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THESIS

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ANAEROBIC TRANSFORMATION OF DDT IN PINE RIVER
SEDIMENTS

presented by

TOMEKA KENYATTA PRIOLEAU

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ANAEROBIC TRANSFORMATIONS OF DDT IN PINE RIVER SEDIMENTS

By

Tomeka Kenyatta Prioleau

A THESIS

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

MASTER OF SCIENCE
Department of Crop & Soil Sciences

2003

ABSTRACT
ANEROBIC TRANSFORMATIONS OF DDT IN PINE RIVER SEDIMENTS
By

Tomeka Kenyatta Prioleau

This study examined the fate of DDT in anaerobic sediments taken from two
locations within the Pine River Superfund site in St. Louis, Michigan. Pine River
sediment microcosms were analyzed for their ability to dechlorinate DDT and its
metabolites DDD, and DDE under anaerobic conditions. Dechlorination of MC-DDT to
14C -DDD occurred in both abiotic (autoclaved) and biologically active sediment slurries.
No transformations of DDD or DDE were observed. Based on recovery of added DDT as
DDD there was a greater amount of dechlorination occurring in bottom (10-20cm) layer
sediments than in the top (0-10 cm) layer. Similar amounts of DDT were recovered as
DDD in sediments from the two locations with different levels of historical
contamination by DDT and co—contaminants. Anaerobic sediment slurry systems known
to dechlorinate PCBs and/or PBBs were tested for their ability to transform DDT. These
consisted of Red Cedar River sediments inoculated with microorganisms eluted from
Pine River, Hudson River, and Silver Lake sediments. All systems supported the
dechlorination of DDT to DDD by both abiotic and biotic processes, with abiotic
transformations accounting for the majority (63-95%) DDT transformation. FeSO4
amendments were also utilized in an attempt to enhance reductive dechlorination of
DDT. The FeSO4 amendment enhanced DDT dechlorination in Hudson River

microcosms, however no further transformations beyond DDD were observed.

Copyright by
TOMEKA KENYATTA PRIOLEAU
2003

DEDICATION
This work is dedicated to my family who has always encouraged and supported me, and

especially to my Mother -thank you for having faith in me.

ACKNOWLEDGEMENTS

First, I would like to thank a few faculty members who were very important to me
here at Michigan State University. I would like to thank Dr. Stephen Boyd for his
patience and guidance and for allowing me the opportunity to work with him. Dr. John
Quensen for his guidance in many aspects of my research but especially the laboratory
portion. I would like to thank Dr. James Tiedje and Dr. Karen Chou for serving on my
guidance committee; your insight is greatly appreciated. I would like to thank Dr. James
Jay for his unending guidance and support in my educational endeavors.

Secondly, I would like to thank all of my classmates and labmates who helped
and encouraged me through the years. Many lifelong friendships were made here and
these friendships I will always hold dear.

Lastly, I would like to thank my family for always encouraging me to go on and

be successful at whatever I seek to accomplish. Thank you all.

TABLE OF CONTENTS

LIST OF TABLES ................................................................................. viii
LIST OF FIGURES ................................................................................... x
CHAPTER 1
INTRODUCTION .................................................................................... 1
History of DDT .............................................................................. 1
Environmental Effects ....................................................................... 3
Degradation Mechanisms of DDT- Dechlorination and Dehydrochlorination .....6
DDT to DDD ................................................................................ 10
DDT to DDE ................................................................................ 13
DDD TO DDMU ........................................................................... 15
DDE to DDMU ............................................................................. 15
Further Transformations ................................................................... 16
REFERENCES ....................................................................................... 17
CHAPTER 2
SITE HISTORY AND METHODOLOGY ...................................................... 21
Site History ................................................................................... 21
Introduction .................................................................................. 26
Objectives ................................................................................... 26
Materials and Methods .................................................................... 27
Materials ........................................................... ‘ ......................... 27
Sediment Sampling and Analyses ....................................................... 27
Methane Analysis .......................................................................... 33
Extraction and TLC Analysis ............................................................ 33
REFERENCES ..................................................................................... 35
CHAPTER 3
RESULTS AND DISCUSSION .................................................................. 36
DDT Transformations ..................................................................... 36
Transformations in biologically active microcosms .......................... 37
Transformations in autoclaved microcosms ................................... 44
Abiotic and Biotic Transformations ..................................................... 49
14C Recovery of DDT ...................................................................... 52
Transformations of DDD and DDE in sediment microcosms ........................ 53
Red Cedar Sediment Microcosms ....................................................... 55
Biologically active microcosms ................................................. 55
Abiotic microcosms ............................................................... 58

vi

Abiotic and Biotic transformations ....................................................... 58

Transformations of DDD and DDE in sediment microcosms ........................ 62
SUMMARY ......................................................................................... 63
REFERENCES ...................................................................................... 65

vii

LIST OF TABLES

Table 1.1. Chemical names and acronyms of DDT and its derivatives used in the text. ...4

Table 2.1. Concentrations of total DDT in skin-off carp ﬁllets from below the St. Louis

Dam and in the St. Louis Impoundment. *Fish collected in 1983 were not analyzed for
DDE and DDD, only DDT (EPA, 1998) ......................................................... 23

Table 3.1. Recovery of 14C DDT and transformation products in Pine River sediments
based on TLC Analysis. Sample microcosms were established using the top (T) and
bottom (B) halves of sediment cores from two different locations (cores 4 and 11). The
standard deviation is reported in parenthesis. *24 week Autoclaved samples were not
setup ................................................................................................... 43

Table 3.2. DDD formation due to biotic and abiotic transformations of DDT in Pine
River Sediment microcosms constructed using top (T) and bottom (B) sections of
sediment cores 4 and 11. The biotic (BI) fraction was determined by dividing the
difference in DDD recovered in live treatments by the DDD recovered in autoclaved
treatments and dividing by the DDD recovered in the live treatment (X 100). The abiotic
(AB1) fraction was determined by dividing the DDD recovered in the autoclaved
sediments by the DDD recovered in the live sediment (X

100) .................................................................................................. 50

Table 3.3. 14C Recovery in Pine River sediment microcosms over 32 weeks. Recoveries
represent the amount of DDX (DDT+DDD+Origin) recovered from aqueous and solvent
extractions of sediment microcosms for top (T) and bottom (B) sections of cores 4 and
11. The standard deviation is reported in parenthesis. *24 week autoclaved samples
were not setup ....................................................................................... 52

Table 3.4. Recovery of added l4C-DDD and -DDE in Pine River sediment microcosms.
Sample microcosms were established using the top (T) and bottom (B) halves of
sediment cores from two different locations (cores 4 and 11). The standard deviation is
reported in parenthesis. *24 week autoclaved samples were not analyzed ................. 54

Table 3.5. Anaerobic transformation of DDT in sterilized Red Cedar (RC) River
sediment microcosms inoculated with microorganisms eluted from Hudson River (HR),
Pine River (PR) or Silver Lake (SL). Some microcosms were also amended with FeSO4
(F). Uninoculated (U) Red Cedar River sediment was also incubated before (live-L) and

viii

after (auto-A) autoclaving. Standard deviations of the means are given in parenthesis.
.......................................................................................................... 59

Table 3.6. DDD formation due to biotic and abiotic transformations of DDT at 32 weeks.
Microcosms were set up using uninoculated Red Cedar River (RCU) sediment, as well as
autoclaved RC sediment slurries which were inoculated with microorganisms eluted from
RC, Hudson River (HR), Pine River (PR) or Silver Lake (SL), and in some instances
amended with FeSO4 (F). Both autoclaved (abiotic) and non-autoclaved (biotic)
microcosms were then incubated under anaerobic conditions for 32 weeks. The biotic
fraction was detemiined by dividing the difference between the DDD recovered in live
and DDD recovered in autoclaved treatments by the DDD recovered in the live treatment
(X100). The abiotic fraction was determined by dividing the DDD recovered in the
autoclaved microcosms by the DDD recovered in the live microcosms (X100) ............ 60

LIST OF FIGURES

Figure 1.1. Transformation pathways of DDT by Aerobacter aerogenes (Wedemeyer,
1967) ................................................................................................... 9

Figure 1.2. Reductive dechlorination of DDT to DDD under anaerobic conditions
(Rockhind and Blackburn, 1986) .................................................................. 10

Figure 1.3. Aerobic dehydroclorination of DDT to DDE (Rockhind and Blackburn,
1986) .................................................................................................. 14

Figure 2.1. The Velsicol Chemical Corporation plant site located on the Pine River
in St. Louis, Michigan. Adapted from Morris et. al., (1993) ................................. 22

Figure 2.2 The Velsicol Chemical Corporation plant site located on the Pine River in St.
Louis, Michigan and sampling points for core 4 and core 11. Adapted from Morris et. al.,
(1993) ................................................................................................. 28

Figure 3.1. Transformation of DDT to DDD in Core 4 top and bottom layer anaerobic
sediment microcosms during a 32 week incubation period .................................... 38

Figure 3.2. Anaerobic dechlorination of DDT to DDD in Core 11 top and bottom layer
anaerobic sediment microcosms over a 32 week incubation period .......................... 40

Figure 3.3. Comparison of anaerobic dechlorination of DDT in Core 4 and 11 top (T)
and bottom (B) sections over a 32 week incubation period .................................... 41

Figure 3.4. Autoradiograms of parent compounds and metabolite zones on TLC plates
for transformations of DDT ........................................................................ 42

Figure 3.5. Transformation of DDT to DDD in Core 4 (top and bottom sections) abiotic
sediment microcosms during a 32 week incubation period .................................... 46

Figure 3.6. Transformation of DDT to DDD in Core 11 (top and bottom sections) abiotic
sediment microcosms during a 32 week incubation period .................................... 47

Figure 3.7. Comparison of anaerobic dechlorination of DDT in abiotic Core 4 and 11 top
(T) and bottom (B) sections during a 32 week incubation period ............................ 48

Figure 3.8. DDD recovered from the abiotic and biotic transformations of DDT in Pine
River sediment microcosms constructed using the top (T) and bottom (B) sections of
sediment cores 4 and 11. The biotic fraction was determined by dividing the difference
of the live and autoclaved treatments by the live treatment. The abiotic fraction was
determined by dividing the DDD recovered in the autoclaved treatments by the DDD
recovered in live treatments. ...................................................................... 51

Figure 3.9. Autoradiograms of parent compounds on TLC plates for transformations of
DDT ................................................................................................... 54

Figure 3.10. Recovery of DDT and DDD under strict anaerobic conditions from Red
Cedar Unamended (RCU), Hudson River (HR), Hudson River with FeSO4 (HRF), Pine
River (PR), Silver Lake (SL), and Silver Lake with F eSO4 (SLF) inocula after 32

weeks ................................................................................................. 57

Figure 3.11. The fraction of DDD recovered resulting from biotic and abiotic
transformations of DDT at 32 weeks. Microcosms were set up using uninoculated Red
Cedar River (RCU) sediment, and autoclaved RC sediment slurries which were
inoculated with microorganisms eluted from RC, Hudson River (HR), Pine River (PR) or
Silver Lake (SL) and in some instances amended with FeSO4 (F). Both autoclaved
(abiotic) and non-autoclaved (biotic) microcosms were then incubated under anaerobic
conditions for 32 weeks. The abiotic fraction was determined by dividing the DDD
recovered in the autoclaved microcosms by the DDD recovered in the live treatments (X
100%). The biotic fraction was determined by dividing the difference of the DDD
recovered in the live microcosms and the DDD recovered in the autoclaved microcosms
by the DDD recovered in the live treatment microcosms (X 100%) ......................... 61

xi

CHAPTER 1

INTRODUCTION

History of DDT

1,1,1-Trichloro-2, 2 bis (p-chlorophenyl) ethane (DDT) was one of the ﬁrst
chlorinated organic insecticides discovered. Synthesis of DDT was reported in 1874 but
its effectiveness as an insecticide was not discovered until 1939 (World Health
Organization, 1979). In 1874 a young chemistry student, Othmar Zeidler at the
University of Strassburg, Germany developed DDT (Zimmerman and Lavine, 1946).
Zeidler was not seeking to develop an insecticide when he discovered DDT during his
studies. However, much later, a team of research scientists at a large chemical company
was working to develop more potent insecticides. In the autumn of 1939, Dr. Paul
Muller, working in the laboratories of J .R. Geigy, A.C., Basle Switzerland, synthesized
the same compound that had been developed by Zeidler years before (Zimmerman and
Lavine, 1946; West and Campbell, 1952; Brooks, 1974). Soon, he and his colleagues,
Dr. Paul Lauger and Dr. Robert Weisman, discovered the insecticidal value of DDT
(West and Campbell, 1952, Monclova, 1946). In 1942, the Geigy Company in New York
received its first shipment of DDT from Geigy, Switzerland and then submitted DDT to
the United States Department of Agriculture for testing. Soon its value to the US Armed
Forces for controlling insect borne diseases (e. g. malaria) would be demonstrated. By
1943, DDT was in commercial production at the Cincinnati Chemical Works, and in
early 1944, DuPont, Merck, and Hercules Powder Company also went into commercial

production of DDT (Zimmerman and Lavine, 1946).

 

I0

DDT was used extensively during the Second World War among allied troops and
proved effective in controlling diseases such as malaria and typhus, which are spread by
insects (ATSDR, 2001). Upon use by the military in 1944, the general public began using
DDT extensively to control pests. As a result, DDT was sprayed directly into homes
including onto the ﬂoors, walls, and beds, and also dusted directly onto the body. This
very effective insecticide was widely used because insects not only transmit disease but
also cause discomfort and destroy property and agricultural crops.

From 1945 until it was banned, DDT was one of the most widely used pesticides
for the control of insects on agricultural crops (ATSDR, 2001). As a result of increased
insect resistance to DDT, and growing environmental concerns, the use of this chemical
in agriculture in the United States began to decline by 1959. In 1946, evidence of
toxicity of DDT to humans was reported (Zimmerman and Lavine, 1946). It began to be
evident through ecological studies that ﬁsh and birds of prey suffered most from the
effects of DDT. Based on ecological considerations the United States Environmental
Protection Agency (EPA) banned its use as an insecticide in 1972 (WHO, 1979). Most
other industrialized countries, including West Germany, also banned the use of DDT in
the early 1970’s. In the former German Democratic Republic (GDR) however, DDT was
still produced and used until the end of the 1980’s (Voldner and Li, 1995; Mitra and
Raghu, 1998). Ultimately, a number of other developed countries restricted the use of
DDT except when it was needed for the protection of health (WHO, 1979); however it
remains in use for controlling mosquito-bome malaria in many nations in Africa, Asia
and Latin America. DDT has been recognized as among the best man made compounds

for use as an insecticide. Many environmental groups have advocated that DDT be

phased out of use worldwide by 2007 because of its toxicity and environmental effects
(Roberts, 2001).

DDD (Ll—Bis (4—chlorophenyl)-2,2-dichloroethane) and DDE (2,2-Bis (4-
chlorophenyl)—l,l-dichloroethene) are transformation products of DDT formed in the
environment (Table 1.1). DDD was used as an insecticide for contact control of leaf
rollers and other insects on vegetables and tobacco, but its use has been banned
(Agrochemicals Desk Reference, 1993). DDE has no commercial use but its presence as
a persistent environmental transformation product of DDT is a major concern (ATSDR,
2000). DDT and its metabolites DDD and DDE are still found in various proportions in
soils and sediments and have been reported at 3,422 of 22,000 sites identiﬁed as posing a
danger to human and animal life by the EPA National Sediment Quality Survey (Quensen

et al, 1998).

Environmental Effects

Environmental contamination of land and water from DDT has occurred due to
past production and disposal processes, and agricultural application. The Pine River
reservoir adjacent to the former Velsicol Chemical Corporation in St. Louis, Michigan is
heavily contaminated with DDT and has been designated a Superfund site by EPA. Due
to its inherent structural stability, and strong adsorption to soil and sediment solids, DDT
is particularly recalcitrant in the environment (EPA, 1986). Some investigations have
reported that DDT will remain present in the soil for 2 years while others have found that

the process of degradation can take more than 15 years (Alexander, 1994).

Table 1.1. Chemical names and acronyms of DDT and its derivatives used in the text.

DDT: 1,1,1 trichloro-2,2-bis (p-chlorophenyl) ethane
DDD: 1,1, dichloro-2, 2-bis (p-chlorophenyl) ethane
DDE: 1,1, dichloro-2, 2-bis (p-chlorophenyl) ethylene
DDMU=1-chloro-2, 2-bis (p-chlorophenyl) ethene
DDMS= l-chloro-2, 2-bis (p—chlorophenyl) ethane
DDNU= 1,1-bis (p-chlorophenyl) ethylene

DDOH= 2,2-bis (p-chlorophenyl) ethanol

DDCN= (bis (p-chlorophenyl)-acetonitrile)

DDA= bis (p—chlorophenyl) acetate

DBP= 4,4’- dichlorobenzophenone

Because of its long half-life, DDT and its derivatives (e. g. DDE, DDD) will continue to
be found in the environment in various proportions several decades after introduction.

A major concern about exposure to DDT and its derivatives is bioaccumulation
which is deﬁned as an increase in the concentration of a chemical in an organism over
time compared to the chemical’s concentration in the environment (EXTOXNET, 2000).
This accumulation occurs mainly through prolonged exposure of invertebrates,
earthworms, ﬁsh, and aquatic vegetation to contaminated soil, sediment and water.
Because of its lipophilic nature DDT accumulates in the fatty tissues of organisms
causing bioaccumulation in the food chain.

Bioaccumulation in the food chain can also lead to biomagniﬁcation.
Biomagniﬁcation is a process that results in an increased concentration of a chemical in
an organism higher than the levels found in its food (EXTOXNET, 1999). For example
DDT levels in soil of 10 ppm manifested concentrations of 141 ppm in earthworms and
444 ppm in robins (EXTOXNET, 1999). Biomagniﬁcation may lead to concentrations
high enough to cause adverse effects on reproduction and even death in animals at the top
of the food chain. Consequently even low levels in soil, water, and air may be an
endangerment to certain species.

Studies have shown that DDT is toxic to birds and aquatic organisms. Birds are
exposed to DDT primarily through the food web when they ingest DDT-contaminated
organisms such as ﬁsh (ATSDR, 2001). One of the major concerns with chronic
exposure of birds to DDT is its effects on reproduction, especially eggshell thinning and
embryo mortality (WHO, 1979). DDT derivatives (primarily DDE) have similar toxic

effects and are responsible for the thinning of eggshells of birds and impaired

reproduction in wildlife; predator birds such as the bald eagle are most sensitive to these
effects WHO, 1979; Karte, 1992; Wiemeyer et. al., 1993). It has been further suggested
that DDE causes additional adverse effects including increased embryo mortality in birds

(Heath et al., 1969).

Degradation Mechanisms of DDT- Dechlorination and Dehydrochlorination

There are several degradation pathways of DDT that suggest sequential
dechlorination of the molecule through two types of reactions: reductive dechlorination
and dehydrochlorination. Also, through oxidative reactions, the carbon skeleton of DDT
can be degraded. Reductive dechlorination is the only known biodegradation mechanism
for certain signiﬁcant pollutants including highly chlorinated polychlorinated biphenyls
(PCBs), hexachlorobenzene, tetrachloroethene and pentachlorophenol (Mohn and Tiedje,
1992). Reductive dehalogenation of DDT has been observed to be the primary
degradation pathway in anaerobic soil, sediment, and sewage waste (Hill and McCarty,
1967, Guenzi and Beard, 1967, Burge, 1971; Jensen et al, 1972; Zoro et al. 1974).

The process of anaerobic dechlorination requires an electron donor (reductant)
and proceeds by the removal of a chlorine atom directly from the biphenyl ring, with the
simultaneous addition of a hydrogen atom to the molecule (Hill and McCarty, 1967;
Morrill et al., 1982). The halogen atoms are then released as halide anions. In an
anaerobic community, the availability of electron acceptors is frequently the limiting
resource and a major determinant of species composition. After depletion of more
common natural electron acceptors (e. g. NO3', S04 2') certain chlorinated aromatic

compounds may be used in a process called halorespiration. Therefore, the population of

reducing organisms depends on the availability of electron acceptors. In principle,
reductive dechlorination can occur by either nucleophilic or free radical substitution but
evidence from natural or model anaerobic systems supports a free radical mechanism
(March, 1985). Reductive dechlorination generally occurs under anaerobic conditions but
is also involved in aerobic degradation of certain highly halogenated compounds (Mohn
and Tiedje, 1992). The transfer of electrons to DDT is the essential process for its
transformation to less chlorinated derivatives which may then be subject to aerobic
metabolism. Dehalogenation generally makes xenobiotic compounds less toxic and more
readily biodegradable (Mohn and Tiedje, 1992). Hence, this mechanism is very important
because of its involvement in the environmental fate of pesticides and industrial
chemicals, and its application to bioremediation of pollutants and hazardous wastes
(Mohn and Tiedje, 1992).

Dehydrochlorination involves the removal of hydrogen and chlorine from
organochlorine insecticides forming a double bond. This reaction typically takes place
between the saturated chlorinated carbon and hydrogen on the neighboring carbon
leaving a carbon-carbon double bond on the substrate (Lal and Saxena, 1982). A number
of studies have shown that under aerobic conditions dehydrochlorination is the dominant
reaction (Lal and Saxena, 1982). A very familiar example of this reaction is the
formation of DDE from DDT.

Chlorine substituents on molecules such as DDT contribute to their environmental
persistence; DDT for example is recalcitrant, whereas its non—chlorinated analogue
diphenylmethane is readily biodegradable (Alexander, 1977). Evidence was presented by

Stemersen, (1965) and Wedemeyer (1966) concerning the capability of microorganisms

to dechlorinate DDT. Both recognized that the degradation of DDT by bacteria was
inversely related to the availability of oxygen in the environment. DDT can be
biodegraded to a series of metabolites such as DDD (1,1-dichloro-2, 2-bis (p-
chlorophenyl) ethane, DDE (1,1-dichloro-2, 2 bis (p-chlorophenyl) ethylene, and DDMU
(1-chloro-2, 2-bis (p-chlorophenyl) ethene. Experiments with marine sediments and
radiolabeled DDT showed that biodegradation could transform DDT into DDD, DDE,
DDOH (2,2-bis (p-chlorophenyl) ethanol and DDNS (2,2-bis (p-chlorophenyl) ethane
(Patil et al., 1972) (Figure 1.1). The next sections will further discuss the individual

transformation pathways of DDT.

Q
E Q
—n-d-:

: Q
Q

Q

0

Q

“ rm
HCl H‘
[1]) ”WV 6_’/
C|.C.H kc.
CIOCOCI
11M)
+2H I

OH
H1." COOH
9 r0] '
QO—C_©—Cl ———> Cl—O—C—O—Cl
DDOH ”DA

Figure 1.1. Transformation pathways of DDT by Aerobacter aerogenes (Wedemeyer,
1967).

DDT to DDD

Reductive dechlorination of DDT to DDD is a common first step in the
environmental transformation of DDT (Figure 1.2). The transfer of electrons to DDT is
the essential process for its degradation to form less chlorinated derivatives (Esaac and
Matsumura, 1980) In the environment, dechlorination of DDT is carried out extensively
by microorganisms, but may also occur abiotically. There are several reports of the
conversion of DDT to DDD under anaerobic conditions (Johnson et al., 1967; Kallman
and Andrews, 1963; Wedermeyer, 1966; Guenzi and Beard, 1967; K0 and Lockwood,

1968; Patil et al., 1970).

C1 H
CI \C’Cl CI' CI \C’C]
. H+e- .
H H
DDT DDD

Figure 1.2. Reductive dechlorination of DDT to DDD under anaerobic conditions
(Rockhind and Blackburn, 1986).

Researchers have isolated microorganisms that carry out the conversion of DDT
to DDD under anaerobic conditions. Johnson and Goodman (1967), reported that of 27
microorganisms examined, 23 bacterial species could convert p,p’ DDT to p,p’ DDD
under anaerobic conditions. Escheria coli and Aerobacter aerogenes isolated from the
gastrointestinal tract of rats were capable of degrading DDT to DDD (Mendel, 1966).

Wedemeyer (1967) reported that extracts of Aerobacter aerogenes catalyze the

degradation of DDT to DDD, DDE, DDMU, DDMS, DDNU, DDA, and DBP. By a
technique of sequential analysis the metabolic pathway was postulated as
DDT—>DDD—+DDMU—>DDMSHDDNU—eDDA—ADBP, or DDT—+DDE (see Figure
1.1).

Several studies have been conducted to investigate the conditions under which
dechlorination of DDT takes place. DDT is reductively converted to DDD in sewage
sludge under anaerobic conditions. (Hill and McCarty, 1967; Jensen et al., 1972; Albone
et al., 1972; Zoro et al., 1974.) Zoro (1974) concluded that direct microbial metabolism
does not account for the reductive dechlorination of DDT in treated sewage sludge. His
results showed that the conversion of DDT to DDD in the environment is mediated by
reduced iron porphyrins. Guenzi et.al (1971) published a wide-ranging study that showed
ﬂooded soils favor the degradation of DDT. Under anaerobic conditions in soil,
conversion of DDT to DDD was more rapid than in moist soils where aerobic conditions
presumably exist. In a study by Guenzi and Beard, (1968), DDT was converted to DDD
by microorganisms in an anaerobic soil system.

Because DDD is a highly toxic compound, the conversion of DDT to DDD in soil
cannot be considered as a detoxiﬁcation step. Moreover, DDD is more stable than DDT
itself in soil. Therefore, this conversion may contribute to the persistence of DDT

residues in soil and water (Ko and Lockwood, 1968).

Il

Abiotic systems

In addition to microbially mediated reactions, the addition of some chemical
agents in anaerobic systems may accelerate the transformation of DDT to DDD (Quirke
et al., 1979). These chemical agents serve as catalysts for reductive dechlorination of
organochlorine insecticides under anaerobic conditions. Studies on such catalytic systems
was reported by Castro ( 1964), Miskus et al., (1965), and Zoro et al. (1974). DDT was
effectively converted to DDD in the presence of an anhydrous, anaerobic solution of
ferrous deuteroporphyrin, and by hemoglobin or hematin in the presence of excess
sodium dithionite under anaerobic conditions (Castro, 1964; Miskus 1965). Addition of a
detergent (Tween 80) or ethanol to the incubation media containing hematin, ferrous
sulfate, and DDT increased degradation (Zoro et al., (1974). It has been shown that DDD
formation is related to the levels of ferrous iron present in the anaerobic environment
(Glass, 1972). Furthermore, DDT dechlorination increases as redox potential decreases

(Burge, 1971; Guenzi et al., 1971; Glass, 1972; Parr and Smith, 1974,).

Biological reductive systems

Reductive degradation of insecticides in soil is believed to be mediated by
microbial activity based on two experimental observations: sterile soil is often devoid of
reductive activity, and organic matter amendments to non-sterile soil stimulates the
reductive processes (Esaac and Matsumura, 1980). Studies by Guenzi and Beard (1967,
1968) established that microbial processes were responsible for the dechlorination of
DDT. This was because there was no detection of DDT metabolism in sterile soil after

anaerobic incubation for 2 and 4 weeks with MC DDT added to soil. Likewise, Pfaender

12

and Alexander (1973) demonstrated the presence of DDT-metabolizing organisms in
fresh water, sewage, and marine environments. The rate of anaerobic conversion of DDT
to DDD was increased with the addition of alfalfa. One possible reason for this was that
alfalfa provided favorable nutrients for the microorganisms stimulating the
transformation and increasing the rate of conversion (Guenzi and Beard, 1968).
Similarly, Burge (1971) found DDD to be the only product after incubation of DDT in
soil amended with alfalfa or alfalfa distillate. Johnsen (1971) reported that after
incubation with or without addition of cattle manure, DDT was degraded readily in soil,
principally to DDD. Conversion of DDT to DDD was more rapid in waterlogged soil
than in moist soil where aerobic conditions presumably existed (Ko and Lockwood,
1968). In addition, Miskus et al (1965) reported that bovine rumen fluid degrades DDT
to DDD. Matsumura et al. (1971) examined the fate of DDT and reported that the
majority of microbial isolates from water and bottom silt of Lake Michigan reductively
dechlorinated DDT to DDD. Likewise, studies of isolated bacteria showed DDD to be the
main metabolic product in lab cultures (Barker et al., 1965; Stemersen, 1965; Subba Rao

and Alexander, 1985; Wedemeyer, 1967).

DDT to DDE

DDE can be formed from DDT via dehydrochlorination, an oxidative process
(Mohn and Tiedje, (1992) (Figure 1.3). Aerobic dehydrochlorination of DDT
simultaneously removes the hydrogen and chlorine from the aliphatic portion of the
molecule resulting in the formation of a carbon-carbon double bond. Patil et al., (1970)

found that in aerobic environments such as ocean water, DDE was the major

13

biodegradation product of DDT. A number of studies have demonstrated the metabolism
of DDT to DDE in soils. Guenzi and Beard, (1968) completed a study that suggests DDT

is probably degraded ﬁrst to DDE in aerobic soil treatments.

Cl\| :10 HCI CI\C,C1
II

DDT

Figure 1.3. Aerobic dehydrochlorination of DDT to DDE (Rockhind and Blackburn,

1986)

Dehydrochlorination of DDT to DDE has been observed in aerobic and anaerobic
environments, and in model systems containing reduced poryphins (Guenzi and Beard,
1967; Burge, 1971; Marci et al., 1978; Quirke et al., 1979). Wedemeyer (1967)
conﬁrmed the formation of DDE after dehydrochlorination of DDT by whole cells or
cell—free extracts of Aerobacter aerogenes. Similarly, studies by Hitch and Day (1992)
and Spencer et al., (1996) indicated aerobic soil metabolism of DDT to DDE. Hitch and
Day (1992) concluded that certain conditions (e. g., metals content, moisture content) of

the soil environment affect the degradation of DDT to DDE.

l4

DDD TO DDMU

The transformation of DDD to DDMU has been proposed in DDT metabolism
studies (Quirke et. al., 1979, Quensen et. al., 1998). This conversion occurs by reductive
dechlorination to DDD followed by dehydrochlorination to DDMU in aerobic and
anaerobic environments. This, and the subsequent conversion of DDMU to DDA and
DBP, was observed by Quirke et. al. (1979). Quensen et al., (1998) observed a trace
amount of DDMU formed from the transformation of DDD in Palos Verdes sediments.
Wedemeyer (1967) observed the metabolic pathway of DDT to be DDT—> DDD—r
DDMU—> DDMS——> DDNU—> DDA—> DBP. These dechlorination reactions occurred
under aerobic and anaerobic conditions and were carried out by microbial enzyme

systems with very speciﬁc environmental requirements to support these reactions.

DDE to DDMU

Typically, DDE has been viewed as a dead end product in the metabolism of DDT
(Hay and Focht, 1998). DDE is a metabolite of DDT that is very persistent in the
environment. It was believed for a long time that DDE was not microbially transformed
but studies have proven reductive dechlorination under anaerobic conditions. Quensen et
al. (1998) reported the formation of DDMU from the anaerobic microbial degradation of
DDE in the Palos Verdes sediments. There were signiﬁcant differences in the rates and
extents of DDE dechlorination among the sediments from the three sites investigated.
Furthermore, another investigation of the factors controlling the rate of DDE
dechlorination to DDMU in the sediments showed that sediment depth and temperature

affect the dechlorination rate of DDE (Quensen et.al., 2001 ).

15

Further Transformations

In studies on the transformation of DDT, the presence of intermediate
metabolites, aside from the ones previously discussed have been observed. DBP, DDA,
and DDCN are some of these metabolites formed from the degradation of DDT in
anaerobic systems. Microorganisms in pure cultures have shown transformation of DDT
to be: DDT—rDDD—rDDMU—>DDMS—rDDNU—>DDA—>DBP or DDT—+DDE
Wedemeyer, 1967). Marci et. al., (1978) found that DDCN was formed during the
incubation of DDT with sewage sludge. These results are very similar to those observed
by Jensen et. al., (1972) which showed the transformation to DDD, DDMU, DBP, and

DDCN in anaerobic sewage Sludge.

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Jensen, 8.; Gothe, R.; Kindertedt, MO. 1972. Bis-(p-Chlorophenyl)-Acetonitrile
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Kallman, B.J., and AK. Andrews, 1963. Reductive dechlorination of DDT to DDD by
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Matsumura, F., Patil, K. C., and Boush, GM. 1971. DDT metabolized by
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Miskus, R. P., Blair, D. P., and Casida, J .E. 1965. J. Agric. Fd. Chem. 13:481.

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Parr, J. F., and Smith, S. 1974. Transformation of DDT in an Everglades muck as
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environments and energy sources on the degradation of DDT. Soil Science. 110:306-
312.

Patil K.C., Matsumura F., Boush GM. 1970. Degradation of Endrin, Aldrin, and DDT
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399.

Pfaender, F.K., and Alexander, M. (1973). Effect of Nutrient Additions on the Apparent
Cometabolism of DDT. J. Agr. Food Chem. 21: 397-399.

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724.

Quirke, J .M.E.; Marci, A. S. M.; Eglinton, G. 1979. The Degradation of DDT and its
degradative products by reduced iron (III) poryphyrins and ammonia. 3: 151-155.

Rochind, M. L., and Blackburn. 1986. Microbial decomposition of chlorinated aromatic
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1996. DDT persistence and volatility as affected by management practices after 23 years.
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19

Stemersen, J. H. V. 1965. DDT metabolism in resistant and susceptible stable ﬂies and
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20

 

165
by

the

CHAPTER 2
SITE HISTORY AND METHODOLOGY

Site History

The Velsicol Chemical Corporation plant site formerly known as Michigan
Chemical is located on the Pine River in St. Louis, Michigan (Figure 2.1). The chemical
plant operated between 1936 and 1978 and manufactured a wide array of chemicals
including 1,1,1- trichloro—2, 2 bis (p-chlorophenyl) ethane (DDT), polybrominated
biphenyls (PBB), hexabromobenzene (HBB), and tris (2,3-dibromopropyl) phosphate
(TRIS). In 1974, the facility became the subject of intense scrutiny after bags of the
white powder PBB were mistakenly shipped instead of the feed additive magnesium
oxide to operators throughout Michigan resulting in feed for dairy cattle that was
contaminated with PBB. This action led to investigations by the Michigan Department of
Natural Resources (DNR) and the US. Environmental Protection Agency (EPA) to
determine if there were other negligent activities by Velsicol that presented a threat to the
environment. They discovered that poor waste management practices such as process
waste discharges directly into the Pine River had led to widespread contamination of the
plant site and the adjoining Pine River Irnpoundment. There was considerable
contamination of site soils and Pine River sediments with DDT, PBB, HBB and Tris.
There were also elevated levels of DDT and other contaminants in ﬁsh (Table 2.1).

DDT levels in the Pine River sediments adjacent to the plant as high as 44,000 ppm have
resulted in DDT levels in ﬁsh that greatly exceed the maximum safe level of 5 ppm as set
by the Food and Drug Administration (FDA). Because of the widespread contamination

the site was deemed to pose a threat to human health and the environment.

21

Pine River,
St. Louis, Michigan

 

 

 

 

Former Plant Site

Mill St.

 

 

£5; Washington St.

 

State St.

Figure 2.1. The Velsicol Chemical Corporation plant site located on the Pine River
in St. Louis, Michigan. Adapted from Morris et. al., (1993).

22

As a result of the investigations the State of Michigan and the US. EPA closed the

facility in 1976 and remedial measures for the contaminated site began in October, 1978.

 

 

Collection Avg. Cone. Max. Cone. Min. Cone.

Location Year (ppm) (ppm) (ppm)
Dam 1983* 0.08 0.22 0.05
Darn 1985 9.66 18.66 5.27
Dam 1994 23.79 47.31 1.65
Dam 1997 26.82 72.56 10.61
Irnpoundment 1989 10.48 39.76 0.06
Irnpoundment 1995 16.15 43.27 0.51
Irnpoundment 1997 34.57 89.92 2.47

 

Table 2.1. Concentrations of total DDT in skin-off carp fillets from below the St. Louis
Dam and in the St. Louis Irnpoundment. *Fish collected in 1983 were not analyzed for
DDE and DDD, only DDT (EPA, 1998).

The initial remedial measures stopped discharges from the plant into the Pine

River and subsequently the buildings and structures on the Site were demolished.

Agreements made between EPA, the State of Michigan and Velsicol included a plan to

stop off-site migration of DDT and the other contaminants. This plan included

excavation of the contaminated on-site soil, isolation of the site with a low-permeability

slurry wall and clay cap, and other measures to control and monitor the boundaries of the

site. After many hearings, a consent judgement-agreement was made between the

Michigan DNR, EPA and Velsicol in 1982. The judgment did not require Velsicol to

treat or remove contaminated sediments from the Pine River/ St. Louis Irnpoundment (the

so-called no-action alternative) but they did pay restitution to the State for studies at the

site. To deal with the issue of ﬁsh contamination in the River, the State of Michigan

issued a no consumption advisory for all species of ﬁsh in the river. Regulators were

23

convinced that with time new sediment deposition would isolate the contaminated
sediments and levels of DDT in ﬁsh would decline.

The Velsicol Chemical Plant Site has been under continuous investigation to
monitor the effectiveness of the slurry wall and cap, as well as the contaminant levels in
ﬁsh. The agencies collected sediment and ﬁsh data throughout the 1980’s and 1990’s.
The studies revealed that contaminant levels in sediments had not decreased over time,
and that there were still elevated or perhaps increasing levels of DDT in ﬁsh well above
the FDA tolerance level of 5 ppm (Table 2.1).

As a result of these ﬁndings, the Pine River reservoir was designated a Superfund
Site in 1986, and US. EPA began emergency remedial action at the site in 1998. The
plan was to immediately remove the most contaminated sediments, and then continue
removal of lesser-contaminated adjacent sediments. The cleanup goal for this project is
5ppm DDT in sediments. This concentration would result in the removal of the majority
of the contamination and therefore result in acceptable risk levels. At the 5 ppm
concentration level the reduction of DDT concentrations in carp was reduced from, ~42.5
ppm to ~1.7 ppm, making the ﬁsh clean enough to eat (EPA, 1998). The remedial plan
chosen to carry this out would remove 533,000 pounds of DDT from the St. Louis
impoundment. This involved physically isolating certain sections of the River with
temporary coffer dams (sheet piling) to reduce the amount of resuspension of
contaminated sediments during the dredging operations and to enhance dewatering.
Accordingly, the plan involved removal of water from the impounded area and treatment
using sand ﬁlters and activated carbon columns; treated water was then pumped back into

the River. To ensure discharge requirements were being met, periodic sampling was

24

performed. The exposed sediments would be solidiﬁed by treating with a strengthening
agent such as kiln dust, excavated and disposed of at on offsite RCRA subtitle C landﬁll.
The proposed date of completion for this work was the end of 2001, but this date has now

been extended.

25

Introduction

Degradation pathways of DDT in the environment have been studied for decades.
These studies have indicated several possible chemical and biological transformations
that lead to the degradation of DDT in the environment. Studies have shown degradation
under aerobic and anaerobic conditions. Microorganisms in soils, sediments, sewage
sludges as well as iron porphyrins have proven effective in degrading DDT. Laboratory
studies have used radiolabeled compounds to help identify the speciﬁc degradation
pathways. These studies have provided important insights but questions still remain
regarding which speciﬁc mechanisms are operative under different environmental
conditions, whether these reactions are chemical or biological in nature, and the exact

sequence of intermediates produced from these reactions.

Objective

In this ﬁrst study we examined the fate of DDT, DDD, and DDE in anaerobic
Pine River sediments contaminated with DDT. The extent of degradation was monitored
over a 32 week time period. The objectives were to determine the pathways of DDT
degradation and determine the abiotic and biotic contributions to the anaerobic
transformation of DDT. An additional objective was to determine if sediment depth
inﬂuenced DDT degradation in the Pine River. A second study sought to evaluate the
dechlorination of DDT by microbial communities with demonstrated capability to
dechlorinate PCBs, as well as to determine if FeSO4 amendments would stimulate the

transformation of DDT.

26

 

\rlr'r

MATERIALS AND METHODS

Materials

The chemicals DDT (99% purity), 1,1-dichloro-2, 2 bis (p-chlorophenyl) ethane DDD
(99%purity), 2,2-bis (p-chlorophenyl) 1,1-dichloroethylene DDE (99.9% purity) were
obtained from Ultra Scientiﬁc, Co., North Kingstown, RI. DDMU (100% purity) was
obtained from AccuStandard Inc., New Haven, CT. Ring labeled l4C-DDT (97 %
radiochemical purity) l4C-DDD (3.6mCi/mmol, 100 % radiochemical purity) l4C-DDE
(13mCi/mmol, 98% radiochemical purity) were obtained from the Sigma Chemical Co.

St. Louis, MO.

Sediment Sampling and Analyses

Pine River Sediments

Sediments were collected from two different Sites on the Pine River at the
Velsicol Chemical Corporation Superfund Site in St. Louis, Michigan (Figure 2.2).
Sediment samples were collected by inserting 5 cm x 90 cm polyvinyl chloride (PVC)
pipes into the sediment as deep as possible. Holed rubber stoppers with an attached
rubber ﬂap were placed at one end of the pipe to release water and gas as the PVC pipes
were inserted. As the pipes were withdrawn the rubber ﬂap sealed to prevent loss of
sediment and overlying water. After withdrawal, the pipes were capped, tightly sealed
with tape to minimize oxygen exposure and transported to the laboratory. The samples

were stored at 4°C until used.

27

Pine River,
St. Louis, Michigan

 

 

 

 

 
   
   

 

Core 1 1

Dam
Former Plant Site

Mill St.

 

 

 

State St.

Figure 2.2- The Velsicol Chemical Corporation plant site located on the Pine
River in St. Louis, Michigan and sampling points for core 4 and core 11. Adapted
from Morris et. al., (1993).

Two sediment cores were selected for study in this experiment. Samples
containing trace levels (~0-5 ppm) of contaminants were collected slightly downstream
(core 11) of the region of highest DDT contamination and samples containing a greater
DDT concentration (~2000-4000 ppm) were obtained slightly upstream of this region
(core 4) (EPA, 1986). Anaerobic sediment slurries were prepared in a 1:1 ratio of
sediment to Reduced Anaerobic Mineral Media (RAMM) (Shelton and Tiedje, 1984) as

described by Quensen et al. (1998). The sediments were removed from the PVC pipes

and separated into two sections (top and bottom sections), each approximately 10 cm

28

long. They were immediately placed into oxygen-free [NZ/C02 (80:20, vol/vol)] ﬂasks,
which were continuously ﬂushed using a Hungate gassing apparatus (Hungate, 1968).
The ﬂasks contained an approximately equal volume of RAMM (600ml) and sediment

which were slurried using a magnetic stirrer.

Pine River Sediment Microcosms

Microcosms were setup using core 11 and core 4 sediments amended with MC
radiolabelled- DDT, -DDD or -DDE as substrates for the assays. A mixture of each
compound was made containing 6.9 mg of l4C-DDT, and 141.3 mg of unlabelled DDT;
8.9 mg of ”C-DDD and 51 mg of unlabelled DDD; and 2.5 mg of ”C-DDE and 56.8 mg
of unlabelled DDE diluted with acetone to 5.2, 2.1, and 2.1 mL of DDT, DDD, and DDE
respectively. This gave a solution concentration of 28.5 mg/mL for DDT, 28.5 mg/mL
for DDD, and 28.2 mg/mL for DDE and an activity of approximately 1.07X 105 dpm/mL
as veriﬁed by liquid scintillation counting (Beckman LS6500). A volume of 7 mL of
sediment slurry (containing about 2 g of sediment on a dry weight basis) was transferred
to N2/COz purged 20 ml glass vials and spiked with 7111 of ”C-labelled-DDT, -DDD, or-
DDE. This gave an activity of 7.5 X 105 dpm/microcosm. The microcosms were mixed
thoroughly after addition of the DDT, DDD, or DDE, capped with Teﬂon stoppers,
sealed with aluminum crimp caps, and stored in the dark at room temperature (22° to 25°
C).

Samples for the determination of biological (live) as well as non-biological
(sterile) transformations occurring in the sediments were set up for this study. Four

replicate samples were sacrificed at intervals of 0, 4, 8, l2, 16, 24, and 32 weeks for live,

29

and 0, 8, l6, and 32 weeks for the sterile samples, for each compound (DDT, DDD,
DDE) and each section of sediment. The sterile microcosms were autoclaved three
consecutive days for 1 hour before the addition of 14C-labeled compounds to the vials.

Upon completion of the speciﬁed incubation period, samples were immediately frozen.

3O

Microcosms amended with Pine River and Silver Lake Inocula

Additional microcosms were setup using inoculated sediment slurries previously
shown to dechlorinate PCBs. N on-PCB-contarninated Red Cedar River sediments were
prc-incubatcd, sterilized, then inoculated with microorganisms eluted from downstream
Pine River and Silver Lake sediments. Speciﬁcally, samples of air-dried Red Cedar
River sediments (2 grams) were weighed into 20 mL glass vials. The vials were
evacuated and reﬁlled with N2 in an anaerobic chamber then sealed with butyl stoppers.
For the pre-incubation step, 1 L of RAMM was prepared with the addition of 1 mL of
ethanol and 25 mL of inocula eluted from the speciﬁed sediment slurry as described by
Quensen et. al., (1988; 1990). The microcosms were ﬁrst tested for their ability to
maintain strict anaerobic conditions. For this, 6 mL of inoculated RAMM were added to
each sample vial with a sterile syringe through the stopper and incubated in the dark for
one week at 37°C. After 7 days, headspace gas was analyzed for methane production
then autoclaved one hour for two consecutive days. The sample vials were then re-
inoculated to determine the DDX dechlorination activity. Using sterile anaerobic
technique the butyl stoppers were removed from the vials. The vials were then ﬂushed
with oxygen-free N2 ICOZ and lmL of eluted inoculum and 7 11L of either l4C-DDT, -
DDD, or -DDE were added. Some of the Silver Lake inoculated microcosms were also
amended with 1 mL of a 10 mM FeSO4 solution. All vials were rescaled with Teﬂon
stoppers and aluminum crimp caps and stored in the dark at room temperature for the
predetermined incubation period. For the sterile (abiotic) samples, the microcosm setup

was the same with the exclusion of inoculum addition.

31

Red Cedar Unamended Microcosms.

Red Cedar sediments were tested for their ability to dechlorinate DDT.
Sediments were collected from the Red Cedar River, placed in glass jars, capped and
transported to the laboratory and stored at 4°C. Subsequently, sediments were removed
from the jars and placed into ﬂasks which were being ﬂushed with oxygen-free N2/C02
(80:20, vol/vol) using a Hungate gassing apparatus (Hungate, 1968), and which contained
an equal volume (600 ml) of RAMM. The sediment-RAMM mixtures were slurried
using a magnetic stirrer. Red Cedar sediment slurry (7 mL) was transferred into
continuously ﬂushed vials, Spiked with 7 ul of either MC radiolabelled- DDT, -DDD or -
DDE, capped with Teﬂon stoppers, sealed with aluminum crimp caps, and then stored for

their incubation period.

Microcosms amended with Hudson River Inocula and FeSOa

Red Cedar sediment microcosms were setup as described above, inoculated with
microorganisms eluted from Hudson River sediments, then amended with FeSO.; to
evaluate the potential stimulatory effects of FcSO4 on dechlorination (Zwiemick ct a1,
1999). Sediment microcosms containing Red Cedar sediment slurried in RAMM were
autoclaved for one hour two consecutive days. The samples were rc-opcned and ﬂushed
with a continuous stream of N2/COz while adding lmL of eluted Hudson River inoculum
along with 7hr of either l4c radiolabelled -DDT, -DDD or -DDE. Some of the Hudson
River microcosms were also amended with lmL of a 10mM FeSO4 solution. The vials
were rescaled with Teﬂon stoppers and aluminum crimp caps and stored in the dark at

room temperature for the predetermined incubation period. Abiotic microcosms were

32

setup in the same fashion, with the exclusion of inoculation, then autoclaved for one hour
on three consecutive days before the addition of l4C-labeled compounds to the vials.
Four replicate samples were sacriﬁced at intervals of 0, 8, 16, 24, and 32 weeks for live,
and 0, 8, 16, and 32 weeks for the autoclaved controls. Upon completion of the speciﬁed

incubation period, samples were frozen until subsequent analysis.

Methane Analysis

After each incubation period, methane analysis was used to demonstrate strict
anaerobic conditions (production of methane), as well as to show the biological activity
in each microcosm. Headspace gas (20 ul) of the microcosms withdrawn was analyzed

for methane using a gas chromatograph equipped with a ﬂame ionization detector.

Extraction and TLC Analysis

According to the methods of Quensen et.al (1998), frozen samples were thawed
and extracted three times by shaking for 10 minutes with 7 ml of petroleum ether and
acetone (5:2,volzvol). Solvent phases were combined and evaporated to a volume of ~500
til under a stream of dry nitrogen gas. Sample extracts (20 til) were spotted on activated
silica gel TLC plates. The plates were developed to 15 cm with a (5:95,vol: vol) of
petroleum ether and hexane in a lined TLC chamber at room temperature.
Autoradiography was used to determine locations of the parent compound and
metabolites on the TLC plates. Kodak Scientiﬁc Imaging Film (X-OMAT AR) was

exposed to the TLC plates for 7 days at negative 20°C before developing.

33

Autoradiography ﬁlms and TLC plates were placed side by side on a light box where the
parent compound and metabolite zones were visualized and marked for scraping. Liquid

scintillation counting was used to determine the '4C activity in the scrapings.

34

REFERENCES

Forba, R.W. US. EPA Pine River Contamination Survey, St. Louis, MI, EPA-330/2-80-
031, 1980.

Forba, R.W. US. EPA Summary of Pine River Reservoir Sediment Sampling Survey, St.
Louis, MI, EPA-330/2-82-001, 1982.

Hungate,R.E. 1968. Adv. Microbiol. 3B,117.

Mom's, P.J., Quensen, J.F III, Tiedje J.M., and Boyd, SA. 1993. An assessment of the
Reductive Debromination of Polybrominatcd Biphenyls in the Pine River reservoir.

Environmental Science and Technology. 27: 1580-1586.

Quensen, J .F III; Tiedje, J .M.; and Boyd, SA. 1988. Reductive dechlorination of
Polychlorinatcd Biphenyls by anaerobic microorganisms from sediments.

Quensen, J .F 111; Boyd S.A.; and Tiedje J .M. 1990. Dechlorination of Four commercial
Polychlorinatcd Biphenyl Mixtures (Aroclors) by Anaerobic Microorganisms from
Sediments. Appl and Environ. Microbiol. 56: 2360-2369.

Quensen, J .F. III, Mueller, S.A., Jain, KM. 1998. Reductive dechlorination of DDE to
DDMU in marine sediment microcosms. Science. 280: 722-724.

Shelton, D.R.; Tiedje,J.M. 1984. Appl. Environ. Microbiol. 47 850.

US. EPA Region 5. 1998. National Remedy Review Board, National Remedy Selection
Brieﬁng: Velsicol Chemical Site, St. Louis, MI.

U.S.EPA Region 5. 1998. Draft Streamlined Remediation Investigation Report:
Velsicol Chemical Site, St. Louis, MI.

Zwiemick, M.J., Quensen J .F. III, Boyd, SA. 1999. FeSO4 amendments stimulate
extensive anaerobic PCB dechlorination.

35

CHAPTER 3

RESULTS AND DISCUSSION

DDT Transformations

DDT, PBBs, HBBs, and heavy metals are some of the compounds that heavily
contaminate sediments at the Pine River Reservoir Superfund Site in St. Louis, Michigan.
Studies on the Pine River Reservoir indicated the presence of DDT in concentrations as
high as 4% by weight of the sediment (Forba et al. 1980; 1982). Levels of total DDT in
ﬁsh sampled between 1985 and 1997 have been alarmingly high and may be increasing.
The average DDX (Z DDT+DDD+DDE) concentrations in ﬁsh taken from below the
Pine River Dam between 1985 and 1997 increased from lOppm to 27 ppm, and between
1989 and 1997 the concentrations below the impoundment increased from lOppm to 35
ppm (EPA, 1998). The continuing high levels of DDT in Pine River sediments and in
ﬁsh taken from the Pine River in proximity to the Velsicol Superfund Site led to
investigations focusing on potential environmental transformation of these compounds in
the sediments. Morris et. al. (1993) investigated the degradation of PBBs in the Pine
River sediments and found evidence of only very limited in situ anaerobic
biobromination of PBBs. Microorganisms capable of PBB debromination were found in
Pine River sediments, however, high concentrations of co-contarninants were believed to
inhibit in situ debrorrrination. Quensen et al. (1998) discovered the presence of DDMU
originating from the dechlorination of DDE in Palos Verdes sediments. These ﬁndings
led to studies by Roberts (2001) to determine if the Pine River could support the
dechlorination of DDT and its metabolites DDD and DDE. Sediment analysis identiﬁed

the DDT metabolites DDD, DDE, and DDMU. Roberts (2001) sought to identify the

36

origin of DDMU detected in the sediments and evaluate the transformations of DDT,
DDD and DDE under strict anaerobic conditions in controlled laboratory experiments. In
these investigations the transformation of DDT to DDD was observed, but no
transformations of DDD or DDE occurred. The present study was undertaken to further
investigate the transformation of DDT, DDD, and DDE in Pine River sediments under
anaerobic conditions and to evaluate the dechlorination of DDT by microbial
communities with demonstrated capability to dechlorinate PCBs. The Pine River
sediments did support the anaerobic dechlorination of DDT to DDD by both biotic and
abiotic transformations, however it did not support transformations of DDD or DDE.
Similar results were obtained in previously non-DDT contaminated Red Cedar sediments

inoculated with microorganisms from Pine River, Hudson River and Silver Lake.

Transformations in biologically active microcosms

Pine River Sediments incubated under anaerobic (methanogenic) conditions
supported dechlorination of DDT to DDD in both sediment cores studied, top and bottom
sections. In core 4 top and bottom sections there was dechlorination of DDT to DDD
(Figure 3.1). The greatest amount of dechlorination was observed at the termination of
the incubation (32 weeks) for both top and bottom sections. The average recovery for
DDT was 45% (of 14C DDT added) for 16 and 24 weeks for the top section and 39% and
32% for 16 and 24 weeks, respectively, for the bottom section. The average DDT
recovery at 32 weeks for the top section was 32% and 19% for the bottom section. Thus,
based on loss of DDT, the greatest amount of dechlorination occurred in the bottom

section of sediment.

37

 

1(1) 1(1)

 

 

 

 

 

 

 

 

 

 

 

TOP
80 80
60 . 6O
:—'
4o \ l40
2m ' .20
'O
a
d)
g 0 0 :E’
32100 100 E;
U I
E
s3 ~80
~60
'40 +1131:
—o—[]])
‘1) +01gin
+314
-0
0 8 16 24 32
ImrbationTrneOhbelG)

Figure3.l. TrarsforrmtimofIDTtoHDDinCue4tqmrﬂbottan

layeramcrobicsedinertrriaooosnsdningaﬂwwkirnbaimpaiai

38

 

In core 11 sediments there was also dechlorination of DDT to DDD (Figure 3.2).
Transformation of DDT occurred for the top and bottom sections of sediments, with the
greatest amounts occurring in the bottom section of sediment. The average recovery of
DDT at 16 and 24 weeks was 23 and 30%, respectively, for the top section and 11 and
9 % for the bottom section of sediment. The greatest amount of dechlorination occurred at
3 2 weeks for the top and bottom sections of sediment. The amount of DDT recovered at
3 2 weeks (termination of experiment) was 18 and 5% for the top and bottom sections of
sediment, respectively (Figure 3.2).

As the amount of DDT recovered decreased overtime, the amount of DDD
recovered increased (Figures 3.1, 3.2, 3.3). For core 4 from 16 to 32 weeks the amount
of DDD recovered ranged from 48 to 56% for the top section of sediment and 52 to 68%
for the bottom section of sediment. For core 11 between 16 and 32 weeks, the recovery

range for the top section of sediments ranged from 58 to 66% and 67 to 72% for the
bottom section of sediment. A Student’s t-tcst was performed to assess the statistical
signiﬁcance of differences in DDD for the top and bottom sections of core 4 and core 1 1.
When comparing the statistical signiﬁcance (P<0.05) between the differences in
recoveries from the top and bottom sections and between the two cores, there was no
Si gni frcant difference between the top and bottom sections in core 4, or in the top and
bottom sections of core 11.
1 Overall a slightly greater amount of dechlorination occurred in core 11 where
72 % of the DDT had been transformed to DDD, as compared to 68% in core 4 (based on
1‘ eCO Verics). However when compared statistically there was no Signiﬁcant difference

betWeen the top section of core 4 and the top section of core 11, or between the bottom

39

% of 14C Recovered

1(1)

 

TOP

 

 

 

 

 

 

 

 

 

Flgm'e3.2 AnaerobicdechlorinationofDDTtoDDDinCae 11 topandbottom
layeranaembicsedinentnicmoosnsovera32weekirmbationperiod

4O

W0 of 14C recovered as DDT and DDD

 

 

+ 4T-DDD
—0— 4B-DDD
+ 4T-DDT
—6— 4B-DUF

 

 

 

 

 

 

Corell

 

-0— llT—DDD
—O- llT—DDT

 

 

 

 

40. ’ + llB-DUI‘
/ . —v— llB-DDD
20 - ' .
.F ‘

Incubation Tum (Weeks)

 

 

Figure 3.3. Corrparisonofanaerobic dechlorinationofDDTinCae4and 11
top(T)andbottom(B) sections overa32 weekincubationperiod.

41

Section of core 4 and the bottom section of core 11. Core 11 was taken upstream from

the region of highest contamination and core 4 that was taken downstream from this
region. Based on previous sediment analyses (EPA, 1998) DDX levels in the proximity
of core 11 were estimated as between ~0 and 5 ppm, and between ~2000 to 4000 ppm for
sediments in proximity to core 4. It is also likely that sediments in the area that core 4
was taken were heavily co-contarrrinated with the products from the outfall, including
PBB, I-[BB and TRIS. Despite these differences in background levels of DDX and co-

contaminants, the transformation of DDT to DDD was very similar in core 4 and core 11
sediments.

Other metabolites were observed on the TLC plates but not analyzed because they
were present in all samples in trace amounts (Figure 3.4). Some dechlorination of DDT
was observed in the time 0 samples. This could have been caused by abiotic reactions
taking place in the freezer or transformations occurring before the samples were frozen.

At 0 weeks DDD was observed in the range of 2-12% (% of DDT added) for all
sediments. (Table 3.1).

6
”W“ “ 552's.- . *4 m : pg m $3 “r

  

”summits-mm

     

F igur e 3.4. Autoradiograms of parent compounds and metabolite zones on TLC plates
for transformations of DDT.

42

T able 3.1. Recovery of 14C DDT and transformation products in Pine River sediments

b ased on TLC Analysis. Sample microcosms were established using the top (T) and

bottom (B) halves of sediment cores from two different locations (cores 4 and 11). The
standard deviation is reported in parenthesis. *24 week Autoclaved samples were not
§;<3tup.

 

 

 

 

 

Incubation Time Core 4T Core 48 Core 1 1T Core 11B
Weeks Live Auto Live Auto Live Auto Live Auto
% Recovered as DDT
0 96 96 92 89 93 94 85 88
(1) (2) (0) (4) (0) (2) (5) (3)
16 45 75 39 74 23 48 ll 47
(1) (6) (7) (6) (16) (5) (3) (6)
24 45 * 32 * 30 * 9 *
(6) * (3) * (9) * (0) *
32 32 63 19 57 18 59 5 73
(9) (7) (4) (9) (5) (9) (1) (5)
% Recovered as DDD
0 2 2 6 7 6 5 l2 9
(1) (0) (0) (1) (0) (1) (5) (2)
16 49 21 52 21 65 40 68 40

(1) (5) (6) (5) (13) (5) (6) (4)

24 48 * 59 * 58 * 72 *
(5) * (2) * (1 1) * (1) *

32 56 28 68 26 66- 34 67 18
(9) (2) (6) (8) (5) (7) (2) (2)

% Recovered as Polar Metabolites

 

 

 

0 0 1 l 4 l 0 2 1
(0) (0) (0) (3) (0) (0) (1) (1)
16 3 4 7 4 8 12 16 13
(1) (0) (2) (0) (3) (2) (3) (3)
24 3 * 7 * 10 * 16 *
(0) * (5) * (3) * (0) *
32 7 8 10 15 l 1 6 24 6
¥ (2) (8) (3) (3) (1) (2) (2) (3)
% Total Recovery
0 98 99 99 100 100 99 99 98
16 97 100 98 99 96 100 95 100
24 96 * 98 * 98 * 97 *

32 95 99 97 98 95 99 96 98

 

43

1 'ransformations in autoclaved microcosms

Transformation of DDT to DDD was observed in the autoclaved treatments
(Figures 3.5, 3.6, 3.7). The amount of DDT transformed in the sediments was less in the
autoclaved sediments than for the live treatments. The amount of DDT recovered in the
Core 4 top layer sediments ranged from about 63% to 96% with the greatest amount of
dechlorination occurring at 32 weeks (63% of added DDT recovered). In the core 4
bottom layer sediments, the DDT recovered ran gcd between 57-89% with the greatest
amount of dechlorination (lowest DDT recovery) occurring at 32 weeks (57% of the
added DDT recovered) (Figure 3.5). In the core 11 top layer sediments, the amount of
DDT recovered ranged from 48% to 94% over the 32 week time period. In the bottom

1 ayer sediments, the DDT recovered ranged between 47% to 88% with the greatest

amount of dechlorination occurring at 16 weeks (47% of added DDT recovered) (Figure
3 - 6).
There was a considerable amount of transformation activity occurring in both

cores, but there was a larger percentage of DDT dechlorination occurring in the Core 11
sediments based on recoveries. T-tests were also used to compare autoclaved
microcosms at 32 weeks. When compared statistically, at a 95% conﬁdence level there
Was a signiﬁcant difference between the bottom section of core 4 and the bottom section
0f core 11. There was also a signiﬁcant difference observed between the top and bottom
Sections of core 1 1. In both sections of core 11 the greatest amount of dechlorination
Occurred at 16 weeks whereas in core 4 the greatest amount of dechlorination occurred at

32 Weeks. Furthermore there was also the increase in DDD as the DDT was transformed

44

A s the percentage of 14C recovered as DDT decreased, the percent recovery of DDD
i ncreased over the 32 week time period (Figure 3.5, 3.6, 3.7).

For the abiotic and live microcosms methane production in the headspace of
rmcrocosms was used as an indicator of biological activity in sediment microcosms. The
l ive microcosms show headspace methane production up to 37% over the 32-wecks,

i ndicating that there was biological activity in the sediments. The methane production in
autoclaved sediments was less than 0.01 % or undetectable, therefore giving no indication
of biological activity in these sediments. The observation of DDD production in the

autoclaved sediments indicates that DDT dechlorination was occurring by abiotic

reactions as well as biologically.

45

 

1(X) ICX)

 

 

 

 

 

 

 

 

 

 

 

 

 

TOP
80‘ 80
60* 60
40~ 40
B 20- 20
i3
>
8
o 0 at
04
U 100 5
3
SS ~80
~60
4O +DD1‘
—O—DDD
20 20 +314
0 e a .0
0 8 16 24 32
IncubationTrrre(Wecks)

Figure 3.5. Transfonmtion ofDDTtoDDDinCore4 (topandbottomseotions)
abiotic sediment m'crooosrrs during a 32 week incubation period

46

 

 

 

 

 

 

 

 

 

 

100‘ 1m
1
TOP
80- - 80
60‘ - 60
40- ~ 40
“O 20‘ b 20
d)
i; i
8 0 e e p 0
32100 100 :E’
O ,_ U
E i
go 80? Bottom 80
60‘ 60
40* ° 40 [1211‘
—o—DDD
20‘ . 20 +0n'gin
—+—CH4
0 9 e 0
0 8 16 24 32
InmbationTrrrc(Weeks)

Figure 3.6. Transfmnation ofDDI‘to DDDin Core 11 (top andbottom sections)
abiotic sedirrmt nicrooosrrs during a 32 wedt incubation period.

47

 

% of 14c recovered as DDT and DDD

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

—O— 4T-DDD
—O— 4B-DDD
20 a —v— 4B-DDT
0
100
_- Core 11
80 -
6O -
/ + llT—DDD
' —O— llB-DDD
40 ‘ ' + llT-DDT
—V-— llB-DDT
20 o
0
0 8 16 24 32
Incubation Time (Weeks)

Figure 3.7. Comparison of anaerobic dechlorination of DDT in abiotic Core 4
and 11 top (T) and bottom (B) sections during a 32 week incubation period.

48

Abiotic and Biotic Transformations

It was evident that transformations other than biotic were also occurring in the
sediment microcosms. Transformations of DDT to DDD occurred in both autoclaved and
non-autoclaved sediments. The contributions of each process were calculated for the 16
and 32 week samples from each core (Figure 3.8 and Table 3.2). The biotic
transformation contributions were calculated based on the difference in recoveries
between live and autoclaved treatments divided by the DDD recovered in the live
microcosms. The abiotic transformations were calculated by dividing the DDD formed in
the autoclaved sediment microcosms by the DDD formed in the live sediment
microcosms. At 16 and 32- week time points both abiotic and biotic processes
contributed signiﬁcantly to the transformation of DDT to DDD in the Pine River
sediments. These contributions differed with each core and each section of sediment.

At 16 weeks, 43 and 40% of DDD formed in core 4 top and bottom sections was
due to abiotic transformation, and 57 and 60% was formed by biotic transformations,
respectively. In core 11 top and bottom sections 62 and 59% of DDD was formed by
abiotic transformations and 38 and 41% by biotic transformations, respectively. Thus
after 16 weeks of incubation biotic processes had a greater contribution in the formation
of DDD in the core 4 sediments, in contrast to core 11 sediments where there was a
greater abiotic contribution to the transformation of DDT to DDD. At 32 weeks the
abiotic contribution to DDD formation was 50 and 38% for core 4 top and bottom
sections, and 52 and 27% core 11 top and bottom sections, respectively. The biotic
contributions were 50 and 62% for core 4 top and bottom sections, and 48 and 73% for

core 11 top and bottom sections, respectively. T-tests were also used to compare

49

Signiﬁcant differences between live and autoclaved samples at 16 and 32 weeks. At a
95% conﬁdence level, all samples were statistically different when compared to

corresponding live samples at 16 and 32 weeks.

 

 

 

Incubation
Time (wk) Core 4T Core 4B Core 1 1T Core 11B
BI ABI BI ABI BI ABI BI ABI
DDD formation (% of total)
16 57 43 60 40 38 62 41 59
32 50 50 62 38 48 52 73 27

 

Table 3.2. DDD formation due to biotic and abiotic transformations of DDT in Pine
River Sediment microcosms constructed using top (T) and bottom (B) sections of
sediment cores 4 and 11. The biotic (BI) fraction was determined by dividing the
difference in DDD recovered in live treatments by the DDD recovered in autoclaved
treatments and dividing by the DDD recovered in the live treatment (X 100). The abiotic
(AB 1) fraction was determined by dividing the DDD recovered in the autoclaved
sediments by the DDD recovered in the live sediment (X 100).

50

100 e

 

   

 

16 weeks
a 80 e
33
“5
§ 60 e
G
.2
23' 40 z
E
0
LL.
g 20 i I abiotic
I biotic
O L
4T 4B 1 1T 1 1B
Sample core
100 e

g 32 weeks
9 80 -
“-1
O
s: 60 —
C:
.9.
a 40 -
E
m I
8 20 - I abiotic
Q I biotic

O _

4T 4B 1 1T 11B
Sample core

Figure 3.8. DDD recovered from the abiotic and biotic transformations of DDT in Pine
River sediment microcosms constructed using the top (T) and bottom (B) sections of
sediment cores 4 and 11. The biotic fraction was determined by dividing the difference
of the live and autoclaved treatments by the live treatment. The abiotic fraction was
determined by dividing the DDD recovered in the autoclaved treatments by the DDD
recovered in live treatments.

1"C Recovery of DDT

The "C recovery gives the amount of DDX (=DDT+ DDD+ origin) recovered
from the aqueous phase and solvent extractions of the sediment microcosms.
Transformations of DDT (to DDD) occurred in all live and autoclaved sediment
microcosms. At time zero total “C recovery in live sample microcosms ranged from 91-
99%, with an average recovery of 95%, and from 87-95% in autoclaved sediment
microcosms with an average recovery of 92%. After 16 weeks incubation, the recovery
totals were considerably lower with an average recovery of 77% in live microcosms and
82% in the autoclaved microcosms. There was not a considerable decrease thereafter. In
the 32 week samples the average recovery ranged from 72-98% in live samples and 77-
82% in autoclaved microcosms. There was some unexplained loss of radioactivity in
these sample microcosms. This loss could be from reasons such as volatilization or loss

during the extraction process, but there is no deﬁnite explanation for this loss (Table 3.3).

 

% ”c Recovery of DDX (DDT+DDD+Origin)

 

 

 

Incubation Core 4T Core 4B Core llT Core 11B
Time
(wks) Live Auto Live Auto Live Auto Live Auto
96 95 99 93 93 92 91 87
0 (5) (4) (1) (2) (3) (3) (4) (2)
84 81 99 87 78 79 71 79
16 (4) (1) (3) (8) (2) (8) (1) (2)
79 * 99 * 82 * 73 *
24 (1) (1) (7) (0)
80 77 98 82 82 78 72 77
32 (4) (5) (3) (9) (1) (3) (0) (0)

 

Table 3.3. 14C Recovery in Pine River sediment microcosms over 32 weeks. Recoveries
represent the amount of DDX (DDT+DDD+Origin) recovered from aqueous and solvent
extractions of sediment microcosms for top (T) and bottom (B) sections of cores 4 and
11. The standard deviation is reported in parenthesis. *24 week autoclaved samples
were not setup.

52

Transformations of DDD and DDE in sediment microcosms

DDD and DDE were not degraded signiﬁcantly in sediment microcosms after 32
weeks of incubation in sterile and non-sterile microcosms (Table 3.4, Figure 3.9). The
average recovery of 14C-DDD over the 32 -week incubation period in live treatments was
97% of the added DDD with a range of 90-100%. The recovery range for the sterile
microcosms ranged from 88-99% DDD with an average recovery of 97%. The recovery
of DDE in the sediment microcosms was similar to that in the DDD treated sediments.
Over the 32- week incubation period, the l4C-recovery ranged from 87-96% for non-
sterile microcosms and 78-97% for sterile microcosms. These results show that after 32
weeks incubation there were no signiﬁcant transformations of DDD and DDE in the Pine

River sediment microcosms.

53

 

Incubation

 

 

 

 

Time Core 4T Core 4B Core llT Core 11B
(weeks) Live Auto Live Auto Live Auto Live Auto
% Recovered as DDD
0 100 97 99 99 * 99 97 99
(0) (0) (0) (0) * (0) (0) (0)
16 99 98 99 97 99 97 97 94
(0) (0) (0) (1) (0) (1) (1) (0)
24 99 * 95 * 94 * 95 *
(0) * (6) * (2) * (0) *
32 96 99 97 99 97 97 90 88
(0) (0) (2) (1) (0) (1) (0) (9)
% Recovered as DDE
0 96 94 92 86 94 93 96 78
(l) (2) (8) (12) (0) (7) (1) (11)
16 93 97 93 95 95 95 95 95
(2) (1) (4) (3) (2) (2) (0) (5)
24 94 * 89 * 92 * 91 *
(1) * (9) * (1) * (1) *
32 91 95 92 92 87 88 87 96
(1) (5) (1) (5) (7) (6) (7) (2)

 

Table 3.4. Recovery of added MC-DDD and -DDE in Pine River sediment microcosms.
Sample microcosms were established using the top (T) and bottom (B) halves of
sediment cores from two different locations (cores 4 and 11). The standard deviation is
reported in parenthesis. *24 week autoclaved samples were not analyzed

 

Figure 3.9. Autoradiogram of parent compound on TLC plate for transformation of
DDD.

54

Red Cedar Sediment Microcosms

Different sediments and potential sources of dechlorinating inocula were studied
in combination to see if they would support the degradation of DDT, DDD, or DDE. Air-
dried sediments from the Red Cedar River (no known previous exposure to DDT or PCBs
above normal background) were slurried with RAMM and treated with inocula eluted
from various sediments, and in some cases FeSOa amendment. The inocula were from
several locations that had previously shown the ability to dechlorinate PCBs or PBBs
(Morris et. al., 1993). These locations are the Red Cedar River (MI), Hudson River
(NY), Pine River (MI), and Silver Lake (MA). In a study conducted by Zwiemick et. al.,
Fe804 was used as an amendment to stimulate dechlorination. Its stimulatory effects
were attributed to two factors: (1) provision of sulfate as an electron acceptor, which
stimulates growth of sulfate-reducing bacteria responsible for dechlorination activity, and
(2) provision of Fe2+ which precipitates sulﬁde formed during sulfate reduction, hence
reducing sulﬁde toxicity. Accordingly, once sulfate is consumed, an increased number of
sulfate reducers utilize PCBs as an alternate electron acceptor, leading to extensive
dechlorination (Zwiemick, 1999). Adding this amendment may also alleviate inhibition

of PCB dechlorination by heavy metals.

Biologically active microcosms

All treatments supported the dechlorination of DDT to DDD, with the greatest
amount occurring in microcosms with Silver Lake inoculum and the least in the Hudson
River inoculated microcosms. Both Hudson River and Silver Lake sediments are

contaminated with PCBs and have a demonstrated ability to dechlorinate PCBs (Quensen

55

et. al., 1988; Quensen et.al., 1990). DDD formation was directly related to loss of DDT.
The dechlorination of DDT to DDD was the only signiﬁcant transformation observed in
the live microcosms. The DDT recovery range for Red Cedar sediment microcosms was
between 18 and 55%. The greatest amount of DDT recovered (least dechlorination)
occurred in the Hudson River inoculated sediments where 55% of the DDT added was
recovered after 32 weeks of incubation; and the least DDT recovered was observed in the
Silver Lake inoculated sediments with 19% of DDT recovered (Figure 3.10).

Hudson River and Silver Lake inoculated sediments were also treated with Fe804
in an attempt to enhance the dechlorination of DDT. There was about 15% more DDD
recovered in the Silver Lake inoculated microcosms without Fe804 added (73%) than in
Silver Lake microcosms with FeSO4 added (59%). However, in the Hudson River
inoculated microcosms, greater DDD recovery was observed in sediments with FeSO4
added (48%with, 36% without). The greatest amount of DDD formed was observed in
Silver Lake inoculated microcosms where 73% of DDT added was recovered as DDD,
and the least occurred in the Hudson River inoculated sediments with 36% recovered as
DDD. Dechlorination of DDT by Pine River and Silver Lake inocula was greater than
that by indigenous Red Cedar River microorganisms. Zwiemick (1999) observed
enhanced dechlorination of PCBs by Hudson River microorganisms with the addition of
FeSOa. This is consistent with results presented here for DDT as indicated by a decrease
in DDT recovery and an increased amount of DDD formed in the Hudson River
inoculated microcosms amended with FeSO4 as compared to the corresponding non-

FeSO4 amended sediments.

56

 

r- 80-

c:

Q

a

'U

0

a

>

O

8

a:

O

: -RCU
.5 -HR
3* -HRF

PR

 

 

 

 

SIF

 

 

 

% of 14C Recovered as DDD

 

 

 

 

 

RCU HR HRF PR SL SIJ=

Figure 3.10. Recovery of DDT and DDD under strict anaerobic conditions from Red
Cedar Unamended (RCU), Hudson River (HR), Hudson River with Fe804 (HRF), Pine
River (PR), Silver Lake (SL), and Silver Lake with FeSOa (SLF) inocula after 32 weeks.

57

Abiotic microcosms

Transformation of DDT to DDD was observed in both live and autoclaved
sediments, however it was greater in the live sediments than in the autoclaved sediments
for the Red Cedar, Hudson River, and Pine River inoculated sediments as shown in Table
3.5. Overall, the Silver Lake inoculated Red Cedar sediments showed the greatest
transformation of DDT to DDD. For the Silver Lake inoculated sediments there was
slightly greater transformation occurring in the autoclaved sediments. After 32 weeks of
incubation, the recovery of DDT ranged from 15-58% for autoclaved samples. The DDD
recovery range was 30-79% after 32 weeks for autoclaved samples. DDD was formed
from DDT by both abiotic and biotic transformations.

As in the Pine River sediment microcosms, methane production was used as an
indicator of biological activity in sediment microcosms. The live microcosms revealed
headspace gas methane production up to 48% over 32 weeks, indicating that there was
biological activity in the sediments. The methane production in autoclaved sediments

was less than 0.01 % or undetectable.

Abiotic and Biotic transformations

Relative contributions of abiotic and biotic transformations of DDT to DDD were
calculated for the Red Cedar sediment microcosms (Table 3.6, Figure 3.11). Biotic
transformation contributions were calculated based on the difference between the DDD
recovered in live microcosms and DDD recovered in autoclaved microcosms divided by
the DDD formed in the live microcosms. Abiotic transformations were calculated by

dividing the DDD recovered in the autoclaved sediments, by the DDD recovered in the

58

 

% Recovered as DDT
____RCU 1.1.13. ____HRF 1313 .SL _S_L_E
Wks L A L A L A L A L A L A

24 55 45 31 27 18 41
(10) (5) (5) (4) (10) (4)

32 44 58 55 58 42 58 31 32 19 15 34 15
(2) (8) (6) (8) (15) (8) (15) (0) (9) (1) (0) (1)

 

 

 

% Recovered as DDD
_RCU & _HRF ﬂ 5L SE
Wks L A L A L A L A L A L A
24 39 45 58 67 76 55
(9) (6) (5) (4) (9) (3)

32 48 30 36 30 48 30 64 61 73 79 59 79
(3) (5) (5) (5) (14) (5) (12) (0) (8) (1) (0) (1)
Table 3.5. Anaerobic transformation of DDT in sterilized Red Cedar (RC) River
sediment microcosms inoculated with microorganisms eluted from Hudson River (HR),
Pine River (PR) or Silver Lake (SL). Some microcosms were also amended with FeSOa

(F). Uninoculated (U) Red Cedar River sediment was also incubated before (live-L) and
after (auto-A) autoclaving. Standard deviations of the means are given in parenthesis.

live sediments. The results showed that abiotic processes were responsible for the
majority of dechlorination occurring in these treatments but there is substantial biotic
dechlorination occurring. The abiotic contributions were 63, 83, 63, and 95% for Red
Cedar unamended, Hudson River, Hudson River with FeSOa and Pine River,
respectively, and the biotic contributions were 38, 17, 38, and 5% for Red Cedar
unamended, Hudson River, Hudson River with FeSOa and Pine River treatments
respectively. Silver Lake abiotic and biotic contributions were 108 and -8%, and Silver
Lake with FeSOa abiotic and biotic contributions were 134 and -34% respectively. These
values are much greater than those exhibited by the other treatments because the value for

autoclaved DDD recovered was greater than non-autoclaved DDD recovered, thus giving

59

a value greater than 100 using the established calculations. In all of the inoculated Red
Cedar sediment microcosms the majority of DDT transformation occurred by abiotic
processes as opposed to the Pine River sediment microcosms in which the abiotic/ biotic

contributions varied across all sites.

 

% of added DDT recovered as DDD

 

 

Inoculum abiotic biotic
RC 63 38
HR 83 17
HRF 63 38
PR 95 5
SL 108 -8
SLF 134 -34

 

Table 3.6. DDD formation due to biotic and abiotic transformations of DDT at 32 weeks.
Microcosms were set up using uninoculated Red Cedar River (RCU) sediment, as well as
autoclaved RC sediment slurries which were inoculated with microorganisms eluted from
RC, Hudson River (HR), Pine River (PR) or Silver Lake (SL), and in some instances
amended with FeS04 (F). Both autoclaved (abiotic) and non-autoclaved (biotic)
microcosms were then incubated under anaerobic conditions for 32 weeks. The biotic
fraction was determined by dividing the difference between the DDD recovered in live
and DDD recovered in autoclaved treatments by the DDD recovered in the live treatment
(X100). The abiotic fraction was determined by dividing the DDD recovered in the
autoclaved microcosms by the DDD recovered in the live microcosms (X100).

60

 

 

r: 80
.9.
‘a‘
E 60 abiotic
8 40 I biotic
Q
Q
S 20

0

RCU HR HRF PR
Treatments

Figure 3.11. The fraction of DDD recovered resulting from biotic and abiotic
transformations of DDT at 32 weeks. Microcosms were set up using uninoculated Red
Cedar River (RCU) sediment, and autoclaved RC sediment slurries which were
inoculated with microorganisms eluted from RC, Hudson River (HR), Pine River (PR) or
Silver Lake (SL) and in some instances amended with FeSO4 (F). Both autoclaved
(abiotic) and non-autoclaved (biotic) microcosms were then incubated under anaerobic
conditions for 32 weeks. The abiotic fraction was determined by dividing the DDD
recovered in the autoclaved microcosms by the DDD recovered in the live treatments (X
100%). The biotic fraction was determined by dividing the difference of the DDD
recovered in the live microcosms and the DDD recovered in the autoclaved microcosms
by the DDD recovered in the live treatment microcosms (X 100%).

61

Transformations of DDD and DDE in sediment microcosms

After a 32-wcek incubation period, there was no evidence of DDD and DDE
degradation in any of the Red Cedar sediment microcosms. The average recovery of 14C
DDD after the 32 -week incubation period in live treatments was 97% of the added DDD
with a range of 87-99%. In the sterile microcosms, DDD recovery ranged from 96-99%
with an average recovery of 98%. The recovery of DDE in Red Cedar sediment
microcosms was less than that observed for DDD. Over the 32- week incubation period,
the recovery of DDE ranged from 81-92% of added DDE for live microcosms, and 82-
89% for sterile microcosms. These results give evidence that after 32 weeks incubation
Red Cedar sediments did not support the transformations of DDD and DDE in laboratory

microcosms, including those inoculated with known dehalogenating populations.

62

SUMMARY

This study was conducted to examine the fate of DDT, DDD, and DDE in the
Pine River sediments. The transformation of DDT to DDD did occur in the Pine River
sediments under anaerobic conditions by both abiotic and biotic processes. Biotic
transformations accounted for the majority of DDT dechlorination in the core 4
sediments, and abiotic transformations accounted for the majority in the core 11
sediments. Based on recovery totals there appeared to be a greater amount of
dechlorination occurring in the bottom layer sediments in both cores. However,
statistically there was no signiﬁcant difference (P<0.05) between the cores or sections.
Similar amounts of DDD were recovered in core 4 and 11 microcosms despite the greater
amount of co-contaminants present at the location of core 4. Our examination did not
establish any other pathways in the Pine River sediment microcosms. Transformations of
DDD or DDE were not observed.

Sediment slurry systems known to dechlorinate PCBs and/or PBBs were tested
for their ability to transform DDT. These consisted of Red Cedar River sediments, and
autoclaved Red Cedar River sediments inoculated with microorganisms eluted from
Hudson River, Silver Lake and Pine River. All treatments supported the dechlorination
of DDT to DDD by both abiotic and biotic processes, with the majority (63 %-95%)
occurring by abiotic processes. There was no observation of DDD or DDE
transformation. Hudson River and Silver Lake microcosms were amended with Fc804 to
test its ability to stimulate reductive dechlorination of DDT. The FeSOa amendment did

enhance DDT dechlorination in the Hudson River microcosms consistent with its

63

stimulatory effects on PCB dechlorination. However, no further transformations beyond
DDD were observed in these sediment microcosms.

These studies suggest that the predominant transformation of DDT in anaerobic
sediments is dechlorination to DDD, and that these processes occur both biologically and
abiotically. These ﬁndings will be beneﬁcial in establishing the processes and conditions

under which DDX transformations take place in the environment.

REFERENCES

Forba, R.W. US. EPA Pine River Contamination Survey, St. Louis, MI, EPA-330/2-80-
031, 1980.

Forba, R.W. US. EPA Summary of Pine River Reservoir Sediment Sampling Survey, St.
Louis, MI, EPA-330/2-82-001, 1982.

Morris, P.J., Quensen, J.F III, Tiedje J .M., and Boyd, SA. 1993. An assessment of the
Reductive Debromination of Polybrominatcd Biphenyls in the Pine River reservoir.
Environmental Science and Technology. 27: 1580-1586.

Quensen, J .F 111; Tiedje, J .M.; and Boyd, SA. 1988. Reductive dechlorination of
Polychlorinatcd Biphenyls by anaerobic microorganisms from sediments.

Quensen, J .F IH; Boyd S.A.; and Tiedje J.M. 1990. Dechlorination of Four commercial
Polychlorinatcd Biphenyl Mixtures (Aroclors) by Anaerobic Microorganisms from
Sediments. Appl and Environ. Microbiol. 56: 2360-2369.

Quensen, J .F. 111, Mueller, S.A., Jain, KM. 1998. Reductive dechlorination of DDE to
DDMU in marine sediment microcosms. Science. 280: 722-724.

Roberts, MG. 2001. Microbial transformation of DDT in Pine River Sediments. Thesis.
Michigan State University.

U.S.EPA Region 5. 1998. Draft Streamlined Remediation Investigation Report:
Velsicol Chemical Site, St. Louis, MI.

Zwiemick, M.J., Quensen J .F. 111, Boyd, SA. 1999. FeSO4 amendments stimulate
extensive anaerobic PCB dechlorination

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