 0.3% for E2; <0.1% for norethindrone,
estrone, 178-estradiol, diethylstilbestrol, hexoestrol and dienoestrol; and <0.01%
for all other steroids and compounds tested. To determine recovery during
reverse-phase HPLC fractionation, known amounts (1, 2, 5, 10, 25, 50, 100, 250,
and 500 pg) of E2 or EE2 were fractionated by HPLC and the masses of the
recovered materials determined. No significant losses occurred during
fractionation. Parallelism between the dose-response relationships for standards
and samples were evaluated by assaying dilutions of HPLC fractions (2.5, 5, 10,
15, and 20 pL HPLC fraction volumes). The standard curves were determined to

be parallel to the dilution curves for HPLC fractions.

Recovery and Precision. Spike/recovery experiments were performed using
reagent water or untreated ground water to determine the accuracy and precision
of the method. Nineteen-L glass carboys were spiked with NP, BP, OP, and EE2
and mixed for at least 1 h. Three 4-L samples were sequentially extracted from

the 19-L mixture using the procedure described as above. Three spikes were

95

also performed to determine the effect of larger volumes by spiking 19 L and
extracting 17 L. Flow rate was optimized by spiking 4 L of water with the
compounds of interest and loading the EmporeTM disks at different rates.
For NPE, quantitation was based on a sum of all oligomers and reported
as total NPE (NPE). The instrument was calibrated using SurfonicTM N-95.
Therefore, all results relative to NPE concentrations are semi-quantitative in
relation to the defined conditions. To assure NPE oligomer distribution integrity,
oligomer distribution of extracted NPE was compared to that of an NPE (as
Surfonicm N-95) standard. The NPE oligomer distribution was determined by
normal phase HPLC using a Phenomenex Phenosphere 5 pm cyanopropyl
column (250 mm x 4.6 mm). The PE binary HPLC instrument and fluorescence
detector have been described previously. The mobile phase solvents consisted
of 35% iso-propanol in MeOH (A) and hexane (B). The elution profile was 3% A
and 97% B for 7 min at a flow rate of 0.5 mUmin isocratic followed by a 20 min
gradient to 95% A and 5% B at a curve of -5 and flow rate of 1 mUmin. The
column was purged a 1 mUmin with 3% A and 97% B to return the column to the
initial conditions. It was determined that the oligomer distribution remained
unchanged throughout the procedure (Figure 5). Oligomer-spec'rfic distributions
were not determined for environmental samples.
To determine losses of EE2, OP, NP, and BP during evaporation
procedure, these compounds were spiked into a 250 mL flat bottom flasks

containing the correct amounts and ratios of solvents used for the EmporeTM

96

 

extraction. Solvents were rotary evaporated and the volume was adjusted as

shown earlier.

Detection Limits. The instrumental detection limits (IDLs) and limits of
quantitation (LOQs) for NP, OP, BP, NPE, and EE2 by reverse phase HPLC with
fluorescence detection were determined by injecting 10 pL of a 50 ngImL mixture
of each of these compounds 5 times. The IDLs and LOQs are defined as 3 times
and 10 times the standard deviation (SD) of the quantified peak for each
compound in the mixture, respectively. The method detection limits (MDLs)
using reverse phase HPLC with fluorescence detection (10 pL injections) were
estimated from the mass of analyte at the LOQ divided by 5 L sample size plus
25% to account for a 75% average recovery. The IDLs for E2 and EE2 using
RIA were determined using lmmunoFitTM EIAIRIA Data Analysis Software
(Beckman, Fullerton, CA). The minimum concentration distinguishable from zero
for E2 and EE2 was 427 and 211 pg/mL extract, respectively. The MDLs for E2
and EE2 using RIA detection were calculated in the same manner as for HPLC

with fluorescence detection.

RESULTS

Recoveries for these compounds at various concentrations are shown

(Table 1). It was determined that recovery did not vary significantly between

untreated ground water and reagent water. The compounds of interested added

97

to a natural water sample from Lake Mead were recovered within the range
determined by laboratory spike-recovery experiments. Flow rates greater than
500 mLImin resulted in poor recoveries (<50%) possibly due to breakthrough.
The greatest average recovery of > 80% for all analytes was achieved using a
flow rate of 50 mLImin; however, this resulted in extraction times > 1.5 h. A flow
rate of 100 mLImin resulted in consistent recoveries of the compounds of interest
with an acceptable average recovery >70% (Table 1). BP was the only
compound with poor recoveries (average 60%). BP was tested for use as a
possible surrogate standard in future studies because it has been used
previously as an internal standard (42). BP has a greater volatility than the other
compounds of interest, which is the likely cause for its lesser recoveries (Table
1). If rotary and nitrogen evaporation temperatures and final volumes were not
carefully monitored, losses of up to 50% were observed. By maintaining the
evaporation temperatures at 30 °C and avoiding over-concentration, evaporation
losses could be kept consistent and less than 10%.

It was determined that EE2 could function as a surrogate for E2.
Recoveries for E2 and EE2 were virtually the same for each experiment
performed. Additionally, E2 required fresh standards weekly, while EE2 was
stable and remained in calibration for at least 6 mo. NPE was also spiked and
recovered 4 times with an average recovery of 71% i 12. In environmental
samples, oligomers greater than NPE4 rarely were observed (qualitatively) by

GC/MS.

98

The IDLs, LOQs, and MDLs for the compounds of interest for each
method of detection are presented in Table 2. The coefficient of variation (CV)
for duplicate RIA analyses was less than 12% in each case.

The compounds of interest were detected in some samples (Tables 3, 4, &
5). NP was detected in 17 of the 23 samples collected (not including replicate
samples) with concentrations ranging from less than detection to 37 pglL. OP
was detected in 13 of the 23 samples analyzed (not including replicate samples)
with concentrations ranging from less than the MDL to 0.67 pglL. NPE was
detected in 16 of the 23 samples analyzed (not including replicate samples) with
concentrations ranging from less than the MDL to 332 pglL. E2 was detected in
16 of the 22 samples analyzed (not including replicate samples) with
concentrations ranging from less than the MDL to 3.7 nglL. EE2 was detected in
8 of the 22 samples analyzed (not including replicate samples) with

concentrations ranging from less than the MDL to 0.76 ng/L.

DISCUSSION

The Trenton Channel of the Detroit River, Michigan, is in an industrialized
and urbanized area with point sources of industrial and municipal waste water
effluents (20, 43, 44). Concentrations of NP and OP at Grosse lle, an island that
forms the Trenton Channel, have been reported to be 0.12 and 0.045 pglL,
respectively, in June, 1994 (20). These concentrations are similar to those

observed in our studies. The Trenton Channel is influenced by these compounds

99

not only in the head waters but by point sources entering the Channel.
Monguagan Creek appears to provide the greatest loading of NP, OP, and NPE
to the Trenton Channel (Table 3). This Creek was previously reported to be a
source for unique tert-alkylphenols such as 4-tert-pentylphenol (45). Many of
these unique alkylphenols were also detected during GCIMS screening in this
study. However, no concentrations are reported here. No trend was observed
from the concentrations of E2 and EE2 detected in the Trenton Channel. The
greatest concentration of E2 and the only detectable concentration of EE2 were
at the Black Lagoon site.

Concentrations of NP, OP, NPE, E2, and EE2 were determined in 4
WWTP effluents in south central Michigan. Samples were also analyzed
upstream from 3 of the WWTP discharges. Concentrations of these compounds
varied greatly among locations (Table 4). NP and E2 were detected in each
effluent sample analyzed. No upstream site contained detectable concentrations
of NP, OP, NPE, or EE2. The only upstream location with a detectable
concentration of E2 was the BV site. This site is located in an agricultural area
with many livestock farms, which may have contributed to the detection of E2,
but we do not have enough data to draw conclusions on the reason for this
occurrence. It should be noted, however, that the BV site uses a lagoon system
for wastewater treatment. Operators of that plant during the October sampling
event indicated that only primary treatment was in operation at that time. This
effect can be seen in the F3 chromatogram (diluted 50x) of the sample collected

during that time period (Figure 4). The NPE oligomer distribution is different from

100

that of the other sites (Figure 4). This is an indication of inefficient treatment.
Removal rates for NP during wastewater treatment have been shown to be 95-
99% (46). Concentrations of NP in municipal wastewater effluents in the US.
have been previously demonstrated to range from less than detection to 4.9 pglL
(46). These concentrations are similar to the concentrations observed in our
studies. In US. rivers, NP has been reported to be undetectable (MDL=0.11
pglL) in 60-75% of samples collected and averaged 0.12 pglL in samples where
it was detected (17).

Concentrations of analytes varied greatly among locations within Lake
Mead (Table 5). The LV Wash site was located at the confluence of VWVTP
effluents from the city of Las Vegas and Clark County Santitation District,
Nevada. This combined effluent stream represents an average of approximately
460 million Uday of tertiary treated wastewater. The LV Bay site is located
approximately 8 km downstream of the LV Wash site and is located in Lake
Mead near the entry point of the LV Wash. The LV Bay site would be expected
to have some mixing of the effluent with Lake Mead water and some dilution from
non-point source ground water entry. Concentrations of NP, OP, and NPE
decreased by nearly 50% from LV Wash to LV Bay, while E2 and EE2
concentrations decreased and increased, respectively. The increase in EE2
could be due to the deconjugation of EE2 conjugates (15, 31). The LV Marina
site lies approximately 2.3 km downstream of the LV Bay site. While only E2 was
detected at this site, it should be noted that previous research by the US. Bureau

of Reclamation has shown an interflow of the dense waste water extending from

101

the Las Vegas Wash to the Hoover Dam (47). This interflow lies on top of the
hypolimnion and its depth changes seasonally. It is likely that our samples
collected at 3 m of depth do not represent the influence of the wastewater
effluent at the LV Marina and Saddle Island sites, thus, offering a possible
explanation of the non-detectable concentrations of the compounds of interest.
Both of these sites lie in the path of the wastewater interflow as it moves towards
the Hoover Dam. Samples collected in September followed a rain event that
resulted in a three-fold increase of the LV Wash average daily flow (personal
communication from Clark County Sanitation District). The addition of large
amounts of storm water in the LV Wash could explain the lesser concentrations
of the compounds of interest from this sampling event. To our knowledge this
paper offers the first reported data for these compounds in the waters of Lake
Mead, Nevada.

This method permits the sensitive detection of NP, OP, NPE, E2 and EE2
without the transportation of large volumes of water. This equates to savings in
time and costs while achieving biologically relevant detection limits. While HPLC
fluorescence provides sensitive detection of E2 and EE2, polar interferences
were encountered in all environmental samples. RIA allowed a very sensitive
detection of E2 and EE2 without a rigorous clean-up. However, RIA is also
sensitive to compounds that are structurally similar to the analyte. Future efforts
will include greater sample volumes and derivatizations to achieve structural

confirmations of E2 and EE2 by GCIMS.

102

Although the sites studied represent only one point in time and were
chosen to demonstrate the utility of the method, rather than the dynamics of the
target compounds, it can be concluded that estrogenic compounds are
widespread contaminants of WWTP effluents. A monitoring program would be
required to better describe the flux and fate of these compounds in the
environment. While the concentrations of these compounds determined in this
study are in the range of those predicted to cause physiological responses in
certain fish (10, 15, 37, 38), no direct link currently exists between the sites

evaluated and a direct cause-effect relationship.

ACKNOWLEDGMENTS

This work was supported by the Chlorine Chemical Council (CCC) of the
Chemical Manufacturer’s Association, the National Institute of Environmental
Health Sciences (NIEHS) Superfund Basic Research Program (ES0491), and the
US. Environmental Protection Agency (EPA) Office of Water Quality (CR
822983-01-0). Technical and material support was provided by Carter Naylor of
Huntsman Corp., Austin TX; Eric Hulsey of Phenomenex Corp.; and Oliver Bell of
3M Corp. We would also like to thank Dr. Matthew Zabik of Michigan State
University for his technical assistance throughout this project. Logistical support
for Lake Mead, NV samples was provided by the Southern Nevada Water

Authority, US. National Park Service, and the US. Bureau of Reclamation.

103

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107

TABLE 1. Recoveries through the analytical procedure (n=3) (%

Recovered : SD)

 

 

Spike cone (nglL) Volume EE2 BP OP NP
240 5 78:6 65:7 76:4 77:7
1128 5 78:4 62:3 72:3 74:5
2374 5 76:5 61:8 74:4 76:5
5934 5 74:3 65:6 73:6 72:8
23737 5 72:7 46:4 68:9 69:4
10761 17 63:7 48:5 67:10 67:9

 

EE2 = ethynylestradiol
BP = butylphenol
OP = octylphenol

NP = nonylphenol

108

TABLE 2. Dection Limits for Selected Xenoestrogens

 

 

Method E2 EE2 BP OP NP NPE
HPLC/ IDL (nglml) NM 5 4 3 13 62
Fluorescence
“ LOQ (nglml) NM 16 13 9 42 208
“ MDL (nglL) NM 4 3 2 11 52
RIA IDL (pg/ml) 427 21 1 NA NA NA NA
“ LOQ (pg/ml) NA NA NA NA NA NA
“ MDL (pg/L) 1 07 53 NA NA NA NA

 

NM = not measured

NA = not applicable

IDL = instrumental detection limit
LOQ = limit of quantitation

MDL = method detection limit

RIA = radioimmunoassay

109

TABLE 3. Concentrations of Selected Xenoestrogens in the Trenton

 

 

Channel
Location Date NP OP NPE E2 EE2
(no/L) (nglL) (nglL) (poll-I (pg/Ll
Lower Transect 8/30/97 993 26 6970 ND ND
Trenton WWTP 8/30/97 479 5 5390 1070 ND
Chemical Plant 8/30/97 862 15 7310 91 1 ND
Power Plant 8/30I97 721 17 8600 ND ND
Elizabeth Park 8/30/97 269 ND 1910 435 ND
Black Lagoon 8/30/97 936 66 8680 1290 359
Monguagan 8/30/97 1 190 81 17800 1060 ND
Creek
Upper Transect 8/30/97 665 23 8330 1280 ND

 

ND = not detectable

NM = not measured

NP = nonylphenol
OP = octylphenol
NPE = nonylphenolethoxylate

E2 = 178-estradiol

EE2 = ethynylestradiol

110

TABLE 4. Concentrations of Selected Xenoestrogens at WWTPs

 

 

Location Date NP OP NPE E2 EE2
(nglL) (HQ/L) (nglL) (Pg/L) (Pg/L)
ER-effluent 5/16/97 806 N D 9020 NM NM
EL-effluent 5/16/97 1020 72 9310 656 248
BV-effluent 5/16/97 22800 249 21800 3230 242
BV-upstream 10l8l97 ND ND ND 711 ND
BV-effluent 10l8l97 37000 673 332000 3660 759
MA-upstream 10l8l97 ND ND ND ND ND
MA-effluent 10/8/97 590 16 4850 905 357
ER-upstream 10l8l97 ND ND ND ND ND
ER-effluent 10l8l97 171 ND ND 477 ND

 

ND = not detectable

NM = not measured

NP = nonylphenol

OP = octylphenol

NPE = nonylphenolethoxylate

E2 = 178-estradiol

EE2 = ethynylestradiol

111

TABLE 5. Concentrations of Selected Xenoestrogens in Lake Mead

 

 

Location Date NP OP NPE E2 EE2
(HI/L) (nglL) (nglL) (Pg/L) (Pg/L)
LV Wash 4/30/97 1 140 43 8990 2670 480
LV Bay 4/30/97 750 27 4850 2210 520
9/5/97 160 ND 3180 188 253
LV Marina 9/5/97 ND ND ND 270 ND
Saddle 4/30/97 ND ND ND ND ND
Island
Callville Bay 9/5/97 ND ND ND ND ND

 

ND = not detectable

NM = not measured

NP = nonylphenol

OP = octylphenol

NPE = nonylphenolethoxylate
E2 = 178-estradiol

EE2 = ethynylestradiol

112

.0 Q

p-t-Octylphenol

HO
Ethinylestradiol

Ho—@—(CQH19)

p-Nonylphenol

17B-Estradiol

Figure 1. Structures of target compounds

113

 

 

5L in situ SPE
extraction

 

 

l

 

 

Vacuum/Solvent
Extraction

 

 

l

 

Concentrate to
1 mL

 

 

 

l

 

 

Normal Phase

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Chromatography
Fractionation
i F1 v F2 V F3
Non-polar Semi-Polar Polar
HAHs and PAHs OP, NP NPE, E2, EE2
Reverse Phase Radioimmunoassay
Fluorescence E2, EE2

Figure 2. Flow diagram of analytical method

 

OP, NP, NPE

 

 

114

 

 

 

BP OP

 

 

 

 

A. NP
B.
C

 

 

Figure 3. Reverse-phase HPLC of F2: (A) spiked standards; (B) Trenton

Channel — Black Lagoon 8/30/97; (C) WWTP - BV effluent 10l8l97

115

EE2

 

 

 

 

 

B.

r—I—II r 1 I I _.'_.
C.

r—I—I—w . r fiT I w—F—r—w—r—u

Figure 4. Reverse-phase HPLC of F3: (A) spiked standards; (B) Trenton-

Channel — Black Lagoon 8/30/97; (C) WWTP — BV effluent 10l8l97

116

 

 

 

Figure 5. Normal-phase HPLC of NPE oligomer distribution: (A) NPE

standard; (B) extracted NPE

117

Chapter 5

TOXICITY IDENTIFICATION AND EVALUATION OF ESTROGENIC AND
DIOXIN-LIKE COMPOUNDS IN WASTEWATER EFFLUENTS

(To be submitted to Environmental Science and Technology)

ABSTRACT

Many xenobiotics enter the aquatic environment from wastewater
discharges. Recent reports have shown that some compounds in wastewater
effluent are estrogenic while others are dioxin-like. The project described here
was designed to screen extractable organic compounds dissolved in water
samples for in vitro estrogen and dioxin-like activity. Samples from 3 municipal
wastewater treatment plants (WWTPs) in south central Michigan (upstream and
effluent); 4 point source locations on the Trenton Channel of the Detroit River,
Michigan; and 5 locations in Lake Mead, Nevada were analyzed. Organic
compounds were extracted from 5 L water samples using solid-phase extraction
disks. Extracts were further fractionated based on polarity. Whole extracts and
fractions were tested for estrogen- and dioxin-like activity using MVLN and H4IIE-
Iuc in vitro bioassays, respectively. No significant dioxin-like activity was
observed. Estrogenic activity was quantitated by comparing the magnitude of
induction elicited by the extract or fraction to the maximum induction caused by

178-estradiol (E2). Several extracts and fractions displayed significant

118

estrogenic activity. In each case, the most polar fraction (F3) was associated
with the greatest estrogenicity. Instrumental analyses and further fractionation
were used to identify compounds associated with estrogenic responses. Results
suggested that the endogenous estrogen, E2, and the synthetic estrogen,
ethynylestradiol (EE2), were the most likely cause of the responses observed.
Alkylphenols (APs) did not appear to contribute significantly to the observed

estrogenicity.

INTRODUCTION

A number of compounds released into the environment by human
activities can modulate endogenous hormone activities and have been termed
endocrine-disrupting compounds (EDCs) (1, 2). EDCs have been defined as
exogenous agents which interfere with the "synthesis, secretion, transport,
binding, action, or elimination of natural hormones in the body that are
responsible for the maintenance of homeostasis, reproduction, development,
and/or behavior" (3). It has been hypothesized that such compounds may elicit a
variety of adverse effects in both humans and wildlife, including promotion of
hormone-dependent cancers, reproductive tract disorders, and reduction in
reproductive fitness (1, 4-10). Endocrine disruption by environmental
contaminants has become a cause for concern (11), and as a result there is an
urgent need to develop methods for monitoring EDCs in the environment (12).

This need is underscored by recent legislation mandating that chemicals and

119

formulations be screened for potential estrogenic activity before they are
manufactured or used in certain processes (Safe Drinking Water Act
Amendments of 1995 - Bill Number S.1316; Food Quality Protection Act of 1996 -
Bill Number PL. 104-170).

Estrogenic and anti-estrogenic compounds are of particular concern.
These compounds have the ability to mimic or block the functions of endogenous
estrogen. Estrogenic effects have been documented in fish exposed to municipal
wastewater treatment plant effluents (13-18). Some marine gastropods exhibited
decreased reproductive success and developed imposex (females with male
genitalia) when exposed to tributyltins (19, 20). Nonylphenol (NP), nonylphenol
polyethoxylates (NPEs), octylphenol (OP), and synthetic and natural steroids are
estrogenic organic constituents of many wastewater effluents (13, 21-24). These
compounds were targeted in this investigation.

Halogenated aromatic hydrocarbons (HAHs) and polycyclic
aromatic hydrocarbons (PAHs) are classes of compounds that have been shown
to exert adverse effects on aquatic organisms. Toxic effects associated with
HAHs and PAHs include mortality, wasting syndrome, hepatotoxicity,
immunotoxiclty, reproductive impairment, and carcinogenicity (25-27). Many of
these effects are the result of an aryl hydrocarbon receptor (AhR)-mediated
mechanism (25). Additionally, HAHs and PAHs also modulate estrogen receptor
(ER)-mediated activities. HAHs such as polychlorinated dibenzo-p-dioxins
(PCDDs), polychlorinated dibenzofurans (PCDFs), and some polychlorinated

biphenyls (PCBs), have been reported to cause anti-estrogenic effects in vitro

120

(28, 29). PAHs have been associated with both estrogenic and anti-estrogenic
responses in vitro (30, 31). As a result, both AhR- and ER-mediated effects of
HAHs and PAHs are of concern when screening surface water samples for
potential (anti)estrogenic compounds.

Although instrumental analyses can be used to detect and quantify
known (anti)estrogenic compounds associated with wastewater treatment plant
(WWT P) effluents, in vitro bioassays provide useful information that can
complement instrumental analyses to provide a more comprehensive
characterization of a sample’s potential to modulate estrogenic responses. In
vitro bioassays measure the ability of a sample to alter a mechanism-specific
biochemical or physiological process. As a result, they provide an indication of
the biological relevance of a sample. Furthermore, this is accomplished in a
manner that integrates the overall potency of complex mixtures. Thus, in vitro
bioassays can account for both unknown compounds and potential non-additive
interactions among compounds. This study involved use of in vitro bioassays to
screen samples for compounds able to modulate ER- or AhR-mediated gene
expression. Successive iterations of sample fractionation coupled with bioassay
and instrumental analysis were employed to identify specific compounds
associated with in vitro responses. Furthermore, bioassay-derived estimates of
estrogenic activity were compared to estimates based on analytical
concentrations of known estrogen agonists and their relative potencies (REPs) in
a mass balance analysis (32, 33). Mass balance analysis was used to generate

hypotheses regarding whether the known composition of a sample could account

121

 

for the response observed or whether unknown compounds or non-additive
interactions between components of the sample influenced the activity. Thus,
instrumental analyses and in vitro bioassays were applied, in a complementary
manner, to screen wastewater outfalls for compounds able to elicit estrogenic or
dioxin-like biological responses and generate hypotheses regarding the probable

cause of the activity observed.

MATERIALS AND METHODS

Sample Collection and Fractionation. A detailed description of the analytical
methodology used in this study was published previously (34). Briefly, 5 L water
samples were extracted at each field site using solid-phase extraction (SPE)
EmporeTM disks. Organic extracts from these SPE disks were separated into 3
fractions based on polarity using normal-phase high-pressure liquid
chromatography (NP-HPLC) (Figure 1). In some cases, EDCs of interest were
isolated further from F2 and F3 by fractionating these again with reverse-phase
HPLC (RP-HPLC), with fractions collected approximately every 3 min (Figure 2).
Chromatographic conditions for RP-HPLC fractionation were described
previously (34). NP-HPLC and RP-HPLC separations were accomplished using

silica and Cu; analytical columns, respectively.

Cell Culture and Bioassay. An MCF-7 human breast carcinoma cell line, stably

transfected with an estrogen receptor-controlled luciferase reporter gene

122

construct (MVLN or MCF-7-luc cells), was developed and characterized by Dr.
M.D. Pons, Institut National de la Sante et de la Recherche Medicale (35).
MVLN cells were cultured in 75-cm2 disposable polyethylene tissue culture flasks
(Corning, Corning NY, USA) containing 20-25 mL of Dulbecco’s Modified Eagle
Medium (DMEM) with Hams F-12 nutrient mixture (Sigma D-2906; St Louis, MO,
USA) supplemented with 10% defined fetal bovine serum (Hyclone, Logan UT,
USA), 27.3 I.U. insulin (Sigma I-1882) IL, and 1.0 mM sodium pyruvate (Sigma).
H4llE-Iuc cells are rat hepatoma cells containing a Iuciferase reporter gene under
control of dioxin responsive enhancers (DREs) (36). They were cultured in 100-
mm disposable petri plates (Corning) containing 12 mL Dulbecco’s Modified
Eagle Medium (DMEM; Sigma D2902). Cells were incubated at 37 °C in a
humidified 95:5 air:COz atmosphere and were passaged when plates became
confluent (generally every 5 d). New cultures were started from frozen stocks
after 30 passages.

In preparation for bioassay, cells were trypsinized from flasks or
plates containing 80-100% confluent cell growths. The number of cells per mL
was determined microscopically by use of a hemacytometer. MVLN cells were
diluted in stripped media [DMEM with Hams F-12 nutrient mixture, supplemented
with 10% dextran-coated charcoal filtered fetal bovine serum (Hyclone), 27.3 I.U.
insulin (Sigma I-1882) IL, and 1.0 mM sodium pyruvate (Sigma)] to a
concentration of approximately 1.5 x 105 cells/mL. H4llE-luc cells were diluted to
the same concentration in DMEM. Cells were seeded into the 60 interior wells of

96-well flat bottom microplates (Packard Instruments 6005181; Meriden CT,

123

USA) at 125 pL per well (15-20,000 cells per well) using a repeating pipettor. To
assure homogeneity, the cell solution was vortexed continuously during seeding.
The 36 exterior wells of each microplate were filled with 125 pL of media. Cells
were dosed after an overnight incubation to allow for cell attachment. Extracts or
fractions were dissolved in the appropriate culture media (stripped media for
MVLN, DMEM for H4llE-luc) to yield a final concentration of 1.0% extract. A 3-
fold dilution of each extract or fraction was also prepared yielding a concentration
of 0.33% extract. Test wells were dosed with 125 pL 1.0% or 0.33% extract in
media to yield final in-well concentrations of 0.50% and 0.165% extract. Solvent
control wells were dosed with 125 pL of media spiked with 1.0% of the
appropriate solvent to yield a final in-well concentration of 0.50% solvent. Blank
wells received 125 pL of the appropriate media. Each plate tested included a
minimum of three solvent control wells, three blank wells, and three replicates of
each fraction tested (at both 0.50% and 0.165% levels). Dosed cells were
exposed for 72 h at standard incubation conditions.

Each test plate was inspected visually and differences in cell
numbers and condition relative to control wells and conditions normally observed
during routine culturing were noted for each well. Culture medium was then
removed and each well was rinsed twice with phosphate buffered saline (PBS)
supplemented with 1.0 mM Ca2+ and Mg2+ using an eight channel vacuum
manifold. Plates were inspected for cell loss during washing. Following
inspection, 75 pL PBS supplemented with Ca2+ and Mg2+ was added to each

well, followed by 75 pL Luc-liteTM reagent (Packard Instruments). Each plate was

124

 

incubated for 10 min at 30 °C then scanned with an ML 3000 microplate reading
luminometer (Dynatech Laboratories, Chantilly, VA, USA). Following the
luminomter scan, 125 pL 1.08 mM fluorescamine (Sigma) in acetonitrile (ACN)
was added to each well and plates were assayed for protein after a 15 min
incubation at room temperature (37). Plates were scanned using a Cytofluor
2300 (excitation 400 nm, emission 460 nm), and responses were compared to a
standard curve consisting of 6 concentrations of bovine serum albumin (BSA)
(Sigma) ranging from 50-1.5 pg per well.

All data were collected electronically and imported into a
spreadsheet (Excel 7.0, Microsoft Inc., Seattle, WA) for data analysis. Protein
content per well was calculated by regression against the BSA standard curve.
Protein data were used as an index of cell number to detect outliers that were not
apparent by visual inspection. Relative luminescence units (RLU) were not
adjusted for protein. Sample responses in RLU were expressed as a percentage
of the mean maximum response observed for E2 or 2,3,7,8-tetrachlorodibenzo-p-
dioxin (T CDD) standard curves developed on the same day (% TCDD-max and
% E2-max, respectively). The greatest response of the two extract dilutions was
reported. However, for each significant response, the greatest response came
from the greater extract concentration (0.5% in the well). In addition, for some
samples, the dose-response relationship was used to determine the total
estrogenic or dioxin-like activity of the samples. Activities derived from the
bioassay were designated E2-EQ and TCDD-EQ for estrogenic and dioxin-like

activities, respectively.

125

 

In addition to determining the total activity of extracts by the above-
mentioned bioanalytical techniques, for some samples the total activity was
estimated by correcting the concentrations of individual compounds present in
the extract for their relative potencies. This was accomplished by multiplying the
concentration of each compound by the corresponding toxic equivalency factor
(TEF) or relative potency factor (REP) and summing the products. The
calculated concentrations were referred to as EEQ and TEQ, respectively. A
mass balance of activities determined by bioanalytical techniques and those
calculated from the measured concentrations of individual compounds could be
calculated. This was done to determine whether there were unidentified
compounds contributing to the total activities measured in the bioassays. To
make an accurate comparison, the potential for antagonistic and synergistic
interactions needed to be addressed. This was done by the fine fractionation of
samples, by which compounds known to be antagonistic to the measurement of

EEO were separated from the active compounds.

RESULTS AND DISCUSSION

Dioxin-Iike Activity. None of the extracts or fractions tested in this study
induced a significant response in the H4llE-luc bioassay (Figure 3). This was not
surprising, because water concentrations of dioxin-like compounds were
expected to be very small. No PCBs or PCDDs/DFs were detected in any of the

extracts analyzed in this study. PAHs were detected at each site; however,

126

summed concentrations did not exceed 40 ng/L at any site. Of the OC pesticides
analyzed, only hexachlorocyclohexanes (HCHs) were observed and only in the
Las Vegas Bay sites. HCH concentrations were less than 20 nglL in all samples
that contained detectable concentrations. The method detection limit (MDL) for
the H4llE-luc bioassay was around 0.2 frnol TCDD-EQlwell (32). Assuming there
were no antagonistic interactions among components of the sample, the H4IIE-
luc results suggest that the water samples analyzed contained less than 10 pg

TCDD-EQ/L.

Estrogen-like Activity. Whole extracts and corresponding fractions were
screened for estrogenic activity using the MVLN in vitro bioassay. Non-polar
compounds such as PAHs, PCBs, and most organochlorine (OC) pesticides, if
present, would have been contained in F1 (Figure 1) (34). Certain OC pesticides
and PAHs such as chrysene, benz[a]anthracene, and benzo[a]pyrene have been
reported to cause weak estrogenic responses in vitro (31, 38). None of the F1
samples; however, induced a significant response in the MVLN assay (Figures 4
- 7). Based on the MDL for E2 in the MVLN assay, concentrations of estrogenic
compounds present in F1 contributed less than 110 pg E2-EQIL. These results
support the conclusion that concentrations of non-polar estrogenic compounds in
the surface waters and effluents examined were small.

Weak estrogen agonists such as NP and OP were present in F2
(Figure 1). Concentrations of E2-EQ in F2 were less than the MDL, despite the

confirmed presence of NP and OP (Figures 4 — 7). These results suggest that

127

the compounds present in F2 contributed less than 110 pg EEQ/L. Estrogenic
potencies of NP and OP, relative to E2, for luciferase induction in MVLN cells
have been reported to be 1.25 x 10'5 and 1.9 x 10'5 for NP and OP, respectively
(33). Concentrations of NP and OP present in the sample extracts (Tables 1 & 2)
were used to calculate EEQ. NP and OP were estimated to contribute less than
15 pg E2-EQ/L for fifteen of the sixteen samples tested. Thus, the lack of
significant induction of the MVLN cells was consistent with the known
concentrations of estrogenic alkylphenols present in the F2 samples. The BV-
effluent sample extract contained approximately 380 pg EEQ/L, which would
correspond to a dose of approximately 8.5 fmol E2-EQ/well in the MVLN
bioassay. Based on regression against an E2 standard curve, this dose could
have elicited a response as great as 53% E2-max. The F2 extract of the BV-
effluent; however, failed to induce a significant MVLN response (Figure 4). This
suggests that F2 of the BV—effluent sample might have contained interfering
compounds that suppressed the estrogenic activity of NP and OP. Therefore,
the F2 extract of the BV-effluent was further fractionated and E2-EQ measured
(Figure 8). No significant estrogen-like responses were observed in this fine
fractionation. The same fine fractionation was applied to the F2 extract from
Black Lagoon (Figure 5). Once again, no significant estrogen-like activity was
observed. In general, however, concentrations of E2-EQ in F2 extracts were in
agreement with the EEO based on the known concentrations and relative

potencies of these compounds.

128

F3 extracts were the most active of the fractions tested. Six of the
sixteen F3 samples elicited significant estrogenic activity in the MVLN bioassay
(Figures 4 — 7). The greatest magnitudes of response for F3 extracts,
approximately 80% E2-max, were observed for samples collected from the LV
Wash and LV Bay in April 1997 (Figure 6). However, samples from these sites
collected in September 1997 did not contain significant concentrations of E2-EQ
(Figure 7). The samples collected in September 1997 were after a large storm
event, which diluted the wastewater entering the LV Wash and LV Bay (34). The
estrogenic potency of EE2 relative to E2 for Iuciferase induction in the MVLN
assay was previously reported to be approximately 0.1 (33). Based on the
concentrations of E2 and EE2, and their corresponding relative potencies,
samples collected from LV Wash and LV Bay in April 1997 were estimated to
contain 2180 and 1810 pg E2/EE2-derived EEQIL, respectively. These
concentrations should have yielded doses of approximately 50 and 41 frnol E2-
EQ/well in the MVLN bioassay. Based on regression against an E2 standard
curve, such doses would be expected to yield responses of approximately 92%
and 88% E2-max, respectively. Given the potential range of uncertainty
associated with the bioassay responses and the relative potency of EE2, the
responses observed for the samples collected from LV Wash and LV Bay in April
1997 were not significantly different from predicted responses. Thus, the known
E2 and EE2 composition of F3 of the samples collected from LV Wash and LV
Bay appeared to account for all the estrogenic activity observed. Further

fractionation of the F3 extracts from LV Wash and LV Bay revealed that all of the

129

 

estrogenicity was associated with the fine fractions (F Fs) 3 and 4, which equates
roughly to the retention times of E2 and EE2 (Figures 10 & 11). FFs 3 and 4
from the F3 extract of the LV Wash sample were collected, combined, and
fractionated again by RP-HPLC using a slower flow rate and solvent gradient to
separate E2 and EE2 (Figure 12). The estrogen-like bioactivity was isolated to
the fractions where E2 and EE2 elute, and the magnitude of induction was similar
to that predicted from the E2-EQs measured in the MVLN bioassay (Tables 1 &
2). The greater estrogenic activity of the water extracts from LV Wash and LV
Bay was most likely due to the fact that the WWTPs discharging into the Las
Vegas Wash represent a larger population of humans.

Significant estrogenic activity also was associated with F3 extracts of
water from three locations on the Trenton Channel of the Detroit River (B.
Lagoon, Chem, and WWTP) (Figure 5), and BV—effluent (Figure 4).
Concentrations of E2-EQ in extracts from B. Lagoon, Chem., WWTP, and BV-
effluent contained 1060, 730, 850, and 2980 pg E2/EE2-derived EEQ/L,
respectively. Based on regression against an E2 standard curve, these
concentrations of E2-EQ were predicted to yield responses of 76%, 67%, 71%,
and 99% E2-max, respectively. Observed MVLN responses for these F3
samples were; however, less than predicted (Figures 4 & 5). Further
fractionation and bioanalysis of F3 extracts from BV-effluent and B. Lagoon
indicated that all observed estrogenicity is contained in FFs 3 and 4. However,
the magnitude of induction of the FFs was significantly different from that of the

corresponding total F3 extract. This suggests that interfering and/or anti-

130

estrogenic compounds present in F3 might have modulated the activity of the
known estrogen agonists.

The potential presence of interfering and/or anti-estrogenic (estrogen
antagonists) compounds also was suggested by the lack of significant response
for several samples. Based on concentrations of E2 and EE2, six additional F3
samples should have elicited significant responses in the MVLN bioassay.
Concentrations of EEQs calculated from concentrations of EE2 and E2 in
samples collected from LV Bay (Sept. 1997), LV Marina, M. Creek, BV-upstream,
MA-effluent, and ER-effluent were estimated to range from 170-850 pg EEQIL.
Regression against an E2 standard curve would indicate, predicted responses of
35-71% E2-max in the MVLN assay for these samples. Thus, the responses
were less than predicted for these samples. The reason for this observation is
unknown at this time.

Concentrations of EEO in unfractionated (total) extracts were similar to
those for F3 fractions. Samples for which F3 elicited a significant response also
elicited a significant whole extract response (Figures 4 - 6). In five of the six
cases, the whole extract response was slightly lesser than the response elicited
by F3. This suggests that F1 and F2 may have contained some interfering or
anti-estrogenic substances that modulated the activity of the known ER agonists
in the samples. However, no significant anti-estrogenic responses were
observed (Figures 4 — 6). Because the decreases were slight; however, the
results suggest that the bulk of potential interfering (antagonistic) compounds

were present in F3. The extract of BV-effluent was the only sample for which the

131

whole-extract response was greater than the corresponding F3 response. It was
also the only sample for which the concentrations of NP and OP in F2 were
predicted to yield significant estrogenic activity. NP and OP accounted for 11%
of the total E2-EQ calculated for the BV-effluent extract. Thus, although F2 of
the BV-effluent sample failed to elicit a significant response, NP and OP may
have contributed to the response of the whole extract, such that the total extract
response was greater than the F3 response. In general, however, NP and OP
accounted for less than 1% of the total concentrations of sample E2-EQs present

in samples.

SUMMARY

The mass balance calculations based on instrumental analyses and
bioassay-directed fractionation support the conclusion that E2 and EE2 were the
dominant environmental estrogens in the samples. Also, interfering or anti-
estrogenic compounds present in samples (predominantly in F3) may have acted
to mask or dampen the activity of the known ER agonists in the MVLN bioassay.
All observed responses in the MVLN bioassay were either less than or
approximately equal to responses predicted based on the concentrations and
relative potencies of known ER agonists measured by instrumental techniques.
NP and OP generally contributed less than 1% of the total E2-EQs. Furthermore,
sample fractions containing NP and OP did not elicit significant activity. For most

samples, fractions containing E2 and EE2 elicited responses slightly greater than

132

the responses of the corresponding whole extracts. Thus, among the ER
agonists detected in the samples, E2 and EE2 appear to be responsible for the
bulk of the activity. The fact that observed responses were generally lower than
predicted suggests the presence of interfering compounds. Concentrations of
E2-EQ in whole extracts were only slightly less than those from F3 extracts. This
suggests that the interfering compounds may have been present. Because there
were few instances where the concentrations of E2-EQ were greater than the
concentrations of EEO in any of the fractions or in the subsequent fine fractions,
the known composition can account for the magnitude of response observed. It
is unlikely that there were additional ER-active compounds of significant
concentration that were not identified.

There are insufficient data to explain differences in bioactivity among the
locations investigated. It should be noted that samples from the Trenton Channel
of the Detroit River in Michigan received less effluent from municipal WWl'Ps
relative to the volume of the receiving water compared to the other sites. Also,
the population served by the WWI'Ps varied greatly among sites. Further
investigations would be necessary to determine the actual loading of bioactive
compounds as a function of population density. Without knowing the available
fractions and bioaccumulation potential of the various compounds and dose-
response relationships for target species, it is not possible to predict the potential

effects of the observed concentrations of environmental estrogens on biota.

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Table 1. Extract Concentrations

 

 

 

Location Date NP
Lake Mead
LV Wash 4/30/97 4560
LV Bay 4/30/97 3000
9/5/97 640
LV Marina 9/5/97 ND
Saddle Island 4/30/97 ND
Callville Bay 9/5/97 ND
Trenton
Channel
WWTP 8l30/97 1916
Chem. 8l30/97 3450
B. Lagoon 8l30/97 3740
M. Creek 8l30/97 4740
WWTPs
BV-upstream 1 0/8/97 ND
BV-effluent 10l8l97 148000
MA-upstream 1 0/8/97 N D
MA-effluent 1 0/8/97 2065
ER-upstream 1 0/8/97 ND
ER-effluent 1 0/8/97 680

OP

172
108
ND
ND
ND
ND

20
60
264
324

ND
2350
ND
54
ND
ND

NPE

35960
19400
12710
ND
ND
ND

21600
29200
34700
71260

ND

1 160000
ND
19400
ND

ND

 

ND = not detectable

NA = not applicable

NP = nonylphenol

OP = octylphenol

NPE = nonylphenolethoxylate

E2 = 178-estradiol

EE2 = ethynylestradiol

138

E2

1 0700

8840
752
1080
ND
ND

4260
3640
51 80
4244

2500

14600

ND
3620
ND
1 900

EE2

(ng/mL) (ng/mL) (ng/mL) (pg/mL) (pg/mL)

1920
2080
1012
ND
ND
ND

ND
ND
1440
ND

ND
3036
ND
1430
ND
ND

 

Table 2. E2-EQs and Predicted Responses

 

Location Date NPIOP- Predicted E2IEE2- Predicted
E2-EQa Responseb E2-EQ° Responsed
(PS/L) (0%)
Lake Mead
LV Wash 4/30/97 12. 1 0 2180 92
LV Bay 4/30/97 7.91 0 1810 88
9/5/97 1.60 0 171 35
LV Marina 9/5/97 NA NA 216 41
Saddle Island 4/30/97 NA NA NA NA
Callville Bay 9/5/97 NA NA NA NA
Trenton
Channel
WWTP 8/30/97 4.87 0 852. 71
Chem. 8l30/97 8.85 0 728 68
B. Lagoon 8130/97 10.4 0 1 060 76
M. Creek 8l30/97 13.1 0 849 71
WWTPs
BV-effluent 1 0/8/97 379 53 2980 99
BV-upstream 1 0/8/97 NA NA 500 59
MA-upstream 1 0/8/97 NA NA NA NA
MA-effluent 1 0/8/97 5.41 0 753 68
ER-upstream 1 0/8/97 NA NA NA NA
ER-effluent 1 0I8/97 1 .70 0 380 53

 

NA = not applicable

3 Nonylphenol and octylphenol-derived 17-B-estradiol equivalents. NPIOP-EZ-EQ = (NPNW.

potency (REP)x Npmnmum) '1' (OPREP X opooncentratlonIo NPREP = 1.25 X 10.5. OPREP = 1.9 X 10.5. A
REP estimate was not available for NPE, therefore it was not considered when deriving E2-EQ
estimates. Values presented were back-calculated to water concentrations (pg E2-EQI L H20).

b MVLN bioassay response magnitudes predicted based on regression of NPIOP-derived E2-EQ
against a 17-B-estradiol standard curve. Units are %-E2-max.

° Estradiol and ethynylestradiol-derived 17-B-estradiol equivalents. E2IEE2-E2-EQ = (EZREp x

EZWO") + (EEZREP X EEanmmjon). EZREP = 1.0. EEZREP = 0..10 values presented were
back-calculated to water concentrations (pg E2-EQ/ L H20).

d MVLN bioassay response magnitudes predicted based on regression of E2IEE2-derived E2-
EQ against a 17-B-estradiol standard curve. Units are %-E2.-max.

Note: Total E2-EQ = NPIOP-EZ-EQ + E2IEE2-E2-EQ. Predicted bioassay response magnitudes
are not additive.

139

 

 

 

5L in situ SPE
extraction

 

 

l

 

 

Vacuum/Solvent
Extraction

 

 

l

 

 

Concentrate to
1 mL

 

 

l

 

 

Normal-phase
Fractionation

 

 

 

in

F2

 

t

 

 

 

Non-polar
PAHs and HAHs

 

 

Semi-Polar
OP and NP

 

 

 

 

 

F3

 

 

Polar
NPE, E2, and EE2

 

 

 

 

 

l

 

 

Reverse-phase
Fractionation

 

 

Figure 1. Flow diagram of analytical method

140

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

1 2 3 4 5 6 7 8 9 10
OP
F2
NP
5152
F3 52 NPE1
I17 3. .. .1,o .fi .. ..20.. .. -

Tlme (min)

Figure 2. Reverse-phase HPLC fine fractionation

141

% TCDD Max

100.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

I Total

80.0 _
. F1

50.0 CI F2 _
5 F3

40.0

20.0

-20.

MA LV-Bay B. Lagoon Blank

Site

Figure 3. Dioxin-Iike bioactivity (various sites): Significance bars

represent 3x SD of the background signal

142

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

80.0 I roan a
I F1
60.0 CI F2 _
I F3
40.0
20.0
0.0
-20.0

MA-up MA BV-up ev ER-up ER Blank

Site

Figure 4. Estrogen-like bioactivity — Michigan WWTPs (October 1997):

Significance bars represent 3x SD of the background signal

143

% E2 Max

 

1 00.0

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

80.0 I roar —
fl F1

60.0 CI F2 F—
m F3

40.0

20.0

0.0

-20.0

B.Lagoon Chem. M.Creek WWTP Blank
Site

Figure 5. Estrogen-like bioactivity — Trenton Channel (August 1997):

Significance bars represent 3x SD of the background signal

144

% E2 Max

 

100.0

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

80.0
60.0
40.0
20.0
0.0
-20.0
LV LV Saddle Blank
Wash Bay Island
* Site

Figure 6. Estrogen-like bioactivity — Lake Mead (April 1997): Significance

bars represent 3x SD of the background signal

145

 

 

 

 

 

 

 

 

80.0 I Total ’—
n F1
50.0 5 F2 _
5 F3
40.0

 

 

 

 

 

 

LV LV Callville Blank
Bay Marina Bay

Site

Figure 7. Estrogen-like bioactivity - Lake Mead (September 1997):

Significance bars represent 3x SD of the background signal

146

%E2Max
d
e 8 3 S 8 8

 

iv
a

Figure 8. Fine fractionation and corresponding estrogen-like bioactivity of

WWTP BV-effluent F2 extract

147

°/o E2 Max

nb

OP

0”
90

N15
00

 

r'o
c

Figure 9. Fine fractionation and corresponding estrogen-like bioactivity of

Black Lagoon, Trenton Channel, F2 extract

148

% E2 Max

 

Figure 10. Fine fractionation and corresponding estrogen-like bioactivity

of LV Wash, Lake Mead (April 1997), F3 extract

149

% E2 Max

 

NPE

 

 

—

10

 

100

 

 

 

40

 

 

20

 

 

Figure 11. Fine fractionation and corresponding estrogen-like bioactivity

 

 

 

 

 

l

 

 

 

i

 

 

 

 

 

of LV Bay, Lake Mead (April 1997), F3 extract

150

 

 

% E2 Max

EE2

E2

100

I
N-tsoaco
Booooo

 

Figure 12. Further fractionation and corresponding estrogen-like

bioactivity of LV Wash, Lake Mead (April 1997), F3 extract

151

°/o E2 Max

 

Figure 13. Fine fractionation and corresponding estrogen-like bioactivity

of VWVTP BV-effluent, F3 extract

152

 

% E2 Max

 

Figure 14. Fine fractionation and corresponding estrogen-like bioactivity

of Black Lagoon, Trenton Channel, F3 extract

153

Chapter 6

BIOACTIVE ORGANIC COMPOUNDS IN THE WATERS OF LAKE MEAD,
NEVADA, USA

(To be submitted to Environmental Science and Technology)

ABSTRACT

Several recent reports have indicated a flux of organic compounds flowing
into the Boulder Basin of Lake Mead, Nevada from the Las Vegas Wash. The
primary objective of this study was to screen for a wide range of organic
contaminants from 5 locations in Lake Mead between May 1998 and June 1999.
Certain pesticides, polychlorinated biphenyls (PCBs), alkylphenols, polycyclic
aromatic hydrocarbons (PAHs), and steroids were target compounds. Mass
spectrometry was utilized to screen for previously unreported compounds.
Samples of 100 L of filtered water were collected in Teflon-lined containers and
were extracted using large-scale solid-phase extraction. Following a series of
elution and extraction procedures, resulting extracts were fractionated into 3
polarity classes. Each fraction was analyzed by gas chromatography with mass
selective detection (GCIMS). DDT, DDE, and DDD (collectively, DDTs) and four
isomers of hexachlorocyclohexane (HCH) were detected at some sites, with
concentrations ranging from less than detection to 28 nglL. Octylphenol,

nonylphenol, nonylphenol monoethoxylate, and nonylphenol diethoxylate were

154

 

detected in some samples, with concentrations ranging from less than detection
to 1500 ngIL. Caffeine, nicotine, oxybenzone, N,N-diethyl-meta-toluamide
(DEET), phosphate-based flame-retardants, and several pharmaceuticals were
detected in some samples with concentrations ranging from less than detection
to 360 nglL. The identities of several of these previously unreported compounds
were verified using high-resolution mass spectrometry. PAHs were detected at
each site and did not vary significantly among the sites in concentration or in
pattern. Total PAH concentrations ranged from 5 to 78 ng/L. Polychlorinated
biphenyls, triazine herbicides, and steroids were not detected in any of the

samples.

INTRODUCTION

With approximately 36.7 x 109 m3 of storage capacity, Lake Mead is the
largest reservoir on the Colorado River system and was formed with the
completion of Hoover Dam in 1935. The lake encompasses approximately 593
km2 of surface area and extends 185 km east from Hoover Dam, up the old
course of the Colorado River. The majority of flow into Lake Mead (97%) comes
from the Colorado River. Approximately 1.5% of the flow originates from the
Virgin and Muddy Rivers, which discharge into the Overton Arm of Lake Mead,
and 1.5% comes from the Las Vegas Wash, which discharges into the Boulder
Basin (Figure 1). Approximately 22 million people depend on water from Lake

Mead and the lower Colorado River for domestic and agricultural usage (1).

155

Las Vegas Wash is a 19 km channel that drains the entire 4100 km2 Las
Vegas Valley Hydrographic Basin. Sources of flow to the wash include storm
water, urban runoff, shallow groundwater, and tertiary treated wastewater. The
average daily flow in Las Vegas Wash is approximately 580 million L per day
(excluding storm water). The drinking water intakes for Southern Nevada lie
approximately 10 km down stream of the Las Vegas Wash confluence. The Las
Vegas Wash provides significant nutrient and particulate loading into the Las
Vegas Bay (1-3). This increased nutrient loading results in frequent
eutrophication of the Las Vegas Bay. Secchi readings of less than 0.5 m and
chlorophyll a concentrations of greater than 300 mg/m3 have been reported (1).
Water from the Las Vegas Wash is warmer and has greater conductivity,
turbidity, and suspended and dissolved solids than main body Lake Mead water.
Despite being warmer, Las Vegas Wash water has a greater density, which
results in an underflow that extends until density equilibrium is met. This
intrusion then can form an interflow above the hypolimnion. At certain times of
the year, the intrusion of the Las Vegas Wash can be detected at various depths
over the 16 km to Hoover Dam (1).

In 1996, the US. Geological Survey (USGS) reported endocrine
disruptive type effects in feral carp from the Las Vegas Wash and Bay. These
carp had significantly different sex steroid and plasma vitellogenin levels than
carp collected from Callville Bay (reference site) in Lake Mead (4). Furthermore,
concentrations of some synthetic organic chemicals were found to be greater in

water, sediment, and fish tissues from the Las Vegas Wash and Bay as

156

 

compared to Callville Bay (4). The presence of alkylphenols and natural and
synthetic estrogen also has been reported in the Las Vegas Wash and Bay (5).
In addition, in vitro bioassays were used to screen extracts of Lake Mead water
samples for estrogenicity. This study indicated that natural and synthetic
estrogen were the most potent xenoestrogens present in Las Vegas Wash and
Bay water extracts (6). These reports indicate an urgent need to identify organic
compounds present in the Las Vegas Bay that may have an impact on the
aquatic organisms which reside there.

In this report we describe a novel method for the sensitive detection
of several classes of organic compounds. The basic methodology is a modified
version of that reported previously (5). Organic compounds from a 100 L sample
of water were concentrated using solid-phase extraction (Figure 2). Compounds
reported previously and known to induce endocrine disruptive type effects in
certain aquatic organisms were targeted. These include organochlorine (OC)
pesticides, polycyclic aromatic hydrocarbons (PAHs), polychlorinated biphenyls
(PCBs), atrazine, simazine, alkylphenols (APs), 178-estradiol (E2), and
ethynylestradiol (EE2). Mass spectrometry was used to quantify and identify
these target compounds and to screen for previously unreported organic
compounds. Gas chromatography with high-resolution mass spectrometry
(GCIHRMS) was used for structural verification for some compounds never
before reported in US. waters. Mass Peak Profiling from Selected Ion Recording
Data (MPPSIRD) and a Profile Generation Model (PGM) were used in

conjunction with GC/HRMS to identify of trace concentrations of organic

157

compounds, including pharmaceuticals, in these complex extracts. The data
presented for novel compounds is semi-quantitative due to the methodologies

applied.

EXPERIMENTAL SECTION

Standards and Reagents. All standards and reagents used were of the highest
purity commercially available. Standards (approximately 98% pure) of p-
nonylphenol (NP) and p-tert octylphenol (OP) were obtained from Schenectady
International (Freeport, TX). Standards of nonylphenol monoethoxylate (NPE1)
and nonylphenol diethoxylate (NPEZ) were obtained from Huntsman Corporation
(Austin, TX). Organochlorine (OC) pesticides, polycyclic aromatic hydrocarbons
(PAHs), polychlorinated biphenyls (PCBs), and triazine herbicide standards were
obtained from AccuStandard Inc. (New Haven, CT). All pharmaceutical and
steroid standards were obtained from Sigma Chemical Company (St. Louis, MO)
with the exception of fluoexetine (Prozacm), which was obtained from US.
Pharrnacopeia (Rockville, MD). Bisphenol A (BPA), DEET, and 2-hydroxy-4-
methoxybenzophenone (oxybenzone) were obtained from Aldrich (Milwaukee,
WI). Pesticide residue grade methanol (MeOH), hexane, dichloromethane
(DCM), iso-octane, and acetone were obtained from Burdick and Jackson
(Muskegon, MI) or Sigma-Aldrich (Milwaukee, WI). Solvents were tested for
interfering compounds by examining 150-fold concentrated solvents using gas

chromatography with mass selective detection (GCIMS). Reagent water was first

158

purified by reverse osmosis (RO) followed by Nanopurem (Barnstead, Dubuque,
IA) treatment. Glass fiber filters, fine grade (GF/F) with 0.7 pm nominal pore size
(293 mm, Whatman, Hillsboro, OR), were heated at 450 °C for 4 h in aluminum
foil prior to use. A 25% salt solution was made by dissolving 250 g sodium
chloride (Baker, Phillipsburg, NJ) in 1 L reagent grade water and liquid-liquid
extracted 3x with DCM to remove contaminants. Anhydrous granular sodium
sulfate (EM Science, Gibbstown, NJ) was heated at 550 °C, overnight then

stored at 125 °C until day of use.

Sample Collection. Water samples were collected between May 1998 and June
1999 from 5 locations in Lake Mead, Nevada (Figure 1). Samples were taken
from 3 m below the surface except at the LV1 site where samples also were
collected at the zone of greatest conductivity as determined using an H20
SondeTM unit connected to a Surveyor 3TM data recorder (Hydrolab Corp., Austin,
TX).

Samples were pumped through one to five 30 cm glass fiber filters
(GF/Fs) in a 293 mm stainless steel parallel filtration system (Pentaplate) using
fluoropolymer (PFA) tubing (3I8” o.d.) and a peristaltic pump fitted with platinum-
cured silicone peristaltic tubing at the field site. The Pentaplate was cleaned with
water and solvents before and after each sample was filtered. At each sampling
site, prior to collecting the samples, the apparatus was purged with sample water
for 10 min by bypassing the filter holder. At sites with predicted concentration

gradients, samples were taken sequentially from the least to greatest

159

concentrations. The initial 4 L portion of filtered sample was discarded, and the
remaining sample was collected in a custom-built clean TeflonTM bag (Berghof
America, Concorde, CA) lining a 113 L high-density polyethylene tank. Once the
TeflonTM bag was filled, the tank was transported to the laboratory.

Field blanks were taken with each sampling event by passing 100 L
of NanopureTM water through the system as previously described. A column
blank was processed with each batch of columns extracted by extracting a
prepared SPE column through which no sample or blank water had passed.
Laboratory blanks were performed on each day of extractions, and instrument
blanks were conducted daily. Flow meter calibrations were verified prior to each
sampling trip by comparing the reported totalizer output to the measured volume

of water exiting the meter.

Extraction. Immediately upon arrival in the laboratory, solid-phase extraction
(SPE) began. Samples were extracted by pumping the samples through a
stainless steel preparative high pressure liquid chromatography (HPLC) column
(250 x 21.2 mm, Phenomenex, Torrance, CA). The HPLC column was dry-
packed with approximately 55 g of pre-cleaned Bond Elut ENVTM (125 pm
particle size, Varian, Harbor City, CA) SPE resin. These assembled SPE
columns were additionally cleaned by pumping sequentially, for one hour each,
acetone, 50% DCM in hexane, methanol, and water through the column at 5

mUmin. Columns were stored at 4 °C until use.

160

Samples were pumped through the SPE columns using a metering pump
(Fluid Metering, Syosset, NY) at a flow rate of 0.1 lein until 100 L total volume
had been extracted (approximately 16.7 h). An electronic flow meter (model 102-
6TP, McMillan, Georgetown, TX), calibrated to i 1% accuracy with National
Institute of Standards and Technology (NIST) traceable standards, was installed
downstream of the SPE column. This digital rate meter and totalizer was used to
monitor flow rate and to record the total volume. Once the desired sample
volume had been reached, the column was removed, capped, and stored at 4 °C
until elution.
The SPE column was eluted using a quaternary HPLC pump
(Perkin Elmer, Series 410, Nonrvalk, CT) at 5 mUmin sequentially for 10 min with
acetone, 40 min with 50% DCM in hexane, and an additional 10 min with
acetone. The effluent of the SPE column was collected in a 1 L separatory
funnel containing 100 mL 25% sodium chloride solution, and the resulting mixture
was liquid-liquid extracted for 5 min, and the aqueous layer (bottom) was
collected into a 1 L separatory funnel. The original aqueous layer was then
extracted twice more with 50 mL DCM and the organic layer (bottom) combined
with the original organic layer. The combined organic layers were then passed at
a rate of 1 drop/sec through a funnel containing 100 g anhydrous sodium sulfate
and collected in a RapidVapTM tube (Labconco, Kansas City, MO). Extracts were
then concentrated to approximately 3 mL in a RapidVapTM N2 evaporation system
(Labconco, Kansas City, M0) at 35 °C and vortex speed of 30. The sample was

quantitatively transferred to a 15 mL centrifuge tube and concentrated to 1 mL

161

under a gentle stream of nitrogen at 30 °C. The extract was diluted to 4 mL with

iso-octane, vortexed, and concentrated to 2.0 mL under nitrogen.

Silica Gel Fractionation. One mL of iso-octane extract was fractionated using a
gravimetric silica gel column prepared by placing a small plug of DCM extracted
glass wool in a glass chromatography column followed by 10 g of acid-activated
granular copper (10 - 40 mesh, Aldrich, Milwaukee, WI), 1 cm of sodium sulfate,
3 g of silica gel (60 — 100 mesh, Aldrich, Milwaukee, WI) in hexane, and an
additional 1 cm of sodium sulfate. This column was rinsed with 25 mL of hexane.
The sample was then loaded onto the column and the column drained until the
sample was adsorbed into the top sodium sulfate layer. The first fraction (F 1)
was eluted with 100 mL of 20% DCM in hexane at 1 drop/sec. A second and
third fraction (F2 and F3) were eluted with 100 mL of 75% DCM in hexane and
100 mL 50% DCM in MeOH, respectively, at 1 drop/sec. F1 contained the least
polar compounds, such as PAHs, PCBs, most organochlorine (OC) pesticides,
and contaminant compounds leached from the SDB resin (Figure 2). F2
contained nonylphenol (NP), octylphenol (OP), and greater polarity OC
pesticides (Figure 2). F3 contained the most polar compounds, including triazine
herbicides, pharmaceuticals, DEET, oxybenzone, flame-retardants, steroids, and
NPE1-2 (Figure 2). Each fraction was collected in a N2 Rapid VapTM tube
(Labconco, Kansas City, MO) and concentrated using a N2 RapidVapTM
(Labconco, Kansas City, MO) as previously described. The fractions were

further concentrated under a gentle stream of nitrogen and solvent-exchanged to

162

1 mL iso-octane (F1 and F2) or MeOH (F3). No internal or surrogate standards
were added to the samples because these could interfere with bioassays to be

used for estrogenic and dioxin-like bioactivity.

Identification and Quantitation. All compounds were identified and quantified
using an HP 5890 series Il GC with an HP 5972 mass selective detector (MSD).
A 30 m DB—5MS capillary column (0.25 mm l.D., 0.25 urn film, J&W Scientific,
Folsom, CA) was used for PAH, OC pesticides, atrazine, simazine, and PCB
analyses. An injection volume of 3 pL was used for these compounds.
Temperature programming was optimized based on that provided in the J&W
catalog (1998, J&W Scientific, Folsom, CA) for these analytes with the DB-5MS
column. NP and OP were analyzed using the same GCIMS system, but using a
30 m DB-17MS capillary column (0.25 mm l.D., 0.25 um film, J&W Scientific,
Folsom, CA). NPE122, pharmaceuticals, steroids, personal care products, and
BPA were analyzed using the same GCIMS system described previously. These
compounds were analyzed using the DB-5MS and DB-17MS described above,
plus a 30 m Rtx-50 capillary column (0.25 mm ID, 0.25 pm film, Restek,
Bellefonte, PA). Injection volumes of 2 uL were used for the analyses of these
compounds. For all compounds quantitated, the MSD was operated in selected
ion monitoring (SIM) mode with 2 -3 ions per compound (Tables 1 - 4).
Identification was determined by proper retention time and ion ratio. Quantitation
was performed using HP ChemStation software. All calibration curves were

linear (r2 > 0.9). All screening was performed with the MSD operating in scan

163

mode, and structural matching was accomplished using the NlST/EPAINIH mass
spectral database (May 1992 release, NIST, Gaithersburg, MD) (Tables 1 - 4).
Complete instrumental parameters are available as supplemental materials upon
request.

Further confirmation of previously unreported compounds from F3
extracts was accomplished using GC/HRMS. A CTC Analytics Model A2208
autosampler (Zwingen, Switzerland) was used to make 1 uL splitless injections
onto a 30 m DB-5 capillary GC column (0.25 mm ID, 0.25 pm film, J&W
Scientific, Folsom, CA) located within the oven of an HP 6890 GC. A Finnigan
MAT 900 S-trap hybrid mass spectrometer (Bremen, Germany) was used for
analysis of the column effluent. Only the double focusing MS stage was used in
these studies. Full scan low resolution (1000) mass spectra over mass ranges of
50-300 Da and 100-400 Da were acquired at 1.8 sec/cycle and 1.1 sec/cycle,
respectively. Elemental compositions of specific ions observed in background-
subtracted mass spectra were determined using MPPSIRD to acquire data and
the PGM to automatically interpret the data. Multiple mlz ratios were monitored
in selected ion recording mode across three to five calibration and analyte mass
peak profiles with a constant magnetic current and jumps in the accelerating and
electrostatic sector potentials. Each mlz ratio was monitored for 20 msec to
provide 1 sec/cycle for 31 monitored mlz ratios. Five mlz ratios each were used
to monitor 40% of the mass range of the lock mass and calibration mass profiles
which bracketed the analyte ion profiles investigated. The remaining 21 mlz

ratios were used to monitor a 2000 parts per million (ppm) wide mass range

164

centered on an estimated mass from a low resolution mass spectrum using 3000
resolution, a single analyte ion with 10 ppm mass increments between mlz ratios
using 10,000 resolution, or 60% of the mass range of three partial analyte
profiles using 25,000 resolution. This system was used only for identification

purposes and no quantitation was attempted.

Recovery and Precision. Spike recovery experiments were performed using
laboratory and Lake Mead water to determine the accuracy and precision of the
method. Target compounds were spiked into laboratory water at various
concentrations several times. During the initial sample collection at Lake Mead,
samples were collected in duplicate at two sites. One sample was spiked with
the target compounds while the other was not. To facilitate sample
homogenization, the standards were spiked into approximately 50 L of water and
mixed thoroughly. An additional 50 L of water was added vigorously and mixed
thoroughly. The analysis procedure was the same as previously described for

water samples.

Detection Limits. Instrumental detection limits (IDLs) were based on the
minimum concentration necessary to achieve a signal 3x the baseline noise. The
method detection limits (MDLs) were estimated based on lDLs and the sample
volume extracted (Tables 1 - 4). Furthermore, signals were not quantified in

sample extracts unless the signal to noise ratio in that sample was greater than

165

10. Detection limits could not be calculated for compounds detected by use of

NIST structural database matching or MPPSIRD and PGM.

RESULTS AND DISCUSSION

Analytical Considerations. Recoveries for the target compounds were
consistent in both laboratory and matrix spikes and ranged from 73 — 107%.
Therefore, no recovery adjustments were made to the data. Some compounds
of interest were detected in trace quantities in field blanks. These concentrations
were consistent and therefore subtracted from reported data. Contamination of
samples was most likely due to the Pentaplate filtration system because it had
great surface area and was difficult to clean thoroughly. Although the SPE
columns should have been able to achieve good recoveries with much faster flow
rates, back-pressure as a result of the small particle size and pressure limitations
of the metering pump (approximately 60 psi) limited the flow rate to 100 mUmin.
F3 extracts caused rapid deterioration of the GCIMS. Therefore, a
20 m guard column of deactivated fused silica (0.32 mm I.D., J&W Scientific,
Folsom, CA) was added, and 1 m was removed when chromatographic
resolution deteriorated. It also was observed that contamination of the injector
liner caused the greatest impact on chromatography. The use of a cyclo double
gooseneck liner (4 mm, Restek, Bellefonte, PA) allowed more injections of F3
extracts without replacement. In order to determine if chromatographic resolution

had deteriorated, a set of standards was analyzed after every 5 sample

166

 

 

injections. If the resulting quantitation varied by greater than 20%, the guard
column was trimmed of 1 to 2 m and the injector liner replaced. Even with these
measures, some polar F3 pharmaceuticals were not particularly amenable to GC
separation, and at times the resulting quantitation varied by as much as 50%.

Also, it was unknown whether these compounds were stable over time. To

reduce sample weathering, all extracts remained cooled at --15 °C until analysis.

Organochlorine Pesticides and PCBs. Concentrations of OC pesticides in the
waters of Lake Mead were generally low (Tables 5 & 6). Concentrations of
HCHs and DDTs ranged from 0.12 to 25.9 nglL and 0.16 to 28.2 nglL,
respectively (Tables 5 & 6). One water blank that was collected following
sampling in Las Vegas Bay contained 4.8 ng DDT/L. Data were not adjusted for
this contamination. The observed patterns of relative concentrations of HCHs
and DDTs indicate leaching from a source that has been under anaerobic
conditions (such as burial). Ground water seepage entering Las Vegas Wash
has been shown contain HCHs with the same isomer pattern observed in this
study (1999 Southern Nevada Water Authority, unpublished data). Historical
manufacturing of OC pesticides in the Las Vegas Valley is the most likely source
of these pesticides in the Las Vegas Bay (Southern Nevada Water Authority
communication). Concentrations of OC pesticides are extremely low in other
areas of Lake Mead.

The US. EPA fresh water chronic exposure to aquatic life

protection value for DDT is 0.001 pglL. The EPA values for bioconcentration to

167

predators (including humans) and human health cancer risk for DDT are 0.024
and 0.59 ng/L, respectively. Therefore, concentrations of DDT in the Las Vegas
Bay may be sufficient to cause adverse affects in certain aquatic organisms
(Federal Register 45-FR-79331).

The chronic aquatic toxicity values for all HCH isomers is 0.08 pglL.
For the protection of predatory species (through biomagnification) is 0.0163,
0.0186, and 0.0123 ug/L for a-, [3-, and y- HCH, respectively (Federal Register
45-FR-79335). Therefore, Las Vegas Bay water concentrations for B-HCH would
have exceeded the recommended limits on two occasions (Tables 5 & 6). This
indicates that HCHs may pose a threat to predator species in Lake Mead.

PCBs were not detected in any water samples from Lake Mead. This is
not surprising since these compounds are extremely lipophilic and very insoluble
in water. The chronic toxicity value for fresh water and water quality criteria for
aquatic life and predators such as humans (including safety factors and
bioaccumulation factors) are 0.065 and 0.00079 ug/L, respectively. Therefore, it
is unlikely that total PCB concentrations in Lake Mead waters pose a significant

ecological threat.

Polycyclic Aromatic Hydrocarbons. PAHs were detected at every site in Lake
Mead. PAHs were also detectable in all water and column blanks. Summed
concentrations of the 16 PAHs investigated did not vary significantly among the
sites and ranged from 14 to 78 ng/L. Patterns of relative concentrations of PAHs

did not vary significantly among locations with the exception of naphthalene,

168

concentrations of which were greater in the Las Vegas Bay. It is possible that
motors from the sample collection vessels may have influenced PAH results
since fuel from the motors occasionally was visible in the water. Ubiquitous
detection of PAHs in Lake Mead is consistent with the great usage of recreational
vehicles on the Lake.

The value of the protection of water and organisms (including
biomagnification potential and is based primarily on human health) for total PAHs
is 0.0028 jig/L. Therefore, most sites tested at Lake Mead exceed this threshold

and may have adverse affects on aquatic organisms.

Alkylphenols and Bisphenol A. NP, OP, NPE1&2, and BPA were detected at
some sites in Lake Mead (I' able 7). Detectable concentrations of NP and OP
ranged from 5.0 to 308 nglL and 1.4 to 23.2 ng/L, respectively, and were
consistently greater in the Las Vegas Bay. NPE1 and NPE2 concentrations
ranged from 40.1 to 1540 ng/L and 10.0 to 702 nglL, respectively.
Concentrations in the Las Vegas Bay were always much greater than
concentrations at the reference sites. BPA was detected in only one sample
from the Las Vegas Bay at 2.4 ng/L.

Acute and chronic toxicity values for NP range from approximately 17 to
3,000 and 4 to 900 pglL for a wide range of aquatic organisms (7).
Concentrations of NP observed in Lake Mead are far below those likely to cause
acute or chronic toxicity (Table 7). Although NP has been shown to be weakly

estrogenic in a variety of both in vitro (8-12) and in vivo bioassays (13-15),

169

 

concentrations to elicit effects are generally greater than those found in the

waters of Lake Mead.

Pharmaceuticals. Several pharmaceuticals were detected in Las Vegas Bay
water samples (T ables 8 - 10). Initially, pharmaceuticals were detected using the
MSD in scan mode. Mass spectra of the suspected pharmaceuticals were
compared to the spectra of authentic standards (Figures 3 & 4). Because these
matches were extremely good, the samples and standards were also analyzed in
SIM mode (Figures 5 & 6). Concentrations of pharmaceuticals detected using
the MSD in SIM mode and calibrated with authenticated standards ranged from
less than the MDLs to 260 ng/L. These data are presented semi-quantitatively.
These concentrations of pharmaceuticals are extremely small.
Because there are no water quality criteria for these compounds, the
toxicological significance needs to be placed into proper perspective.
Furthermore, because persons using these compounds are exposed to
pham'iacologically appropriate doses, it is inappropriate to compare exposures to
maximum tolerated or toxic doses. Instead, the relevance of the dose that would
be received by persons consuming untreated water was assesses by calculating
the volume of water that would need to be consumed to obtain a single daily
dose. For instance, a typical human dose of phenytoin is 50 mglday. If one were
to drink the water from the location in Las Vegas Bay with the greatest
concentration of phenytoin (260 nglL), it would require drinking approximately

190,000 L of this water to achieve a therapeutic dosage. The average daily

170

 

 

water intake of humans is 2 Ud. Thus, if the entire volume of water consumed
contained phenytoin at the maximum concentration observed in surface waters of
Las Vegas Bay, a person would receive 0.001% of a therapeutic dose. While
these calculations do not represent a rigorous risk assessment, it does indicate
that the concentrations of pharmaceuticals in the waters of Lake Mead probably
do not present a significant risk to humans. However, the potential of the
observed concentrations of pharmaceuticals directly on wildlife is unknown and
can not be accurately assessed due to a lack of information of the effects of
these compounds on wildlife species. A thorough review of pharmaceuticals in

the environment has been published (16).

Other Compounds. Several other classes of novel compounds were detected
in Lake Mead water samples (Tables 1 & 6). Initial validation of these
compounds was performed using the MSD in scan mode using authenticated
standards or the NIST mass spectral database (Figures 7 & 8). Concentrations
of compounds analyzed by GCIMS SIM for which authentic standards were
available are presented (Table 11). The resolution and reproducibility of DEET
and oxybenzone were such that 2 significant figures are provided. Whereas, due
to poorer analytical resolution, concentrations of nicotine and caffeine are
reported to only one significant figure. Compounds identified only by spectral
matching with the NIST mass spectral database are presented (Tables 1 - 4).
These compounds were observed only in water samples from the Las Vegas

Bay.

171

 

High-resolution Mass Spectrometry. Analytical tools have been developed
recently that utilize GC/HRMS for identification of unknown organic compounds
in environmental samples. For example, several isomers of acrylonitrile-styrene,
which are byproducts of a polymerization process, were identified in water from a
municipal well near Toms River, NJ, where an increased incidence of childhood
cancer was observed (17). Several novel compounds were verified and
identified using MPPSIRD and PGM (Table 12). These data are considered to

be qualitative only and were detected only Las Vegas Bay.

Summary. This paper provides information on a methodology that used large
volume water sampling with solid-phase extraction to screen for a broad range of
classes of organic compounds. Several classes of organic compounds were
determined to be entering Lake Mead via the Las Vegas Wash. However, the
Virgin, Muddy, and Colorado Rivers entering Lake Mead were not tested.
Concentrations of OC pesticides and halogenated aromatic hydrocarbons were
relatively small. A number of previously unreported compounds including
pharmaceuticals, health care products, and flame-retardants were identified.
This study may serve as a basis for further research that could validate and

better quantitate these novel compounds.

ACKNOWLEDGMENTS

172

 

 

This work was supported by grants and donations from the Southern
Nevada Water Authority, US. Department of the Interior - Bureau of
Reclamation, Clark County Sanitation District, and the National Park Service.
Technical and instrumental support was provided by US. Environmental
Protection Agency - National Environmental Research Laboratory, Las Vegas,
NV. The authors would like to thank Fred DeSiIva and the rest of the National

Park Service Volunteers who operated and organized boats for this project.

REFERENCES

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(2) Prentki, R. T.; Paulson, L. J. In Aquatic Resources Management of the
Colorado River Ecosystem; Adams, V. D., Lamarra, V. A., Eds.; Ann Arbor
Science: Ann Arbor,Ml, 1983, pp 105-123.

(3) Paulson, L. J.; Baker, J. R. In Proceedings of the Symposium on Surface
Water lmpoundments; Stefan, H. G., Ed.; American Society of Civil
Engineering: New York, 1981, pp 1647-1658.

(4) Bevans, H. E.; Goodbred, S. L.; Miesner, J. F.; Watkins, S. A.; Gross, T.
S.; Denslow, N. D.; Schoeb, T. Water-Resources Investigations Report
96-4266 1996.

(5) Snyder, S. A.; Keith, T. L.; Verbrugge, D. A.; Snyder, E. M.; Gross, T. S.;

Kannan, K.; Giesy, J. P. Environ. Sci. Technol. 1999, 33, 2814-2820.

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(7)

(8)
(9)

(10)

(11)
(12)

(13)

(14)

(15)

Snyder, S. A.; Snyder, E.; Villeneuve, D.; Kurunthachalam, K.; Villalobos,
A.; Blankenship, A.; Giesy, J. In Analysis of Environmental Endocrine
Disruptors; Keith, L. H., Jones-Lepp, T. L., Needham, L. L., Eds.;
American Chemical Society: Washington,DC, 2000, pp 73-95.

Staples, C. A.; Weeks, J.; Hall, J.; Naylor, C. Environ. Toxicol. Chem.
1998, 17, 2470-2480.

Mueller, G. C.; Kim, U. H. Endo. 1978, 102, 1429-1435.

Soto, A. M.; Tien-Min, L.; Honorato, J.; Silvia, R. M.; Sonnenschein, C. In
Chemically-induced alterations in sexual and functional development: The

wildlife/human connection; Mehlman, M. A., Ed.; Princeton Scientific

 

Publishing Co.,lnc: Princeton,NJ, 1992; Vol. 21.

Desbrow, C.; Routledge, E. J.; Brighty, G. C.; Sumpter, J. P.; Waldock, M.
Environ. Sci. Technol. 1998, 32, 1549-1558.

Jobling, S.; Sumpter, J. P. Aquat. Toxicol. 1993, 27, 361-372.

White, R.; Jobling, S.; Hoare, S. A.; Sumpter, J. R; Parker, M. G. Endo.
1994, 135, 175-182.

Jobling, S.; Sheahan, D.; Osborne, J. A.; Matthiessen, P.; Sumpter, J. P.

 

Environ. Toxicol. Chem. 1996, 15, 194-202.

Giesy, J. P.; Pierens, S. L.; Snyder, E. M.; Miles-Richardson, S.; Kramer,
V. J.; Snyder, S. A.; Nichols, K. M.; Villeneuve, D. A. Environ. Toxicol.
Chem. 2000, (in press).

Nimrod, A. C.; Benson, W. H. Crit. Rev. Toxicol. 1996, 26, 335-364.

 

174

(16) Daughton, C. G.; Ternes, T. A. Environ. Health Perspect. 1999, 107, 907-
938.
(17) Grange, A. H.; Sovocool, G. W. Rapid Commun. Mass Spectrom. 1998,

1161-1169.

 

175

 

TABLE 1. F1 and F2 Compounds Detected, Estimated Method Detection
Limits (nglL), and Ions Monitored

 

 

MDL Ions Usage/Source
E1
HCHs' 0.10 Various Insect control
DDT,DDE, and DDD 0.15 Various Insect Control
PAHs“ 4.0 Various Petroleum-based products
3
NP 5.0 107, 135 Degradation product of NPEs
OP 1.0 107, 135 Degradation product of OPEs

 

176

 

 

TABLE 2. Pharmaceuticals Compounds (F3) Detected, Estimated Method
Detection Limits (nglL), and Ions Monitored

 

 

\

Compound MDL Ions UsagaISource
Carbamazepine 10 165, 193, 236 Pharmaceutical — anti-seizure
Primidone 20 1 17, 146, 190 Pharmaceutical - anti-seizure
Phenytoin 50 180, 223, 209 Pharmaceutical — anti-seizure
Phenobarbital 10 204, 117 Pharmaceutical — anti-seizure
Diazapam 1 256, 283, 284 Pharmaceutical - anxiety disorders
Meprobamate NA Scan Pharmaceutical - anxiety disorders
Guaifenesin 20 109, 124, 198 Pharmaceutical - expectorant
Codeine 10 162, 229, 299 Pharmaceutical — pain relief
Hydrocodone 4 185, 242, 299 Pharmaceutical — pain relief
Pentoxifylline 5 180, 193, 221 Pharmaceutical — blood thinner
Carisoprodol NA Scan Pharmaceutical - muscle spasms
Methocarbamol NA Scan Pharmaceutical — muscle spasms
Mequinol NA Scan Pharmaceutical - pigmentation

 

NA = not applicable

177

 

TABLE 3. F3 Compounds Detected, Estimated Method Detection Limits
(nglL), and Ions Monitored - Part I

 

 

 

Compound MDL Ions Usage/Source
NPE1 30 219, 223, 228 Surfactants
NPE2 30 107, 223, 237 Surfactants
Nicotine 4 84, 133, 161 Tobacco products
Caffeine 7 67, 109, 194 Coffee, tea, etc.
DEET 2 91, 1 19, 190 Insect repellant
BPA 2 213, 219, 228 Plasticizer
Oxybenzone 10 151 , 227, 228 Sunscreen
Homomenthyl salicylate NA Scan Sunscreen
o-Methoxyacetophenone NA Scan Fragrance
Vanillin NA Scan Fragrance
Terbuthylazine NA Scan Weed control

 

NA = not applicable

178

 

TABLE 4. F3 Compounds Detected, Estimated Method Detection Limits
(nglL), and Ions Monitored - Part II

 

 

Compound MDL Ions Usage/Source

p-Chloroaniline NA Scan Dye

Acridine NA Scan Dye

Triclosan NA Scan Anti-fungal

F ulvicin NA Scan Anti-fungal

Tris(2-chloroethyl)phosphate NA Scan F lame-retardant I
Tris(2-chloropropyl)phosphate NA Scan Flame-retardant

 

Tris(1,3-dichIoroisopropyl)phosphate NA Scan Flame-retardant

Triphenylphosphate NA Scan Industrial chemical
Triethylphosphate NA Scan Industrial chemical
Ethyl citrate NA Scan Food additive
Mequinol NA Scan Anti-pigmentation
Cholesterol NA Scan Biomolecule

 

NA = not applicable

179

 

TABLE 5. Concentrations of HCHs in Lake Mead (nglL)

 

 

Location Date a-HCH B-HCH S-HCH y-HCH
LV Wash 6/23/98 7.94 25.91 23.53 7.43
3/26/99 1.43 3.18 2.26 1.95
5/05/99 1.88 5.70 3.34 2.86
6I09/99 2.96 8.07 5.20 4.17
LV-0.5 3/26/99 1 .06 2.23 1 .45 1.58
5/05/99 1.15 2.61 1.75 1.66
6/09/99 1.21 0.80 2.11 1.32
LV-1 (plume) 8/03l98 3.25 13.3 10.8 4.88
3/26/99 6.88 20.3 12.3 10.6
5/05/99 2.45 6.84 4.56 3.49
LV-1 (surface) 6/23/98 1 .49 4.56 3.32 2.33
8/03/98 0.42 2.40 0.85 ND
Water Barge 6/22I98 ND ND ND ND
8/03/98 ND ND ND ND
3/26/99 ND ND ND ND
5/05/99 ND 0.26 ND 0.21
6I09/99 ND ND ND ND
Moon Cove 6/22/98 0.12 0.38 0.31 ND
3/26/99 ND 0.47 ND 0.27
5/05/99 ND 0.26 ND 0.24
6/09/99 ND 0.24 ND 0.23

 

ND = not detectable

180

 

 

TABLE 6. Concentrations of DDTs Detected in Lake Mead (nglL)

 

 

Location Date DDE DDD DDT
LV Wash 6/23/98 ND 0.39 1.09
3/26/99 ND 0.24 0.48
5/05/99 ND 0.42 4.26
6/09/99 ND 0.52 9.61
LV-0.5 3/26/99 ND 0.47 12.0
5105/99 ND 0.28 ND
6/09/99 ND ND ND
LV-1 (plume) 8/03/98 0.18 1.15 19.5
3/26/99 ND 0.36 28.2
5/05/99 0.29 0.44 25.7
LV-1 (surface) 6/23/98 ND 0.16 1.81
8l03l98 ND 0.46 4.62
Water Barge 6/22/98 ND ND ND
8/03/98 ND ND ND
3/26/99 ND ND 0.31
5/05/99 ND ND ND
6/09/99 ND ND ND
Moon Cove 6122/98 ND ND ND
3/26/99 ND ND 0.46
5/05/99 ND ND 0.45
6/09/99 ND ND 0.50

 

ND = not detectable

181

 

 

TABLE 7. Concentrations of Alkylphenols and Bisphenol A in

 

 

Lake Mead (nglL)
Location Date OP NP NPE. NPE2 BPA
LV Wash 6/23/98 22.0 308 1000 663 ND
3/26/99 3.7 32.3 193 138 ND
5I05/99 ND ND 192 94 ND
6/09/99 7.1 61 .9 496 350 ND
LV-0.5 3/26/99 1 .4 13.6 129 33.0 ND
5/05/99 4.0 40.2 198 58 2.4
6/09/99 1.8 18.3 117 57 ND
LV-1 (plume) 8l03l98 15.1 172 398 222 ND
3/26/99 23.2 1 96 1 540 702 ND
5I05/99 20.7 232 978 468 ND
LV-1 (surface) 6/23/98 ND 78.7 677 489 ND
8/03/98 ND 10.1 47 ND ND
Water Barge 6/22/98 ND ND ND ND ND
8l03l98 ND ND ND ND ND
3/26/99 ND ND ND ND ND
5I05/99 ND ND ND ND ND
6/09/99 ND ND ND ND ND
Moon Cove 6/22/98 ND 5.6 40.1 10.0 ND
3/26/99 ND 5.0 ND ND ND
5/05/99 ND ND ND ND ND
6/09/99 ND ND ND 29.3 ND

 

ND = not detectable

182

 

 

TABLE 8. Concentrations of Pharmaceuticals in Lake Mead (nglL), Part I

 

 

Location Date Phenytoin Codeine Primidone
LV Wash 6/23/98 100 20 80
3l26l99 70 ND 40
5/05/99 80 30 40
6/09/99 260 ND 80
LV-0.5 3l26l99 50 20 20
5I05I99 ND 20 40
6/09/99 70 30 40
LV-1 (plume) 8l03l98 250 100 80
3l26l99 230 120 50
5/05/99 250 ND 1 30
LV-1 (surface) 6/23/98 ND ND 10
8/03/98 ND ND 20
Water Barge 6/22/98 ND ND ND
8/03/98 ND ND ND
3/26/99 ND ND ND
5/05/99 ND ND ND
6I09/99 ND ND ND
Moon Cove 6/22/98 ND ND ND
3l26l99 ND ND ND
5I05/99 ND ND ND
6/09/99 ND ND ND

 

ND = not detectable

183

 

 

 

TABLE 9. Concentrations of Pharmaceuticals in Lake Mead (nglL), Part II

 

 

Location Date Pentoxifylline Carbamazepine. Guaifenisen.
LV Wash 6/23/98 10 30 40
3/26/99 1 0 30 40
5/05/99 ND 20 30
6/09/99 10 30 50
LV-0.5 3/26/99 ND ND ND
5I05/99 ND 40 20
6/09I99 1 0 30 30
LV-1 (plume) 8/03/98 ND 40 40
3l26l99 ND ND ND
5/05/99 ND ND ND
LV-1 (surface) 6/23/98 ND 20 ND
8/03/98 ND 10 ND
Water Barge 6/22/98 ND ND ND
8/03/98 ND ND ND
3l26l99 ND ND ND
5/05/99 ND ND ND
6/09l99 ND ND ND
Moon Cove 6/22/98 ND ND ND
3/26/99 ND ND ND
5/05/99 ND ND ND
6/09/99 ND ND ND

 

ND = not detectable

184

 

TABLE 10. Concentrations of Pharmaceuticals in Lake Mead (nglL), Part III

 

 

Location Date Phenobarbital Hydrocodone Diazepam
LV Wash 6/23/98 1 0 ND ND
3l26l99 ND ND 50
5/05/99 ND ND 60
6/09/99 ND 10 ND
LV—0.5 3l26l99 ND ND ND
5/05/99 ND ND ND
6/09/99 ND 10 ND
LV-1 (plume) 8/03/98 30 10 ND
3l26l99 40 ND ND
5I05/99 40 10 10
LV-1 (surface) 6l23/98 ND ND ND
8/03/98 ND ND ND
Water Barge 6/22/98 ND ND ND
8/03/98 ND ND ND
3/26/99 ND ND ND
5/05/99 ND ND ND
6/09/99 ND ND ND
Moon Cove 6/22/98 ND ND ND
3l26l99 ND ND ND
5/05/99 ND ND ND
6I09I99 ND ND ND

 

ND = not detectable

185

 

 

TABLE 11. Concentrations of Other Compounds in Lake Mead (nglL)

 

 

Location Date DEET Oxybenzone Caffeine Nicotine
LV Wash 6/23/98 270 200 10 1 0
3l26l99 20 56 30 1 0
5/05/99 24 1 50 30 1 0
6I09I99 87 82 30 1 0
LV-0.5 3l26l99 12 140 20 ND
5I05l99 1 1 120 20 10
6I09I99 45 210 20 10
LV-1 (plume) 8/03/98 164 220 30 10
3/26/99 51 120 30 ND
5I05l99 67 58 50 20
LV-1 (surface) 6/23/98 57 360 10 ND
8l03l98 17 200 ND ND
Water Barge 6/22/98 ND 73 ND ND
8/03/98 4 79 ND ND
3l26l99 N D 38 ND ND
5/05/99 ND 53 1 0 ND
6/09/99 3 78 ND ND
Moon Cove 6/22l98 3 38 ND ND
3l26l99 ND 180 ND ND
5I05l99 ND 30 ND ND
6/09/99 ND 59 ND ND

 

ND = not detectable

186

 

TABLE 12. Compounds Identified in Las Vegas Bay by HRMS

Phenobarbital

Phenytoin

Oxybenzone

Trichlorophenol

Ethyl citrate
Tris(2-chloroethyl)phosphate
Tris(2-chloropropyl)phosphate

Tris(1 ,3-dichIoroisopropyl)phosphate

Tertbutylazine

187

 

 

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c r-, )
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Cove ‘- k, ,1‘\_,'
‘ I
I
Government Wash . ‘ r t. ,u 7 95‘. 2t,
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m LV-t -- 1, ml (061°
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Q
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I I
D 'P‘ ' ’
I I
\\ ' I \ ' ls
" -- o 34
I a
09 ‘

9, ‘
ye.

. I
'Las Vegas Bay ' . 4, , :~ '
Boat Harbor

. Boulder .' r'
’1 "“ Moon ' la
Fish Hatchery“ ' v Cove Basrn ' '
Pumping :‘\ Saddle ." '_ ("
Statlons 4\ Island (S, . ‘ _

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Boulder
Boat Harbor ‘

\‘ ‘h‘ ‘1 ‘ \
Boulder ‘~ 2 3? :
Beach ‘-~, I, ’ 3’
\\"‘~ I ’1’)

\

Figure 1. Map of the Boulder Basin study area of Lake Mead, Nevada

188

 

 

 

130 L filtered
in situ

I

100 L solid-phase
extracted (SPE)

I

SPE eluted, liquid/liquid extracted, and
concentrated to 2 mL

I

1 mL silica gel
fractionated

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

iF1 vF2 lF3

 

 

 

 

Non—polar Semi-Polar Polar

 

 

 

 

 

 

 

 

I I I

 

 

 

 

PAHs NP NPE122
OC Pesticides A OP Atrazine
PCBs OC Pesticides Simazine
Estrone

 

 

 

 

 

1 78-estradiol
Ethynylestradiol
Progesterone
Pharmaceuticals
DEET
Nicotine
Caffeine
Oxybenzone
Bisphenol A

 

 

 

Figure 2. Flow diagram of analytical method

189

 

 

180

A. 77 10“ e 209 223
l 165 l 252
147 l
I J . u L I. .1. l l ,7 l

104
Be 77 223

209
I65 252

193
147 236

 

 

 

 

 

 

mlz

Figure 3. Mass spectra of phenytoin (Dilantin): (A) authentic standard;

(B). LV Bay water extract

190

 

 

 

 

 

 

 

 

204
O O
A N N
. T
117 O
77 “’4 161
91 146 II 174 i1 232
II "I In I I I
204
B.
117
104
77 91 146 161 174 i 232
All A. - . J] I.
m/z

Figure 4. Mass spectral scan of phenobarbital: (A) authentic standard;

(B) LV Bay water extract

191

 

 

DEET Oxybenzone

 

 

 

 

 

 

 

Phcnacetin
A Guaifenesin Bromphenlramine
O
Chlorcyclizinc Pentoxifylline
Phenytoin
r l I I A f
B .
I I I T '1 _ I
18 21 24 27 30 33 36

RT (min)

Figure 5. Mass spectral SIM chromatograms I: (A) authentic standards;

(B) LV Bay water extract

192

 

 

 

 

 

 

 

 

 

 

 

 

 

f I 1 I
9 12 15 18 21 24
RT (min)

Figure 6. Mass spectral SIM chromatograms II: (A) standards; (B) LV Bay
water extract. Peak # 1. DEET; 2. Guafenesin; 3. Acetominophen; 4.

Meprobamate; 5. Caffeine; 6. Oxybenzone; 7. Bisphenol A; 8. A8 THC; 9.

A9 THC; 10. Primidone; 11. Carbamazepine; 12. Hydrocodone

193

 

 

119

<“"\

A. 91 s

 

 

b3
i—
=—
[:_——

119

It .I .---I . .I

Figure 7. Mass spectral scan of DEET: (A) authentic standard; (B) LV

 

 

Bay water extract

194

 

 

 

 

 

 

 

 

 

CI
75 CI
Cl Cl
99 \__2 O S—/
II
0— 1'2— 0
A. 191 o
\CC|
155 209 C' 381
u 243 321
1. 1.
75
99
B- .9.
155 209 m
I L 321
1. . L ..
m/z

Figure 8. Mass spectral scan of tris(1,3-dichloroisopropyl)phosphate: (A)

NIST database spectra; (B) LV Bay water extract 1

195