AN ABSTRACT OF THE THESIS OF
David M. Doughty for the degree of Master of Science in Microbiology presented
on January 6, 2003. Title: Metabolism of Dichioroethenes by the Butaneoxidizing Bacterium 'Pseudomonas butanovora'.
Redacted for privacy
Abstract approved:__
Peter J. Bottomlèy
Reductive dechlorination of chlorinated ethenes is dependent upon suitable
substrates promoting microbial activity and creating anaerobic conditions. At the
periphery of active reductive dechiorinating zones combinations of lesser
chlorinated ethenes should exist along with end products of the anaerobic
metabolism that is driving reductive dechlorination. Potential end-products of
anaerobic metabolism were investigated for their ability to stimulate oxidative
cometabolism of dichloro ethenes (DCEs) by the butane-degrading bacterium,
Pseudomonas butanovora. Organic acids that supported butane-monooxygenase
(BMO) activity were acetate, propionate, lactate, and butyrate. Lactate
consistently supported and sustained greater rates of cooxidation than did the other
organic acids. When propane replaced butane as the growth substrate, lactate
remained the superior electron donor, while the ability of butyrate and acetate to
support BMO activity decreased. In contrast, propionate-supported cooxidation
was only observed in propane-grown cells.
Lactate supported the degradation of 1,2-trans dichioroethylene (1,2-trans
DCE), 1,2-cis dichloroethylene (1,2-cis DCE) and 1,1-dichioroethylene (1,1-DCE)
in butane-grown P. butanovora. 78 nmoles (25 tM) of 1,2-cis DCE were
completely degraded by butane-grown P. butanovora. In contrast, smaller
amounts of 1,1 -DCE and 1,2-trans DCE were degraded over the twenty minuet
time course. Decreasing rates of cooxidation over time were observed for of all
three DCEs, and 50% of BMO activity was irreversibly lost after 15 mm, 6 mm,
and 0.5 mm exposures to 1,2-cis DCE, 1,2 trans-DCE, and 1,1-DCE respectively.
Cell viability decreased by over 90, 95, and 99.95% during the transformation of
25 nmoles/mg protein of 1,2-cis DCE, 1,2-trans DCE and 1,1-DCE. These results
indicate that cellular viability was more sensitive to cooxidation of 1 ,2-cis DCE
and 1,2-trans DCE than was BMO.
1,2-cis DCE and 1,2-trans DCE induced BMO activity to 25 and 45% of
the butane control, respectively. Induction by 1,2-trans DCE was observed at a
threshold of about 20 M and higher concentrations did not increase BMO
activity. Fusion of lacZ to the BMO catabolic promoter, with consequent knock
out of BMO activity, provided the opportunity to assess substrate induction
without the confounding effects of enzyme inactivation and product induction.
While BMO substrates, butane, I ,2-cis DCE, and ethylene, were unable to induce
lacZ activity the BMO products, 1 -butanol, and ethylene oxide, effectively induced
lac Z activity. 1,2-trans DCE was unique among the BMO substrates tested in it's
ability to induce expression of lacZ, 2-fold above background, in the reporter
strain. A wide range of concentrations induced lacZ activities (10 to 100 SM), and
low levels of 1,2-trans DCE achieved high levels of induction after 4 hrs.
However, lac Z activities were limited to an induction of about four-fold above
background and this limit allowed lower concentrations of 1,2-trans DCE to
eventually produce equal levels of beta-galactosidease. These data provide proofof-concept that BMO-dependent cometabolism can occur independently of butane
as an inducer and electron donor for BMO gene expression and activity.
©Copyright by David M. Doughty
January 6, 2004
All Rights Reserved
Metabolism of Dichioroethenes by the Butane-oxidizing Bacterium
'Pseudomonas butanovora'.
by
David M. Doughty
A THESIS
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Master of Science
Presented January 6, 2004
Commencement June, 2004
Master of Science thesis of David M. Doughty presented on January 6, 2003.
APPROVED:
Redacted for privacy
Major Professor,
Microbiology
Redacted for privacy
Head of the Department of$obiology
Redacted for privacy
Dean of th
Ilri6uate School
I understand that my thesis will become part of a permanent collection of Oregon
State University libraries. My signature below authorizes release of my thesis to
any reader upon request.
Redacted for privacy
DaviMt1hty, Author
ACKNOWLEDGEMENTS
I would like to express my sincere gratitude to Dr. Peter J. Bottomley.
Without your experience and patience I would not have been able to complete my
project. The questions you have asked have directed my work and development as
a scientist.
I have appreciated the opportunity to work so closely with Dr. Daniel Arp
and his lab. Thank you for your commitment. It has been a pleasure working with
you.
A special thanks to Dr. Luis Sayavedra-Soto who selflessly provided his
time and assistance throughout this degree. I have enjoyed our discussions of
science and appreciate your encouragement in my professional pursuits. To the
rest of the Arp and Bottomley labs, I have enjoyed working with all of you. Thank
you for the creative and interesting discussions that we have had.
Finally, I would like to thank my parents Michael and Lillian Doughty for
their encouragement and support.
CONTRIBUTION OF AUTHORS
Peter J. Bottomley, Daniel J. Arp aided in the experimental designs and the
creation of this manuscript. Luis A. Sayavedra-Soto provided bacterial strains and
aided in the design of experiments. All experiments and analysis were conducted
in the laboratories of Dr. Arp or Dr. Bottomley at Oregon State University.
TABLE OF CONTENTS
Chapter 1. ThTRODUCTION ................................................ 1
1.1. DCEs-industrial use, primary contaminants, and
daughter products of reductive dechlorination .................
1.1.1. Industrial use of chlorinated ethenes .....................
1.1.2. Primary ground water pollution by DCEs ...............
1.1.3. Reductive dechlorination and the creation of
DCEs .........................................................
1.2. Cooxidation .........................................................
1.2.1. Electron donation, monooxygenase induction
and the competition of natural substrates in
cooxidation ..................................................
1.2.2. Electron donation by alternative substrates .............
1.2.3. Induction of non-specific oxygenases by chlorinated
ethenes .......................................................
1.3. Alkane oxidizing bacteria and multicomponent
oxygenases ..........................................................
1
1
3
3
3
4
5
6
1.3.1. Induction of butane metabolism in P.
butanovora ...................................................
8
1.3.2. Inductions of pathways involved in the metabolism of
alternative electron donors ..................................
8
1.4. Research objectives .....................................................
9
Chapter 2. MATERIALS AND METHODS ............................
10
2.1. Bacterial strains and growth conditions .........................
10
2.2. Ethene oxidation assay .............................................
11
2.3. Degradation of chlorinated ethenes ..............................
11
TABLE OF CONTENTS (CONTINUED)
Page
2.4. Irreversible effects of DCE cooxidation on BMO ............
12
2.5. Inhibition of 02 uptake by DCEs ................................
13
2.6. Determination of cell viability after cooxidation ..............
14
2.7. Induction of BMO by chlorinated ethenes .....................
14
2.8. Induction of the LacZ reporter strain ...........................
15
2.9. Analytical and other methods ....................................
16
Chapter 3. RESULTS .......................................................
17
3.1. Reductant enhanced cooxidation and metabolic fitness in
P. butanovora ......................................................
17
3.2. Characteristics of chlorinated ethene degradation in P.
butanovora ..........................................................
22
3.3. Induction of BMO activity in P. butanovora by
chlorinated ethenes ................................................
32
Chapter 4. DISCUSSION...................................................
38
BIBLIOGRAPHY ............................................................
45
LIST OF FIGURES
Figure
3.1.
3.2.
Page
Concentration effects of electron donor on the cooxidation
of ethene by butane-grown P. butanovora ........................
Effect of the alternative electron donors, 5mM butyrate or
lactate, on the consumption of 1 ,2-cis DCE by butane-grown P.
butanovora
3.3.
3.4.
3.5.
3.6.
3.7.
3.8.
19
.............................................................
20
Time course of 1,2-cis DCE, 1,2-trans DCE, and 1,1-DCE
consumption by butane-grown P. butanovora ..................
25
Time course of the irreversible loss of BMO activity during
the consumption of 1,2-cis DCE, 1,2-trans DCE, and 1,1
DCE by butane-grown P. butanovora ...........................
27
Time course of lactate-dependent oxygen consumption by
butane-grown P. butanovora during the cooxidation of 1,2cis DCE, 1,2-trans DCE, and 1,1-DCE ...........................
29
The effect of equimolar (25nmoles/mg protein) consumption
of 1 ,2-cis DCE, 1,2-trans DCE, 1,1 DCE on cell viability in
butane-grown P. butanovora ......................................
31
Induction of BMO activity in lactate-grown cells of P.
butanovora in response to exposure to butane or varying
concentrations of 1,2-trans DCE ..................................
34
Induction of lac Z in the reporter strain by varying
concentrations ofi -butanol, 1,2-trans DCE, 1 ,2-cis DCE,
and1,1-DCE .........................................................
35
LIST OF TABLES
Table
3.1
3.2.
Page
Reduc ant enhanced oxidation of ethene to ethene oxide
by butane and propane-grown P. butanovora...................
21
Induction of BMO activity in P. butanovora and lacZ activity
in the lacZ-BMO fusion in response to various substrates
and products of BMO ................................................
37
Metabolism of Dichioroethenes by the Butane-oxidizing Bacterium
'Pseudomonas butanovora'.
Chapter 1. INTRODUCTION
1.1.
DCEs - industrial use, primary contaminants, and daughter products
of reductive dechlorination
1.1.1. Industrial use of chlorinated ethenes
Chlorinated ethenes are used in a wide range of industrial processes and in
the creation of consumer goods. Vinyl chloride (VC), 1,1 dichloroethene (1,1
DCE), trichioroethene (TCE) , and tetrachloroethene (PCE) are commonly used
chlorinated ethenes. TCE and PCE are used as organic solvents for the degreasing
of metal machinery, and as dry cleaning fluids. Vinyl chloride and 1,1 -DCE will
undergo polymerization to polyvinyl chloride and polyvinylidene chloride
respectively, and are used in furniture coverings and food wraps. Mixing these
compounds with each other or TCE will cause different chain linking in the
polymer and produce plastics with different properties. Industrial demands for
TCE in the United States alone topped 171 million pounds in 1998 and continues
to grow by 7-9% a year (Agency for Toxic Substances and Disease Registry
2003).
2
1.1.2
Primary pround water pollution by DCEs
Chlorinated ethenes are suspected carcinogens and known neuro
depressants (Agency for Toxic Substances and Disease Registry, 1995). The EPA
has regulated the concentration of chlorinated ethenes in drinking water since 1994
(Agency for Toxic Substances and Disease Registry 2003). Prior to their
regulation, industrial wastes containing high levels of chlorinated ethenes were
disposed of haphazardly. These disposal practices made chlorinated ethenes
common ground water contaminants in urban areas. EPA standards for safe
drinking water limit the concentration of chlorinated ethenes to 2ppb, 5ppb, and
7ppb for vinyl chloride, 1,1-DCE, and TCE respectively; EPA standards for 1,2-cis
DCE and 1,2- trans DCE are about ten-fold higher (Agency for Toxic Substances
and Disease Registry, 1997).
1.1.3. Reductive dechlorination and the creation of DCEs
Many different organisms are known to metabolize and cometabolize
chlorinated ethenes (Chauhan et al. 1998; Coleman et al. 2002 a and b; Major et al.
2002; Sullivan et al. 1998; Verce 2001; Verce 2002; Yeager et al. 2001). Some
organisms are able to use chlorinated ethenes as terminal electron acceptors during
growth under anaerobic conditions. This ability results in stepwise reductive
dechlorination according to the following pathway: PCE, TCE, DCEs, VC, and
ethene. The degradation of chlorinated ethenes becomes stereochemically and
thermodynamically restricted resulting in buildup of lesser-chlorinated ethenes
3
(Semprini, 1995). The persistence of DCE and VC leads to their eventual
diffusion away from the contaminated site. As a result, low concentrations of
DCE and VC may persist in the groundwater adjacent to sites experiencing
reductive dechlorination of more highly chlorinated ethenes (Freedman et al. 2001;
Kastner 1991).
1.2.
Cooxidation
Aerobic cooxidation occurs when a non-specific oxygenase enzyme
converts chlorinated ethenes into epoxides or alcohols. Several characteristics of
cooxidation limit this process. First, enzymatic attack by oxygenases on
chlorinated ethenes drains the reductant needed for other cellular processes.
Second enzyme inactivation brought about by successful cooxidation can limit the
amount of chlorinated ethenes degraded. As a result the sustained cooxidation of
chlorinated ethenes requires the presence of an appropriate substrate to induce
monooxygenase activity and requires a suitable electron donor to support
monooxygenase activity (Arp et al. 2001).
1.2.1. Electron donation, monooxygenase induction and competition
of natural substrates in cooxidation
Induction of non-specific oxygenases required for the cooxidation of DCEs
usually involves the natural substrate. Under these conditions, the natural substrate
also serves as the growth substrate, and indirectly as the electron donor for
chlorinated ethene cooxidation. However, the metabolism of natural substrates
supplied to enhance cooxidation of chlorinated ethenes competes with the
cooxidative process. Kim et al. (2002a) showed that the degradation of 1,2-cis
DCE and 1,1 -DCE by a butane-grown mixed culture proceeded more slowly in the
presence of butane because the transformation of the growth substrate limited the
allocation of reductant to the cooxidation reaction (Alvarez-Cohen and McCarty
1991 a and b; Kim et al. a and b). Under these conditions the transformation yield,
the ratio of electrons allocated to cooxidation versus other cellular processes, may
be low.
1.2.2. Electron donation by alternative substrates
Conceptually, competition between natural substrates and cosubstrates may
be averted by supplying cells with an alternative electron donor. The
transformation yield in these systems may be very high as long as cosubstrate
availability and enzyme activity are not limiting. This concept has been
highlighted in methanotrophic systems where formate, a downstream intermediate
of methane metabolism, was used to support oxidative cometabolism (Alvarez-
Cohen and McCarty 1991a; Arp et al. 2001). In these systems, alternative electron
donors allowed kinetic and cytotoxic effects to be studied without the confounding
effects of inhibition by the natural substrate. Exogenous electron donors may lead
to faster initial rates of degradation, however, enzyme activity will decline because
of either protein turnover, or inhibition of the enzyme by the cosubstrate.
1.2.3. Induction of non-specific oxygenases by chlorinated ethenes
Induction of non-specific monooxygenase enzymes by their cosubstrates
has been reported in a variety of organisms (Ensign 1996; Ryoo et al. 2001;
Yeager et al. In Press 2004). Chlorinated ethene induction of a monooxygenase
circumvents the need for a "natural" substrate, and, conceptually, relieves the
competition between the natural substrate and the cosubstrate. This permits a
greater percentage of reductant to be allocated to chlorinated ethene cooxidation
and supports a higher transformation yield. Enzyme induction by chlorinated
ethenes has been most extensively studied in toluene-oxidizing bacteria.
A 3-fold induction of monooxygenase activity was observed following
exposure of Pseudomonas stutzeri OXI to 100 pM TCE or PCE. Toluene
monooxygenase activity was not observed following exposure to other chlorinated
ethenes including DCEs. Transfer of the inducible monooxygenase system to a
heterologous host produced an inducible monooxygenase system that responded to
the natural substrate, but did not respond to PCE or TCE. It was concluded that
factors outside the previously known genes and gene products were required for
induction by TCE or PCE relative to toluene (Ryoo et al. 2001). Four tolueneoxidizing bacteria were evaluated for the range of electron donors able to support
the degradation of TCE when TCE was also used as the inducing compound. The
ability of Pseudomonas mendocina KR1, Pseudomonasputida Fl, and Raistonia
pickettii PKOI to degrade TCE depended on the growth-supporting substrate.
Differences in the ability of electron donors to support cooxidation were likely due
to catabolite repression or sensitivity to the cooxidative process. Various carbon
sources supported different rates of degradation and growth, leading to speculation
that the allocation of reductant varied by organism and by electron donor. P.
mendocina KR1 degraded TCE in the presence of lactate, glucose, fructose, and
glutamate, but not in the presence of acetate or Luria-Bertani broth. These results
showed P. mendocina KR1 to be the most versatile in its ability to use growthsupporting substrates as electron donors for the cooxidation of TCE (Yeager et al.
In Press, 2004).
1.3. Alkane oxidizing bacteria and multicomponent oxygenases
Alkane oxidizing bacteria may be grouped empirically according to the
chain length of the alkanes used for growth. Methanotrophic bacteria comprise the
first group, and two representative organisms, Methylococcus capsulatus Bath and
Methylosinus trichosporium OB3b, have been the subject of intense study
(Stafford et al. 2003; Csaki 2003; Coufal et al. 2000). Particulate methane
monooxygenase (pMMO) is a membrane-bound enzyme capable of converting
methane to methanol. Under conditions of copper limitation methanotrophs may
express a soluble cytoplasmic methane monooxygenase (sMMO). pMMO and
sMMO are representative of two families of non-specific oxygenases that have
been adapted by organisms for growth on a variety of substrates and have been
exploited for bioremediation of chlorinated ethenes (Sullivan et al. 1998). Alkanes
7
of C2 to C4 in chain length are growth substrates for another group of alkane
oxidizing bacteria (Hamamura et al. 1999). Bacteria in this group use both sMMO
and pMMO homologs to oxidize alkanes. P. butanovora is a gram negative
representative of this group and the subject of study in this thesis. At the other
extreme, alkane oxidizing bacteria also exist that grow on liquid alkanes of chain
lengths C5-C 12. Pseudomonas putida GPo 1 (formerly Pseudomonas oleovorans)
relies on a membrane bound enzyme analogous to pMMO and has been studied as
a model system for the expression and regulation of alkane-oxidizing genes
(Dinamarca et al. 2002).
Sluis et al. (2002) sequenced the BMO gene cluster and compared the
deduced amino acid sequences with other multi-component monooxygenases.
BMO subunits shared as much as 64% identity with sMMO. The high sequence
identity suggests that BMO is a member of a family of non-specific oxygenases
called three component hydroxylases. The substrate binding pocket and active site
are located on the hydroxylase subunit (Coufal et al. 2000), while the regulatory
subunit is associated with substrate sensing and regio-specificity of hydroxylation
(Astier et al. 2003). Both subunits are conserved features of this family of
oxygenase enzymes. Despite similarities in sequence between sMMO and BMO,
methane oxidation and the production of 2-butanol during butane oxidation are
characteristics of sMMO not shared with BMO (Sluis et al. 2002). These results
suggest that substrate range of BMO is more restricted than for sMMO. The
reflection of these trends in the kinetics of cooxidation of the three DCE isomers
will be discussed later in this thesis.
1.3.1. Induction of butane metabolism in P.
Pseudomonas butanovora
butanovora
is able to use butane as the sole source of carbon
and energy necessary for growth. Although lactate or citrate-grown P.
butanovora
cultures do not consume butane, butane consumption may be induced by butane
(Sayavedra-Soto et al. 2001). Exposure of a BMO-minus mutant to butane,
however, did not stimulate the consumption of 1-butanol, a down-stream
intermediate in butane metabolism. These results indicate that the enzymes
involved in the downstream oxidation of butane are regulated by mechanisms
different from the BMO gene cluster (Sayavedra-Soto et al. 2001).
1.3.2. Induction of pathways involved in the metabolism of alternative
electron donors
Inducible metabolic pathways will influence the sources of reductant that
may be used to drive monooxygenase activity. Two inducible alcohol
dehydrogenases expressed during growth on butane were determined to be
controlled by separate regulatory mechanisms (Vangnai et al. 2001; Vangnai et al.
2002 a and b) The first alcohol dehydrogenase was expressed in response to 1butanol or butane, the second alcohol dehydrogenase was expressed in response to
butane or lactate. The regulation of other metabolic pathways in P.
have not been elucidated.
butanovora
1.4.
Research objectives
TCE and DCEs are some of the most common ground water pollutants in
the United States. Reductive dechlorination of TCE and PCE has enjoyed
considerable popularity because it is a sustainable and growth supporting process
(Maymo-Gatell et al. 2001; Beeman and Bleckman 2002; Major et al. 2002).
However, the partially dechlorinated end products of reductive dechlorination can
persist as toxic contaminants of ground water. Successful reductive dechlorination
depends on intense microbial metabolism that creates anaerobic conditions, which
may result in the formation of organic acids and alcohols as metabolic end
products (Kastner 1991). The objectives of the current work were to determine the
extent that aerobic cooxidation may be supported by putative fermentative end
products of anaerobic processes, and to examine the range of chlorinated aliphatics
that induce BMO in P.
butanovora.
'[C
Chapter 2. MATERIALS AND METHODS
2.1. Bacterial strains and growth conditions
P. butanovora
liquid cultures were grown in 750 ml flasks that were sealed
with butyl rubber stoppers and contained 200 ml of liquid medium. Flasks were
incubated at 300 C with constant circular shaking (180 rpm) for two days. Butane
and propane gas (99% Airgas Randor, Penn.), were sterilized by passing through a
0.2 jm filter. When butane or propane served as the growth substrate, cultures
were supplied with an overpressure of 7% volume! total volume (or 10%
headspace) of the gas. When organic acids served as the carbon source for growth,
a concentration of 10 mM was used. Minimal media used during growth on either
alkanes or organic acids consisted of, per liter, 2.7 g, (NH4)2HPO4; 4.3 g,
Na2HPO4; 4.2 g, KH2PO4; 0.5 g, MgSO4*7H20; 0.06 g, CaC12*2H20; and lml
trace element solution (Wiegant and de Bont, 1980) and adjusted to pH 7.1.
Cells were sedimented by centrifugation (10 mm at 6000 rpm and 10°C)
after cultures reached early stationary phase, and washed three times in 50 mM
phosphate buffer (as described for the media except potassium and sodium
phosphate concentrations were adjusted to compensate for the omission of
(NH4)2HPO4). Harvested cells, kept as a concentrated cell suspension on ice,
retained ethene-dependent ethene oxide production for greater than 3 hours
without observable loss in activity. Fresh cells were harvested daily or when
needed.
11
2.2. Ethene oxidation assay
Ethene dependent ethene oxide production served as the standard measure
of BMO activity (Sayavedra-Soto et al. 2001). Vials (10 ml) sealed with butyl
rubber stoppers and aluminum crimp caps served as the reaction chamber for this
assay. Vials contained 1 ml of whole cell suspensions (no more than 0.35 mg of
cellular protein), the specified electron donor, and 20% (v/v) ethylene gas was
added as over pressure to start the assay (Hamamura et al. 1999). Vials were
agitated (150 rpm) at 300 C in a covered water-bath shaker. Ethene oxide
formation was followed by gas chromatography with a Shimadzu GC-8A equipped
with a flame ionization detector and a 120 cm Porapak Q column (Ailtech
Associates, Inc.). The colunm temperature was set at 120° C and the injector and
detector were maintained at 200 °C as previously described (Hamamura et al.
1999). Samples (100 jtl) of the headspace were taken with a gas tight syringe at
the time intervals of 0, 2.5, 5, and 7.5 mm. Assays that determined BMO activity
were limited to less than 10 mm because the product ethene oxide has been
associated with BMO inactivation.
2.3. Degradation of chlorinated ethenes
The degradation of chlorinated ethenes was followed by measuring their
disappearance from the headspaces by gas chromatography. Abiotic controls,
which did not receive P. butanovora, and butane-grown P. butanovora cells
pretreated with acetylene, a mechanism based inhibitor of BMO (Hamamura et al.
12
1997), were used to ensure that the observed disappearance of the chlorinated
ethene was due to the activity of the BMO enzyme. Vials (10 ml) were prepared
containing phosphate buffer (900 p1) and the specified electron donor. Aliquots of
a DCE saturated solution were injected into the vials sealed with Teflon-lined
butyl rubber stoppers and allowed to equilibrate by shaking at 150 rpm and 30° C
for 30 mm prior to the start of the assay. Assays were initiated by the injection of
100
/Ll
of concentrated cell suspension. Protein concentration in the cell
suspensions were determined using the biuret assay (Gornall et al. 1949) and
usually ranged from 0.7 to 1 mg protein per vial.
2.4. Irreversible effects of BCE cooxidation on BMO
Vials were prepared as described above. Accurate assessment of timedependent irreversible inactivation required the prompt interruption of the
cooxidation reaction. To accomplish this, butane was flushed into the headspace
of the vial. Conceptually this prevented the continued degradation of the DCE
during the washing procedure by purging oxygen and DCE out of the vial while
providing the natural substrate to compete for the BMO active site. Two 60 ml
syringes were used, the first syringe was kept empty with the plunger suppressed,
and the second syringe was filled with butane. Both syringes were inserted into
the vial and pulling back the plunger of the empty syringe pulled the butane from
the second syringe through the vial. Cells were then washed three times by
microcentrifuge, resuspended in phosphate buffer and tested for residual BMO
13
activity using the ethene oxidation assay. Control vials that did not receive a
chloroethene were also flushed with butane.
2.5. Inhibition of 02 uptake by DCEs
Lactate-dependent 02 uptake was followed using a Clark-style 02 electrode
with glass reaction vial equipped with an YSI Model 5300 Biological Oxygen
Monitor (Yellow Springs Co., Yellow Springs, OH). Temperature of the reaction
chamber was maintained at 30° C by a circulating water jacket. Phosphate buffer
(1.75 ml, 50mM) was allowed to equilibrate with air for 5 mm before the start of
each assay, and the reaction chamber was washed 20 times with water and twice
with phosphate buffer between assays to remove residual carbon sources and other
contaminants. The reaction chamber was capped, and additions were made by
injection in the following order: butane-grown P. butanovora cells (0.2 mg
protein), lactate (2mM in electrode chamber), and the indicated cosubstrate (25
M in electrode chamber). This order of addition allowed for the sequential
determination of abiotic 02 consumption, resting cell 02 consumption, lactate
dependent 02 consumption, and the effects of the cosubstrate. Control reactions
received the same treatments except cells were first treated with acetylene to
inactivate BMO.
14
2.6. Determination of cell viability after cooxidation
Cell viability after treatment with chlorinated ethenes was determined by
dilution plating, on R2A agar, pH 7.2 (Becton, Dickerson and Company, Sparks
MD). Cells were treated as described for the degradation of chlorinated ethenes,
except, a sterile 0.5 ml microfuge tube was inserted into the vials to serve as a
reservoir for the nonsterile DCEs. DCE solutions were injected into the open end
of the microcentrifuge tube and allowed to equilibrate with the phosphate buffer
outside of the tube. Assays were initiated by injecting cells, and incubated with
constant shaking for 20 mm. Controls included acetylene-treated cells to
inactivate BMO, and control cells that did not receive DCE.
2.7. Induction of BMO by chlorinated ethenes
Cells were grown in 10 mM lactate and the minimal medium described
above. Induction vials received aliquots of the indicated compounds and were
allowed to reach equilibrium by circular shaking at 30° C for 30 mm prior to the
start of the experiment. Uninduced cells were resuspended in fresh ammonium
phosphate buffer plus minimal medium and were aliquoted into 10 ml vials with
900
/Ll
of growth media plus ammonium phosphate buffer, sealed with Teflon
coated butyl rubber stoppers and aluminum crimp caps. Unless otherwise
indicated, all inductions were 4 h in length, and included positive butane and
negative buffer controls. After 4 h, cells were harvested in a microcentriflige and
tested for BMO activity using the ethene oxidation assay described above.
15
2.8. Induction of the LacZ reporter strain
Dr. Sayavedra-Soto provided a reporter strain with a bicistronic expression
system in which kanamycin resistance was constitutive and the BMO catabolic
promoter controlled lacZ expression. The lacZ fusion provided multiple in-frame
transcriptional stop codons before the beginning of the BMO structural genes that
effectively eliminated BMO activity. The reporter strain was grown on citrate in
the same growth medium and buffer as the wild type cells except that kanamycin
sulfate (1 jzglml) was added to the medium to maintain the insert containing the j3-
galactosidase gene. Inductions were performed in 1 OmI vials with Imi of the
reporter strain culture diluted with growth media to an OD600 of 0.3. All vials
were sealed with butyl rubber stoppers and aluminum crimp caps. Inductions were
2, 4, or 6 hours in length and were initiated by the injection of cells. j3galactosidase activity was determined by adding 2 jil SDS (10%), 2 jil 3mercaptoethanol, 4 jig ortho-nitrophenyl j3-D-galactopyranoside (ONPG), and a
drop of chloroform are expressed in Miller units (Miller 1972). /3-galactosidase
assays were incubated at 36°C and stopped by the addition of 200 jil 5% calcium
carbonate. All activities were calculated using the following equation to
compensate differences in the duration of fl-galactosidase assays and the number
of cells! ml that were assayed. The variables are, OD 420
ONPG absorption; OD
600
the wavelength of
the cell density during induction; OD
turbidity after cell lysis; and time is in hours (Miller 1972).
540
is the
16
1
= Miller Units
OOD420 -1.75(OD54
L
2.9. Analytical and other methods
Analytical/HPLC grade PCE, TCE, 1,2-trans DCE, 1,2-cis DCE, 1,1-DCE
were acquired from Sigma and Aldrich (St. Louis, Mo.). PCE, TCE, 1 ,2-cis DCE,
1,2-trans DCE, and 1,1 -DCE saturated solutions were prepared by agitating two
phase solutions (aqueous/chlorinated ethene) for 24 h with constant circular
shaking. The saturated aqueous phase was used to aliquot chlorinated ethenes into
the reaction vials. Concentrations of chlorinated ethenes in the liquid phase of
reaction vials were assumed to follow the unitless Henry's constant (Hc) displayed
in equation 1 (Gossett 1987) with volume (V) in liters and concentration (C) in
molarity. The unsaturated DCE solutions were created based on calculation given
in equation 2.
Equation 1:
CI
Cg= HC
Equation 2:
Moles
VICI+ V9C9
Moles
C9
HcVg+Vi
17
Chapter 3. RESULTS
3.1. Reductant enhanced cooxidation and metabolic fitness in P. butanovora.
An earlier report indicated that the cooxidation of 1 ,2-cis DCE by butane-
grown P. butanovora was dependent upon exogenous reductants, but did not
evaluate differential effects of alternative electron donors on oxidative
cometabolism (Hamamura et al. 1997). Cooxidation of 1,2-cis DCE by butanegrown P. butanovora was enhanced to different extents by 5mM lactate or
butyrate. The cooxidation of 1,2-cis DCE was enhanced less by butyrate than by
lactate in butane-grown cells (figure 3.2), and these differences became more
exaggerated over time (figure 3.2). For example, 10 to 15 nmoles of 1,2-cis DCE
were degraded in 20 mm. when butane-grown cells were supplied butyrate. In
contrast, > 40 nmoles of 1 ,2-cis DCE were degraded over the same time interval,
when cells were provided with the same concentration of lactate. The electron
donors, lactate and butyrate, were tested across a concentration range from 2 to 10
mM (figure 3.1). The differences between the two compounds in supporting
ethene dependent ethene oxide production over the ten mm assay was independent
of concentration.
Evidence was obtained for differences among the pathways responsible for
allocating reductant to BMO from different electron donors. In butane-grown
cells, acetate, lactate, and butyrate enhanced the rate of ethylene cooxidation, from
an endogenous rate of 3.6 nmol/mmn*mg protein' to 17, 20, and 13 nmol/min*mg
EI
protein', respectively (Table 3.1). The same organic acids were tested for their
ability to enhance ethene dependent ethene oxide production in cells grown on
propane. The cooxidation of ethene by propane-grown cells was enhanced from
an endogenous rate of 1.5 nmol/min*mg protein' to 7.8, 17.2, 8.2, and 5.4
nmol/min*mg protein1 by acetate, lactate, propionate, and butyrate, respectively
(Table 3.1). These data clearly illustrate that lactate supported similar rates of
cooxidation in both types of cells. In contrast, specific differences in the ability of
the other organic acids to support ethene cooxidation were observed between
butane and propane-grown cells. Propionate-supported cooxidation was observed
to be 5.5 times greater than the background in propane-grown cells, but did not
support cooxidation above the background rate in butane-grown cells. Thus, equal
concentrations of different organic acids produced unequal rates of cooxidation,
and for several organic acids these trends were affected by the alkane used as a
growth substrate. In contrast, lactate-supported cooxidation displayed consistent
rates of activity regardless of which alkane the cells were grown on.
19
20
215
p
10
-
o
E
>1
5
Butyrate (mM)
Lactate (mM)
Figure 3.1. Concentration effects of electron donor on the cooxidation of ethene
by butane-grown P. butanovora. Paired bars represent different replicates
performed with the same culture.
25
C)
a.
05
E
V
S.-.
C)
VlS
0)
C)
w
C-)
010
U)
9
c.l
U)
C)
E
0
0 tolO mm.
10 to 20 mm.
Figure 3.2. Effect of the alternative electron donors, 5mM butyrate (black) or
lactate (gray), on the consumption of 1 ,2-cis DCE by butane-grown P. butanovora.
The Y-axis depicts the amount of 1 ,2-cis DCE degraded within the two selected
time intervals labeled on the X-axis.
21
TABLE 3.1. Reducant enhanced oxidation of ethene to ethene oxide by butane
and propane-grown P. butanovora.
Reductant
(5mM)
Ethene oxide produced (nmol/min*mg of protein')
(Values in parentheses represent
fold-increase above background)
Butane-grown
Buffer
Formate
Acetate
Propionate
Lactate
Butyrate
3.6
0.7
17.0
3.4
20.1
13.2
Propane-grown
1.5
0
(4.7)
(1.0)
(5.9)
(3.7)
7.8
8.2
17.2
5.4
* Each reaction mixture contained ethylene (25% vol/vol) and the indicated
reductant source.
(5.2)
(5.5)
(11.5)
(3.6)
22
3.2. Characteristics of chlorinated ethene degradation in P. butanovora.
Butane-grown P. butanovora consumed 1,2-cis DCE 1,1-DCE and 1,2trans DCE (Figure 3.3). Lactate (5mM)-supported cooxidation proceeded at
different rates for each of the DCE isomers. The rate of degradation of 1,2-cis
DCE and 1,1 -DCE and within 45 mm the degradation of each compound had
ceased. Reversible inhibition of oxidative cometabolism was distinguished from
irreversible inactivation by resuspending cells in fresh buffer and assaying BMO
activity using the ethene oxidation assay described previously. However,
decreases in BMO activity due to damage to the BMO enzyme and damage to
other cellular processes could not be distinguished. Time-dependent irreversible
BMO inactivation was observed upon exposure of butane-grown cells to each
DCE. Cooxidation of each of the DCE isomers produced different rates of
irreversible inactivation. Half of BMO activity was irreversibly lost after 15 mm,
6 mm, and 0.5 mm exposures to 1,2-cis DCE, 1,2-trans DCE and 1,1-DCE,
respectively (figure 3.4). 1,2-cis DCE and 1,2-trans DCE did not completely
inactivate BMO.
1 ,2-cis DCE and 1,2-trans DCE were degraded in larger amounts, albeit at
different rates, .than was 1,1 DCE. To compensate for these differences among the
DCEs, only 25 nmoles/mg protein of each DCE was allowed to be degraded
during the determination of non-specific cytotoxic effects. Two aspects of
cellular health, namely, lactate-dependent 02 consumption and viability were used
to evaluate these cytotoxic effects in butane-grown P. butanovora. Initial rates of
23
lactate dependent 02 consumption were consistently 69 nmoles/min*mg protein
in cells either expressing active BMO or inactive BMO (acetylene treated cells).
Initial rates of 02 consumption were maintained for 20 mm in cells expressing
active BMO in the absence of a cosubstrate, and in acetylene treated cells after the
addition of 1 ,2-cis DCE, 1,2-trans DCE and 1,1 -DCE. In contrast, the addition of
DCEs increased the rate of lactate-dependent 02 consumption from 27 to 80
nmoles/min*mg protein', or within the range of monooxygenase activity found in
butane-grown cells. Following the initial increase in 02 consumption, lactatedependent 02 consumption decreased rapidly after exposure to 1,1 -DCE and no
lactate-dependent 02 consumption remained after 20 mm. Lactate-dependent 02
consumption also decreased rapidly during the cooxidation of 1,2-trans DCE and
after 20 mm lactate-dependent 02 consumption had dropped from an initial rate of
69 to about 15 nmoles mirilmg protein. In contrast, cells that cooxidized 25 LM
1,2-cis DCE retained 30 nmoles/ mmn*mg protein' of lactate-dependent 02
consumption after 20 mm (figure 3.5). Nonetheless, when 1,2-cis DCE 1,2-trans
DCF or 1,1 -DCE was provided as a cosubstrate 90, 95 and >99% of cell viability
were lost, respectively. Pretreatment with acetylene eliminated DCE degradation
by butane-grown P.
butanovora
and prevented the loss of viability of cells during
exposure to 1,2-cis DCE, 1,2-trans DCE, and 1,1 DCE. The quantities of 1,2-cis
DCE and 1,2-trans DCE degraded during the determination of cellular viability
was less than the amount degraded in figure 3.1 by the same concentration of cells.
These results for 1 ,2-cis DCE and 1,2-trans DCE are consistent with the
24
observations of Yeager et al. (2001), who described the continued cooxidation of
TCE by non-viable Burkholderia cepacia G4. In contrast, this method could not
identify separate thresholds for the transformation capacity of 1,1 DCE and the
loss in cell viability during 1,1 DCE degradation by P. butanovora.
25
Figure 3.3. Time course of 1,2-cis DCE (A), 1,2-trans DCE (B), and 1,1-DCE (C)
consumption by butane-grown P.
butanovora
(closed squares). Assays were
performed in 10 ml vials containing 1 ml of phosphate buffer amended with 5mM
lactate and 25p.M of the indicated cosubstrate. All assays were initiated with the
injection of 100
jLl
of concentrated cell suspension. Butane-grown cells pretreated
with acetylene, a mechanism based inhibitor of BMO served as the BMO negative
control (closed circles). Loss of cosubstrates from the negative control likely
represents leakage of the cosubstrate from the vial. Data points represent the mean
of three replicates and the error bars represent standard deviations.
26
Fig. 3.3
30
25
w
20
9 15
('4
10
L
5
10
15
20
15
20
15
20
Mm.
F',
30
w
0
25
020
(0
15
'-10
5
5
10
Mm.
30
25
20
0
'-
15
1
-10
5
0
0
5
10
Mm.
27
Figure 3.4. Time course of the irreversible loss of BMO activity during the
consumption of 1 ,2-cis DCE (A), 1,2-trans DCE (B), and 1,1 DCE (C) by butanegrown P.
butanovora.
Incubations were performed as described for the
consumption of DCEs except 50 M concentrations were used to ensure that
cosubstrate was not limiting. Incubations were stopped by flushing the vials with
butane at select time points, and the chlorinated ethenes were subsequently
removed by washing the cells three times with phosphate buffer. Cells were then
assayed for BMO activity by measuring ethene dependent ethene oxide
production. Both the cooxidation of the chlorinated compounds and the ethene
were driven by providing 5mM lactate as an electron donor. Butane-grown cells
that did not receive cosubstrates served as BMO positive controls (closed circles)
and indicated losses of BMO activity that were not attributed to the consumption
of cosubstrates. Butane-grown cells that received cosubstrate are represented as
closed squares. Data points are the mean of three replicates and error bars
represent the standard deviations.
0
N)
(Ti
(J1
0
0
(71
0
Cl
-
0
N)
/mg protein)
(TI
N)
(nmoles ethene oxide/mm
Remaining BMO activity
0
()
0
N)
cJ1
0
0
0
(TI
0
(TI
-
0
N)
(71
N)
Remaining BMO activity
(nmoles ethene oxide/mm
1mg protein)
0
()
0
N)
()i
r
(J1
0
0
Remaining BMO activity
(TI
0
-
(Ti
-
0
N)
/mg protein)
(TI
N)
(nmoles ethene oxide/mm
0
()
29
Figure 3.5. Time course of lactate-dependent 02 consumption by butane-grown
P. butanovora
during the cooxidation of 1,2-cis DCE (A), 1,2-trans DCE (B), and
1,1 -DCE (C). Assays were performed in a Clark style 02 electrode containing
1.75 ml of air-saturated phosphate buffer. Additions to the electrode after capping,
were made by injections in the following order: butane-grown cells, lactate, and
cosubstrate. This allowed for the sequential determination of abiotic 02
consumption (0 nmoles/min), resting cell 02 consumption (about 40 nmole 1mm
*mg protein), lactate dependent 02 consumption (69 nmoles/ min*mg protein1),
and the effect of cosubstrate over the 20 mm time interval. Addition of the
cosubstrate occurred immediately following the determination of lactate-dependent
02 consumption at time zero. Cooxidation-dependent effects may be
discriminated from DCE-dependent effects by comparing butane-grown cells
incubated with cosubstrate (squares) with the acetylene treated, BMO negative
controls (closed circles). Furthermore, DCE dependent effects may be
discriminated from DCE independent effects by comparison with the buffer
control (open circles) that consists of butane-grown cells not supplied with
cosubstrate.
Ni
C
01
-
C
CI
-
0)
CD
Ni
C C
C C C CC 0Ni
Lactate dependent oxygen consumption
(nmoles consumed! min*mg protein1)
C'j
C
NJ
-s
01
C
01
.
0)
0)
-
Ni
C C
C C C CC CNi
_&
actate aepenaent oxygen consumption
(nmoles consumed! min*mg protein1)
Ni
0
0
0
0)
0)
Ni
C 0
C C 0 CC 0Ni
Lactate dependent oxygen consumption
(nmoles consumed! min*mg protein1)
w
e
32
3.3. Induction of BMO activity in P. butanovora by chlorinated ethenes.
Residual BMO activity was preserved better than cell viability during the
degradation of 1,2-trans DCE. The possibility that 1,2-trans DCE induced
production of new BMO enzyme was explored. The same concentrations of 1,2trans DCE that were used to characterize cytotoxic effects (25-50 jIM) were tested
for their ability to induce BMO using a resting cell assay (Sayavedra-Soto et al.
2001). Lactate-grown P. butanovora, with no detectable BMO activity, gained
BMO activity after exposure to 1,2-trans DCE concentrations between 20 and 50
M (figure 3.7). Concentrations of 1,2-trans DCE between 20 and 50 M induced
BMO expression to about 50 % that of the butane control. The range of
chlorinated compounds able to induce BMO activity was examined using the
resting cell assay. 50 /LM 1,2-trans DCE, 1 ,2-cis DCE, and 1 ,2-DCA were found
to induce 47, 25, and 29% of the activity of butane controls, respectively (see table
3.2). In contrast, PCE, TCE and 1,1 DCE did not induce expression of BMO.
Fusion of lacZ to the BMO promoter, with consequent knock out of BMO
activity, provided the opportunity to assess gene expression without the
confounding effects of BMO inactivation or product induction of BMO gene
expression. These data provide evidence that products of the BMO enzyme (1butanol and ethene oxide) may act as inducers of BMO gene expression. The
natural substrate, butane, and the chlorinated cosubstrates, 1,2 DCA and 1 ,2-cis
DCE induced the expression of BMO in the wild type but were unable to induce
expression in the lacZ. These data indicate that the products of cooxidation may
33
be responsible for the induction of the BMO enzyme by 1,2 DCA and 1 ,2-cis
DCE. In contrast, 1,2-trans DCE was able to induce lacZ expression without the
involvement of BMO activity. Butane could not induce expression from the BMO
catabolic promoter in the reporter strain (figure 3.8). In contrast, 1 -butanol, a
downstream intermediate in butane metabolism, was able to induce lacZ
expression (figure 3.8). 1,2-trans DCE was the only chlorinated ethene, and the
only BMO substrate, able to induce the reporter strain. Increasing concentrations
of 1,2-trans DCE produced increasing levels of induction in the reporter strain, and
the induction phenomenon was observed at concentrations of 1,2-trans DCE <20
iiM. Lac Z activity levels in the reporter strain were limited to about a five-fold
induction over background even at high concentrations of DCE. This provided an
upper limit to the induction assays that could eventually be reached even by lower
concentrations of 1,2-trans DCE (see figure 3.8).
34
0.5
0.4
w
0
0.
0.3
C
I0.2
0
E
C
0.1
Butane Buffer
5
10
20
30
40
50
1,2-trans DCE (pM)
Figure 3.7. Induction of BMO activity in lactate-grown cells of P. butanovora in
response to exposure to butane or varying concentrations of 1,2-trans DCE. BMO
activity was measured after four hours using the ethene oxidation assay and is
expressed on the Y-axis as nmoles EtO produced/mini mg protein.
35
Figure 3.8. Induction of lac Z in the reporter strain by varying concentrations of
1-butanol (.), 1,2-trans DCE (.), 1,2-cis DCE (o), and 1,1-DCE (A). Betagalactosidase activities were determined by end-point assays taken at select time
intervals of 2 h (A) and 4 h (B) as described in the materials and methods. Data
points are the means of three replicates and error bars represent standard deviation.
36
A
Fig. 3.8
35000
30000
25000
Cl)
20000
:D
15000
10000
5000
0
0
20
40
60
80
100
60
80
100
pM
B
35000
30000
25000
U)
20000
15000
10000
5000
0
0
20
40
pM
37
Table 3.2. Induction of BMO activity in P. butanovora and lacZ activity in the
lacZ-BMO fusion in response to various substrates and products of BMO.
Compound
Concentration
BMO Activity as
a % of Butane
control
50
50
100
>10
15880
0
Butane
1-Butanol
Butyrate
Reporter Strain Miller
Units Above Background
0
50
0
% headspace
10 % headspace
0
0
ND
4751
50
0
0
50
0
0
1,2-trans DCE
50
47
5068
1 ,2-cis DCE
50
25
0
1,1-DCE
50
50
0
0
29
0
Ethylene
Ethylene Oxide
PCE
TCE
1 ,2-DCA
10
Chapter 4. DISCUSSION
Previous reports have shown that the roles of natural substrates as inducers
of monooxygenase genes and generators of reductant to support monooxygenase
activity may be replaced by chlorinated ethenes and other organic sources (Heald
and Jenkins 1994; Leahy et al. 1996; McClay et al. 1995; Ryoo et al. 2001; Yeager
Ct al In press 2004). This study extends the concept to cooxidation of DCEs by the
alkane-oxidizing bacterium, 'Pseudomonas butanovora'. We believe this
particular example could have applied significance because it is well recognized
that partly dechlorinated products like DCEs may accumulate in sites experiencing
reductive dechlorination, (Freedman et al. 2001; Kastner 1991; Semprini 1995). In
situ generation of fermentative end products might facilitate further degradation of
DCEs by oxidative cometabolism in downstream parts of the contaminated plume
where conditions are aerobic.
The use of exogenous electron donors has allowed researchers to examine
the substrate range and kinetic parameters of oxidative cometabolism, while
avoiding the competitive effects that occur when natural substrates are provided as
electron donors (Dolan and McCarty 1995; Chu and Alvarez-Cohen 1999;
Hamamura et al. 1997, Hamamura et al. 1999, Yeager et al. Biodegradation In
press). For example, whole cell kinetic studies of TCE degradation in
methanotrophic bacteria used formate as the alternative electron donor (Dolan and
McCarty 1995; Chu and Alvarez-Cohen 1999; Henry and Grbic-Galic 1991; Johan
et al. 1997), and by analogy, studies carried out on butane-oxidizing bacteria
routinely used butyrate as an electron donor (Hamamura et al. 1997; Hamamura et
al. 1999). My data showed that while various organic acids enhanced BMO
activity in P. butanovora, differences existed in the maximum rate of cooxidation
that could be supported by different organic acids and in the duration that those
rates of cooxidation could be sustained. Indeed, the choice of butyrate as electron
donor might have compromised interpretation of previously published data from
our own laboratory that indicated 1, 2-trans DCE was not degraded by butane-
grown P. butanovora when butyrate was provided as an alternative electron donor
(Hamamura et al. 1997). In the current study the data clearly showed that 1,2-trans
DCE was degraded with lactate as the electron donor. Although it remains unclear
why lactate is a superior reductant for driving cooxidation in P. butanovora,
several possibilities should be mentioned. First, the rate of butyrate-supported 1,2cis DCE cooxidation decreased more rapidly than did lactate-supported
cooxidation, indicating that enzymes involved in generating reductant from
butyrate might be differentially sensitive to the cooxidation process. In this
context, it is interesting that pyruvate, an intermediate in lactate metabolism and
known antioxidant, was able to offset some of the damage that occurred during
TCE cooxidation in B. cepacia G4 (Yeager et al. 2001). Secondly, initial reactions
in the metabolism of lactate in P. butanovora, and formate in methanotrophic
bacteria directly generate reductant, whereas, initial energy investments are
required in butyrate metabolism before net generation of reductant occurs.
Furthermore, it is possible that butyrate is preferentially metabolized for
biosynthetic purposes or carbon storage and not to oxidative metabolism via a
monooxygenase. Thus, while removal of the natural substrate conceptually
relieves competition between substrate and cosubstrate for the BMO active site,
oxidative cometabolism remains in competition with other aspects of metabolism
for reductant.
Propionate was not able to enhance the cooxidation of ethene in butanegrown P. butanovora, but did enhance cooxidation in propane-grown cells.
Competitive inhibition between the intermediates of butane metabolism (1butanol, butyraldehyde, and butyrate) and three carbon structural analogs suggests
that enzymes involved in butane metabolism have high affinities for 1-propanol,
propanaldehyde, and propionate, (Arp 1999) and it is likely that the oxidation of
propane and butane to propionate and butyrate, respectively, is carried out by the
same enzymes. Although the alkane substrates, butane and propane, appear to be
oxidized to their respective acids by the same pathway, different downstream
pathways are probably involved in the continued metabolism of the organic acids.
Although acetate and formate are known products of propane metabolism in M
vaccae (Coleman and Perry 1885; Vanderberg and Perryl98S), propane-grown P.
butanovora cells could not use formate to support BMO activity and acetate
supported BMO activity was less in propane than butane-grown cells.
Additionally, propionate was not an identified intermediate in propane metabolism
41
in Lvi. vaccae, propionate supported BMO activity was specifically observed in
propane-grown P. butanovora. Further investigation lead to several observations
that were inconsistent with the model of propane metabolism described in M.
vaccae. These results suggest that P. butanovora uses both even and odd chained
13-oxidation pathways in the metabolism of alkanes, and that even and odd chained
13-oxidation pathways are regulated in response to the chain-length of the alkane
used for growth.
Although the substrate range of BMO overlaps with the substrate range of
other non-specific oxygenases (Arp et al. 2001; Sluis et al. 2002), kinetic
parameters of substrate oxidation often differ between enzymes. For example,
acetylene is a potent mechanism-based inhibitor of BMO (Hamamura et al. 1999),
but the related toluene monooxygenase of B. cepacia G4 is unaffected by
acetylene at similar concentrations (Yeager 1999). Cooxidation of 1,1-DCE, 1,2trans DCE, and 1 ,2-cis DCE led to different cytotoxic effects in P. butanovora.
Cytotoxicity associated with the cooxidation of chlorinated ethenes was observed
in several organisms including Burkholderia cepacia G4 (Yeager et al. 2001), a
mixed methanotrophic culture (Dolan and McCarty 1995), and in Methylosinus
trichosporium OB3b (Johan et al. 1997). B. cepacia G4 demonstrated a toxicity
threshold in which cell viability decreased before TCE cooxidation capacity was
reached (Yeager et al. 2001). Underlying metabolic processes responsible for the
allocation of reductant to the monooxygenase remained intact longer than did the
viability of the cells, making the B. cepacia G4 example unique.
1,2-trans DCE and 1 ,2-cis DCE, might represent potential tools for
comparative structure function studies in these enzymes, and to address aspects of
gene regulation. In the case of 1,2-cis DCE, Johan et al. (1997) determined that its
cooxidation by M trichosporium OB3b was more toxic to cell viability than the
degradation of 1,2-trans DCE. The authors suspected that decreased viability and
enzyme activity appeared to be associated with the turnover of 1 ,2-cis DCE
epoxide, while 1,2-trans DCE epoxide was not actively degraded. In contrast, P.
butanovora incurred loss of viability >90% after the cooxidation of only 25
nmoles /mg protein of either 1,2-trans DCE, 1,2-cis DCE, or 1,1 DCE.
Discrepancies between M trichosporium OB3b and P. butanovora could be
attributed to differential activity of the monooxygenase enzymes towards the
epoxides or differential sensitivity of the organisms to oxidative metabolism.
TCE consumption by toluene-grown B. cepacia G4 was not enhanced by
exogenous electron donors. However, the overall respiration of toluene-grown
cells could be enhanced by acetate. During TCE cooxidation B. cepacia G4
retained the ability to allocate reductant from endogenous energy stores to
oxidative cometabolism longer than acetate was able to enhance the overall
respiration of the cells (Yeager et al. 2001). This indicated differential sensitivity
of these metabolic pathways to the cooxidative process. Similarly, lactatedependent oxygen consumption (figure 3.5) and cell viability (figure 3.6)
decreased more rapidly than did the rate of DCE consumption (figure 3.3) or BMO
activity (figure 3.4) in cells treated with either 1 ,2-cis DCE or 1,2-trans DCE.
43
Previous research from our laboratory showed that butane-monooxygenase
can be up-regulated and down regulated in P. butanovora. Sayavedra-Soto et al.
(2001) demonstrated that butane was the most effective inducer of BMO activity.
However, downstream metabolites of butane oxidation, 1-butanol and
butyraldehyde, were also inducers of BMO and it remains unclear if the substrate
or products are the more significant inducers. Xanthobacter Py2 possesses a
related monooxygenase (Zhou et al 1999) enzyme that is induced during growth
on alkenes and during the cooxidation of some chlorinated ethenes (Ensign 1996;
Krum and Ensign 2000). Correlation between the substrate range and the range of
compounds able to act as inducers of the monooxygenase lead to the proposal that
products induced the monooxygenase of Xanthobacter Py2 and not the substrates
themselves (Ensign 1996). In my study, the range of compounds able to induce
monooxygenase activity correlated with the substrate range of BMO. A reporter
strain, BMO insertional knockout, highlighted the trend of product induction in P.
butanovora. Butane, 1 ,2-cis DCE, and ethylene were unable to induce the
reporter. In contrast, the BMO products, 1-butanol and ethylene oxide induced the
reporter strain. 1,2-trans DCE was unique among these compounds in its ability to
induce expression of 3-ga1actosidease from the BMO promoter of the reporter
strain. Although a clear explanation for reporter strain induction by 1,2-trans DCE
cannot be made, others have proposed that separate mechanisms were responsible
for the induction of toluene-monooxygenase in response to toluene and TCE in P.
stutzeri OX1 (Ryoo 2001). Researchers in our lab have speculated that
44
downstream regulatory proteins may also bind upstream of the BMO promoter
(Sayavedra-Soto et al. 2001), and thereby provide an independent mechanism of
induction.
In summary, the findings presented in this thesis have provided some new
insights into the properties of hydrocarbon oxidation in P. butanovora regarding
the molecular regulation of hydrocarbon oxidation genes in P. butanovora, and
into the mechanisms that control the competing requirements for reductant of
monooxygenase reductant and biosynthesis. Furthermore, from an ecological
perspective it raises the possibility that cometabolism might be involved in the fate
of DCEs transported into aerobic zones immediately adjacent to more heavily
contaminated anaerobic areas.
45
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