AN ABSTRACT OF THE DISSERTATION OF
Sarun Tejasen for the degree of Doctor of Philosophy in Civil Engineering presented
on June 27, 2003. Title: Aerobic Biotransformation of Chlorinated Aliphatic
Hydrocarbons by a Benzyl Alcohol Grown Mixed Culture: Cometabolism,
Mechanisms, Kinetics, and Modeling
Redacted for privacy
Abstract approved:
s Semprini
The aerobic transformation of TCE and cis-DCE by a tetrabutoxysilane-grown
microorganism (Vancheeswaran et al., 1999) led to the investigation of novel
substrates, including benzyl alcohol, for promoting cometabolism. The culture grew
on carboxylic.compounds and alcohols, but did not grow on formate, methanol,
methane, propane, butane, ethylene, benzene, toluene, or p-xylene. Cis-DCE
transformation was observed when the culture grew on butyrate, glucose, I -propanol,
1 -butanol, ethanol, benzyl alcohol, and phenol, and effectively transformed TCE, cis-
DCE, and vinyl chloride when grown on phenol or benzyl alcohol.
Several cycles of growth on benzyl alcohol led to increases in ICE
transformation rates and transformation capacities. Products of benzyl alcohol
degradation shifted from benzaldehyde to 2-hydroxy benzyl alcohol (2HBA) during
the several cycles of growth. In resting cells studies, 2HBA production rates were
highly correlated with TCE transformation rates. TCE transformation and 2HBA
production rates doubled when the culture was grown on phenol and rates of TCE
transformation were correlated with 2HBA production rates. Benzyl alcohol- and
phenol-grown cells oxidized toluene to o-cresol, which indicated the similarity
between benzyl alcohol ortho-monooxygenase, phenol hydroxylase, and toluene
ortho-monooxygenase. 2-Butyne and 1 -hexyne (but not acetylene) inhibited benzyl
alcohol- and phenol-grown cells similarly, indicating the same ortho-monooxygenase
was responsible for ICE cometabolism.
Resting cell kinetic studies were performed with cells grown on phenol or
benzyl alcohol. Benzyl alcohol degradation followed a Monod kinetics while phenol
degradation followed a Haldane kinetics. The maximum transformation rates (km) of
TCE, cis-DCE, and VC achieved by phenol-grown cells were about a factor of two
higher than achieved with benzyl alcohol-grown cells, while the half-saturation
constants (Ks) were in a similar range. Transformation capacities (Tc) for TCE, cis-
DCE, and YC were about a factor of two to four higher with phenol-grown cells. The
modeling of TCE, cis-DCE, and VC transformation using independently measured
kmax
and K values matched well with observed data from batch tests. Benzyl alcohol
was shown to be an effective novel substrate for the aerobic cometabolism of TCE,
cis-DCE, and vinyl chloride. Being a non-regulated compound, it might have
applications for in-situ bioremediation.
©Copyright by Sarun Tejasen
June 27, 2003
All Rights Reserved
Aerobic Biotransformation of Chlorinated Aliphatic Hydrocarbons by a Benzyl
Alcohol Grown Mixed Culture: Cometabolism, Mechanisms, Kinetics, and Modeling
by
Sarun Tejasen
A DISSERTATION
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Doctor of Philosophy
Presented June 27, 2003
Commencement June 2004
Doctor of Philosophy dissertation of Sarun Tejasen presented on June 27, 2003.
APPROVED
Redacted for privacy
Major Professor, representing Civil Engineering
Redacted for privacy
Head Department of Civil, Construction, and Environmental Engineering
Redacted for privacy
Dean of Grduate School
I understand that my dissertation will become part of the permanent collection of
Oregon State University libraries. My signature below authorizes release of my
dissertation to any reader upon request.
Redacted for privacy
Sarun Tejasen, Author
ACKNOWLEDGEMENTS
The completion of my doctoral studies would not have been possible without
the help and support of many people in my professional and personal life.
My doctoral studies were financially supported by the Thai Ministry of
University Affairs and Chulalongkom University, Bangkok, Thailand. This research
was supported by a grant from the US Department of Defense sponsored Strategic
Environmental Research and Development Program.
I would like to thank my Ph.D. advisor, Dr. Lewis Semprini, for his help,
encouragement, advice, and financial support over the course of my studies. I also
thank my committee members, Dr. Kenneth J. Williamson, Dr. Daniel J. Arp, Dr. Jack
Istok, and Dr. Thomas McLain, for putting their time, energy, and insights into my
work. I would like to thank Dr. Mohammad Azizian for his help on every problem
with instruments, and Dr. Mark Dolan for his advice on the experimental methods. I
also thank Dr. Young Kim for his advice on kinetic test, and Sanjay Vancheeswaran
for the starting of my research. I thank Adriana Martinez-Prado, Seungho Yu,
Adisorn Tovanabootr, and George Pon for their valuable discussions and friendship.
Lastly, I also would like to thank my wonderful parents and families in
Thailand for their support and encouragement. I thank my Merryfield and Thai
friends, including Chulanee Thianthai, for making my time in Corvallis a pleasurable
and memorable one.
CONTRIBUTION OF AUTHORS
Dr. Lewis Semprini assisted in the concept and writing of each manuscript.
Dr. Mark E. Dolan assisted in the monitoring of microbial communities presented in
Chapter 4. Dr. Daniel J. Arp assisted in the concept and the design of inhibition
experiments presented in Chapter 4. Dr. Kermeth J. Williamson, Dr. Jack Istok, and
Dr. Thomas McLain provided helpful insights and advice.
TABLE OF CONTENTS
CHAPTER 1: Introduction
1
OBJECTIVES
8
REFERENCES
9
CHAPTER 2: Literature Review
14
DEFINITION OF COMETABOLISM
14
CHLORINATED ALIPHATIC HYDROCARBONS
15
1,2-Dichioroethylene
Vinyl Chloride
AEROBIC COMETABOLISM OF TCE, CIS-DCE, AND VC
Methane
Phenol and Toluene
Non-Regulated Substrates
16
17
18
19
20
22
25
PHENOL-GROWN MICROORGANISMS
27
ENZYMES INHIBITORS
28
KINETICS OF AEROBIC COMETABOLISM OF CAHS
30
Michalis-Menten Enzyme Kinetics
Haldane Kinetics
Transformation capacity
Active cells mass
Transformation yield
Kinetic parameters
30
31
31
32
32
32
SUMMARY
36
REFERENCES
37
TABLE OF CONTENTS (CONTINUED)
CHAPTER 3: Aerobic Biotransformation of Chlorinated Ethenes
by a Mixed Culture Grown on a Broad Range of Substrate
47
ABSTRACT
48
INTRODUCTION
49
METHODS
52
RESULTS
57
Cometabolic transformation of TCE and cis-DCE during
growth on TBOS and 1 -butanol
Growth on a broad range of substrates and the cometabolic
transformation of TCE and cis-DCE
Test of cometabolic growth substrates for VC, cis-DCE,
and TCE transformation
57
59
64
DISCUSSION AND CONCLUSION
66
ACKNOWLEDGEMENTS
71
REFERENCES
72
CHAPTER 4: Induction of 2-Hydroxy-Benzyl Alcohol Production and
Trichioroethylene Cometabolism by a Benzyl Alcohol-Grown
77
ABSTRACT
78
INTRODUCTION
79
METHODS
81
TABLE OF CONTENTS (CONTINUED)
RESULTS
Cometabolic transformation of TCE during growth
on benzyl
Transformation TCE and the degradation of benzyl alcohol
by resting cells grown on benzyl alcohol
TCE Induction
Microbial communities monitoring
Benzyl alcohol degradation and TCE Transformation
by resting cells grown on phenol
Toluene oxidation
Inhibition studies
90
90
93
95
96
97
101
102
DISCUSSION AND CONCLUSION
106
ACKNOWLEDGEMENTS
110
REFERENCES
III
CHAPTER 5: Kinetic and Modeling of Vinyl Chloride, cis-Dichioroethylene,
and Trichloroethylene Transformations by an Aerobic Enrichment Grown
on Phenol and Benzyl Alcohol
114
ABSTRACT
115
INTRODUCTION
116
METHODS
118
RESULTS
126
CAHs cometabolism by benzyl alcohol-grown cells
Substrate utilization kinetic
TCE, cis-DCE, and VC transformation kinetic
Transformation capacity
Modeling of CAHs transformation
126
127
129
132
133
TABLE OF CONTENTS (CONTINUED)
DISCUSSION
137
REFERENCES
145
CHAPTER 6: Engineering Significance and Conclusion
150
CONCLUSIONS
150
ENGINEERING SIGNIFICANCE
154
FUTURE WORK
155
REFERENCES
156
BIBLIOGRAPHY
157
APPENDICES
170
LIST OF FIGURES
Figure
Page
2.1 Chemical structure of TCE, cis-DCE, and VC
16
2.2 TCE breakdown mechanism by a methanotrophs-isolate strain 46-1
21
2.3 Summary of toluene and phenol degradation pathways
24
3.1 Comparison of TCE transformation by a culture growing
on TBOS and 1-butanol
58
3.2 Cis-DCE transformation and growth of the enrichment grown
on formate, acetate, propionate, butyrate, and 1-butanol
60
3.3 Transformation of VC, cis-DCE, and TCE, and growth after addition
of butyrate, glucose, and benzyl alcohol
65
4.1 Cometabolic transformation of TCE during the growth on
multiple and single additions of benzyl alcohol
91
4.2 The degradation of benzyl alcohol and the transformation of TCE
by benzyl alcohol-grown cells under resting cells condition
94
4.3 TRFLP analysis of the benzyl alcohol grown culture for
differ growth cycles
97
4.4 The benzyl alcohol degradation and the TCE transformation
by phenol-grown cells under resting cells condition
98
4.5 Correlation of 2HBA production and TCE transformation
by benzyl alcohol-grown cells
99
4.6 The benzyl alcohol degradation and TCE transformation by resting cells
grown on the combination of phenol and benzyl alcohol
101
4.7 Time course of inactivation by 1-hexyne
103
4.8 Observed pathways of benzyl alcohol degradation
103
LIST OF FIGURES (CONTiNUED)
Figure
Page
5.1 Resting cell transformation of CAHs by benzyl alcohol-grown
126
5.2 Degradation kinetic of phenol and benzyl
128
5.3 TCE transformation by resting cells grown on benzyl alcohol and phenol
129
5.4 TCE transformation by resting cells grown on benzyl alcohol
130
5.5 Cis-DCE transformation by resting cells grown on benzyl and phenol
131
5.6 VC transformation by resting cells grown on benzyl alcohol and phenol
132
5.7 Modeling of TCE, cis-DCE, and VC transformation
by phenol-grown resting cells
134
5.8 Modeling of TCE, cis-DCE, and VC transformation
by benzyl alcohol-grown resting cells
135
LIST OF TABLES
Table
Page
2.1 Kinetic parameters of phenol degradation
33
2.2 Comparison of microorganisms and CAlls transformation kinetics
35
3.1 Growth substrates and cometabolic conditions of the batch experiments
55
3.2 Substrates tested for growth and cis-DCE and TCE transformation
62
4.1 Effects of inhibitors on the resting cells activities
103
5.1 Summary of kinetic parameters from the resting cells studies
128
5.2 Ratio of kmax over Tc for CAHs transformation
133
5.3 Experimental conditions for modeling of resting cells CAlls transformation
136
5.4 Comparison of microorganisms and phenol degradation kinetics
139
5.5 Comparison of microorganisms and CABs transformation kinetics
140
LIST OF APPENDICES
Appendix
A: HPLC
171
B: Inhibition of Phenol on Benzyl Alcohol Degradation and 2FIBA
174
C: Haldane Kinetic of TCE Transformation by Benzyl Alcohol-Grown
176
D: Significance of Decay Term in the Transformation
177
LIST OF APPENDIX FIGURES
Figure
Al. Chromatograms benzyl alcohol, 2-, 3-, and 4-hydroxy benzyl alcohol
173
BI. Effects of phenol on benzyl alcohol degradation and 2HBA production
by phenol-grown cells
175
B2. Effects of phenol on benzyl alcohol degradation and 2HBA production
by benzyl alcohol-grown cells
175
Cl. Haldane kinetic of TCE transformation by resting cells
grown on benzyl alcohol
176
DI. Effects of decay term on the modeling of TCE transformation
by benzyl alcohol grown cells
177
LIST OF APPENDIX TABLES
Table
Al. Approximate HPLC Retention time and peak area
of compounds of interest
172
Cl. Kinetic parameters of Haldane model for TCE transformation
by benzyl alcohol-grown cells
176
DEDICATION
I would like to dedicate this dissertation to my parents, Jumsak and Pranat
Tejasen, who are happier and more proud of my success than I am.
Aerobic Biotransformation of Chlorinated Aliphatic Hydrocarbons by a Benzyl
Alcohol Grown Mixed Culture: Cometabolism, Mechanisms, Kinetics, and Modeling
CHAPTER 1: Introduction
Trichioroethylene (TCE) is a suspected human carcinogen that is one of the
most frequently found organic contaminants in groundwater. The Environmental
Protection Agency (EPA) reported finding TCE in over 700 Superfund sites and
estimated that 9-34% of the United States groundwater may be contaminated with
TCE (U.S. EPA, 2003). Upon improper use or leakage, TCE can evaporate into the
air or migrate through the soil and into groundwater. TCE is an organic solvent,
heavier than water, and has a solubility of about 1 gIL, therefore it can penetrate the
groundwater, adsorb into underground sediments, and be a long term contaminant
source in groundwater. Chemical treatment of TCE is often prohibitively expensive.
Physical treatments such as carbon adsorption requires further disposal of the
contaminant and adsorbent, and air stripping releases the contaminant into the
atmosphere, where it may pose alternative environmental and health concerns.
Therefore, biological treatments, which degrade the contaminant without generating
toxic wastes, may be more suitable for cleaning large volumes of contaminated
groundwater.
There are many reports of anaerobic bioremediation of TCE, but the anaerobic
process generally leads to the accumulation of harmful byproducts, such as cisdichloroethylene (cis-DCE) and vinyl chloride (VC) (Gibson and SeweIl, 1992;
2
Semprini, I 997a; Vogel and McCarty, 1985). On the contrary, numerous studies have
demonstrated that aerobic bacteria can transform TCE via cometabolic transformation
to harmless end products, such as chloride ion and carbon dioxide (Arp et aL, 2001;
McCarty, 1997; McCarty and Semprini, 1994; Semprini, I 997b). Cometabolic
transformations are reactions that are catalyzed by existing microbial enzymes and do
not yield carbon or energy to the transforming cells (Alvarez-Cohen and Speitel, 2001;
Horvath, 1972). A growth substrate is therefore required to provide an energy source,
and induce production of the cometabolic enzymes. Growth substrates for these
bacteria include methane (Chang and Alvarez-Cohen, 1995; Oldenhuis et al., 1989;
Wilson and Wilson, 1985), phenol (Folsom et al., 1990; Nelson et al., 1987), toluene
(McCarty et al., I 998a; Nelson et al., 1987; Wackett and Gibson, 1988), ammonia
(Arciero et al., 1989), ethylene (Ensign et al., 1992), isoprene (Ewers et aL, 1991),
propane (Wackett et al., 1989), propylene (Ensign et al., 1992), and butane (Kim et al.,
2000).
Among cometabolic substrates, toluene and phenol are well known as growth
substrates supporting the cometabolism of TCE. In-situ microorganisms grown on
aromatic substrates, such as phenol and toluene, have been reported to cometabolize
TCE better than those grown on methane (Bielefeldt et al., 1995; Hopkins et al.,
1993). In a demonstration test of in-situ cometabolism conducted at Edward Air Force
Base (CA), toluene was chosen as the cometabolic growth substrate over phenol, since
it has a lower odor threshold than phenol and as a liquid it could be easily handled.
Phenol also has the potential of forming chlorinated phenols upon chlorination
ci
(McCarty et al., 1998a). The U.S. Environmental Protection Agency (EPA), however,
reported that inhalation of toluene can affect the human nervous system, the kidneys,
the liver, and the heart. Toluene is listed as a groundwater contaminant with a
recommended maximum contaminant level (MCL) in drinking water of I mg/I (U.S.
EPA, 2001). Thus, as a regulated compound, obtaining regulatory approval for in-situ
use may prove difficult in some cases.
A number of studies have focused on finding non-regulated substrates for use
in the in-situ treatment of TCE and other CABs. Gao and Skeen (1999) reported cisDCE transformation in an aerobic groundwater/soil microcosm fed glucose. Recently
a microorganism was isolated that can be aerobically grown on cis-DCE as a primary
substrate (Coleman et al., 2002). These results indicate that aerobic microorganisms
that grow on other substrates might be able to gain energy from cis-DCE
transformation. Glucose was also evaluated as a growth substrate for Xanthobacter
strain Py2 (Ensign, 1996). Upon the induction with propylene, the glucose-grown
strain Py2 expressed alkene monooxygenase which is a well-known enzyme
responsible for TCE transformation (Ensign, 1996). Fan and Scow (1993) reported
moderate TCE transformation in soil microcosms when methanol was added.
Fructose was used in combination with phenol and dichlorophenoxyacetic acid to
maintain the TCE transformation activity of A icaligenes eutrophus JMP 134 (Muller
and Babel, 1995, 1 996b). A phenol-grown Rastonia eutropha also exhibited TCE
transformation when grown on sodium citrate (Ayoubi and Harker, 1998). Also, there
are some reports of using genetically engineered microorganisms (GEMs) grown on
non-regulated substrates to transform TCE. Genetically engineered Tn5- induced phi
mutant from the strain A. eutrophus JMP 134 grew on ethanol and was capable of TCE
transformation (Kim et al., 1996). Genetically altered Burkholderia cepacia strain
PR1301
was capable of TCE transformation when grown on lactate; however, with
long-term stimulation TCE transformation ability was lost (McCarty Ct al., 1998b;
Munakata-Marr et al., 1997; Munakata-Marr et al., 1996).
In 1999, Vancheeswaran et al. reported an enrichment culture having TCE and
cis-DCE transformation ability when grown on tetrabutoxysilane (TBOS). This
culture rapidly hydrolyzed TBOS to I -butanol, which was used as a growth substrate.
The formation of cis-DCE epoxide was observed, indicating cometabolic
transformation was occurring (Vancheeswaran et al., 1999).
This dissertation study started from surveying the cometabolic potential of the
enrichment obtained by Vancheeswaran et al. In Chapter 3, the TBOS enrichment
culture was tested for its ability to grow on a broad range of substrates and
cometabolize cis-DCE and TCE. Transformation yield (Tv) was used to assess
cometabolic cis-DCE and ICE transformation upon growth on these substrates. T
represents the mass of chlorinated aliphatic hydrocarbon (CAH) transformed per unit
mass of primary substrate consumed (McCarty, 1997). Substrates tested included
silicon-based organic compounds (TBOS, tetrapropoxysilane (TPOS), and
tetraphenoxysilane), phenolic compounds (phenyl acetate, p-cresol, and phenol),
alcohols (methanol, ethanol, 1 -propanol, I -butanol, 4-methyl-benzyl alcohol, 3-buten2-ol, 3-buten-1 -ol, 2-buten-1 -ol, and benzyl alcohol), organic acids (formate, acetate,
propionate, butyrate, and benzoate), aromatic compounds (benzene, toluene, and pxylene), saturated and unsaturated hydrocarbons (methane, propane, butane, and
ethylene), acetone, glucose, and methyl-tert-butyl-ether (MTBE).
Among these substrates, benzyl alcohol was found to be an effective substrate
to cometabolize TCE, cis-DCE, and VC. Benzyl alcohol is a non-regulated
compound, is non toxic, and is commonly used as food flavoring agent (Mallinckrodt
Baker Inc., 2000). It is also a liquid that can be easily handled like toluene. Thus,
benzyl alcohol may be a promising substrate for TCE cometabolism at contamination
sites.
In Chapter 4, TCE cometabolism by a benzyl alcohol-grown culture was
investigated. The culture status was monitored using molecular methods during
successive growth on benzyl alcohol. The parameters used to compare the
cometabolic transformation of TCE were the transformation rates, transformation
capacity
(Ta),
and transformation yield (Tv). Transformation capacity
(Ta) defines the
mass of TCE that can be transformed by a given mass of resting cells, while the
transformation yield (Tv) gives the maximum amount of TCE that can be transformed
per mass of growth substrate utilized (Arp et al., 2001).
Successive growth on benzyl alcohol also led to changes in the benzyl alcohol
degradation pathway. Evidence of changes in the utilization pathway upon substrate
induction have been reported.
Alcaligenes eutrophus
strain JMP 134 when degrading
phenol was reported to change from ortho- to meta-cleavage pathway as growth rate
increased (Muller and Babel, I 996a). Different enzymes can also be induced when
bacteria are exposed to different compounds.
Xanthobacter
strain Py2 grew on
glucose without evidence of alkene monooxygenase or epoxidase, and was able to
produce both enzymes upon the introduction of propylene (Ensign, 1996).
Chlorinated compounds such as TCE and cis-DCE have also been reported to induce
both toluene degradation and TCE transformation (Leahy et al., 1996; McClay et al.,
1995; Shingleton et al., 1998). Successive growth on benzyl alcohol and the
possibility of TCE induction were evaluated to determine changes in T, TCE
transformation rate, benzyl alcohol degradation rate, and the production rate of
byproduct from benzyl alcohol degradation. Correlations of changes in TCE
transformation with product production were evaluated. Since the mixed culture does
not grow on toluene, phenol was compared to benzyl alcohol as a growth substrate.
Toluene, was oxidized by both benzyl alcohol- and phenol-grown cells, and was used
to investigate byproducts and enzymes involved.
Acetylene has been reported as a monooxygenase inhibitor and has been used
to indicate the involvement of a monooxygenase (Hamamura et al., 1997; Hyman and
Wood, 1985; Verce et al., 2000; Vlieg et al., 1996). Yeager et al. (1999) reported that
acetylene, however; is not an effective enzymes inhibitor for aromatic substrates
utilization. They found that the more effective inhibitors are the longer-chain alkynes,
such as butynes or hexynes (Yeager et al., 1999). In chapter 4, inhibitors, including
acetylene, 2-butyne, and 1 -hexyne, were studied and their effects on benzyl alcohol
degradation, toluene oxidation, TCE transformation by both benzyl alcohol- and
7
phenol-grown cells were compared. The potential of using benzyl alcohol as a phenol
substitute in the aerobic cometabolism of TCE was also investigated.
In Chapter 5, a benzyl alcohol-grown culture was tested for cometabolic
potential of other CAHs. These CABs included chlorinated etbanes, such as
chloroform (CF), 1,1 -dichloroethane (DCA), and 1,1,1 -trichloroethane (TCA), and
chlorinated ethenes, such as TCE, cis-DCE, 1,1 -dichloroethylene (1,1 -DCE), and VC.
This study is the first, to our knowledge, to report all kinetic values for TCE, cis-DCE,
and VC by mixed culture grown on aromatic substrates such as phenol and benzyl
alcohol. The resting cells kinetic studies were performed on the benzyl alcohol
degradation and the transformation of TCE, cis-DCE, and VC. The kinetic parameters
include growth yield (Y), maximum degradation rate
(Ks), and transformation capacity
(km),
half-saturation constant
(Ta). To compare the effectiveness of benzyl
alcohol, phenol was also studied as a growth substrate since this culture does not grow
on toluene. Kinetics of phenol degradation and the transformation of TCE, cis-DCE,
and VC by phenol grown cells are also reported. Modeling of resting cells
transformation of TCE, cis-DCE, and VC with the achieved kinetic parameters from
both benzyl alcohol and phenol-grown cells was also performed.
8
OBJECTIVES
In summary, the following objectives were developed for this dissertation:
I. To find alternative substrates for the aerobic cometabolism of TCE, cis-DCE,
andVC.
2. To study the mechanism and pathway of benzyl alcohol degradation that
results in an effective cometabolic transformation of TCE.
3. To evaluate the effectiveness of benzyl alcohol as growth substrate for TCE,
cis-DCE, and VC transformations, including the determination of kinetic
parameters and comparison to values achieved by phenol-grown culture.
4. To determine the ability of kinetic parameters to predict results from batch
reactor experiments of TCE, cis-DCE, and VC cometabolic transformations by
resting cells grown on benzyl alcohol or phenol.
REFERENCES
Alvarez-Cohen, L., and Speitel, G. E. J. (2001). Kinetics of Aerobic Cometabolism of
Chlorinated Solvents. Biodegradation 12(2), 105-126.
Arciero, D., Varmelli, T., Logan, M., and Hooper, A. B. (1989). Degradation of
Trichloroethylene by the Ammonia-Oxidizing Bacterium Nitrosomonas
Europaea. Biochem. Biophys. Res. Comm. 159(2), 640-643.
Arp, D. 3., Yeager, C. M., and I-lyman, M. R. (2001). Molecular and Cellular
Fundamentals of Aerobic Cometabolism of Trichioroethylene. Biodegradation
12(2), 81-103.
Ayoubi, P. J., and Harker, A. R. (1998). Whole-Cell Kinetics of Trichioroethylene
Degradation by Phenol Hydroxylase in a Rastonia Eutropha Jmp 134
Derivative. App!. Environ. Microbiol. 64(11), 4353-4356.
Bielefeldt, A. R., Stensel, H. D., and Strand, S. E. (1995). Cometabolic Degradation of
Tee and Dee without Intermediate Toxicity. Journal of Environmental
Engineering 121(11), 791-797.
Chang, H.-L., and Alvarez-Cohen, L. (1995). Transformation Capacities of
Chlorinated Organics by Mixed Cultures Enriched on Methane, Propane,
Toluene, or Phenol. Biotechnol. Bioeng. 45, 440-449.
Coleman, N. V., Mattes, T. E., Gossett, 3. M., and Spain, J. C. (2002). Biodegradation
of Cis-Dichloroethene as the Sole Carbon Source by a B-Proteobacterium.
AppI. Environ. Microbiol. 68(6), 2726-2730.
Ensign, S. A. (1996). Aliphatic and Chlorinated Alkenes and Epoxides as Inducers of
Alkene Monooxygenase and Epoxidase Activities in Xanthobacter Strain Py2.
Appi. Environ. Microbiol. 62(1), 61-66.
Ensign, S. A., Hyman, M. R., and Arp, D. 3. (1992). Cometabolic Degradation of
Chlorinated Alkenes by Alkene Monooxygenase in a Propylene-Grown
Xanthobacter Strain. App!. Environ. Microbiol. 58(9), 3038-3 046.
Ewers, 3., Clemens, W., and Knackmuss, H. J., Eds. (1991). Biodegradation of
Chioroethenes Using Isoprene as Co-Substrate. Vol. 1. International
Symposium on Environmental Biotechnology. Ostend, Belgium: Royal
Flemish Society of Engineers.
Folsom, B. R., Chapman, P. J., and Pritchard, P. H. (1990). Phenol and
Trichioroethylene Degradation by Pseudomonas Cepacia G4: Kinetics and
Interactions between Substrates. App!. Environ. Microbiol. 56(5), 1279-1285.
10
Gibson, S. A., and Sewell, G. W. (1992). Stimulation of Reductive Dechlorination of
Tetrachioroethene in Anaerobic Aquifer Microcosms by Addition of ShortChain Organic Acids or Alcohols. App!. Environ. Microbiol. 58(4), 1392-1393.
Hamamura, N., Page, C., Long, T., Semprini, L., and Arp, D. J. (1997). Chloroform
Cometabolism by Butane-Grown Cf8, Pseudomonas Butanovora, and
Mycobacterium Vaccae Job5 and Methane-Grown Methylosinus
Trichosporium Ob3b. App!. Environ. Microbiol. 63(9), 3607-3613.
Hopkins, G. D., Semprini, L., and McCarty, P. L. (1993). Microcosm and in Situ Field
Studies of Enhanced Biotransformation of Trichloroethylene by PhenolUtilizing Microorganisms. App!. Environ. Microbiol. 59(7), 2277-2285.
Horvath, R. (1972). Microbial Co-Metabolism and the Degradation of Organic
Compounds in Nature. Bacteriol. Rev. 36, 146-155.
Hyman, M. R., and Wood, P. M. (1985). Suicidal Inactiation and Labelling of
Ammonia Mono-Oxygenase by Acetylene. Biochem. J 227, 719-725.
Kim, Y., Arp, D. J., and Semprini, L. (2000). Chlorinated Solvent Cometabolism by
Butane-Grown Mixed Culture. Journal of Environmental Engineering
October, 934-942.
Kim, Y., Ayoubi, P., and Harker, A. R. (1996). Constitutive Expression of the Cloned
Phenol Hydroxylase Gene(S) from Alcaligenes Eutrophus Jmpl 34 and
Concomitant Trichiorethylene Oxidation. App!. Environ. Microbiol. 62(9),
3227-3233.
Leahy, J., Byrne, A., and Olsen, R. (1996). Comparison of Factors Influencing
Trichloroethylene Degradation by Toluene-Oxidizing Bacteria. App!. Environ.
Microbiol. 62(3), 825-833.
Mallinckrodt Baker Inc. (2000). Material Safety Data Sheet (Msds): Benzyl Alcohol.,
www.jtbaker.conilrnsds/bl 885.htm.
McCarty, P. L. (1997). Aerobic Cometabolism of Chlorinated Aliphatic
Hydrocarbons. In "Subsurface Restoration" (C. H. Ward, J. A. Cherry, and M.
R. Scaif, Eds.), pp. 373-395. Ann Arbor Press, Inc., Chelsea, Michigan.
McCarty, P. L., Goltz, M. N., Hopkins, G. D., Dolan, M. E., Allan, J. P., Kawakami,
B. T., and Carrothers, T. J. (1998a). Full-Scale Evaluation of in Situ
Cometabolic Degradation of Trichloroethylene in Groundwater through
Toluene Injection. Environ. Sci. Technol. 32(1), 88-100.
McCarty, P. L., Hopkins, G. D., Munakata-Marr, J., Matheson, V. G., Dolan, M. E.,
Dion, L. B., Shields, M., Forney, L. J., and Tiedje, J. M. (1998b).
11
Bioaugmentation with Burkholderia Cepacia Pr1301 for in Situ Bioremediation
of Trichioroethylene Contaminated Groundwater. EPA (April), 1-1 1.
McCarty, P. L., and Semprini, L. (1994). Ground-Water Treatment for Chlorinated
Solvents. In "Handbook of Bioremediation." (J. E. Matthews, Ed.), pp. 87-116.
Lewis Publishers, Ann Arbor.
McClay, K., Streger, S. H., and Steffan, R. J. (1995). Induction of Toluene Oxidation
Activity in Pseudomonas Mendocina Krl and Pseudomonas Sp. Strain Envpc5
by Chlorinated Solvents and Alkanes. Appi. Environ. Microbiol. 61(9), 34793481.
Muller, R. H., and Babel, W. (1995). Determination of the Ks Values During the
Growth of Alcaligenes Eutrophus on Phenol, 2,4-Dichiorophenoxyacetic Acid
and Fructose. Acta Biotechnol. 15(4), 347-3 53.
Muller, R. H., and Babel, W. (1 996a). Growth Rate-Dependent Expression of PhenolAssimilation Pathways in Alcaligenes Eutrophus Jmp 134 - the Influence of
Formate as as Auxiliary Energy Source on Phenol Conversion Characteristics.
App!. Micro biol. Biotechnol. 46(2), 156-162.
Muller, R. H., and Babel, W. (1996b). Measurement of Growth at Very Low Rates
(M>0), an Approach to Study the Energy Requirement for the Surial of
Alcaligenes Eutrophus Jmp134. Appi. Environ. Microbiol. 62(1), 147-151.
Munakata-Marr, J., Matheson, V. G., Fomey, L. J., Tiedje, J. M., and McCarty, P. L.
(1997). Long-Term Biodegradation of Trichioroethylene Influenced by
Bioaugmentation and Dissolved Oxygen in Aquifer Microcosms. Environ. Sci.
Technol. 3 1(3), 786-791.
Munakata-Marr, J., McCarty, P. L., Shields, M. S., Reagin, M., and Francesconi, S. C.
(1996). Enhancement of Trichloroethylene Degradation in Aquifer
Microcosms Bioaugmented with Wild Type and Genetically Altered
Burkholderia (Pseudomonas) Cepacia G4 and Pri. Environ. Sci. Technol.
30(6), 2045-2052.
Nelson, M. J. K., Montgomery, 5. 0., Mahaffey, w. R., and Pritchard, P. H. (1987).
Biode gradation of Trichloroethylene and Involvement of an Aromatic
Biodegradative Pathway. App!. Environ. Microbiol. 53(5), 949-954.
Oldenhuis, R., Vink, R. L. J. M., Janssen, D. B., and Witholt, B. (1989). Degradation
of Chlorinated Aliphatic Hydrocarbons by Methylosinus Trichosporium Ob3b
Expressing Soluble Methane Monooxygenase. App!. Environ. Microbiol. 55,
2819-2826.
12
Semprini, L. (1 997a). In Situ Transformation of Halogenated Aliphatic Compounds
under Anaerobic Conditions. In "Subsurface Restoration" (C. H. Ward, J. A.
Cherry, and M. R. Scaif, Eds.), pp. 429-450. Ann Arbor Press, Inc., Chelsea,
Michigan.
Semprini, L. (1 997b). Strategies for the Aerobic Co-Metabolism of Chlorinated
Solvents. Curr. Opin. Biotechnol. 8, 296-3 08.
Shingleton, J. t., Applegate, B. M., Nagel, A. C., Bienkowski, P. R., and Sayler, G. S.
(1998). Induction of the TodOperon by Trichioroethylene in Pseudomonas
Putida Tva8. Appi. Environ. Microbiol. 64(12), 5049-5052.
U.S. EPA. (2001). National Primary Drinking Water Regulations. Office of water,
U.S. Environmental Protection Agency,
http://www.bren.ucsb.edu/fac staff/fac/keller/courses/esm223/MCL.html.
U.S. EPA. (2003). Consumer Factsheet On: Trichioroethylene. U.S. Environmental
Protection Agency, http://www.epa.gov/safewater/dwhlc-voc/trichlor.html.
Vancheeswaran, S., Halden, R. U., Williamson, K. J., James D. Ingle, J., and
Semprini, L. (1999). Abiotic and Biological Transformation of
Tetraalkoxysilanes and Trichioroethene/Cis- I ,2-Dichloroethene Cometabolism
Driven by Tetrabutoxysilane-Degrading Microorganisms. Environ Sci.
Technol. 33(7), 1077-1085.
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2000). Characterization of an Isolate
That Uses Vinyl Chloride as a Growth Substrate under Aerobic Conditions.
App!. Environ. Microbiol. 66(8), 3535-3542.
Vlieg, J. E. T. v. H., Koning, W. d., and Janssen, D. B. (1996). Transformation
Kinetics of Chlorinated Ethenes by Methylosinus Trichosporium Ob3b and
Detection of Unstable Epoxides by on-Line Gas Chromatography. Appi.
Environ. Microbiol. 62(9), 3304-3312.
Vogel, T. M., and McCarty, P. L. (1985). Biotransformation of Tetrachloroethylene to
Trichioroethylene, Dichloroethylene, Vinyl Chloride and Carbon Dioxide
under Methanogenic Conditions. App!. Environ. Microbiol. 49(5), 1080-1083.
Wackett, L. P., Brusseau, G. A., Householder, S. R., and Hanson, R. 5. (1989). Survey
of Microbial Oxygenases Trichloroethylene Degradation by PropaneOxidizing Bacteria. Appi. Environ. Microbiol. 55, 2960-2964.
Wackett, L. P., and Gibson, D. T. (1988). Degradation of Trichioroethylene by
Toluene Dioxygenase in Whole-Cell Studies with Pseudomonas Putida Fl.
App!. Environ. Microbiol. 54(7), 1703-1708.
13
Wilson, J. T., and Wilson, B. II. (1985). Biotransformation of Trichioroethylene. Appi.
Environ. Microbiol. 49(1), 242-243.
Yeager, C. M., Bottomley, P. J., Arp, D. J., and Flyman, M. R. (1999). Inactivation of
Toluene 2-Monooxygenase in Burkholderia Cepacia G4 by Alkynes. Appi.
Environ. Microbiol. 65(2), 632-639.
14
CHAPTER 2: Literature Review
DEFINITIONS OF COMETABOLISM
The study of cometabolism process started since 1950s, and has focused on the
microbial degradation of a wide range of industrial chemicals including aromatics,
chlorinated organics, pesticides, and petroleum hydrocarbons (Arp et al., 2001).
Definitions of cometabolism include:
McCarty (1997): "Cometabolism is the fortuitous transformation of an organic
compound by enzymes or other biomolecules produced by organisms for other
purposes" (McCarty, 1997).
Semprini (1997): "Co-metabolism describes the metabolism of a substrate not
required for growth in which no apparent benefit is accrued by the metabolizing
organism" (Semprini, 1997).
Alvarez-Cohen and Speitel (2001): "Cometabolic transformations are reactions
that are catalyzed by existing microbial enzymes and that yield no carbon or energy
benefits to the transforming cells" (Alvarez-Cohen and Speitel, 2001).
A general concept is that microorganisms can degrade compounds that have no
benefit to them by using energy acquired from metabolism of growth substrate. To
illustrate this concept, the cometabolism of trichioroethene (ICE) by methane-grown
microorganism (McCarty and Semprini, 1994) is shown below.
15
Methane Oxidation:
CH4
>
ICOOH
CH3 OH
NADH, 02
Synthesis
_co2
NADH
NADH
TCE Transformation:
TCE epoxide
TCE
/0\
CC12 = CHC1
>C12CCHC1
>
>CO2,C1,H70
NADH, 02
To utilize methane as substrate, methanotrophs (methane-grown
microorganisms) use methane monooxygenase (MMO) to catalyze the oxidation of
methane to methanol, which requires energy in the form of NADH. MMO also
transforms TCE into TCE epoxide, which is unstable and abiotically transformed
rapidly. In this process methane metabolism results in cell synthesis and MMO
production which catalyzes TCE transformation.
CHLORINATED ALIPHATIC HYDROCARBONS (CAHs)
The CABs discussed in this dissertation are TCE and its anaerobic byproducts
including I ,2-dichloroethene (1 ,2-DCE) and vinyl chloride (VC). The chemical
structures of these compounds are shown in Figure 2.1.
16
TCE
cis-1,2-DCE
H
CI
c=C
H
/
/
/
C=C
H
/
CI
C
VC
\
C
H
H
/
c=C\
/
H
C
Figure 2.1 Chemical structure of TCE, cis-DCE, and VC.
Trichioroethene (TCE)
TCE is a man-made, clear liquid used mainly as a solvent to remove oils and
grease from metal during manufacture or maintenance and as an ingredient in
adhesives, paint removers, typewriter correction fluids, and spot remover (U.S. EPA,
2003b). Because of the variety of past uses and disposal practices, it is also one of the
most commonly found contaminants in groundwater. The Environmental Protection
Agency (EPA) reported finding TCE in over 700 Superfund sites and estimated that 934% of the United States groundwater may be contaminated with TCE (U.S. EPA,
2003b). Upon improper use or leakage, TCE can evaporate into the air or migrate
through the soil and into groundwater. TCE is an organic, heavier than water, and has
a solubility of about 1 gIL, therefore it can penetrate the groundwater, adsorb into
underground sediments, and be a long term contaminant source in groundwater.
TCE exposure is associated with several adverse health effects, including
neurotoxicity, immunotoxicity, developmental toxicity, liver toxicity, kidney toxicity,
endocrine effects, and several forms of cancer (National Center for Environmental
Assessment, 2001). The U.S. EPA has set a maximum contaminant level (MCL) for
17
TCE in drinking water at 5 tg!L and a maximum contaminant level goal (MCLG) at
zero (U.S. EPA, 2003b). With its wide distribution and toxicity, TCE has received a
great deal of attention in the field of bioremediation.
There have not been any reports of TCE as a microbial growth substrate.
However, TCE can be biologically transformed by both aerobic cometabolism and
anaerobic halorespiration. Under anaerobic halorespiration, chloride atoms in TCE
are replaced with hydrogen (reductive dechlorination) and TCE is transformed into
I ,2-dichloroethylene (1 ,2-DCE), vinyl chloride (VC), and ethene, accordingly. But
the rates of anaerobic transformation are much greater for TCE than for DCE or VC,
so that DCE and VC tend to persist longer in the environment (McCarty and Semprini,
1994). VC is a carcinogenic compound with a recommended MCL in drinking water
of 2 tg!L and a MCLG of zero. Thus one concern of anaerobic treatment is the
creation of VC. In contrast to anaerobic halorespiration, numerous studies have
demonstrated that TCE can be aerobically transformed via cometabolic oxidation to
harmless end products, such as chloride ion and carbon dioxide (McCarty, 1997;
McCarty and Semprini, 1994).
1 ,2-Dichloroethene (1 ,2-DCE)
1 ,2-DCE is an odorless organic liquid that normally is used as a solvent for
waxes and resins; in the extraction of rubbers; as a refrigerant; in the manufacture of
pharmaceuticals and artificial pearls; in the extraction of oils and fats from fish and
meat; and in making other organics (U.S. EPA, 2003a). It has two forms, "cis" and
18
"trans". Cis-DCE is often observed as the main isomer product from the anaerobic
transformation of TCE and perchloroethylene (PCE) (Vogel and McCarty, 1985). The
EPA has set an MCLG for cis-DCE and trans-DCE in drinking water at 70 and 100
ig!L, respectively. Exposure to I ,2-DCE above MCL can cause central nervous
system depression and damage in liver, circulatory, and nervous system. Most of the
aerobic microorganisms that cometabolize TCE can also cometabolize cis-DCE. In
addition, cis-DCE has recently been reported as a growth substrate for aerobic
microcosms (Bradley and Chapelle, 2000; Coleman et al., 2002a).
Vinyl Chloride (VC)
VC is a colorless organic gas used in the manufacture of numerous products in
building and construction, automotive industry, electrical wire insulation and cables,
piping, industrial and household equipment, medical supplies, and is depended upon
heavily by the rubber, paper, and glass industries (U.S. EPA, 2003c). Its major release
is as emissions and in wastewater at polyvinyl chloride (PVC) plastic production and
manufacturing facilities. It is also a main byproduct of anaerobic transformation of
TCE and PCE. VC is a known carcinogen and a long-term exposure to VC above
MCL can cause cancer in the liver and nervous system. The EPA has set an MCLG of
VC in a drinking water at zero and an MCL at 2 ig/L due to the detection ability and
treatment technologies (U.S. EPA, 2003c). VC has much less potential for anaerobic
bioremediation than cis-DCE or TCE. It, however, is readily biodegradable under
aerobic conditions. Growth substrates yielding VC cometabolism include ethene,
19
ethane, methane, propane, propylene, isopropene, and ammonia (Verce et aL, 2000).
VC also has been reported as a growth substrate for some microorganisms. (Coleman
et al., 2002b; Hartmans and Bont, 1992; Verce et at., 2000, 2001)
AEROBIC COMETABOLISM OF TCE, 1,2-DCE, AND VC
Reported substrates for microorganisms that have cometabolism potential are:
methane (Chang and Alvarez-Cohen, 1996; Oldenhuis et al., 1991; Semprini et al.,
1990), phenol (Ayoubi and Harker, 1998; Folsom et al., 1990; Hopkins et at., 1993),
toluene (Nelson et al., 1987; Shields et al., 1989; Wackett and Gibson, 1988),
ammonia (Arciero et al., 1989), butane (Kim et aL, 2000), cresol (Folsom et al., 1990;
Nelson et al., 1988; Wackett and Gibson, 1988), dichlorophenoxyacetic acid (Barker
and Kim, 1990), ethylene (Ensign et al., 1992), isoprene (Ewers et al., 1991),
isopropylbenzene (Dabrock et al., 1992), propane (Tovanabootr and Semprini, 1998;
Wackett et at., I 989), and propene (Ensign et al., 1992). Among these substrates,
methane is the most studied substrate with a broad range of TCE transformation
activities reported. In-situ microorganisms grown on aromatic substrates, such as
toluene and phenol have been reported to cometabolize TCE better than those grown
on methane (Bielefeldt et at., 1995; Hopkins et al., 1993). Phenol was also used as
growth substrate for the in-situ bioaugmentation for the removal of TCE, cis-DCE and
VC (Steffan et al., 1999). In a demonstration test of in-situ cometabolism conducted
at Edward Air Force Base (CA), toluene was chosen as the cometabolic growth
substrate over phenol, since it has a lower odor threshold than phenol and as a liquid it
20
could be easily handled and phenol also has the potential of forming chlorinated
phenols upon chlorination (McCarty et al., 1998a). However, toluene is also a
common groundwater contaminant with a recommended maximum contaminant level
(MCL) in drinking water of I mg/i (U.S. EPA, 2001). Thus, as a regulated compound,
obtaining regulatory approval for in-situ use may prove difficult in some cases.
This dissertation will focus on the transformation of CAlls by microorganisms
grown on aromatic substrates, such as phenol and toluene. However, since methane is
the most studied substrate, a review of CAlls cometabolism by rnethanotrophs is also
included.
Methane
In 1985, Wilson and Wilson reported that TCE could be aerobically
transformed to carbon dioxide by a microcosm fed with natural gas (Wilson and
Wilson, 1985). Fogel et al. (1986) confirmed that a methane-grown mixed culture was
not only responsible for the biodegradation of TCE, but it also could transform VC,
cis-DCE, and trans-DCE (Fogel et al., 1986). Little et al. (1988) isolated a pure
culture of methanotrophs capable of TCE transformation and suggested that TCE was
transformed by methane monooxygenase (MMO) into TCE epoxide, which broke
down spontaneously to dichioroacetic acid, glyoxylic acid, formate, and carbon
monoxide, before they were converted to carbon dioxide (Figure 2.2) (Little et al.,
1988). Oldenhuis et al. (1989) and Tsien et al. (1989) found a well-known
Methylosinus trichosporium
strain OB3b, which they described as containing a soluble
21
H
CI
CC
/
/
1MMO
C
7
\/\/
/
C __C
C
ICE EPDXIDE
C\
'.... \ INTERMEDIATE
H20
H
CCH
/
TCEDIOL
C
CC--H
/
OH
C
c
/
OH
OH
j
+ c=o
H"
FORMATE
C
CC
Cl/
H
CARBON
MONOXIDE
0
OH
/
CC
H
/
7,
0
OH
DICHLOROACETIC
ACID
0
GLYOXYLIC
ACID
CO2
Figure 2.2. TCE breakdown pathway by a methanotroph-isolate strain 46-1
(adapted from Little et al., 1988).
22
methane monooxygenase (sMMO) that could transform TCE, VC, cisDCE, transDCE, and 1,1-DCE (Oldenhuis et al., 1989; Tsien et al., 1989). They also suggested
that TCE or TCE epoxide might be toxic to the microorganism. DiSpirito et aL (1992)
reported that M tric/wsporium OB3b could express a particulate methane
monooxygenase (pMMO) which could transform TCE at a lower rate than sMMO
(DiSpirito et al., 1992). pMMO was later shown to be capable of transforming transDCE, cis-DCE, and VC, although the transformation rates were much lower than
sMMO (van Hylckama Vlieg et al., 1996). The disadvantage of sMMO is that it is
found in a small subset of methanotrophs and is expressed only under conditions of Cu
limitation, while pMMO is found in all methanotrophs and is expressed under
conditions of Cu sufficiency (Arp et al., 2001).
Phenol and Toluene
In 1986, Nelson et al. reported an isolate, identified later as Pseudomonas
cepacia G4 (Shields et al., 1991), which was capable of transforming TCE into carbon
dioxide when it was grown on a specific water sample (Nelson et al., 1986). They
found phenol was the key substrate, and that toluene, o-cresol, or m-cresol, but not p.
cresol could replace phenol as a growth substrate for the strain G4 to cometabolize
TCE (Nelson et al., 1987). They suggested that phenol and toluene induced the same
aromatic degradation pathway through catechol-2,3-dioxygenase and cleaved the
aromatic ring by meta fission (Nelson et al., 1987). The strain P. cepacia G4 was later
found to catabolize toluene by successive monooxygenations at the ortho and then
23
meta positions (Shields et aL, 1989). In 1988, the same research group reported two
strains of Pseudomonas putida from natural environments containing toluene
dioxygenase capable of transforming TCE when grown on phenol or toluene (Nelson
et al., 1988). They also observed that a P. putida strain mt-2 (pWWO), which
oxidized toluene at the methyl group, could not transform TCE (Nelson et aL, 1988).
Wackett and Gibson (1988) reported that P. putida Fl expressing toluene dioxygenase
can transform TCE and cis-DCE at significant rates, but can not transform
tetrachioroethylene (PCE), VC, or ethylene (Wackett and Gibson, 1988). In addition,
Wackett and Householder (1989) suggested that TCE transformation has a cytotoxic
effect on P. putida Fl (Wackett and Householder, 1989). The toluene dioxygenase
was also reported on Pseudornonas sp. strain JS 150 with a TCE transformation
capability and a substrate range similar to P. putida (Haigler et al., 1992).
In 1989, Winter et al. reported a Pseudomonas mendocina strain KRI, which
catalyzed toluene by monooxygenase at para position and resulted in TCE
transformation (Winter et al., 1989). Whited and Gibson (1991) later confirmed this
result and designated this enzyme as toluene-4-monooxygenase (Whited and Gibson,
1991b). They also suggested that the strain KR1 degraded toluene through p-cresol, phydroxybenzoate, protocatechuate, and then cleaved the aromatic ring by ortho fission
(Whited and Gibson, I 99 Ia). Another monooxygenase which catalyzed toluene at the
meta position was reported on Pseudomonaspickettii PKO1 by Kaphammer et al.
(1991). They also proposed the toluene catabolic pathway through m-cresol, 3methylcatechol, and then cleaved the aromatic ring by meta fission, which is similar to
24
the proposed pathway of P. cepacia G4 (Kaphamrner et al., 1991; Kukor and Olsen,
1991; Shields et al., 1991). The degradation pathways of toluene and phenol are
summarized in Figure 2.3.
OOH
OH
o
47
meta
rOH fission
CH3
QH3
Toluene
P.
2,OH
putidaFl,JFI5C}
(TOD)
OH
meta
fission
OH
H3
OH
OH
Phenol
OOH
OH
COON
y
ON
OH
P cepacia4
me/a
fission
Figure 2.3. Summary of toluene and phenol degradation pathways.
ortho
fission
OH
Non-Regulated Substrates
A number of studies have focused on finding non-regulated substrates for use
in the in-situ treatment of TCE and other CAHs. Gao and Skeen (1999) observed cisDCE transformation in a glucose-induced microcosm and suggested that some
enzymes in the multiple pathway of glucose utilization are responsible for cis-DCE
transformation. They reasoned that substrates like glucose can be aerobically
degraded through multiple pathways. For example, glucose can be catabolized
through the Embden-Meyerhof-Pamas (EMP) pathway, the hexose monophosphate
(HMP) pathway, and tricarboxylic acid (TCA) cycle (Gao and Skeen, 1999; Voet and
Voet, 1990). The enzymes involved in these pathways might be responsible for the
cis-DCE transformation. Cellulose and hemicellulose can also be hydrolyzed or
degraded into glucose. Therefore, there may be substrates produced from the
breakdown of natural organic matter that would promote the aerobic cometabolism of
cis-DCE. Recently a microorganism was isolated that can be aerobically grown on
cis-DCE as a primary substrate (Coleman et al., 2002a). These results indicate that
microorganisms can also gain energy from cis-DCE transformation.
Glucose was also evaluated as a growth substrate for Xanthobacter strain Py2
(Ensign, 1996). Upon the induction with propylene, the glucose-grown strain Py2
expressed alkene monooxygenase, a well-known enzyme responsible for TCE
transformation. Fan and Scow (1993) reported moderate TCE transformation in soil
microcosms when methanol was added (Fan and Scow, 1993). In a microcosm study
with aquifer solids from the Moffett Field site (CA), lactate enhanced the TCE
26
transformation in both Burkholderia cepacia G4 and its mutant (PR1301) augmented
microcosms (Munakata-Marr et aL, 1996). Genetically altered Burkholderia cepacia
strain
PR1301
was capable of TCE transformation when grown on lactate; however,
with long-term stimulation TCE transformation ability was lost (McCarty et al.,
1998b; Munakata-Marr et al., 1997; Munakata-Marr et al., 1996). Sodium citrate was
used as a growth substrate to study TCE transformation kinetics by the resting cells of
Raistonia eutropha JMP134 derivative (AEK3O1/pYK3O21) (Ayoubi and Barker,
1998). Fructose was also used in combination with phenol and dichiorophenoxyacetic
acid to maintain the TCE transformation activity of Alcaligenes eutrophus JMP 134
(Muller and Babel, 1995, 1996). Genetically engineered Tn5- induced phi mutant
from the strain JMPI 34 grew on ethanol and was capable of TCE transformation (Kim
etal., 1996).
In 1999, Vancheeswaran et al. reported an enrichment culture having TCE and
cis-DCE transformation ability when grown on tetrabutoxysilane (TBOS). This
culture rapidly hydrolyzed TBOS to 1 -butanol, which was used as a growth substrate.
The formation of cis-DCE epoxide was observed, indicating cometabolic
transformation was occurring (Vancheeswaran et al., 1999). The result of this work
led to the start of this dissertation which was surveying of other potential nonregulated growth substrates for this mixed culture to cometabolize TCE and the
discovery of benzyl alcohol as an effective substrate for the cornetabolism of TCE,
cis-DCE, and VC.
27
PHENOL-GROWN MICROORGANISMS
One of the interesting characteristics of TBOS-grown culture (BA-i) is its
ability to grow on phenol, but not on toluene (Chapter 3). Based on the growth
substrates tested, the enrichment culture has growth characteristics similar to
Rhodococcus strain R-22 (Fairlee et al., 1997). This Rhodococcus strain is able to
grow on acetone, phenol, and benzyl alcohol, but did not grow on benzene or toluene.
One difference is that R-22 can also grow on propane, but BA-I could not. We have
not found any reports of chlorinated ethene cometabolism by Rhodococcus R-22.
Rhodococcus strains reported to have cometabolic transformation ability were R.
rhodochrous, which also grew on propane and cometabolized TCE and VC
(Malachowsky et al., 1994), R. corallinus, which grew on propene and cometabolized
TCE (Saeki et al., 1999), and R. erythropolis BD2, which grew on isopropylbenzene
and exhibited enzyme similar to toluene dioxygenase (Dabrock et al., 1994).
Xanthobacter strain Py2 was also reported to grow on phenol but not on toluene (Zhou
et al., 1999). Py2 expressed alkene monooxygenase, which was similar to toluene
monooxygenase, and could oxidize toluene to o-, m-, and p-cresols. Py2 was also
reported to cometabolize TCE upon growth on propene (Reij et al., 1995).
Several isolates for Moffett Field groundwater were able to grow on phenol,
but not on toluene, and exhibited enzymes similar to toluene ortho-monooxygenase
(T2MO) (Fries et al., 1997). Among these isolates, one was matched to Variovorax
paradoxus, which is of interest, since there was a report of TCE cometabolism by
phenol-grown Variovorax strain (Futamata et al., 2001b). However, another Moffett
28
isolate was also matched to V. paradoxus that was able to grow on both phenol and
toluene, and transformed TCE (Fries et al., 1997). Variovorax species were also found
in many contamination sites with many reported activities, such as growth on 2,4dichiorophenoxyacetic acid (Kamagata et al., 1997), anaerobic growth on polycyclic
aromatic hydrocarbon (Eriksson et al., 2003) and benzene (Rooney-Varga et al.,
1999), and coculture with methanotrophs (Dunfield et al., 1999).
Other reported phenol-grown microorganisms with TCE cometabolism
included Raistonia (Alcaligenes) eutropha, Raistonia pIckettii, Burkholderia cepacia,
Pseudoinonasputida, and Comamonas testosterone. Most of them can grow on
toluene, oxidize phenol at the ortho position and exhibit enzymes similar to toluene
monooxygenases (Solyanikova and Golovieva, 1999). R. eutropha JMP 134 and R.
pickettii PKO1 were reported to express enzymes similar to T2MO (Solyanikova and
Golovleva, 1999) and toluene meta-monooxygenase (T3MO) (Kim et al., 1996). P.
putida JS 150 showed multiple enzymes components which are similar to T2MO,
toluene para-monooxygenase (T4MO), and toluene dioxygenase (TOD) (Haigler et
al., 1992; Johnson and Olsen, 1995, 1997).
ENZYME INHIBITORS
Acetylene has been reported as a monooxygenase inhibitor and is often used to
indicate the involvement of a monooxygenase enzyme (Harnamura et al., 1997;
Hyman and Wood, 1985; Verce et al., 2000; Vlieg et al., 1996). Inhibition is a
reversible process causing temporary blocking of enzyme activity, while inactivation
29
is an irreversible process causing permanent damage to bacterial cells. Therefore, the
term inhibition or inhibitor is generally used when there is no evidence of permanent
cell damage or of an irreversible process. Inhibition or inactivation by acetylene has
been reported with ammonia monooxygenase (Hyman and Wood, 1985), soluble and
particulate methane monooxygenases (Vlieg et al., 1996), propane monooxygenase
(Smith et al., 2003), butane monooxygenase (Hamamura et al., 1997; Kim et al.,
2000), and alkene monooxygenase (Verce et al., 2001). Yeager et al. (1999) reported
that acetylene, however, is not an effective inactivator of toluene monooxygenase.
They found that the more effective inhibitors are the longer-chain alkynes, such as
butynes or hexynes, and the inhibition effect was correlated to time of exposure
(Yeager et al., 1999). In chapter 4, the effects of acetylene, 2-butyne, and 1-hexyne on
benzyl alcohol degradation, toluene oxidation, and TCE transformation by both benzyl
alcohol- and phenol-grown cells were compared.
30
KINETICS OF AEROBIC COMETABOLISM OF CAHs
The kinetics of cometabolism is an important consideration in the application
of bioremediation. Kinetic values are needed to estimate the amount of growth
substrate needed, the oxygen demand, and predict the duration required for the
bioremediation process of the concerned CAHs. These factors greatly affect the type
of treatment, project costs, and system operation. The parameters generally required
for the design of cometabolism processes include growth yield (Y), maximum
degradation rate (km), half-saturation constant (Ks), inhibition coefficient (K1),
transformation capacity (Ta), transformation yield (Tv), and cell decay (b) (AlvarexCohen and McCarty, 1991; Chang and Alvarez-Cohen, 1 995a; Semprini et al., 1998).
Michaelis-Menten enzyme kinetics
The aerobic cometabolism of CAlls is an enzyme-responsible mechanism and
therefore generally described using the Michaelis-Menten enzyme kinetics.
I
A
(2.1)
KSA+A
where Mx is cells mass (mg); ra, rate of compound A degradation (mg/day); kma,
maximum specific degradation rate of compound A (mg-A/mg-cells/day);
saturation concentration (mg/L); A, concentration of compound A (mg/L).
KSA,
half-
311
Haldane kinetics
For compounds that exhibit toxicity, such as phenol, degradation rates are
reduced due to the inhibition effect at high concentrations. A Haldane kinetic model
(Bailey and 011is, 1986) is generally applied:
1
kmax a
A
(2.2)
A
where Kh is the Haldane inhibitory constant (mg!L).
Transformation capacity (Ta)
The toxicity of CAl-I transformation byproducts has been observed to cause
cell activity to decrease in proportion to the amount of CABs transformed. In 1991,
Alvarez-Cohen and McCarty introduced a transformation capacity (Ta) term; a
constant representing the amount of CAB transformed divided by the amount of cells
inactivated (Alvarex-Cohen and McCarty, 1991):
TC
dM
dM
(2.3)
where T is the transformation capacity for CAlls (mg-CABs/mg-cells) and M is the
CABs transformed amount (mg). The implication of this approach is that the toxic
effects function to decrease overall cellular functions rather than affecting specific
enzyme activity alone (Alvarez-Cohen and Speitel, 2001).
32
Active cells mass (Mx)
The rate of cells mass production is a function of cell growth due to
consumption of growth substrate, cells inactivation due to toxicity of CABs
transformation, and cellular decay (Anderson and McCarty, 1996; Chang and Alvarez-
Cohen, l995a, b).
Cells mass production:
rx
=Y.
r
b
(2.4)
where r is the cells mass production rate (mg-ceils!d); Y, growth yield (mg-cells/mgsubstrate); b, cellular decay (lid).
Transformation yield (Tv)
Transformation yield (Tv) represents the maximum amount of CAHs that can
be transformed per mass of growth substrate utilized (Arp et aL, 2001). T is a
practical parameter for the estimation of the substrate amount needed in the
cometabolic treatment, which also indicates the oxygen demand in the system. It is
related to T as follow:
T
dMdMdM
dM
dM dM
=YT
(2.5)
where T is the transformation yield (mg-CABs/mg-cells); M, substrate amount (mg).
KINETIC PARAMETERS
The determination of kinetic parameters for microorganisms capable of
cometabolism has been of great interest. Cometabolism may occur relatively slowly
33
in comparison to the metabolism of the growth substrate. Therefore, the kinetics of
cometabolism can be an important consideration in bioremediation applications. For
in-situ remediation, project costs and duration can be greatly influenced by the kinetics
of the dominant biological reactions. Kinetic expressions are also important
components of fate and transport models, which are used to plan and monitor site
remediation, and to conduct risk and exposure assessment. Since this dissertation
focuses on cometabolism of CAHs by microorganisms grown on benzyl alcohol and
phenol, kinetic parameters of phenol degradation and CAlls transformation are
reviewed in Tables 2.1 and 2.2.
Table 2.1. Kinetic parameters of phenol degradation
Microorganism
K5
km
Kh
i/day mg/L
Reference
Burkholderia cepacia El
3.4
0.05
mg/L
20
Comamonas testosteroni R2
C. testosteroni E6
8.8
0.08
23
2.7
0.01
16
C. testosteroni R5
18.3
0.04
11
Pseudomonas putida P-2
P. putida P-6
0.9
0.39
291
1.1
0.37
43
P. putida P-8
1.6
0.50
649
P. putidaP35X
Actinomycetes
Raistonia eutropha
B. cepacia G4
P. putida
R. eutropha
Mixed culture
R. eutropha JMP134
Mixed culture
6.2
0.34
874
}
10.2
0.34
>15
(Lee et al., 2000)
8.6
0.94
31.5
0.80
42
(Folsom et al., 1990)
11.5
<1
470
(Hill and Robinson, 1975)
9.8
2
350
(Leonard et al., 1999)
9.3
<3.3
9.4
5.55
6.0
11
188
-
338
(Futamata et al., 200 Ia)
(Leonard and Lindley, 1999)
(Shurtliffet al., 1996)
(Muller and Babel, 1995)
348
(Goudar et al., 2000)
These phenol-grown microorganisms can be grouped by the values of Ks; low
(<0.1 mg/L), intermediate (0.1
I mg/L), and high (>1 mg/L). Low-Ks are correlated
with effective enzyme induction at low concentration, and these low-Ks
microorganisms were reported to have faster rates of TCE transformation (Futamata et
al., 2001a, b). The km of these microorganisms are in the same range (1
10mg-
phenol/mg-cells/day) with some exceptions of strains R5 and G4 (18 32 mg-
phenol/mg-cells/day). From similarity in
kmax
and variety in K5, K is likely a
controlling parameter of phenol degradation. However, most low-K5 microorganisms
also have low Kh values. The Kh values indicate the tolerance of microorganisms to
toxicity of high phenol concentration. Since lower Kh results in lower degradation rate
(Eq. 2.3), it is another important parameter for phenol degradation kinetics.
Table 2.2 shows CAHs transformation kinetics for microorganisms grown on
phenol and toluene. The
km
values are in comparable range (0.1
0.3 mg-CA}1/mg-
cells/day) with some exceptions of strains 04 and JMP 134 which have kmax about an
order of magnitude higher. The K5 values varied significantly from 0.1
16 mg/L.
Many microorganisms having low K for TCE transformation also have low Ks for
phenol degradation. The first-order rate constants
of both parameters. The variety in K5 resulted
(k1)
ink1
were used to compare the effect
varying significantly from 0.02
4 L/mg-cells/day. Therefore, K5 is an important parameter for CAHs transformation.
Another important parameter is the transformation capacity (Ta). T is defined as the
ratio of the mass of CAH transformed over cell mass used for the transformation.
Table 2.2. TCE transformation kinetics for phenol- and toluene-Qrown microorganisms.
CAHs
Phenol
ICE
Toluene
TCE
Phenol
TCE
Toluene
km,
K5
k1
(1/d)
Ic
(mg/L)
(L/mg-d)
(mg/mg)
0.2-0.5
0,15
0,14
1.1
0.59
0.95
0,80
1.2
0.66-6.6
1.5
0.39
3,8
0.03
ICE
1-30
0,17
8,64
0.02
0.007
Phenol
ICE
1-30
0.21
2.04
0.10
0.03
Mixed culture
Phenol
ICE
1-25
0.33
ii
0.03
0.08
RalstoniaeutrophajMPl34
Phenol
TCE
2.1-210
2.14
83
0.03
B. cepacia El
Phenol
0-80
0.23
3.0
0.6
Comamonas testosteroni R2
Phenol
ICE
ICE
0-80
0.13
4.2
0.5
C. testosteroni E6
Phenol
TCE
0-80
0.25
4,6
0.6
C. testosteroni R5
Phenol
TCE
0-80
0.19
1.5
0.8
P. putida P35X2
Phenol
0-200
0.10
15.8
0.06
Mixed culture (filamentous)
Phenol
ICE
ICE
5-25
0.18
0.3-0.5
cis-DCE
10-80
0.3-1.5
0.15
Actinomycetes
Burkholderia cepacia G4
Mixed culture
2
Growth
Substrate
CAHs cone.
(mg/L)
Microorganism
Reference
(Lee et al,, 2000)
(Landa et at, 1994)
(Folsometal., 1990)
(Chang and Alvarez-Cohen,
1995)
(Shurtliffet al., 1996)
(Ayoubi and Harker, 1998)
(Futamata et al., 2001)
(Bielefeldt et a!,, 1995)
Biomass reported in mg dry cell mass; units conversions assumed dry cell mass is 50% protein.
Reported kinetics were fitted to a Haldane model with Kh = 97 mg/L.
U.)
c1
36
Phenol- and toluene-grown cultures have T ranging from 0.01
0.08 mg-TCE/mg-
cell. The Phenol-grown actinomycete operated in suspended growth gas treatment
reactor had a reported T value as high as 0.6 mg-TCE/mg-cell (Lee et al., 2000).
Bielefeldt et al. (1995) reported a phenol-grown filamentous mixed culture operated in
batch reactor having T of 0.3
0.5 mg-TCE/mg-cell. In chapter 5, we determined
these kinetic parameters with our phenol- and benzyl alcohol-grown culture and more
discussion is provided.
SUMMARY
Results of TCE cometabolism by a TB OS-grown mixed culture and evidence
for TCE cometabolism by other non-regulated substrates led to this dissertation work.
Initial surveys of the other potential non-regulated growth substrates for this mixed
culture lead to the discovery of benzyl alcohol as an effective substrate for the
cometabolism of TCE, cis-DCE, and VC (Chapter 3). Pathways of benzyl alcohol
degradation were studied and compared to reported pathways of toluene and phenol
degradation (Chapter 4). Inhibition studies of acetylene, 2-butyne, and 1 -hexyne, were
performed and compared to other reports in Chapter 4. TCE transformation by benzyl
alcohol- and phenol-grown cells were studied in Chapter 4. The kinetic parameters of
benzyl alcohol and phenol degradation and TCE, cis-DCE, and VC transformation by
benzyl alcohol- and phenol-grown cells were determined and compared to other
reported kinetic values in Chapter 5.
37
REFERENCE:
Alvarez-Cohen, L., and McCarty, P. L. (1991). A Cometabolic Biotransformation
Model for Halogenated Aliphatic Compounds Exhibiting Product Toxicity.
Environ. Sci. Technol. 25(8), 1381-1387.
Alvarez-Cohen, L., and Speitel, G. E. J. (2001). Kinetics of Aerobic Cometabolism of
Chlorinated Solvents. Biodegradation 12(2), 105-126.
Anderson, J. E., and McCarty, P. L. (1996). Effect of Three Chlorinated Ethenes on
Growth Rates for a Methanotrophic Mixed Culture. Environ. Sci. Technol.
30(12), 3517-3524.
Arciero, D., Vannelli, T., Logan, M., and Hooper, A. B. (1989). Degradation of
Trichioroethylene by the Ammonia-Oxidizing Bacterium Nitrosomonas
Europaea. Biochem. Biophys. Res. Comm. 159(2), 640-643.
Arp, D. J., Yeager, C. M., and Hyman, M. R. (2001). Molecular and Cellular
Fundamentals of Aerobic Cometabolism of Trichloroethylene. Biodegradation
12(2), 81-103.
Ayoubi, P. J., and Harker, A. R. (1998). Whole-Cell Kinetics of Trichloroethylene
Degradation by Phenol Hydroxylase in a Rastonia Eutropha Jmp 134
Derivative. App!. Environ. Microbiol. 64(11), 4353-4356.
Bailey, J., and 011is, D. (1986). "Biochemical Engineering Fundamentals." McGrawHill Plublishing Co., New York.
Bielefeldt, A. R., Stensel, H. D., and Strand, S. E. (1995). Cometabolic Degradation of
Tce and Dce without Intermediate Toxicity. Journal of Environmental
Engineering 121(11), 79 1-797.
Bradley, P. M., and Chapelle, F. H. (2000). Aerobic Microbial Mineralization of
Dichloroethene as Sole Carbon Substrate. Environ. Sci. Technol. 34(1), 221223.
Chang, H.-L., and Alvarez-Cohen, L. (1995a). Model for the Cometabolic
Biodegradation of Chlorinated Organics. Environ. Sci. Technol. 29(9), 23 572367.
Chang, H.-L., and Alvarez-Cohen, L. (1 995b). Transformation Capacities of
Chlorinated Organics by Mixed Cultures Enriched on Methane, Propane,
Toluene, or Phenol. BiotechnoL Bioeng. 45,440-449.
38
Chang, H.-L., and Alvarez-Cohen, L. (1996). Biodegradation of Individual and
Multiple Chlorinated Aliphatics by Mixed and Pure Methane Oxidizing
Cultures. App!. Environ. Microbiol. 62(9), 337 1-3377.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain, J. C. (2002a).
Biodegradation of Cis-Dichloroethene as the Sole Carbon Source by a BProteobacterium. App!. Environ. Microbiol. 68(6), 2726-2730.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain, J. C. (2002b). Phylogenetic
and Kinetic Diversity of Aerobic Vinyl Chloride-Assimilating Bacteria from
Contaminated Sites. App!. Environ. Microbiol. 68(12), 6162-6171.
Dabrock, B., Kebeler, M., Averhoff, R., and Gottschalk, G. (1994). Identification and
Characterization of a Transmissible Linear Plasmid from Rhodococcus
Erythropolis Bd2 That Encodes Isopropylbenzene and Trichloroethene
Catabolism. App!. Environ. Microblo!. 60(3), 853-860.
Dabrock, B., Riedel, J., Bertram, J., and Gottschalk, G. (1992). lsopropylbenzene
(Cumene)- a New Substrate for the Isolation of Trichiorothene-Degrading
Bacteria. Arch. Microbiol. 158, 9-13.
DiSpirito, A. A., Gulledge, J., Shiemke, A. K., Murrel, J. C., Lidstrom, M. E., and
Krema, C. L. (1992). Trichioroethylene Oxidation by the MembraneAssociated Methane Monooxygenase in Type I, Type Ii and Type X
Methanotrophs. Biodegradation 2, 151-164.
Dunfield, P. F., Liesack, W., Henckel, T., Knowles, R., and Conrad, R. (1999). HighAffinity Methane Oxidation by a Soil Enrichment Culture Containing a Type Ii
Methanotroph. App!. Environ. Microbiol. 65(3), 1009-1014.
Ensign, S. A. (1996). Aliphatic and Chlorinated Alkenes and Epoxides as Inducers of
Alkene Monooxygenase and Epoxidase Activities in Xanthobacter Strain Py2.
App!. Environ. Microbiol. 62(1), 61-66.
Ensign, S. A., Hyman, M. R., and Arp, D. J. (1992). Cometabolic Degradation of
Chlorinated Alkenes by Alkene Monooxygenase in a Propylene-Grown
Xanthobacter Strain. Appi. Environ. Microbio!. 58(9), 3038-3046.
Eriksson, M., Sodersten, E., Yu, Z., Daihammar, G., and Mohn, W. W. (2003).
Degradation of Polycyclic Aromatic Hydrocarbons at Low Temperature under
Aerobic and Nitrate-Reducing Conditions in Enrichment Cultures from
Northern Soils. App!. Environ. Microbiol. 69(1), 275-284.
Ewers, J., Clemens, W., and Knackmuss, H. J., Eds. (1991). Biodegradation of
Chioroethenes Using Isoprene as Co-Substrate. Vol. 1. International
39
Symposium on Environmental Biotechnology. Ostend, Belgium: Royal
Flemish Society of Engineers.
Fairlee, J. r., Burback, B. L., and Perry, J. J. (1997). Biodegradation of Groundwater
Pollutants by a Combined Culture of Mycobacterium Vaccae and a
Rhodococcus Sp. Can. .1 Microbiol. 43, 841-846.
Fan, S., and Scow, K. (1993). Biodegradation of Trichioroethylene and Toluene by
Indigenous Microbial Populations in Soil. App!. Environ. Microbiol. 59(6),
1911-1918.
Fogel, M. M., Taddeo, A. R., and Fogel, S. (1986). Biodegradation of Chlorinated
Ethenes by a Methane-Utilizing Mixed Culture. App!. Environ. Microbiol.
51(4), 720-724.
Folsom, B. R., Chapman, P. J., and Pritchard, P. H. (1990). Phenol and
Trichloroethylene Degradation by Pseudomonas Cepacia G4: Kinetics and
Interactions between Substrates. Appi. Environ. Microbiol. 56(5), 1279-1285.
Fries, M. R., Forney, L. J., and Tiedje, J. M. (1997). Phenol- and Toluene-Degrading
Microbial Populations from an Aquifer in Which Successful Trichloroethene
Cometabolism Occurred. App!. Environ. Microbiol. 63(4), 1523-1530.
Futamata, H., Harayama, S., and Watanabe, K. (2001a). Diversity in Kinetics of
Trichloroethylene-Degrading Activities Exhibited by Phenol-Degrading
Bacteria. Appi. Microbiol. Biotechnol. 55, 248-253.
Futamata, 1-1., Harayama, S., and Watanabe, K. (2001 b). Group-Specific Monitoring of
Phenol Hydroxylase Genes for a Functional Assessment of Phenol-Stimulated
Trichioroethylene Bioremediation. App!. Environ. Microbiol. 67(10), 46714677.
Gao, J., and Skeen, R. S. (1999). Glucose-Induced Biodegradation of CisDichloroethylene under Aerobic Conditions. Wat. Res. 33(12), 2789-2796.
Goudar, C. T., Ganji, S. H., Pujar, B. G., and Strevett, K. A. (2000). Substrate
Inhibition Kinetics of Phenol Biodegradation. Water Environment Research
72(1), 50-55.
Haigler, B. E., Pettigrew, C. A., and Spain, J. C. (1992). Biodegradation of Mixtures
of Substituted Benzenes by Pseudomonas Sp. Strain Js150. App!. Environ.
Microbiol. 58(7), 223 7-2244.
Hamamura, N., Page, C., Long, T., Semprini, L., and Arp, D. J. (1997). Chloroform
Cometabolism by Butane-Grown Cf8, Pseudomonas Butanovora, and
Mycobacterium Vaccae Job5 and Methane-Grown Methylosinus
Trichosporium Ob3b. Appi. Environ. Microbiol. 63(9), 3607-36 13.
Harker, A. R., and Kim, Y. (1990). Trichioroethylene Degradation by Two
Independent Aromatic-Degrading Pathways in Alcaligenes Eutrophus Jmp134.
App!. Environ. Microbiol. 56(4), 1179-1181.
Hartmans, S., and Bont, J. A. M. d. (1992). Aerobic Vinyl Chloride Metabolism in
Mycobacterium Aurum LI. Appi. Environ. Microbiol. 58(4), 1220-1226.
Hill, 0. A., and Robinson, C. W. (1975). Substrate Inhibition Kinetics: Phenol
Degradation by Pseudomonas Putida. Biotechnol. Bioeng. 17, 1599-1615.
Hopkins, C. D., Semprini, L., and McCarty, P. L. (1993). Microcosm and in Situ Field
Studies of Enhanced Biotransformation of Trichioroethylene by PhenolUtilizing Microorganisms. App!. Environ. Microbiol. 59(7), 2277-2285.
Hyman, M. R., and Wood, P. M. (1985). Suicidal Inactiation and Labelling of
Ammonia Mono-Oxygenase by Acetylene. Biochem. J. 227, 7 19-725.
Johnson, G., and Olsen, R. (1995). Nucleotide Sequence Analysis of Genes Encoding
a Toluene/Benzene-2- Monooxygenase from Pseudomonas Sp. Strain Jsl 50.
App!. Environ. MicrobioL 61(9), 3336-3346.
Johnson, G., and Olsen, R. (1997). Multiple Pathways for Toluene Degradation in
Burkho!deria Sp. Strain Jsl 50. Appi. Environ. Microbiol. 63(10), 4047-4052.
Kamagata, Y., Fuithorpe, R. R., Tamura, K., Takami, H., Fomey, L. J., and Tiedje, J.
M. (1997). Pristine Environments Harbor a New Group of Oligotrophic 2,4Dichlorophenoxyacetic Acid-Degrading Bacteria. Appi. Environ. Microbiol.
63(6), 2266-2272.
Kaphammer, B., Kukor, J. J., and Olsen, R. H. (1991). Cloning and Characterization
of a Novel Toluene Degradative Pathway from Pseudomonas Pickettii Pko 1. In
"Biodeterioration and Biodegradation 8" (H. W. Rossmoore, Ed.). Elsevier
Applied Science, London.
Kim, Y., Arp, D. J., and Semprini, L. (2000). Chlorinated Solvent Cometabolism by
Butane-Grown Mixed Culture. Journal of Environmental Engineering
October, 934-942.
Kim, Y., Ayoubi, P., and Harker, A. R. (1996). Constitutive Expression of the Cloned
Phenol Hydroxylase Gene(S) from Alcaligenes Eutrophus Jmpl 34 and
Concomitant Trichlorethylene Oxidation. Appi. Environ. Microbiol. 62(9),
3227-3233.
41
Kukor, J. J., and Olsen, R. H. (1991). Molecular Cloning and Regulation of Genes
Encoding Meta-Cleavage of Catechols Produced from Catabolism of Phenol
and M-Cresol by Pseudomonas Pickettii Pkol. In "Biodeterioration and
Biodegradation 8" (H. W. Rossmoore, Ed.). Elsevier Applied Science, London.
Lee, S.-B., Strand, S. E., and Stensel, H. D. (2000). Sustained Degradation of
Trichioroethylene in a Suspended Growth Gas Treatment Reactor by an
Actinomycetes Enrichment. Environ. Sci. Technol. 34(15), 3261-3268.
Leonard, D., and Lindley, N. d. (1999). Growth of Raistonia Eutropha on Inhibitory
Concentrations of Phenol: Diminished Growth Can Be Attributed to
Hydrophobic Perturbation of Phenol Hydroxylase Activity. Enzyme and
Microbial Technol. 25,271-277.
Leonard, D., Youssef, C. B., Destruhaut, C., Lindley, N. D., and Queinnec, 1. (1999).
Phenol Degradation by Raistonia Eutropba: Colorimetric Determination of 2Hydroxymuconate Semialdehyde Accumulation to Control Feed Strategy in
Fed-Batch Fermentations. Biotechnol. Bioeng. 65(4), 407-415.
Little, D. C., Palumbo, A. V., Herbes, S. E., Lidstrom, M. E., Tyndall, R. L., and
Gilmer, P. J. (1988). Trichioroethylene Biodegradation by a MethaneOxidizing Bacterium. App!. Environ. Microbiol. 54(4), 951-956.
Malachowsky, K., Phelps, T., Teboli, A., Minnikin, D., and White, D. (1994). Aerobic
Mineralization of Trichioroethylene, Vinyl Chloride, and Aromatic
Compounds by Rhodococcus Species. App!. Environ. Microbiol. 60(2), 542548.
McCarty, P. L. (1997). Aerobic Cometabolism of Chlorinated Aliphatic
Hydrocarbons. In "Subsurface Restoration" (C. H. Ward, J. A. Cherry, and M.
R. Scalf, Eds.), pp. 373-395. Ann Arbor Press, Inc., Chelsea, Michigan.
McCarty, P. L., Goltz, M. N., Hopkins, G. D., Dolan, M. E., Allan, J. P., Kawakami,
B. T., and Carrothers, T. J. (1998a). Full-Scale Evaluation of in Situ
Cometabolic Degradation of Trichioroethylene in Groundwater through
Toluene Injection. Environ. Sd. Technol. 32(1), 88-100.
McCarty, P. L., Hopkins, G. D., Munakata-Marr, J., Matheson, V. G., Dolan, M. E.,
Dion, L. B., Shields, M., Fomey, L. J., and Tiedje, J. M. (1998b).
Bioaugmentation with Burkholderia Cepacia Prl3cn for in Situ Bioremediation
of Trichioroethylene Contaminated Groundwater. EPA(April), 1-11.
McCarty, P. L., and Semprini, L. (1994). Ground-Water Treatment for Chlorinated
Solvents. In "Handbook of Bioremediation." (J. E. Matthews, Ed.), pp. 87-116.
Lewis Publishers, Ann Arbor.
42
Muller, R. H., and Babel, W. (1995). Determination of the Ks Values During the
Growth of Alcaligenes Eutrophus on Phenol, 2,4-Dichiorophenoxyacetic Acid
and Fructose. Acta Biotechnol. 15(4), 347-353.
Muller, R. I-I., and Babel, W. (1996). Measurement of Growth at Very Low Rates
(M>0), an Approach to Study the Energy Requirement for the Surial of
A icaligenes Eutrophus Jmp 134. App!. Environ. Microbiol. 62(1), 147-151.
Munakata-Marr, J., Matheson, V. G., Forney, L. J., Tiedje, J. M., and McCarty, P. L.
(1997). Long-Term Biodegradation of Trichioroethylene Influenced by
Bioaugmentation and Dissolved Oxygen in Aquifer Microcosms. Environ. Sci.
Technol. 31(3), 786-791.
Munakata-Marr, J., McCarty, P. L., Shields, M. S., Reagin, M., and Francesconi, S. C.
(1996). Enhancement of Trichloroethyiene Degradation in Aquifer
Microcosms Bioaugmented with Wild Type and Genetically Altered
Burkholderia (Pseudomonas) Cepacia G4 and Pri. Environ. Sci. Technol.
30(6), 2045-2052.
National Center for Environmental Assessment. (2001). Trichloroethylene Health Risk
Assessment. U.S. Environmental Protection Agency,
http://cfpub.epa.gov/ncea.
Nelson, M. J. K., Montgomery, S. 0., Mahaffey, w. R., and Pritchard, P. H. (1987).
Biodegradation of Trichioroethylene and Involvement of an Aromatic
Biodegradative Pathway. App!. Environ. Microbiol. 53(5), 949-954.
Nelson, M. J. K., Montgomery, S. 0., O'Neill, E. J., and Pritchard, P. FL (1986).
Aerobic Metabolism of Trichioroethylene by a Bacterial Isolate. Appi.
Environ. Microbiol. 52(2), 383-384.
Nelson, M. J. K., Montgomery, S. 0., and Pritchard, P. H. (1988). Trichloroetbylene
Metabolism by Microorganisms That Degrade Aromatic Compounds. App!.
Environ. Microbiol. 54(2), 604-606.
Oldenhuis, R., Oedzes, J. Y., Waarde, v. d. J. J., and Janssen, D. B. (1991). Kinetics of
Chlorinated Hydrocarbon Degradation by Methylosinus Trichosporium Ob3b
and Toxicity of Trichloroethylene. App!. Environ. Microbiol. 57, 7-14.
Oldenhuis, R., Vink, R. L. J. M., Janssen, D. B., and Witholt, B. (1989). Degradation
of Chlorinated Aliphatic Hydrocarbons by Methylosinus Trichosporium Ob3b
Expressing Soluble Methane Monooxygenase. App!. Environ. Microbiol. 55,
28 19-2826.
43
Reij, M., Kieboom, J., Bont, J. d., and Hartmans, S. (1995). Continuous Degradation
of Trichioroethylene by Xanthobacter Sp. Strain Py2 During Growth on
Propene. Appi. Environ. Microbiol. 61(8), 2936-2942.
Rooney-Varga, J. N., Anderson, R. T., Fraga, J. L., Ringelberg, D., and Lovley, D. R.
(1999). Microbial Communities Associated with Anaerobic Benzene
Degradation in a Petroleum-Contaminated Aquifer. Appi. Environ. Microbiol.
65(7), 3056-3063.
Saeki, H., Akira, M., Furuhashi, K., Averhoff, B., and Gottschalk, G. (1999).
Degradation of Trichioroethene by a Linear-Plasmid-Encoded Alkene
Monooxygenase in Rhodococcus Corallinus (Nocardia Corallina) B-276.
Micro biol. 145, 1721-1730.
Semprini, L. (1997). Strategies for the Aerobic Co-Metabolism of Chlorinated
Solvents. Curr. Opin. Biotechnol. 8, 296-308.
Semprini, L., Ely, R. L., and Lang, M. M. (1998). Modeling of Cometabolism for the
in Situ Biodegradation of Trichioroethylene and Other Chlorinated Aliphatic
Hydrocarbons. In "Bioremediation: Principles and Practice Volume I,
Fundamentals and Applications" (S. K. Sikdar, and R. L. Irvine, Eds), pp. 89134. Technomic Publishing, Lancaster.
Semprini, L., Robert, P. V., Hopkins, G. D., and McCarty, P. L. (1990). A Field
Evaluation of in-Situ Biodegradation of Chlorinated Ethenes: Part 2.
Cometabolic Transformations. Ground Water 30(1), 37-44.
Shields, M. S., Montgomery, S. 0., Chapman, P. J., Cuskey, S. M., and Pritchard, P.
H. (1989). Novel Pathway of Toluene Catabolism in the TrichioroethyleneDegrading Bacterium G4. Appi. Environ. Microbiol. 55(6), 1624-1629.
Shields, M. S., Montgomery, S. 0., Cuskey, S. M., Chapman, P. J., and Pritchard, P.
H. (1991). Mutants of Pseudomonas Cepacia G4 Defective in Catabolism of
Aromatic Compounds and Trichloroethylene. Appi. Environ. Microbiol. 57(7),
1935-1941.
Shurtliff, M. M., Parkin, G. F., Weathers, L. J., and Gibson, D. T. (1996).
Biotransformation of Trichloroethylene by a Phenol-Induced Mixed Culture.
Journal ofEnvironmental Engineering 122(7), 581-589.
Smith, C. A., OReil1y, K. T., and Hyman, M. R. (2003). Characterization of the Initial
Reactions During the Cometabolic Oxidation of Methyl Tert-Butyl Ether by
Propane-Grown Mycobacterium Vaccae Job5. App!. Environ. Microbiol.
69(2), 796-804.
Solyanikova, I. P., and Golovieva, L. A. (1999). Phenol Hydroxylases: An Update.
Biochemistry (Moscow) 64(4), 437-446.
Steffan, R. J., Sperry, K. L., Walsh, M. T., Vainberg, S., and Condee, C. W. (1999).
Field-Scale Evaluation of in Situ Bioaugmentation for Remediation of
Chlorinated Sovients in Groundwater. Environ. Sci. Technol. 33(16), 277 1278 1.
Tovanabootr, A., and Semprini, L. (1998). Comparison of Tce Transformation
Abilities of Methane- and Propane-Utilizing Microorganisms. Bioremediation
J 2(2), 105-124.
Tsien, H. C., Brusseau, G. A., Hanson, R. S., and Wackett, L. P. (1989).
Biodegradation of Trichioroethylene by Methyl osinus Trichosporium Ob3b.
App!. Environ. Microbiol. 55(12), 3155-3161.
U.S. EPA. (2001). Chemical Summary for Toluene. Office of Pollution Prevention
and Toxics, U.S. Environmental Protection Agency,
http://www.epa.gov!opptintr/chemfactlstoluene.txt.
U.S. EPA. (2003a). Consumer Factsheet On: 1 ,2-Dichloroethylene. U.S.
Environmental Protection Agency, http://www.epa.gov/safewater/dwhlcvoc/1 2-dich2 .html.
U.S. EPA. (2003b). Consumer Factsheet On: Trichloroethylene. U.S. Environmental
Protection Agency, http://www.epa.gov/safewater/dwhlc-voc/trichlor.html.
U.S. EPA. (2003c). Consumer Factsheet On: Vinyl Chloride. U.S. Environmental
Protection Agency,
http://www.epa.gov/safewater/contaminants/dw_contamfs/vinylchl ,htmL
van Hyickama Vlieg, J. E. T., de Koning, W., and Janssen, D. B. (1996).
Transformation Kinetics of Chlorinated Ethenes by Met hylosinus
Trichosporium Ob3b and Detection of Unstable Epoxides by on-Line Gas
Chromatography. App!. Environ. Microbiol. 62, 3304-3312.
Vancheeswaran, S., Halden, R. U., Williamson, K. J., James D. Ingle, J., and
Semprini, L. (1999). Abiotic and Biological Transformation of
Tetraalkoxysilanes and Trichioroethene/Cis- I ,2-Dichloroethene Cometabolism
Driven by Tetrabutoxysilane-Degrading Microorganisms. Environ Sci.
Technol. 33(7), 1077-1085.
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2000). Characterization of an Isolate
That Uses Vinyl Chloride as a Growth Substrate under Aerobic Conditions.
Appi. Environ. Microbiol. 66(8), 3535-3542.
45
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2001). Transition from Cometabolic
to Growth-Linked Biodegradation of Vinyl Chloride by a Pseudomonas Sp.
Isolated on Ethene. Environ. Sci. Technol. 35(21), 4242-4251.
Vlieg, J. E. T. v. H., Koning, W. d., and Janssen, D. B. (1996). Transformation
Kinetics of Chlorinated Ethenes by Methylosinus Trichosporium Ob3b and
Detection of Unstable Epoxides by on-Line Gas Chromatography. App!.
Environ. Microbiol. 62(9), 3304-3312.
Voet, D., and Voet, J. (1990). "Biochemistry." John Wiley and Sons, New York.
Vogel, T. M., and McCarty, P. L. (1985). Biotransformation of Tetrachloroethylene to
Trichioroethylene, Dichloroethylene, Vinyl Chloride and Carbon Dioxide
under Methanogenic Conditions. App!. Environ. Microbiol. 49(5), 1080-1083.
Wackett, L. P., Brusseau, G. A., Householder, S. R., and Hanson, R. S. (1989). Survey
of Microbial Oxygenases Trichioroethylene Degradation by PropaneOxidizing Bacteria. App!. Environ. Microbiol. 55, 2960-2964.
Wackett, L. P., and Gibson, D. T. (1988). Degradation of Trichioroethylene by
Toluene Dioxygenase in Whole-Cell Studies with Pseudomonas Putida Fl.
Appi. Environ. Micro biol. 54(7), 1703-1708.
Wackett, L. P., and Householder, S. R. (1989). Toxicity of Trichioroethylene to
Pseudomonas Putida Fl Is Mediated by Toluene Dioxygenase. Appl. Environ.
Microbiol. 55, 2723-2725.
Whited, G. M., and Gibson, D. T. (1991a). Separation and Partial Characterization of
the Enzymes of the Toluene-4-Monooxygenase Catabolic Pathway in
Pseudomonas Mendocina Krl. J. Bacteriol. 173(9), 30 17-3020.
Whited, G. M., and Gibson, D. T. (1991b). Toluene-4-Monooxygenase, a ThreeComponent Enzyme System That Catalyzes the Oxidation of Toluene to PCresol in Pseudomonas Mendocina Krl. I Bacteriol. 173(9), 3010-3016.
Wilson, J. T., and Wilson, B. H. (1985). Biotransformation of Trichloroethylene. Appi.
Environ. Microbiol. 49(1), 242-243.
Winter, R. B., Yen, K.-M., and Ensley, B. D. (1989). Efficient Degradation of
Trichioroethylene by a Recombinant Escherichia Coli. Bio Technology 7(3),
282-285.
Yeager, C. M., Bottomley, P. J., Arp, D. J., and Hyman, M. R. (1999). Inactivation of
Toluene 2-Monooxygenase in Burkholderia Cepacia G4 by Alkynes. App!.
Environ. Microbiol. 65(2), 632-639.
Zhou, N.-Y., Jenkins, A., Chion, C. K. N. C. K., and Leak, D. J. (1999). The Alkene
Monooxygenase from Xanthobacter Strain Py2 Is Closely Related to Aromatic
Monooxygenases and Catalyzes Aromatic Monohydroxylation of Benzene,
Toluene, and Phenol. App!. Environ. Microbiol. 65(4), 1589-1595.
47
Chapter 3
AEROBIC BIOTRANSFORMATION OF CHLORINATED ETHENES BY A
MIXED CULTURE GROWN ON A BROAD RANGE OF SUBSTRATES
Sarun Tejasen and Lewis Semprini
To be submitted to:
Water Research
International Association on Water Quality, London, U.K.
The aerobic transformation of TCE and cis-DCE by tetrabutoxysilane-grown
microorganism (Vancheeswaran et al., 1999) led to the investigation of other
alternative substrates, including benzyl alcohol, a novel substrate for promoting
cometabolism. The culture grew on all carboxylic compounds and alcohols tested,
except single carbon substrates, such as formate and methanol. The levels of cis-DCE
and TCE transformations achieved were dependent on the growth substrate.'
Compounds that did not serve as growth substrates included saturated hydrocarbons
(methane, propane, and butane), unsaturated hydrocarbons (ethylene), and non-
oxygenated aromatics (benzene, toluene, and p-xylene). The culture did grow on
alcohols such as 1-propanol, 1-butanol, ethanol, benzyl alcohol, and phenol, and
demonstrated cis-DCE transformation. Among carboxylic compounds, butyrate was a
very effective substrate for supporting cis-DCE transformation. Transformation of
cis-DCE was also achieved when the culture was grown on glucose. The culture when
grown on benzyl alcohol was equally effective in promoting TCE, cis-DCE, and vinyl
chloride transformations, as when grown on phenol, a well-known TCE cometabolic
substrate. The results indicate benzyl alcohol as a novel effective substrate for the
aerobic cometabolism of TCE, cis-DCE, and vinyl chloride. Since benzyl alcohol is
not a regulated compound, such as phenol and toluene, it may have use in applications
for in-situ bioremediation.
riLeJ
INTRODUCTION
Trichloroethylene (TCE) is one of the most frequently found organic
contaminants in groundwater. One of the promising remediation methods is in-situ
biological treatment. Numerous studies have demonstrated that aerobic bacteria can
transform TCE via cometabolic oxidation to harmless end products, such as chloride
ion and carbon dioxide (Arp et al., 2001; McCarty, 1997; McCarty and Semprini,
1994; Semprini, 1997b). Reported here are results of a study where varieties of
substrates were tested for inducing TCE and cis-dichloroethylene (cis-DCE) aerobic
cometabolic activity with an enrichment culture. Among these substrates, benzyl
alcohol was found to be an effective substrate to support cometabolism of TCE, cisDCE, and vinyl chloride (VC).
Wilson and Wilson (1985) first noted aerobic transformation of TCE through
the use of soil microbial communities fed natural gas. Since then, much interest has
focused on finding different microorganisms that could cometabolize chlorinated
aliphatic hydrocarbons (CAH5). Microorganisms that grow on the following
substrates have been shown to have cometabolism potential: ammonia (Arciero et al.,
1989), butane (Kim et al., 2000), cresol (Folsom et al., 1990; Nelson et al., 1988;
Wackett and Gibson, 1988), dichlorophenoxyacetic acid (Harker and Kim, 1990),
ethylene (Ensign et al., 1992), isoprene (Ewers et al., 1991), isopropylbenzene
(Dabrock et aL, 1992), methane (Chang and Alvarez-Cohen, 1996; Oldenhuis et al.,
1991; Semprini et al., 1990), phenol (Ayoubi and Harker, 1998; Folsom et al., 1990;
Hopkins et al., 1993), propane (Tovanabootr and Semprini, 1998; Wackett et al.,
50
1989), propene (Ensign et al., 1992), and toluene (Nelson et al., 1987; Shields et al.,
1989; Wackett and Gibson, 1988).
One of the parameters used to assess cometabolic TCE transformation is
transformation yield (Tv) (Alvarez-Cohen and Speitel, 2001; McCarty, 1997;
Semprini, I 997b). T represents the mass of CAH transformed per unit mass of
primary substrate consumed (McCarty, 1997). Among those primary substrates
mentioned above, methane is the most studied substrate with a broad range of TCE
transformation activities reported. Reported T of methanotrophs for TCE ranges
from 0.015-0.034 mg-TCE/mg-methane (without formate supplement) (Alvarez-
Cohen and Speitel, 2001), while higher T (0.052-0.222 mg-TCE/mg-phenol) were
reported with mixed culture grown on phenol as substrate (Jenal-Wanner and
McCarty, 1997; Shurtliffet al., 1996). In addition, in-situ microorganisms had a
similar T when they were grown on toluene or phenol. Microorganisms grown on
these aromatic compounds have been reported to cometabolize TCE better than those
grown on methane (Bielefeldt et al., 1995; Hopkins et al., 1993).
In a demonstration test of in-situ cometabolism conducted at Edward Air Force
Base (CA), toluene was chosen as the cometabolic growth substrate over phenol, since
it has a lower odor threshold than phenol and as a liquid it could be easily handled.
Phenol also has the potential of forming chlorinated phenols upon chlorination
(McCarty et al., I 998a). However, toluene is also a common groundwater
contaminant with a recommended maximum contaminant level (MCL) in drinking
51
water of I mg/i (U.S. EPA, 2001). Thus, as a regulated compound, obtaining
regulatory approval for in-situ use may prove difficult in some cases.
A number of studies have focused on finding non-regulated substrates for use
in the in-situ treatment of TCE and other CAHs. Gao and Skeen (1999) reported cisDCE transformation in a glucose induced microcosm. Recently a microorganism was
isolated that can aerobically grow on cis-DCE as a primary substrate (Coleman et al.,
2002). These results indicate that microorganisms can gain energy for growth from
cis-DCE. Thus, aerobic microorganisms that grow on a broad range of substrates and
transform cis-DCE are not unreasonable. Glucose was also evaluated as a growth
substrate for Xanthobacter strain Py2 (Ensign, 1996). Upon the induction with
propylene, the glucose-grown strain Py2 expressed alkene monooxygenase, a wellknown enzyme responsible for TCE transformation.
Fan and Scow (1993) reported moderate TCE transformation in soil
microcosms when methanol was added. Fructose was used in combination with
phenol and dichiorophenoxyacetic acid to maintain the TCE transformation activity of
Alcaligenes eutrophus JMP134 (Muller and Babel, 1995, 1996). Genetically
engineered Tn5- induced phi mutant from the strain JMPI34 grew on ethanol and was
capable of TCE transformation (Kim et al., 1996). Genetically altered Burkholderia
cepacia strain
PR1301
was capable of TCE transformation when grown on lactate;
however, with long-term stimulation TCE transformation ability was lost (McCarty et
al., I 998b; Munakata-Marr et al., 1997; Munakata-Marr et al., 1996).
52
Vancheeswaran et al. (1999) reported an enrichment culture having TCE and
cis-DCE transformation ability when grown on tetrabutoxysilane (TBOS). This
culture rapidly hydrolyzed TBOS to I -butanol, which was used as a growth substrate.
The formation of cis-DCE epoxide was observed, indicating cometabolic
transformation was occurring (Vancheeswaran et al., 1999).
In this study, the TBOS enrichment culture was tested for its ability to grow on
a broad range of substrates and cometabolize cis-DCE and TCE. Substrates tested
included silicon-based organic compounds (TBOS, tetrapropoxysilane (TPOS), and
tetraphenoxysilane), phenolic compounds (phenyl acetate, p-cresol, and phenol),
alcohols (methanol, ethanol, I -propanol, I -butanol, 4-methyl-benzyl alcohol, 3-buten2-ol, 3-buten-1 -ol, 2-buten-1 -ol, and benzyl alcohol), organic acids (formate, acetate,
propionate, butyrate, and benzoate), aromatic compounds (benzene, toluene, and pxylene), saturated and unsaturated hydrocarbons (methane, propane, butane, and
ethylene), acetone, glucose, and methyl-tert-butyl-ether (MTBE).
MATERIALS AND METHODS
Bacterial Culture
An aerobic mixed culture was obtained from a TBOS-grown culture that had
cis-DCE and TCE transformation ability (Vancheeswaran et al., 1999), which was
originally started by using activated sludge of Wastewater Treatment plant in
Corvallis, OR. To produce a reproducible culture for these studies, the enriched
53
mixed culture was grown in 710-mi batch serum bottles (Wheaton Glass Co.,
Miliville, NJ) on 400-mg/i TBOS in 400-mi basal salt medium (BSM)
(Vancheeswaran et al., 1999) to an optical density of 1.0 at 550 nm (OD(550)). The
culture was centrifuged, rinsed with BSM, and stored at an approximate concentration
of 3 mg/mi at 80°C in 7% dimethylsulfoxide (DMSO).
To grow cells for the batch reactor tests, the frozen cells were thawed, rinsed
with fresh BSM, and batch-grown on 400 mg/I TBOS or 1-butanol (Table 3.1) to an
OD(550) of 0.6. Cells were centrifuged, rinsed with BSM, and diluted to
concentrations of approximately 0.3 mg/mi for use in growth and cometabolic
transformation tests.
Chemicals
Tetrabutoxysilane (TBOS) (98% purity) and tetrapropoxysilane (TPOS) (98%
purity) were obtained from Gelest, Inc. (Tullytown, PA). Tetraphenoxysilane (98%
purity), 4-methyl-benzyl alcohol (99% purity), 3-buten-2-ol (97% purity), 3-buten-1-oI
(99% purity), and 2-buten- 1-ol(97% purity) were purchased from Lancaster, Inc.
(Peiham, NH). Phenyl acetate (99% purity), I -butanol (99.8% purity), sodium acetate
(99% purity), sodium propionate (99% purity), sodium butyrate (98% purity), sodium
benzoate (98% purity), propane (98% purity), vinyl chloride (VC) (99.5% purity), cis-
dichloroethylene (cis-DCE) (99.9 % purity), and ethylene (99% purity) were
purchased from Aldrich Chemical, Inc. (Milwaukee, WI). D-glucose (99% purity)
was purchased from Spectrum Chemical Corp. (Gardena, CA). Phenol, HPLC-grade
54
acetone, and spectrophotometric-.grade methyl alcohol were purchased from
Mallinckrodt Baker, Inc. (Parid, KY). p-cresol (98% purity) was obtained from
Matheson Coleman & Bell Manufacturing Chemists (Norwood, OH). Ethanol was
purchased from AAPER Alcohol and Chemical Co. (Shelbyville, KY). Certified
A.C.S.-grade 1-propanol and trichioroethylene (99.9 % purity) were purchased from
Fisher Scientific (Fair Lawn, NJ). Sodium formate (99% purity), benzyl alcohol (99%
purity), benzene (99% purity), toluene (99% purity), and p-xylene (99% purity) were
purchased from Acros Organics (NJ). Butane (98% purity) was purchased from
Specialty Products & Equipment (Houston, TX).
Batch Reactors
The experiments were conducted at 20°C in batch reactors consisting of 60-mI
BSM in 125-mI crimp-sealed bottles (Wheaton Glass Co., Miliville, NJ) with
headspace air to maintain aerobic conditions. The reactors were autoclaved to prevent
contamination, and BSM was added. After the addition of substrates and CABs,
bottles were inverted and shaken at 200 rpm overnight prior to cell addition.
Simultaneous growth and cometabolism tests were initiated by adding 0.3 mg of cells
(as described above) into batch reactors. Preparations and conditions of batch reactors
are shown in Table 3.1. Substrates, CAHs, and OD(550) were monitored periodically
until OD(550) and CAHs concentration remained constant. The batch reactors were
operated in triplicate unless indicated otherwise. Pure oxygen was added when
negative pressure developed in the reactors. TCE and cis-DCE were added using a
55
saturated stock solution in deionized water. VC was added as a pure gas. Control
bottles containing CAHs alone and with resting cells (-M.3 mg-cells without substrate)
were monitored along with active bottles.
Table 3.1. Growth Substrates and Cometabolic Conditions of the Batch Experiments
Experiments
Cometabolic transformation
of ICE during growth on
TBOS and I -butanol
Substrate tested for growth
and TCE and cis-DCE
transformation
Substrates tested for VC,
cis-DCE, and TCE
cometabolic transformation
Cell
Inoculate
Growth
Substrate
CAH Transformation During
Growth o1pfic Substrate
Growth substrate
CAH
TBOS
TBOS (0-30 jtmo1),
I -butanol (0-160 iimol)
TCE (0.16 mo1)
I -butanol
See Table 3.2
cis-DCE (15 imo1),
TCE (1.8 lImo])
I -butanol
Butyrate (97 lImoI),
Glucose (81 imo1),
Benzyl alcohol (57 lImoI)
VC (11 iimol),
cis-DCE (4.1 lImol),
TCE (0.7 lImol)
56
Analytical methods
Calibration curves for all compounds were developed using external standards.
The aqueous concentration of TBOS and I -butanol was determined by
dichloromethane (DCM) extraction and GC-FID analysis. Aqueous samples were
extracted with DCM in a 1:1 volume ratio, by vigorously mixing for 5 mm on a vortex
mixer and centrifuging at 14,000 rpm for 3 mm. The DCM extract (1 .il) was injected
by a Hewlett Packard HP 7673A automatic sampler to a Hewlett Packard 5890 gas
chromatograph (GC) equipped with a flame ionization detector (FID).
Chromatographic separation was achieved with a 30 m
x
0.32 mm
x
0.25 im RTX-5
column (Restek Corporation, Bellefonte, PA) and operated at the following
temperature profile: 5 mm at 35°C; gradient, 40°C/mm; and 5 mm at 300°C. Carrier
gas was helium at a flow rate of 1.5 mI/mm.
Headspace concentrations of cis-DCE and VC were determined on a HP5 890GC using a 30 m
x
0.53 mm GSQ-PLOT column (J&W Scientific, CA), operated at
140°C, with a FID detector. The carrier gas was helium at a flow rate of 1.5 mI/mm.
TCE headspace concentration was determined on a HP5890-GC using a 30 m x 0.25
mm
x
1.4 tm HP-624 column (Hewlett Packard, Wilmington, DE) operated at 140°C,
with an electron conductivity detector (01 Analytical, College Station, TX), The
carrier gas was helium at a flow rate of 1.5 mi/mm with an argon/methane mixture
(95%:5%) for make-up gas. The total mass of chlorinated compounds in the serum
bottle was calculated by using published Henry's constants (Gossett, 1987) and
57
equilibrium mass balances using the measured headspace concentration and the
volumes of the gas and liquid phases.
The cell concentration was determined as total suspended solids (TSS) (1985),
using 0.2-pm membrane filter (Micro Separation Inc., MA). Cell growth was
observed by monitoring the optical density at 550 nm (OD(550)) using a Hewlett
Packard
8453
UV-Visible spectrophotometer.
RESULTS
Cometabolic Transformation of TCE During Growth on TBOS and I -Butanol
In a first series of tests, TBOS (Si(0C4H9)4) and its hydrolysis byproduct, I butanol (C4H9OH), were compared as growth substrates and inducers of TCE
transformation. The enrichment culture used to inoculate the batch reactors was
grown on TBOS. The mass used to inoculate the culture was small so that TCE
cometabolism would be the result from microbes grown on the substrate. The amount
of substrate added varied from 0-30 tmoI TBOS and 0-160 jimol I -butanol to
investigate the TCE transformation yields that resulted with an initial TCE mass of
0.16 imol. The constant TBOS concentration in the control bottle indicated that the
abiotic hydrolysis of TBOS was insignificant over the time scale of the growth
experiments (data not shown). TCE was transformed during the degradation of both
TBOS and I -butanol. While greater amounts of TBOS and I -butanol consumption led
to more transformation of TCE, the relationship was not linear. Hydrolysis of one mol
of TBOS produces 4 mol of 1-butanol and similar TCE transformations (0.10-0.12
imol) were observed when normalized this way (Figure 3.1). The results indicate that
growth on 1 -butanol was responsible for TCE cometabolism, and TBOS mainly
served as a source of I -butanol upon hydrolysis.
Butanol Consumption (jimol)
0
40
0
10
80
120
160
30
40
0.14
0.12
0.10
0.08
C,)
F-.
0.04
0.02
[.IIII
20
TBOS Consumption (1imol)
Figure 3.1. Comparison of TCE transformation by a culture growing on
TBOS and 1-butanol. TCE mass 0.16 jtmol.
59
Growth on a Broad Range of Substrates and the Cometabolic Transformation of TCE
and cis-DCE
Other potential growth substrates for this culture were evaluated along with the
culture's ability to transform TCE and cis-DCE when grown on each substrate. The
amount of each substrate added to the batch growth reactors was based on an equal
electron donor potential equivalent to 81 tmol 1-butanol (Table 3.2). Enrichments of
1-butanol-grown cells (0.3 mg dry weight) were added to triplicates of 125-mi bottles
containing 60 ml media with 15 tmol cis-DCE or 1.8 imo1 TCE and the specific
growth substrate. Growth was observed by monitoring optical density at 550 nm
(OD(550)). Initial OD(550) from the enrichment additions were less than 0.1 in all
bottles. Results of cis-DCE transformation studies with formate (973 imol), acetate
(243 imol), propionate (139 !lmol), butyrate (97 imol), and 1-butanol (81 tmol), as
growth substrates, are shown in Figure 3.2. The culture grew to OD(550) of 0.3-0.4
on acetate, propionate, butyrate, and 1-butanol, but no growth occurred with formate.
Transformations of cis-DCE were observed after OD(550) reached the maximum
point indicating the possible inhibition of these substrate on cis-DCE transformation.
Higher amounts cis-DCE transformed (13.9 - 14.7 imol) were achieved with growth
on I -butanol and butyrate. Growth on acetate, and propionate resulted in 11.0 and 6.2
tmo1 cis-DCE transformed, respectively. Conservative estimates of T on each of the
substrates are as follow: 0.18 mol cis-DCE/mol I -butanol, 0.14 mol cis-DCE/mol
butyrate, 0.045 mol cis-DCE/mol acetate, and 0.045 mol cis-DCE/mol propionate.
These estimates are conservative since cis-DCE was almost completely transformed
with several substrates and exposure to more cis-DCE may have resulted in continued
transformation.
20
forrmte
j15
0
0.6
U)
w10
0.4
C,
.-
0
C-,
0
0
20
40
60
002
0
80
20
0
100
40
60
80
100
Time (hi)
Time(hr)
0.8
0
U)
1h=.
--------------------
-
0.6
acetate
0.4
Time (hr)
Time (hr)
0.8
te
propionate
04
54
C)
0
0
20
40
60
80
0
100
40
60
80
100
Time (hi)
Time (hi)
0.8-
20
fr.
0.6
butyrate
w10
0
0.
5
0
20
'butyrate
.
40
60
80
0
100
20
40
60
80
100
Time (hr)
Time (hr)
0.8
- ------------------------------------
------------
U)
butanut
002
resting cells central
0
0
Time (hr)
20
40
60
80
lime (hr)
Figure 3.2. cis-DCE transformation and growth (as observed by
OD(550)) of the enrichment grown on formate, acetate, propionate,
butyrate, and 1 -butanol, compared to resting cells control (x) with no
substrate added.
100
61
The results for all substrates are summarized in Table 3.2. The mixed culture
grew on tetrapropoxysilane (TPOS,
Si(0C3H7)4)
and showed similar transformation of
cis-DCE and TCE as when grown on 1-propanol (C3H7OH), a TPOS hydrolysis
byproduct. Tetraphenoxysilane (Si(0C6H5)4) and phenyl acetate (C6H5CH2COOH)
were as effective as their hydrolysis byproduct, phenol (C6H5OH), in serving as
growth substrates for cis-DCE and TCE cometabolism. It is interesting to note that the
optical density achieved with phenol and tetraphenoxysilane were equal, while phenyl
acetate was higher likely due to presence of acetate that is also generated during the
hydrolysis of this compound. The culture also grew on glucose with effective
transformation of cis-DCE and some TCE transformation. Butyrate and benzoate
were shown to be effective substrates for cis-DCE cometabolism, although they did
not induce much of the TCE transformation. Benzyl alcohol (C61-I5CH2OH) was an
effective substrate that resulted in complete transformation of cis-DCE and TCE.
Optical densities achieved with benzyl alcohol were in the range of that observed with
growth on phenol.
Some interesting trends were observed from the tests with the different growth
substrates (Table 3.2). Most of the growth-supporting substrates are very soluble in
water. The culture did not grow on methanol or formate, suggesting the requirement
of substrate having two or more carbon atoms. The culture also did not grow on
saturated hydrocarbons (methane, propane and butane), unsaturated hydrocarbon
(ethylene), or non-oxygenated aromatics (benzene, toluene, and p-xylene). The
culture, however, grew on oxygenated ringed compound including phenol, p-cresol
62
Table 3.2. Substrates tested for growth and cis-DCE and TCE transformation
Substrate
Amount1
tmol
Tetrabutoxysilane
Tetrapropoxysilane
Tetraphenoxysilane
Phenyl acetate
Methanol
Ethanol
1-Propanol
1-Butanol
Formate
Acetate
Propionate
Butyrate
Benzoate
Acetone
Phenol
p-cresol
4-Methyl-benzyl alcohol
3-buten-2-ol
3-buten-1-oi
2-buten-l-ol
Glucose
Benzyl alcohol
Methane
Propane
Butane
Ethylene
Benzene
Toluene
p-Xylene
MTBE
20
27
17
54
324
162
108
81
973
243
139
97
65
122
69
57
49
88
88
88
81
57
243
97
75
162
65
54
46
65
Final
cis-DCE
OD(550) % Removal2
T3
0.34
45%
0.34
0.29
40%
0.22
0.21
100%
0.88
0.27
100%
0.28
0.02
0%
ND
0.24
32%
0.03
0.24
44%
0.06
0.24
98%
0.18
0.02
0%
ND
0.33
73%
0.05
0.30
42%
0.04
0.40
93%
0.14
0.24
84%
0.19
0.22
58%
0.07
0.21
100%
0.22
0.19
100%
0.26
0.05
0%
ND
0.03
0%
ND
0.22
89%
0.15
0.23
67%
0.11
0.40
93%
0.17
0.18
100%
0.26
0.07
0%
ND
0.04
0%
ND
0.04
0%
ND
0.02
0%
ND
0.04
0%
ND
0.02
0%
ND
0.03
0%
ND
0.03
0%
ND
ICE________
T3
% Removal2
26%
19%
100%
81%
ND
ND
ND
0.023
0.012
0.106
0.027
ND
ND
ND
16%
0.004
ND
ND
ND
9%
6%
ND
ND
ND
ND
0.002
0.002
ND
0.026
0.032
ND
ND
100%
100%
ND
ND
21%
20%
44%
100%
ND
ND
ND
ND
ND
ND
ND
ND
1. Substrate amounts were based on the equal electrons transferred in aerobic utilization.
2. % Removal based on a single addition of 15 tmoI cis-DCE or 1.8 lImo! TCE
3. Transformation yield (Tv) in .tmol CAHs per moi substrate
ND = not determined
0.004
0.004
0.010
0.032
ND
ND
ND
ND
ND
ND
ND
ND
63
(CH3C6H4OH), benzyl alcohol, and benzoate with benzoate the least effective in
supporting cis-DCE and TCE cometabolism. The culture did not grow on 4-methylbenzyl alcohol (CH3C61-14C1-120H), suggesting a specific enzymatic pathway. The
culture also grew on I -propanol, I -butanol, acetone, acetate, and propionate, with aIls
showing some cis-DCE transformation. Growth occurred on 3-buten-1-ol and 2buten-1 -ol, but not on 3-buten-2-ol, indicating the important characteristic of having
hydroxyl group at the terminal carbon.
Growth on acetate, butyrate, and I -butanol gave a similar percent
transformation of cis-DCE (73-98%) (Table 3.2). Effective cis-DCE transformation
(100%) was correlated with good TCE transformation ability, while limited cis-DCE
transformation (less than 50%) was correlated with limited TCE transformation
potential. Effective transformation of both cis-DCE (100%) and TCE (100%) was
achieved with the aromatic substrates, phenol, tetraphenoxysilane (that hydrolyzes to
phenol), p-cresol, and benzyl alcohol.
Further tests compared acetate, butyrate, and 1 -butanol with cells exposed to
higher amounts of cis-DCE (20.5 46 tmol) (data not shown). Acetate and butyrate
were rapidly degraded like 1 -butanol, indicating that the microbial population could
rapidly metabolize these substrates. The greatest amount of cis-DCE was transformed
with cells grown on butyrate, followed by I -butanol and acetate, giving the 1,, of
0.423, 0.230, and 0.038 tmol cis-DCE transformed per tmol substrate utilized,
respectively. The results show the ability to achieve high transformation yields of cisDCE on butyrate and I -butanol grown cells.
Test of cometabolic growth substrates for VC, cis-DCE, and TCE transformation
Butyrate, glucose, and benzyl alcohol were chosen as growth substrates to
compare the cometabolic transformation of VC, cis-DCE and TCE based on the initial
screening showing cis-DCE and TCE transformation potential (Table 3.2) and the
novelty of their promoting cometabolism of cis-DCE and TCE compared to phenol
which has been well studied. The initial amounts of butyrate, glucose, benzyl alcohol,
VC, cis-DCE, and TCE are shown in Table 3.1. Initially, butyrate was added as
growth substrate to all the batch reactors. After the initial growth on butyrate, glucose
and then benzyl alcohol were added to all batch reactors after 2.1 and 6.1 days,
respectively (Figure 3). The amount of these substrates added was based on equal
electron donor equivalents for growth. Similar increases in optical density were
observed with growth on all the substrates. During growth on butyrate, no
transformation of VC was observed, while effective cis-DCE and limited TCE
transformation was observed. The total transformation of cis-DCE and TCE was 5
imol and 0.2 imol, respectively, with growth on butyrate. The more effective
transformation of cis-DCE is consistent with the earlier results with butyrate (Table
3.2).
65
15
1.2
10
C)
0.9
U)
2. 0.6
0
0.3
0
2
4
6
8
10
0
days
2
4
6
8
10
6
8
10
days
8
1.2
0
E6
C
w4
0
0.9
0
LiV)
0
06
0.3
0
0
0
2
4
6
8
10
0
days
2
4
days
1.5
1.2
1.2
C
U)
0.9
0.9
0?- 0.6
0 0.3
w 0.6
0.3
0
0
0
2
4
days
6
8
10
0
2
4
6
days
8
Figure 3.3. Transformation of VC, cis-DCE, and TCE, and growth
observed by OD(550) after addition of butyrate, glucose, and benzyl
alcohol at 0, 2.1, and 6.1 days respectively. Controls were: cells w/o
substrate, x; and no-cells, -.
10
Utilization of glucose, as a substrate, did not yield significant transformation of
VC, however, 6 imol of cis-DCE and 0.4 mo1 of TCE were transformed. VC, cisDCE, and TCE transformation was observed with the addition of benzyl alcohol at 6.1
days, and three repeated additions of VC, cis-DCE, and TCE were transformed
without further addition of benzyl alcohol. Approximate transformation yields based
on the amount of benzyl alcohol utilized (mg of CAHs transformed per mg of
substrate utilized) for VC, cis-DCE, and TCE was 0.354, 0.251, and 0.053,
respectively.
DISCUSSION AND CONCLUSION
The tests of growth and cis-DCE and TCE cometabolism on a broad range of
substrates show that this culture can transform cis-DCE upon growth on most of the
substrates tested that had at least two carbons and were substituted with oxygen.
Among tested substrates, 1-butanol, butyrate, glucose, and benzyl alcohol were non-
regulated compounds and achieved more than 90% transformation of initial amount of
cis-DCE present. TCE, however, was less effectively transformed. The only nonregulated compound to promote 100% transformation of TCE was benzyl alcohol.
Phenol, p-cresol, and phenol-release compounds (tetraphenoxysilane and phenyl
acetate) were very effective in both cis-DCE and TCE transformation, but both phenol
and p-cresol are regulated chemicals that may not be acceptable for release in
environment, even though they are readily biodegraded.
67
The mechanism by which cells when grown on simple substrates, such as 1butanol, butyrate, and glucose, and promote cis-DCE cometabolism is not known.
Gao and Skeen (1999) observed cis-DCE transformation in a glucose-induced
microcosm and suggested that some enzymes in the multiple pathway of glucose
utilization are responsible for cis-DCE transformation. They reasoned that substrates
like glucose can be aerobically degraded through multiple pathways. For example,
glucose can be catabolized through the Ernbden-Meyerhof-Pamas (EMP) pathway, the
hexose monophosphate (HMP) pathway, and tricarboxylic acid (TCA) cycle (Gao and
Skeen, 1999; Voet and Voet, 1990). The enzymes involved in these pathways might
be responsible for the cis-DCE transformation. Cellulose and hemicellulose can also
be hydrolyzed or degraded into glucose. Therefore, there may be substrates produced
from the breakdown Of natural organic matter that would promote the aerobic
cometabolism of cis-DCE. In addition, Verce et al. (2002) reported the cometabolic
transformation of cis-DCE by a VC-grown culture, and Coleman et al. (2002) recently
reported a microcosm that can grow on cis-DCE and cometabolize VC and TCE.
These results indicate that there are multiple natural-mechanisms for the removal of
these chloroethenes in the aerobic environment.
The ability of the culture to transform cis-DCE with microbes grown on simple
organic acids, such as butyrate and acetate, has potential implications on the intrinsic
transformation processes. Anaerobic transformation of PCE and TCE often do not
proceed beyond cis-DCE (Bouwer and McCarty, 1983; Lee et al., 1998; McCarty and
Semprini, 1994; Semprini, 1997a; Vogel and McCarty, 1985). Acetate, propionate,
r4
and butyrate may be produced by fermentation reactions in anaerobic portions of
plume during the breakdown of hydrocarbons and other organic compounds (Fennell
and Gossett, 1997, 1998). At the distal parts of contaminant plume the growth of
organisms on residual organic acids under aerobic conditions might promote intrinsic
treatment of cis-DCE. The results presented here indicate more research is needed on
this potential process.
The constitutive induction of oxygenase enzymes with addition of harmless
substrates has also been reported. In a microcosm study with aquifer solids from the
Moffet Field site (CA), lactate enhanced the TCE transformation in both Burkholderia
cepacia G4 and its mutant (PR1301) augmented microcosms (Munakata-Marr et al.,
1996). Sodium citrate was used as a growth substrate to study TCE transformation
kinetics by the resting cells of Raistonia eutropha JMP 134 derivative
(AEK3OI!pYK3O21) (Ayoubi and Harker, 1998). It may be that with this mixed
culture, oxygenase enzymes are being stimulated to various degrees by the broad
range of substrates.
The important characteristics of growth substrates for this enrichment culture
are 1) high solubility; 2) the substrate must have two or more carbon atoms; and 3)
have some oxygen or hydroxide substitution. Based on the growth substrates tested,
the enrichment culture has growth characteristic similar to Rhodococcus strain R-22
(Fairlee et al., 1997). This Rhodococcus strain is able to grow on acetone, phenol, and
benzyl alcohol, but did not grow on benzene or toluene. One difference is that R-22
can also grow on propane, but our enrichment culture could not. We have not found
any reports of the study of chlorinated ethene cometabolism by R-22. The
Rhodococcus strain reported to have TCE transformation ability was R. erythropolis
BD2, which was grown on isopropylbenzene and exhibited enzyme similar to toluene
dioxygenase (Dabrock et al., 1994). Currently we are attempting to isolate a pure
benzyl alcohol-grown culture from the mixed culture that has the TCE, cis-DCE, and
VC cometabolism abilities.
The enrichment also has the ability to hydrolyze and utilize complex
substrates, including TBOS, TPOS, tetraphenoxysilane and phenyl acetate. Similar
effectiveness in the cometabolic transformation of cis-DCE and TCE was achieved as
when their hydrolysis byproducts (1 -butanol, 1 -propanol, and phenol) were directly
used. This result might be useful for in-situ bioremediation where phenol could be
replaced with relatively inert compounds such as tetraphenoxysilane and phenyl
acetate.
Among the growth substrates tested, benzyl alcohol is of special interest since
it is non-toxic, commonly used as food-flavoring agent (Mallinckrodt Baker Inc.,
2000), and it is very effective in promoting the transformation of VC, cis-DCE, and
TCE. These compounds are important since VC and cis-DCE are often present as
anaerobic transformation products of perchioroethene (PCE) and TCE. When cells
were grown on butyrate and glucose, and then switched to benzyl alcohol, high
transformation yields for vinyl chloride, cis-DCE, and TCE were achieved, and
essentially no lag in the transformation was observed. It is likely that cells grown on
butyrate and glucose were induced on benzyl alcohol to transform these compounds.
70
The cometabolism of vinyl chloride was reported by both soluble and particulate
methane monooxygenase (Chang and Alvarez-Cohen, 1996), alkene monooxygenase
(Ensign et al., 1992), and in in-situ field tests when toluene ortho-monooxygenase was
the dominant enzyme present (Hopkins and McCarty, 1995; Jenal-Wanner and
McCarty, 1997). Toluene dioxygenase, expressed by Pseudomonas putida strain Fl,
did not transform vinyl chloride, while it transformed both TCE and cis-DCE
(Wackett and Gibson, 1988). The enzyme expressed for benzyl alcohol degradation is
more likely a monooxygenase than dioxygenase. Current studies are underway to
identify the oxygenase enzymes involved.
Benzyl alcohol was found to be an effective substrate for the cometabolism of
VC, cis-DCE, and TCE. Benzyl alcohol might be a potential substitute for toluene or
phenol for in-situ bioremediation. Benzyl alcohol is a liquid phase at room
temperature, like toluene, with many advantages in physical, chemical, and
toxicological properties (National Technical Information Service, 1989; National
Toxicology Program, 2002). It has a higher boiling point (205°C), lower vapor
pressure (0.15 mmHg @ 25°C), and higher solubility (40 g/l @ 20°C), compared to
toluene (110.6°C, 22.0 mmHg @ 20°C, and 0.5 15 gIl @ 20°C), which potentially
makes it more suitable in field site applications. For the safety of handling and
transportation, benzyl alcohol has a moderate flammability with irritation effect in
respiratory tract, while toluene is highly flammable with severe central nervous system
effect from inhalation. For in-situ application, toluene is a common groundwater
contaminant, with a recommended maximum contamination level (MCL) of 1 mgll,
71
while benzyl alcohol is a non-regulated compound and commonly used as food
flavoring agent (Mallinckrodt Baker Inc., 2000). More work is needed with direct
comparison of this culture with phenol as a substrate. A broader evaluation is also
needed on the microorganisms that can grow on benzyl alcohol and cometabolize
chlorinated ethenes.
ACKNOWLEDGEMENTS
This research was supported by a Ph.D. scholarship from Thai Ministry of
University Affairs (Bangkok, Thailand) and a research grant from the US Department
of Defense sponsored Strategic Environmental Research and Development Program
under agreement CU-I 127. This article has not been reviewed by the agency, and no
official endorsement should be inferred.
72
REFERENCE
Alvarez-Cohen, L., and Speitel, G. E. J. (2001). Kinetics of Aerobic Cometabolism of
Chlorinated Solvents. Biodegradation 12(2), 105-126.
American Public Health Association. (1985). Standard Methods for the Examination
of Water and Wastewater. 16th ed. APHA, New York.
Arciero, D., Vannelli, T., Logan, M., and Hooper, A. B. (1989). Degradation of
Trichioroethylene by the Ammonia-Oxidizing Bacterium Nitrosomonas
Europaea. Biochem. Biophys. Res. Comm. 159(2), 640-643.
Arp, D. J., Yeager, C. M., and Hyman, M. R. (2001). Molecular and Cellular
Fundamentals of Aerobic Cometabolism of Trichloroethylene. Biodegradation
12(2), 81-103.
Ayoubi, P. J., and Harker, A. R. (1998). Whole-Cell Kinetics of Trichloroethylene
Degradation by Phenol Hydroxylase in a Rastonia Eutropha Jmp134
Derivative. App!. Environ. Microbiol. 64(11), 4353-4356.
Bielefeldt, A. R., Stensel, H. D., and Strand, S. E. (1995). Cometabolic Degradation of
Tce and Dee without Intermediate Toxicity. Journal ofEnvironmental
Engineering 121(11), 79 1-797.
Bouwer, E. J., and McCarty, P. L. (1983). Transformation of 1- and 2-Carbon
Halogenated Aliphatic Organic Compounds under Methanogenic Conditions.
Appi. Environ. Microbiol. 45, 1286-1294.
Chang, H.-L., and Alvarez-Cohen, L. (1996). Biodegradation of Individual and
Multiple Chlorinated Aliphatics by Mixed and Pure Methane Oxidizing
Cultures. Appi. Environ. Microbiol. 62(9), 337 1-3377.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain, J. C. (2002). Biodegradation
of Cis-Dichloroethene as the Sole Carbon Source by a B-Proteobacterium.
App!. Environ. Microbiol. 68(6), 2726-2730.
Dabrock, B., Kebeler, M., Averhoff, R., and Gottschalk, G. (1994). Identification and
Characterization of a Transmissible Linear Plasmid from Rhodococcus
Erythropolis Bd2 That Encodes Isopropylbenzene and Trichioroethene
Catabolism. Appi. Environ. Microbiol. 60(3), 853-860.
Dabrock, B., Riedel, J., Bertram, J., and Gottschalk, G. (1992). Isopropylbenzene
(Cumene)- a New Substrate for the Isolation of Trichiorothene-Degrading
Bacteria. Arch. Microbiol. 158, 9-13.
73
Ensign, S. A. (1996). Aliphatic and Chlorinated Alkenes and Epoxides as Inducers of
Alkene Monooxygenase and Epoxidase Activities in Xanthobacter Strain Py2.
App!. Environ. Microbiol. 62(1), 61-66.
Ensign, S. A., Hyman, M. R., and Arp, D. J. (1992). Cometabolic Degradation of
Chlorinated Alkenes by Alkene Monooxygenase in a Propylene-Grown
Xanthobacter Strain. App!. Environ. Microbiol. 58(9), 3038-3046.
Ewers, J., Clemens, W., and Knackmuss, H. J., Eds. (1991). Biodegradation of
Chioroethenes Using Isoprene as Co-Substrate. Vol. 1. International
Symposium on Environmental Biotechnology. Ostend, Belgium: Royal
Flemish Society of Engineers.
Fairlee, J. r., Burback, B. L., and Perry, J. J. (1997). Biodegradation of Groundwater
Pollutants by a Combined Culture of Mycobacterium Vaccae and a
Rhodococcus Sp. Can. .1. Microbiol. 43, 841-846.
Fennell, D. E., and Gossett, J. M. (1997). Comparison of Butyric Acid, Ethanol, Lactic
Acid, and Propionic Acid as Hydrogen Donors for the Reductive
Dechlorination of Tetrachioroethene. Environ. Sci. Technol. 31(3), 918-926.
Fennell, D. E., and Gossett, J. M. (1998). Modeling the Production of and Competition
for Hydrogen in a Dechiorinating Culture. Environ. Sd. Technol. 32(16), 245 02460.
Folsom, B. R., Chapman, P. J., and Pritchard, P. H. (1990). Phenol and
Trichloroethylene Degradation by Pseudomonas Cepacia G4: Kinetics and
Interactions between Substrates. App!. Environ. Microbiol. 56(5), 1279-1285.
Gao, J., and Skeen, R. 5. (1999). Glucose-Induced Biodegradation of CisDichioroethylene under Aerobic Conditions. Wal. Res. 33(12), 2789-2796.
Gossett, J. M. (1987). Measurement of Henry's Law Constants for C and C2
Chlorinated Hydrocarbons. Environ. Sci. Technol. 21(2), 202-208.
Harker, A. R., and Kim, Y. (1990). Trichloroethylene Degradation by Two
Independent Aromatic-Degrading Pathways in Alcaligenes Eutrophus Jmp 134.
App!. Environ. Microbiol. 56(4), 1179-1181.
Hopkins, G. D., and McCarty, P. L. (1995). Field Evaluation of in Situ Aerobic
Cometabolism of Trichloroethylene and Three Dichloroethylene Isomers
Using Phenol and Toluene as the Primary Substrates. Environ. Sci. Technol.
29(6), 1628-1637.
74
Hopkins, 0. D., Semprini, L., and McCarty, P. L. (1993). Microcosm and in Situ Field
Studies of Enhanced Biotransformation of Trichioroethylene by PhenolUtilizing Microorganisms. Appi. Environ. Microbiol. 59(7), 2277-2285.
Jenal-Wanner, U., and McCarty, P. L. (1997). Development and Evaluation of
Semicontinuous Slurry Microcosms to Simulate in Situ Biodegradation of
Trichioroethylene in Contaminated Aquifers. Environ. Sd. Technol. 31(10),
2915-2922.
Kim, Y., Arp, D. J., and Semprini, L. (2000). Chlorinated Solvent Cometabolism by
Butane-Grown Mixed Culture. Journal of Environmental Engineering
October, 93 4-942.
Kim, Y., Ayoubi, P., and Harker, A. R. (1996). Constitutive Expression of the Cloned
Phenol Hydroxylase Gene(S) from Alcaligenes Eutrophus Jmpl 34 and
Concomitant Trichiorethylene Oxidation. App!. Environ. Microbiol. 62(9),
3227-3233.
Lee, M. D., Odom, J. M., and Buchanan Jr., R. J. (1998). New Perspectives on
Microbial Dehalogenation of Chlorinated Solvents: Insights from the Field.
Annu. Rev. Microbiol. 52, 423-452.
Mallinckrodt Baker Inc. (2000). Material Safety Data Sheet (Msds): Benzyl Alcohol.,
www.jtbaker.com/msds/bl885.htm.
McCarty, P. L. (1997). Aerobic Cometabolism of Chlorinated Aliphatic
Hydrocarbons. In "Subsurface Restoration" (C. H. Ward, J. A. Cherry, and M.
R. Scalf, Eds.), pp. 373-3 95. Ann Arbor Press, Inc., Chelsea, Michigan.
McCarty, P. L., Goltz, M. N., Hopkins, G. D., Dolan, M. E., Allan, J. P., Kawakami,
B. T., and Carrothers, T. J. (1 998a). Full-Scale Evaluation of in Situ
Cometabolic Degradation of Trichloroethylene in Groundwater through
Toluene Injection. Environ. Sci. Technol. 32(1), 88-100.
McCarty, P. L., Hopkins, G. D., Munakata-Marr, J., Matheson, V. G., Dolan, M. E.,
Dion, L. B., Shields, M., Forney, L. J., and Tiedje, J. M. (l998b).
Bioaugmentation with Burkholderia Cepacia Pr1301 for in Situ Bioremediation
of Trichioroethylene Contaminated Groundwater. EPA(April), 1-11.
McCarty, P. L., and Semprini, L. (1994). Ground-Water Treatment for Chlorinated
Solvents. In "Handbook of Bioremediation." (J. E. Matthews, Ed.), pp. 87-116.
Lewis Publishers, Ann Arbor.
Muller, R. H., and Babel, W. (1995). Detennination of the Ks Values During the
Growth of Alcaligenes Eutrophus on Phenol, 2,4-Dichiorophenoxyacetic Acid
and Fructose. Acta Biotechnol. 15(4), 347-3 53.
75
Muller, R. H., and Babel, W. (1996). Measurement of Growth at Very Low Rates
(M>.0), an Approach to Study the Energy Requirement for the Surial of
Alcaligenes Eutrophus Jmp 134. App!. Environ. Microbiol. 62(1), 147-151.
Munakata-Marr, J., Matheson, V. G., Fomey, L. J., Tiedje, J. M., and McCarty, P. L.
(1997). Long-Term Biodegradation of Trichioroethylene Influenced by
Bioaugmentation and Dissolved Oxygen in Aquifer Microcosms. Environ. Sci.
Technol. 31(3), 786-791.
Munakata-Marr, J., McCarty, P. L., Shields, M. S., Reagin, M., and Francesconi, S. C.
(1996). Enhancement of Trichioroethylene Degradation in Aquifer
Microcosms Bioaugmented with Wild Type and Genetically Altered
Burkholderia (Pseudomonas) Cepacia 04 and Pri. Environ. Sci. Technol.
30(6), 2045-2052.
National Technical Information Service. (1989). Tr-343: Toxicology and
Carcinogenesis Studies of Benzyl Alcohol (Cas No. 100-51-6) in F344/N Rats
and B6c3f1 Mice (Gavage Studies)., Vol. 2001. National Institue of
Environmental Health Sciences, http://ntp-server.niehs.nih.gov.
National Toxicology Program. (2002). H&S: Benzyl Alcohol 100-51-6, Vol. 2001.
National Institue of Environmental Health Sciences, http:Ilntpserver.niehs.nih. gov.
Nelson, M. J. K., Montgomery, S. 0., Mahaffey, w. R., and Pritchard, P. H. (1987).
Biodegradation of Trichloroethylene and Involvement of an Aromatic
Biodegradative Pathway. App!. Environ. Microbiol. 53(5), 949-954.
Nelson, M. J. K., Montgomery, S. 0., and Pritchard, P. H. (1988). Trichloroethylene
Metabolism by Microorganisms That Degrade Aromatic Compounds. App!.
Environ. Microbiol. 54(2), 604-606.
Oldenhuis, R., Oedzes, J. Y., Waarde, v. d. J. J., and Janssen, D. B. (1991). Kinetics of
Chlorinated Hydrocarbon Degradation by Methylosinus Trichosporium Ob3b
and Toxicity of Trichioroethylene. Appi. Environ. Microbiol. 57, 7-14.
Semprini, L. (l997a). In Situ Transformation of Halogenated Aliphatic Compounds
under Anaerobic Conditions. In "Subsurface Restoration" (C. H. Ward, 3. A.
Cherry, and M. R. Scaif, Eds.), pp. 429-450. Ann Arbor Press, Inc., Chelsea,
Michigan.
Semprini, L. (1997b). Strategies for the Aerobic Co-Metabolism of Chlorinated
Solvents. Curr. Opin. Biotechnol. 8, 296-308.
76
Semprini, L., Robert, P. V., Hopkins, G. D., and McCarty, P. L. (1990). A Field
Evaluation of in-Situ Biodegradation of Chlorinated Ethenes: Part 2.
Cometabolic Transformations. Ground Water 30(1), 37-44.
Shields, M. S., Montgomery, S. 0., Chapman, P. J., Cuskey, S. M., and Pritchard, P.
H. (1989). Novel Pathway of Toluene Catabolism in the TrichloroethyleneDegrading Bacterium G4. Appi. Environ. Microbiol. 55(6), 1624-1629.
Shurtliff, M. M., Parkin, G. F., Weathers, L. J., and Gibson, D. T. (1996).
Biotransformation of Trichioroethylene by a Phenol-Induced Mixed Culture.
Journal of Environmental Engineering 122(7), 581-589.
Tovanabootr, A., and Semprini, L. (1998). Comparison of Tee Transformation
Abilities of Methane- and Propane-Utilizing Microorganisms. Bioremediation
J. 2(2), 105-124.
U.S. EPA. (2001). Chemical Summary for Toluene. Office of Pollution Prevention
and Toxics, U.S. Environmental Protection Agency,
http://www.epa.gov/opptintr/chemfact!s_toluene.txt.
Vancheeswaran, S., Halden, R. U., Williamson, K. J., James D. Ingle, J., and
Semprini, L. (1999). Abiotic and Biological Transformation of
Tetraalkoxysilanes and Trichioroethene/Cis- I ,2-Dichloroethene Cometabolism
Driven by Tetrabutoxysilane-Degrading Microorganisms. Environ Sci.
Technol. 33(7), 1077-1085.
Voet, D., and Voet, J. (1990). "Biochemistry." John Wiley and Sons, New York.
Vogel, T. M., and McCarty, P. L. (1985). Biotransformation of Tetrachloroethylene to
Trichloroethylene, Dichloroethylene, Vinyl Chloride and Carbon Dioxide
under Methanogenic Conditions. App!. Environ. Microbiol. 49(5), 1080-1083.
Wackett, L. P., Brusseau, G. A., Householder, S. R., and Hanson, R. S. (1989). Survey
of Microbial Oxygenases Trichloroethylene Degradation by PropaneOxidizing Bacteria. App!. Environ. Microbiol. 55,2960-2964.
Wackett, L. P., and Gibson, D. T. (1988). Degradation of Trichioroethylene by
Toluene Dioxygenase in Whole-Cell Studies with Pseudomonas Putida Fl.
Appi. Environ. Microbiol. 54(7), 1703-1708.
77
Chapter 4
INDUCTION OF 2-HYDROXY-BENZYL ALCOHOL PRODUCTION
AND TRICHLOROETHYLENE COMETABOLISM BY
A BENZYL ALCOHOL GROWN CULTURE
Sarun Tejasen, Mark E. Dolan, and Lewis Semprini
78
ABSTRACT
Benzyl alcohol was found to be an effective non-regulated substrate for
tricbloroethylene (TCE) aerobic cometabolism. Multiple additions of benzyl alcohol
led to an increase in TCE transformation rate up to 0.05 mg-TCE/mg-cells/day and
transformation capacity
(Ta) of 0.035 mg-TCEImg-cells. The transformation rate and
T were not improved with cells previously exposed to TCE. By-product formation
from benzyl alcohol degradation changed from benzaldehyde to 2-hydroxy benzyl
alcohol (2HBA) with multiple additions of benzyl alcohol, indicating a change in the
degradation pathway. Resting cells studies showed the production rate of 2HBA was
correlated to the TCE transformation rate. When the culture was grown on phenol,
TCE transformation and 2HBA production rates doubled and were stable during long-
term additions. Benzyl alcohol- and phenol-grown cells both oxidized toluene to ocresol, which indicated the similarity of benzyl alcohol ortho-monooxygenase, phenol
hydroxylase, and toluene ortho-monooxygenase. Inhibition studies with acetylene, 2butyne, and 1 -hexyne indicated the same enzyme was expressed by the benzyl
alcohol- and phenol-grown cells. The results suggested that an ortho-monooxygenase
was responsible for TCE cometabolism, and benzyl alcohol may be a promising
substrate for TCE cometabolism at contamination sites.
79
INTRODUCTION
Trichioroethylene (TCE) is a suspected human carcinogen that is a frequently
detected subsurface contaminant. Toluene and phenol are well studied as effective
substrates for growing aerobic microorganism capable of cometabolizing TCE. An insitu field demonstration showed microorganisms grown on aromatic substrates, such
as phenol and toluene, cometabolized TCE better than those grown on methane
(Bielefeldt et al., 1995; Hopkins et al., 1993). In a full scale field demonstration,
toluene was added to stimulate indigenous microorganisms, since it can be easily
handled as a pure liquid, and phenol in groundwater can form chlorinated phenols
upon chlorination (McCarty et al., 1998). Toluene is however listed as a groundwater
contaminant with a recommended maximum contaminant level (MCL) in drinking
water of I mg/I (U.S. EPA, 2001). Thus obtaining regulatory approval to add toluene
to the subsurface could be problematic in some cases.
In a previous study, we found that benzyl alcohol is an effective substrate for
the aerobic cometabolism of TCE, cis-DCE, and VC (Tejasen and Semprini, 2003).
Benzyl alcohol is a non-regulated compound, is non toxic, and is commonly used as
food flavoring agent (Mallinckrodt Baker Inc., 2000). It is also a pure liquid that can
be easily handled like toluene. Thus, TCE cometabolism by a benzyl alcohol-grown
culture, benzyl alcohol degradation pathway, and its relationship to TCE
cometabolism were evaluated in this study. The mixed culture status was also
monitored using molecular methods during prolonged growth on benzyl alcohol.
[1L
Evidence for changes in the utilization pathway with substrate induction has
been previously reported. Alcaligenes eutrophus strain JMP 134 when degrading
phenol changed from ortho- to meta-cleavage pathway as the growth rate increased
(Muller and Babel, 1996). Different enzymes can also be induced when bacteria are
exposed to different compounds. Xanthobacter strain Py2 grew on glucose without
evidence of alkene monooxygenase or epoxidase, and was able to produce both
enzymes upon the induction of propylene (Ensign, 1996). Chlorinated compounds
such as TCE and cis-DCE have also been reported to induce both toluene degradation
and TCE transformation (Leahy et al., 1996; McClay et al., 1995; Shingleton et al.,
1998).
Since our mixed culture does not grow on toluene (Tejasen and Semprini,
2003), phenol was compared to benzyl alcohol as a growth substrate. The ability of
toluene to be oxidized by both benzyl alcohol- and phenol-grown cells was
investigated, and used to evaluate byproducts and enzymes involved. Enzyme
inhibitors were also used to investigate the enzymes involved in benzyl alcohol
degradation and toluene oxidation. Acetylene has been used as a monooxygenase
inhibitor (Hamamura et al., 1997; Hyman and Wood, 1985; Verce et al., 2000; Vlieg
et al., 1996). Yeager et al. (1999) reported that acetylene however is not an effective
enzymes inhibitor of aromatic substrates utilization. They found that the longer-chain
alkynes, such as butynes or hexynes are more effective inhibitors (Yeager et al., 1999).
In this study, the effect of acetylene, 2-butyne, and 1-hexyne on benzyl alcohol
81
degradation and toluene oxidation was studied and compared between benzyl alcoholand phenol-grown enrichments.
In addition, benzyl alcohol was also compared to phenol as growth substrate
for the resting cells transformation of TCE. TCE transformation rates, TCE
transformation capacities (Tc), benzyl alcohol degradation rates, and byproduct
production rates were compared.
METHODS
Microbial Culture
The aerobic enrichment culture was obtained from a tetrabutoxysilane
(TBOS)-grown culture that was able to cometabolize cis-DCE and TCE
(Vancheeswaran et al., 1999). Earlier studies showed that this mixed culture when
grown on benzyl alcohol could cometabolize vinyl chloride, cis-DCE and TCE
(Tejasen and Semprini, 2003). Isolation of a pure culture was attempted using
traditional aseptic plating techniques (Madigan et aL, 1996). The benzyl alcohol
grown mixed culture was plated in 1.0% agar in basal salt medium (BSM), as
described by Vancheeswaran et al. (1999) (Vancheeswaran et al., 1999), with benzyl
alcohol (200 mg/I) as the growth substrate. An isolated colony was picked and then
grown in batch serum bottles on 400-mg/I benzyl alcohol in BSM media to an optical
density of 1.0 (at 550 urn), then centrifuged and washed to an approximate
concentration of 3 mg/mI (dry weight). Despite repeat efforts to obtain a pure culture,
a I 6S-DNA based method indicated at least two dominant microorganisms remained
in the culture. Thus the experiments were performed with a highly enriched mixed
culture. To provide a uniform innoculum for growing cells in batch reactors, the
culture was stored at 80 °C in 7% dimethylsulfoxide (DMSO).
Monitoring of Microbial Communities
Terminal Restriction Fragment Length Polymorphisms (T-RFLP) analysis was
performed to monitor the microbial communities. The microbial samples were
obtained after each cycle of continuous growth on benzyl alcohol in the resting cells
tests described above, and stored at -20°C. DNA was extracted from 200-j.tl microbial
samples using FastDNA SPIN kits for soil (Bio 101, Calsbad, CA). PCR reactions
were conducted with FAM-labeled 27F and 338R (e-coli numbering) primers. T-
RFLP analyses were conducted in a maimer similar to that of Liu et aL (Liu et al.,
1997) using restriction enzymes Mnll and Hin6I (MBI Fermentas Inc., MD),
Chemicals
Benzyl alcohol (99% purity), benzaldehyde (98+% purity), toluene (99%
purity), and o-cresol (98% purity) were purchased from Acros Organics (NJ). Phenol
(99.1 % purity) was purchased from Mallinckrodt Baker, Inc. (Parid, KY). 2-hydroxy-
benzyl alcohol (99% purity), 2-butyne (98% purity), and I -hexyne (97% purity) were
purchased from Lancaster, Inc. (Peiham, NH). Acetylene (99.6%) was purchased
from AIRCO (Vancouver, WA). Trichloroethylene (99.9 % purity) was purchased
from Fisher Scientific (Fair Lawn, NJ).
Batch Reactor Construction
All the experiments were performed in batch reactors of different sizes (160-mi
and 710-mi serum bottles or 27-mi vials, Wheaton Glass Co., Miliville, NJ). The
reactors were autoclaved and filled with a combination of basal salt media
(Vancheeswaran
Ct
al., 1999) and headspace air to maintain aerobic conditions.
Benzyl alcohol as a growth substrate was added as a pure liquid. Phenol was added
using a high-concentration stock (40 mg/mi) in deionized (Dl) water. TCE was added
using saturated TCE in DI water at 20°C. Bottles and vials were inverted to prevent
headspace leakage, incubated at 20°C, and shaken at 200 rpm to provide good mass
transfer. Non-oxygen control bottles were purged with nitrogen gas, which was
passed through tube furnace at 600°C to remove oxygen.
Cometabolic Transformation of ICE during Growth on Benzyl Alcohol
This study was to compare ICE transformation that resulted from a single and
multiple additions of benzyl alcohol for microbial growth. The frozen culture was
thawed at room temperature and washed with media to remove the DMSO. The
culture was batch-grown on benzyl alcohol (250 mg/I) to the optical density of 0.53.
The liquid culture (1.3 mg in 10 ml) was added to 160-mi serum bottles containing 50-
ml BSM. Tests were performed with a single addition of benzyl alcohol at a
84
concentration of 400 mg/I, compared to multi-addition tests where benzyl alcohol was
added at the concentration of 100, 100 and 200 mg/I, respectively. The initial
concentrations of TCE were about 0.6-0.8 mg/I. Additional TCE was added when the
TCE was completely transformed to determIne T.
Preparation of Resting Cell Studies
To investigate changes in benzyl alcohol degradation and TCE transformation
upon successive growth without effects from any possible byproducts, resting cell
studies were performed. The frozen culture, upon thawing, was grown on benzyl
alcohol or phenol for up to 13 growth cycles with 100 mg/I sequential additions. Afler
complete substrate degradation within the specific growth cycle, the liquid culture was
harvested and washed with fresh media to cell concentrations of approximately 2.5
mg/mi. An equivalent cell mass was added to each batch reactor. The cells were
assayed as described below. Liquid cultures from each growth cycle were also kept
frozen at -2 0°C for T-RFLP analysis to monitor for changes in the microbial
population.
Resting Cells Transformation Assays
In resting cells transformation assays we determined TCE transformation rates,
TCE transformation capacities, benzyl alcohol degradation rates, and byproduct
production rates from benzyl alcohol degradation. 1-mi of liquid culture (2.5 mg) was
added into two triplicate sets of 27-mi vials. The first set contained 5 ml of media
85
with a TCE concentration of 4 mg/I. The second set contained 9 ml of media with a
benzyl alcohol concentration of 50 mg/I. TCE transformation rates, benzyl alcohol
degradation rates, and byproduct production rates were determined from monitoring
the concentration of each compound over a 2.0 - 3.5 hour period after cell addition.
After the initial rates were determined, prolonged monitoring of TCE was also
performed to determine the TCE transformation capacity (To) of cells for each growth
cycle. T was determined by monitoring the TCE concentration until transformation
stopped without the addition of an energy source. Multiple additions of TCE were
often required. Controls were also included which contained I) no-cells, 2) killedcells, and 3) no-oxygen.
TCE Induction
The effect of TCE exposure to changes in TCE transformation and benzyl
alcohol degradation were investigated. The mixed culture was successively grown on
benzyl alcohol, as described in the preparation of resting cells studies, in a presence of
1 mg/I TCE. A liquid culture (2.5 mg/mI) was prepared in a similar manner and
assayed for the resting cells transformation as described above.
Identification of Enzymes Involved in Benzyl Alcohol Degradation
Three methods were used to investigate enzymes involved in degradation of
benzyl alcohol. The first method evaluated the products of toluene oxidation by
benzyl alcohol-grown cells. The second method was to identify the products of benzyl
rei
alcohol degradation. The third method was to use acetylene, 2-butyne, and 1-hexyne
as enzymes inhibitors and study the effects of these compounds to the degradation
rates of toluene oxidation and benzyl alcohol degradation. These tests were performed
as described in resting cell transformation assay for determining benzyl alcohol
degradation rates. Phenol and benzyl alcohol were compared as growth substrates for
the resting cells.
Toluene Oxidation
Previous study (Tejasen and Semprini, 2003) showed the culture did not grow
on toluene. Studies here evaluated if the culture could cometabolize toluene, and if so,
what products were formed. Cells were grown on benzyl alcohol or phenol for 8
cycles of growth (successive addition of 100 mg/I of substrate), harvested, and washed
to achieve a cell density of 2.5 mg/mi. The enrichment cells (1 ml) were added to 27ml crimped top vials containing 9 ml of media with 50 mg/i of toluene. Samples were
taken temporally and analyzed via HPLC to observe product formation.
Identification of Byproducts from Benzyl Alcohol Degradation and Toluene Oxidation
Identification of 2-hydroxy benzyl alcohol (2HBA) production from the
oxidation of benzyl alcohol by both benzyl alcohol- and phenol-grown cultures was
performed using three different HPLC methods. An EPS C-I 8 column (Alltech
Assoc. Inc., IL) was operated with two mobile phases: acetonitrile and DI water (1:3),
and methanol and 0.05 M KH2PO4, pH 3, (1:3). These two conditions resulted in
87
2HBA retention times of 2.7 and 3.9 minutes respectively, that were confirmed with
the known standard. The third method used a 250 mm
x
4.6 mm
x
5 pm Prevail
Organic Acid column (Ailtech Assoc. Inc., IL) with mobile phase of 0.71 M
acetonitrile in 25 mM KH2PO4 adjusted with phosphoric acid to pH 1.5. These
conditions resulted in a retention time of 2HBA of 29 mm. The production of 2HBA
from benzyl alcohol degradation was further confirmed by standard addition to
sample, which resulted in a single peak for all the methods described above.
Identification of o-cresol from toluene oxidation and benzaldehyde from benzyl
alcohol degradation were achieved under the two operating conditions used on the
EPS C-18 column.
Inhibition Studies
Acetylene, 2-butyne, and I -hexyne were evaluated as potential inhibitors of
benzyl alcohol degradation and TCE transformation. Resting cells grown on benzyl
alcohol or phenol were prepared similarly to cells used in the toluene tests. Benzyl
alcohol degradation and byproduct production test vials contained 9 ml of media with
50 mg/I of benzyl alcohol. TCE transformation test vials contained 5 ml of media
with an initial TCE concentration of 4 mg/I. Acetylene was added as pure gas and 2butyne and 1-hexyne were added as pure liquids. Acetylene (0.18 mg), 2-butyne (0.38
mg), and 1 -hexyne (0.57 mg) were added separately, and the vials were inverted and
shaken over night. Cells (2.5 mg) were then added to initiate the transformation tests.
Initial rates of benzyl alcohol degradation, byproduct production, and TCE
88
transformation, were determined over 1.5 hour period for cells exposed to the inhibitor
and uninhibited cells.
Yeager et al. (1999) and Kim (2000) reported that the effect of inhibitors was
dependent on exposure time (Kim, 2000; Yeager et al., 1999). In this study, the timedependent inhibition was performed to study the enzymes produced by phenol- and
benzyl alcohol-grown enrichments. I -Hexyne was used as the inhibitor and added in a
lower concentration to prevent the complete inhibition. 1 -Hexyne (0.18 mg) was
added to 27-mi vials containing 10 ml of resting cells. The vials then were inverted
and shaken at 200 rpm for 60 mm. Cells (1 ml) were removed at selected times,
washed twice with fresh media, and essayed for rates of benzyl alcohol degradation
and toluene oxidation. The rates were plotted against time of exposure and were fitted
to a first-order exponential decay curve to determine the apparent rate constant for
activity loss (k0b).
Analytical Methods
Calibration curves for all compounds were developed using external standards.
Aqueous concentrations of benzyl alcohol, phenol, toluene, 2-hydroxy benzyl alcohol,
benzaldehyde, and o-cresol were determined by injecting 50 il of aqueous sample into
a Dionex 2000i high performance liquid chromatograph (HPLC) connected to an UV
detector operated at 254 nm. Chromatographic separation was achieved using a 150
mm
x
4.6 mm
x
5 im platinum EPS C-i 8 colunm (Alitech Assoc. Inc., IL) with 0.05
89
M KH2PO4 (pH 3) and methanol (3:1 volume ratio) as a mobile phase flowing at a rate
of I mi/mm.
Headspace concentrations of toluene were determined by injecting 100
tl
of
the headspace sample into a HP5890-GC connected to flame ionization detector (FID)
at 250°C. The GC was operated at the following conditions: oven temperature, 140°C;
carrier gas (He) flow, 15 mI/mm; H2 flow to detectors, 35 mi/mm; airflow to detectors,
165 mI/mm; and FID detector makeup gas (He) flow, 15 mI/mm. Chromatographic
separation was performed with a 30 m
x
0.53 mm GSQ-PLOT column (J&W
Scientific, CA). Headspace concentrations of TCE were determined by injecting 100
tl
of the headspace sample into a HP5890-GC connected to an electron conductivity
detector (ECD) at 250°C. The GC was operated at the following conditions: oven
temperature, 140°C; carrier gas (He) flow, 15 mI/mm; and ECD makeup gas
(argon/methane mixture, 95%:5%) flow, 15 mI/mm. Chromatographic separation was
performed with a 30 m x 0.25 mm
x
1.4 im HP-624 column from Hewlett Packard
(Wilmington, DE). The total mass of chlorinated compounds in the serum bottle was
calculated by using published Henry's constants (Gossett, 1987) and mass balances
using the measured headspace concentration and the volumes of the gas and liquid
phases.
The cell concentration was determined as total suspended solids (TSS)
(American Public Health Association, 1985), using 0.2-jim membrane filter (Micro
Separation Inc., MA). Cell growth was observed by monitoring the optical density at
550 nm (OD(550)) using a Hewlett Packard 8453 UV-Visible spectrophotometer.
RESULTS
Cometabolic Transformation of TCE during the Growth on Benzyl Alcohol
The objectives of this experiment were to evaluate TCE transformation with
single and multi-additions of benzyl alcohol as a growth substrate. These initial tests
would indicate whether more effective TCE transformation resulted with prolonged
exposure to benzyl alcohol. Batch serum bottles (160 ml) were filled with 60 ml BSM
media and 100 ml of headspace air. The benzyl alcohol grown cells (1.3 mg) were
added to the bottle along with 54ig of TCE to achieve a TCE aqueous concentration
of 0.6 mg!l. Benzyl alcohol was successively added three times (6 mg, 6 mg, and 12
mg) to achieve concentrations of 100, 100, and 200 mg/i, respectively (Fig. 4.la). The
first addition of benzyl alcohol was rapidly utilized within one day and a
corresponding increase in optical density was observed. TCE however was not
transformed upon this first utilization and growth on benzyi alcohol (Fig. 4.lc). After
the second addition and utilization of benzyl alcohol, TCE was transformed after most
of benzyl alcohol was utilized, potentially indicating the inhibition of benzyl alcohol
on TCE transformation. The third addition of benzyl alcohol was rapidly consumed,
and TCE was again transformed after benzyl alcohol was reduced to low
concentrations. TCE was then added several times. In the absence of benzyl alcohol,
TCE was more rapidly transformed, likely indicating inhibition of benzyl alcohol on
TCE transformation. TCE transformation eventually slowed indicating a finite
91
capacity to transform TCE. Shown in Figure 4.lb and 4.ld are results from a single
addition of an equivalent amount of benzyl alcohol (20 mg). Multiple additions of
benzyl alcohol resulted in faster TCE transformation rates than the single addition.
The overall TCE transformed by multiple additions of benzyl alcohol was 336 jig,
while the one addition of benzyl alcohol resulted in only 36 jig of TCE transformed.
20
1.2
20
l5
09
!15
g 10
0.6
1.2
0.9-
C
10
O
0.6
>'
0
3
6
9
0
12
3
6
9
12
Time (days)
Time (days)
80
80
60
60
C
LI40
U
F-
d).
p
0
0
3
6
Time (days)
9
12
0
3
6
9
Time (days)
Figure 4.1. Cometabolic transformation of TCE during
the growth on multiple and single additions of benzyl
alcohol. Symbols: benzyl alcohol, ; TCE, X;
OD(550)4.
12
0
The increase in rate and more TCE transformed could be associated with
changes in the benzyi alcohol utilization pathway, gradual enzyme induction, or the
microbial population over time. The final optical densities indicated the
approximately equal amounts of cells were produced through single or multiple
substrate additions. The results indicated that TCE cometabolism was enhanced with
prolonged addition of benzyl alcohol.
Results of modeling studies (results not shown) indicated that the increase in
TCE transformation rates could not be explained by an increase in cell mass due to the
growth of benzyl alcohol. During the utilization of the initial addition (100 mg/i) of
benzyl alcohol (C6H5-CH2OH), benzaldehyde (C6H5-COH) accumulated. It was
degraded after the benzyl alcohol was almost completely consumed (data not shown).
The same observation was observed in reactor where 400-mg/I benzyl alcohol was
utilized. Benzaldehyde however was not observed during the degradation of the
second addition, or in a third addition of benzyi alcohol. A new byproduct was
observed during HPLC analysis, which was identified as 2-hydroxy benzyl alcohol
(2HBA). This observation indicates the enhancement in TCE transformation may be
associated with the production of 21-TIBA, and a possible change in degradation
pathway of benzyi alcohol.
TCE Transformation yields (Tv) show the improvement in TCE transformation
ability with successive exposure to benzyl alcohol. Little transformation of TCE
occurred with the first exposure, while with the second and third consumption of
93
benzyl alcohol, the T increased to 0.01 and 0.03 mg-TCE transformed per mg-benzyl
alcohol consumption respectively.
Transformation of TCE and the Degradation of Benzyl Alcohol under Resting Cells
Conditions
This study focused on determining if TCE transformation rates and
transformation capacities increased as cells were exposed to benzyl alcohol
repetitively. The benzyl alcohol cells were batch-grown with sequential addition of
100 mg/I benzyl alcohol up to 13 cycles. After each indicated cycle, cells were
harvested, washed, and assayed for benzyl alcohol degradation rates and TCE
transformation, as described above. The specific benzyl alcohol degradation rate
decreased slightly from 1.8 to 1.2 mg-benzyl alcohol/mg-cells/day over 13 cycles of
cells growth (Fig. 4.2a). Since cell mass was estimated on a total suspended solid
basis, more inert content was likely present as the growth cycles increased, which is
one possible reason for the lower specific rate as time proceeded. The specific rate of
2HBA production increased with each cycle and stabilized after 7 cycles of growth
(Fig. 4.2b). The rate of 2HBA production was about 10% of the rate of benzyl alcohol
degradation. The specific rate of TCE transformation and the TCE transformation
capacity increased with each growth cycle and also stabilized after about 7 cycles of
growth (Fig 4.2c and 4.2d). The maximum observed specific TCE transformation rate
and transformation capacity was 0.051 mg-TCE/mg-cells/day and 0.035 mg-TCE/mgcells, respectively.
2.5
0.15
)
ci
= 0]
(ID
L5
4
4
0
I
0
V
<
0.
O.5
a).
O0
b).
OF
5
0
'-s
5
Growth cycles
Growth
0.06
c
10
15
cycles
0.04
c
-
-
0.03
0.04
O.03
O.O2
A
A
A
0.02
'xx
L)
H
A
H0.Ol
H
0
0
5
Growth
10
cycles
15
d).
A
0-
0
5
Growth
10
15
cycles
Figure 4.2. The degradation of benzyl alcohol and the transformation of TCE
by benzyl alcohol-grown cells under resting cells condition: a) benzyl alcohol
(BA) degradation rate; b). 2HBA production rate; c). TCE transformation
rate; and d) TCE transformation capacity.
95
The increase in TCE transformation rates and transformation capacities with
the prolonged growth on benzyl alcohol should be noted. One possibility is that the
degradation pathway through 2HBA is responsible for TCE transformation and was
gradually induced in the successive growth on benzyl alcohol. Another is that the
microbial population gradually shifts to one that is more effective in transforming
TCE and transforms benzyl alcohol through a pathway where 21-IBA is produced.
TCE Induction
Exposure to TCE has been reported to induce higher transformation rates of
TCE (Ensign, 1996; Ryoo et al., 2001; Shingleton et al., 1998). The rates of TCE
transformation and benzyl alcohol degradation by cells previously exposed to TCE
were measured to determine if exposure to TCE during cell growth induced TCE
transformation. Cells were grown for up to 4 cycles of 100 mg/I benzyl alcohol in the
presence of 1 mg/l TCE. At the end of each cycle, cells were harvested, washed, and
assayed as described in resting cells transformation. Rates of benzyl alcohol
degradation, 2HBA production, TCE transformation, and the TCE transformation
capacity were similar to cells grown in the absence of TCE. Benzyl alcohol (BA)
degradation rates decreased from 1.9 to 1.5 mg-BA/mg-cells/day. 2HBA production
rates increased from 0 to 0.14 rng-2HBAImg-cells/day, over the 4 cycles of growth.
TCE transformation rates and transformation capacities increased from 0.003 to 0.05
mg-TCE/mg-cells/day and from 0.002 to 0.03 mg-TCE/mg-cells. This result agreed
with our previous observations (data not shown) that TCE itself was not likely
inducing the enzyme responsible for its transformation.
Microbial Communities Monitoring
To monitor for changes in microbial population, liquid samples were obtained
after each growth cycle, kept frozen at -20°C, and later analyzed by the T-RFLP
method. The results showed that there are at least two dominant microorganisms
present in the culture (Fig. 4.3). The microbial community remained stable throughout
the 13 cycles of cell growth; with two dominant peaks present in the T-RFLP patterns.
The results indicate that improvement of TCE transformation rates did not likely result
from changes in microbial community. Enzyme induction, or enhancement in enzyme
activity, was likely responsible for the improvement in TCE transformation with the
repeated exposures to benzyl alcohol.
97
GeneScw3) 3.1.2
Gel osog& Oiepey-3
After 1 growth cycle
After 2 growth cycles
Figure 4.3. TRFLP analysis of the benzyl alcohol grown culture for differ
growth cycles.
Benzyl Alcohol Degradation and TCE Transformation by Resting Cells Grown on
Phenol
Phenol was compared to benzyl alcohol as a growth substrate to induce TCE
cometabolism. Cells were batch-grown on 100 mg/I phenol up to 13 cycles of growth.
At the end of each cycle, cells were harvested, washed, and assayed for rates of TCE
transformation and benzyl alcohol degradation. Benzyl alcohol degradation by
phenol-grown cells also resulted in the production of 2HBA. Unlike the benzyl
alcohol grown cells, specific rates of benzyl alcohol degradation, 2HBA production,
98
and TCE transformation were stable during the sequential 13 growth cycles (Fig. 4.4).
The specific rates of benzyl alcohol degradation were about a factor of three lower
than rates achieved by the benzyl alcohol-grown culture. Rates of 2IUBA production
and TUE transformation by phenol-grown cells were both about a factor of two higher
than rates achieved by benzyl alcohol-grown cells. TCE transformation capacities by
phenol-grown cells were about a factor of three higher than those achieved by benzylalcohol-grown cells.
flA
-
a) >
-- a)
.2
>
!C = 0.3
0.4
Cl,
OG)
= C)
< 6) 0.1
tOE
00.1
0'0
('4
5
0
15
Growth cycles
a)
5
10
Growth cycles
15
a.' 4
u.1j
>s
Cl)
a)Q)
9.
a)
0.09
0)
bO.06
.
w
0
H
i-,-- 0.03
OE
H'
0.09
0.06
0)
0.03
d).
0
0
5
10
Growth cycles
15
0
5
10
Growth cycles
Figure 4.4. The benzyl alcohol degradation and the TCE transformation by
phenol-grown cells under resting cells condition: a) benzyl alcohol (BA)
degradation rate; b) 2HBA production rate; c) TCE transformation rate; and d)
TCE transformation capacity.
15
The TCE transformation rates were correlated to 2HBA production rates in
both benzyl alcohol- and phenol-grown cells (Figure 4.5). The correlation between
the rates of production of the byproduct and the rate of TCE transformation (r2 = 0.94)
indicates that the enzyme associated with producing 2HBA was likely responsible for
TCE transformation.
The result shows higher TCE transformation rates are achieved
by phenol grown cells and phenol is a better inducer of enzyme activity than benzyl
alcohol.
0.12
CC
+
0.1
008
II:::
E--0.Q2
0
0.1
0.2
0.3
2HBA production rate (mg-2HBA/mg-cells/day)
Figure 4.5. Correlation of 2HBA production and TCE
transformation by benzyl alcohol-grown cells (o) and
phenol-grown cells (.).
0.4
100
Benzyl alcohol degradation and TCE transformation activities were evaluated
when growth substrate was switched from phenol to benzyl alcohol. The culture was
grown on 100 mg/I phenol for two cycles, and then switched to 100 mg/I benzyl
alcohol for up to eight cycles of growth. Cells were harvested at each of the indicated
cycles and essayed for resting cells activities. After two cycles of growth on phenol,
the resting cells activities were consistent with previous experiment: benzyl alcohol
degradation rates of 0.4 day1, 2HBA production rates of 0.3 day', TCE
transformation rates of 0.1 day', and T of 0.08 mg-TCE/mg-cell (Figure 4.6). The
activities however changed when the substrate was switched to benzyl alcohol.
Benzyl alcohol degradation rates increased to 0.8 day' while 2HBA production rates,
TCE transformation rates, and T reduced to about 0.2 day', 0.08 day', and 0.04 mgTCE/mg-cell, respectively (Fig. 4.6). The 2HBA production and TCE transformation
rates were almost a factor of two higher than those achieved from cells grown on
benzyl alcohol alone. The results indicated that the mixed culture maintained high
enzyme activities when benzyl alcohol was substituted as a substrate for phenol.
10:1
I)
;;-
=
0.8
02
04
0.2
''0
b).
a).
0
3
6
9
Growth cycles
0
12
6
3
9
12
Growth cycles
0.12
tJ
0.09
1)
0
X(
x
0006
A
o.o3
0.03
ii,
1
0
3
6
Growth cycles
9
c).
d).
12
0
3
6
9
I
12
Growth cycles
Figure 4.6. The benzyl alcohol degradation and TCE transformation by resting
cells grown on the combination of phenol (first 2 growth cycles) and benzyl
alcohol (8 growth cycles): a) BA degradation rate; b) 2HBA production rate;
TCE transformation rate; and d) TCE transformation capacity.
Toluene Oxidation
The toluene oxidation rates achieved from resting cells grown on benzyl
alcohol and phenol were 0.27 and 0.71 mg-toluene/mg-cell/day, respectively. Both
benzyl alcohol- and phenol-grown cells oxidized toluene to o-cresol, which was
identified as described in analytical methods. This result agreed with the observation
of 2IIBA from benzyl alcohol degradation that an ortho-monooxygenase enzyme is
being expressed in both benzyl alcohol- and phenol-grown cells. The higher toluene
102
oxidation rates were also related to higher TCE transformation rates and 2HBA
production rates of phenol-grown cells than those of benzyl alcohol-grown cells. It is
interesting to note that the benzyl alcohol and phenol culture can not grow on toluene,
by express enzyme activity of an ortho-monooxygenase of benzyl alcohol degradation
and TCE transformation.
Inhibition Studies
Acetylene, 2-butyne, and I -hexyie inhibition were performed with benzyl
alcohol and phenol grown cells. Acetylene (0.18 mg), 2-butyne (0.38 mg), and 1hexyne (0.57 mg) were added separately into benzyl alcohol and TCE test vials, with
either benzyl alcohol or phenol grown cells (2.5 mg) added. Acetylene (1% headspace
concentration) did not affect benzyl alcohol degradation, 2HBA production, or TCE
transformation rates in both benzyl alcohol- or phenol-grown cells (Table 4.1). The
presence of 2-butyne and I -hexyne (same molar amount as acetylene) resulted in a
slight decrease in benzyl alcohol degradation rate, but completely inhibited 21-IBA
production and TCE transformation.
Table 4.1. Effects of inhibitors on the resting cells activities
Growth substrate
Inhibitors
BA degradation rate
(7.1 imol)
(mg-BA/mg-cells/day)
None
1.13
Acetylene
1.13
Benzyl alcohol
2-butyne
0.95
2HBA production rate
(mg-2HBA/mg-cells/day)
0.18
0.18
L00
1-hexyne
0.87
0.00
None
0.60
0.30
Acetylene
0.62
0.28
Phenol
2-butyne
0.40
0.00
1-hexyne
0.49
0.00
Reported values are an average of duplicates.
Initial benzyl alcohol (BA) and TCE concentrations are 50 and 4 mg/i, respectively.
TCE transformation rate
(mg-TCE/mg-cells/day)
0.054
0.053
0.00
0.00
0.100
0.093
0.00
0.00
104
The time-dependent loss of resting cells activity from the exposure to a lower
amount of I -hexyne (0.18 mg) was also determined. The exposure of I -hexyne had a
great impact on the resting cells activities (Fig. 4.7). At a 2-mm exposure time, benzyl
alcohol-grown cells lost about 30% of the initial benzyl alcohol degradation rate, 85%
of the 2HBA production rate, and 60% of the toluene oxidation rate. Phenol-grown
cells also lost about 60% of benzyl alcohol degradation, 21-IBA production, and
toluene oxidation rates within 2-mm exposure time. Previous observations found that
benzyl alcohol can be degraded in at least two pathways, through benzaldebyde and
2HBA (Fig. 4.8). The greater initial-loss of benzyl alcohol degradation rates in
phenol-grown cells than in benzyl alcohol-grown cells (Fig. 4.7a) indicates that 1hexyne blocked the ortho-monooxygenase, but not the alcohol-dehydrogenase
enzyme. The similar initial-loss in benzyl alcohol degradation rate and 2HBA
production rate in phenol grown cells (j- 60%) suggested that phenol-grown cells
exhibited mainly ortho-monooxygenase enzyme for benzyl alcohol degradation.
Prolonged exposure of I -hexyne led to additional loss of cells activities in both
phenol and benzyl alcohol grown cells (Fig. 7b). When the results were fitted with
first-order inactivation, the 95% confidence range of observed inactivation rates
in 2HBA production of phenol-grown cells (0.006
(kobs)
0.014 min') is similar to the
range achieved from benzyl alcohol-grown cells (0.004 - 0.012 min1). This result
suggests that the ortho-monooxygenase produced by phenol-grown cells might be the
same enzyme produced by benzyl alcohol-grown cells.
105
___-
100
65.338e0025Xj
b).
y =36.118e°°°9
y = 37 873
I
-O.008x
10
10
10 20 30 40 50 60 70
0
0
Exposure time (mm)
10 20 30 40 50 60 70
Exposure time (miii)
100
c).
Figure 4.7. Time course of
inactivation by I -hexyne; a)
BA degradation; b) 2HBA
production; and c) tohuene
oxidation by benzyl alcohol-
C)
00
grown cells (C>) and phenol10
grown cells (u).
0
10 20 30 40 50 60 70
o
Exposure time (miii)
Benzyl alcohol
CH2OH
L
+
OH
rxygenas
Benzaldehyde
2-hydroxy benzyl alcohol
Figure 4.8. Observed pathways of benzyl alcohol degradation.
106
The inactivation effect by I -hexyne on toluene oxidation was similar for both
benzyl alcohol-grown cells and phenol-grown cells, although the initial toluene
oxidation rate of phenol-grown cells was almost a factor of 3 higher. This result also
suggests that the toluene oxidation enzymes from both benzyl alcohol- and phenol-
grown cells are similar. Toluene oxidation rates remained constant at 35 - 40% of the
initial rate (rate without 1 -hexyne) with the prolong exposure to 1 -hexyne (up to 60
mm) for both benzyl alcohol- and phenol-grown cells (Fig. 4.7c).
The reason for this
is not known since 2HBA production activity was associated with ortho-
monooxygenase activity. Thus a similar loss in toluene and 2HBA rates would be
expected over this period.
DISCUSSION AND CONCLUSION
This is the first report to our knowledge that TCE can be aerobically
transformed by an ortho-monooxygenase expressed by a benzyl alcohol-grown
culture. TCE transformation improved with prolonged growth on benzyl alcohol.
Although TCE induction of enzymes for cometabolism has been reported (Ensign,
1996; Ryoo et al., 2001; Shingleton et al., 1998), it did not induce this culture using
the methods that were applied. The benzyl alcohol culture studied here was a highly
enriched mixed culture, with T-RFLP analysis showing two dominant peaks, with no
evidence of changing in bacterial community structure. More studies are needed to
107
determine if bacterial communities and TCE transformation ability can be maintained
with prolong incubations with TCE.
Two pathways of benzyl alcohol degradation were observed; through
benzaldehyde and 2HBA (Fig. 4.8). The increase in 2HBA production with
successive incubations on benzyl alcohol, with no major change in microbial
communities indicates a gradual induction of the 2HBA pathway. For benzyl alcoholgrown cells, the 2HBA production rate was much lower than benzyl alcohol
degradation rate. One possible explanation is that 2HBA was degraded while it was
produced. Another possibility is that benzyl alcohol was degraded through multiple
pathways. Phenol-grown cells had similar rates of benzyl alcohol degradation and
2HBA production, suggesting that an ortho-monooxygenase was the main enzyme
expressed. It is also possible that a downstream pathway differs for benzyl alcohol
and phenol grown cells, and 2HBA is being removed at slower rates in the phenolgrown cells.
The mixed culture did not grow on toluene, but both benzyl alcohol- and
phenol-grown resting cells could oxidize toluene to o-cresol, indicating the induction
of ortho-monooxygenase enzyme. The studies with toluene also showed that TCE
transformation was related to toluene oxidation. The similar effect of 1 -hexyne on
toluene oxidation rates by benzyl alcohol- and phenol-grown cells showed the similar
enzyme expressed. Yeager et al. (1999) reported the inactivation of toluene orthomonooxygenase (TOM) by long-chain alkynes, such as butynes and hexynes, but not
by acetylene (Yeager et al., 1999). Our studies with acetylene, 2-butyne, and 1-
108
hexyne showed similar results. The comparable inactivation rate constants (kobs) of 1hexyne on 2HBA production by phenol- and benzyl alcohol-grown cells support the
induction of the same enzyme. This agrees with our earlier finding that the production
of 2HBA was inhibited by the presence of phenol in both phenol- and benzyl alcoholgrown enrichments (data not shown). Future studies with pure cultures isolated from
this enrichment will permit a better understanding of the induction of this enzyme by
benzyl alcohol.
Degradation of benzyl alcohol through benzaldehyde did not result in effective
TCE transformation. TCE transformation was related to 2HBA production, indicating
that ortho-monooxygenase was responsible for TCE transformation. This result
agrees with other reports. The oxidation of methyl group of toluene, forming
benzaldehyde and benzoate, did not enhance the transformation of TCE (Duetz et al.,
1994; Nelson et al., 1988). The toluene degradation pathways that result in TCE
cometabolism are from the monooxygenase oxidation of the aromatic ring in the
ortho, meta, orpara positions, or by dioxygenase oxidation (Duetz et al., 1994;
Kaphammer et aL, 1991; Landa et al., 1994; Shields et al., 1989; Wackett and Gibson,
1988; Whited and Gibson, 1991).
Our goal was to establish and maintain an effective TCE cometabolism without
using regulated aromatic substrates, such as phenol or toluene. Since phenol yields
higher TCE transformation rates and capacities, we first grew the cells on phenol
before switching to benzyl alcohol. After switching to benzyl alcohol, the TCE
transformation rates decreased about 25% (Figure 4.6) and remained fairly stable
109
through 10 cycles of growth. The TCE transformation capacities (Ta) also were
decreased by 50%. The TCE transformation rates were still almost a factor of two
higher than those of cells grown on benzyl alcohol alone, aJthough T were
comparable. Another advantage was that the maximum TCE transformation rates, and
T were achieved sooner than with growth on benzyl alcohol alone.
In this study, TCE transformation rates were determined at a single TCE
concentration. TCE transformation kinetic tests need to be performed to determine
kinetic parameters including the K and
kma
values for both benzyl alcohol- and
phenol-grown cells. Our previous studies have shown that the benzyl alcohol grown
culture can transform both cis-dichioroethylene and vinyl chloride, which are
commonly observed anaerobic transformation products of perchioroethylene (PCE)
and TCE. Kinetic parameters of these compounds should also be determined.
In summary, benzyl alcohol was shown to be an effective substrate for the
cometabolism of TCE. The TCE transformation rate (0.051 mg-TCE/mg-cells/day) is
in the range of reported values (0.004 - 0.8 mg-TCE/mg-cells/day) (McCarty, 1997),
and the transformation capacity (0.04 mg-TCE/mg-cells) and transformation yield
(0.03 mg-TCE/mg-benzyl alcohol) indicate effective TCE transformation might be
achieved. The enhancement of TCE transformation ability with repeated growth on
benzyl alcohol is also a positive characteristic of this culture. This culture when
grown on phenol achieved a higher TCE transformation rate (0.085 mg-TCE/mg-
cells/day) and transformation capacity (0.11 mg-TCE/mg-cells). However, phenol and
toluene are regulated compounds, while the benzyl alcohol is not. The National
110
Institute of Environmental Health Sciences (NIEHS) reported that there was no
evidence of carcinogenic activity with rats and mice with high dose of benzyl alcohol
in long-term studies (National Technical Information Service, 1989). It is also
commonly used as a food-flavoring agent (National Technical Information Service,
1989). Therefore, benzyl alcohol may be suitable for use as a substrate for in-situ
aerobic cometabolism where the injection of phenol and toluene is prohibited. A
broader screening of microorganisms is needed to determine whether cometabolic
potential can be achieved on benzyl alcohol.
ACKNOWLEDGEMENTS
This research was supported by a Ph.D. scholarship from the Thai Ministry of
University Affairs (Bangkok, Thailand) and a research grant from the US Department
of Defense sponsored Strategic Environmental Research and Development Program
under agreement CU-i 127. This article has not been reviewed by the agency, and no
official endorsement should be inferred.
111
REFERENCE
American Public Health Association. (1985). Standard Methods for the Examination
of Water and Wastewater. 16th ed. APHA, New York.
Bielefeldt, A. R., Stensel, H. D., and Strand, S. E. (1995). Cometabolic Degradation of
Tce and Dee without Intermediate Toxicity. Journal of Environmental
Engineering 121(11), 791-797.
Duetz, W., Jong, C. d., Williams, P., and Andel, J. v. (1994). Competition in
Chemostat Culture between Pseudomonas Strains That Use Different Pathways
for the Degradation of Toluene. App!. Environ. MIcrobiol. 60(8), 2858-2863.
Ensign, S. A. (1996). Aliphatic and Chlorinated Alkenes and Epoxides as Inducers of
Alkene Monooxygenase and Epoxidase Activities in Xanthobacter Strain Py2.
App!. Environ. Microbiol. 62(1), 61-66.
Gossett, J. M. (1987). Measurement of Henry's Law Constants for C1 and C2
Chlorinated Hydrocarbons. Environ. Sd. Technol. 21(2), 202-208.
Hamamura, N., Page, C., Long. T., Semprini, L., and Arp, D. J. (1997). Chloroform
Cometabolism by Butane-Grown Cf8, Pseudomonas Butanovora, and
Mycobacterium Vaccae Job5 and Methane-Grown Methylosinus
Trichosporium Ob3b. App!. Environ. Microbiol. 63(9), 3607-3613.
Hopkins, G. D., Semprini, L., and McCarty, P. L. (1993). Microcosm and in Situ Field
Studies of Enhanced Biotransformation of Trichioroethylene by PhenolUtilizing Microorganisms. Appi. Environ. Microbiol. 59(7), 2277-2285.
Hyman, M. R., and Wood, P. M. (1985). Suicidal Inactiation and Labelling of
Ammonia Mono-Oxygenase by Acetylene. Biochem. J. 227, 719-725.
Kaphammer, B., Kukor, J. J., and Olsen, R. H. (1991). Cloning and Characterization
of a Novel Toluene Degradative Pathway from Pseudornonas Pickettii Pko 1. In
"Biodeterioration and Biodegradation 8" (H. W. Rossmoore, Ed.). Elsevier
Applied Science, London.
Kim, Y. (2000). PhD dissertation. Oregon State University, Corvallis, Oregon.
Landa, A. S., Sipkema, E. M., Weijma, J., Beenackers, A. A. C. M., Dolfing, J., and
Jannssen, D. B. (1994). Cometabolic Degradation of Trichioroethylene by
Pseudomonas Cepacia G4 in a Chemostat with Toluene as the Primary
Substrate. App!. Environ. Microbiol. 60(9), 3368-3374.
112
Leahy, J., Byrne, A., and Olsen, R. (1996). Comparison of Factors Influencing
Trichioroethylene Degradation by Toluene-Oxidizing Bacteria. App!. Environ.
Microbiol. 62(3), 825-833.
Liu, W.-T., Marsh, T. L., Cheng, H., and Fomey, L. J. (1997). Characterization of
Microbial Diversity by Determining Terminal Restriction Fracment Length
Polymorphisms of Genes Encoding I 6s Rma. Appi. Environ. Microbiol.
63(11), 45 16-4522.
Madigan, M. T., Martinko, J. M., and Parker, J., Eds. (1996). Brock Biology of
Microorganisms. 8 ed. Upper Saddle River, NJ: Prentice Hall.
Mallinckrodt Baker Inc. (2000). Material Safety Data Sheet (Msds): Benzyl Alcohol.,
www.jtbaker.comlmsdslbl 885.htm.
McCarty, P. L. (1997). Aerobic Cometabolism of Chlorinated Aliphatic
Hydrocarbons. In "Subsurface Restoration" (C. H. Ward, J. A. Cherry, and M.
R. Scalf, Eds.), pp. 373-395. Ann Arbor Press, Inc., Chelsea, Michigan.
McCarty, P. L., Goltz, M. N., Hopkins, G. D., Dolan, M. E., Allan, 3. P., Kawakami,
B. T., and Carrotbers, T. J. (1998). Full-Scale Evaluation of in Situ
Cometabolic Degradation of Trichioroethylene in Groundwater through
Toluene Injection. Environ. Sci. Technol. 32(1), 88-100.
McClay, K., Streger, S. H., and Steffan, R. J. (1995). Induction of Toluene Oxidation
Activity in Pseudomonas Mendocina Krl and Pseudornonas Sp. Strain Envpc5
by Chlorinated Solvents and Alkanes. App!. Environ. Microbiol. 61(9), 34793481.
Muller, R. H., and Babel, W. (1996). Growth Rate-Dependent Expression of PhenolAssimilation Pathways in Alcaligenes Eutrophus Jmp 134 the Influence of
Formate as as Auxiliary Energy Source on Phenol Conversion Characteristics.
App!. Microbiol. Biotechnol. 46(2), 156-162.
National Technical Information Service. (11989). Tr-343: Toxicology and
Carcinogenesis Studies of Benzyl Alcohol (Cas No. 100-51-6) in F3441N Rats
and B6c3f1 Mice (Gavage Studies)., Vol. 2001. National Institue of
Environmental Health Sciences, http ://ntp-server.niehs.nih.gov.
Nelson, M. J. K., Montgomery, S. 0., and Pritchard, P. H. (1988). Trichloroethylene
Metabolism by Microorganisms That Degrade Aromatic Compounds. App!.
Environ. Microbiol. 54(2), 604-606.
Ryoo, D., Shim, H., Arenghi, F. L. g., Barbieri, P., and Wood, T. K. (2001).
Tetrachioroethylene, Trichioroethylene, and Chlorinated Phenols Induce
113
Toluene-O-Xylene Monooxygenase Activity in Pseudomonas Stutzeri Ox 1.
App!. Microbial. Biotechnol. 56(3/4), 545-549.
Shields, M. S., Montgomery, S. 0., Chapman, P. J., Cuskey, S. M., and Pritchard, P.
H. (1989). Novel Pathway of Toluene Catabolism in the TrichioroethyleneDegrading Bacterium G4. App!. Environ. Microbial. 55(6), 1624-1629.
Shingleton, J. t., Applegate, B. M., Nagel, A. C., Bienkowski, P. R., and Sayler, G. S.
(1998). Induction of the Tod Operon by Trichioroethylene in Pseudomonas
Putida Tva8. AppI. Environ. Microbial. 64(12), 5049-5052.
Tejasen, S., and Semprini, L. (2003). Aerobic Biotransformation of Chlorinated
Ethenes by a Mixed Culture Grown on a Broad Range of Substrates. Wat. Res.
submitted for publication.
U.S. EPA. (2001). National Primary Drinking Water Regulations. Office of water,
U.S. Environmental Protection Agency,
http://www.bren.ucsb.edulfac_staff/fac/keller/courses/esm223/MCL.htmL
Vancheeswaran, S., Halden, R. U., Williamson, K. J., James D. Ingle, J., and
Semprini, L. (1999). Abiotic and Biological Transformation of
Tetraalkoxysilanes and Trichioroethene/Cis- I ,2-Dichloroethene Cometabolism
Driven by Tetrabutoxysilane-Degrading Microorganisms. Environ Sci.
Technol. 33(7), 1077-1085.
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2000). Characterization of an Isolate
That Uses Vinyl Chloride as a Growth Substrate under Aerobic Conditions.
Appi. Environ. Microbtol. 66(8), 3535-3542.
Vlieg, J. E. T. v. H., Koning, W. d., and Janssen, D. B. (1996). Transformation
Kinetics of Chlorinated Ethenes by Methylosinus Trichosporium Ob3b and
Detection of Unstable Epoxides by on-Line Gas Chromatography. App!.
Environ. Microbiol. 62(9), 3304-33 12.
Wackett, L. P., and Gibson, D. T. (1988). Degradation of Trichioroethylene by
Toluene Dioxygenase in Whole-Cell Studies with Pseudomonas Putida Fl.
Appi. Environ. Microbiol. 54(7), 1703-1708.
Whited, G. M., and Gibson, D. T. (1991). Toluene-4-Monooxygenase, a ThreeComponent Enzyme System That Catalyzes the Oxidation of Toluene to PCresol in Pseudomonas Mendocina Krl. J. Bacteriol. 173(9), 3010-30 16.
Yeager, C. M., Bottomley, P. J., Arp, D. J., and Hyman, M. R. (1999). Inactivation of
Toluene 2-Monooxygenase in Burkholderia Cepacia G4 by Alkynes. App!.
Environ. Microbial. 65(2), 632-639.
114
Chapter 5
KINETIC AND MODELING OF VINYL CHLORIDE,
CIS-DICHLOROETHYLENE, AND TRICHLOROETHYLENE
TRANSFORMATIONS BY AN AEROBIC ENRICHMENT
GROWN ON PHENOL OR BENZYL ALCOHOL
Sarun Tejasen and Lewis Semprini
115
ABSTRACT
Benzyl alcohol was found to be an effective substrate for the aerobic
cometabolism of trichioroethene (TCE), cis-dichloroethene (cis-DCE) and vinyl
chloride (VC). Benzyl alcohol-grown culture did not cometabolize chloroform, 1,1dichloroethane, 1,1,1 -trichloroethane, or 1,1 -dichloroethene. Phenol was compared to
benzyl alcohol as growth substrate for the resting cells kinetic studies. Degradation
kinetics of benzyl alcohol followed Monod kinetic model with specific maximum
degradation rate (k!fl) and half-saturation constant (Ks) of 1.03 mg-benzyl
alcohol/mg-cell/day and 3.94 mg/L, while degradation kinetics of phenol followed
Haldane kinetic model with kmax, Ks, and Haldane inhibition constant (Kb) of 3.45 mg-
phenol/mg-cell/day, 0.07, and 113 mg/L, respectively. The km of TCE, cis-DCE, and
VC bybenzyl alcohol-grown cells were 0.08, 1.12, and 0.12, and by phenol-grown
cells were 0.16, 1.08, and 0.33 mg-CAH/mg-cell/day, respectively. The Ks of TCE,
cis-DCE, and VC by benzyl alcohol-grown cells were 0.33, 0.77, and 0.21, and by
phenol-grown cells were 0.30, 0.57, and 0.69 mg/L, respectively. Transformation
capacities (Tc) of TCE, cis-DCE, and VC achieved by benzyl alcohol-grown cells
were 0.03, 0.22, and 0.09, and by phenol-grown cells were 0.11, 0.80, and 0.17 mg-
CAll/mg-cell, respectively. The modeling of TCE, cis-DCE, and VC transformation
using independently measured
from batch tests.
km
and Ks values matched well with observed data
116
INTRODUCTION
A number of studies have confirmed that aerobic bacteria can transform
chlorinated aliphatic hydrocarbons (CAlls) by means of cometabolic transformation to
harmless end products, such as chloride ion and carbon dioxide (McCarty, 1997;
McCarty and Semprini, 1994; Wilson and Wilson, 1985). Cometabolic
transformations are reactions that are catalyzed by existing microbial enzymes and do
not yield carbon or energy to the transforming cells (Alvarez-Cohen and Speitel, 2001;
Horvath, 1972). A growth substrate is therefore required to provide an energy source,
and induce production of the cornetabolic enzymes. Growth substrates that have been
evaluated for the CAHs cometabolism include methane (Chang and Alvarez-Cohen,
1995; Oldenhuis et al., 1989; Wilson and Wilson, 1985), propane (Tovanabootr and
Semprini, 1998; Wackett et al., 1989), phenol (Ayoubi and Harker, 1998; Folsom et
aL, 1990), and toluene (McCarty et al., 1998; Nelson et al., 1986; Wackett and Gibson,
1988). The most effective substrates for in-situ cometabolism of trichloroethylene
(TCE) that have been evaluated in controlled field tests include phenol and toluene
(Hopkins and McCarty, 1995; Hopkins et al., 1993a; McCarty et al., 1998). Toluene,
which was used as the growth substrate in the Edward Field Demonstration (McCarty
et al., 1998), is a regulated compound, with a recommended maximum contamination
level (MCL) for drinking water of I mg/L. Being a regulated compound, obtaining
regulatory approval for toluene addition might prove difficult in some cases.
117
Although there are many reports of kinetic parameters for TCE by toluene or
phenol-grown microorganisms, few data have been reported for cis-DCE and VC. cisDCE and VC are often observed as the main byproducts from the anaerobic
transformation of TCE and perchloroethylene (PCE) (McCarty and Semprini, 1994;
Semprini, 1 997a; Vogel and McCarty, 1985). In the in-situ study at Moffett field site,
high removal rates of TCE, cis-DCE and VC were observed when phenol and toluene
were added as growth substrates (Hopkins and McCarty, 1995; Hopkins et al., 1993b).
Phenol was also used as growth substrate for the in-situ bioaugmentation for the
removal of TCE, cis-DCE and VC (Steffan et al., 1999). There are limited reports of
cis-DCE and VC kinetics by aromatic degraders. Most cis-DCE and VC kinetics were
conducted with methanotrophs (Chang and Alvarez-Cohen, 1996; Dolan and McCarty,
1995; van Hyickama Vlieg et al., 1996). Bielefeldt et al. (1995) reported kinetics of
TCE and DCE by phenol-grown filamentous enrichment (Bielefeldt et al., 1995) but
not VC. However, both cis-DCE and VC have been reported as growth substrates for
aerobic microorganisms and their kinetics have been included (Coleman et al., 2002a;
Verce et al., 2000).
In previous studies, we reported an aerobic mixed culture that grew on benzyl
alcohol and cometabolized TCE, cis-dichloroethene (cis-DCE), and vinyl chloride
(VC) (Tejasen and Semprini, 2003). The mixed culture also grew on phenol and
exhibited high transformation of TCE and cis-DCE, but it did not grow on toluene.
Benzyl alcohol is a non-regulated compound, has no long-term toxicity, and is
commonly used as food flavoring agent (Mallinckrodt Baker Inc., 2000; National
118
Technical Information Service, 1989). It is also a liquid phase at room temperature
that can be easily handled like toluene. Thus benzyl alcohol may be a promising
substrate for TCE, cis-DCE, and VC cometabolism at contamination sites.
Reported here are the results of a survey of the CAHs that could be
transformed by a benzyl alcohol-grown culture as well as the determination of kinetic
constants for TCE, cis-DCE, and VC. Since our mixed culture does not grow on
toluene (Tejasen and Semprini, 2003), kinetic parameters are also determined for cells
grown on phenol to permit comparison between benzyl alcohol and phenol. The
CAl-Is included chloroform (CF), chlorinated ethanes, such as 1,1 -dichloroethane (1,1 DCA), and 1,1,1 -trichloroethane (1,1,1 -TCA), and chlorinated ethenes, such as TCE,
cis-DCE, VC, and 1,1 -dichloroethylene (1,1 -DCE). The kinetic parameters measured
include growth yield (Y), maximum degradation rate (k), half-saturation constant
(K5), and transformation capacity
(Ta). T (mg-CAHs/mg-ceils) represents the ratio of
the total amount of CAlls transformed over the cells mass added (Arp et al., 2001).
METHODS
Microbial Culture
The aerobic enrichment culture (BA-i) was obtained from a tetrabutoxysilane
(TBOS)-grown culture that was able to cometabolize cis-DCE and TCE
(Vancheeswaran et al., 1999). BA-i did not grow on toluene, but grew on phenol or
benzyl alcohol and transformed VC, cis-DCE, and TCE (Tejasen and Semprini, 2003).
119
To obtain a reproducible innoculum culture for these studies, BA-i was grown in 710mL batch serum bottles (Wheaton Glass Co., MilIville, NJ) with eight sequential
additions of 100-mg/I benzyl alcohol in 400-mL basal salt medium (BSM) described
by Vancheeswaran et al. (1999). The culture was centrifuged, washed with BSM to an
approximate concentration of 3 mg/mL, and stored at 80 °C in 7% dimethylsulfoxide
(DMSO).
Preparation of Resting Cells
Benzyl alcohol or phenol was used as growth substrate. Previous studies
showed that TCE transformation rates and transformation capacities of benzyl alcoholgrown culture increased with successive batch growth on benzyl alcohol and stabilized
after 7 cycles of growth (Tejasen et al., 2003). In order to achieve stable and
maximum transformation rates, the frozen culture was thawed and grown on eight
growth cycles with 100 mg/I sequential of benzyl alcohol additions. Upon completion
of the eight growth cycles, the liquid culture was centrifuged and rinsed with fresh
media to cell concentrations in range of 5-10 mg/mL for the resting cell tests.
Chemicals
Benzyl alcohol (99% purity) and 1,1 -DCA (99% purity) were purchased from
Acros Organics (Pittsburgh, PA). Phenol (99.1% purity) was purchased from
Mallinckrodt Baker, Inc. (Parid, KY). VC (99.5% purity), cis-DCE (99.9 % purity),
I,1-DCE (99% purity), and 1,I,I-TCA (99.5% anhydrous) were purchased from
120
Aldrich Chemical, Inc. (Milwaukee, WI). TCE (99.9 % purity) was purchased from
Fisher Scientific (Fair Lawn, NJ).
Experimental Conditions
Resting cell CAll transformation studies were conducted in 27-mL vials
(Wheaton Glass Co., Miliville, NJ) filled with 5 mL BSM, while for the substrate
degradation studies 9 mL BSM was used. The vials and reactors were autoclaved and
filled with a combination of BSM and headspace air to maintain aerobic conditions.
Benzyl alcohol and phenol were added using a high-concentration stock (40 mg!mL)
in deionized (DI) water. TCE and cis-DCE were added using a saturated solution
(20°C) in DI water. VC was added as a pure gas. Varying amounts of cell mass were
added depending on the kinetic test being performed. Bottles and vials were inverted
to prevent headspace leakage, incubated at 20°C, and shaken at 260 rpm to assure the
equilibrium condition of mass transfer between gas and aqueous phases (Kim et al.,
2002). Poisoned control bottles were prepared in the same manners as active bottles
with the addition of mercuric chloride to the concentration of 25 mg/L.
CAHs Cometabolism by Benzyl Alcohol-Grown Cells
A survey of benzyl alcohol grown cells ability to transform a broad range of
CAlls included CF, 1,1 -DCA, I ,1,1 - TCA, VC, cis-DCE, 1,1 -DCE, and TCE was
performed. Benzyl alcohol-grown resting cells (4.9 mg) were tested with each CAH
121
(0.25 tmol) separately in triplicate, representing a CAH to cell mass ratio of
approximated 0.005 mg-CAl-I/mg-cells.
Determination of Kinetic Parameters (km, Ks, Kb, and T)
Resting cells kinetic studies included km, K5 and T for VC, cis-DCE, and
TCE, and km and K values for phenol and benzyl alcohol. Haldane inhibitory
constant (Kh) was also determined for phenol degradation and TCE transformation by
benzyl alcohol-grown cells. The mass of CAH or substrate were varied to observe
rates over a wide range of concentrations in order to determine km and K values.
The mass of culture was varied from 0.3 - 10 mg to achieve 60-90% CAlls
transformation or substrate degradation within 0.5-1.5 hours. The mass ratio of cell
mass to CAll mass was maintained high (>300) to insure the cell capacity to transform
the CAl-I remained essentially unchanged during the kinetic test. The initial rates were
determined by linear regression of concentration versus time for a minimum of 5 data
points.
Yield (Y) was calculated as the ratio of total cell mass produced (rng) over
total substrate consumed (mg), using average from at least 10 independent
experiments. The transformation capacity (Ta) was determined by adding a lower
cells mass (5-10 times of CAHs mass) and monitoring the CAlls concentration until
transformation stopped. Successive additions of CAll were often required. No
endogenous energy source was added.
122
Analytical Methods
The CAH headspace concentrations were determined by gas chromatography
analysis. The temporal compound mass in the batch serum bottle was calculated using
the headspace concentration, liquid and gas volumes, and published Henry's constants
(Gossett, 1987). The high speed shaking (260 rpm) assured equilibrium partitioning
which was confirmed by Kim et al. (2002). Calibration curves for all compounds
were developed using external standards.
Headspace concentrations of VC, cis-DCE, 1,1 -dichloroethane (DCA), and
TCE were determined by injecting 100 tL of the headspace sample into a HP5 890
series gas chromatography (GC) connected to a photo ionization detector (PID),
followed by a flame ionization detector (FID) at 250°C. The GC was operated at the
following conditions: oven temperature, 140°C; carrier gas (He) flow, 15 mL/min; H2
flow to detectors, 35 mL/min; airflow to detectors, 165 mL/min; and FID detector
makeup gas (He) flow, 15 mL/min. Chromatographic separation was performed with
a 30 m
x
0.53 mm GSQ-PLOT column from J&W Scientific (Folsom, CA).
Headspace concentrations of chloroform (CF), 1,1,1 -trichloroethane (TCA), 1,1 -
dichioroethylene (1,l-DCE), and TCE were determined by injecting 100 jiL of the
headspace sample into a flP5890-GC connected to an electron conductivity detector
(ECD) at 250°C. The GC was operated at the following conditions: oven temperature,
140°C; carrier gas (He) flow, 15 mL/min; and ECD makeup gas (argonlmethane
mixture, 95%:5%) flow, 15 mL/min. Chromatographic separation was performed with
a 30 m
x
0.25 mm
x
1.4 tm HP-624 column from Hewlett Packard (Wilmington, DE).
123
Aqueous concentrations of benzyl alcohol and phenol were determined by
injecting 50 tL of aqueous sample into a Dionex 2000i high performance liquid
chromatograph (HPLC) connected to an UV detector operated at 254 nm.
Chromatographic separation was achieved using a 150 mm
x
4.6 mm
x
5 tm platinum
EPS C-18 column (Alltech Assoc. Inc., IL) with 0.05 M KH2PO4 (pH 3) and methanol
(3:1 volume ratio) as a mobile phase flowing at a rate of I mL/min.
The cell concentration was determined as total suspended solids (TSS)
(American Public Health Association, 1985), using 0.2-tm membrane filter (Micro
Separation Inc., MA).
Determination of Kinetic Parameters
For all compounds, except phenol, initial rates and concentrations were plotted
according to Michaelis-Menten enzyme kinetics:
I
M
C
=
.('
(5.1)
K+C
where Mx is cells mass added (mg); r, initial rates of degradation or transformation of
the interest compound (mg/day); km, maximum specific rate of the compound
degradation or transformation (mg-compound/mg-cells/day); Ks, half-saturation
concentration (mg/L); C, the initial aqueous concentration (mg/L).
The values of aqueous concentration
(Cag) and r of CAHs were calculated
from temporal mass at equilibrium condition as described above:
Caq
;=HccCg
(5.2)
124
ForCAHs,
dM
r
=(V+V.H)0
C
(5.3)
di'
For phenol and benzyl alcohol, r
where
Caq
dM
dCaq
C
di''
(54)
di'
is aqueous concentration (mgIL); VI, liquid volume (L); Vg, headspace
volume (L); M, CAH mass (mg); t, time (day).
For TCE transformation by benzyl alcohol-grown cells and phenol
degradation, initial rates and concentrations were plotted according to a Haldane
kinetic model (Bailey and 011is, 1986), which incorporates inhibition effects at high
concentrations:
1
M
=
kmaxC
(5.5)r
Kj
where Kh is the Haldane inhibitory constant (mg/L).
The values of kmax, Ks, and Kh were determined by fitting the specific initial
rates and concentrations to models using a non-linear regression analysis by a
statistical package of SPSS (Chicago, IL), as described by Kim et al. (2002).
Modeling of CAHs Transformation
The values of T, km, and K were used to model the transformation of CAHs
by resting cells grown on either phenol or benzyl alcohol. Another required variable is
the active cells mass (Mx). The modeling approach applied was that described by
Chang and Alvarez-Cohen (1995) and Anderson and McCarty (1996) where the rate
125
of change of cell mass presented in Eq. 5.6 for cell growth including terms of cell
inactivation due to product toxicity and cell decay:
dM
=Yr1rbM
(5f1)
where r is the rate of cell production (mg-cells/d); Y, growth yield (mg-cells/mgsubstrate); rg, substrate consumption rate (mg-substrate/d); r, CAMs transformation
rate (mg-CAHs/d); b, cellular decay (1/d); Mx, active cells mass (mg).
Under the resting cell conditions when no substrate is present, the growth term
(Yrg) is equal to zero. Also, in our batch reactor system, the product toxicity is
substantially greater than cell decay; cell decay in equation 3 can be neglected
(Alvarez-Cohen and Speitel, 2001). The equations used for the modeling are:
CAHs transformation,
=
di
Changes in active cells mass,
(k+VgHcr)
dX= 1
di
XC
K5+C
kiax
-
(Vi+VgHcc) dC
.
T
(57)
(5.8)
di
V,
where X is cell concentration (mg/L).
The modeling was performed according to equation 7 and 8 by using both
spreadsheet program (Microsoft® Excel 2002) and Aquasirn (EAWAG, Switzerland).
The model sensitivity for T was also performed using values of T + 110%. The fit of
the model to experimental data was evaluated based on the standard error of estimate
(SEE) (Holman, 2001):
I
(5.9)
n-2
J
126
where SSE is standard error of estimate; C1, experimental data;
,
model prediction;
and n, number of data points.
RESULTS
CAHs Cometabolism by Benzyl Alcohol-Grown Cells
The results of the survey of benzyl alcohol-grown cells ability to transform a
broad range of CAHs are shown in Figure 5.11. Benzyl alcohol-grown cells were able
to transform VC, cis-DCE, and TCE, with VC was most rapidly transformed, followed
by TCE and then cis-DCE. Comparison of active bottles with controls (not shown)
shows no significant transformation of CF, 1,1 -DCA, 1,1,1 -TCA, or 1,1 -DCE at the
ratio of cell mass to CAll ratio used of approximately 0.005 mg-CAH/mg-cells.
1.20
0
0)
0.60
I
0.40
C.
0.20
I.e
0
20
40
60
80
Time (hr)
Figure 5.1. Resting cell transformation of CAlls by benzyl alcohol
grown cells. Symbols: 0, VC; LI, TCE; A, cis-DCE; 4, CF; A, 1,1DCE; , DCA; s, TCA. Initial CAH mass (M0) 0.25 j.xmol.
127
Substrate Utilization Kinetics
The specific rates of phenol and benzyl alcohol utilizations versus
concentrations are presented in Figure 5.2. The kmax of phenol was about a factor of
three higher than
km
of benzyl alcohol (Table 5.1). The specific utilization rates of
phenol were well fit by a Haldane kinetic model (Eq. 5.5) with rates of phenol
utilization decreasing for phenol concentrations above 10 mg/L. The statistical
analysis of phenol degradation rates was also performed at low phenol concentrations
(<5 mg/L) fitted to a Monod kinetic (equation 5.1). Similar km and K5 were
achieved (95% confidence interval of 3.1
3.6 mg-phenol/mg-cell/day and 0.03
0.13 mgIL, respectively), indicating that Kh did not affect the estimation of
kmax
and
K. Maximum rates of benzyl alcohol utilization were lower, and there was no
evidence of inhibition at higher concentrations up to 180 mg/L (data not shown).
Statistical analysis of benzyl alcohol degradation at low concentrations (<35 mgJL)
showed similar
kmax
and K5 (95% confidence interval of 0.95
alcohol/mg-cell/day and 3.4
8.3 mg/L), suggesting that the
not biased by high benzyl alcohol concentrations.
1.25 mg-benzyl
km
and K values were
128
-..
4
C',
.;
3.5
9
a,
2.5
C',
2
C)
a)
E 1.5
c
7-
1
0
20
40
60
80
100
Substrate Concentration (mg/L)
Figure 5.2. Degradation kinetic of phenol (S) and benzyl
alcohol (0). Solid and dotted lines represent Haldane and Monod
kinetic fits to the data, respectively.
Table 5.1. Summary of kinetic parameters from the resting cells studies
Growth substrate
Y
Phenol
Phenol*
Benzyl alcohol
Benzylalcohol**
0.55 + 0.02
Growth substrate
CAlls
Phenol
Benzyl alcohol
0.53 + 0.02
TCE
cis-DCE
VC
TCE
cis-DCE
VC
kmax
K5
(mg.-CAWmg-cefls/d)
(mg/L)
(mg!L)
3.45 ± 0.11
0.07 ± 0.02
112.6 ± 20.4
3.35±0.13
0.08+0.03
1.06± 0.03
5.29 ± 0.68
1.10± 0.08
5.85± 1.23
km,
K5
(mgCAWmg-cell/d)
(mg/L)
(mg-CAll/mg-cell)
0.16±0.01
0.30±0.06
0.11±0.002
1.08 ± 0.07
0.33 ± 0.02
0.08 ± 0.004
1.12 ± 0.12
0.12 + 0.003
0.57 ± 0.12
0.69 ± 0.08
0.33 ± 0.05
0.77 ± 0.18
0.22 ± 0.02
0.80 ±
0.17 ±
0.03 ±
0.22 ±
0.09 ±
Values shown in average ± I standard deviation
* Phenol fit to phenol concentration below 5 mg!L.
** Benzyl alcohol fit to benzyl alcohol concentration below 35 mgIL
0.004
0.0 14
0.003
0.004
0.004
129
TCE, cis-DCE, and VC Transformation Kinetics
Figure 5.3 shows the specific rates of TCE transformation versus concentration
for phenol- or benzyl alcohol-grown cells. The K5 values for TCE transformation by
phenol- and benzyl alcohol-growr cells were essentially equal, statistical analysis
showed the 95% confidence interval of 0.18
0.43 and 0.21 - 0.44 mg/L,
respectively. The km for TCE by phenol-grown cells was a factor of two higher than
by that of benzyl alcohol-grown cells (Table 5.1). Kinetic tests with benzyl alcoholgrown cells were performed over a higher concentration range than phenol-grown
cells. TCE transformation followed Haldane kinetics (Figure 5.4). The 95%
confidence intervals of K, km, and Kh achieved by benzyl alcohol-grown cells were
0.1
0.2 mg/L, 0.4
1.1 mg-TCE/mg-cell/d, and 1.4
3.4 mg/L, respectively.
0.16
0.14
0
9
0.12
C)
w
C-)
0.08
E
0
C.)
0°
/
o 0.06
0
0
0.04
-1J
9-
/10-...-
C.)
a 0.02
Cl)
0
0
0.5
1
1.5
TCE conc. (mgIL)
Figure 5.3. TCE Transformation by resting cells grown on
benzyl alcohol (LI) and phenol (i). Solid and dotted lines
represent Monod model fits to the data.
2
130
0
0.07
0.06
0.05
I-;.
0.04
a, 0.03
0.02
0.01
0.
sJ
I
10
15
ICE (mgIL)
Figure 5.4. TCE transformation by resting cells grown on benzyl
alcohol. Line represents Haldane model fits to the data.
The specific rates for cis-DCE transformation versus concentrations are shown
in Figure 5.5. Comparable kinetics were achieved (Table 5.1), the 95% confidence
interval Of kmac achieved by phenol- and benzyl alcohol-grown cells were 1.04
1.40
and 0.96 1.5 mg-cis-DCE/mg-cells!day, and those of K5 were 0.37 1.2 and 0.340.79 mg/L, respectively. The km values for cis-DCE transformation were about an
order of magnitude higher than those for TCE transformation for both phenol- and
benzyl alcohol-grown cells.
131
1
a
U) 09
a)
9 0.8
a,
0.7
LU
C)
00.5
0)
E 0.4
P-3
2 0.2
o 0.1
a.
U)
0
0
1
2
3
4
cis-DCE conc. (mg/L)
Figure 5.5. cis-DCE Transformation by resting cells grown on
benzyl alcohol (A) and phenol (A). Solid and dotted lines
represent Monod model fits to the data of phenol- and benzyl
alcohol-grown cells, respectively.
The specific rates for VC transformation versus concentration are shown in
Figure 5.6. Both
km
and K5 for VC of phenol-grown cells were about a factor of
three higher than benzyl alcohol-grown cells (Table 5.1). The 95% confidence
interval showed different ranges of K5 achieved by phenol-grown cells (0.51 0.86
mg/L) and by benzyl alcohol-grown cells (0.18-0.26 mgIL). The km values for VC
were lower than those for cis-DCE, but higher than those for TCE transformations by
both phenol- and benzyl alcohol-grown cells. The maximum specific rates for VC
transformation by benzyl alcohol-grown cells remained about 0.12 mg-VC/mgcell/day at VC concentrations up to 5 mg/L.
132
0.3
0.25
0
C;)
02
0
>
8)
0.15
0.05
a.
Cl)
0
0.5
1.5
2
2.5
3
VC conc. (mg/L)
Figure 5.6. VC Transformation by resting cells grown on
benzy! alcohol (0) and phenol (+). Solid and dotted lines
represent Monod model fits to the data.
Transformation capacity (Tn)
Transformation capacity values for TCE, cis-DCE, and VC by both phenoland benzyl alcohol-grown cells are presented in Table 5.1. Transformation capacity
values achieved by phenol-grown cells were factors of three higher than benzyl
alcohol-grown cells, for both TCE and cis-DCE, and a factor of two higher for VC. In
both enrichments, transformation of cis-DCE had the highest T, about 8 times that of
TCE and 3-4 times that of VC. The similarity in the ratio of T between CAlls also
supported the idea of similar enzymes expression in phenol-grown cells and benzyl
alcohol-grown cells. Table 5.2 showed ratios of
km
over T from independent
measures for CAHs transformations. Similar values of these ratios suggested that
133
higher T was correlated with higher km, with the exception of cis-DCE
transformation by benzyl alcohol-grown cells, where the ratio was about a factor of
two higher than the others.
Table 5.2. Ratio of kmax over T for CAHs transformation
kmiTc (l/d)
Growth substrate
TCE
Phenol
Benzyl alcohol
1.5
2.4
1
{
cis-DCE
VC
1.4
1.9
1.3
4.7
1
j
Modeling of CAlls transformation
The model for CAll transformation by phenol and benzyl alcohol cells was
compared to the independent experimental data from the study of transformation
capacity
(Ta)
(Figure 5.7 and 5.8). The models were performed using equations 5.7
and 5.8. A sensitivity analysis to the T was also performed. The fit of the model to
the experimental data was evaluated using equation 9. Model input parameters for
equations 5.7 and 5.8 in Table 5.3, along with the kinetic values in Table 5.1 were
used to simulate these temporal results. The input for T values were calculated from
these tests and were not independently measured. These experiments were done for
resting cells conditions, without extra substrate or energy added, so the growth term
(Yrg) was not included. The product toxicity in the batch reactor was assumed to be
much greater than cell decay, so the decay term (bMx) was also not included. The
initial concentrations of TCE, cis-DCE, or VC were about an order of magnitude
higher than K, in order to cover the range of concentrations effected by Monod
kinetics. Initial cell mass was also independently determined.
134
' 4
SEE = 0.093
( (A
tiO
0.03
0
0
0
0.02
0H
0.01
0
0
0
1
2
3
Time (d)
4
'-S
SEE = 5.52
O.3
( 30
0
O.2
0
0.1
0
10
00
0
fl
0
1
2
3
4
Time (d)
10
0.2
-
0.15
0
0
0
0.1
02
>
0.05
0
E
0
>
0
0
1
2
3
4
Time (d)
Figure 5.7. Modeling of TCE, cis-DCE, and VC transformation by phenolgrown resting cells. Solid lines represent model simulation and data represent
mean and standard deviation of triplicate values. Dotted lines represent a model
sensitivity analysis to T ± 10%. The initial cell mass (M0) were: 0.44, 0.76,
and 2.00 mg for TCE, cis-DCE, and VC tests, respectively.
135
6
SEE= 0.33!
b05
C.)
0
02
EE
01
H
0
I
0
0
2
1
3
4
5
6
Time (d)
8
0.04
SEE = 0.40
0.03
0.02
U
0.01
C.)
0
0
0.5
1
1.5
2
Time (d)
15
'J.
A
Aio
'3.
C.)
05
0
(I)
O.lo
U
0
0
0
1
2
3
Time (d)
4
5
Figure 5.8. Modeling of TCE, cis-DCE, and VC transformation by benzyl
alcohol-grown resting cells. Solid lines represent model simulation and
data represent mean and standard deviation of triplicate values. Dotted
lines represent a model sensitivity analysis to Tc ± 10%. The initial cell
mass (MX0) were: 2.7, 0.3, and 13.3 mg for TCE, cis-DCE, and VC tests,
respectively.
136
Table 5.3. Experimental conditions used for modeling of resting cells CAHs transformation
Growth substrate
Phenol
CAHs
TCE
cis-DCE
VC
TCE
cis-DCE
VC
VI
(ml)
Vg (ml)
Mo (mg)
21.9
21.9
21.8
0.296
0.44
0.122
0.76
5.2
0.903
1.99
Benzyl alcohol
6
21
0.296
2.7
3.1
23.9
0.122
0.303
7
20
0.903
13.33
All experiments were performed at 20°C. Henry's constants (H) were calculated
according to Gossett (1987). V1, liquid volume; Vg, headspace volume; Mx0, initial
cell mass.
5.1
5.1
Under the experimental conditions, the models matched well the observed
data. The significance of decay term was evaluated with model of TCE
transformation by benzyl alcohol-grown cells, which had the longest experimental
period (6 days). A conventional decay of 0.06 l7d (Metcalf& Eddy Inc., 1991) was
applied, and the model showed little differences (4%). The calculated SEE values
are about 3
8 % of initial concentrations, except for the cis-DCE transformation by
phenol-grown culture (14%). The model of cis-DCE transformation by phenolgrown cells did not match with the observed data during 0
I day, but it matched well
with later data after 1.5 day. The reason for this is not known. If the errors came from
the effects of high cis-DCE concentrations, they would also apply to the model during
second addition of cis-DCE as well. Another possibility would be mass transfer
limitation because of low }1 of cis-DCE. However, the model of cis-DCE
transformation by benzyl alcohol-grown cells matched well with the observed data
137
under the same environmental conditions (temperature, volume ratio of liquid to
headspace, and shaker speed).
A sensitivity analysis to the T is shown in Figures 5.7 and 5.8. The results are
shown to be very sensitive to the T, with 10% variation covering the range of
observations at the latter stage of the tests. This result indicated that the reported T
matched well the observation. The greater error variation in VC transformation by
benzyl alcohol-grown cells was a result from the higher VC transformation capacity
with the amount of cell mass used. The ratios of the error over total VC transformed
by benzyl alcohol- and phenol-grown cells were about the same (0.004). Overall the
simulation shows a good fit with the independently measured parameters.
DISCUSSION
The benzyl alcohol grown culture was able to transform three chlorinated
ethenes (TCE, cis-DCE, and VC), but not chlorinated ethanes (DCA and TCA), CF, or
1,1 -DCE. Chlorinated ethenes are generally more effectively cometabolized by
microbes grown on aromatic hydrocarbons, such as phenol or toluene, while
chlorinated ethanes are more effectively transformed when microbes grown on
saturated hydrocarbons, such as methane, propane, or butane (Semprini, 1997b). The
ability of this culture to transform chlorinated ethenes, but not TCA, DCA, CF, or 1,1 DCE is consistent with previous reported of phenol- and toluene-grown mixed culture
(Bielefeldt etal., 1995; Chang and Alvarez-Cohen, 1995). Our previous report
indicated that the enzyme expressed by benzyl alcohol enrichment is similar to ortho-
138
monooxygenase expressed by toluene-grown culture or a culture expressing phenol
hydroxylase enzymes (Futamata et al., 2001 b; Tejasen et aL, 2003; Yeager et al.,
1999). In a survey of microorganisms in Moffett Field site, several TCE
cometabolizing strains also grew on phenol, not on toluene, and exhibited enzymes
similar to toluene ortho-monooxygenase (Fries et al., 1997). The characteristics of
cultures grown on phenol, but not on toluene has also been reported with Rhodococcus
strain R-22 expressing propane monooxygenase (Fairlee et al., 1997) and
Xanthobactor
strain Py2 expressing alkene monooxygenase (Zhou et al., 1999). Our
culture however does not grow on propane or ethylene.
This mixed culture has a growth yield (Y) on phenol similar to other reported
cultures (0.5-0.6 mg-cell/mg-phenol) (Chang and Alvarez-Cohen, 1995; Hill and
Robinson, 1975; Shurtliffet al., 1996). A comparison of phenol degradation kinetics
is presented in Table 5.4. These microorganisms can be grouped by K values; low
(<0.1 mg/L), intermediate (0.1
I mg/L), and high (1
11 mg/L). Our culture is in
the low-Ks group. Futamata et al. (2001a, b) reported strains P2, P6, and P8 as highKs microorganisms, however, the K of these strains are comparable to intermediateK5 group in Table 5.4.
The
km
of our culture is within the reported values (1
11
mg-phenol/mg-cell/day) of other cultures, with a few exceptions of strains R5 and G4
where km
are 20
30 mg-phenol/mg-cell/day. Haldane kinetics have been reported
for many cultures growing on phenol. Our culture has an intermediate Kh (113 mg/L)
compared to others (11
900 mgJL), indicating a moderate tolerance to toxicity of
high phenol concentrations.
139
Table 5.4. Comparison of microorganisms and phenol degradation kinetics
Microorganisms
km
K5
1/day mgIL
Reference
Kh
mg/L
This study
Mixed culture
3.5
0.07
113
Burkholderia cepacia El
3.4
0.05
20
Comamonas testosteroni R2
8.8
0.08
23
C. testosteroni E6
2.7
0.01
16
C. testosteroni R5
18.3
0.04
11
Pseudomonas putida P-2
0.9
0.39
291
P. putidaP-6
P. putida P-8
P. putida P35X
Actinomycetes
1.1
0.37
43
1.6
0.50
649
6.2
0.34
874
10.2
0.34
>15
Raistonia eutropha
8.6
0.94
B. cepacia G4
31.5
0.80
42
(Folsom et al., 1990)
(Hill and Robinson,
P. putida
11.5
<1
470
1975)
R. eutropha
9.8
2
350
(Leonard et al., 1999)
Mixed culture
9.3
<3.3
(Shurtliffet al., 1996)
(Muller and Babel,
R.eutrophaJMPl34
9.4
5.55
1995)
Mixed culture
6.0
11
188
-
338
348
(Futamata et al.,
2001a, b)
(Lee et al., 2000)
(Leonard and
Lindley, 1999)
(Goudar et al., 2000)
The Ks of phenol was about a factor of fifty lower than benzyl alcohol,
indicating higher affinity for the enzyme and it likely being a better inducer of the
enzyme. Correlation of K5 for phenol degradation and TCE transformation was
reported with strains El, R2, E6, and R5 (Futamata et al., 200la, b). These strains
also have the first-order TCE transformation rates (k1) of 0.5
0.8 L!mglday, which
are comparable to our phenol-grown culture (0.53 L/mg/day), presented in Table 5.5.
Phenol was observed to inhibit benzyl alcohol degradation in our previous studies
2
I
VC
-
VC
II
TCE
eis-DCE
VC
TCE
cis-DCE
VC
TCE
cis-DCE
VC
TCE
cis-DCE
ICE
TCE
TCE
25
1.9-6.3
58
5-25
10-80
1.0-3.1
0-80
0-80
0-200
0.13-2.0
0.2-3.5
0.1-2.5
0.13-2.0
0.2-1.6
0,1.5.0
0-59
0-34
0-22
0.2-0.5
0-80
0-80
ICE
TCE
TCE
1-25
2.1-210
0.17
1-30
1-30
0.23
0.026
0.96
0.18
0.3-1.5
0.88
8.3
5.6
0,12
9.6
1.12
0,25
0,19
0.10
0.16
1.08
0.33
0.084
0.33
0,15
0,23
0.13
2,14
0.21
1.5
095
(mg/mg-d)'
0.66-6.6
TCE
TCE
TCE
TCE
TCE
TCE
CAHs
1.1
0.073
0.016
0.20
0.16
1.6
6.2
3.0
3,5
3.1
1.6
4.8
5.7
1.6
2.7
0.6
1,5
0.5
0.3
1.9
0.6
0.8
0.06
0.5
0.6
0.5
0.80
0.17
0.03
0.22
0.09
0.12
0.18
0.38
0.3-0.5
0.15
0.11
0,08
0.59
(Verce et al., 2001)
(Verce et al., 2000)
1992)
(Hartmans and Bont,
(Coleman et at., 2002)
(Bielefeldt et at., 1995)
(Chang and AlvarezCohen, 1996)
This study
This study
(Futamata et at., 2001)
(Shurtliffet al., 1996)
(Leeetal., 2000)
1998)
(Folsom et al,, 1990)
(Chang and AlvarezCohen, 1995)
(Ayoubi and Harker,
0.03
(Landaetal., 1994)
Reference
1.2
0.03
0.007
0.03
Tc
(mg/mg)1
3.8
0.02
0.10
0.03
(L/mg-d)'
k1
n,th'z
0.21
0.30
0.57
0,69
0.33
0.77
1.5
15.8
4.6
0,14
3.0
4.2
11
83
0.80
0.39
8,64
2.04
(mg/L)
CAL-Is trin.cfnrmtinn
wviss tpuru in mg ury ceu mass; units conversions assumed dry cell mass is 50% protein.
Reported kinetics were fitted to a Haldane model with K11 = 97 mg/L.
Pseudomonas
aeruginosa MFI
P. aeruginosa DLI
aurumLI
1
cis-DCE
13-proteobacterium
JS666
VC
Phenol
Mixedcujture
(filamentous)
Mycobacterium
Methane
Benzyl
alcohol
Phenol
Phenol
Phenol
Phenol
Phenol
Phenol
Phenol
Phenol
Toluene
Phenol
Toluene
Phenol
Phenol
Growth
Substrate
Mixed culture
Mixed culture
Mixed culture
P. putidaP3sX2
Mixed culture
Actinomycetes
B. cepacia El
Comamonas
testosteroni R2
C. testosteroni E6
C. testosteroni R5
Ralstoniaeutropha
Burkholderja
cepacia 04
Mixed culture
Microorganism
CAl-Is
Table 5.5. Comparison of microorganisms and
0
(,.
CD
-
0-
s1
-
-.
-
C
C
-I
CD
-
C
CD
CD
-
II
C
0
CD
.,<
CD
CD
141
Reported here are the first studies where km, K5, and T are determined for
TCE, cis-DCE, and VC by an enrichment grown on aromatic substrates. Table 5.5
permits a comparison with other reported values. Among toluene- and phenol-grown
cultures, our culture when grown on phenol had km value for TCE transformation
about an order of magnitude lower than those ofF. cepacia G4 and R. eutropha
JMP 134, and the
kmax
for benzyl alcohol-grown cells is half that of phenol-grown
cells. The K values for TCE transformation are in the same range of phenol-grown P.
cepacia G4. The TCE transformation capacities of our culture when grown on either
phenol or benzyl alcohol are among the highest values reported, except phenol-grown
filamentous and actinomycetes reported by Bielefeldt et al. (1995) and Lee et al.
(2000). This might indicate a tolerance of this mixed culture to TCE transformation
toxicity.
The
km
and K5 for cis-DCE transformation are similar when the mixed
culture is grown on either phenol or benzyl alcohol. To our knowledge there are no
earlier reports of both
km,,
and K values for cis-DCE of cultures grown on aromatic
substrates. The km of BA-i is in the range of values reported by phenol-grown
mixed culture (Bielefeldt et al., 1995); although it is almost an order of magnitude
lower than those of methanotrophs (Chang and Alvarez-Cohen, 1996). The K5 for cisDCE transformation of this culture is about a factor of 5 lower than those of
methanotrophs. The cis-DCE T of BA-I when grown on benzyl alcohol is similar to
previous reports with phenol utilizing (Bielefeldt et al., 1995) and methane utilizing
cultures (Chang and Alvarez-Cohen, 1996), while the T is much higher when BA-I is
142
grown on phenol. Both phenol- and benzyl alcohol-grown cultures have a
comparative low km, but very high T. Thus, high T for cis-DCE by phenol-grown
cells indicates low transformation product toxicity. Whether the cells can obtain
energy from this transformation is not known.
Coleman Ct al. (2002) recently isolated a f3-proteobacterium JS666 grown on
cis-DCE as a carbon and energy source. This discovery brings up the possibility that
other microorganisms may have ability to obtain energy for cis-DCE utilization, and
this needs to be examined further with our culture. Our data to date do not show
evidence for growth on cis-DCE. One similar growth characteristic of this mixed
culture and the strain JS666 is that both do not grow on ethylene.
Reported here is the first determination Of km and Ks values of VC
cometabolism by microbes grown on an aromatic compound. High removal of VC
was observed along with TCE and cis-DCE when phenol and toluene were injected as
substrate in the in-situ study at Moffett field site (Hopkins and McCarty, 1995;
Hopkins et al., 1993b). Bioaugmentation of phenol-grown cells also resulted in high
TCE, cis-DCE, and VC removal rates (Steffan et al., 1999). Growth substrates that
induce VC cometabolism include ethene, ethane, methane, propane, propylene,
isoprene, and ammonia (Verce et al., 2000). A few microorganisms have been
reported of ability to use VC as a sole growth substrate (Coleman et al., 2002b;
Hartmans and Bont, 1992; Verce et al., 2000, 2001). BA-I has a kmax for VC
transformation in the lower range of reported by these VC-grown strains. The K5 for
VC of our culture is in the higher range of values reported for VC-grown strains
143
(Hartmans and Bont, 1992; Verce et al., 2000, 2001). When VC concentrations are
lower than K, then first-order kinetics apply and the first order rate constant, k1
kmIK5 (L/mglday), is important. The k1 values for phenol- and benzyl alcohol-grown
cells are almost identical (0.5 L/mg/day), indicating similar rates of VC
transformation would be achieved with growth of the culture on either substrate.
BA-I is less effective towards VC transformation than cis-DCE. The
transformation capacities for VC are about a factor of 4 lower than those for cis-DCE,
and
km
values are about an order of magnitude lower. The Ks values for TCE and
cis-DCE transformation are similar for phenol- and benzyl alcohol-grown cells, while
the Ks value for VC is significantly higher for the phenol-grown cells.
CAHs transformation by resting cells fit well with equations 7 and 8, except
for cis-DCE transformation by phenol-grown culture. The cis-DCE transformation
rate was lower than predicted. This might be because of the toxicity from high cis-
DCE concentration. The toxicity of higher CAHs concentrations resulted in lower
rates of CAl-Is transformation is frequently observed (Alvarex-Cohen and McCarty,
1991; Chang and Alvarez-Cohen, 1995; Wackett and Householder, 1989; Yeager et
al., 2001).
In this study phenol-grown cells achieved better TCE, cis-DCE, and VC
transformations than benzyl alcohol-grown cells. This result indicates that phenol is
more effective than benzyl alcohol in being growth substrate in the transformation of
TCE, cis-DCE, and VC. However, phenol is a regulated compound (OSHA, ACGIH,
and NIOSH) and listed as material extremely hazardous to health (National
144
Toxicology Program, 2001), while benzyl alcohol is a non-regulated compound and
listed as moderate hazardous (National Toxicology Program, 2002). Therefore, benzyl
alcohol has potential in the application of these CAHs cometabolism treatment. More
work is needed on evaluating benzyl alcohol as a growth substrate for well studied
TCE cometabolizing microorganisms, such as Pseudomonas cepacia G4. Isolation of
pure cultures from this mixed culture is also needed.
145
REFERENCE
Alvarez-Cohen, L., and McCarty, P. L. (1991). A Cometabolic Biotransformation
Model for Halogenated Aliphatic Compounds Exhibiting Product Toxicity.
Environ. Sci. Technol. 25(8), 1381-1387.
Alvarez-Cohen, L., and Speitel, G. E. J. (2001). Kinetics of Aerobic Cometabolism of
Chlorinated Solvents. Biodegradation 12(2), 105-126.
American Public Health Association. (1985). Standard Methods for the Examination
of Water and Wastewater. 16th ed. APHA, New York.
Arp, D. J., Yeager, C. M., and Hyman, M. R. (2001). Molecular and Cellular
Fundamentals of Aerobic Cometabolism of Trichioroethylene. Biodegradation
12(2), 81-103.
Ayoubi, P. J., and Harker, A. R. (1998). Whole-Cell Kinetics of Trichloroethylene
Degradation by Phenol Hydroxylase in a Rastonia Eutropha Jmp 134
Derivative. App!. Environ. Microbiol. 64(11), 43 53-4356.
Bailey, J., and 011is, D. (1986). "Biochemical Engineering Fundamentals." McGrawHill Plublishing Co., New York.
Bielefeldt, A. R., Stensel, H. D., and Strand, S. E. (1995). Cornetabolic Degradation of
Tee and Dee without Intermediate Toxicity. Journal of Environmental
Engineering 121(11), 791-797.
Chang, H.-L., and Alvarez-Cohen, L. (1995). Transformation Capacities of
Chlorinated Organics by Mixed Cultures Enriched on Methane, Propane,
Toluene, or Phenol. Biotechnol. Bioeng. 45, 440-449.
Chang, H.-L., and Alvarez-Cohen, L. (1996). Biodegradation of Individual and
Multiple Chlorinated Aliphatics by Mixed and Pure Methane Oxidizing
Cultures. App!. Environ. Microbiol. 62(9), 3371-3377.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain, J. C. (2002a).
Biodegradation of Cis-Dichioroethene as the Sole Carbon Source by a BProteobacterium. App!. Environ. Microbiol. 68(6), 2726-2730.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain, J. C. (2002b). Phylogenetic
and Kinetic Diversity of Aerobic Vinyl Chloride-Assimilating Bacteria from
Contaminated Sites. Appi. Environ. Microbiol. 68(12), 6162-6171.
Dolan, M. E., and McCarty, P. L. (1995). Methanotrophic Chloroethene
Transformation Capacities and 1,1 -Dichloroethene Transformation in Aquifer
Samples. Environ. Sci. Technol. 29, 1892-1897.
146
Fairlee, J. r., Burback, B. L., and Perry, J. J. (1997). Biodegradation of Groundwater
Pollutants by a Combined Culture of Mycobacterium Vaccae and a
Rhodococcus Sp. Can. J. Microbiol. 43, 841-846.
Folsom, B. R., Chapman, P. J., and Pritchard, P. H. (1990). Phenol and
Trichioroethylene Degradation by Pseudomonas Cepacia G4: Kinetics and
Interactions between Substrates. Appi. Environ. Micro biol. 56(5), 1279-1285.
Fries, M. R., Fomey, L. J., and Tiedje, J. M. (1997). Phenol- and Toluene-Degrading
Microbial Populations from an Aquifer in Which Successful Trichloroethene
Cometabolism Occurred. Appi. Environ. Microbiol. 63(4), 1523-1530.
Futamata, H., Harayama, S., and Watanabe, K. (2001 a). Diversity in Kinetics of
Trichloroethylene-Degrading Activities Exhibited by Phenol-Degrading
Bacteria. App!. Microbiol. BIotechnol. 55, 248-253.
Futamata, H., Harayama, S., and Watanabe, K. (2001b). Group-Specific Monitoring of
Phenol Hydroxylase Genes for a Functional Assessment of Phenol-Stimulated
Trichloroethylene Bioremediation. App!. Environ. Microbiol. 67(10), 46714677.
Gossett, J. M. (1987). Measurement of Henry's Law Constants for C1 and C2
Chlorinated Hydrocarbons. Environ. Sd. Technol. 21(2), 202-208.
Hartmans, S., and Bont, J. A. M. d. (1992). Aerobic Vinyl Chloride Metabolism in
Mycobacterium Aurum Li. App!. Environ. Microbiol. 58(4), 1220-1226.
Hill, G. A., and Robinson, C. W. (1975). Substrate Inhibition Kinetics: Phenol
Degradation by Pseudomonas Putida. Biotechnol. Bioeng. 17, 1599-1615.
Holman, J. P. (2001). "Experimental Methods for Engineers." Seventh ed. McGrawHill, Boston.
Hopkins, G. D., and McCarty, P. L. (1995). Field Evaluation of in Situ Aerobic
Cometabolism of Trichioroethylene and Three Dichioroethylene Isomers
Using Phenol and Toluene as the Primary Substrates. Environ. Sci. Technol.
29(6), 1628-1637.
Hopkins, G. D., Munakata, J., Semprini, L., and McCarty, P. L. (1993a).
Trichioroethylene Concentration Effects on Pilot Field-Scale in-Situ
Groundwater Bioremediation by Phenol-Oxidizing Microorganisms. Environ.
Sd. Technol. 27(12), 2542-2547.
Hopkins, G. D., Semprini, L., and McCarty, P. L. (1993b). Microcosm and in Situ
Field Studies of Enhanced Biotransformation of Trichloroethylene by PhenolUtilizing Microorganisms. App!. Environ. Microbiol. 59(7), 2277-2285.
147
Horvath, R. (1972). Microbial Co-Metabolism and the Degradation of Organic
Compounds in Nature. Bacteriol. Rev. 36, 146-155.
Kim, Y., Arp, D. J., and Semprini, L. (2002). A Combined Method for Determining
Inhibition Type, Kinetic Parameters, and Inhibition Coefficients for Aerobic
Cometabolism of 1,1,1-Trichioroethane by a Butane-Grown Mixed Culture.
Biotechnol. Bioeng. 77(5), 564-576.
Mallinckrodt Baker Inc. (2000). Material Safety Data Sheet (Msds): Benzyl Alcohol.,
www.jtbaker.com/msds/b.htm.
McCarty, P. L. (1997). Aerobic Cometabolism of Chlorinated Aliphatic
Hydrocarbons. In "Subsurface Restoration" (C. H. Ward, J. A. Cherry, and M.
R. Scaif, Eds.), pp. 373-395. Ann Arbor Press, Inc., Chelsea, Michigan.
McCarty, P. L., Goltz, M. N., Hopkins, G. D., Dolan, M. E., Allan, J. P., Kawakarni,
B. T., and Carrothers, T. J. (1998). Full-Scale Evaluation of in Situ
Cometabolic Degradation of Trichioroethylene in Groundwater through
Toluene Injection. Environ. Sci. Technol. 32(1), 88-100.
McCarty, P. L., and Semprini, L. (1994). Ground-Water Treatment for Chlorinated
Solvents. In "Handbook of Bioremediation." (J. E. Matthews, Ed.), pp. 87-116.
Lewis Publishers, Ann Arbor.
Metcalf & Eddy Inc. (1991). "Wastewater Engineering: Treatment, Disposal, and
Reuse." (G. Tchobanoglous, and F. L. Burton, Eds.) McGraw-HIll, mc, New
York.
National Technical Information Service. (1989). Tr-343: Toxicology and
Carcinogenesis Studies of Benzyl Alcohol (Cas No. 100-51-6) in F344/N Rats
and B6c3fl Mice (Gavage Studies)., Vol. 2001. National Institue of
Environmental Health Sciences, http://ntp-server.niehs.nih.gov.
National Toxicology Program. (2001). H&S: Phenol 108-95-2, Vol. 2001. National
Institue of Environmental Health Sciences, http://ntp.-server.niehs.nih.gov.
National Toxicology Program. (2002). H&S: Benzyl Alcohol 100-51-6, Vol. 2001.
National Institue of Environmental Health Sciences, http://ntpserver.niehs.nih.gov.
Nelson, M. J. K., Montgomery, S. 0., O'Neill, E. J., and Pritchard, P. H. (1986).
Aerobic Metabolism of Trichloroethylene by a Bacterial Isolate. Appi.
Environ. Microbiol. 52(2), 383-384.
Oldenhuis, R., Vink, R. L. J. M., Janssen, D. B., and Witholt, B. (1989). Degradation
of Chlorinated Aliphatic Hydrocarbons by Methylosinus Trichosporium Ob3b
Expressing Soluble Methane Monooxygenase. Appi. Environ. Microbiol. 55,
2819-2826.
Semprini, L. (1997a). In Situ Transformation of Halogenated Aliphatic Compounds
under Anaerobic Conditions. In "Subsurface Restoration" (C. H. Ward, J. A.
Cherry, and M. R. Scaif, Eds.), pp. 429-450. Ann Arbor Press, Inc., Chelsea,
Michigan.
Semprini, L. (1997b). Strategies for the Aerobic Co-Metabolism of Chlorinated
Solvents. Curr. Opin. Biotechnol. 8, 296-308.
Shurtliff, M. M., Parkin, G. F., Weathers, L. J., and Gibson, D. T. (1996).
Biotransformation of Trichioroethylene by a Phenol-Induced Mixed Culture.
Journal of Environmental Engineering 122(7), 581-589.
Steffan, R. J., Sperry, K. L., Walsh, M. T., Vainberg, S., and Condee, C. W. (1999).
Field-Scale Evaluation of in Situ Bioaugmentation for Remediation of
Chlorinated Sovlents in Groundwater. Environ. Sci. Technol. 33(16), 27712781.
Tejasen, S., Dolan, M. E., and Semprini, L. (2003). Induction of 2-Hydroxy-Benzyl
Alcohol Production and Trichioroethylene Cometabolism by Benzyl Alcohol.
In Progress.
Tejasen, S., and Semprini, L. (2003). Aerobic Biotransformation of Chlorinated
Ethenes by a Mixed Culture Grown on a Broad Range of Substrates. Wat. Res.
submitted for publication.
Tovanabootr, A., and Semprini, L. (1998). Comparison of Tce Transformation
Abilities of Methane- and Propane-Utilizing Microorganisms. Bioremediation
J 2(2), 105-124.
van Hyickama Vlieg, J. E. T., de Koning, W., and Janssen, D. B. (1996).
Transformation Kinetics of Chlorinated Ethenes by Met hylosinus
Trichosporium Ob3b and Detection of Unstable Epoxides by on-Line Gas
Chromatography. Appi. Environ. Microbiol. 62, 3304-33 12.
Vancheeswaran, S., Halden, R. U., Williamson, K. J., James D. Ingle, J., and
Semprini, L. (1999). Abiotic and Biological Transformation of
Tetraalkoxysilanes and Trichloroethene/Cis- 1 ,2-Dichloroethene Cometabolism
Driven by Tetrabutoxysilane-Degrading Microorganisms. Environ Sci
Technol. 33(7), 1077-1085.
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2000). Characterization of an Isolate
That Uses Vinyl Chloride as a Growth Substrate under Aerobic Conditions.
Appi. Environ. Microbiol. 66(8), 3535-3542.
149
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2001). Transition from Cometabolic
to Growth-Linked Biodegradation of Vinyl Chloride by a Pseudomonas Sp.
Isolated on Ethene. Environ. Sci. Technol. 35(21), 4242-4251.
Vogel, T. M., and McCarty, P. L. (1985). Biotransformation of Tetrachioroethylene to
Trichioroethylene, Dichioroethylene, Vinyl Chloride and Carbon Dioxide
under Methanogenic Conditions. App!. Environ. Microbiol. 49(5), 1080-1083.
Wackett, L. P., Brusseau, G. A., Householder, S. R., and Hanson, R. S. (1989). Survey
of Microbial Oxygenases Trichioroethylene Degradation by PropaneOxidizing Bacteria. AppI. Environ. Microbiol. 55, 2960-2964.
Wackett, L. P., and Gibson, D. T. (1988). Degradation of Trichioroethylene by
Toluene Dioxygenase in Whole-Cell Studies with Pseudomonas Putida Fl.
App!. Environ. Microbiol. 54(7), 1703-1708.
Wackett, L. P., and Householder, S. R. (1989). Toxicity of Trichloroethylene to
Pseudomonas Putida Fl Is Mediated by Toluene Dioxygenase. App!. Environ.
Microbiol. 55, 2723-2725.
Wilson, J. T., and Wilson, B. H. (1985). Biotransformation of Trichioroethylene. App!.
Environ. Microbiol. 49(1), 242-243.
Yeager, C. M., Bottomley, P. J., and Arp, D. J. (2001). Cytotoxicity Associated with
Trichioroethylene Oxidation in Burkholderia Cepacia G4. App!. Environ.
Microbiol. 67(5), 2107-2115.
Yeager, C. M., Bottomley, P. J., Arp, D. I., and Hyman, M. R. (1999). Inactivation of
Toluene 2-Monooxygenase in Burkho!deria Cepacia G4 by Alkynes. Appl.
Environ. Microbio!. 65(2), 632-639.
Zhou, N.-Y., Jenkins, A., Chion, C. K. N. C. K., and Leak, D. J. (1999). The Alkene
Monooxygenase from Xanthobacter Strain Py2 Is Closely Related to Aromatic
Monooxygenases and Catalyzes Aromatic Monohydroxylation of Benzene,
Toluene, and Phenol. Appi. Environ. Microbiol. 65(4), 1589-1595.
150
CHAPTER 6: Conclusions and Engineering Significance
CONCLUSIONS
The enrichment culture was able of transforming cis-DCE upon growth on
most of the substrates tested that had at least two carbons and were substituted with
oxygen. TCE was also transformed when the culture effectively transformed cis-DCE.
Benzyl alcohol is of special interest since it is non-regulated, commonly used as foodflavoring agent (Mallinckrodt Baker Inc., 2000; National Technical Information
Service, 1989; National Toxicology Program, 2002), and was found to be very
effective in promoting the trarisformafion of VC, cis-DCE, and TCE. Other nonregulated substrates such as I -butanol, butyrate, and glucose were effectively served
as growth substrates for cis-DCE cometabolism.
The important characteristics of the growth substrates for this enrichment
culture are 1) high solubility; 2) the substrate must have two or more carbon atoms;
and 3) have some oxygen or hydroxide substitution. The enrichment also has the
ability to hydrolyze and utilize complex substrates, including TBOS, TPOS,
tetraphenoxysilane and phenyl acetate. Similar effectiveness in the cometabolic
transformation of cis-DCE and TCE was achieved when their hydrolysis byproducts
(1 -butanol, 1 -propanol, and phenol) were directly used. This result might be useful for
in-situ bioremediation where phenol could be replaced with relatively inert
compounds such as tetraphenoxysilane and phenyl acetate.
151
Two pathways of benzyl alcohol degradation were observed; through
benzaldehyde and 2-hydroxy benzyl alcohol (2HBA). Degradation of benzyl alcohol
through berizaldehyde did not result in effective TCE transfonnation. Successive
growth on benzyl alcohol led to increases in TCE transformation rates, TCE
transformation capacities, and 2HBA production rates. T-RFLP analysis indicated
that microbial communities remained unchanged during successive growth on benzyl
alcohol. A gradual induction of the 21-IBA pathway is one possible explanation for the
increase in performance. Rates of TCE transformation were correlated to rates of
2HBA production, indicating that ortho-monooxygenase was responsible for TCE
transformation. Exposure to TCE during growth on benzyl alcohol did not induce
enzymes for TCE cometabolism. Phenol-grown cells had higher TCE transformation
rates, TCE transformation capacities, and 21-JBA production rates than benzyl alcohol-
grown cells. When the mixed culture grew on benzyl alcohol after phenol, TCE
transformation rates were improved, but TCE transformation capacities were in the
range of those achieved from growth on benzyl alcohol alone. Another advantage was
that higher TCE transformation rates were achieved instantly when the cells were first
grown on phenol.
For benzyl alcohol-grown cells, the 2HBA production rate was much lower
than benzyl alcohol degradation rate, indicating that benzyl alcohol was likely
degraded through multiple pathways. Phenol-grown cells had similar rates of benzyl
alcohol degradation and 2HBA production, indicating an ortho-monooxygenase was
the dominant pathway. It is also possible that a downstream pathway differs for
152
benzyl alcohol and phenol grown cells, and 2HBA is being removed at slower rates in
the phenol-grown cells.
The mixed culture did not grow on toluene, but both benzyl alcohol- and
phenol-grown resting cells could oxidize toluene to o-cresol, indicating a similarity of
ortho-monooxygenase and phenol hydroxylase enzymes. Faster rates of TCE
transformation was related to faster rates of toluene oxidation. Inhibition and
inactivation studies with acetylene, 2-butyne, and I -hexyne indicated that both phenol-
and benzyl alcohol-grown cells expressed the same enzyme, which is similar to
toluene ortho-monooxygenase or phenol hydroxylase.
A benzyl alcohol grown culture was able to transform TCE, cis-DCE, and VC,
but not chloroform, 1,1 -dichloroethane, 1,1 -dichloroethene, or 1,1,1 -trichloroethane.
Degradation kinetics of benzyi alcohol followed the Monod kinetic model while
degradation kinetic of phenol followed the flaldane kinetic model. The specific
maximum degradation rate (k) of benzyl alcohol was a factor of three lower than
that of phenol. The half-saturation constant (Ks) of benzyl alcohol was about a factor
of fifty higher than that of phenol, indicating that phenol had a much higher affinity
for the enzyme, which is consistent with it being a better substrate for enzyme
induction. The Haldane inhibitory constant of phenol was 113 mg/L, indicating a
moderate tolerance to toxicity at high phenol concentrations. The km of TCE,
CjS
DCE, and VC by benzyl alcohol-grown cells was about a factor of two lower than
those of phenol-grown cells, while K values were in a similar range. Transformation
capacities
(Ta) for TCE, cis-DCE, and VC were about a factor of three higher with
153
phenol-grown cells. The modeling of these CAlls transformation using independently
measured
kmax
and K values matched well with observed data from batch studies.
In this study phenol-grown cells achieved better TCE, cis-DCE, and VC
transformations than benzyl alcohol-grown cells. This result indicates that phenol is
more effective than benzyl alcohol in being growth substrate in the transformation of
TCE, cis-DCE, and VC. However, phenol is a regulated compound (OSHA, ACGIH,
and NIOSH) and listed as material extremely hazardous to health (National
Toxicology Program, 2001), while benzyl alcohol is a non-regulated compound and
listed as moderate hazardous (National Toxicology Program, 2002). Benzyl alcohol
was also re-evaluated and approved in 2002 as a food flavoring agent (European
Commission,2O02). The kinetic parameters of TCE, cis-DCE, and VC
transformations achieved by benzyl alcohol-grown cells are among values reported by
other studies with toluene or phenol, indicating that benzyl alcohol is an effective
substrate for cometabolism of these compounds. Benzyl alcohol is also a liquid phase
at room temperature with a moderate flammability. Therefore, it might have potential
for in-situ bioremediation.
154
ENGINEERING SIGNIFICANCE
I. Benzyl alcohol, a non-regulated compound, was found to be an effective
substrate for the aerobic bioremediation of trichioroethylene, cisdichioroethylene, and vinyl chloride.
2. This mixed culture showed a high transformation of cis-dichioroethylene when
grown on simple substrates, such as 1 -butanol and butyrate. Therefore, it can
be used for bioaugmentation for cis-dichioroethylene bioremediation.
3. The enzyme involved in benzyl alcohol degradation was similar to toluene
ortho-monooxygenase, indicating the potential of replacing benzyl alcohol for
phenol or toluene in the in-situ bioremediation.
4. Kinetics of benzyl alcohol and phenol degradation and trichioroethylene, cis-
dichloroethylene, and vinyl chloride cometabolic transformation can be
important parameters for the design of the bioremediation system.
155
FUTURE WORK
Recommended future work includes:
1.
Isolation of a pure benzyl alcohol-culture capable of cometabolizing VC, cisDCE, and TCE.
2. Evaluate the induction of ortho-monooxygenase enzyme with successive
growth on benzyl alcohol by the pure culture.
3. Evaluate benzyl alcohol as a growth substrate for other TCE cometabolizing
microorganisms.
4. Conduct a long term study of VC, cis-DCE, and TCE cometabolism with
growth on benzyl alcohol.
156
REFERENCE
European Commission. (2002). Opinion of the Scientific Committee on Food on
Benzyly Alcohol, pp. 1-15. Health & Consumer Protection DirectorateGeneral, Brussel, Belgium.
Mallinckrodt Baker Inc. (2000). Material Safety Data Sheet (Msds): Benzyl Alcohol.,
www.jtbaker.comlmsdslbl 885 .htm.
National Technical Information Service. (1989). Tr-343: Toxicology and
Carcinogenesis Studies of Benzyl Alcohol (Cas No. 100-51-6) in F344/N Rats
and B6c3fl Mice (Gavage Studies)., Vol. 2001. National Institue of
Environmental Health Sciences, http://ntp-server.niehs.nih.gov.
National Toxicology Program. (2001). H&S: Phenol 108-95-2, Vol. 2001.National
Institue of Environmental Health Sciences, http://ntp-server.niehs.nih.gov.
National Toxicology Program. (2002). H&S: Benzyl Alcohol 100-51-6, Vol. 2001.
National Institue of Environmental Health Sciences, http:/Intpserver.niehs.nih.gov.
157
BIBLIOGRAPHY
Alvarez-Cohen, L., and McCarty, P. L. (1991). A Cometabolic Biotransformation
Model for Halogenated Aliphatic Compounds Exhibiting Product Toxicity.
Environ. Sci. Technol. 25(8), 1381-1387.
Alvarez-Cohen, L., arid Speitel, 0. E. J. (2001). Kinetics of Aerobic Cometabolism of
Chlorinated Solvents. Biodegradation 12(2), 105-126.
American Public Health Association. (1985). Standard Methods for the Examination
of Water and Wastewater. 16th ed. APHA, New York.
Anderson, J. E., and McCarty, P. L. (1996). Effect of Three Chlorinated Ethenes on
Growth Rates for a Methanotrophic Mixed Culture. Environ. Sci. Technol.
30(12), 3517-3524.
Arciero, D., Vaimelli, T., Logan, M., and Hooper, A. B. (1989). Degradation of
Trichioroethylene by the Ammonia-Oxidizing Bacterium Nitrosomonas
Europaea. Biochem. Biophys. Res. Comm. 159(2), 640-643.
Arp, D. J., Yeager, C. M., and I-lyman, M. R. (2001). Molecular and Cellular
Fundamentals of Aerobic Cometabolism of Trichioroethylene. Biodegradation
12(2), 81-1 03.
Ayoubi, P. J., and Harker, A. R. (1998). Whole-Cell Kinetics of Trichloroethylene
Degradation by Phenol Hydroxylase in a Rastonia Eutropha Jmp134
Derivative. App!. Environ. Microbiol. 64(11), 4353-4356.
Bailey, J., and 011is, D. (1986). "Biochemical Engineering Fundamentals." McGrawHill Plublishing Co., New York.
Bielefeldt, A. R., Stensel, H. D., and Strand, S. E. (1995). Cometabolic Degradation of
Tee and Dee without Intermediate Toxicity. Journal ofEnvironmental
Engineering 121(11), 791-797.
Bouwer, E. J., and McCarty, P. L. (1983). Transformation of 1- and 2-Carbon
Halogenated Aliphatic Organic Compounds under Methanogenic Conditions.
Appl. Environ. Microbiol. 45, 1286-1294.
Bradley, P. M., and Chapelle, F. Fl. (2000). Aerobic Microbial Mineralization of
Dichloroethene as Sole Carbon Substrate. Environ. Sci. Technol. 34(1), 221223.
158
Chang, H.-L., and Alvarez-Cohen, L. (1995 a). Model for the Cometabolic
Biodegradation of Chlorinated Organics. Environ. Sci. Technol. 29(9), 23572367.
Chang, H.-L., and Alvarez-Cohen, L. (1 995b). Transformation Capacities of
Chlorinated Organics by Mixed Cultures Enriched on Methane, Propane,
Toluene, or Phenol. Biotechnol. Bioeng. 45,440-449.
Chang, H.-L., and Alvarez-Cohen, L. (1996). Biodegradation of Individual and
Multiple Chlorinated Aliphatics by Mixed and Pure Methane Oxidizing
Cultures. App!. Environ. Microbiol. 62(9), 3371-3377.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain. J. C. (2002a).
Biodegradation of Cis-Dichioroethene as the Sole Carbon Source by a BProteobacterium. Appi. Environ. Microbiol. 68(6), 2726-2730.
Coleman, N. V., Mattes, T. E., Gossett, J. M., and Spain, J. C. (2002b). Phylogenetic
and Kinetic Diversity of Aerobic Vinyl Chloride-Assimilating Bacteria from
Contaminated Sites. App!. Environ. Microbiol. 68(12), 6162-6171.
Dabrock, B., Kebeler, M., Averhoff, R., and Gottschalk, G. (1994). Identification and
Characterization of a Transmissible Linear Plasmid from Rhodococcus
Erythropolis Bd2 That Encodes Isopropylbenzene and Trichloroethene
Catabolism. Appi. Environ. Microbiol. 60(3), 853-860.
Dabrock, B., Riedel, J., Bertram, J., and Gottschalk, G. (1992). Isopropylbenzene
(Cumene)- a New Substrate for the Isolation of Trichlorothene-Degrading
Bacteria. Arch. Microbiol. 158, 9-13.
Dagley, S., and Gibson, D. T. (1965). The Bacterial Degradation of Catechol. Bichem.
95, 466-474.
DiSpirito, A. A., Gulledge, J., Shiemke, A. K., Murrel, J. C., Lidstrom, M. E., and
Krerna, C. L. (1992). Trichloroethylene Oxidation by the MembraneAssociated Methane Monooxygenase in Type 1, Type Ii and Type X
Methanotrophs. Biodegradation 2, 151-164.
Dolan, M. E., and McCarty, P. L. (1995). Methanotrophic Chioroethene
Transformation Capacities and 1,1 -Dichloroethene Transformation in Aquifer
Samples. Environ. Sci. Technol. 29, 1892-1897.
Duetz, W., Jong, C. d., Williams, P., and Andel, J. v. (1994). Competition in
Chemostat Culture between Pseudomonas Strains That Use Different Pathways
for the Degradation of Toluene. Appi. Environ. Microbiol. 60(8), 2858-2863.
159
Dunfield, P. F., Liesack, W., Henckel, T., Knowles, R., and Conrad, R. (1999). HighAffinity Methane Oxidation by a Soil Enrichment Culture Containing a Type Ii
Methanotroph. Appi. Environ. Microbiol. 65(3), 1009-1014.
Ensign, S. A. (1996). Aliphatic and Chlorinated Alkenes and Epoxides as Inducers of
Alkene Monooxygenase and Epoxidase Activities in Xanthobacter Strain Py2.
App!. Environ. Microbiol. 62(1), 61-66.
Ensign, S. A., Hyman, M. R., and Arp, D. J. (1992). Cometabolic Degradation of
Chlorinated Alkenes by Alkene Monooxygenase in a Propylene-Grown
Xanthobacter Strain. Appi. Environ. Microbiol. 58(9), 3038-3046.
Eriksson, M., Sodersten, E., Yu, Z., Daihammar, G., and Mohn, W. W. (2003).
Degradation of Polycyclic Aromatic Hydrocarbons at Low Temperature under
Aerobic and Nitrate-Reducing Conditions in Enrichment Cultures from
Northern Soils. App!. Environ. Microbiol. 69(1), 275-284.
European Commission. (2002). Opinion of the Scientific Committee on Food on
Benzyly Alcohol, pp. 1-15. Health & Consumer Protection DirectorateGeneral, Brussel, Belgium.
Ewers, J., Clemens, W., and Knackmuss, H. J., Eds. (1991). Biodegradation of
Chloroethenes Using Isoprene as Co-Substrate. Vol. 1. International
Symposium on Environmental Biotechnology. Ostend, Belgium: Royal
Flemish Society of Engineers.
Fairlee, J. r., Burback, B. L., and Perry, J. J. (1997). Biodegradation of Groundwater
Pollutants by a Combined Culture of Mycobacterium Vaccae and a
Rhodococcus Sp. Can. I Microbiol. 43, 841-846.
Fan, S., and Scow, K. (1993). Biodegradation of Trichloroethylene and Toluene by
Indigenous Microbial Populations in Soil. Appi. Environ. Microbiol. 59(6),
1911-1918.
Fennell, D. E., and Gossett, J. M. (1997). Comparison of Butyric Acid, Ethanol, Lactic
Acid, and Propionic Acid as Hydrogen Donors for the Reductive
Dechlorination of Tetrachioroethene. Environ. Sci. Technol. 31(3), 918-926.
Fennell, D. E., and Gossett, J. M. (1998). Modeling the Production of and Competition
for Hydrogen in a Dechlorinating Culture. Environ. Sd. Technol. 32(16), 24502460.
Fogel, M. M., Taddeo, A. R., and Fogel, 5. (1986). Biodegradation of Chlorinated
Ethenes by a Methane-Utilizing Mixed Culture. Appi. Environ. Microbiol.
51(4), 720-724.
160
Folsom, B. R., Chapman, P. J., and Pritchard, P. H. (1990). Phenol and
Trichioroethylene Degradation by Pseudomonas Cepacia G4: Kinetics and
Interactions between Substrates. Appi. Environ. Microbiol. 56(5), 1279-1285.
Fries, M. R., Forney, L. J., and Tiedje, J. M. (1997). Phenol- and Toluene-Degrading
Microbial Populations from an Aquifer in Which Successful Trichioroethene
Cometabolism Occurred. App!. Environ. Microbiol. 63(4), 1523-1530.
Futamata, H., Harayama, S., and Watanabe, K. (2001a). Diversity in Kinetics of
Trichioroethylene-Degrading Activities Exhibited by Phenol-Degrading
Bacteria. Appi. Microbiol. Biotechnol. 55, 248-253.
Futamata, H., Harayama, S., and Watanabe, K. (2001b). Group-Specific Monitoring of
Phenol Hydroxylase Genes for a Functional Assessment of Phenol-Stimulated
Trichioroethylene Bioremediation. App!. Environ. Microbiol. 67(10), 46714677.
Gao, J., and Skeen, R. S. (1999). Glucose-Induced Biodegradation of CisDichioroethylene under Aerobic Conditions. Wat. Res. 33(12), 2789-2796.
Gibson, S. A., and Sewell, G. W. (1992). Stimulation of Reductive Dechlorination of
Tetrachioroethene in Anaerobic Aquifer Microcosms by Addition of ShortChain Organic Acids or Alcohols. Appi. Environ. Microbiol. 58(4), 1392-1393.
Gossett, J. M. (1987). Measurement of Henry's Law Constants for C1 and C2
Chlorinated Hydrocarbons. Environ. Sci. Technol. 21(2), 202-208.
Goudar, C. T., Ganji, S. H., Pujar, B. G., and Strevett, K. A. (2000). Substrate
Inhibition Kinetics of Phenol Biodegradation. Water Environment Research
72(1), 50-55.
1-laigler, B. E., Pettigrew, C. A., and Spain, J. C. (1992). Biodegradation of Mixtures
of Substituted Benzenes by Pseudomonas Sp. Strain Jsl 50. App!. Environ.
Microbiol. 58(7), 2237-2244.
Hamamura, N., Page, C., Long, T., Semprini, L., and Arp, D. J. (1997). Chlorofonn
Cometabolism by Butane-Grown Cf8, Pseudomonas Butanovora, and
Mycobacterium Vaccae Job5 and Methane-Grown Methylosinus
Trichosporium Ob3b. App!. Environ. Microbiol. 63(9), 3607-36 13.
Harker, A. R., and Kim, Y. (1990). Trichloroethylene Degradation by Two
Independent Aromatic-Degrading Pathways in Alcaligenes Eutrophus Jmp134.
AppI. Environ. Microbiol. 56(4), 1179-1181.
Hartmans, S., and Bont, J. A. M. d. (1992). Aerobic Vinyl Chloride Metabolism in
Mycobacterium Aurum Li. Appi. Environ. Microbiol 58(4), 1220-1226.
161
Hill, G. A., and Robinson, C. W. (1975). Substrate Inhibition Kinetics: Phenol
Degradation by Pseudomonas Putida. Biotechnol. Bioeng. 17, 1599-1615.
Holman, J. P. (2001). "Experimental Methods for Engineers." Seventh ed. McGrawHill, Boston.
Hopkins, G. D., and McCarty, P. L. (1995). Field Evaluation of in Situ Aerobic
Cometabolism of Trichioroethylene and Three Dichioroethylene Isomers
Using Phenol and Toluene as the Primary Substrates. Environ. Sd. Technol.
29(6), 1628-1637.
Hopkins, G. D., Munakata, J., Semprini, L., and McCarty, P. L. (1993a).
Trichloroethylene Concentration Effects on Pilot Field-Scale in-Situ
Groundwater Bioremediation by Phenol-Oxidizing Microorganisms. Environ.
Sd. Technol. 27(12), 2542-2547.
Hopkins, G. D., Semprini, L., and McCarty, P. L. (1993b). Microcosm and in Situ
Field Studies of Enhanced Biotransformation of Trichioroethylene by PhenolUtilizing Microorganisms. App!. Environ. Microbiol. 59(7), 2277-2285.
Horvath, R. (1972). Microbial Co-Metabolism and the Degradation of Organic
Compounds in Nature. Bacteriol. Rev. 36, 146-155.
Hyman, M. R., and Wood, P. M. (1985). Suicidal Inactiation and Labelling of
Ammonia Mono-Oxygenase by Acetylene. Biochem. .J. 227,719-725.
Jenal-Wanrier, U., and McCarty, P. L. (1997). Development and Evaluation of
Semicontinuous Slurry Microcosms to Simulate in Situ Biodegradation of
Trichloroethylene in Contaminated Aquifers. Environ. Sci. Technol. 3 1(10),
2915-2922.
Johnson, G., and Olsen, R. (1995). Nucleotide Sequence Analysis of Genes Encoding
a Toluene/Benzene-2- Monooxygenase from Pseudomonas Sp. Strain Jsl 50.
App!. Environ. Microbiol. 61(9), 3336-3346.
Johnson, G., and Olsen, R. (1997). Multiple Pathways for Toluene Degradation in
Burkholderia Sp. Strain JslSO. Appi. Environ. Microbiol. 63(10), 4047-4052.
Kamagata, Y., Fuithorpe, R. R., Tamura, K., Takami, H., Fomey, L. J., and Tiedje, J.
M. (1997). Pristine Environments Harbor a New Group of Oligotrophic 2,4Dichlorophenoxyacetic Acid-Degrading Bacteria. App!. Envfron. Microbiol.
63(6), 2266-2272.
Kaphammer, B., Kukor, J. J., and Olsen, R. H. (1991). Cloning and Characterization
of a Novel Toluene Degradative Pathway from Pseudomonas Pickettii Pko 1. In
162
"Biodeterioration and Biodegradation 8" (H. W. Rossmoore, Ed.). Elsevier
Applied Science, London.
Kim, Y., Arp, D. J., and Semprini, L. (2000). Chlorinated Solvent Cometabolism by
Butane-Grown Mixed Culture. Journal ofEnvironmental Engineering
October, 934-942.
Kim, Y., Arp, D. J., and Semprini, L. (2002). A Combined Method for Determining
Inhibition Type, Kinetic Parameters, and Inhibition Coefficients for Aerobic
Cometabolism of 1,1,1-Trichloroethane by a Butane-Grown Mixed Culture.
Biotechnol. Bioeng. 77(5), 564-576.
Kim, Y., Ayoubi, P., and Harker, A. R. (1996). Constitutive Expression of the Cloned
Phenol Hydroxylase Gene(S) from Alcaligenes Eutrophus Jmp 134 and
Concomitant Trichiorethylene Oxidation. Appi. Environ. Micro biol. 62(9),
3227-3233.
Kukor, J. J., and Olsen, R. H. (1991). Molecular Cloning and Regulation of Genes
Encoding Meta-Cleavage of Catechols Produced from Catabolism of Phenol
and M-Cresol by Pseudomonas Pickettii Pkol. In "Biodeterioration and
Biodegradation 8" (H. W. Rossmoore, Ed.). Elsevier Applied Science, London.
Landa, A. S., Sipkema, E. M., Weijma, J., Beenackers, A. A. C. M., Dolfing, J., and
Jannssen, D. B. (1994). Cometabolic Degradation of Trichloroethylene by
Pseudomonas Cepacia G4 in a Chemostat with Toluene as the Primary
Substrate. App!. Environ. Microbiol. 60(9), 3368-3374.
Leahy, J., Byrne, A., and Olsen, R. (1996). Comparison of Factors Influencing
Trichioroethylene Degradation by Toluene-Oxidizing Bacteria. App!. Environ.
MicrobioL 62(3), 825-833.
Lee, M. D., Odom, J. M., and Buchanan Jr., R. J. (1998). New Perspectives on
Microbial Dehalogenation of Chlorinated Solvents: Insights from the Field.
Annu. Rev. Microbiol. 52, 423-452.
Lee, S.-B., Strand, S. E., and Stensel, H. D. (2000). Sustained Degradation of
Trichloroethylene in a Suspended Growth Gas Treatment Reactor by an
Actinomycetes Enrichment. Environ. Sci. Technol. 34(15), 3261-3268.
Leonard, D., Youssef, C. B., Destruhaut, C., Lindley, N. D., and Queinnec, 1. (1999).
Phenol Degradation by Raistonia Eutropba: Colorimetric Determination of 2Flydroxymuconate Semialdehyde Accumulation to Control Feed Strategy in
Fed-Batch Fermentations. Biotechnol. Bioeng. 65(4), 407-415.
163
Little, D. C., Palumbo, A. V., Herbes, S. E., Lidstrom, M. E., Tyndall, R. L., and
Gilmer, P. J. (1988). Trichloroethylene Biodegradation by a MethaneOxidizing Bacterium. App!. Environ. Microbiol. 54(4), 951-956.
Liu, W.-T., Marsh, T. L., Cheng, H., and Forney, L. J. (1997). Characterization of
Microbial Diversity by Determining Terminal Restriction Fracment Length
Polymorphisms of Genes Encoding 1 6s Rrna. App!. Environ. MicrobioL
63(11), 45 16-4522.
Madigan, M. T., Martinko, J. M., and Parker, J., Eds. (1996). Brock Biology of
Microorganisms. 8 ed. Upper Saddle River, NJ: Prentice Hall.
Malachowsky, K., Phelps, T., Teboli, A., Minnikin, D., and White, D. (1994). Aerobic
Mineralization of Trichioroethylene, Vinyl Chloride, and Aromatic
Compounds by Rhodococcus Species. App!. Environ. Microbiol. 60(2), 542548.
Mallinckrodt Baker Inc. (2000). Material Safety Data Sheet (Msds): Benzyl Alcohol.,
www.jtbaker.comlmsds/bl 885 .htm.
McCarty, P. L. (1997). Aerobic Cometabolism of Chlorinated Aliphatic
Hydrocarbons. In "Subsurface Restoration" (C. H. Ward, J. A. Cherry, and M.
R. Scaif, Eds.), pp. 373-395. Ann Arbor Press, Inc., Chelsea, Michigan.
McCarty, P. L., Goltz, M. N., Hopkins, G. D., Dolan, M. E., Allan, J. P., Kawakami,
B. T., and Carrothers, T. J. (1998a). Full-Scale Evaluation of in Situ
Cometabolic Degradation of Trichloroethylene in Groundwater through
Toluene Injection. Environ. Sci. Technol. 32(1), 88-100.
McCarty, P. L., Hopkins, G. D., Munakata-Marr, J., Matheson, V. G., Dolan, M. E.,
Dion, L. B., Shields, M., Forney, L. J., and Tiedje, J. M. (1998b).
Bioaugmentation with Burkholderia Cepacia Pr 13 for in Situ Bioremediation
of Trichioroethylene Contaminated Groundwater. EPA(April), 1-1 1.
McCarty, P. L., and Semprini, L. (1994). Ground-Water Treatment for Chlorinated
Solvents. In "Handbook of Bioremediation." (J. E. Matthews, Ed.), pp. 87-116.
Lewis Publishers, Ann Arbor.
McClay, K., Streger, S. H., and Steffan, R. J. (1995). Induction of Toluene Oxidation
Activity in Pseudomonas Mendocina Krl and Pseudomonas Sp. Strain Envpc5
by Chlorinated Solvents and Alkanes. App!. Environ. Microbiol. 61(9), 3479348 1.
Metcalf & Eddy Inc. (1991). "Wastewater Engineering: Treatment, Disposal, and
Reuse." (G. Tchobanoglous, and F. L. Burton, Eds.) McGraw-HIll, mc, New
York.
164
Muller, R. H., and Babel, W. (1995). Determination of the Ks Values During the
Growth of Alcaligenes Eutrophus on Phenol, 2,4-Dichiorophenoxyacetic Acid
and Fructose. Acta Biotechnol. 15(4), 347-353.
Muller, R. H., and Babel, W. (1996a). Growth Rate-Dependent Expression of PhenolAssimilation Pathways in Alcaligenes Eutrophus Jmp 134 - the Influence of
Formate as as Auxiliary Energy Source on Phenol Conversion Characteristics.
Appi. Microbiol. Biotechnol. 46(2), 156-162.
Muller, R. FL, and Babel, W. (1996b). Measurement of Growth at Very Low Rates
(M>0), an Approach to Study the Energy Requirement for the Surial of
Alcaligenes Eutrophus Jmpl 34. Appl. Environ. Microbiol. 62(1), 147-151.
Munakata-Marr, J., Matheson, V. G., Fomey, L. J., Tiedje, J. M., and McCarty, P. L.
(1997). Long-Term Biodegradation of Trichioroethylene Influenced by
Bioaugmentation and Dissolved Oxygen in Aquifer Microcosms. Environ. Sci.
Technol. 31(3), 786-791.
Munakata-Marr, J., McCarty, P. L., Shields, M. S., Reagin, M., and Francesconi, S. C.
(1996). Enhancement of Trichloroethylene Degradation in Aquifer
Microcosms Bioaugmented with Wild Type and Genetically Altered
Burkholderia (Pseudomonas) Cepacia G4 and Pri. Environ. Sd. Technol.
30(6), 2045-2052.
National Center for Environmental Assessment. (20011). Trichloroethylene Health Risk
Assessment. U.S. Environmental Protection Agency,
http://cfpub.epa.gov/ncea.
National Technical Information Service. (1989). Tr-343: Toxicology and
Carcinogenesis Studies of Benzyl Alcohol (Cas No. 100-51-6) in F344/N Rats
and B6c3f1 Mice (Gavage Studies)., Vol. 2001. National Institue of
Environmental Health Sciences, http ://ntp-server.niehs.nih.gov.
National Toxicology Program. (2001). H&S: Phenol 108-95-2, Vol. 2001. National
Institue of Environmental Health Sciences, http://ntp-server.niehs.nih.gov.
National Toxicology Program. (2002). H&S: Benzyl Alcohol 100-51-6, Vol. 2001.
National Institue of Environmental Health Sciences, http:!/ntpserver.niehs.nih.gov.
Nelson, M. J. K., Montgomery, 5. 0., Mahaffey, w. R., and Pritchard, P. H. (1987).
Biodegradation of Trichloroethylene and Involvement of an Aromatic
Biodegradative Pathway. Appi. Environ. Microbiol. 53(5), 949-954.
165
Nelson, M. J. K., Montgomery, S. 0., O'Neill, E. J., and Pritchard, P. Fl. (1986).
Aerobic Metabolism of Trichioroethylene by a Bacterial Isolate. App!.
Environ. Microbiol. 52(2), 383-384.
Nelson, M. J. K., Montgomery, S. 0., and Pritchard, P. H. (1988). Trichloroethylene
Metabolism by Microorganisms That Degrade Aromatic Compounds. Appi.
Environ. Microbiol. 54(2), 604-606.
Oldenhuis, R., Oedzes, J. Y., Waarde, v. d. J. J., and Janssen, D. B. (1991). Kinetics of
Chlorinated Hydrocarbon Degradation by Methylosinus Trichosporium Ob3b
and Toxicity of Trichloroethylene. App!. Environ. Microbiol. 57, 7-14.
Oldenhuis, R., Vink, R. L. J. M., Janssen, D. B., and Witholt, B. (1989). Degradation
of Chlorinated Aliphatic Hydrocarbons by Methylosinus Trichosporiurn Ob3b
Expressing Soluble Methane Monooxygenase. Appi. Environ. Microbiol. 55,
28 19-2826.
Reij, M., Kieboom, J., Bont, J. d., and Hartmans, S. (1995). Continuous Degradation
of Trichioroethylene by Xanthobacter Sp. Strain Py2 During Growth on
Propene. Appi. Environ. Microbiol. 61(8), 2936-2942.
Rooney-Varga, J. N., Anderson, R. T., Fraga, J. L., Ringelberg, D., and Lovley, D. R.
(1999). Microbial Communities Associated with Anaerobic Benzene
Degradation in a Petroleum-Contaminated Aquifer. App!. Environ. Microbiol.
65(7), 3056-3063.
Ryoo, D., Shim, H., Arenghi, F. L. g., Barbieri, P., and Wood, T. K. (2001).
Tetrachioroethylene, Trichloroethylene, and Chlorinated Phenols Induce
Toluene-O-Xylene Monooxygenase Activity in Pseudomonas Stutzeri Oxi.
AppL Microbiol. Biotechnol. 56(3/4), 545-549.
Saeki, H., Akira, M., Furuhashi, K., Averhoff, B., and Gottschalk, G. (1999).
Degradation of Trichioroethene by a Linear-Plasmid-Encoded Alkene
Monooxygenase in Rhodococcus Corallinus (Nocardia Coral/ma,) B-276.
Microbiol. 145, 1721-1730.
Semprini, L. (1997a). In Situ Transformation of Halogenated Aliphatic Compounds
under Anaerobic Conditions. In "Subsurface Restoration" (C. H. Ward, J. A.
Cherry, and M. R. Scaif, Eds.), pp. 429-450. Ann Arbor Press, Inc., Chelsea,
Michigan.
Semprini, L. (1 997b). Strategies for the Aerobic Co-Metabolism of Chlorinated
Solvents. Curr. Opin. Biotechnol. 8, 296-3 08.
Semprini, L., Ely, R. L., and Lang, M. M. (1998). Modeling of Cometabolism for the
in Situ Biodegradation of Trichloroethylene and Other Chlorinated Aliphatic
166
Hydrocarbons. In "Bioremediation: Principles and Practice Volume 1,
Fundamentals and Applications" (S. K. Sikdar, and R. L. Irvine, Eds.), pp. 89134. Technomic Publishing, Lancaster.
Semprini, L., Robert, P. V., Hopkins, G. D., and McCarty, P. L. (1990). A Field
Evaluation of in-Situ Biodegradation of Chlorinated Ethenes: Part 2.
Cometabolic Transformations. Ground Water 30(1), 37-44.
Shields, M. S., Montgomery, S. 0., Chapman, P. J., Cuskey, S. M., and Pritchard, P.
H. (1989). Novel Pathway of Toluene Catabolism in the TrichloroethyleneDegrading Bacterium G4. Appi. Environ. Microbiol. 55(6), 1624-1629.
Shields, M. S., Montgomery, S. 0., Cuskey, S. M., Chapman, P. J., and Pritchard, P.
H. (1991). Mutants of Pseudomonas Cepacia G4 Defective in Catabolism of
Aromatic Compounds and Trichioroethylene. App!. Environ. Microbiol. 57(7),
1935-1941.
Shingleton, J. t., Applegate, B. M., Nagel, A. C., Bienkowski, P. R., and Sayler, G. S.
(1998). Induction of the Tod Operon by Trichioroethylene in Pseudomonas
Putida Tva8. Appi. Environ. Microbiol. 64(12), 5049-5052.
Shurtliff, M. M., Parkin, G. F., Weathers, L. J., and Gibson, D. T. (1996).
Biotransformation of Trichloroethylene by a Phenol-Induced Mixed Culture.
Journal of Environmental Engineering 122(7), 581-589.
Smith, C. A., O'Reilly, K. T., and Hyman, M. R. (2003). Characterization of the Initial
Reactions During the Cometabolic Oxidation of Methyl Tert-Butyl Ether by
Propane-Grown Mycobacterium Vaccae JobS. App!. Environ. Microbiol.
69(2), 796-804.
Solyanikova, I. P., and Golovieva, L. A. (1999). Phenol Hydroxylases: An Update.
Biochemistry (Moscow) 64(4), 437-446.
Steffan, R. J., Sperry, K. L., Walsh, M. T., Vainberg, S., and Condee, C. W. (1999).
Field-Scale Evaluation of in Situ Bioaugmentation for Remediation of
Chlorinated Sovlents in Groundwater. Environ. Sci. Technol. 33(16), 2771278 1.
Tejasen, S., Dolan, M. F., and Semprini, L. (2003). Induction of 2-Hydroxy-Benzyl
Alcohol Production and Trichioroethylene Cometabolism by Benzyl Alcohol.
In Progress.
Tejasen, S., and Semprini, L. (2003). Aerobic Biotransformation of Chlorinated
Ethenes by a Mixed Culture Grown on a Broad Range of Substrates. Wat Res.
submitted for publication.
167
Tovanabootr, A., and Semprini, L. (1998). Comparison of Tce Transformation
Abilities of Methane- and Propane-Utilizing Microorganisms. Bioreniediation
J 2(2), 105-124.
Tsien, H. C., Brusseau, G. A., Hanson, R. S., and Wackett, L. P. (1989).
Biodegradation of Trichioroethylene by Methylosinus Trichosporium Ob3b.
AppI. Environ. Microbiol. 55(12), 3155-3161.
U.S. EPA. (2001a). Chemical Summary for Toluene. Office of Pollution Prevention
and Toxics, U.S. Environmental Protection Agency,
http://www.epa.gov/opptintr/chemfact/s_toluene.txt.
U.S. EPA. (2001b). National Primary Drinking Water Regulations. Office of water,
U.S. Environmental Protection Agency,
http://www.bren.ucsb.edulfac_staff!faclkeller/courses/esm223/MCL.html.
U.S. EPA. (2002a). Consumer Factsheet On: I ,2-Dichloroethylene. U.S.
Environmental Protection Agency, http://www.epa.gov/safewater/dwhlcvoc/1 2-dich2.html.
U.S. EPA. (2002b). Consumer Factsheet On: Trichioroethylene. U.S. Environmental
Protection Agency, http://www.epa.gov/safewater/dwhlc-voc/trichlor.html.
U.S. EPA. (2003). Consumer Factsheet On: Vinyl Chloride. U.S. Environmental
Protection Agency,
http://www.epa.gov/safewater/contaminants/dw_contamfs/vinylchl .html.
van Hyickama Vlieg, J. E. T., de Koning, W., and Janssen, D. B. (1996).
Transformation Kinetics of Chlorinated Ethenes by Met hylosinus
Trichosporium Ob3b and Detection of Unstable Epoxides by on-Line Gas
Chromatography. Appl. Environ. Microbiol. 62, 3304-3312.
Vancheeswaran, S., Halden, R. U., Williamson, K. J., James D. Ingle, J., and
Semprini, L. (1999). Abiotic and Biological Transformation of
Tetraalkoxysilanes and Trichioroethene/Cis- I ,2-Dichloroethene Cometabolism
Driven by Tetrabutoxysilane-Degrading Microorganisms. Environ Sci.
Technol. 33(7), 1077-1085.
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2000). Characterization of an Isolate
That Uses Vinyl Chloride as a Growth Substrate under Aerobic Conditions.
Appi. Environ. Microbiol. 66(8), 3535-3542.
Verce, M. F., Ulrich, R. L., and Freedman, D. L. (2001). Transition from Cometabolic
to Growth-Linked Biodegradation of Vinyl Chloride by a Pseudomonas Sp.
Isolated on Ethene. Environ. Sci. Technol. 35(21), 4242-4251.
168
Vlieg, J. E. T. v. H., Koning, W. d., and Janssen, D. B. (1996). Transformation
Kinetics of Chlorinated Ethenes by Methylosinus Trichosporium Ob3b and
Detection of Unstable Epoxides by on-Line Gas Chromatography. Appi.
Environ. Microbiol. 62(9), 3304-33 12.
Voet, D., and Voet, J. (1990). "Biochemistry." John Wiley and Sons, New York.
Vogel, T. M., and McCarty, P. L. (1985). Biotransformation of Tetrachloroethylene to
Trichioroethylene, Dichloroethylene, Vinyl Chloride and Carbon Dioxide
under Methanogenic Conditions. Appi. Environ. Microbiol. 49(5), 1080-1083.
Wackett, L. P., Brusseau, G. A., Householder, S. R., and Hanson, R. S. (1989). Survey
of Microbial Oxygenases Trichioroethylene Degradation by PropaneOxidizing Bacteria. App!. Environ. Microbiol. 55, 2960-2964.
Wackett, L. P., and Gibson, D. T. (1988). Degradation of Trichloroethylene by
Toluene Dioxygenase in Whole-Cell Studies with Pseudomonas Putida Fl.
Appi. Environ. Microbiol. 54(7), 1703-1708.
Wackett, L. P., and Householder, S. R. (1989). Toxicity of Trichioroethylene to
Pseudomonas Putida Fl Is Mediated by Toluene Dioxygenase. App!. Environ.
Microbiol. 55, 2723-2725.
Whited, G. M., and Gibson, D. T. (199 Ia). Separation and Partial Characterization of
the Enzymes of the Toluene-4-Monooxygenase Catabolic Pathway in
Pseudomonas Mendocina Krl. J. Bacteriol. 173(9), 30 17-3020.
Whited, G. M., and Gibson, D. T. (1991b). Toluene-4-Monooxygenase, a ThreeComponent Enzyme System That Catalyzes the Oxidation of Toluene to PCresol in Pseudomonas Mendocina Krl. J. Bacteriol. 173(9), 30 10-3016.
Wilson, J. T., and Wilson, B. H. (1985). Biotransformation of Trichioroethylene. Appi.
Environ. Microbiol. 49(1), 242-243.
Winter, R. B., Yen, K.-M., and Ensley, B. D. (1989). Efficient Degradation of
Trichioroethylene by a Recombinant Escherichia Co/i. Bio Technology 7(3),
282-285.
Yeager, C. M., Bottomley, P. J., and Arp, D. J. (2001). Cytotoxicity Associated with
Trichioroethylene Oxidation in Burkholderia Cepacia G4. App!. Environ.
Microbiol. 67(5), 2107-2115.
Yeager, C. M., Bottomley, P. J., Arp, D. J., and Hyman, M. R. (1999). Inactivation of
Toluene 2-Monooxygenase in Burkhoideria Cepacia G4 by Alkynes. App!.
Environ. Microbiol. 65(2), 632-639.
169
Zhou, N.-Y., Jenkins, A., Chion, C. K. N. C. K., and Leak, D. J. (1999). The Alkene
Monooxygenase from Xanthobacter Strain Py2 Is Closely Related to Aromatic
Monooxygenases and Catalyzes Aromatic Monohydroxylation of Benzene,
Toluene, and Phenol. App!. Environ. Microbiol. 65(4), 1589-1595.
170
APPENDICES
171
Appendix A: HPLC Operation
This method is applicable for the determination of aromatic compounds such
as phenol, toluene, benzyl alcohol, and their degradation byproducts by using a high
performance liquid chromatography (HPLC).
Eluent
The eluent is a mixture of methanol and 0.05 M KH2PO4 (adjusted to pH 3
with HCL) in a ratio of 1:3.
Sample
A liquid sample must be centrifuged before the injection. Approximately 0.25
mL of sample is needed to inject into a 50 tL-1oop injection port. Air bubble in the
syringe must be removed before the injection.
Column
The column used for chromatograph separation is a 150 mm
x
4.6 mm
x
5 pm
platinum EPS C-18 column (Alitech Assoc. Inc., IL). A guard column of the same
resin must be installed prior to the main column.
172
HPLC (Dionex 2000i)
1.
Set the pumping rate to 1.0 mL/min.
2. Turn on the UV detector.
3. Set the wavelength at 254 nm.
4. Set the output range at 0.2 AU.
Integrator (HP3396 Series II)
The setting parameters for the integrator are
1.
Attenuation (ATT 2'") = 8
2. Chart Speed (CHT SP)
1.0
3. Area Rejected (AR REJ) = 10000
4. Threshold (THRSH) = 4
5. Peak width (PK WD) = 0.04
Chromatogram
Separation of chromatograms is shown in Table Al and Figure Al.
Table Al. Approximate HPLC Retention time and peak area of compounds of interest
Retention Time (mm) Approx. Area of 50 mgIL conc.
Compounds
6.5e106
Benzyl alcohol
3.7
21.8e106
2-hydroxybenzyl alcohol
2.7
17.5e106
3-hydroxy benzyl alcohol
2.4
25.7e106
4-hydroxy benzyl alcohol
2.3
RUN
81441
067
*
DEC
2004
4,
11135450
START
IF
c).50mg/L3.HydroxyBenzy1Mcoh
a). 100 mg/L Benzyl alcohol
l,S3
Benzaldehyde
5.032
jR,665
"19ETA0LE STOP
T'IMEIRRLE STOP
PURR
DEC
4,
oøi
DEC IS.
065
200!
1012414
11435154'
AREA",
04:66
N
4080 TYPE
74
3651440
84
11995$
134:2536
P6
2042430
88
PT
AREA ropE
48
473133
11539644 SF0
AT
1.142
0,424
04014
.079'
.044
4,504
2.049
3,698
5.632
WREAN
3,69314'
96,34404
TOTAL ARER*!,8218E,O7
8UL FWCT,D0!.8440E,44
WITTY,
.107
.066
.115
.175
40806
10,47804
.61811
69.09160
4,71641
TOTAL AREU*! .92060+07
MUL FACTOR*I.00600+0R
*
RUN 0
START
868
DEC II, 2001
71105103
+
PUS
866
8
Y
DEC 16,
2221
11:27139
II
4,684
.1
0.720
!
d). 50 mgIL 4. Hydroxy Benzyl Alcohol
.....................
b). 50 mg/L 2-1-lydroxy Benzyl Alcohol
_9====1 2. 316
__,.,,.2ZCD==tO:'==O4===' . 0r'
2. 3D!
8,040
3.643
6.870
5,131
9.460
'TDKETROLE 3101
TIAETW8LE 4TOP
8014*
864
DEC IS,
2441
11,45:23
08046
465
DEC 14. 2001
11,27189
AREA',
RI
o
66410
1.345
2.316
0.931
WRE4 TYPE
460663
88
25662540 OPS
124445
SF8
41056
.402
48003
.7543!
.111
.872
9717198
.47412
RI
1.166
.865
2,458
3,080
AREA TYPE
08
169039
448904
88
21784564 SF8
64427
SF8
TOTAL 08E62.60488*07
AU!, F4CTOR*l.8000E000
TOT0L 6866.2,26774+07
WIDTH
.87!
.450
.494
.807
AREAS
.74544
2.74748
94.48486
.46005'
-4
2. 65!
174
Appendix B: Inhibition of Phenol on Benzyl Alcohol Degradation
and 2HBA Production
Degradation of benzyl alcohol (BA) and production of 2HBA by phenol-grown
cells were investigated in the present of phenol 0, 5, and 45 mg!L. When phenol was
not present, rates of BA degradation and 2HBA production were 0.51 and 0.35
mg!mg-cell/d, respectively (Figure B 1-a). In the presence of 5 mgIL phenol (Figure
B1-b, 0 - 70 mm), BA rate reduced to 0.11 mg!mg-cell/d and 2HBA was not observed.
After phenol was all degraded (Figure B 1-b, 70
200 mm), BA rate increased to 0.41
mg!mg-cellld and 2HBA was observed with production rate of 0.17 mg/mg-cell/d. In
the presence of 45 mg!L phenol, BA rate was 0.15 mg/mg-cell/d and 2HBA was not
observed within experimental time of 200 mm (Figure B 1-c). Phenol degradation
rates were 1.8 and 2.4 mg!mg-cell/d when the initial concentrations were 5 and 45
rng/L respectively.
Similar observations of the phenol inhibition of 2HBA production were
observed with benzyl alcohol-grown cells (Fig. B2). The 2HBA production rate in an
absence of phenol was 0.15 mg!mg-cell/d, while in the presence of 45 mg/L phenol,
the 2HBA production rate was reduced to 0.03 mg/mg-cell/d. The BA degradation
rate was also reduced from 1.3 mg!mg-cell/d in an absence of phenol to 1.0 mg/mgcell/d in the presence of 45 mg/L phenol.
These results showed that phenol inhibited BA degradation and 2IIBA
production and indicated the similarity of phenol hydroxylase and benzyl alcohol
175
ortho-monooxygenase. Inhibition of phenol on BA degradation also suggested the
lower value of K5 in phenol degradation than K in BA degradation.
C
50
5
40
4,_
-
5
40
4-
30
3E
-J
E
30
20
50
0)
a. 20
2
2
0
0
0
50
100
Fig. b).
0
0
200
150
0
50
50
40
4
30
2
0.
0
< 10
1
Fig.
C).
0
g)
150
200
.
Figure B1. Effects of phenol (A) on
benzyl alcohol () degradation and
2HBA (D) production by phenolgrown cells: a) no phenol present;
b) with 5 mg/L phenol; and c) with
mgIL phenol.
0
0
oC
100
Time (mm)
Time (mm)
oC
50
100
150
Time (mm)
200
50
5
4-
40
4-
3E
30
3E
20
2
50
5
40
30
-j
0)
C
a)
20
-J
0)
10
io
0
0
0
C
I
('4
<10
10
50
100
Time (mm)
150
200
0
50
5
40
4-
30
3E
20
2
-j
0)
10
0
0
50
100
Time (mm)
150
200
I
50
100
Time (mm)
150
200
Figure B2. Effects of phenol (A) on
benzyl alcohol (0) degradation and
2HBA (o) production by benzyl
alcohol-grown cells: a) no phenol
present; b) with 5 mgIL phenol; and
c) with 45 mgIL phenol.
176
Appendix C: Haldane Kinetic of TCE Transformation by
Benzyl Alcohol-Grown Cells
TCE transformation by benzyl alcohol-grown cells was investigated at high
TCE concentrations up to 12 mg/L. TCE transformation followed Haldane kinetics.
The achieved kinetic parameters are shown in Table B 1. The specific rates versus
concentration are shown below.
0
0.07
0.06
0.05
IT
0.04
0.03
0.02
0.01
Cl)
0
10
15
TCE (mg/L)
Figure Cl. Haldane kinetic of TCE transformation by resting cells
grown on benzyl alcohol. Line represents Haldane model fits to the
data
Table Cl. Kinetic parameters of Haldane model for TCE transformation by benzyl
alcohol-grown cells.
average ± I std. dev. 95% confidence
0.10 0.18
0.14 ± 0.02
kmax (mg-TCE/mg-cell/d)
0.38-1.10
0.74±0.18
Ks(mg/L)
1.38-3.41
Kh(mg/L)
2.39±0.50
177
Appendix D: Significance of Decay Term in the Transformation Model
The significance of decay term was evaluated with model of TCE transformation
by benzyl alcohol-grown cells, which had the longest experimental period (6 days).
The equations used for the modeling are:
dC
TCE transformation,
+
(
Changes in active cells mass,
kmaxXC
V1
Vg H)
dX= 1-. (Vj+VgHcc)
dt
1.
V1
(Dl)
K + C
dC
.
bX
(D2)
dt
where b is the cellular decay (lid).
A typical decay value of 0.06 1 /d (Metcalf and Eddy, 1991) was applied in the
Equation D2. Applying decay term resulted in less TCE transformed. The difference
in the total amount of TCE transformed between models with and without cellular
decay was about 0.003 8 mg. This amount can be considered insignificant since it was
only 4% of the total TCE transformed (0.09 mg).
5
4
0.05
-J
C)
C)
0.04 g
C.)
0.03
0
C-,
Lii
0.02
'1
0.01
0
I-
ri
0
1
2
3
4
5
6
Time (d)
Figure Dl. Effects of decay term on the modeling of TCE transformation by
benzyl alcohol grown cells. Solid and dotted lines represents model
simulations with b = 0 and 0.06 lid, respectively. Data represent mean and
standard deviation of triplicate values.