Sediment acute toxicity testing utilizing short-term bioassays

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Sediment acute toxicity testing utilizing short-term bioassays
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Thesis (Ph. D.)--University of Florida, 1991.
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Includes bibliographical references (leaves 145-157).
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by Marjorie G.S. Campbell.
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SEDIMENT ACUTE TOXICITY TESTING
UTILIZING SHORT-TERM BIOASSAYS














By

MARJORIE G. S. CAMPBELL


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


1991





























Copyright 1991
by
Marjorie G. S. Campbell
























To my children, Adrian and Demeka,
in hopes that this work will be
inspirational in their educational pursuits















ACKNOWLEDGEMENTS

To God be the glory for the great things He has done!

It is because of the blessings of God that I have been

surrounded by individuals who have assisted me in this

educational endeavor. I wish to express my sincere

gratitude to Dr. Gabriel Bitton for his suggestions,

guidance and encouragement during this investigation.

Special thanks is extended to Dr. Thomas Crisman for his

positive advice. Gratitude is extended to other members of

my supervisory committee, Dr. Ben Koopman, Dr. Joseph

Delfino and Dr. Sam Farrah for their guidance during this

study.

The unmeasureable support of my husband, Willie, is

especially acknowledged. He has been the "rock" I have

stood on, the "shoulder" I have cried on and the "force"

that kept me striving.

My entire family is acknowledged for their positive and

inspirational support.

I also wish to thank my fellow students, in particular

Mr. Mark Jacobs and Mr. Shanshin Ton for their help and

guidance during a portion of this study. Additional thanks

is extended to the entire "toxics project" group for the

collection of sediment samples used in this study. Special









thanks is also extended to Mrs. Sybil Stephens Brown and Ms.

Paula Rooks for their invaluable assistance.

Deep appreciation is extended to Ms. Shirley A. Buford

for her patience and skills in the typing of a portion of

this dissertation.

Finally, deep appreciation is extended to the Florida

Endowment Fund and its president Dr. Israel Tribble. This

work would not have been possible without the financial and

moral support of this outstanding organization.

Partial funding of this work was provided through

Contract No. WM266 from the Florida Department of

Environmental Regulation, Dr. J. J. Delfino, principal

investigator and Dr. R. Patton, project officer.















TABLE OF CONTENTS


page


ACKNOWLEDGEMENTS . .

ABSTRACT . . .


. iv

*. ix


CHAPTERS


1 INTRODUCTION . .... .. 1

2 LITERATURE REVIEW . . 7

Sediment Contamination . 7
Toxicity Bioassays . .. 10
Bacterial Bioassays.. . ... 15
Battery-of-Test Approach To Toxicity
Testing of Sediments ...... .. 20
Rationale for the use of the
B. licheniformis Inhibition of Enzyme
Biosynthesis Assay in Toxicity Screening
of Sediments . . ... .24
Rationale for the use of
Chydorus sphaericus in Sediment
Toxicity Tests . ... .27

3 TOXICITY TESTING OF SEDIMENT ELUTRIATES
BASED ON INHIBITION OF ALPHA-GLUCOSIDASE
BIOSYNTHESIS IN BACILLUS LICHENIFORMIS .. .31

Introduction . ... .. .31
Materials and Methods . ... 34
Test Chemicals, Reagents and Media 34
Sediment Samples . ... 35
Preparation of Sediment Elutriates ... .35
Test Bacteria ................. .36
Assay for Alpha-Glucosidase Biosynthesis ... 36
The Microtox Assay . .... 46
Metal Analyses . . 48
MetPAD . ..... .......... .48
Data Analysis . ... 49
Results .. . 50
Toxicity Testing of Sediment Elutriates:
Comparison of the Alpha-Glucosidase
Biosynthesis and Microtox Bioassays .. .50









Heavy Metal Toxicity of Some Sediment
Elutriates as Determined by MetPAD 65
Metal Concentrations in Elutriate Samples 70
Discussion . . ... .74


4 COMPARISON OF SEDIMENT EXTRACTION
PROCEDURES AND EXCHANGE SOLVENTS
FOR HYDROPHOBIC COMPOUNDS BASED ON
INHIBITION OF BIOLUMINESCENCE .

Introduction . .
Materials and Methods . .
Chemicals and Reagents .
Sediment Samples . .
Percentage Organic Carbon .
Extraction Procedure .
The Microtox Assay .
The Bacillus licheniformis
Alpha-Glucosidase Assay (AGB Assay)
Data Analysis . .
Results
Shaking vs Sonication for Extracting
Sediment Samples .
Methanol vs DMSO as Exchange Solvent
EC50s of Shaking Extracts Exchanged
Into DMSO . .
Alpha-Glucosidase Biosynthesis Assay
Discussion . .

5 THE CHYDORUS SPHAERICUS ACUTE TOXICITY
BIOASSAY . .


. 81


. 89
. 92


Introduction . .
Materials and Methods .
Starting of Chydorus sphaericus Cultures
Culture and Dilution Water .
Food . . .
Culture Maintenance .. .
Test Chemicals and Solutions .
Bioassay Methodology . .
Data Analysis ... .
Results . . .
Discussion . .

6 ACUTE TOXICITY SCREENING OF SEDIMENTS
UTILIZING CHYDORUS SPHAERICUS .


Introduction . .
Materials and Method .... .
The Chydorus sphaericus Miniaturized
Solid Phase Sediment and Water
Toxicity Test . .


104

104
106
106
107
108
108
109
110
112
113
119


124

124
125


. 125


vii


. .
. .
. .
* *


. .



r








Liquid Phase Elutriate Test with
C. sphaericus and Ceriodaphnia dubia .126
Data Analysis . ... 127
Results. . ........ 127
The Miniaturized Solid Phase Sediment
and Water Toxicity Test . .. .127
Liquid Phase Elutriate Test with
C. sphaericus and Ceriodaphnia dubia .129
Discussion . . .. .. 136

7 CONCLUSIONS . . .. 140

REFERENCES . .. ... 145

BIOGRAPHICAL SKETCH . ... 158


viii















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy


SEDIMENT ACUTE TOXICITY TESTING
UTILIZING SHORT-TERM BIOASSAYS

By

Marjorie G. S. Campbell


May 1991

Chairman: Gabriel Bitton
Major Department: Environmental Engineering Sciences

The purpose of this study was to investigate the

usefulness of two new bioassays for acute toxicity

assessments of sediments. A bacterial bioassay based on

inhibition of alpha-glucosidase biosynthesis in Bacillus

licheniformis and a 48-hour lethality bioassay employing the

benthic cladoceran, Chvdorus sphaericus, were evaluated by

direct comparisons with standard bioassays, using sediment

samples collected from various sites in Florida.

This study showed that the bioassay based on inhibition

of alpha-glucosidase biosynthesis in Bacillus licheniformis

was useful in the acute toxicity screening of sediment

elutriates. In regards to Escambia County, Florida samples,

the assay was comparable with the Microtox assay and was

especially sensitive for samples containing metals.

ix









To determine an appropriate procedure for assessing

hydrophobic contaminants of sediments in the B.

licheniformis bioassay, two extracting procedures were

compared. Based on the responses in the Microtox bioassay,

shaking (24 hours) sediment samples in methylene chloride

produced extracts that were significantly higher in toxicity

than extracts obtained by sonication (15 minutes with

methylene chloride) for eight of the ten sediment samples

tested. Comparisons of methanol and dimethyl sulfoxide

(DMSO) as exchange solvents revealed that there was

generally no significant difference between these solvents

in terms of toxicity in the Microtox assay. Solvent

extracts prepared by shaking and exchanged into methanol

showed lower toxicity in the B. licheniformis bioassay than

in the Microtox assay. Observed sediment toxicity in both

bioassays was expressed in terms of the equivalent dry

weight concentration of sediment causing 50% inhibition of

the assay organism.

Investigations of the 48 hour lethality bioassay

employing Chvdorus sphaericus showed that it can be used

successfully in the acute toxicity screening of whole

sediments and sediment elutriates. The bioassay is

ecologically relevant because the organism is ubiquitous and

it lives associated with sediments in freshwater aquatic

environments. Most importantly, Chydorus sphaericus was









found to be just as sensitive to contaminants as other

organisms that are commonly used in sediment acute toxicity

bioassays.















CHAPTER 1
INTRODUCTION


The generation of large quantities of chemicals in

today's industrialized society has resulted in the release

of a variety of hazardous pollutants into the environment.

Some sources of such pollutants include: effluents from

waste water treatment plants; illegal dumping sites; sites

of accidental chemical spills; poorly designed landfills;

uncontrolled hazardous waste sites; and sites of deliberate

chemical applications. Release of hazardous pollutants from

such sources has resulted in environmental contamination

that is more visible today than in any period of the history

of our nation. Many hazardous pollutants are persistent and

have a tendency to concentrate in the food chain. Such

pollutants may interfere with vital bio-processes and can

pose a direct threat to human health. In response to this

threat, the U.S. Government has passed environmental

legislation (e.g. the Clean Water Act, the Clean Air Act,

the Resource Conservation and Recovery Act, and the

Superfund Act) over the past two decades that has motivated

the development of innovative methods for monitoring the

potential impacts and toxicity of hazardous pollutants to











the environment. Many of these methods include a variety of

bioassays and chemical procedures.

In general, the toxicity of environmental media

contaminated by hazardous pollutants can be estimated using

two approaches: a toxicity-based approach and a chemistry-

based approach. The toxicity-based approach employs

bioassays that were developed for measuring and regulating

the toxicity of complex effluents discharged to surface

waters and for the identification of toxic waste under the

Resource Conservation and Recovery Act (U.S. EPA 1986) and

the Superfund Act (U.S. EPA 1987). Bioassays directly

measure toxic effects and, in general, they measure

biological effects associated with exposure to hazardous

pollutants.

The chemistry-based approach utilizes chemical analyses

primarily to establish criteria values for concentrations of

chemicals in the environment. This approach involves

complex methods and requires costly equipment, well-equipped

analytical laboratories and trained personnel. Most

importantly, chemical data alone are not always reliable

indicators of biological effects (Long and Chapman 1985).

Pollutants in aquatic environments are of concern

because water is essential to all life, yet, it can also

serve as a transporter of toxic pollutants. Because of this

fact, bioassays utilizing aquatic species have been used

extensively in the formulation of water quality criteria and











standards. However, in spite of monitoring efforts, there

is evidence of environmental degradation in areas where

surface water quality criteria and standards are not being

exceeded. Specifically, organisms living in or near the

sediments are being adversely affected, apparently by

chemical contaminants associated with sediments (Chapman

1989).

It was once accepted that contaminants in sediments are

effectively bound and unavailable, but recent studies have

shown that polluted sediments can be a significant source of

contaminants to aquatic organisms. In general, sediments

are not only a repository for trace organic and heavy

metals, but can also serve as a potential source for the re-

introduction of contaminants into the water column. Impacts

of contaminated sediments related to toxicity,

bioaccumulation, biomagnification and transfer through the

food chain to higher trophic levels are of concern. Given

the range of potential problems associated with contaminated

sediments, it is evident that knowledge of sediment

contamination and toxicity is important in understanding and

ultimately predicting the long-term fate and effects of

contaminants in aquatic environments.

Until the 1980s, freshwater studies assessing sediment

quality by means of toxicity testing were few in number

(Burton et al. 1989). Most testing has consisted primarily

of single species testing using Chironomus tentans (midge),











Hyalella azteca (amphipod), Gammarus lacustris (amphipod),

Hexacenia limbata (mayfly), and Daphnia magna (cladoceran)

(Nebeker et al. 1984). In a limited number of sediment

toxicity studies, bacterial bioluminescence (Giesy et al.

1988, Schiewe et al. 1985, True and Heyward 1990),

indigenous populations of bacteria (Barnhart and Vestal

1983, Giesy and Hoke 1988, Burton and Lanza 1985),

protozoans (Ross and Henebry 1989, Maciorowski et al. 1981),

and algae (McFeters et al. 1983) have been used in

bioassays.

The limited research associated with sediment toxicity

testing has given rise to many questions such as: which

test assays should be used with a particular class of

sediment contaminants; should whole sediment, elutriate

testing, or interstitial water be used; and what are the

most appropriate test methods and organisms? Obviously

there is a need for basic experimental data to help answer

these questions.

In accordance with the need for basic experimental data

related to sediment toxicity testing, the goals of this

study were to evaluate the applicability of: 1) a new

bacterial acute toxicity test based on the inhibition of

alpha-glucosidase biosynthesis in Bacillus licheniformis in

the screening and acute toxicity testing of sediment

elutriates and solvent extracts and 2) the benthic

cladoceran, Chydorus sphaericus, in sediment solid phase











screening and elutriate acute toxicity testing. Specific

objectives and their coverage in this dissertation are as

follows:

1. To refine and apply the alpha-glucosidase biosynthesis

(AGB) assay in the screening of sediment elutriates (Chapter

3).

2. To compare two solvent extraction methods (shaking vs.

sonication) for sediments to determine the better extraction

method to be used in conjunction with the AGB assay (Chapter

4).

3. To compare results of the AGB assay with results of the

Microtox assay to determine possible correlation between the

two bioassays (Chapter 3 and 4).

4. To develop laboratory culture techniques for the

maintenance of cultures of Chydorus sphaericus and determine

its relative sensitivity to select inorganic and organic

chemicals (Chapter 5).

5. To compare the toxicity of sediment elutriates to

Chydorus sphaericus and Ceriodaphnia dubia. (Chapter 6).

6. To use Chydorus sphaericus in the solid-phase and

elutriate acute toxicity screening of freshwater sediment

samples (Chapter 6).

These objectives were based on the following two

hypotheses:

1. The gram positive bacterium Bacillus licheniformis, when

employed in the alpha-glucosidase biosynthesis assay, is as









6

sensitive or more sensitive than the gram negative bacterium

Photobacterium phosphoreum that is employed in the Microtox

assay.

2. The ubiquitous benthic invertebrate Chydorus sphaericus,

is as sensitive or more sensitive than Ceriodaphnia dubia

for toxicity screening tests of aqueous extracts of

sediments.















CHAPTER 2
LITERATURE REVIEW


Sediment Contamination


Many studies have shown that xenobiotics of varying

chemical structure discharged into aquatic environments can

effectively become adsorbed to particulates and sediment

constituents (Wakeham and Carpenter 1976; Wu and Gschwend

1986; Ozretich and Schroeder 1986). Adsorbed chemicals or

residues may consist of the parent compound and/or its

reaction products, degradation products and metabolites.

The presence of a variety of residues in the environment is

of major concern to environmental scientists because the

persistence, and fate of residue chemicals are major factors

in determining their impact on the environment.

Several mechanisms have been proposed for adsorption of

chemicals to particulates and sediment constituents. Two or

more mechanisms may occur simultaneously depending on the

chemical and the solid surface. These mechanisms as

summarized by Khan (1980) are: (1) Van der Waals

attraction; (2) hydrophobic bonding; (3) hydrogen bonding;

(4) charge transfer; and (5) ion exchange. Other factors

related to adsorption of chemicals to sediment constituents

include: organic content (Edwards 1970, Voice and Weber

7











1983, and Garbarini and Lion 1986), clay content (Johnsen

and Starr 1972), inorganic detrital content (Windom et al.

1989), and acidity (Edwards 1970). It is clear that

physical, chemical, and biological factors affect the

adsorption and persistence of chemical contaminants in

sediments.

One of the primary factors that influences the

adsorption process in aquatic environments is the water

solubility of the chemical. Nonpolar chemicals that have

extremely low solubility in water have a strong tendency to

adsorb to suspended matter and sediments. According to

Weber et al. (1983), sorption reactions between hydrophobic

pollutants and sediments or suspended solids in aqueous

systems are rapid and probably rate limited. Lopez-Avila

and Hites (1980) studied the transport of organic compounds

into sediments and found that the aqueous concentration of

the various compounds follows the rules of simple dilution

and that those compounds with the highest octanol-water

partition coefficient are strongly associated with suspended

particulate matter and sediment in the water.

Metals, in contrast to organic compounds, are often

introduced initially into the aqueous environment in

solution and accumulate on suspended sediments and on

inorganic and organic colloidal particles. Trace metal

contaminants from anthropogenic sources may also be

introduced in large particles (Windom et al. 1989). In











general, absorbents for metals in the aquatic environment

include clays, organic detritus, algae, and the hydrous

oxides of iron and manganese (Persaud and Lomas 1987).

Metals may also be coprecipatated with hydrous iron,

manganese oxides, and iron sulphides. Metals that are

adsorbed onto or coprecipitated with particulate matter tend

to accumulate in bottom sediments and may reach

concentrations that are very toxic to aquatic organisms

(Persaud and Lomas 1987).

Although the ultimate fate of adsorbed chemicals in

sediments is not clear, several researchers have shown that

adsorbed chemicals can be released from sediments. Oliver

(1985) investigated the desorption of chlorinated

hydrocarbons from sediments and concluded that: (1)

chemicals in bottom sediments can desorb into pore water and

subsequently diffuse into overlying waters and (2) bottom

sediment particles may be resuspended into the water column

where desorption of chemicals can occur.

The release of metals from sediments is governed by the

physicochemical conditions of the environment. When metals

are adsorbed onto particulates mainly by electrostatic

forces, there is a continuous exchange of cations between

the sorbing particles and the surrounding water (Persaud and

Lomas 1987).

There are numerous reports in the literature of

contamination of both marine and freshwater sediments.











Sediments contain a variety of pollutants that include:

pesticide residues (Miles and Harris 1978), aliphatic

hydrocarbons (Wakeham and Carpenter 1976, Solanas et al.

1982, and Grimalt et al. 1984), aromatic hydrocarbons

including chlorophenols and chloroguaiacols (Eder and Weber

1980, Xie 1983, and Catallo and Gambrell 1987),

polychlorinated biphenyls (Christensen and Lo 1986, Delfino

1979), and heavy metals such as lead mercury and cadmium

(Delfino 1979, Edgington and Robbins 1976). Such widespread

contamination has prompted research concerned with the

toxicological biomonitoring of polluted sediments.

Toxicity Bioassays

Polluted water and sediments are of concern because of

potential impacts to aquatic organisms. This concern has

led to the development of toxicity biossays that employ a

variety of test organisms. Toxicity biossays can provide

data on the short-term (acute) and long-term (chronic)

toxicity of contaminated samples to aquatic organisms.

(Rand and Petrocelli 1985). According to Buikema et al.

(1982), the selection of test organisms for any bioassay

should ideally be based on four criteria: 1) the organism

should be representative of an ecologically important group

in terms of taxonomy, trophic level, or niche; 2) the

organism should occupy a position within a food chain

leading to man; 3) the organism should be widely available,

amenable to laboratory testing, easily maintained and









11
genetically stable so uniform populations can be tested and

4) there should be adequate background data on the organism

in regards to its physiology, genetics, taxonomy and role in

natural environments. Additionally, the selection of tests

species should depend on the objective of the study (Buikema

et al. 1982).

Studies with the objective of assessing contaminated

sediments have been limited by the complexity of sediment-

water column and sediment-biotic interactions (Giesy and

Hoke 1989). Most of the early studies concerned with the

assessment of sediment quality were based on chemical

characterization of whole sediment concentrations of

inorganic and organic compounds (Burton et al. 1989).

However, chemical characterization does not in itself prove

an ecological hazard exists. Therefore, in recent years,

toxicity bioassays have been applied in the quality

assessments of sediments.

Characteristics for ideal sediment bioassays have been

proposed by several researchers. Giesy and Hoke (1989) made

the following recommendations: 1) the response of the

organisms should be predictable and constant; 2) the assay

organisms should have responses similar to many classes of

toxicants; 3) the results of the assays should be related to

ecologically relevant processes under field conditions; 4)

the results of the assays should be related to sediment or

water quality standards and criteria; 5) the bioassays











should be applicable to a number of sediment types and

environments; 6) the assays should be rapid, replicable,

inexpensive, and easily implemented; 7) the assays should be

standardized to facilitate widespread use; and 8) the assays

should be sensitive. It should be noted that these

recommendations refer to single species toxicity bioassays.

Such bioassays have generated valuable information that has

been used in the management of pollution and regulation of

discharges (Buikema et al. 1982).

Most single species toxicity tests used for sediment

assessments have used aquatic invertebrates that live in the

water column. The selection of such species is based on the

fact that exposure to sediment-associated pollutants for

many organisms is habitat related. When aquatic organisms

come in contact with sediment-associated pollutants,

biological effects may include such responses as mortality,

bioconcentration, reproductive impairments, pathological

effects, alteration in distribution or behavior, and

biochemical or physiological modifications that offset

various biological processes (Anderson et al. 1987). Many

of these responses have been the endpoints of a variety of

sediment toxicity bioassays employing invertebrates and fish

as noted in the following studies.

A variety of invertebrates as well as fish and

amphibian embryo-larval stages have been used in sediment

toxicity bioassays. Prater and Anderson (1977a) used











Hexagenia limbata, Asellus communis, Daphnia magna, and

Pimephales promelas in a 96-hour bioassay using whole

sediment samples from sites in Lake Superior harbors. The

same procedure was used in another study by Prater and

Anderson (1977b) of contaminated sediments from Otto Creek,

Ohio. Results of their studies indicated that sediment

associated chemicals can pose a threat to aquatic ecosystems

and sediment bioassays employing invertebrates and fish can

successfully be used in hazard assessments.

Wentsel et al. (1977 and 1978) exposed larvae of the

midge, Chironomus tentans to whole sediment samples

containing heavy metal contamination. Test endpoints for

the larvae were based on growth and adult emergence.

Results of the two studies indicated a good correlation

between the sizes of the midge larvae and the degree of

metal contamination in the sediment samples. These studies

led the researchers to conclude that chironomids are

sensitive indicators of the presence of toxic substances in

sediments. In other studies, chironomids have been used

successfully as indicators of acute toxicity of sediments

(Nebeker et al. 1984).

Birge et al. (1987) used freshwater fish and amphibian

embryo-larval stages to monitor sediment toxicity. The test

organisms were exposed to sediments in environmental

chambers where endpoints such as hatchability and mortality

were observed. Teratogenesis and toxicant bioconcentration









14
in the test organisms were also observed. These researchers

showed that the test organisms could be used in a simple

cost-effective test system and would give clear detectable

endpoints. In a similar study, Landrum et al. (1987)

exposed the amphipod Pontoporeia hovi to whole sediments in

static chambers. The invertebrate, which is a major

component of the benthic fauna in the Great Lakes, was used

to determine the bioavailability of organic contaminants in

the sediment samples. Results of the study revealed that

the test could be a good indicator of the bioavailability of

sediment-associated contaminants.

Several marine organisms have been used in bioassays to

determine marine sediment toxicity. Some of the organisms

used include: oyster larvae (Chapman and Morgan 1983), the

polychaete Nereis virens (McLeese et al. 1982), shrimp

Crangon septemspiriosa (McLeese et al. 1982), marine fish

(Swartz 1987), soft-shell clams (Chu-Fa et al. 1979), and

several other marine species (Swartz 1987). The study

conducted by Chu-Fa et al. (1979) assessed sediments in the

northern Chesapeake Bay using fish and the soft-shell clam

in acute 24-hour and 48-hour bioassays. Mortality of the

test organisms increased proportionately with concentrations

of metals and organic pollutants in the sediment samples.

These results give support to the rationale that the

use of invertebrates in sediment bioassays constitutes a

feasible and economical approach for assessing the potential











toxicological effects of polluted sediments on aquatic

biota. However, the use of a wide variety of invertebrates

in sediment toxicity testing has been limited by the lack of

culture methods for most species. According to Buikema et

al. (1982), from over 165 proposed test organisms for

toxicity testing, relatively few are invertebrates, and even

fewer have ever been cultured in the laboratory.

Consequently, standard stocks of aquatic invertebrates often

are not available throughout the year. Clearly there is a

need for continued basic research into the ecological

requirements of invertebrate species brought into the

laboratory for culture and sediment toxicity tests.

Bacterial Bioassays

Since sediment bioassays have been practiced for a

limited time, methodology continues to evolve with time and

research needs. Examples of toxicity bioassays which

diverge from the traditional bioassays employing

invertebrates and fish are bacterial bioassays. The use of

bacteria as assay organisms in toxicity tests is based on

the fact that bacteria have many biochemical and

physiological systems that are similar to higher organisms.

Therefore, when toxicants elicit a response in bacteria, it

is due to the interaction of the toxicant with biomolecules

that are similar in many different organisms (Giesy and Hoke

1989). The importance of bacteria as toxicity assay

organisms is also related to the key role that bacteria play











in the geochemical cycling processes of nutrients and the

fact that bacteria represent the first level at the base of

the food chain (Atlas and Bartha 1987).

In general, bacterial assays are suitable for use in

the rapid toxicity screening of aqueous samples because of

their ease-in-handling, speed, reproducibility of results

and low cost (Bitton and Dutka 1986, Liu and Dutka 1984).

Some endpoints of bacteria toxicity tests include: (1)

measurement of growth inhibition, (2) colony formation, (3)

oxygen uptake, (4) ATP content, (5) inhibition of

bioluminescence, (6) enzyme activity (Liu and Dutka 1984,

and Dutton 1988) and (7) inhibition of enzyme biosynthesis.

According to Bitton (1983), the various bacterial toxicity

screening tests can be divided into two main categories:

(1) assays based on the measurement of viability or growth

of specific bacteria or specific groups of bacteria; and (2)

assays based on the measurement of the effect of toxic

chemicals upon bacterial activity such as respiration and

heterotrophic potential.

Some of the earliest applications of bacteria as

indicators of environmental toxicity were made by Dawson and

Jenkins (1950) and Malaney et al. (1959). In these studies

manometric methods were used to determine the effect of

metal ion toxicity on the biological oxidation processes

which occurred at municipal wastewater treatment plants.

Over the years bacterial bioassays have continued to be











employed in the assessment of chemical contamination in

water but limited attention has been given to the

application of bacterial bioassays in toxicity assessments

of sediments.

Most bacterial bioassays applied in toxicity

assessments of sediments have been based on measurements of

the effect of sediments on the activity of indigenous mixed

bacterial populations (Burton and Lanza 1985). However, in

recent years, more attention has been given to the

application of the Microtox System for toxicity assessment

of sediment-associated contaminants (Chang et al. 1981,

Schiewe et al. 1985, and True and Heyward 1990).

The Microtox System (Microbics Corp. 1982) utilizes the

marine luminescent bacterium, Photobacterium phosphoreum, as

the bioassay organism for measuring acute toxicity in

aqueous samples (Bulich 1984). A suspension of the

bioluminescent organisms is exposed to several

concentrations of a toxic sample. The Microtox Toxicity

Analyzer is used to measure quantitatively the light output

of the organisms before and after they are exposed to the

toxicant sample. A reduction in light output reflects a

deterioration in the state of health of the organisms,

thereby signifying the presence of toxicants in the sample.

The use of a large population of bacteria gives Microtox

results statistical significance (Fuller and Warrwick 1985,

Bulich 1984).











In spite of the limited application of bacterial

bioassays in sediment toxicity testing, a number of studies

have focused on the toxicological effects of various

chemicals on bacteria. Many of these studies have shown

that bacteria can be sensitive indicators of toxicity.

Bauer et al. (1981) found that pentachlorophenol

affected dissolved oxygen depletion by a mixed bacterial

population during short-term exposure. They also found that

following a 22 hour exposure period, organic compounds such

as atrazine, trichloroacetic acid, diuron and

dimethylformamide were toxic to the bacteria with effects

ranging from slight to extreme. Murray and Guthrie (1980)

studied the response of aquatic bacterial populations to

insecticides such as sevin and malathion using respirometry.

They concluded that the method was a useful and rapid one

for detecting the effect of pollutants on aquatic microbial

populations. A study that tested the toxicity of methanol-

dichloromethane extracts of sediments that had been solvent-

exchanged into ethanol revealed that Photobacterium

phosphoreum was sensitive to aromatic and chlorinated

hydrocarbons (Schiewe 1985).

In comparative studies of bacteria with other toxicity

bioassay test organisms, variable responses to test

chemicals have been observed. Babich and Stotzky (1985)

found that bacteria in general are equally or less sensitive

to metals than are plant or animal cells. For example,











Photobacterium phosphoreum have been shown to be less

sensitive to both mercury and cadmium than is Daphnia magna,

but very sensitive to the toxic effects of copper (Giesy and

Hoke 1989). Remacle and Houba (1980) showed that cadmium

sensitive saprophytic bacteria do not grow in media

containing 8 mg/L cadmium, but some resistant strains could

tolerate concentrations of cadmium as high as 300 mg/L.

Bacteria have also been shown to be very tolerant to

some organic compounds which are extremely toxic to

crustaceans and fish. The LC50 for malathion is 1.8 g/L for

Daphnia magna while a solution of 10 g/L promoted growth of

aquatic bacteria (Giesy and Hoke 1989). This differential

response may be explained by the fact that malathion is an

organophosphate and its primary target is the cholinesterase

system which is absent in bacteria (Bauer et al. 1981).

Comparative toxicity studies of bacteria as well as

other bioassay test organisms have shown variable responses

to chemicals. Such responses are not unusual because there

are certain limitations of each individual biosystem.

Several researchers have concluded that data obtained from

toxicity tests of complex chemical mixtures become more

reliable when a battery of short-term tests are used (Dutka

et al. 1988, Dutka and Kwan 1988, Plokin and Ram 1984 and

Giesy and Hoke 1989).











Battery-of-Test Approach To Toxicity Testing of Sediments

The use of multiple bioassays in the toxicological

assessment of pollutants in aquatic environments has been

emphasized by numerous researchers (Dutka et al. 1988, Giesy

and Hoke 1989, Plokin and Ram 1984, Malueg et al. 1984).

Such an approach is emphasized because, as noted earlier,

single species bioassays give variable results due to

differing modes of action and different matabolic processes

in the test organisms. In addition, the sensitivity of an

ecosystem to pollutants is influenced by several factors

such as indigenous species sensitivity, physical and

biological transformation of the pollutants, and

environmental factors that affect various species

differently. (Burton et al. 1989). Therefore, bioassay

tests performed independently do not provide realistic

evaluations of areas of concern (Dutka et al. 1988).

Confidence in positive or negative results of toxicity

bioassays is increased when the data are confirmed in

different test systems using either different biological end

points or indicator organisms or different activation

systems (Bartsh et al. 1980).

In regards to sediment toxicity, a variety of indicator

organisms, representing the major classes of aquatic

organisms, have been used in the "battery of tests"

approach. Some of these organisms include: fathead

minnows, zooplankton, luminescent bacteria, spiral bacteria,











indigenous bacterial population, protozoa, midges,

amphipods, mayflies, oligochaetes, and green algae (Plotkin

and Ram 1984, Dutka and Kwan 1988, Burton and Stemmer 1988,

and Giesy and Hoke 1989). Results of several investigations

have emphasized that no one single species assay

consistently detects toxicant impacts. Therefore, a battery

of indicator tests is necessary for reliable ecotoxicity

assessments of sediments. Several studies give support to

this observation.

Giesy et al. (1988) evaluated the toxicities of Detroit

River whole sediments and sediment pore water using Daphnia

macrna, Photobacterium phosphoreum, and Chironomus tentans.

The results of this study showed that the use of several

assay organisms increases the probability of correctly

identifying sediments that would be expected to be toxic to

a community of benthic organisms under field conditions.

Burton and Stemmer (1988) used a battery of tests which

consisted of microbial activity assays, macrofaunal assays,

amphipod assays and the fathead minnow assay. Waters,

leachates and sediments were tested for toxicity from sites

in five states. Results of the microbial activity tests

were significantly correlated with stream profiles of

biological and chemical parameters.

Burton et al. (1989) used five assays to evaluate the

toxicity of sediments taken from Waukegan Harbor, Illinois

and Indiana Harbor sites. The assays included: the Daphnia











magna and Hyalella azteca 48-hour survival assays,

Selenastrum capricornutum 48-hour growth inhibition assay,

and an indigenous microbial activity test. The data

collected also supported the premise that multiple test

assays are necessary to both detect sediment toxicity and

differentiate degrees of toxicity to various organisms.

Additionally, the results demonstrated the importance of

conducting several toxicity tests using organisms from

different trophic levels to assess the potential impact of

contaminated sediments.

Dutka et al. (1988) conducted a study to evaluate the

suitability of a variety of microbiological, biochemical,

and toxicant screening tests. The tests employed in their

study were used to identify contaminated sediments and water

in the St. John River Basin, Canada. Results indicated that

the Microtox test was one of the least sensitive screening

tests used in the study. Such results reemphasize the fact

that individual bacteriological and toxicant screening tests

do not provide sufficient data for realistic

ecotoxicological management decisions, while the "battery of

tests" approach can facilitate the prioritization of further

investigations that could lead to more realistic management

decisions.

In a comprehensive review of procedures used in

assessing sediment toxicity, Giesy and Hoke (1989) proposed

the use of a battery of assays which consists of tests which











have a range of sensitivities, are simple, reproducible,

inexpensive, ecologically relevant and can be related to

regulatory criteria. Specific tests such as the Microtox

assay, the 10-day Chironomus tentans growth assay, and the

48-hour Daphnia magna acute lethality assay were recommended

for inclusion in the battery of assays. This approach

enables larger numbers of sediments to be analyzed and

decreases the probability of incorrectly identifying a toxic

sediment as a non-toxic sediment (Giesy and Hoke 1989).

The investigations reported here represent some of the

current knowledge concerning the use of bioassays for the

assessment of sediment toxicity. It is evident that a

number of assays can be used for the toxicity assessment of

sediments. Many of the assays have characteristics that

favor their inclusion in the "battery of tests". Based on

several studies, this approach seems to be the most

appropriate and reliable means of assessing sediment

toxicity. This approach applies a variety of assays, that

may use tests organisms from several trophic levels, in the

toxicological screening of sediments. Practical application

of the "battery of tests" approach in sediment quality

assessments can facilitate the prioritization of further

investigations and permit the more judicious use of more

expensive procedures (Giesy and Hoke 1989). The general

consensus among researchers is that the "ideal" sediment

toxicity bioassay or "battery of assays" has not been











developed. Therefore, there is still a need for continued

research focused on sediment toxicity bioassays.

Rationale For The Use of the B. licheniformis
Inhibition of Enzyme Biosynthesis Assay
In Toxicity Screening of Sediments

As noted earlier, bacteria are suitable as bioassay

organisms because they are inexpensive to cultivate, grow

rapidly, and contain physiological and enzymatic processes

also found in higher organisms (Liu and Dutka 1984). Most

bacterial toxicity assays can be divided into three

categories: 1) assays based on bacterial luminescence, 2)

assays based on the measurement of viability or growth, and

3) ecological effect assays. Bacterial luminescence and

viability or growth assays typically use gram-negative

bacteria (i.e. Photobacterium phosphoreum and Escherichia

coli) which possess a thick outer cell membrane consisting

of lipopolysaccharides and lipoproteins (Van Demark and

Batzing 1987). The outer membrane helps protect these

bacteria from certain chemicals by acting as a hydrophobic

barrier (Bitton et al. 1988).

Ecological effects assays usually involve the isolation

of indigenous bacterial populations and the measurement of

some metabolic end product (C02, 02, enzymes) after exposure

to a toxicant (Burton and Stemmer 1988). Results of such

assays may show variable sensitivities, especially if the

indigenous bacterial populations were isolated from areas

which had received waste inputs from urban or industrial









25
sources (Barnhart and Vestal 1983). There is evidence that

spontaneous mutants with resistance to certain xenobiotics

can develop in natural populations exposed to toxic

environmental contaminants (Atlas and Bartha 1987).

Bacillus licheniformis is different from the typical

bacterial species used in bioassays. This organism is a

gram positive bacterium lacking the outer membrane barrier

found in gram negative bacteria. In gram positive bacteria

the cell wall is an exposed organelle that may offer the

first encounter between a bacterium and a molecule in its

environment (Collins and Stotzky 1989). The absence of the

outer membrane layer in Bacillus licheniformis should make

the organism more sensitive to certain hydrophobic toxicants

than a gram negative bacterium.

Dutton (1988) investigated the usefulness of enzyme

biosynthesis in bacteria as a basis for toxicity testing

using beta-galactosidase and tryptophanase biosynthesis in

Escherichia coli and alpha-glucosidase biosynthesis in

Bacillus licheniformis. He showed that the assay based on

inhibition of alpha-glucosidase in Bacillus licheniformis

was more sensitive than inhibition of tryptophanase or beta-

galactosidase in Escherichia coli. The differences were

attributed to differences in cell permeability.

The Bacillus licheniformis bioassay is based on

inhibition of alpha-glucosidase biosynthesis in the

bacterium. Alpha-glucosidase is an extracellular enzyme









26
induced by the presence of maltose in the environment of the

cell. The enzyme degrades maltose into glucose residues

acting at the 1, 4-alpha-glycoside linkages. The formation

of the enzyme is initiated when genetically regulated

repressors are inactivated (Dutton 1988).

The basic steps of the assay according to Dutton (1988)

are: 1) cell growth and preparation (washing), 2) cell

exposure to toxicants, 3) enzyme induction and 3)

colorimetric measurement of enzyme production. During the

assay, induction follows exposure of cells to toxicants;

therefore enzyme production at the beginning of the toxicant

contact period is negligible. At the end of the exposure

period, an inducer is added and additional time is allowed

for biosynthesis of alpha-glucosidase. Differences between

final enzyme levels are attributed mainly to the toxicant

action on the operon for alpha-glucosidase biosynthesis

(Dutton 1988).

Application of the assay to environmental samples has

not been done and research using this assay to test

environmental samples such as sediments is needed. The

Bacillus licheniformis bioassay has the potential to be more

sensitive than other bacterial assays that have been applied

in the toxicity screening of environmental samples. The use

of a pure culture of cells makes the assay dependable.

Also, the assay can be completed in a relatively short











period of time, and it is inexpensive when compared with

other bacterial bioassays such as the Microtox assay.

Rationale for the use of Chvdorus sphaericus in
Sediment Toxicity Tests
Out of over 165 species of organisms that have been

proposed for use in toxicity bioassays, only a few are

invertebrates and even fewer have ever been cultured in the

laboratory (Buikema et al. 1982). Many of the invertebrates

that have been applied in sediment toxicity tests are not

benthic organisms and possess few characteristics of the

"ideal" sediment bioassay organism. Some tests species have

limited ecological ranges (i.e. Daphnia magna); some may not

be widely available for testing; and many are not easily

maintained in the laboratory. Genetically stable and/or

uniform populations are often difficult to obtain (Buikema

et al. 1982). In addition, some traditional sediment

toxicity tests utilize organisms that spend no part or only

part of their life cycle in contact with sediment

constituents, and therefore lack, in some degree, biological

realism.

Oligochaetes, midges, mayflies and amphipods have been

used most often as assay organisms in freshwater sediment

toxicity bioassays (Nebeker et al. 1984). The cladoceran,

Daphnia magna has also been used (Giesy et al. 1985). The

use of these organisms has generated information that has

been useful in assessing sediments. However, several

drawbacks are associated with these assays.









28
Oligochaetes are difficult to isolate initially from

sediments and difficult to handle without injury in toxicity

tests (Buikema 1982). Similar difficulties may be

encountered with amphipods in regards to collecting enough

organisms of the appropriate age. Viable laboratory

populations of amphipods are difficult to maintain, and

culture and the long life cycle of most amphipods can delay

tests results (Giesy et al. 1988). Delayed test results are

also associated with the use of midges and mayflies in

sediment toxicity assays. The use of Daphnia magna in

sediment toxicity tests lacks biological realism because in

nature the organism inhabits the water column and may rarely

come in contact with sediment constituents.

Little attention has been given to investigations that

apply a variety of benthic invertebrate species in toxicity

tests, and sediment toxicological studies using benthic

cladocera are virtually nonexistent. Chydorus sphaericus is

a benthic cladoceran that occurs abundantly associated with

submerged macrophytes and sediments of lakes and ponds. The

species exists in a wide range of environments from arctic-

alpine regions to the tropics and is often considered to be

ubiquitous in its distribution (Frey 1979). The organism

ranges in size from 0.1-0.2mm (Haney 1973) and has modified

legs that are somewhat prehensile in scraping up larger

pieces of sediments or detrital material. Filter feeding

also has been noted (Wetzel 1983). Reproduction occurs











parthenogenetically and continuously during most of the

year. In many instances, populations reach high numbers in

environments of extreme eutrophication. Parthenogenic

females carry no more than two eggs in their brood pouch and

young are released every two days. In nature, this

reproductive rate is dependent upon several factors,

especially food availability, temperature, and age of the

organism (Shan 1968). Under low food conditions, limnetic

individuals of Chydorus sphaericus have been observed

carrying one egg. Also, older organisms often produce one

egg (Keen 1973). The released young pass through two non-

reproductive instars, becoming reproductive about the fourth

day after they have molted to the third instar. Thus, the

development from a parthenogenetic egg to an adult female

requires about six days. The life span of the organism

under ideal conditions at 250C is 9-24 days (Shan 1969).

Although satisfactory culture methods for several

species of chydorid cladocera have been worked out (Shan

1967), studies applying Chydorus sphaericus in sediment

toxicity tests are nonexistent. Based on general

characteristics, several advantages can be associated with

the use of Chvdorus sphaericus in sediment toxicological

studies. Some of these advantages are: 1) a genetically

stable and uniform population can be obtained because the

organism reproduces primarily by parthenogenesis; 2) testing

at 25C is possible because laboratory cultures have been











maintained at this temperature (Shan 1967); 3) smaller

volumes of test solutions or sediments can be used because

of the small size of the organism; and 4) some degree of

biological realism can be incorporated into a sediment

toxicity bioassay using C. sphaericus because the organism

is associated with sediments in nature. Advantages

associated with traditional tests should be considered if

the organism is used in static acute toxicity tests.

Traditional static acute toxicity tests are usually rapid,

simple and inexpensive, requiring a minimum amount of

laboratory space (Buikema et al. 1982). It is evident that

an investigation of Chydorus sphaericus as a test organism

in sediment toxicity studies is relevant.















CHAPTER 3
TOXICITY TESTING OF SEDIMENT ELUTRIATES BASED ON INHIBITION
OF ALPHA-GLUCOSIDASE BIOSYNTHESIS IN
BACILLUS LICHENIFORMIS


Introduction

The role of aquatic sediments as a repository for and

source of large quantities of contaminants has led to the

development of a wide variety of bioassays for the

toxicological assessment of sediments. Although there have

been limited applications of bacterial bioassays in

assessments of environmental samples such as sediments,

recent advances in toxicity testing using bacteria have

contributed information about bacterial characteristics that

make their application in sediment bioassays feasible.

Bacteria are suitable for use in the rapid screening of

environmental toxicity mainly because of their ease of

handling, speed, reproducibility of results and low cost.

Endpoints of bacterial bioassays include: (1) measurement

of growth inhibition; (2) colony formation; (3) oxygen

uptake; (4) ATP content; (5) inhibition of bioluminescence

and (6) enzyme activity and biosynthesis (Bitton and Dutka

1986). The endpoints of the bacterial assays used in this

study are inhibition of enzyme (alpha-glucosidase)

biosynthesis and inhibition of bioluminescence.











A comprehensive study of the effectiveness of enzyme

biosynthesis in bacteria as a basis for toxicity testing

conducted by Dutton (1988) showed that the bioassay based on

inhibition of alpha-glucosidase biosynthesis in Bacillus

licheniformis was more sensitive to certain toxicants when

compared to similar bioassays using gram negative bacteria.

Differences in sensitivity were attributed to differences in

cell permeability.

Bacillus licheniformis is a gram positive bacterium

that lacks the outer lipopolysaccharide membrane barrier

found in gram negative bacteria. Absence of the outer

membrane allows easier interaction between the bacterium and

toxicant molecules in the cell environment, thus increasing

the cell sensitivity to toxicants.

Biosynthesis of the extracellular enzyme alpha-

glucosidase in B. licheniformis is induced by the presence

of maltose in the cells growth medium. The enzyme degrades

maltose into glucose residues acting at the l,-4-alpha-

glycoside linkage (Dutton 1988). Biosynthesis of the enzyme

is initiated when genetically regulated repressor genes are

inactivated by the presence of maltose in the cell

environment. The assay consists of exposing a suspension of

cells to a toxicant. At the end of the exposure period, the

inducer, maltose, is added and additional time is allowed

for biosynthesis of alpha-glucosidase. At the end of

biosynthesis, a chromogen substrate is added. Enzyme











hydrolysis of the chromogen results in the liberation of a

product which gives the medium an amber color. Variations

in color intensity, which reflect differences in enzyme

levels, are measured with a spectrophotometer. Differences

in final enzyme levels are mainly attributed to toxicant

action on the metabolism of the cells (Dutton 1988).

The sensitivity of this assay combined with other

advantages of bacterial bioassays makes this assay ideal for

testing environmental samples such as sediment elutriates or

extracts. An elutriate can be defined as the

centrifuged/filtered clear liquid containing the soluble

fraction of a sediment sample (Ludwig et al. 1989).

The standard elutriate test was jointly developed in

the early 1970s by the U.S. Army Corps of Engineers and the

U.S. Environmental Protection Agency to monitor the soluble

release of contaminants into the water column during open-

water disposal of dredged sediments (Ludwig et al. 1989,

Daniel et al. 1989). This approach is used in sediment

studies to simulate processes that might disturb the

sediment and bring contaminants into the water column (Ross

and Henebry 1989). Toxicity testing of sediment elutriates

is important because dissolved forms of pollutants are more

bioavailable to aquatic biota for uptake and are the primary

cause of adverse impacts in aquatic ecosystems (Ludwig et

al. 1989).











The primary objective of the study reported in this

chapter was to use the alpha-glucosidase biosynthesis assay

to screen sediment elutriates prepared from sixty-six

sediment samples collected from various sites in Florida.

The same elutriate samples were also screened using the

Microtox (Microbics Corp. 1982) assay. Statistical

comparisons were made between the percentage inhibitions of

alpha-glucosidase biosynthesis and bioluminescence in the

Microtox assay. Analyses of concentrations of selected

metals in the elutriates were also made in an attempt to

correlate metal concentrations with percentage inhibitions

in the alpha-glucosidase bioassay.

Materials and Methods

Test Chemicals. Reagents and Media

Unless otherwise indicated, all chemicals and reagents

were obtained from Sigma Corporation, St. Louis Missouri.

The Z-buffer solution used in the alpha-glucosidase

biosynthesis assay contained the following components:

Na2PO4-7H2O, 16.1 g/L; Na2H2PO4H20, 5.5 g/L; KC1, 0.75 g/L;

and MgSO4-7H20, 0.25 g/L. Para-nitrophenyl-alpha-D-glucoside

chromogen solution was prepared by dissolving 0.5 g in 100

mL of Z-buffer. This solution was filter sterilized and

stored at 40C in an amber bottle. Other reagents used in

the assay were sodium dodecyl sulfate (SDS) 0.7% (w/v) and

Na2CO3, IM.










The growth medium for B. licheniformis contained

trypticase soy-broth without dextrose, 27.5 g/L, yeast

extract, 5 g/L and polyoxyethylene sorbitan monooleate

(Tween 80) 10 g/L (w/v). The inducer solution, maltose, was

dissolved in distilled water (4%; w/v) and then autoclaved.

Sediment Samples

Elutriates were prepared from sixty-six sediment

samples that were collected from Hillsborough (HIL),

Escambia (ESC), Palm Beach (PAL), and Monroe (MON) counties,

Florida. The samples were collected during the summer of

1989 by the "toxics project" group of the Department of

Environmental Engineering Sciences, University of Florida,

Gainesville, Florida. A description of the sites and their

location by latitude and longitude are presented in Table 3-

1. Sediments were collected by Ponar dredge to a maximum

depth of 2-8 cm. The sediments were mixed in an enamel

coated pan, placed in plastic bags, stored on ice and

transported to the laboratory where they were stored at

40C. Elutriates of the sediment samples were prepared

within 30 days of collection.

Preparation of Sediment Elutriates

The methods developed for evaluating toxicity of dredged

sediment (Nebeker et al. 1984) were modified and used to

prepare the sediment elutriates. The steps of the procedure

summarized in Figure 3-1 involved (1) mixing 25 grams (wet

weight) of sediment with 100 mL of deionized water (Milli-Q











water, Millipore Corp., New Bedford, MA); (2) shaking the

sediment-water slurry on a Eberbach Shaker overnight for 18

hours at low speed; (3) blending the mixture for five

minutes on high speed in a Waring blender (stainless steel);

(4) centrifugation of the sediment slurry for 15 minutes at

8,000 revolutions per minute and (5) separation of the

supernatant from the sediment. The decanted supernatants

were stored in deionized water-rinsed plastic bottles at 4C

until used in the screening tests.

Test Bacteria

Stock cultures of Bacillus licheniformis (strain 749)

were used in the alpha-glucosidase assay. Original cultures

were obtained from the Bacillus Genetic Stock Center (BGSC

#5A20), Ohio State University, Columbus, OH. The strain was

maintained in 40% glycerol at -150C.

Lyophilized cultures of the marine bacterium

Photobacterium phosphoreum were reconstituted according to

procedures outlined in the Microtox Operating Manual

(Microbics Corp.1982) for use in the Microtox assay.

Assay for Alpha-Glucosidase Biosynthesis

Optimal assay conditions were based on previous work

conducted by Dutton (1988). Preliminary experiments to

determine optimum conditions for enzyme production showed

that cultures grown to an absorbance greater than 0.700, at

550 nm gave higher enzyme levels. Furthermore, the use of

SDS in the assay caused the release of more enzyme. These











I. MIX sediment with deionized water
in a volumetric sediment-to-water
ratio of 1:4



Nj

II. SHAKE the sediment slurry
overnight (18 hours) on a
shaker


III. BLEND the sediment slurry
in a Waring Blender on high
speed for 5 minutes




IV. CENTRIFUGE the blended sediment slurry
15 minutes at 8,000 rpms


V. DECANT the supernatant carefully


Fig. 3-1. Protocol for preparing sediment elutriates.












Table 3-1. Location by latitude and longitude and
description of sites where sediment samples were collected.


SITE LAT./LONG. DESCRIPTION


ESC-05-01




ESC-05-02






ESC-05-03






ESC-05-04


30 23 30 / 87 13 00




30 23 25 /87 13 43






30 23 32 / 87 13 52






30 23 45 /87 14 08


America Creosote Corp./
Penascola Bay near Main
St. STP effluent boil
[7.3 m deep]

American Creosote
Corp./Pensacola Bay at
Light No. 2 near
end of Bayou Chico
Approach Channel [3.9 m
deep]

American Creosote
Corp./Pensacola
Bay at CM 4 in
Bayou Chico
Approach Channel
[4.8 m deep]

American Creosote
Corp./Pensacola Bay
at CM 6 in Bayou Chico
Approach Channel
(4.8 m deep]


ESC-05-10


ESC-05-05





ESC-05-06


Field Duplicate of
ESC-05-04


30 21 57 /87 14 21





30 23 50 /87 14 25


American Creosote
Corp./Pensacola Bay at
CM 8 in Bayou Chico
Approach Channel
[3.6 m deep]

American Creosote
Corp./Pensacola
Mouth of Bayou
Chico at Hwy 292
(Barrancas Ave.) Bridge
[3.6 m deep]











Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


ESC-05-07







ESC-05-08





ESC-05-09







ESC-06-01




ESC-06-02




ESC-06-03




ESC-06-04


30 23 58 / 87 14 11







30 24 02 / 97 14 05





30 23 59 / 87 14 16







30 25 00 / 87 15 20




30 24 45 / 87 15 35




30 24 45 / 87 15 45




30 24 00 / 97 15 60


American Creosote
Corp./ Pensacola
Bay E of Mouth of Bayou
Chico at End of Drainage
Ditch from Am. Creosote
Site [0.6 m deep]


American Creosote
Corp./Pensacola Bay off
E End of Sanders Beach
60m E of Station 07
[0.9 m deep]

American Creosote
Corp./Pensacola; Mouth of
Mouth of drainage ditch
from Am. Creosote Site at
Entry to Penascola Bay
adj. USGF Gaugeing Sta.
[0.75 m deep]

Reichold Chemical Co./
Pensacola NE Arm Bayou
Chico 150m W of "W" St
[0.9 m deep]

Reichold Chemical Co./
Pensacola NC Arm Bayou
Chico 200m N of US 98 Br.
[1.5 m deep]

Reichold Chemical Co./
Pensacola NW Arm Bayou
Chico W of Sta. 02
[0.9 m deep]

Reichold Chemical Co./
Pensacola S of Sta.
02 Mid-Way btw. US 98 and
Frisco RR Bridges
[1.5 m deep]











Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


ESC-06-05


30 24 30 / 87 15 30


Reichold Chemical Co./
Pensacola Bayou Chico S
of Frisco RR Br. 14.2 m
deep]


ESC-06-10


ESC-06-06




ESC-06-07




ESC-06-08




ESC-06-09




ESC-07-01



ESC-07-02





ESC-07-03


Field Duplicate of
ESC-06-05


30 24 15 / 87 15 25




30 23 55 / 87 16 05




30 24 00 / 87 14 50




30 23 55 / 87 14 45




30 28 33 / 87 21 25



30 28 05 / 87 21 46





30 28 05 / 87 21 55


Reichold Chemical Co./
Pensacola Middle of Bayou
Chico at CM 17 [2.4 m
deep]

Reichold Chemical Co./
Pensacola SW Arm Bayou
Chico 200m W of Old
Coryfield Rd [0.7 m deep]

Reichold Chemical Co./
Pensacola SE Arm Bayou
adj. Auto Shredding Co.
[4.2 m deep]

Reichold Chemical Co./
Pensacola Bayou Chico W
of Hwy 292 (Barrancas
Rd.) Br. [4.5 m deep]

Champion Paper Co./Eleven
Mile Cr. W of USN Saufley
Field [1.8 m deep]

Champion Paper Co./Eleven
Mile Cr. Eleven Mile Cr.
d/s Sta 01 just u/s
Confl. with Hurst Br. Cr.
[0.6m deep]

Champion Paper Co./Eleven
Mile Cr. Eleven Mile Cr.
d/s Sta. 02 on inside of
wide curve [4.5 m deep]












Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


ESC-07-04




ESC-07-05





ESC-07-06




ESC-07-07





ESC-07-08




ESC-07-09


30 27 22 / 87 22 36




30 27 00 / 87 22 15





30 34 59 / 87 19 42




30 34 22 / 87 19 18





30 34 21 / 87 19 18




30 32 29 / 87 19 48


Champion Paper Co./Eleven
Mile Cr. Mouth of Eleven
Mile Cr. at Perdido Bay
[2.1 m deep]

Champion Paper Co./Eleven
Mile Cr. 200 m into
Perdido Bay SE of mouth
of Eleven Mile Creek
[1.5 m deep]

Champion Paper Co./Eleven
Mile Cr. NW Tributary to
Eleven Mile Cr. at Hwy
297A Br. [0.4 m deep]

Champion Paper Co./Eleven
Mile Cr. 15 m N of Hwy 86
Br. just u/s of
Cantonment STP Outfall
[0.75 m deep]

Champion Paper Co./Eleven
Mile Cr. 100 m d/s Hwy 86
Br. at Champion Discharge
"Boil" [0.9 m deep]

DuBose Oil Prod./Jacks
Br. Perdido R. 100m d/s
Hwy 90 Br. [5.1 m deep]


ESC-07-10


ESC-08-01



ESC-08-02


Field duplicate of
ESC-07-05


30 31 17 / 87 26 51



30 27 00 / 87 23 21


DuBose Oil Prod./Jacks
Br. Perdido R. 100m d/s
Hwy 90 Br. [5.1 m deep]

DuBose Oil Prod./Jacks
Br. Mouth of Perdido R.
at Perdido Bay
[2.4 m deep]












Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


ESC-08-03





PAL-09-01




PAL-09-02




PAL-09-03




PAL-09-04



PAL-09-05




PAL-09-06




PAL-09-07



PAL-09-08


PAL-09-09


30 37 12 / 87 22 30





26 47 05 / 80 31 31




26 47 05 / 80 31 29




26 47 21 / 80 31 59




26 47 11 / 80 31 45



26 46 59 / 80 31 30




26 46 59 / 80 31 30




26 46 58 / 80 31 29



26 45 50 / 80 31 20


26 45 50 / 80 29 57


DuBose Oil Prod./Jacks
Br. Drainage Ditch from
Dubose at rear of
Whitehurst Property 701
Hwy 97 [7.6 cm deep]

Chem Air Spray/WPB Canal
L-10 Drainage Ditch N of
Chem Air Spray Bldg
[0.75 m deep]

Chem Air Spray/WPB Canal
L-10 Drainage Canal E of
Sta. ol E of Culvert
[0.6 m deep]

Chem Air Spray/WPB Canal
L-10 800 m NW u/s of
Bldgs. adj. Drainage
Ditch [4.5 m deep]

Chem Air Spray/WPB Canal
L-10 400 m SE d/s of Sta.
03 [4.5 m deep]

Chem Air Spray/WPB Canal
L-10 L-10 SW of Site Near
Pump House on SW Shore
[4.5 m deep]

Chem Air Spray/WPB Canal
L-10 Outlet of Drainage
Canal S of Sta. 02
[4 m deep]

Chem Air Spray/WPB Canal
L-10 L-10 20 m SE d/s of
Sta. 06 [4.5 m deep]

Chem Air Spray/WPB Canal
L-10 L-10 200 m SE d/s
Sta. 07 [5 m deep]
Chem Air Spray/WPB Canal
L-10 3200+ m SE d/s Sta.
0.8 at Small Br.
[1.75 m deep]











Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


PAL-09-10


MON-12-02



MON-12-03



MON-12-04


MON-12-05


MON-12-06


MON-12-07


MON-12-08




MON-12-09




MON-12-10


HIL-02-01


26 46 58 / 80 31 30


24 33 42 / 81 48 00



24 33 45 / 81 48 00



24 33 55 / 81 48 03


24 33 25


/ 81 48 40


24 32 86 / 81 48 51


24 34 33


/ 81 45 06


24 33 39 / 81 44 27




24 33 46 / 81 44 12




24 33 39 / 81 44 27


27 57 25 / 82 23 05


Field Duplicate of
PAL-09-06

Key West/Trumbo Pt. Tanks
Key West Bight Channel E
of Pier D-1 [7.8 m deep]

Key West/Trumbo Pt. Tanks
USCG Station E of Pier
D-2 [10 m deep]

Key West/Trumbo Pt. Tanks
USCG Station E of Pier
D-3 [9.5 m deep]

Key West/Ocean Key House
Basin at SW End of Duval
St. [4.8 m deep]

Key West/Truman Annex
Basin Mid-Basin SW of
Bulkhead Construction
[10 m deep]

Key West/Stock Island
Landfill & Marina Marina
Mid-Basin SE of Landfill
[9 m deep]

Key West/Stock
Island/Safe Harbor Safe
Harbor Slip W of Alex's
Junk Yard [9 m deep]

Key West/Stock
Island/Safe Harbor
Mid-Basin 200 m W of
Power Plant [9 m deep]

Field Duplicate of
MON-12-08

Florida Steel Corp.
SW Property Boundary on
CSX RR Property [dry
ditch soil]











Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


HIL-02-03




HIL-02-06




HIL-02-10


HIL-02-11


HIL-03-01





HIL-03-03





HIL-03-04




HIL-03-06



HIL-04-02


27 57 25 / 82 22 52




27 57 25 / 82 21 55




27 57 35 / 82 23 38


27 57 36 /


82 23 37


27 58 57 / 82 21 11





27 57 45 / 82 22 05





27 57 41 / 82 22 05




27 57 35 / 82 23 38



27 52 57 / 82 32 00


Florida Steel Corp. S
Property Fence at Survey
Stake CN34c
[dry ditch soil]

Florida Steel Corp.
SE Property Boundary on
CSX RR Property
[dry soil]

Field Duplicate of
HIL-02-08

Drainage Ditch d/s
Sta. 03 [11 cm deep]

Tampa Bypass Canal/Six
Mile Creek U/s of
SR 574 Br. at Flow
Control Structure
[8 m deep]

Tampa Bypass Canal/Six
Mile Creek D/s Sta. 02
adj. Stauffer Chemical
[7.8 m deep]


Tampa Bypass Canal/Six
Mile Creek D/s Sta. 03
adj. Stauffer Chemical
pond [8.5 m deep]

Drainage Ditch at SW
Corner of FMC Property
[7 cm deep]

Old Tampa Bay
Between E Shore and
First Island near
Abandoned Westinghouse
Facility [3.8 m deep]











Table 3-1 continued.


SITE LAT./LONG. DESCRIPTION


HIL-04-06






HIL-04-07






HIL-04-08





HIL-04-09







HIL-04-11


27 53 11 / 82 31 38






27 53 12 / 82 31 48






27 53 11 / 82 31 23





27 53 11 / 82 31 35







27 57 38 / 82 23 25


Roto Rooter Drain and
Sewer Service Culvert W
Side of Westshore Blvd
across Street from SM
Boundary of Roto Rooter
Facility [0.2 m deep]


Drainage Ditch adj.
Tyson Ave. Terminus
Drainage Ditch from
Rooter Facility ca.
NE of Sta. 01
[2.5 cm deep]


5001
of
Roto
100 m


Roto Tooter Drain and
Sewer Service Drainage
Ditch 100 m u/s of Roto
Rooter SE Property
Boundary [5 cm deep]

Roto Rooter Drain and
Sewer Service Drainage
Ditch at Entry Point
to Buried Culvert 60 m
East of Westshore Blvd.
adj. S Boundary of Roto
Rooter [ 0.15 m deep]

Drainage ditch on NE
side of N 57th St.
[5 cm deep]


Source: Delfino 1989











conditions were used in the assay protocol outlined in

Figure 3-2. The bacteria were grown overnight in the growth

medium that had been inoculated with 50 uL of stock glycerol

culture. The cultures were incubated in a shaking incubator

(100 rpm) at 290- 30C. The overnight cultures were diluted

with fresh growth medium to A550=0.2 and re-incubated for

growth up to A55,=0.750- 0.800.

The basic steps of the assay include: (1) cell growth;

(2) washing of cells; (3) exposure to toxicant (sediment

elutriate at 45% dilution); (4) induction of enzyme

biosynthesis; and (5) assay of alpha-glucosidase by

measuring the absorption at 420 nm on a spectrophotometer

(Spectronic 20) of the p-nitrophenol that is liberated by

enzyme hydrolysis of the chromogen, p-nitrophenyl-alpha-D-

glucoside.

The Microtox Assay

Reconstituted cultures of Photobacterium phosphoreum

were exposed to a 45% dilution of the elutriate samples.

The assays were carried out at 150C with a 15 minute contact

period using standard procedures outlined in the Beckman

Microtox System Operating Manual (Microbics Co. 1982).

Reduction in bioluminescence was measured after the 15

minute incubation period with a Microtox Toxicity Analyzer

(Model 2500, Microbics Co., Carlsbad, CA). Ten elutriate

samples that gave percentage inhibitions of bioluminescence

greater than 50% were further tested in definitive assays to










o CELL GROWTH: grow Bacillus licheniformis
overnight at 30uC


Nt

o CELL PREPARATION: dilute cells with
fresh medium to absorbance @550 to 0.2;
allow to grow up to absorbance @550 > 0.7;
wash cells once in distilled water.


'V

o EXPOSURE TO TOXICANT: add 0.9 mL toxicant
to 1.0 mL cells; incubate for 30 min.


'V
o ENZYME INDUCTION: add 0.4 mL of buffer,
0.1 mL maltose and 0.5 mL fresh medium;
incubate for 60 min.


'V

o ALPHA-GLUCOSIDASE MEASUREMENT: add 0.2 mL of
chromogen solution and incubate
until color develops (30-60 min);
stop reaction with 1 mL of sodium carbonate;
measure absorbance at 420nm.









Figure 3-2. Summary of the B. licheniformis alpha-
glucosidase biosynthesis (AGB) assay.









48
determine their EC50s (the concentration of extract causing

a 50% reduction of bioluminescence). Four to eight

dilutions were used to make this determination.

Metal Analyses

Total concentrations of lead, cadmium, zinc and copper

were determined for each elutriate sample using a Perkin-

Elmer atomic absorption spectrophotometer, Model 5000.

Single element lamps were used, and E.P.A. protocols (U.S.

EPA 1983) were followed. Concentrations of metals were

determined against a standard curve calculated from known

absorbances and concentrations using linear regression

analysis.

MetPAD

MetPAD is an assay developed at the Department of

Environmental Engineering Sciences, University of Florida,

Gainesville, Florida. It is specifically sensitive to

metals in aqueous samples. Components of the assay include

freeze-dried bacteria (Escherichia coli), a diluent, buffer

and an assay pad containing an enzyme substrate. The

bacteria are reconstituted in distilled water, and 0.1 mL

of the bacterial suspension is incubated with 0.9 mL of the

toxicant sample. After incubation, the mixture is buffered,

and 10 ul drops are applied to a yellow pad containing an

enzyme substrate. Results are read by observing the change

in color on the yellow pad. The range is from purple (non-

toxic) to white (very toxic) (TREEO News 1989).









49
This assay was used to determine if elutriate toxicity

observed with the Microtox and alpha-glucosidase

biosynthesis assays were due to metals in the samples.

MetPAD testing of several very toxic and non-toxic elutriate

samples were done before and after filtration through a

metal chelating resin to confirm the presence or absence of

metals in the elutriate samples. Metal chelating resin

columns were prepared for each elutriate sample by packing

0.2-0.4 g of resin (Sigma Corp. St. Louis, MO) in flint

glass pasteur pipets (4 mL volume, 150 mm length) without

tips. Two-inch rubber tubes with screw clamps were attached

to the pipets to control the flow of elutriate through the

column. The metal chelating resin was preconditioned by

passing 25 mL of Milli-Q water through the column.

Following this step, 5 mL of 1M NaCI was passed through the

columns. Each elutriate sample was passed through a

separate column. The filtrate was collected and assayed

using MetPAD. Results of the MetPAD assay indicated metal

toxicity in several of the elutriate samples and thus led to

the determination of metal concentrations in all sixty-six

elutriates.

Data Analysis

The degree of inhibition of enzyme biosynthesis by

sediment elutriates was determined by measuring absorbance

with respect to a control (distilled water). Absorbances of

blanks were subtracted from the elutriate samples absorbance











readings to correct for background enzyme or elutriate

sample color. Blanks consisted of all assay components

except the inducer. EC50s were derived from linear

regression analysis in terms of percent inhibition versus

percentage concentration of elutriate. All elutriate

samples were tested in duplicate. Four to eight dilutions

were made to determine the EC50s for ten of the more toxic

elutriate samples.

Reduction in bioluminescence in the Microtox assay was

determined by measuring light output of the bacteria before

and after exposure to the elutriate samples. The degree of

inhibition was determined with respect to the controls. All

elutriate samples were tested in duplicate in the Microtox

assay. EC50s were determined using four to eight dilutions

of the elutriates.

All statistical analyses except calculations of EC50s

utilized the SAS statistical package (SAS 1986) available at

the Northeast Regional Data Center, University of Florida,

Gainesville. The analyses consisted of one-way analysis of

variance (ANOVA), t-test and linear regression analyses.

Results

Toxicity Testing of Sediment Elutriates: Comparison of The
Alpha-Glucosidase Biosynthesis and Microtox Bioassavs

Results of the Alpha-Glucosidase Biosynthesis and

Mocrotox screening assays for sixty-six sediments are

presented in Table 3-2. A statistical comparison of the

percentage inhibitions obtained with the two assays yielded












Table 3-2. Results of toxicity screening of sediment
elutriates using Microtox and the B. licheniformis alpha-
glucosidase biosynthesis assays.


SITE MICROTOX ALPHA-GLU. pH CONDUCTIVITY
%INHIB.a %INHIB.8 AemHOS/cm


ESC-05-01
ESC-05-02
ESC-05-03
ESC-05-04
ESC-05-05
ESC-05-06
ESC-05-07
ESC-05-08
ESC-05-09
ESC-05-10
ESC-06-01
ESC-06-02
ESC-06-03
ESC-06-04
ESC-06-05
ESC-06-06
ESC-06-07
ESC-06-08
ESC-06-09
ESC-06-10
ESC-07-01
ESC-07-02
ESC-07-03
ESC-07-04
ESC-07-05
ESC-07-06
ESC-07-08
ESC-07-09
ESC-07-10
ESC-08-01
ESC-08-02
ESC-08-03
PAL-09-01
PAL-09-02
PAL-09-03
PAL-09-04
PAL-09-05
PAL-09-06
PAL-09-07
PAL-09-08
PAL-09-09
PAL-09-10
MON-12-02


-0.13
74.80
48.04
44.80
46.01
95.62
100.00
100.00
100.00
52.31
92.67
45.55
100.00
65.13
-0.0012
70.67
40.05
85.96
62.65
48.11
99.31
100.00
98.88
44.06
11.70
99.20
64.46
78.97
33.91
100.00
25.69
63.61
22.10
26.94
21.11
24.36
27.46
17.44
23.09
25.23
27.46
25.76
52.60


23.13
100.00
0.88
22.91
41.10
100.00
100.00
100.00
100.00
100.00
49.12
27.53
100.00
100.00
-12.78
57.49
45.59
100.00
55.51
48.68
100.00
100.00
100.00
45.81
53.52
99.34
100.00.
100.00
50.44
100.00
-9.47
-1.10
2.64
-6.94
25.11
-2.10
15.64
13.77
-15.86
8.92
11.01
5.07
36.12


7.60
7.58
7.55
7.55
7.35
7.10
7.10
7.38
6.95
7.05
6.24
5.50
6.88
3.74
7.03
7.35
6.10
7.85
7.01
6.88
7.04
7.05
6.18
6.17
7.35
7.68
7.30
7.09
7.35
6.45
5.30
6.18
7.86
8.07
7.97
7.88
8.06
7.70
7.80
7.60
7.69
7.50
7.25


7650.00
2100.00
4250.00
3110.00
1929.00
1407.00
374.00
510.00
122.00
3700.00
322.00
3770.00
405.00
4560.00
3950.00
4200.00
604.00
2190.00
5040.00
3950.00
60.90
75.70
142.00
1117.00
552.00
183.00
245.00
86.20
528.00
51.20
1830.00
42.80
443.00
348.00
355.00
403.00
476.00
472.00
356.00
389.00
400.00
484.00
7670.00












Table 3-2. continued


SITE MICROTOX ALPHA-GLU. pH CONDUCTIVITY
%INHIB.a %INHIB.a A/mHOS/cm


MON-12-U0
MON-12-04
MON-12-05
MON-12-06
MON-12-07
MON-12-08
MON-12-09
MON-12-10
HIM-02-01
HIL-02-03
HIL-02-06
HIL-02-10
HIL-02-11
HIL-03-01
HIL-03-03
HIL-03-04
HIL-03-06
HIL-04-02
HIL-04-06
HIL-04-07
HIL-04-08
HIL-04-09
HIL-04-11


75.50
30.47
12.27
51.31
-0.05
12.12
49.00
-0.25
29.83
26.96
30.74
34.40
64.68
32.43
38.75
29.98
49.19
20.15
31.79
4.18
46.23
36.21
42.98


80.18
12.78
2.64
90.09
79.96
90.09
68.06
84.80
-23.13
-23.57
19.16
-4.41
16.30
-6.61
-13.44
15.20
-27.53
45.82
4.63
-4.63
-14.98
-33.70
0.44


7.25
7.40
7.45
7.20
7.82
7.61
7.67
7.73
7.50
7.85
7.32
7.45
6.45
7.49
7.60
7.70
7.30
7.85
7.75
7.59
6.92
7.50
7.20


7570.00
7360.00
6320.00
6980.00
8880.00
9290.00
7940.00
9700.00
352.00
945.00
260.00
151.50
111.70
360.00
255.00
240.00
105.20
7470.00
722.00
7690.00
254.00
181.70
347.00


a Percentage inhibition at 45% dilution










53
Table 3-3. Regression analysis of the relationship between
the alpha-glucosidase (x) and Microtox (y) assays"


Model

F-Value


y = 31.31 + .42x

42.83


P-Value


R-Square Value

Pearsons Correlation
Coefficient


.0001


.40


.63


ON = 66 sites (Four Florida


Counties)










a Pearson Correlation Coefficient (r) of 0.63 with an

associated p-value of 0.0001 (Table 3-3). These results

reflect a similar trend in the percentage inhibition

elicited by elutriates in the two assays. The elutriate

samples responsible for a percentage inhibition greater than

90% in both assays were all from sites in Escambia County,

Florida. The number of elutriates found to have percentage

inhibitions greater than 90% were 18 or 27% by the Alpha-

glucosidase biosynthesis assay and 11 or 17% by the Microtox

Assay. All of these elutriates were categorized as very

toxic. A comparison of the number of elutriates causing

inhibitory response of 50% or greater revealed that 39% (26)

of all the elutriates elicited this response in the Alpha-

Glucosidase Biosynthesis assay, while 35% (23) of all the

elutriates elicited this response in the Microtox assay.

EC50 values were determined for ten elutriates samples

that caused very high percentage inhibitions in the two

assays. All of the sediment elutriates were prepared with

sediments from Escambia County, Florida. These results are

presented in Table 3-4. The Alpha-Glucosidase Biosynthesis

assay was more sensitive (i. e. lower EC50s) than the

Microtox assay for every elutriate sample tested (Figure 3-

3). However, there was still a very good correlation, r =

0.88, between the two assays, with an associated p-value of

0.0003.











Table 3-4. EC50a values and equivalent wet and
concentrations for ten sediment elutriates.


55

dry sediment


SITE


MICROTOX
%ELUTRIATE


ESCO5-06

ESCO5-07

ESCO5-08

ESCO5-09

ESCO6-01

ESC06-03

ESC07-01

ESC07-02

ESC07-07

ESC08-01


21.8

2.9

5.7

2.5

29.0

1.6

2.8

2.45

4.7

5.4


EQUIV. DRY AGBb
SED. CONC. %ELUTRIATE
(mg/L)

44690 5.8

5873 1.7

11685 1.0

5063 1.8

55100 25.7

3240 1.2

5740 0.9

5063 1.0

9518 1.1

10935 1.0


EQUIV. DRY
SED. CONC.
(mg/L)

11890

3443

2050

3645

48830

2430

1845

2025

2228

2025


aPecentage concentration of elutriate causing 50%
inhibition of bioluminesence or enzyme biosynthesis
b The B. licheniformis alpha-glucosidase biosynthesis assay













% conc. of elutriate
30-



25



20



15



10-



5-




6-06 6-07 6-08 6-09 6-01 6-03 7-01 7-02 7-07 8-01
Escambla County, FL Sites


M Microtox M Alpha-Glucosidase

Figure 3-3. Comparisons of EC50 values for ten Escambia
County, Florida, sediment elutriates.









57
Classification and ranking of the EC50 values according

to Sanchez et al. (1988) were made using the following

criteria based on the percentage concentrations of

elutriates: at less than 25%, the sample was ranked #1 and

classified as very toxic; at 25%-50%, the sample was ranked

#2 and classified as moderately toxic; at 51%-75%, the

sample was ranked #3 and classified as toxic; at greater

than 75%, the sample was ranked as #4 and classified as

slightly toxic; and elutriates samples causing little or no

effect were ranked #5. By using this method of ranking and

classification, it was found that out of the ten elutriate

samples, nine were very toxic, and one (ESC-06-01) was

moderately toxic based on responses in both bioassays (Table

3-4). These results also showed good agreement between the

two assays.

Graphical comparisons of the two assays for each site

are presented in Figure 3-4 through Figure 3-9. These

figures show a good agreement between the two assays, which

is especially noticeable in the samples from Escambia

County, Florida. Statistically, these samples had the

highest correlation coefficient values among all the

samples. The correlation between the two assays for each

county is presented in Table 3-5. The lowest degree of

correlation between the two assays was obtained with the

Monroe County elutriates. Also, the negative correlations

achieved with the Hillsborough-03 and -04 sites indicated a









% Inhibition


120 -


100-


80-


60-


40 -


20-


0-


-20


-I ~ I I I I I I


01 02 03


04 06 06 07 08 09 10


Esoambla County, FL 05 Sites


M Alpha-Gluoosldase


E Miorotox


Figure 3-4. Comparisons of percent inhibitions in the
Microtox and alpha-glucosidase biosynthesis assays by
sediment elutriates from Escambia County, Florida (sub-sites
05).


S.............................................................. I ... F I- I ........


[


........... ...........................................~....... ............ ........-


...I.













% Inhibition


120 -



100 -



80 -



60-


Figure 3-5. Comparisons of percent inhibitions in
Microtox and alpha-glucosidase biosynthesis assays
sediment elutriates from Escambia County, Florida
sites 06).


the
by
(sub-


r I I I I I I i
02 03 0 0 0 0 0 07 08 09
Escambia County, FL 06 Sites


Alpha-Glucosldase E5 MIcrotox


40-


20 -



0-



-20 -



-40 -


I
01


jf


i


. ................. -.........


I









% Inhibition


120 -


100 -


80 -


60 -


40 -


20 -1


o4


-2 0 1 i 1 1 1 1 1 1 1 1 1 1 1
01 02 03 04 06 07 08 09 10 8-01 8-02 8-03
Escambia County, FL 07 and 08 Sites

SAlpha-Glucoaldase E Microtox

Figure 3-6. Comparisons of percent inhibitions in the
Microtox and alpha-glucosidase biosynthesis assays by
sediment elutriates from Escambia County, Florida (sub-sites
07 and 08).


.................. ......................................


....................................


...............................


.. .. ........


.. .. ..........


. .

I..........













%Inhibition


12-02 12-03 12-04 12-06 12-06 12-07 12-08 12-09 12-10
Monroe County, FL Sites


Alpha-Gluooildase K MIorotox




Figure 3-7. Comparisons of percent inhibitions in the
Microtox and alpha-glucosidase biosynthesis assays by
sediment elutriates from Monroe County, Florida.


100


80





60





40





20





0*





-20 -











30 -





20 -





10


% Inhibition





.... ... ... .... ............... .. ....... .. ..........




j ll \\(\\


-10 -


I I I I I
01 02 03 04 06


I 0I 0I I I
06 07 08 09 10


Palm Beach County, FL 09 Sites

SAlpha-Glucooldase E Miorotox

Negative numbers Indclate stimulation.

Figure 3-8. Comparisons of percent inhibitions in the
Microtox and alpha-glucosidase biosynthesis assays by
sediment elutriates from Palm Beach County, Florida.


-20


v------------------------------


0 1






% Inhibition


s0 -


so -

40 -







- -

-h. -


I


I'


2-1 2-3 2-6 2-10 2-11 3-1 3-3 3-4 3-6 4-2 4-6 4-7 4-8 4-9 4-11
Hillaborough County, FL Sites
-Alpha- Glucosidase E Mlorotox
Negative numbers indicate stimulation
Figure 3-9. Comparisons of percent inhibitions in the
Microtox and alpha-glucosidase biosynthesis assays by
sediment elutriates from Hillsborough County, Florida.


.j
I


.............


. .


...............................................................i












Table 3-5. Relationship between the Microtox and the B.
licheniformis alpha-glucosidase biosynthesis assays based on
percentage inhibition of sediment elutriates.

CORRELATION
COUNTY N COEFFICIENT
(r)

ESCAMBIA-05 10 0.76

ESCAMBIA-06 10 0.88

ESCAMBIA-07 9 0.89

ESCAMBIA-08 3 0.89

PALM BEACH-09 10 0.52

MONROE-12 9 0.04

HILLSBOROUGH-02 5 0.59

HILLSBOROUGH-03 4 -0.91

HILLSBOROUGH-04 6 -0.37

ALL COUNTIES 66 0.63









65
strong inverse relationship between the two assays which is

especially noticeable in Figure 3-8.

In summary, the responses of the alpha-glucosidase

biosynthesis assay to sediment elutriates is very similar to

responses in the Microtox assay. There were good

correlations between the assays for the Escambia County,

Florida sediment elutriates. These elutriates were the most

toxic among all elutriates tested.

Heavy Metal Toxicity of Some Sediment Elutriates As
Determined by MetPAD

MetPAD was used to test the toxicity of sediment

elutriates and the results are presented in Figure 3-10

through Figure 3-12. The purple color intensity on the pad

indicate the degree of toxicity. The control purple spots

were made from samples of the cell suspension exposed only

to distilled water. The white spots, as seen on the pad

labeled Cd or Cu, were made from the cell suspension exposed

to 1 mg/L of cadmium or 25 mg/L of copper. The white spots

indicated strong metal toxicity in the samples.

Samples used on the MetPAD in Figure 3-10 were diluted

to 90% and 45%. All of the elutriate sample dilutions

prepared from Escambia County, Florida, sediments were

toxic, giving white to very light purple spots (spots la and

5a ). The two elutriate samples prepared from Palm Beach

County, Florida sediments showed no toxicity on the MetPAD

as seen by the purple spots numbered lla, llb, 12a, and 12b.

These results indicate the presence of metals in the











Escambia County elutriates and led to additional MetPAD

tests to confirm this assumption.

The spots in Figure 3-11 show results obtained with 90%

dilutions of elutriates prepared from Escambia County,

Florida sediments before and after filtration through metal

chelating columns. Toxicity was noticeable for most of the

elutriate samples labeled 'b' in Figure 3-11 before

filtration through the chelating resin columns. Spots

resulting from elutriates that had passed through the

chelating resin appear purple for all of the samples except

9a (ESC-07-07). A replicate of this sample showed the same

results, no reduction in toxicity after filtration through

the chelating resin column. However, a second test of this

sample showed a reduction in toxicity and can be seen in

Figure 3-12.

To compare MetPAD results with the alpha-glucosidase

biosynthesis assay, 45% dilutions (labeled with "a" suffix)

of the resin-filtered elutriate samples were compared with a

90% dilution (labeled with "b" suffix) of the samples as

seen in Figure 3-12. No toxicity was indicated in either of

the dilutions. The elutrate sample from the ESC 07-07 site

that showed toxicity after filtration in Figure 3-10 was

non-toxic at both a 90% and 45% dilution. This observation

indicated perhaps a mix up in the samples during the test.

These results confirmed the presence of metals in the

Escambia County, Florida, elutriates. Any metals present in















1Cd

1S 2a 3a 4a
Ib 2b 3b 4b


C,

5a 6a 7a 8a

5b 6b 7b 8b


0 eCd

9a 1Oa
9b 10b









Figure 3-10. MetPAD results using ten toxic and two non-
toxic sediment elutriates. Key: 1, ESC-05-02; 2, ESC-05-07;
3, ESC-05-08; 4, ESC-05--09; 5, ESC-06-01; 6, ESC-06-03;
7, ESC-07-01; 8, ESC-07-02; 9, ESC-07-07; 10, ESC-08-01; 11,
PAL-09-02; and 12, PAL-09-04. The "a" suffix denotes cell
exposure to a 45% dilution of the elutriate. The "b" suffix
denotes cell exposure to a 90% dilution of the elutriate.





















CM CO


20. .34


I b 2b 3b


S. C.





, 6b 7b Ob


Cu

100


Figure 3-11. MetPAD results obtained with toxic sediment
elutriates before and after filtration through metal
chelating resin columns (90% dilution of elutriates). Key:
1, ESC-05-02; 2, ESC-05-07; 3, ESC-05-08; 4, ESC-05-09; 5,
ESC-06-01; 6, ESC-06-03; 7,ESC-07-01; 8, ESC-07-02; 9, ESC-
07-07; and 10, ESC-08-01. The "a" suffix denotes cell
exposure to elutriates that were filtered through chelating
resin columns. The "b" suffix denotes cell exposure to
elutriates before filtration through metal chelating
columns.


















i*jtge
^0 0S^


, g i

14Ql s)


9b lOb












Figure 3-12. MetPAD results obtained with sediment
elutriates filtered through metal chelating resin columns.
Key: 1, ESC-05-02; 2, ESC-05-02; 3, ESC-05-08; 4, ESC-05-09;
5, ESC-06-01; 6, ESC-06-03; 7, ESC-07-01; 8, ESC-07-02;
9, ESC-07-07 and 10, ESC-08-01. The "a" suffix denotes cell
exposure to a 45% dilution of the elutriate. The b" suffix
denotes cell exposure to a 90% dilution of elutriate.











the samples would be chelated by the resin resulting in a

considerable decrease in the amount of metals in the

elutriates and thus a change in toxic response of MetPAD.

These results led to the determinations of metal

concentrations in all of the elutriate samples, which are

discussed in the next section.

Metal Concentrations In Elutriate Samples

The concentrations of cadmium, copper, lead and zinc in

the elutriate samples are recorded in Table 3-7. Detection

of high concentrations of metals in many of the elutriate

samples confirmed results of the MetPAD test. The highest

concentrations of metals were found in elutriates prepared

from sediments collected from sites in Escambia County,

Florida. The elutriate prepared from the ESC 07-02 site had

the highest combined concentration of the four metals (10,

648.3 ug/L). The ESC-06-03 elutriate followed with a

combined concentration of 10,357.3 ug/L. Seven elutriate

samples having the lowest concentrations of metals were from

Palm Beach (4 samples) Hillsborough (2 samples) and Monroe

(one sample) Counties, Florida. Each elutriate contained

2.7 ug/L of one of the four metals.

Metals in the highest concentrations in the elutriates

were copper and zinc. The concentrations of zinc and copper

(these metals have synergistic toxicity ) showed a strong

correlation (r=0.79) in the elutriate samples. Cadmium and

lead were detected at low concentrations in the elutriate












Table 3-7. Concentrations of four metals in sediment
elutriates.

Metal Concentrations in ug/L

Site Cd Cu Pb Zn Total


ESC-05-01 5 3 8
ESC-05-02 5 663 708 1376
ESC-05-03 10 179 171 360
ESC-05-04 10 201 296 507
ESC-05-05 5 3 83 91
ESC-05-06 10 993 3976 4979
ESC-05-07 10 5128 100 4183 9421
ESC-05-08 5 3259 100 3487 6850
ESC-05-09 5 2247 10 3913 6174
ESC-05-10 10 201 484 694
ESC-06-01 5 179 10 2961 3155
ESC-06-02 10 3 100 2628 2741
ESC-06-03 5 1917 2920 5516 10,357
ESC-06-04 5 47 241 1353 1646
ESC-06-05 5 3 100 107
ESC-06-06 5 245 241 203 693
ESC-06-07 5 69 241 348 663
ESC-06-08 5 1477 241 1951 3674
ESC-06-09 5 267 241 41 554
ESC-06-10 5 157 100 10 271
ESC-07-01 3545 665 3267 7476
ESC-07-02 3545 1089 6015 10,648
ESC-07-03 773 241 1296 2309
ESC-07-04 157 100 671 928
ESC-07-05 47 100 171 318
ESC-07-07 1301 241 2436 3977
ESC-07-08 355 100 390 844
ESC-07-09 619 241 770 1630
ESC-07-10 157 100 317 574
ESC-08-01 2885 10 1124 4019
ESC-08-02 135 879 1014
ESC-08-03 465 328 792
PAL-09-01 113 113
PAL-09-02 3 3
PAL-09-03 *
PAL-09-04 3 3
PAL-09-05 91 91
PAL-09-06 *
PAL-09-07 *
PAL-09-08 3 3












Table 3-7. Continued.


Metal Concentrations in uQ/L
SITE Cd Cu Pb Zn Total


PAL-09-09 3 3
PAL-09-10 47 47
HIL-02-01 47 15 62
HIL-02-03 135 83 218
HIL-02-06 91 91
HIL-02-10 91 140 231
HIL-02-11 333 10 260 603
HIL-03-01 3 135 138
HIL-03-03 3 317 320
HIL-03-04 3 640 643
HIL-03-06 201 78 278
HIL-04-02 3 3
HIL-04-06 3 3
HIL-04-07 3 31 34
HIL-04-08 91 109 200
HIL-04-09 113 255 367
HIL-04-11 157 400 557
MON-12-02 25 73 97
MON-12-03 25 15 40
MON-12-04 3 15 18
MON-12-05 69 36 105
MON-12-06 3 21 23
MON-12-07 3 3
MON-12-08 3 109 112
MON-12-09 25 41 66
MON-12-10 3 88 91

* Denotes concentrations below instrument detection level









73

samples. Cadmium was absent in most of the elutriates, but

concentrations ranging from 5-10 ug/L were detected in the

Escambia 05 and 06 elutriate samples. Lead occurred more

frequently than cadmium and at higher concentrations. The

highest concentration of lead occurred in the ESC-06-03

elutriate sample.

Seventeen elutriate samples had combined concentrations

of the four metals greater than 1000 ug/L. Thirteen

elutriate samples had concentrations between 500 and 1000

ug/L; 14 had concentrations below 100 and 500 ug/L and 22

samples had metal concentration ranging from 0-100 ug/L.

All of the Escambia County elutriates that had very

high combined metal concentrations displayed 100% inhibition

in the alpha-glucosidase biosynthesis assay. Elutriate

samples that showed a toxic response in the MetPAD tests

also had very high metal concentrations. Comparisons of

metal concentrations and responses with the alpha-

glucosidase, Microtox and MetPAD assays are presented in

Table 3-8.

Statistical correlations between the metal

concentrations and the responses of the alpha-glucosidase

biosynthesis assay were made. There was good correlation

between the combined concentrations of the four metals and

responses in the alpha-glucosidase biosynthesis assay (r =

0.57). There was also good correlation between the

concentrations of copper (r = 0.55) and zinc (r = 0.53) and











responses in the assay. A summary of the comparisons are

presented in Table 3-9. These results indicated that the

alpha-glucosidase biosynthesis assay is sensitive to metal

toxicity in aqueous samples.

Discussion

The primary objective of this study was to evaluate the

B. licheniformis alpha-glucosidase biosynthesis (AGB) assay

for use in toxicity screening of sediment elutriates. The

evaluation was based on comparisons of inhibitory responses

in the B. licheniformis AGB assay and the Microtox assay

which is a standard assay used to evaluate toxicity of

environmental samples.

In the first part of the study, elutriates of sediment

samples were prepared and tested by both assays.

Elutriates were diluted to 45% and used at this level in

both assays where percentage inhibitions of alpha-

glucosidase biosynthesis in B. licheniformis and

bioluminescence in P. phosphoreum were indicators of acute

toxicity. Agreement between the two bioassays was greatest

for elutriate samples that were toxic. The most striking

divergence in the responses of the assays were obtained with

elutriate samples with little or no toxicity.

Very little correlation was obtained from a comparison

of responses elicited by elutriates from Monroe County,

Florida, in the assays. The alpha-glucosidase biosynthesis

assay gave percentage inhibitions greater than 50% for six











Table 3-8. EC50 values, metal concentration, and MetPad
results for ten sediment elutriates.

SITE METAL CONC.a MICROTOX AGB0 METPAfD
(ug/L) EC50d EC50

ESC-05-06 4979 21.8 5.82 ++++



ESC-05-07 9421 2.94 1.70 +++++


ESC-05-08 6850 5.70 1.00 +++++


ESC-05-09 6174 2.54 1.80 +++++


ESC-06-01 3155 29.00 25.7 +++


ESC-06-03 10,357 1.60 1.15 +++++


ESC-07-01 7476 2.82 0.93 +++++


ESC-07-02 10,648 2.45 0.98 +++++


ESC-07-07 3977 4.66 1.13 +++++


ESC-08-01 4019 5.42 1.00 +++++


a Total metal concentration of four metals: Cd, Cu, Pb, and
Zn
b The B. licheniformis Alpha-Glucosidase Biosynthesis Assay
cMetPAD results based on color change on pads
dPercentage concentration of elutriate
+ purple (no toxicity)
++ medium purple
+++ light purple
++++ pale purple
+++++ white (very toxic)











Table 3-9. Correlation between percentage inhibitions of
the B. licheniformis alpha-glucosidase biosynthesis assay
and metal concentrations in sixty-six sediment elutriate
samples.


CORRELATION
(r)


Cadmium

Copper

Lead

Zinc

Four Metals Combined


SThe total number of samples tested was 66.
b Significant at p<0.05


METALa











of the nine sites, while Microtox showed similar responses

for three of the sites. Differences in responses were first

thought to be related to the high conductivity of the

samples. Sediment samples from the Monroe county sites were

all marine and had high conductivity. However, the

relationship between responses elicited in the B.

licheniformis AGB assay, and conductivity was negligible.

This observation indicated some other factor, such as the

metal concentrations, contributing to the differential

responses of the assays.

Differential responses between Microtox and other

bioassay test organisms have been observed in other studies.

Dutka et al. (1988) tested water extracts of sediments using

the Microtox test and several other bioassays. The samples

were found to be negative for toxicant activity with the

Microtox test. However, in another toxicant screening test

using an invertebrate species, the same samples showed

toxicant activity. These results support the use of a

battery of tests to evaluate samples for the presence of

toxicants.

Negative correlation coefficients were obtained for

Hillsborough-03 and -04 sites (Table 3-6 ). Enzyme

stimulation was observed in the AGB assay, but none in the

Microtox assay. Again, the two test systems were responding

differently to the water soluble constituents in the

elutriates. Similar observations of bacterial stimulation by











organic constituents have been observed in other studies.

(Giesy and Hoke 1989).

Such divergence in results of two test systems is not

unusual. Upon comparing two different test species, Sanchez

et al. (1988) concluded that a direct comparison of the

results is difficult and complicated by the inherent

differences between the two systems. In spite of the

differences, an overall comparison of the results of the two

assays showed good agreement, giving a correlation

coefficient (r) of 0.63 with an associated p-value of .0001.

Comparison of EC50s for Escambia County elutriates

indicated a higher degree of sensitivity in the alpha-

glucosidase biosynthesis assay when compared with the

Microtox assay. However, there was good correlation between

the EC50s for the two assays, r = 0.88 with an associated p-

value of 0.0003.

Results obtained with MetPAD indicated that this assay

can be useful as a rapid tool to detect metal toxicity. The

MetPAD results were confirmed by analysis of the elutriate

samples for metals. There was also a good correlation

between the total (four) metals in all 66 elutriate samples

and responses of the alpha-glucosidase biosynthesis assay

(r = 0.76).

The sensitivity of the AGB assay to metals can be

associated with the structure of the gram positive cell

wall. The interaction between metals and a gram-positive











bacterium involves the positive and negative charged sites

in the cell wall (Collins and Stotzky 1989). The penetration

of the cell wall by metal ions and their ultimate

incorporation into membrane or cytoplasmic proteins can

involve several distinct kinetic steps and can be a factor

in the response of the bacterium to specific metals (Collins

and Stotzky, 1989). The concentration of metals in the cells

environment is also a factor in toxic effects exerted on the

bacterium.

The concentrations of copper and zinc in several of the

sediment eultriates were relatively high and occurred at

levels that contributed to a toxic effect on B.

licheniformis. For example, the zinc concentrations in

elutriates from several of the Escambia county sediments

ranged from 2 to 6 times the maximum concentration of zinc

found in surface waters, which is 1183 ug/liter (Lamb 1985).

Copper concentrations were even higher, ranging from 1.6-18

times the concentrations found in surface waters (280

ug/liter). Such high concentrations of these metals is not

unusual for areas like Escambia County which receive waste

waters from chemical and pulp and paper industries (Table 3-

1).

There was a good correlation, r = 0.79 between the

concentrations of zinc and copper in the Escambia County

elutriates. These results indicate a potential threat to

the aquatic environment in this area which may especially











become apparent if sediments are disturbed. It often has

been noted that the simultaneous presence of copper and zinc

in water may be synergistic and may cause toxicity to fish

that is more serious than would be predicted by considering

the effects of these chemicals individually (Lamb 1988).

In summary, the data presented in this chapter show

that the B. licheniformis alpha-glucosidase biosynthesis

assay can be useful in the toxicant screening of sediment

elutriates. The assay is comparable with the Microtox assay

and appears to be sensitive for samples containing toxic

metals. The large number of samples tested and the variable

nature of the sediments used provided statistical validity

to the study. The assay is simple and very economical when

compared with the Microtox test system. These

characteristics makes this assay suitable for consideration

for inclusion in a battery of screening tests for toxicity

in aqueous samples.















CHAPTER 4
COMPARISON OF SEDIMENT EXTRACTION PROCEDURES AND EXCHANGE
SOLVENTS FOR HYDROPHOBIC COMPOUNDS BASED ON INHIBITION
OF BIOLUMINESCENCE



Introduction

The use of sediment elutriates in bioassays as

demonstrated in Chapter 3 gives results that indicate the

effects of bioavailable or water-soluble compounds to the

test organisms. However, it has been shown that many

hydrophobic compounds that are tightly sorbed to sediment

are not completely unavailable to aquatic organisms and can

have an impact on them (Reynoldson 1987; Nielson et al.

1986; Delfino 1979). The recognition of toxic effects

associated with hydrophobic residues in sediments has led to

an increase in the use of sediment solvent extracts in

conjunction with bioassays. The extraction efficiency for

toxicants plays an important role in obtaining reproducible

results in the toxicity assessment of contaminated sediment

samples.

There is a broad base of well developed procedures used

to extract chemicals from sediments for analytical purposes.

However, many of these procedures can not be used with

bioassays, since some of the solvents are toxic to the test

organisms. Therefore, researchers that use organic solvent

81











extraction procedures in conjunction with bacterial

bioassays are limited to the use of solvents that exert the

least toxic effect on the test organisms.

Some solvents that have been used in conjunction with

bacterial bioasssays include: dimethyl sulfoxide, methanol,

acetone and dichloromethane (methylene chloride) (Dutka and

Kwan 1988; Schiewe et al. 1985; and Donnelly et al. 1987).

The extraction methods involve shaking, tumbling, sonifying

or blending sediment or soil samples in the presence of a

specific solvent for a specified period of time.

Investigators considered shaking periods ranging from 15

minutes (Larson 1989) to 18-24 hours (Dutka and Kwan 1988).

Schiewe et al. (1985) extracted various marine sediments

by tumbling and compared the effect of different solvents on

Photobacterium phosphoreum in the Microtox assay (Bulich et

al 1984). True and Heyward (1990) extracted marine sediments

using sonication. The compounds extracted were analyzed

along with interstitial water (pore water) samples using the

Microtox assay. They concluded that the toxicity of solvent

extracts was more indicative of potential toxicity to

organisms affected by hydrophobic toxins than results

obtained using only interstitial water.

Thus, it is obvious that the extraction of hydrophobic

residues from sediment samples is very important for

toxicity assessment of sediment samples. However, most

methods for preparing solvent extracts of sediment samples











for bioassays have not been standardized. Subsequently,

there is need for more research associated with extraction

of sediment samples in conjunction with bioassays.

The purpose of this study was to compare shaking and

sonication procedures utilizing methylene chloride as the

extracting solvent for sediment samples. A comparison of

methanol and dimethyl sulfoxide (DMSO) as exchange solvents

was also made. The efficiencies of the extraction

procedures were based on the inhibition of bioluminescence

in the Microtox assay. Extracts obtained by the most

efficient procedure were used in the B. licheniformis alpha-

glucosidase biosynthesis (AGB) assay, and responses of the

two assays were compared for possible correlations.

Materials and Methods

Chemicals and Reagents

All solvents used were HPLC grade and included:

methylene chloride, dimethyl sulfoxide and methanol.

Reagent grade chemicals were used for determinations of

organic carbon and included the following reagents:

standard IN K2Cr207 prepared by dissolving exactly 49.04g in

distilled water diluted to 1 liter; Diphenylamine indicator;

0.5g dissolved in 20 mL distilled water and 100 mL of

concentrated H2S04; and in ferrous ion solution prepared by

dissolving 278.Og of FeSO4-7HO in 1 liter distilled water

containing 15 mL of concentrated H2SO4. An 85% solution of

phosphoric acid (H3PO4) was prepared also.











Sediment Samples

Sediment samples used in this study were selected based

on results obtained from the initial toxicity screening of

sixty-six sediment elutriates deionizedd water extracts) of

sediment samples collected from various sites in the state

of Florida. A description of the sites and their location

by latitude and longitude are presented in Table 3-1 in

Chapter 3. Five toxic (percentage inhibition of

bioluminescence greater than 50%) and five non-toxic

(percentage inhibition of bioluminescence less than 50%)

samples as determined by the elutriate screening of the

samples by the Microtox assay were used.

The collection and storage of sediment samples are

described in Chapter 3. The sediments extracted in this

study were collected from Escambia (ESC) and Hillsborough

(HIL) counties of Florida. These sediment samples were

frozen for six months before being extracted. The frozen

samples were defrosted at 40C and room temperature for 1-2

hours prior to weighing. The sediment samples extracted in

this study were not dried so that some volatile as well as

non-volatile organic could be extracted. Extraction of wet

sediment samples also allowed the determination of the toxic

effects of both sediment particulates and their associated

interstitial water. Percentage water concentration and dry

weight for each sediment sample are presented in Table 4-1.











Percentage Organic Carbon

The percentage organic carbon content of each sediment

sample was determined by the Walkey-Black method (1934).

The steps of the procedure were as follows: 1) 5g of dry

sediment was placed in a 500ml conical flask; 2) exactly

10ml of IN K2Cr20O was pipetted into the flask and gently

mixed with the sediment; 3) 20mL of concentrated H2SO4 was

added and gently mixed for 1 minute; 4) the mixture was

allowed to stand 20-30 minutes; 5) after standing the

solution was diluted to 200ml with distilled water; 6) 10ml

of 85% phosphoric acid was added to the solution (in a

hood); 7) 30 drops of diphenylamine was added to the

solution; 8) the solution was back titrated with IN ferrous

sulfate; and 9) the solution was back titrated until its

initial color of dull green shifted to a turbid blue color,

and finally at the end point to a brilliant green. A

standardization blank was prepared without sediment and

tested in the same manner. Results were calculated using

the following equation:

% OM = 10(1-T/S) x 1.34 where

S = standarization blank titration, mL
ferrous sulfate solution
T = sample titration, mL ferrous sulfate solution

Extraction Procedure

Solvent Extraction by Sonication. Sediment extractions with

dichloromethane followed the general procedure of True and

Heyward (1990) with some modifications. Triplicate 10 gram










86
Table 4-1. Water content, dry weight, and percentage organic
matter of sediment samples.


Site % Waters Dry weight % Organic
(in grams)" Matter

ESC-05-07 18.56 8.14 0.43


ESC-05-09 19.29 8.07 0.61


ESC-07-01 18.18 8.18 0.40


ESC-07-02 18.93 8.10 0.18


ESC-07-03 25.96 7.40 0.98


ESC-05-01 67.49 3.25 5.44


HIL-02-10 20.53 7.95 4.92


HIL-03-04 65.24 3.50 6.45


HIL-04-06 18.63 8.14 0.92


HIL-04-09 25.58 7.44 4.61


abased on 10 gram wet weight sediment sample









87
(wet weight) samples from each site were placed in solvent-

rinsed 250 mL stainless steel or glass centrifuge tubes and

mixed with 20 grams of sodium sulfate to remove water.

Seventy-five milliliters of methylene chloride were added

and mixed with the sediment sample. The sediment-solvent

slurry was stirred with a stainless steel spatula and

sonicated (50 duty cycle, pulse rate; W-375 Heat Systems-

Ultrasonics Inc.) for five minutes. The solvent was

decanted carefully, and the procedure was repeated twice. A

total solvent extract volume of 225 mL was collected and

poured through sodium sulfate. This extract was concentrated

in a Rotary Evaporator (Brinkman R110) to a volume of 6 mL.

This extract was further concentrated by evaporation under a

gentle stream of nitrogen gas in a water bath (30 40C) to

final volume of 1 mL. One hundred microliters of the final

concentrated extract was added to 2.25 mL of an exchange

solvent (dimethyl sulfoxide or methanol) and placed under

nitrogen gas in a water bath (42C) for 30 minutes to

facilitate solvent exchange. The 2.25 mL volume extract

represented 10% (or 1 gram) of the original 10 gram wet

sediment sample. A 1% dilution (representing .01 gram of

the original wet sediment sample) was made with distilled

deionized water for use with the Microtox bioassays.

Solvent Extraction by Shaking. Three separate 10 gram (wet

weight) samples from each site were prepared in the same

manner described for sonication. However, the sediment









88
samples were placed in methylene chloride-rinsed flasks (500

ml volume) and mixed with 20 grams of sodium sulfate. A

total volume of 225 ml of methylene chloride was added to

the flask and mixed with the sediment sample. The samples

were allowed to shake for 24 hours at low speed. The

extracts were collected and concentrated in the same manner

described previously for the sonicated samples.

A method blank (no sediment) was prepared in the same

manner described for each procedure to determine background

toxicity for the solvents.

The Microtox Assay

Lyophilized samples of the gram negative marine bacterium

Photobacterium phosphoreum were reconstituted according to

procedures outlined in the Microtox Operating Manual

(Microbics 1982) for use in the tests. Two replicates of a

45% dilution of the 1% solvent extracts of each sample were

tested (this dilution represented 0.0045 g {wet weight} of

the original sediment sample). All assays were carried out

at 150C with a 15 minute contact period. Samples that gave

percentage inhibitions of bioluminescense greater than 50%

were further tested in definitive assays to determine their

EC50s (the concentration of extract causing a 50% reduction

of bioluminescence). Four to eight dilutions were done for

each sample to make this determination. Data were tabulated

and reduced according to the Microtox Operating Manual

(Microbics 1982). Background toxicity determined with the











solvent blanks was subtracted from the initial percentage

inhibitions for each sample, thus, the percentage

inhibitions reported represent the toxic effects of only the

extracted toxicants.

The Bacillus licheniformis Alpha-Glucosidase Assay
(AGB Assay)

The AGB assay was performed as described in Chapter 3

using the 1% Methanol extracts. A 45% dilution was used in

the assay. Duplicates of each sample and blanks were

prepared for each sample.

Data Analysis

EC50s for the Microtox assay were derived using linear

regression analysis. Statistical analysis, consisting of

Analysis of variance (ANOVA) and Student's t-test, of the

percentage inhibitions obtained from extracts by both

extraction procedures with DMSO as the exchange solvent were

made. The best procedure determined by the statistical

analyses was used to compare DMSO and methanol as exchange

solvents.

In summary, comparisons were made between shaking and

sonication for extracting sediment samples, between methanol

and DMSO as exchange solvents and between responses of the

two assays.

Results
Shaking vs Sonication for Extracting Sediment Samples

Results of the Microtox assay evaluating the toxicity of

the solvent extracts prepared by shaking and sonication with