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CONTRIBUTION OF METHANOTROPHIC GROUNDWATER AND
RHIZOSPHERE BACTERIA TO PHYTOREMEDIATION
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
This dissertation is dedicated to my parents.
I thank Dr. Gabriel Bitton, Dr. Robin L. Brigmon, Dr. Donald L. Rockwood, Dr.
Willian R. Wise, and Dr. Angela S. Lindner for their time and assistance offered as my
advisory committee. I especially thank Dr. Angela S. Lindner, my advisor professor, for
her mentoring, her patience, and, especially, for sharing her enthusiasm towards teaching
and scientific research that will always be a source of inspiration. Also, I would like to
thank Dr. Jud Isebrands and Dr. Madeline Rasche for their absolute support. I also thank
our research group for all their support; Jessica Strate, Chance Lauderdale, and Callie
Whitfield. Finally, I thank my family and friends for their unconditional support, without
which none of this would have been possible.
TABLE OF CONTENTS
I W Le
ACKNOWLEDGMENT S ................. ................. 4..............
LIST OF TABLES ................. ..............8........... .....
LI ST OF FIGURE S ................. ................. 9......... ...
AB STRACT .........._.._.._ ................ 1...._._. 1...
1 INTRODUCTION ................ ................. 13..............
Significance of the Study ................ ................. 16.............
Literature Review............... ................ 17
Chlorinated Compounds ................. .......... ......... ..... .....1
Phytoremediation. ........._.. ..... ._ ._ .............._ 20...
The Rhizosphere ........._.. ..... ._ ._ .............._ 24...
M onoterpenes ................ ................. 29..............
Methanotrophic Bacteria............... .. .. ..............3
Ecology and habitats of methanotrophs. ........._..._.. ...._.._ ........._.... 34
Environmental factors affecting methanotrophs. ........._..._.. ......_..._.......36
Methanotrophs. and chlorinated compounds ........._..._.. ......_._. ..........37
Methanotrophs. and plants ........._..._.._ ...._._. ...._._ ............3
Phylogenetics of methanotrophs. .....__. ............... ........._.._...... 40
Molecular analysis of methanotrophs. ........._.._.... ........._._. ........._.........41
Methods Used to Assess Rhizodegradation Potential in Phytoremediation .......44
Culture-dependent techniques .....___.....__.___ .......____ .............4
Culture-independent techniques .....__.___ ........_._ ........__.........46
Study Hypothesis.............. ............... 50
Study Objectives ........._.. ..... ._ ._ .............._ 51...
Broad Obj ectives ........._.. ..... ._ ._ .............._ 5 1..
Specific Obj ectives ........._.. ..... ._ ._ .............._ 51...
2 M~ETHYLOCYSTIS ALDRICHII SP. NOV., A NOVEL METHANOTROPH
ISOLATED FROM A GROUNDWATER AQUIFER.............. ............ .... 57
Introduction............... ............. 57
M materials and M ethods ................. ................. 58......... ...
Results .............. .. ..............64.......... ......
Di scus si on ................ ..............67. ..............
3 EFFECTS OF ALPHA-PINENE AND TRICHLOROETHYLENE ON
OXIDATION POTENTIALS OF METHANOTROPHIC BACTERIA ............... 78
Introduction............... ............. 78
Materials and Methods ............. ...... .............. 80...
Results and Discussion ............. ...... ..............82...
4 STABLE ISOTOPE PROBING FOR CHARACTERIZATION OF
METHANOTROPHIC BACTERIA INT THE RHIZOSPHERE OF
PHYTOREMEDIATING PLANT S ......... ............... ......... ............8
Introduction............... ............. 88
M material s and M ethod s ................. ................. 90......... ...
Site Description ................ ................. 90..............
Sam pling .................. ........ ......... .......... ............ 9
Stable Isotope Probing (SIP) Soil Microcosms.................... ........... 92
Denaturing Gradient Gel Electrophoresis Analysis (DGGE), Sequencing, and
Phylogenetic Analysis ................ ................. 95..............
Statistics ................ ................. 97..............
Results .............. ... ........... ................. 97....
SIP Protocol Implementation ................ ... ........... ..... .............. 97....
Methanotroph Activity and Composition in the TCE Site ........._.._... ..............98
Methanotroph Activity and Composition in the PCE Site. ........._.._... ............ 101
Discussion ........._.._.. ...._..._ ............... 102..
5 CHARACTERIZATION OF RHIZOSPHERE METHANOTROPHIC
BACTERIA INT TCE PHYTOREMEDIATION: IMPACT OF THE DESIGN...... 111
Introducti on ............. ...... ._ ............._ 111...
Materials and Methods ............. ...... __ ............._ 114..
Site Description ............. ...... __ ............._ 114...
Sampling ............. ...... __ ............._ 116...
Soil Characterization ............. ....._ __ .....__ ... ......17
M icrobial Counts .............. ......_ ............._ 118...
Characterization of Enrichments ...._._._._ ..... ..__.. ....._............. 1
Stable Isotope Probing (SIP) Soil Microcosms.............. .............. 120
Phylogenetic Analysis of Enrichments and SIP Microcosms............._.._.. ....... 121
Statistics ........._.._.. ...._..._ ............... 123..
Results ........._.... ......_.._ ............... 124...
Description of Sites ........._.._.. ...._..._ ............... 124..
Microbial Counts ........._.._.. ...._..._ ............... 125..
Root Biomass ........._.._.. ...._..._ ............... 126..
Enrichments Activity ........._.._.._ ...._.._......_._ ..........12
Phylogenetics of Enrichments ................ ................ 129........ ....
SIP Soil M icrocosms .........._.._.. .........._ __ ............._. 131...
Principal Component Analysis (PCA).............. ................. 134
Discussion .........._.. .. ...._.. ............._. 135...
Microbial Abundance ............... ..... ..._ _....._ .... .. .....13
Activity and Phylogenetics of Enrichments. ...._.._.._ ..... ...... ............ 136
Activity and Phylogenetics of SIP Soil Microcosms ................. ................ 139
Principal Component Analysis (PCA).............. ................. 142
6 CONCLUSIONS ................ ................ 152........ .....
APPENDIX ADDITIONAL TABLES AND FIGURES ................ .................... 157
LIST OF REFERENCES.............. .............. 163
BIOGRAPHICAL SKETCH ................. ................ 185........ ....
LIST OF TABLES
1-1. Physical and chemical properties of TCE and PCE .............. ......_ ............53
1-2. Physical and chemical properties of a-pinene. ......___. .... ..._. .............. 54
1-3. Characteristics of different methanotroph types. ........._.._.. ...._.._ ........._.... 55
2-1. Phenotypic characteristics differentiating Strain CSC1 from M~ethylosinus
trichosporium, M~ethylocystis echinoides, and M\~ethylocystis parvus ................... .. 75
5-1. General characteristics of the different phytoremediation plots at the SRS and
LaSalle sites. ........._._.. ...._._._ ............... 145...
5-2. Summary of phylogenetic assignments (BLAST search) of the pmoA gene
sequences of the active methanotroph populations (13C-DNA fraction) from the
SIP soil microcosms in each phytoremediation plot.............. ................. 150
Al. Soil characteristics of the SRS and LaSalle phytoremediation plots from high-
contaminant regions.* ........... 157.......................
A2. Analysis of variance results (P-values) for the effect of time and depth on
rhizosphere (RH) and rhizoplane (RP) microbial abundance of the LaSalle
phytoremediation plots. ........._.._ ...._._ .....___ ............ 5
LIST OF FIGURES
1-1. Schematic of processes in a phytoremediation system. .........._.__ ........._._ .... 53
1-2. Cl metabolism by methanotrophs and methylotrophs as described by Wackett
(1995). .............. .. ................. 54..............
1-3. Oxidation of TCE by aerobic methanotroph degradation (A) and anaerobic
reductive dehalogenation (B).............. .................. 56
2-1. 16S rRNA phylogeny of Strain CSC1 and related M~ethylosinus and M~ethylocystis
species.. ........._ ......._. ..............73....
2-2. Functional genes phylogenies of Strain CSC1 ................ ................ ......... 74
2-3. Transmission electron microscopy photographs of Strain CSC1 and
M~ethylocystis echinoides.. ................ ......... ......... .... ......7
2-4. Electron microscope cytochemistry of the S-layer of Strain CSC1 ................... ...... 77
3-1. Normalized rate of oxygen uptake by the representative methanotrophs in the
presence of varying concentrations of TCE (*) and (R)-a-pinene. ........._............. 86
3-2. Change in the normalized oxygen uptake rate by representative methanotrophs
observed in the presence of 20 ppm TCE at varying concentrations of (R)-a-
pinene.. ........._.._._ ............... 87..._._. ....
4-1. Equilibrium centrifugation of isotopically labeled DNA in CsCl density gradient
colum ns............... ............... 107
4-2. Initial 13CH4 depletion rates (bars) observed in SIP microcosms after the three
sampling periods at the TCE Site (A) and PCE Site (B).............. .................. 108
4-3. DGGE gels of pmoA PCR products derived from the 13C-DNA fraction of SIP
microcosms at the TCE Site (A-C) and PCE Site (D). ................ ................ ..109
4-4. Neighbor joining phylogenetic tree of pmoA sequences derived from the 13C
DNA fraction of 13CH4 SIP microcosms. ........._.._.. ..........__ ....._.. ... 110
5-1. Location of phytoremediation sites and diagram of sampling areas at the (A)
Savannah River Site (SRS), S.C., and (B) LaSalle, IL. ................ ................. 144
5-2. Microbial counts per tree type from the different phytoremediation plots at SRS
and LaSalle. .......___..........___ .......___..........14
5-3. Oxygen uptake rates of enrichments from different tree types in the presence of
CH 4. ........._.. ......___ ............._ 147...
5-4. Frequency of phylum affiliations per tree type of the NMS with Cu enrichment
components. ................ ................ 148........ .....
5-5. Phylogenetic analysis of the SRS SIP soil microcosms .................... ............. 149
5-6. Principal component analysis (PCA) of culture-dependent and culture-
independent measurements. ................ ................ 151........ .....
Al. DGGE gel of PCR-amplified partial pmoA fragments of different methanotroph
types. ............. .................... 159
A2. Effect of depth on oxygen uptake rates of NMS with Cu enrichments. ................. 160
A3. DGGE gels of 16S rDNA partial sequences from NMS with Cu enrichments....... 161
A4. Phylogenetic tree of 16S rDNA partial sequences from NMS with Cu
enrichments. ................ ................ 162........ .....
Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
CONTRIBUTION OF METHANOTROPHIC GROUNDWATER AND
RHIZOSPHERE BACTERIA TO PHYTOREMEDIATION
Chair: Angela S. Lindner
Major Department: Environmental Engineering Sciences
Trichloroethylene (TCE), a widely used solvent and ubiquitous contaminant, is
effectively removed from soil and groundwater by the use of plants (phytoremediation).
Rapid removal has been reported at the root-zone (rhizosphere), where methanotrophs
(methane-oxidizing bacteria) capable of co-oxidizing TCE are present. The objective of
the study was to determine, by the development of an adequate protocol, how plant type,
system design, and environmental conditions present at two phytoremediation sites
impacts methanotroph's biodegradation potential. To develop characterization methods,
phenotypic and genotypic analyses of an uncharacterized methanotroph, Strain CSC1,
isolated from an uncontaminated groundwater aquifer, were performed. Field sites
represented an engineered system, with poplar and willow trees, and a natural loblolly
pine re-growth area. Laboratory studies were conducted to assess the ability of
methanotrophs to oxidize pine exudates (monoterpenes) and its effects on TCE oxidation.
Field samples were analyzed by culture-dependent microbial counts and enrichments, and
culture-independent stable isotope probing (SIP) microcosms and molecular methods.
Strain CSC1 possessed a unique spiny S-layer and was shown to be a novel strain of the
genus M~ethylocystis and was named M~ethylocystis aldrichii sp. nov. Characterization
methods developed with Strain CSC1 were successfully applied to phytoremediation field
samples and isolates. Different types of methanotrophs were capable of oxidizing
monoterpenes (a-pinene) and, in the presence of TCE; antagonistic and synergistic
responses were observed depending on methanotroph type. Rhizosphere samples
analyzed by culture-dependent methods confirmed the presence of methanotrophs at both
sites; however, enrichments were biased towards type II methanotrophs and did not
correspond with the active populations. Active populations were more diverse and
abundant in the planted samples and strongly influenced by the design, especially the use
of planting material that resulted in a dominance of thermotolerant methanotrophs.
Variable results between the engineered and natural settings highlight the importance of
measuring oxidation potentials and diversity of rhizosphere methanotrophs at any
phytoremediation site, especially if monoterpene-releasing plants are contemplated for
use. Also, study of active populations was shown to be the most accurate
characterization method. Phylogenetic analysis combined with SIP microcosms offers
powerful analytical tools that can ultimately aid practitioners in optimizing
phytoremediation for more effective treatment.
Chlorinated solvents, such as trichloroethylene (TCE), are a major source of
groundwater and soil pollution throughout the United States. It is well known that soil
microorganisms in the presence and absence of oxygen are capable of degrading these
compounds (Barrio-Lage et al., 1986; Fox et al., 1990; Hanson and Hanson, 1996). TCE
can be metabolized to vinyl chloride, a potent carcinogen, if oxygen is not present
(Barrio-Lage et al., 1986; Ensley, 1991). Therefore, this pathway of degradation is
undesirable given conditions where vinyl chloride can accumulate with no further
breakdown. In contrast, methane-oxidizing bacteria (methanotrophs), aerobic
microorganisms known for their bioremediation potential and prevalence in the
environment, can co-metabolize TCE to CO2 at higher rates than other microorganisms
(Wilson and Wilson, 1985; Little et al., 1988; Fox et al., 1990). However, the pathway of
TCE degradation under aerobic conditions is not without risk of forming toxic
intermediates, including chloral hydrate, dichloroacetate, and ethylene glycol (Oldenhuis
et al., 1989; Alvarez-Cohen and McCarty, 1991a; Stacpoole et al., 1998; Lash et al.,
Recently, the possibility of using vegetation to enhance degradation of organic
contaminants in soil systems (phytoremediation) has received attention as an attractive
low-cost alternative to the traditional engineering approaches of soil excavation and
incineration, air stripping, and pump-and-treat (EPA, 2000b; EPA, 2001; McCutcheon
and Schnoor, 2003). When plants are used for this purpose, a series of mechanisms are
involved, including phyto-volatilization, -accumulation, -degradation, -stabilization, and
rhizodegradation. However, the role and contribution of each of these processes to the
overall remediation system has not been accurately characterized (Orchard et al., 2000b;
Shang et al., 2003).
It is well known that the microenvironment surrounding the root-zone of plants
(rhizosphere) is characterized by higher numbers of bacteria and increased microbial
activity (Curl and Truelove, 1986). Therefore, rhizosphere metabolism can significantly
contribute or govern the remedial potential of vegetation. Pilot studies of
phytoremediation systems are being tested in the field in areas contaminated with
chlorinated compounds. The two sites studied in this project are the Savannah River Site
(SRS) in Aiken, South Carolina, and the former LaSalle Electrical Utilities in LaSalle,
Illinois, both involving active phytoremediation of TCE and PCE (Brigmon et al., 2001;
EPA, 2002; Lange, 2004). Research conducted at the SRS has demonstrated that TCE
degradation occurs faster in the rhizosphere of trees (Walton and Anderson, 1990;
Anderson and Walton, 1995; Brigmon et al., 2001). Thus, vegetation may be used to
actively promote microbial restoration of contaminated soils and enhanced hazardous
In these sites several tree species are being studied for their phytoremediation
potential. Loblolly pine (Pinus taeda) is being tested as a promising species that is
capable of up to 90% TCE removal from soils and groundwater at the SRS (Brigmon et
al., 2001). This species is characterized by the production of significant quantities of oil
extracts, composed mainly of monoterpenes (Amaral et al., 1998; Savithiry et al., 1998;
Phillips et al., 1999). Therefore, these compounds may influence the microbial processes
occurring in the rhizosphere either as root exudates or leachate from the decaying foliage
in the surface soil layers. The importance and occurrence of these interactions on
microbial metabolism have been addressed previously in several studies (White, 1986;
Misra et al., 1996; Amaral and Knowles, 1997; Ward et al., 1997). Conflicting findings
on the role of plant exudates on microbial natural processes in the upper soil layers, such
as nitrification and methane consumption, have claimed inhibition as well as stimulation
by these compounds (White, 1986; Misra et al., 1996; Amaral and Knowles, 1997; Ward
et al., 1997). In planted soils at phytoremediation sites, there is no evidence that connects
the increased microbial mineralization of contaminants with the presence of plant
The lack of understanding of the potential roles that rhizosphere bacteria can
assume in the overall removal of contaminants is hindered by the inability to directly
assess the activity and diversity of microorganisms in situ using traditional culture-
dependent methods (Fry, 2004; Smalla, 2004). The recent development of culture-
independent methods that involve soil microcosms and labeled substrates combined with
molecular techniques have enabled scientists to more effectively test in situ conditions
and, more importantly, accurately identify and characterize active microbial populations
(Radajewski et al., 2000).
The main purpose of this study is to determine, by the development of an adequate
protocol, how plant type, system design, and environmental conditions present at two
phytoremediation sites impact the potential ability of methanotrophic bacteria to achieve
biodegradation of chlorinated solvents in contaminated rhizosphere soils and
groundwater. The characterization of this mechanism and the provision of an adequate
technique specific to the rhizosphere can ultimately lead to more efficient
phytoremediation for more effective environmental restoration.
Significance of the Study
Any substance that poses a significant threat to human health and the environment
deserves prioritized attention not only to the study of its toxicology profile but also to its
environmental fate and development of remediation technologies. The U.S. EPA has
classified chlorinated solvents, TCE and PCE, as priority pollutants on the basis of its
widespread contamination in groundwater, its possible carcinogenic nature, and its
potential to be biologically converted to the more potent carcinogen vinyl chloride under
anaerobic conditions. Therefore, the majority of the National Priority List (NPL) sites
are dealing with this type of contamination.
Groundwater and soil contamination by chlorinated solvents presents unique
challenges to remediation technologies. Due to the chemical and physical properties of
these compounds, small amounts of these solvents can contaminate a large volume of
groundwater. The remediation of contaminated soil usually involves excavation and
disposal of the impacted media. However, if the contaminant has reached the
groundwater, the risk to the public, the remedial cost, and the amount of time required to
remove the contaminants can increase substantially (Cheremisinoff, 2001).
Alternatively, in situ remediation technologies, such as bioremediation and
phytoremediation are being tested at several NPL field sites. One example is the SRS
where the potential for phytoremediation of chlorinated solvents has been demonstrated
with loblolly pines capable of up to 90% TCE removal in soil and groundwater (Walton
and Anderson, 1990; Anderson and Walton, 1995). These results are encouraging for the
application of more sustainable remediation technologies that do not require large
amounts of inputs and promote the application of biological systems already in nature to
environmental problems created by humans.
In order to better understand and monitor these biological processes occurring
during phytoremediation, this study combined a laboratory and field approach. The focus
of the study is on the potential contribution of methanotrophic bacteria, capable of co-
oxidizing chlorinated compounds, to the rhizodegradation mechanism in
phytoremediation systems. Additionally, the proj ect concentrates efforts on method
development specific to the rhizosphere environment and compares traditionally used
methodologies with more recent culture-independent methods. Laboratory-based studies
involved characterization of isolated pure and mixed cultures, including a groundwater
isolate, Strain CSC1, which served as a means to develop phenotypic and genotypic
protocols used in the phytoremediation portion of this study. In addition, the role of plant
exudates and their impact on TCE biodegradation was determined with representative
methanotrophs. in the laboratory. Field-based studies involved characterization of the
methanotrophic community in rhizosphere samples from two current TCE- and PCE-
contaminated sites undergoing phytoremediation with different tree types.
The widespread use of chlorinated hydrocarbons as solvents and degreasers in the
metal and dry cleaning industries and their indiscriminate disposal have resulted in a
significant adverse effect on the environment. Trichloroethylene (TCE) and
tetrachloroethylene (PCE), primary chlorinated solvents found at hazardous waste sites,
occupied the 16th and 31s~t position, respectively, in the Priority List of the United States
Comprehensive Environmental Response, Compensation, and Liability Act (CERCLA),
known as Superfund (ATSDR, 2006). The pollutants rank according to their presence at
National Priority List facilities, possible carcinogenic nature, and potential to be
converted to more toxic byproducts, as vinyl chloride that occupies the 4th pOSition in the
CERCLA Priority List (Vogel and McCarty, 1985). A recent report from the National
Academies of Science's Committee on Human Health Risks of Trichloroethylene
concluded that the evidence of TCE' s carcinogenic risk has increased since 2001 (NAS,
2006). Consequently, these compounds are heavily regulated by federal and state
standards. The Safe Drinking Water Act regulates the national maximum contaminant
level (MCL) of TCE and PCE in drinking water at 5 ppb, with a zero maximum
contaminant level goal (MCLG) (EPA, 2000a).
The fate of TCE and PCE released into the environment through a variety of waste
streams will be dictated by their physical and chemical properties (Table 1-1). Because
their densities are greater than 1 g ml l, PCE and TCE are considered dense nonaqueous
phase liquids (DNAPLs). These compounds, considered volatile organic compounds
(VOCs), because of their high values of vapor pressure (74-18.5 mm Hg) and Henry's
law constants (0.011-0.018 atm m3 mOl-1), when released into the atmosphere or surface
water and soil, will volatilize into the atmosphere. In the atmosphere, both compounds
are subj ected to photooxidation with a half-life of a couple of months to days. Also, they
both would be predicted to reach the groundwater given their low partitioning coeffieient
values (log Koc and log Kow of 2-3), high specific gravity (>1), and resulting low
tendencies to adsorb to sediments or soils and to bioconcentrate in animals and plants.
Nevertheless, specific site conditions such as organic soil content can readily contribute
to transient sorption of TCE (Brigmon et al., 1998; Sheremata et al., 2000). PCE may
move slower than TCE in soil infiltration processes, because of its lower water solubility
(150 mg 1- ) compared to TCE (1,366 mg 1- ) (ATSDR, 1999).
As a result of their chemical and physical characteristics, groundwater
contamination by chlorinated compounds presents several challenges for remediation.
When TCE and PCE reach the groundwater, they are anticipated to sink deeper into the
subsurface until they reach a less permeable stratum (confining layer). In this layer they
will spread out or escape through fractures of the rock or clay (Kueper and McWhorter,
1991). Therefore, remediation is more difficult than spills of light NAPLs (LNAPLs),
such as gasoline fuels, that float near the surface of the water table as a compact mass and
do not act as a slow-releasing, continuous source of pollution (Cheremisinoff, 2001).
Chlorinated compounds in the environment are prone to microbial degradation;
however, the rate and extent of oxidation in the presence of oxygen is inversely related to
the chlorine-to-carbon ratio (Hanson and Hanson, 1996). Highly chlorinated
hydrocarbons, such as PCE, are not degraded aerobically. PCE is reductively
dehalogenated under anaerobic conditions (Uchiyama et al., 1989; Bowman et al.,
In aerobic environments, TCE and the metabolites of the reductive dehalogenation
of PCE and TCE, such as dichloroethylene (DCE) and vinyl chloride (VC), are
cometabolically oxidized to CO2 by bacteria that possess oxygenase enzymes. Some of
these enzymes are the methane monooxygenase (MMO) of methanotrophs, toluene
dioxygenase (TDO) of Pseudomona~sputida F l, toluene 2-monooxygenases (TMO) of
Burkholderia cepacia G4, propane monoxygenase of Mycobacterium vaccae JOB5,
phenol hydroxylase (PH) of Alcaligenes eutrophus JMPl34 and Burkholderia cepacia
G4, alkene monooxygenase (AMO) ofAlcaligenes denitrificans spp., ammonia
monooxygenase of Nitrosomona~s europaea, and isopropylbenzene dioxygenase (IPB) of
Pseudomona~s sp. JR1 (Arciero et al., 1989; Wackett et al., 1989; Ewers et al., 1990;
Folsom et al., 1990; Fox et al., 1990; Dabrock et al., 1992; Kim et al., 1996; Smith et al.,
1997). From this cometabolic process, microorganisms do not gain energy or carbon of
the oxidized pollutant. Therefore, an external source of carbon (electron donor) must be
present apart from oxygen that serves as the electron acceptor in the reaction.
In anaerobic conditions, pathways of degradation occur via the process of
dehalorespiration catalyzed by the reductive dehalogenase enzyme. The chlorinated
compound functions as the electron acceptor and, commonly, hydrogen as the electron
donor. The only known microorganism that performs reductive dechlorination of TCE
and PCE to completion is Dehalococcoides ethenogenes)11) strain 195 (Maymo-Gatell et al.,
1999). Other anaerobes and facultative anaerobes, such as sulfate reducers and
methanogens, degrade TCE and PCE incompletely to cis-DCE and VC (Holliger et al.,
1998). Desulfuromona~s chloroethenica, a sulfur-reducing bacterium utilizes pyruvate or
acetate as the electron donor and degrades PCE or TCE to cis-DCE (Krumholz, 1997).
Phytoremediation, the use of plants to remediate contaminated sites, takes
advantage of the ability of plants to extract, sequester or degrade pollutants by the
mechanisms of phyto-extraction, -volatilization, -degradation, -stabilization, and
rhizodegradation (Fig. 1-1). The last mechanism is of special interest because it involves
plant-microbe interactions occurring in the root system (rhizosphere). The role of
rhizosphere microorganisms in the overall breakdown and removal of pollutants is
influenced by the type of contaminant and plant species utilized. Rhizodegradation has
been reported as the main process in organic pollutant remediation of toluene, phenol,
and TCE (Narayanan et al., 1999).
The use of plants represents an alternative technology to traditional waste
management practices, such as incineration, excavation and landfilling, and pump-and-
treat-systems. The effectiveness of phytoremediation has been demonstrated in a wide
range of applications, such as herbicides, petroleum hydrocarbons, metals, radionuclides,
leachates from landfills and sewage, nutrients, pentachlorophenol, polycyclic aromatic
hydrocarbons, and chlorinated solvents. Phytoremediation offers multiple advantages,
including being a low cost in situ technology that is environment-friendly and publicly
accepted. Most importantly there is no need to disturb the site and, after the treatment,
the soil is left fertile for further use. However, some limitations and concerns dictate the
potential applications of this technology, including the time necessary for acceptable
effects to take place, the limited depth of the root system, the sensitivity of plants and
microbes towards the contaminant, the seasonal variability in the rate of treatment, and
the potential of contaminant bioaccumulation or transport into the food chain.
Nevertheless, some of these limitations can be overcome by selecting the appropriate
plant species or by combining other technologies, such as pumping and irrigating the
trees with the deeper contaminated groundwater (EPA, 2000b; McCutcheon and Schnoor,
Phytoremediation efficiency is still limited by a lack of knowledge of many basic
plant processes and interactions with other organisms such as bacteria and fungi.
Pollutant degradation by bacteria and fungi have been studied extensively and, even
though plants can also express similar metabolic pathways, it is only recently that efforts
have been concentrated towards understanding the plant system. Most enzymes involved
in organic xenobiotic degradation, such as cytochrome P450 oxidases, peroxidases, and
glutathione-S-transferase, are known to be present in both microorganisms and plants
(Sandermann, 1994; Shang et al., 2003; Chaudhry et al., 2005).
Phytoremediation of chlorinated solvents from groundwater and soil have reported
up to 90% contaminant removal by the use of different plant species (Walton and
Anderson, 1990; Newman et al., 1999; Brigmon et al., 2001; Nevius et al., 2004).
However, when assessing the responsible mechanisms of contaminant removal, studies
are not consistent. The main contradiction in phytoremediation of chlorinated solvents
regards the magnitude of plant uptake, phytovolatilization, and rhizodegradation
(Orchard et al., 2000a). Several studies have reported that TCE disappearance in planted
systems is mainly due to plant uptake, followed by phytovolatilization and diffusion
through the stem and/or metabolism by the plant (Schroll et al., 1994; Anderson and
Walton, 1995; Newman et al., 1997; Burken and Schnoor, 1998). On the contrary, other
studies have observed TCE degradation occurring mainly as a result of rhizosphere
microbial metabolism (Walton and Anderson, 1990; Anderson et al., 1993; Schnabel et
al., 1997; Orchard et al., 2000a). Contradictory results may be the outcome of
experimental artifacts caused by high exposure to TCE concentrations, use of co-
solvents, the short duration of many studies, and plant stress originated by the use of
static chambers to assess a mass balance of the system. Additionally, problems exist in
the separation of the above- and below-ground compartments, selection of adequate
controls, and lack of methods to correlate bench-scale studies to the field (Orchard et al.,
2000b; Orchard et al., 2000a).
Poplar and willow trees are the preferred plant species in temperate climates for
TCE and PCE phytoremediation. They have also been used for the remediation of heavy
metals, salts, pesticides, explosives, radionuclides, hydrocarbons, and landfill leachates
(Isebrands and Karnosky, 2001). Valuable poplar and willow characteristics that make
them ideal for this application are that they are fast-growing, easily propagated, tolerant
to high levels of contaminants (<550 ppm TCE), resistant to saturated conditions, and
they are phreatophytes (deep-rooted plant where water uptake is mainly from the
groundwater) (Isebrands and Karnosky, 2001; Pilon-Smits, 2005). In particular, willows
have been found to consistently utilize groundwater sources even during periods of
rainfall (Snyder and Williams, 2000). Additionally, poplar and willow trees possess
specialized root vessels (aerenchyma) that may comprise up to 60% of the intracellular
volume and mediate oxygen diffusion deeper into the soil profile (Chaudhry et al., 2005).
It has been hypothesized that willow trees may contain a higher concentration of
oxidative enzymes. When poplar and willow trees were dosed with PCE, only by-
products of degradation were found in willow and no TCE was detected, as it was
commonly found in poplar tissue and its rhizosphere (Nzengung and Jeffers, 2001).
The large surface area and porous wood of poplar trees allows water transport
through the entire cross-section of the stem, which can result in 3 m year- growth under
optimal conditions (Landmeyer, 2001). Transpiration rates can increase from 19 to 200-
1000 L of water day-l in young to mature trees (Newman et al., 1997; Pilon-Smits, 2005).
These high transpiration rates can extract enough water to depress the water table locally,
inducing flow toward the trees and, consequently, containing the contaminant plume
(hydraulic control). Additionally, poplar trees possess endophytic bacteria, including
methanotrophs, that live symbiotically within the plant. Some of these bacteria isolated
from plants are known for their bioremediation potential, including members of
Pseudomona~s sp., Enterobacter-Clostridium species, and methylotroph species such as
M~ethylobacterium populi sp. nov. (Brigmon et al., 1999; Van Aken et al., 2004a; Van
Aken et al., 2004b).
The effectiveness of chlorinated solvent phytoremediation by poplar and willow
trees is strongly influenced by the choice of genotypes (clones). Consideration of the
adequate clone is an essential selection criterion, as the choice must be compatible with
the intended use, the site characteristics (soil type, microclimate, pests and diseases) and
with the local opinion concerning use of native versus exotic trees (Isebrands and
Other potential tree species that have been studied for chlorinated solvent
phytoremediation are conifers, in particular the loblolly pine (Anderson and Walton,
1995; Punshon et al., 2002; Brigmon et al., 2003). In a study where pine, willow, and
poplar trees were compared for their TCE phytoremediation potential, undegraded TCE
was found primarily in the vascular system and leaves of pine, whereas plant metabolites
of TCE were found within the leaf tissue of poplar and willow trees, suggesting plant
degradation potential of these type of trees (Punshon et al., 2002). Meanwhile, for pines,
it has been postulated that rhizodegradation is the main phytoremediation mechanism
(Anderson and Walton, 1995).
The rhizosphere is the root-zone under the influence of the plant (Curl and
Truelove, 1986). This zone is constantly enriched with a variety of plant-derived
compounds, and, as a result, higher microbial densities (5-20 times) and rates of activity
(2-3 orders) occurred in this area compared to non-vegetated soil (Walton et al., 1994).
In rhizosphere studies, the "rhizoplane" is defined as the soil adhered to the roots, the
"rhizosphere" as the soil under the influence of the plant, and the "rhizosphere effect"
(R/S ratio) as the ratio between the abundance of microbial populations in the rhizosphere
to that in bulk soil. However, the effect of the plant is not only translated in higher
abundance but also in higher activity. Therefore, when plants are present, selective
enrichment of populations may or may not translate to higher R/S ratios, although higher
degradation activity is observed (Haby and Crowley, 1996).
Up to 10-40% of the assimilated carbon may be exuded by plants into the
rhizosphere (rhizodeposition) in the form of compounds that are readily utilized by
microorganisms (Whipps and Lynch, 1983). Plant exudates include sugars, amino acids,
organic acids, nucleotides, flavonones, phenolic compounds, terpenes, and certain
enzymes. The rate of exudation depends on the age of the plant, soil nutrient availability,
presence of contaminants, and seasonality. These compounds have been shown to be
released into the rhizosphere in greater amounts at the end of the growing season during
leaf senescence (Hegde and Fletcher, 1996). During this period, about 58% of the
produced fine root biomass dies (root turnover), and, as a result, an increase of up to 2-
fold in phenolic compounds has been observed at the rhizosphere (Leigh et al., 2002).
These compounds are known to stimulate polychlorinated biphenyl (PCB) biodegradation
(Donnelly et al., 1994). Apart from the variety of carbon sources, the rhizosphere
provides steady redox conditions and ideal attachment sites for bacterial proliferation
(Curl and Truelove, 1986; Shim et al., 2000).
Plants benefit from the presence of rhizosphere microorganisms because they can
increase nutrient availability through biosurfactant production (solubilizes soil-bound
nutrients) and N2 Eixation, produce hormones that promote plant growth, suppress
deleterious microorganisms by the production of antibiotics, and degrade phytotoxic soil
contaminants (Smalla et al., 2001). Thus, there is also considerable interest in
characterizing the structure and function of rhizosphere microbial communities for the
advantageous effects to plants.
Phytoremediation may exploit the beneficial effect of moderate plant stress
(Barocsi et al., 2003; Chaudhry et al., 2005). Certain levels of nutrient and water
deficiencies and chemical toxicity may induce stress adaptation, root proliferation and
exudation, and enhance root hair density. For example, P or K deficiency is known to
stimulate exudation of organic acids and certain enzymes. Meanwhile, Fe or Zn
deficiency induces the production of metal chelators (phytosiderophores) (Chaudhry et
al., 2005). Plant tolerance to heavy metals was enhanced when a synthetic chelate
(ethylenediamintetraacetic acid, EDTA), which rapidly increases metal bioavailability,
was applied in several low doses avoiding plant detrimental effects and securing time for
plant adaptation (Blaylock et al., 1997; Barocsi et al., 2003).
Within the diversity of rhizosphere microorganisms, there are strains capable of
degrading xenobiotic compounds (Curl and Truelove, 1986; Walton and Anderson, 1990;
Walton et al., 1994; Anderson and Walton, 1995; Brigmon et al., 1999). The diversity of
heterotroph microorganisms may enhance stepwise transformation of contaminants by
microbial consortium and/or provide an environment that is favorable for genetic
exchange and gene rearrangements of the degradative traits. The presence of structural
analogs to contaminants in root exudates, cell wall components, and lysates, as well as
secondary products of degradation of these materials, might fortuitously select for
microbes that metabolize (accompanied by energy gain) or cometabolize (involving no
energy gain) xenobiotics. Terpenes (secondary plant metabolites) and PCBs plant
analogs (phenolic compounds) have been reported to play an important role in activating
or transforming specific bacterial habitats by inducing biphenyl dioxygenase in PCB-
degrading bacteria and increase populations of this degraders by up to 100-fold (Donnelly
et al., 1994; Fletcher and Hegde, 1995; Haby and Crowley, 1996).
The rhizosphere provides stable sources of oxygen and methane that can support
the activity of methane-oxidizing bacteria (methanotrophs), known to cometabolically
oxidize TCE at higher rates than other bacteria (Little et al., 1988; Fox et al., 1990;
Brigmon et al., 1999). It has been demonstrated that TCE degradation occurs faster in the
rhizosphere of plants (Walton and Anderson, 1990; Anderson and Walton, 1995), where
the presence and density of methanotrophs has been shown to play an important role in
TCE degradation (Brigmon et al., 1999).
Microbial degradation of contaminants is usually not driven by energy needs, but
by a necessity to reduce toxicity for which microbes may experience an energy deficit.
Therefore, the process may be assisted and driven by the abundant energy available in the
rhizosphere environment as root exudates and accumulated plant biomass. The type of
compounds, the species of plant, and the degree of contamination may have the potential
to exert pressure and thus select for specialized degrading bacterial populations.
Consequently, rhizosphere microbial populations may change considerably with time in
response to the type and degree of contamination (Fletcher and Hegde, 1995; Hernandez
et al., 1997; Brigmon et al., 1999; Kozdroj and van Elsas, 2000). However, there is a
lack of information on specific plant characteristics that promote microbial degradation
of organic pollutants (Chaudhry et al., 2005).
Another unexplored area of rhizosphere microenvironments is the interaction of
plant and microbial populations with mycorrhizae. Mycorrhizae, symbiotic root-fungi,
play an important role in plant establishment and survival. Some of these symbiotic
associations are specific to plant species, such as with loblolly pine trees used in
phytoremediation. In these pines higher densities of methanotrophic bacteria were
observed to be associated with the fungi (Brigmon et al., 1999). Therefore, mycorrhizae
may contribute significantly to the remediation potential of several plant species. The
fungi provides the plant and rhizosphere bacteria protection against drought and toxic
pollutants because of the physical barrier created by their extensive hyphae network.
Also, this network can increase the surface area over which the plants and associated
microorganisms explore for water, nutrients, and pollutant uptake. Additionally,
mycorrhizae is known for the extraction of heavy metals and degradation of organic
pollutants from soil, including 2-4-D, atrazine, and PCBs (Donnelly and Fletcher, 1995;
Meharg and Cairney, 2000; Chaudhry et al., 2005).
Rhizosphere-enhanced microbial degradation processes are poorly understood and
certainly vary according to soil conditions, plant species, and type of contaminant (Haby
and Crowley, 1996). It is relevant to study these processes, as they have the potential to
completely mineralize contaminants. As a result, contaminants are not transported into
the plant, reducing the possibility of passing the toxic compound into other organisms in
the food chain and the release of potentially harmful pollutants into the atmosphere. This
scenario may represent the ideal in situ remediation system, where the role of the plant is
to support and stimulate microorganisms capable of contaminant degradation.
More than 70 hydrocarbons, including isoprene, mono- and sesqui-terpenes and a
substantial number of oxygenated organic, are the predominant chemical species emitted
by vegetation (Benjamin et al., 1996). Monoterpenes and isoprene are the major natural
volatile organic compounds (VOCs) and a- and (S-pinene are the representative
monoterpenes (Kim, 2001). Monoterpenes are the simplest constituents of the plant
essential oils and the maj or non-methane hydrocarbon emitted to the atmosphere (4.8 X
1014 -1yar), which contributes to the formation of tropospheric ozone (Zimmerman et
Emissions from conifer forests are predominantly monoterpenes (Amaral and
Knowles, 1998; Savithiry et al., 1998). For Pinus taeda (loblolly pine), monoterpene
emissions are 5.1 Epg g leaf dw-l h-l with greater than 60% represented by a-pinene
(Benjamin et al., 1996; Kim, 2001). On the contrary, broad leaf species, such as Populus
deltoides (poplar) and Salix nigra (willow), are among the high isoprene-emitting species
(Geron et al., 2001) with 37.0 and 25.2 Epg leaf dw-l h- respectively, with no detected
monoterpene emissions (Lamb et al., 1985).
Apart from tree emissions, monoterpenes can be released into the environment
from discharge effluents of the pulp-manufacturing industry, as monoterpenes are the
predominant component of turpentine (Kleinheinz et al., 1999). Monoterpenes are also
being used in the food, perfume, pharmaceutical industries, and, recently, at a larger scale
in an effort to substitute for chlorofluorocarbons and halogenated solvents (Amaral et al.,
1998). Therefore, their environmental fate and interactions with other substances are of
These compounds possess a broad range of functions in nature, from ecological
interactions that extend from allelopathy agents antimicrobialss and fungicides) to
pollinator attractants (Tooker et al., 2002). Potential sources of monoterpenes in soils
include leachate from leaf litter and canopy leaves, root exudation, and deposition from
the atmosphere. The role of monoterpenes and their effect on soil microbial communities
is complex and has not been fully elucidated. It is known that certain microbial enzymes
are stimulated by the presence of monoterpenes and that several microorganisms,
including Pseudomona~s sp., Alcaligenes xylosoxidans,~~~~ddd~~~~ddd and Bacillus sp., can use these
compounds as carbon and energy sources (Vokou et al., 1984; Misra et al., 1996; Vokou
and Liotiri, 1999; Yoo et al., 2001). Also, it has been reported that monoterpenes, first
introduced as decaying plant material or exudates of monoterpene-releasing plants, may
enhance biotransformation of PCBs (Hernandez et al., 1997). However, there are also
reports of inhibition of different microbial processes by these compounds (Vokou et al.,
1984; White, 1986; Ward et al., 1997).
Nitrogen mineralization and nitriaication is inhibited in the presence of
monoterpenes, but the precise mode of action has not yet been elucidated (White, 1986;
White, 1988; White, 1994). White (1988) proposed that monoterpenes hinder
nitrifieation by inhibiting the enzymatic activity of ammonium monooxygenase (AMO),
the first enzyme in the ammonia oxidation pathway, and that the degree of inhibition was
determined by the structure of the compound. These results have provoked other studies
on the effect of monoterpenes on methane oxidation because of the similarity between the
monooxygenase enzymes of these two groups of bacteria (Amaral and Knowles, 1997;
Amaral et al., 1998). The speculation is that monoterpenes inhibit IVMO similarly to
AMO, resulting in the inhibition of CH4 uptake. The authors also support this hypothesis
by mentioning that methanotrophs are not that commonly found in the surface layers of
forest soils, where monoterpene concentrations are the highest (Amaral and Knowles,
1997; Amaral et al., 1998). However, methanotrophs in soil surfaces oxidized
atmospheric CH4 (at concentrations of 1.7 ppm), and their isolation has proven to be
difficult because of the competitive advantage of low affinity methanotrophs in generally
used laboratory conditions at high CH4 COncentrations (usually 20% (v/v) CH4). To date,
high affinity methanotrophs have been phylogenetically identified but not isolated
(Holmes et al., 1999; Jensen et al., 2000). Consequently, the low abundance of
methanotrophs in soil surfaces may be the result of inadequate cultivation techniques.
Alpha-pinene, one of the most abundant monoterpenes, exists predominantly in
North America as the right enantiomer, (+)-a-pinene (Savithiry et al., 1998). This
compound is a bicyclic alkene composed of two isoprene units-CsHs. When released into
the environment, the fate of this monoterpene is dictated by its physical and chemical
properties (Table 1-2). Alpha-pinene released into the atmosphere exists solely as a
vapor that can be degraded by the reaction with photochemically produced hydroxyl
radicals (half-life= 4 h), ozone (half-life= 40 min), and nitrate radicals and in a nighttime
reaction (half-life= 6 min). In soils, a-pinene shows low mobility because it adsorbs to
soil particles (Koc of 1 200 and log Kow of 4.83). However, in moist soil surfaces,
volatilization is expected to be an important process based on its Henry's law constant
(0.107 atm m3 mOl-1). In water, a-pinene will adsorb to suspended solids and sediments
and exhibit a high potential for bioconcentration in aquatic organisms (bioconcentration
factor of 2, 800) (HSDB, 1999). Biodegradation of a-pinene occurs in soils, whereas a
variety of bacteria (Pseudomonas sp., Alcaligenes xylosoxid ans,~~~~ddd~~~~ddd Bacillus sp.) and fungi
(Cladosporium sp.) partially degrade this compound in both aerobic and anaerobic
conditions (Harder and Probian, 1995; Misra et al., 1996; Misra and Pavlostathis, 1997;
Kleinheinz et al., 1999; Pavlostathis and Misra, 1999; Yoo et al., 2001).
Methanotrophs belong to the physiological group of methylotrophs. Methylotrophs
are aerobic microorganisms that utilize as their sole source of carbon and energy reduced
carbon substrates with no C-C bonds (C1 compounds) and assimilate carbon via
formaldehyde (Fig. 1-2) (Hanson and Hanson, 1996). Methanotrophs. are considered
obligate methylotrophs because they only grow on C1 compounds, including methane and
methanol (Lidstrom, 2001). However, recently, a new species was described with the
capability of facultative growth on multi-carbon compounds, M~ethylocella silvestris BL2
(Theisen et al., 2005).
The ability to grow on CH4 is almost exclusive to methanotrophs, except for a
gram-positive methylotroph of the genus M~ycobacterium (Reed and Dugan, 1987).
Methanotrophs. possess complex intracytoplasmic membrane systems, which appear to be
involved in CH4 uptake. The configuration of these membranes apart from other
characteristics separates methanotrophs. into two groups. Those that possess membranes
as bundles of disks stacked throughout the center of the cell (type I) and those with
membranes arranged as rings at the periphery of the cell (type II) (Table 1-3). Other
characteristics that correlate to the type classification include DNA GC content, pathways
of C-assimilation, rosette formation, types of cysts, and ability to fix N2. A small number
of methanotrophs from the genus M~ethylococcus possess characteristics of both groups;
therefore, they have been classified into the type X category (Hanson and Hanson, 1996;
Graham et al., 2002). The existence of this type X grouping is, however, a point of
debate within the Cl research community.
The mechanism by which methanotrophs oxidize CH4 to methanol and
cometabolize (the microorganism gains no carbon or energy from the substrate it
oxidizes, as previously defined) many other compounds including chlorinated solvents is
facilitated by the enzyme methane monooxygenase (1VMVO), unique to methanotrophs.
1V1VO exists in the soluble (slVMVO) or particulate (plV1VO) form depending on the
bioavailability of copper in the environment. plV1VO is a Cu- and Fe-containing enzyme
bound to the intracytoplasmic membrane (Nguyen et al., 1994; Lieberman and
Rosenzweig, 2004), whereas slVMVO, with a unique di-iron site at its catalytic center, is
located in the cytoplasm (Lipscomb, 1994; Kopp and Lippard, 2002). Although both
forms of 1V1VO exhibit a lack of substrate specifieity, the soluble form has been shown to
display a broader range, including alkanes, alkenes, and aromatic compounds. slV1VO is
among the most nonspecific enzymes known to date and exhibit high substrate turnover
rates. Therefore, slVMVO is more suitable for the degradation of a wider variety of
contaminants. However, slVMVO is only synthesized by type II and X methanotrophs in
environments with Cu concentrations less than 50 nM (< 0.89 -1 Epmol Cu gl dw cells)
(Oldenhuis et al., 1989; Hanson and Hanson, 1996).
The oxygen and methane levels also influence the expression of either form of
1V1VO. In environments with abundant oxygen and limiting concentrations of CH4,
methanotrophs express plV1VO, regardless of whether Cu is limiting. On the contrary, in
oxygen-limited environments with high CH4 COncentrations, the expression of either
enzyme is dictated solely by the Cu availability. At low Cu concentrations and high cell
densities, sMMO is expressed. Cells that express pMMO have higher growth yields and
greater affinity for CH4 because pMMO employs an abundant high-energy electron donor
for CH4 Oxidation. Meanwhile, sMMO possesses a high-energy demand, because of the
involvement of NADH+H' as an electron donor that catalyzes this reaction (Hanson and
Hanson, 1996; Sullivan et al., 1998).
Ecology and habitats of methanotrophs
Methanotrophs. have been widely studied for their role in the carbon cycle. They
intercept and oxidize CH4 that escapes from anaerobic environments, thus preventing
large quantities from escaping into the atmosphere (Hanson and Hanson, 1996). Thus,
methanotrophs. are considered the principal biological sink of atmospheric CH4 by
regulating the amount of CH4 preSent in the atmosphere and, consequently, decreasing
the impact that CH4, 23 times more potent than CO2, has on global warming (Houghton
et al., 2001).
Methanotrophs. are widespread in nature, found in any environment where CH4 and
oxygen are present. Under flooded conditions, such as rice paddies, wetland soils,
swamps, and bogs, they are restricted to the soil surface layers and to the rhizosphere of
plants where they intercept the CH4 being produced nearby under anaerobic conditions.
Meanwhile, in upland soils, non-flooded habitats, such as forest, grasslands, and arable
land, methanotrophs. are found in the top soil layers where they oxidize atmospheric CH4
(Hanson and Hanson, 1996; Horz et al., 2002). Also, in these habitats, they are found
deeper in the soil profile, stratified in a narrow band at the oxic-anoxic interface where
concentrations of CH4 and oxygen are the highest. Methanotrophs. have been isolated
from marine, freshwater, and terrestrial habitats, under conditions of high and low pH,
and temperatures up to 550C. They exist as symbionts with invertebrates and plants.
However, little attention has been paid to symbiotic relationships between plants and
methanotrophs, even though the first methanotroph isolated was from the leaves of a
macrophyte in 1906 by Soihngen (Hanson and Hanson, 1996).
It is well known that different types of methanotrophs. adapt better to different
environmental conditions. Methane, oxygen, and nitrogen concentrations are the primary
determinants of the type of methanotroph present in an environment. Type I
methanotrophs. outcompete type II species at low CH4 COncentrations (<2 ppmy in soils),
whereas growth of type II methanotrophs. is favored under low oxygen (<0.2 ppm in deep
waters) and high CH4 COnditions (>1 000 ppmy in sediments) (Hanson and Hanson,
1996). However, because of habitat heterogeneity or differences in experimental
techniques used, a consistent pattern concerning the competitive dominance of certain
types of methanotrophs. has been difficult to discern. Type II methanotrophs. have been
reported to be dominant in soils; however, an abundance of type I or of both types has
also been reported in the soil environment (Vecherskaya et al., 1993; Brusseau et al.,
1994; Sundh et al., 1995; Hanson and Hanson, 1996; Seghers et al., 2005). On the
contrary, type I methanotrophs. appear to prevail in aquatic environments such as lake
water, sediments, and groundwater. Apart from these contradictory results, some genera,
including M~ethylobacter and M~ethylocystis, representatives of type I and II
methanotrophs, respectively, have been detected in a wide range of habitats. It has been
speculated that their ability to produce resistant cysts enables these strains to persist in a
wide range of habitats (Knief et al., 2003; Bodelier et al., 2005).
Environmental factors affecting methanotrophs
The effect of several environmental variables on methanotroph composition,
community structure, and activity has been studied in a variety of habitats with some
inconsistent results. The outcome of these studies seems to depend on the type of habitat
being evaluated and on the variety of methodologies used. Soil type has been reported as
the primary determinant of the methanotroph community structure in agricultural soils
(Girvan et al., 2003; Seghers et al., 2005). However, in forest soils pH value has been
postulated as the primary factor affecting methanotroph distribution (Knief et al., 2003).
Atmospheric CH4 Oxidation activity has been reported to depend on plant cover and land
use, where activity has been shown to decrease with an increase in degree of disturbance
(woodland>grasslands>farmland) (Willison et al., 1995; Knief et al., 2003). Also,
management practices, including fertilizer type (organic versus mineral) and type of tree
in forest stands, have been reported to influence methanotroph activity and abundance
(Reay et al., 2001; Girvan et al., 2003). However, some genera, M~ethylocaldum,
M~ethylosinus, and M~ethylocystis, are universally observed in different soils, independent
of land use or plant cover (Knief et al., 2003).
In the presence of plants, mainly in saturated environments, the spatial distribution
of methanotrophs is determined by the soil compartment (rhizosphere> bulk soil> bare
soil) or the position of the soil-water interface (Gilbert and Frenzel, 1998; Dubey and
Singh, 2001; Macalady et al., 2002). It has been proposed that, because plants differ in
their ability to transport oxygen to the rhizosphere, different factors control their
associated methanotroph populations (King, 1994; Macalady et al., 2002). Spatial
changes in the methanotroph community have also been observed in forest soils
depending on season and soil depth (Henckel et al., 2000; Bodelier et al., 2005). In
winter, atmospheric CH4 Oxidation occurs in a well-defined subsurface layer (6-14 cm
deep), and, during summer, the complete soil core (0-26 cm deep) is active. However, no
seasonal shift in community composition was detected, the same methanotroph
population was identified in summer and winter (Henckel et al., 2000).
Other environmental factors known to affect methanotrophs are the potential
inhibitory effects of ammonium and/or nitrite that act as competitive substrates for MMO
(Dunfield and Knowles, 1995; Hanson and Hanson, 1996). However, recent studies with
rice plants have shown that nitrogen fertilization increases CH4 Oxidation in densely
rooted soils because rhizosphere methanotrophs face intense plant and microbial
competition for nitrogen (Macalady et al., 2002; Eller et al., 2005).
Methanotrophs and chlorinated compounds
Methanotrophs oxidize the less-chlorinated hydrocarbons at very different rates
depending on the form of MMO expressed (Leadbetter and Foster, 1959; Little et al.,
1988; Fox et al., 1990; Alvarez-Cohen and McCarty, 1991a; Alvarez-Cohen and
McCarty, 1991b; Henry and Grbic-Galic, 1991). TCE oxidation by sMMO is
comparable to that of CH4 and up to 700-fold higher than that reported for other MMO
microbial enzymes (toluene 4-monoxygenase, ammonia monooxygenase, and propane
monooxygenase) (Fox et al., 1990). However, TCE oxidation catalyzed by pMMO
occurs at much lower rates than sMMO (DiSpirito et al., 1992).
sMMO oxidizes TCE to TCE epoxide (95%) and chloral hydrate (5%) (Fig. 1-3A)
(Oldenhuis et al., 1989; Newman and Wackett, 1991; Fox et al., 1990). TCE epoxide
rapidly undergoes spontaneous decomposition; meanwhile, chloral hydrate is more stable
and undergoes biological transformation within 1 to 24 h of incubation to trichloroethanol
and trichloroacetic acid. At high temperature (600C) and pH of 9.0, chloral hydrate is
easily decomposed to chloroform and formic acid (Newman and Wackett, 1991). Since
TCE degradation is strictly a cometabolic process, no energy or carbon gain results from
its oxidation; therefore, the presence of a cosubstrate is necessary to maintain cell
biomass and regenerate reductant supply. Although CH4 Oxidation is required for growth
and can provide electrons, it also functions as a competitive inhibitor of TCE
transformation (Henry and Grbic-Galic, 1991). Byproduct toxicity also occurs as a result
of this reaction, with a concomitant decrease in CH4 Oxidation rates, respiratory activity,
and TCE degradation rates (Alvarez-Cohen and McCarty, 1991b; Hanson and Hanson,
1996; Chu and Alvarez-Cohen, 1999). Additionally, TCE metabolites can bind
nonspecifically to cell proteins and inactivate MMO activity (Fox et al., 1990). TCE
epoxide has been postulated as the responsible compound for the observed toxicity due to
its reactivity or that of its degradation products (Fox et al., 1990; Chang and Alvarez-
Cohen, 1996; Vlieg et al., 1996; Sullivan et al., 1998). Intermediate toxicity can be
reduced by the addition of an external supply of reducing equivalent such as format
(Alvarez-Cohen and McCarty, 1991b). However, TCE oxidation toxicity appears to have
a selective effect over different species of methanotrophs based on observations of
distinct rates of recovery (Henry and Grbic-Galic, 1991).
Under anaerobic conditions chlorinated compounds readily undergo reductive
dechlorination (Fig. 1-3B). PCE and TCE are degraded to dichloroethene isomers (cis-
and trans-1,2-DCE),tr~r~r~r~r~r~ 1,1-DCE, vinyl chloride, ethene, and ethane. DCE isomers and
vinyl chloride in the presence of TCE and no oxygen often persist in the environment
because their dechloronation yields less energy than that of their parent compound (Fox
et al., 1990; Hanson and Hanson, 1996). Considerable concern exists over the biological
production of vinyl chloride, a known human carcinogen; however, this product is readily
oxidized by sMMO in aerobic environments (Fox et al., 1990). Additionally, when
consortia of bacteria (methanotrophs and heterotrophs) are present, further oxidization of
chloral hydrate, chlorinated acetic acids, and vinyl chloride has been observed along with
a provision of additional reducing power for the process (Alvarez-Cohen et al., 1992;
Uchiyama et al., 1992; Chang and Alvarez-Cohen, 1996).
Methanotrophs and plants
Plant-methanotroph associations studied to date have considered mainly rice Hields
and wetlands because of their importance as maj or areas of CH4 prOduction. DeBont et
al. (1978) was the first to report CH4 Oxidation associated with rice roots, he noticed that
most of the CH4 diffused through the rhizosphere was oxidized. This observation was of
relevance because any small change in oxidation processes occurring at the rhizosphere
could have a global impact because rice Hields contribute to approximately 25% of the
current CH4 flux to the atmosphere. However, studies to date on these interactions show
high unexplained variability within plant species and between environments. It has been
observed that plant species, known to oxidize CH4 in their rhizospheres, when planted in
a different environment, CH4 COnSumption ranged from detected to no oxidation (King,
Root surfaces and their interior, zones where CH4 is transported from the
methanogenic sediments to the atmosphere and where atmospheric oxygen is transported
to the sediments, both support methanotroph populations in saturated environments
(King, 1996; Gilbert et al., 1998; Eller et al., 2005). However, methanotrophs and
methylotrophs have also been detected in these locations in poplar and pine trees in non-
saturated environments (Brigmon et al., 1999; Pilon-Smits, 2005). Methylotrophs
permanently associated with the plant are often encountered in the phyllosphere (leaf
surface) and rhizosphere. Plants can also benefit from these associations. For example,
methanotrophs can excrete or expel by cell lysis phytohormones (cytokinins and auxins)
and other bioactive compounds. Additionally, type II and X methanotrophs can Eix
nitrogen and, therefore, can be considered phytosymbionts on the surface and inside plant
tissues (Doronina et al., 2004).
The pattern of methanotroph root colonization has been studied in rice plants
(Gilbert et al., 1998; Gilbert and Frenzel, 1998). The colonization is spatially very
heterogeneous; some roots are not colonized at all, while others possess microcolonies as
clumps or thick bacterial layers. As known for other types of bacteria, methanotroph root
colonization followed the pattern of cell wall formation, potentially due to the exudation
of organic substrates and oxygen leakage at these sites. While, methanotrophs cannot
utilize complex organic substrates for growth, they do utilize some amino acids as
nitrogen sources (Gilbert and Frenzel, 1998).
Phylogenetics of methanotrophs
Methylotrophs are scattered among the Proteobacteria within the a-, (3-, and y-
subdivisions, not forming an evolutionary coherent group. Multi-gene operons appear to
be rare among its members, and, on the contrary, plasmids are common. However, no
functions have been ascribed to these plasmids, and they are entirely cryptic.
Methanotrophs cluster into the a- and y-Proteobacteria and are considered ideal
microorganisms for molecular biology studies. Methanotroph phylogeny and their
phenotypic and eco-physiology characterization into types I, II, and X validate each other
(Lidstrom, 2001). The type classification, initially proposed by Whittenbury et al.
(1970), has been supported by analysis of 5S and 16S rRNA genes. Recently, with the
incorporation of molecular methods to methanotroph studies, novel strains are being
recognized that do not grow in standard laboratory conditions (enriched solid and liquid
media with high concentrations of CH4). For example, the upland methanotroph soil
clusters (USC-a and USC-y) that oxidize atmospheric CH4 in forest soils. Another
example is the genus M~ethylocella, sensitive to salts in regular cultivation media (Holmes
et al., 1999; Henckel et al., 2000; Jensen et al., 2000; Bourne et al., 2001; Knief et al.,
2003; Theisen et al., 2005).
Molecular analysis of methanotrophs
Methanotroph phylogenetic studies have been conducted with both phylogenetic
and functional gene markers. Functional gene markers detect the active-site subunit of
both MMO forms, pnzoA for pMMO and namoX for sMMO, and of methanol
dehydrogenase by the nzxa~F gene. The use of these markers enables assessment of the
potential functional diversity of methylotrophs and methanotrophs within an
environment. The universal phylogenetic 16S rDNA (rRNA) primer set amplifies the
variable V3 region of the gene, extensively studied to enable inference of phylogenetic
relationships among microorganisms. Phylogenetic analysis of methanotrophs usually
considers both pnzoA and 16S rDNA analysis due to the fact that most methanotrophs
express the pMMO gene and phylogenies between these two primer sets are closely
related to each other (Bowman, 2000).
Interpretation of pnoA phylogenetic analysis must take into consideration that
multiple copies of the gene can exist in one organism; therefore, novel clusters of pnoA
sequences do not necessarily indicate that novel groups of uncultivated methanotrophs
exist. Copies of the pnoA gene (pnzoA2) can show less than 80% identity to the
previously known pnoA gene (pnoA1). Also, in some cases, there is no correlation
between the 16S rDNA and pnoA phylogenies, and, for the genus M~ethylobacter, some
strains poorly amplify the pnzoA gene with the standard primer sets, underestimating the
methanotroph diversity. Finally, one must keep in mind that the genus M~ethylocella, the
only known exception to the universality among methanotrophs of the pnoA gene, must
be detected using a different primer set. Other primer sets that could be used are the
namoX or nadh that amplify the sMMO and methanol dehydrogenase active site,
respectively (Dedysh et al., 2000).
It is of importance to recognize that the success in gene retrieval from
environmental samples depends on the quality of the primer sets used. Different levels of
methanotroph diversity have been reported with different primers sets (Bourne et al.,
2001). For example, Hutchens et al. (2004) reported that, by using the pnoA primer set
Al89f/A682r, only 8 operational taxonomic units (OTUs) were detected, but, with the
Al89f/mb661r primer set, 12 OTUs were retrieved (Hutchens et al., 2004). The pnoA
primer set Al89f/mb661r detects almost all methanotrophic bacteria, except sequences of
M~ethylonona~s, Methylocaldunt, and the reported forest clone clusters, but it does exclude
all known a~noA sequences of ammonia-oxidizing bacteria, except for Nitrosococcus
(Kolb et al., 2003).
With the incorporation of molecular techniques to the study of microbial ecology,
one of the most intriguing questions is the relationship between what has been reported
previously by community assessment based on traditional culturing and what has recently
been described by culture-independent techniques. Several methanotroph studies that
implemented denaturing gradient gel electrophoresis (DGGE) point out misleading
results of previous traditional culture-dependent methods. For example, in grassland
soils the culture-dependent most probable number (MPN) technique was compared to
direct soil sample DGGE analysis (Horz et al., 2002). While MPN analysis detected only
one methanotroph strain, DGGE revealed a more diverse and dynamic methanotroph
community. In a similar matter, enrichments characterized by DGGE were compared to
results of morphological observations and strain isolation from agricultural soil (Jensen et
al., 1998). The DGGE profile of the enrichments showed higher diversity (13-14 bands)
than the morphological observation and isolation, where only 2 to 4 dominant
morphological types were detected and only one colony was isolated (Jensen et al.,
Another methodology that is revolutionizing methanotroph studies by linking
microbial identity to biological function under conditions approaching those in situ is
stable isotope probing (SIP) (Radajewski et al., 2000). A labeled substrate, a less
naturally frequent isotope, is incorporated into the active microbial biomass. In the case
of methanotrophs, 13CH4 has been used to label the DNA of active organisms during
DNA synthesis and replication. The heavier DNA (13C-DNA) can then be separated from
the naturally occurring 12C-DNA. The methodology has been used to study
methanotroph communities of peat soils (Morris et al., 2002), acidic forest soils
(Radajewski et al., 2002), cave water (Hutchens et al., 2004), and soda lake sediments
(Lin et al., 2004).
Results of this SIP method confirmed that most methanotroph communities in the
environment are active and constitute a small fraction of the entire population responsible
for CH4 Oxidation. Soil community fractions revealed that only a small percentage or
possibly no methanotrophs were present in the 12C-DNA fraction, while the 13C-DNA
fraction was composed of 32% or 96% methanotroph, in peat and forest soils,
respectively (Morris et al, 2002; Radajewski et al., 2002). Interestingly, sequences have
been found that may represent novel methanotrophs and methylotrophs, suggesting that
these bacteria most probably assimilated methanol (13CH30H) excreted by
methanotrophs during 13CH4 Oxidation. However, in some cases, the affiliation to
methanotrophs of the retrieved sequences ((p-Proteobacteria) can not be explained,
suggesting the possibility that bacteria not previously considered to be involved in CH4
oxidation may derive a significant proportion of their carbon from products of
methanotroph metabolism or possibly even from CH4 itself.
When the functional pmoA gene has been examined before and after SIP
experiments, the diversity was lower in the 13C-DNA fraction indicating that not all
methanotrophs in an environment are active (Morris et al., 2002; Radajewski et al., 2002;
Lin et al., 2004). Overall these studies using SIP methods have revealed that the active
methanotroph community in peat and acidic forest soils was dominated by type II
methanotrophs (Morris et al., 2002; Radajewski et al., 2002) and in soda lake sediments
by type I methanotrophs (Lin et al., 2004). These results give insight into the ecological
niches occupied by each methanotroph type.
Methods Used to Assess Rhizodegradation Potential in Phytoremediation
The microbial composition of the rhizosphere is known to differ both qualitatively
and quantitatively from that in a non-planted soil. However, a precise determination of
the microbial diversity in soil or the rhizosphere compartment remains to be established,
as only up to 10% of soil microbial species can currently be cultured in the laboratory
(Fry, 2004). Although the ability to culture the yet-uncultured bacteria is of importance,
a number of indirect methods are currently used to establish the biodegradation potential
of soil microorganisms. By the use of these indirect methods, not only is the microbial
composition and structure of a specific habitat being determined but also it is possible to
link function to activity by the use of labeled substrates.
Microbial counts. Historically, colonies forming units (CFU) and most probable
number (MPN) technique have been used to enumerate selected microorganisms and
assess the microbial composition of a site. However, it is recognized that only a small
portion of bacteria can form colonies when traditional plating techniques are used. The
proportion appears to be determined by the oligotrophic extent of the evaluated
environment, where the more oligotrophic the environment, the higher portion of bacteria
that do not grow under standard cultivation conditions (Smalla, 2004). Culturability,
defined as the percentage of culturable bacteria to total cell counts (microscopically
assess), has been determined to be around 0.3% in soils (Amann et al., 1995). Further
limitations represent organisms that, under environmental stress, enter the viable but
nonculturable state and bacteria strongly attached to soil particles that cannot be
dislodged (Smalla, 2004). While microbial counts are widely used and easy to prepare,
they are time-consuming and require multiple replicates and cultivation periods of weeks
or months. Additionally, they do not discern relationships among bacteria, are highly
selective, and inaccurate, underestimating the abundance of the microbial populations
Enrichments. Another commonly used approach for microbial characterization of
environmental samples is to obtain enrichments of selected groups of bacteria. Usually,
the highest dilution of the MPN technique is used as an inoculum for further cultivation.
This procedure avoids the selection of only the fast-growing and less-numerous bacteria,
which benefit from the fact that some abundant bacteria do not grow directly on
conventional media (Fry, 2004). Enrichments offer the possibility of preserving
syntrophic relationships among bacteria and obtaining environmental isolates from which
physiological and phylogenetic characterization can be performed (Wise et al., 1999).
Additionally, cultures can be used to assess potential microbial activity, optimum
conditions for degradation, and microbial diversity of a particular sample. However,
conditions for enrichment do not resemble the Hield, and they are highly selective, which
makes extrapolation of results to the Hield difficult. Furthermore, obtaining a stable
culture can take months, and, when studying the culture's phylogenetics, there is no real
indication of gene expression in situ, which is ultimately what determines environmental
impact at the Hield. Because this technique relies on the culturability of the members of a
particular sample, it is a common finding that isolates represent only a few of the most
abundant bacteria. However, in some environments, isolates can represent higher
numbers. In seawater, isolates represented 7-69% of the total bacterial clones obtained
from culture-independent methods (Fry, 2004).
Molecular methods. Since 1990, microbial ecologists have been studying
bacterial diversity by isolating community DNA, amplifying their 16S rRNA genes,
cloning the fragments, and sequencing the clones. The development of molecular
methods over the past two decades has helped resolve difficulties inherent in studying
diversity using traditional approaches that are based on observations of physiology and
morphology. It has led to an increase in the numbers of identified bacteria divisions to
greater than 40, in which only 23 divisions are represented by isolates (Smalla, 2004).
Therefore, most of the bacteria in culture collections that grow on conventional media are
not the most abundant in natural habitats. For this reason, there is a necessity to isolate
the ecologically relevant bacteria, the as-yet-uncultured bacteria, and study their
physiology. The basic problem is that many numerically abundant bacteria grow more
slowly than the less-dominant bacteria on most laboratory media (Fry, 2004).
In spite of the advances, some challenges still remain for the molecular microbial
ecologist. The most pressing challenges are obtaining nucleic acids suitable for
molecular analysis and access to sufficiently large, high quality databases (Smalla, 2004).
Extraction problems when organic are high in an environment, such as in the
rhizosphere, still constitute a maj or draw back of this technique. Also, the effectiveness
of oligonucleotide probes to detect organisms may be uncertain because of the possibility
of encountering new genes or genes that are not conserved in a similar matter within
related groups of bacteria, and, as a result, genes obtained from cultured organisms may
not be sufficiently similar to genes in the environment (Hanson and Hanson, 1996).
Also, there is a lack of rDNA sequences of many described species. Another limitation is
that some molecular applications do not allow conclusions about the metabolically active
populations or on gene expression because they do not distinguish between active and
non-active organisms, thus limiting the use of these methods. Nevertheless, this
information might be obtained from RNA analysis or by the use of labeled substrates.
DGGE analysis. The technique is based on the separation of PCR fragments of the
same length in polyacrylamide gels containing a linearly increasing gradient of chemical
denaturants (urea and formamide) (Muyzer et al., 1993; Muyzer et al., 2004). Separation
is based on the electrophoretic mobility of the partially melted DNA molecule, which is
lower compared to that of the completely helical form of the molecule. The different
fragments melt in discrete melting domains (stretches of base pairs with an identical
melting temperature). Once the domain with the lowest melting temperature reaches its
denaturing concentration at a particular position in the gel, a transition from helical to
partially melted molecule occurs, and the molecule will stop migrating. Therefore,
sequence variants (different in base pairs) will stop migrating at different positions from
which DNA fragments are differentiated and excised for sequence analysis (Muyzer et
al., 1993). A GC-rich sequence (GC clamp) is incorporated into one of the primers to
modify its melting behavior to the extent to which close to 100% of all possible sequence
variations can be detected. The resulting banding pattern represents a profile of the
populations in the sample, and the relative intensity of each band and position represents
the relative abundance of a particular member of the community (Muyzer et al., 1993).
The main advantage of DGGE is that it permits high-resolution phylogenetic
analysis of a complete community by its diversity pattern in a qualitative and semi-
quantitative matter. Large numbers of samples can be quickly analyzed and compared,
permitting temporal and spatial analysis within and between communities. The only
comparable technique at the moment is terminal restriction fragment length
polymorphism (T-RFLP) (Bodelier et al., 2005). However, interpretation of T-RFLP
data requires constructing a clone library that can be time-consuming due to the cloning
step, and less abundant species are not always detected. On the contrary, DGGE can
detect species represented by as low as 1% of the population, and bands are directly
excised from the gel, reamplified, and sequenced without the need of cloning (Muyzer et
al., 1993). DGGE cagn detect up to 95% of all possible single base substitutions among
sequences of up to 1, 000 bp in length, and it can be adjusted to narrowed denaturant
gradients to provide higher resolution. Also, DGGE profies can be transferred to
hybridization membranes and probed with specific oligonucleotides (Vallaeys et al.,
However, the main constraint of DGGE is the amount of phylogenetic information
in the length of the commonly amplified fragments (< 500 bp). These partial sequences
may not be sufficient to discriminate among strains (Boon et al., 2002; Bodelier et al.,
2005). Another limitation is the production of multiple bands by one organism because
of multiple heterogeneous operons or copies of the target gene, or due to the use of
degenerate primers. Also, if the target sequences in a sample are present at dissimilar
concentrations, the less abundant sequences may not amplified sufficiently to be
visualized as bands, underestimating the diversity of the sample (Boon et al., 2002).
Another problem of DGGE is co-migration of bands and bands at identical positions that
are not necessarily derived from the same species; however, these bands can be screened
by reducing the denaturant gradient, and, when necessary, bands in similar positions may
require multiple sequencing. Finally, in community analysis of highly related
phylogenetic clusters, bands can represent heteroduplexes, PCR artifacts from mixed
DNA templates that result from two similar, but not corresponding strands, annealing
together. These artifacts can be detected because they produce bands at low denaturant
concentrations (Wise et al., 1999).
Stable isotope probing microcosms (SIP). SIP microcosms permit the
identification of organisms responsible for certain in situ transformation processes by the
use of a labeled substrate (a less naturally frequent isotope) that will be incorporated into
the active microbial biomass (Radajewski et al., 2000; McDonald et al., 2005).
Subsequently, the labeled DNA fraction is separated from the naturally occurring fraction
by CsCl density gradient centrifugation. The labeled fraction is then analyzed to identify
the active microbial community by cloning, followed by restriction fragment length
polymorphism (T-RFLP) or by DGGE analysis.
The maj or limitation of the SIP methodology is the dilution of the labeled substrate
before its assimilation and incorporation into the active organisms, which can happen if
simultaneous growth on an unlabeled substrate is occurring in the microcosms. Other
constraints may be the relative long incubation periods needed to label sufficient biomass
and, consequently, the potential for the use of labeled metabolites by non-target
organisms (cross-feeding). Also, the artificial spike and relatively high concentrations
used of the labeled substrate, may stimulate microorganisms that were not active in situ,
and, as a result, the analysis may represent the potential active population, rather than the
active microbial community at the time of sampling (Lin et al., 2004; McDonald et al.,
2005). Therefore, SIP studies should provide a rational basis for the application of
molecular biological techniques to study the role of specific organisms that are likely to
be involved in a defined process (Radajewski et al., 2003). Finally, the technique is
expensive and requires certain expertise, but it is one of the most powerful molecular
techniques available, providing information on the active microbial populations of an
environmental sample and linking function to identity.
The diversity and activity of methanotroph populations associated with the
rhizosphere of plants used in phytoremediation processes are impacted by plant type,
system design, and environmental conditions present at the site.
*Provide full characterization of a well-known, pure methanotroph, Strain CSC1,
isolated from a groundwater aquifer known to oxidize TCE as a means of method
development of phytoremediation studies.
*Assess the effects of plant exudates, specifically monoterpenes, on TCE
cometabolism by methanotroph bacteria.
*Develop a protocol using stable isotope probing (SIP) methods specific to
*Assess differences in methanotroph abundance, activity, and diversity observed in
rhizosphere samples from several plant species used in phytoremediation.
*Determine the effectiveness of culture-dependent and culture-independent methods
to characterize potential microbial degraders.
*Ultimately provide guidance for the phytoremediation practitioner to more
accurately predict the extent of TCE rhizodegradation when using monoterpene-
and non-monoterpene releasing plants.
*Characterize the methanotroph Strain CSC1 using phenotypic and physiological
descriptions, phylogenetics of the 16S rDNA and multiple functional genes (mmoX,
pmoA, mxaF), and DNA-DNA hybridization.
*Determine the effect of (R)-a-pinene on TCE cometabolism by pure cultures of
representative type I, II, and X methanotrophs using oxygen uptake analysis.
*Combine and implement SIP methods with molecular fingerprints techniques, as
denaturing gradient gel electrophoresis (DGGE), using the 16S rDNA and
functional pmoA genes, to develop a precise methodology for methanotroph
*Determine by culture-dependent microbial counts the abundance of the heterotroph
and methanotroph communities from rhizosphere soil compartments of two
*Enrich for and characterize methanotroph mixed cultures from rhizosphere soil
compartments of two phytoremediation sites by their oxidation potential using
oxygen uptake analysis, presence and activity of soluble methane monoxygenase,
and phylogenetics of 16S rDNA-DGGE analysis.
*Determine the effect of several environmental variables (location, time, tree type,
contaminant type and concentration, depth, and system design) on the
methanotroph populations of different rhizosphere soil compartments at two current
Table 1-1. Physical and chemical properties of TCE and PCE.1
Molecular formula C2HCl3 2 C4
Molecular weight (g mol l) 131.4 165.8
Specific gravity (at 20oC) 1.465 1.623
Vapor pressure (at 25oC) (mm Hg) 74 18.5
Water solubility (at 25oC) (mg I 1) 1 366 150
log Koc 2.03-2.66 2.20-2.70
log Kow 2.42 3.40
Henry's law constant (at 25oC) (atm m3 mOl-1) 0.011 0.018
(removal of contaminants
and release to the
(plant metabolism of
(extraction of contaminants,
can result in plant
Phytostabilization of contaminants in the
(soil/vegetation binding or rhizosphere)
containment of contaminant plume
by plant water uptake)
Figure 1-1. Schematic of processes in a phytoremediation system (McCutcheon and
Table 1-2. Physical and chemical properties of a-pinene.l
Molecular formula CloH16
Molecular weight (g mol l) 136.24
Specific gravity (at 20oC) 0.8592
Vapor pressure (at 25oC) (mm Hg) 4.75
Water solubility (at 25oC) (mg 1- ) 2 2.5
Koc 1 200
Log Kow 4.83
Henry's law constant (at 25oC) (atm m3 mOl-1) 0.107
(1HSBD, 1999; 2Li et al., 1998)
Monooxygenas e Formaldehyde
(MMO) Dehydrogenase Formate
H20 Methanol Dehydrogenase
02 Dehydrogenase H20 rHOHAH++0
CH4 /C3H HH CO O
H +NADH NAD ""5 HH
2H+ 2H+ A
Figure 1-2. Cl metabolism by methanotrophs and methylotrophs as described by
Table 1-3. Characteristics of different methanotroph types .
Characteri sti c Type I Type II Type X
Recognized genera M~ethylomicrobium M~ethylococcus
Short rods, some Rods, crescent- or
Cellular shape Cocci
cocci or ellipsoids pear-shaped
Azotobacter-type Exospores or lipid Azotobacter-type
cysts cysts cysts
Paired, parallel to Disc-shaped
membranes bundles of vesicles .h yolsatc bnlso
pathway assimilation RuMP2 Serine u (ao)
TCA cycle Competencomlet
(one exception)Cmlt Icmlt
DNA G+C content 50-54% 62.5% 62.5%
phospholipid fatty 16 C-atoms 18 C-atoms 16 C-atoms
-Methane pMMO2 pMMO/sMMO2 pMMO/sMMO
-3-hexulose phosphate + +
-Hydroxypyruvate + +
-Nitrogenase + +
-Isocitrate NAD /NADP' NADP' NAD
Growth temperature 40oC> 40oC> >45oC
Phylogeny 6 Proteobacteria at Proteobacteria 6 Proteobacteria
'(Hanson and Hanson, 1996: Sullivan et al., 1998: Graham et al., 2002).
2RuMP= ribulose monophosphate cycle: TCA= ricarboxylic acid cycle: pMMO= particulate
methane monooxygenase: sMMO= soluble methane monooxygenase: NAD = nicotinamide
adenine dinucleotide: NADP = nicotinamide adeni le dinucleotide phosphate.
A. C~HPc H
Figure 1-3. Oxidation of TCE by aerobic methanotroph degradation (A) and anaerobic
reductive dehalogenation (B) (4Barrio-Lage et al., 1986; 3Fox et al., 1990;
1Vlieg et al., 1996; 2Lontoh et al., 2000).
CI CI O
tr chloroacetate tric loroethanH
HOCH + CO2
M~ETHYLOCYSTIS ALDRICHII SP. NOV., A NOVEL IVETHANOTROPH ISOLATED
FROM A GROUNDWATER AQUIFER
Note: Manuscript submitted to the International Journal of Systematic and Evolutionary
Lindner, A.S., Pacheco, A., Aldrich, H.C., Costello Staniec, A., Uz, I. and Hodson, D.J.
2006. M~ethylocystis aldrichii sp. nov., a novel methanotroph isolated from a
groundwater aquifer. International Journal of Systematic and Evolutionary
Microbiology XX: XXX-XXX.
Species of the genus M~ethylocystis are strictly aerobic, gram-negative bacteria that
are able to grow on one-carbon compounds (e.g., methane or methanol) (Bowman et al.,
1993a). The genus M~ethylocystis belongs to the alpha-subclass of the Proteobacteria and
currently consists of 2 species with standing in nomenclature, M\~ethylocystis parvus and
M~ethylocystis echinoides (Whittenbury et al., 1970; Gal'chenko et al., 1977; Bowman et
al., 1993a). Numerous M~ethylocystis strains have been identified in a variety of
environments, including lake, ocean, marsh, and creek sediments and water, coal mine
drainage water, the roots of plants, etc. (Whittenbury et al., 1970; Gal'chenko et al.,
1977; Bowman et al., 1993a; Hanson and Hanson, 1996; Calhoun and King, 1998; Heyer
et al., 2002).
Species of the genus M~ethylocystis are Type II methanotrophs, classified, in part,
by their possession of paired membranes aligned with the cell periphery, the serine
pathway, and predominant fatty acids with 18 carbons (Hanson and Hanson, 1996;
Graham et al., 2002). All known Type II methanotrophs, including the M~ethylocystis
species, express the particulate form of methane monooxygenase (plV1VO), and, with the
exception of 2ethylocystis parvus, all express the soluble form of methane
monooxygenase (sMMO) under conditions of low copper concentrations (Stanley et al.,
1983; Prior and Dalton, 1985; Choi et al., 2003). M~ethylocystis parvus does not possess
genes encoding for sMMO (Tsien and Hanson, 1992; McDonald et al., 1997; Lloyd et al.,
1999) and is, therefore, incapable of oxidizing aromatic compounds. All M~ethylocystis
species produce oxidase and catalase, are nonmotile and are capable of fixing
atmospheric nitrogen (Hanson and Hanson, 1996).
The focus of this paper is Strain CSC1, a group II methanotroph previously isolated
from an uncontaminated groundwater aquifer at Moffet Naval Air Station in Mountain
View, CA, USA (Henry and Grbid-Galid, 1990). This methanotroph expresses sMMO
under copper-limiting conditions and is capable of oxidizing aliphatic and aromatic
compounds (Henry and Grbid-Galid, 1991; Adriaens and Grbid-Galid, 1994; Adriaens,
1994; Hriak, 1996; Hriak and Begonja, 1998). Despite its being the focus of these
numerous studies aimed primarily towards contaminant degradation potential, Strain
CSC1 has not been characterized and differentiated from other known Type II
methanotrophs. This study provides phenotypic and genotypic analysis of this
groundwater isolate. The formal taxonomic description of this novel M~ethylocystis
bacterium, M~ethylocystis aldrichii sp. nov. strain CSC1, is reported. Differences in
various characteristics of Strain CSC1 compared to other known methanotrophs are
described, and its unique surface features broaden the observed physiological traits of
Materials and Methods
Strain CSC1 was obtained from Dr. Dubravka Hriak at the Rudj er Boskovic
Institute in Zagreb, Croatia, and M~ethylosinus trichosporium was obtained from Dr.
Jeremy Semrau in the Department of Civil and Environmental Engineering at the
University of Michigan, Ann Arbor, USA. M\~ethylocystis parvus and M~ethylocystis
echinoides were obtained from NCIMB (Aberdeen, England). The basal medium used
for growth when culturing for sMMO expression was nitrate mineral salts (NMS)
medium with no added copper, as described previously (Whittenbury et al., 1970; Lontoh
and Semrau, 1998). Ten Cpmol 1-1 copper nitrate (Cu(NO3)2) WAS added to the NMS
medium to provide conditions for pMMO expression. Liquid cultures were routinely
grown at 250 rpm and 300C in either 50- or 500-ml batches in 250-ml Erlenmeyer or
2800-ml Fernbach flasks, respectively. The flasks were fitted with rubber stoppers
(Fisher Scientifie, Pittsburgh, PA, USA) equipped with a resealable glass tube filled with
glass wool to allow headspace removal and filling. A portion of the air headspace was
removed and refilled with methane of 99.99% purity (Strate Welding, Jacksonville, FL,
USA) using a vacuum pump assembly to achieve a headspace concentration of air with
20% (v/v) methane.
For solid culturing, 1.5% (w/v) of Bacto agar (Difco Laboratories, Detroit, MI,
USA) was added to the NMS medium. All plates were incubated in a sealed desiccator,
containing anhydrous CaSO4 (Drierite, W.A. Hammond Drierite Company, Xenia, OH,
USA) under an atmosphere of 20% methane and 80% air (by volume) at 300C that was
refreshed every four to Hyve days. Purity of the cultures was verified by routine streaking
on 2% (w/v) nutrient agar in doubly deionized water.
sMMO expression was qualitatively verified by a naphthalene assay modified from
Brusseau et al. (1990). Four negative controls--autoclaved cells, cells cultured with 10
Cpmol 1-1 Cu(NO3)2 (fOr expression of pMMO), cell-free, and cells that have been
subj ected to addition of one to two ml of acetylene gas (a known inhibitor of MMO, Prior
and Dalton (1985))--were included with three live samples of active test culture diluted
to an absorbance of 0.2 (at a wavelength of 600 nm) and transferred to autoclaved 10-ml
capped test tubes. Seventy mg of crushed naphthalene (Sigma, St. Louis, MO, USA)
were added to each tube. After incubation at 300C and 250 rpm for a minimum of one
hour, 0. 1 ml of freshly prepared 4.21 mmol 11 tetrazotized ortho-dianisidine (Sigma, St.
Louis, MO, USA) solution was added. A subsequent pink-to-purple color formation in
the tubes indicated positive sMMO activity that was verified using spectrophotometry
(Fisher Scientifie, Pittsburgh, PA, USA) at 550 nm.
Genomic DNA was isolated from Strain CSC1, grown to exponential phase, by a
standard method (Ausubel et al., 1989). 16S rRNA gene was amplified by PCR using the
universal bacterial primers 27f and 1492r (Lane, 1991). PCR primers used for sMMO
were mmoXA (5 '-ACCAAGGARCARTTCAAG-3 ') and mmoXB (5'-
TGGCACTCRTARCGCTC-3 ') (Auman et al., 2000); for methanol dehydrogenase
(MDH), mxa fl003 (5' -GCGGCACCAACTGGGGCTGGT-3 ') and mxa rl561 (5'-
GGGCAGCATGAAGGGCTCCC-3 ') (McDonald and Murrell, 1997); and for pMMO,
Al89f (5' -GGNGACTGGGACTTCTGG-3 ') and A682r (5'-
GAASGCNGAGAAGAASGC-3 ') (Holmes et al., 1995).
All PCR reactions were carried out in a PTC-200 Thermo Cycler (MJ Research,
MA, USA) using 25 Cl1 reactions and Premix Taq polymerase (Takara, Otusu, Shiga,
Japan). Conditions used for the primer sets have been described previously (Holmes et
al., 1995; Costello and Lidstrom, 1999; Auman et al., 2000). The PCR amplification
products were ligated to vector pCR2. 1 (Invitrogen, Carlsbad, CA, USA) and
transformed to competent E. coli cells (TOP 10F') according to the vendor' s instructions.
Plasmid DNA from transformants was isolated and the inserts sequenced by the
Biotechnology Resource Center at Cornell University (Ithaca, NY, USA).
Sequences were compared with previously identified sequences in the National
Center for Biotechnology Information (NCBI) database using BLAST (Altschul et al.,
1990). The 16S rRNA gene from Strain CSC1 was also aligned with sequences obtained
from the Sequence Match program provided by the Ribosomal Database Proj ect II (RDP-
II) (Cole et al., 2005). Phylogenetic trees were generated using PHYLIP version 3.6
(Felsenstein, 2004) and viewed using Treeview (Page, 1996). The GenBank accession
numbers for the 16S rRNA, MDH, sMMO and pMMO gene sequences obtained in this
study are DQ364433, DQ664499, DQ664498, and DQ364434, respectively.
DNA-DNA hybridization was performed on Strain CSC1 by DSMZ
(Braunschweig, Germany) against M~ethylocystis echinoides strain IMET 10491 was
performed using 2 x SSC buffer (0.3 M NaC1, 0.03 M sodium citrate, pH 7.0) + 10%
(v/v) formamide at an optimal renaturation temperature of 680C.
Earlier studies reported Strain CSC1 as gram-negative, non-motile, coccobacillus,
possessing an internal membrane structure characteristic of Type II methanotrophs
(paired membranes inside the periphery of the cell), and forming lipid inclusions (Henry
and Grbic-Galic, 1990; Henry and Grbic-Galic, 1991; Hrliak and Begonja, 1998). Fang et
al. (2000) concluded that the intact phospholipids of Strain CSC1 clustered within the
Type II grouping, clearly distinct from groupings of Type I methanotrophs. This study
extended the previous phenotypic characterization studies by assessing exospore and
rosette formation, growth at 370C, the presence of a surface (S-) layer, carbon and
nitrogen source utilization, and lysis by 2% (w/v) sodium dodecyl sulfate (SDS), all
identified by Bowman et al. (1993a) or Hanson and Hanson (1996) as differentiating
characteristics among Type II methanotrophic species.
Exospore formation was determined with one- to two-week-old broth cultures
grown as previously described following methods of Smibert and Krieg (1981). Five ml
of culture were transferred in duplicate to fresh NMS medium for controls. A second set
of duplicates was heated in a water bath at 800C for 20 min for pasteurization. Growth
was monitored after streaking the controls and treated cultures onto solid NMS plates and
incubation (as previously described) for 21 days. Exospores were monitored using light
microscopy, also used to determine rosette formation (Norris and Ribbons, 1971).
Growth in liquid culture was monitored using 250-ml nephlos flasks with the same
stopper assembly described above and a spectrophotometer (Fisher Scientific, Pittsburgh,
PA, USA) at a wavelength of 600 nm.
Nitrogen and carbon sources were tested using NMS basal medium. To test for
alternate nitrogen sources, KNO3 WAS replaced with 0. 1% (w/v) of anhydrous L-
asparagine (MP Biomedicals, Irvine, CA, USA), L-aspartate (Pfaltz & Bauer, Waterbury,
CT, USA), or L-glutamine (MP Biomedicals, Irvine, CA, USA) (all shown to support
growth of Methylocystis echinoides and M\~ethylocystis parvus, Bowman et al., 1993a) or
L-lysine monohydrochloride (Sigma-Aldrich, St. Louis, MO, USA), L-ornithine
hydrochloride (MP Biomedicals, Irvine, CA, USA), or putrescine (MP Biomedicals,
Irvine, CA, USA) (all shown to support growth of Methylosinus trichosporium, Bowman
et al., 1993a). NMS medium with KNO3 and without a nitrogen source served as positive
and negative controls, respectively, and the latter control also served as a test to fix
atmospheric nitrogen. To test for alternate carbon sources, 0.2% (w/v) of methylamine
hydrochloride (Alfa Aesar, Ward Hill, MA, USA), dimethyl sulfoxide, methanol, or
glucose (Fisher Scientifie, Pittsburgh, PA, USA) was added. Over the 30-day test period,
flasks were prepared in duplicate, and transfers were made to fresh medium and nitrogen
or carbon source every 4 days. Growth measurements were performed as described
Lysis by 2% (w/v) SDS (Fisher Scientifie, Pittsburgh, PA, USA) was determined
by direct microscopic observation using cells harvested at %/-log phase. Cells were
centrifuged at 2460 x g for 20 min, resuspended in the 2% SDS stock solution for
approximately 2 hours, and observed using an oil immersion phase contrast microscope
(Zeiss, Oberkochen, Germany).
Transmission electron microscopy was used to observe cells of Strain CSC1
expressing MMO, lipid inclusions, and other Eine structural features, including S-layers.
Liquid cultures were incubated for two to three days and were Eixed for 30 min at room
temperature with cacodylate-buffered glutaraldehyde both with and without 0. 1% Alcian
blue (Fassel et al., 1992), stained for 30 min at room temperature with 1% cacodylate-
buffered osmium tetroxide, and then stained for 50 min in 1% aqueous uranyl acetate.
After dehydrating in increasing strengths of ethanol, cells were embedded in both Spurr' s
and Epon resins (Dykstra, 1993). Thin sections were prepared and stained with lead
citrate and examined on a Zeiss EM-10CA transmission electron microscope.
M~ethylocystis echinoides was observed by negative stain using 1% aqueous uranyl
acetate applied to cell suspensions on Formvar-coated grids.
In order to provide evidence that the observed S-layer is glycoprotein, two
additional cytochemical approaches were utilized. Images of Alcian blue-stained
specimens were compared to those with no Alcian blue in the glutaraldehyde Eixative,
since Alcian blue stains polysaccharide moieties (Lewis and Knight, 1977). Secondly,
thin Epon sections on Formvar-coated nickel grids were first exposed to 3% hydrogen
peroxide (H202) for 15 min at room temperature to remove osmium and then exposed to
1% aqueous pronase solution (Sigma Chemical Company, St. Louis, MO, USA) for 60-
90 min at 350C to remove protein components from the section (Monneron and Bernhard,
1966; Lewis and Knight, 1977). Controls included H202 alOne and water substituted for
the pronase step.
The phylogenies of the 16S rRNA, sMMO, MDH, and pMMO genes, shown in
Fig. 2-1 and Fig. 2-2 (a-c), are consistent with placement of Strain CSC1 with other
known Type II methanotrophs. The 16S rRNA phylogeny of Strain CSC1 clearly places
it within a branch of the alpha-Proteobacteria dominated by M~ethylocystis species. This
methanotroph shares 98% 16S rRNA gene sequence similarity with its nearest defined
relatives, an uncultured member of the M~ethylocystaceae (AF358021), as well as two
cultured organisms: M~ethylocystis sp. L32 (AJ83 1522) and M~ethylocystis sp. SC2
(AJ4313 84), although 16S rRNA gene similarity is not sufficient to place Strain CSC1 to
the species level. The DNA-DNA hybridization results showed that Strain CSC1
possesses a 3.8% DNA-DNA similarity with 2ethylocystis echinoides strain IMET
As shown in Table 2-1, rosette formation by cells of Strain CSC1 was not
observed. No growth was evident after pasteurization, indicating that this methanotroph
is not resistant to heat, and growth was also not observed at 370C. Optimum growth was
observed at approximately 300C. Strain CSC1 was not lysed by a 2% solution (w/v) of
SDS, but a 10% solution (w/v) of SDS did lyse the cells. It was shown to be capable of
growing on alternate nitrogen sources of L-asparagine, L-aspartate, L-glutamine, L-
ornithine and putrescine; however, no growth was visible in the presence of L-lysine. Of
the four alternate carbon sources of methylamine, dimethylsulfide, methanol, and glucose
tested, only methanol supported growth of Strain CSC1.
The expression of sMMO upon culturing Strain CSC1 in NMS medium with no
copper was confirmed by formation of a purple color after incubation with naphthalene
and addition of ortho-dianisidine, whereas controls with acetylene and with cells cultured
in the presence of copper yielded no color. These results strongly suggest that sMMO
was expressed in Strain CSC1 when grown without copper and was responsible for
Transmission electron micrographs of Strain CSC1 grown in the presence of copper
verify the Type II membrane structure of paired membrane lamellae in the peripheral
cytoplasm (Fig. 2-3a,b). In thin section, a variety of cell shapes were visible at low
magnifieation (Fig. 2-3a), but elongated or dumbbell shapes of cells predominated.
Many of the other profies could represent dumbbell shapes sectioned in different planes.
Cells grown without copper contained only a few internal membranous lamellae (data not
shown). Polyphosphate bodies and lipid inclusions were common.
As shown in Figs. 2-3(a) and 2-3(b), distinctively striking S-layers, likely
composed of glycoprotein, were revealed with transmission electron microscopy of
ultrathin sections of Strain CSC1 Eixed with Alcian blue. These spiked S-layer structures,
50-75 nm in height, covered the entire surface of the cell wall. We have seen that the
cytoplasm of cells of Strain CSC1 embedded in Spurr resin will sometimes shrink away
from the wall, lending support to the idea that the S-layer is more rigid than the rest of the
wall (data not shown). This shrinkage does not occur in cells embedded in Epon resin
(e.g., cells in Fig. 2-3a,b).
S-layers have been observed in both Type I and II methanotrophs isolated from a
wide range of environments, including the genera M~ethylomicrobium, Methylomona~s,
M~ethylosinus, and M~ethylocystis (Fassel et al., 1992; Sorokin et al., 2000; Trotsenko and
Khmelenina, 2002). Type II 2ethylosinus trichosporium OB3b was found to have bead-
like S-layer structures and occasional filamentous material in the outer envelope (Fassel
et al., 1990; Fassel et al., 1992). Similar bead-like S-layer structures were observed in the
cell envelope of2~ethylocystis sp. strain Lake Washington but not in M\~ethylocystis paris,
and the authors concluded that the absence of these structures in the latter species could
be a species variation. Both M~ethylocystis species possessed considerable filamentous
material, however (Fassel et al., 1992). M~ethylocystis echinoides strain IlVET 10491 was
reported to have rigid tubular structures arranged radially at the cell surface (Gal'chenko
et al., 1977), features that are absent from M\~ethylocystis parvus (Heyer et al., 2002). In
this study, negative stain preparations of2~ethylocystis echinoides show ellipsoid cells
with square-ended tubular proj sections (Fig. 2-3c) that appear striated at high
magnification (Fig. 2-3d). This striation was not reported in previous studies of this
strain (Gal'chenko et al., 1977; Bowman et al. 1993a), and the tubular appearance of this
S-layer is much different from the solid-sharp spines of Strain CSC1.
To further elucidate the nature of the spiked S-layer in Strain CSC1 cells Eixed with
glutaraldehyde alone (Fig. 2-4a) were compared with those Eixed with an Alcian
blue/glutaraldehyde mixture (Fig. 2-4b). Alcian blue is a differential stain for
polysaccharide (Lewis and Knight, 1977), and the spines in Fig. 2-4b were considerably
darker, longer, and more distinct than the same structures in Fig. 2-4a, even though Fig.
2-4b is at a lower magnification. This strongly indicates polysaccharide content. After
treatment with H202 to remove osmium from the Epon sections, sections treated with
pronase, a broad-spectrum protease, lost the entire S-layer (Fig. 2-4c,d), indicating that
the layer contains considerable protein.
Sequence analysis of the 16S rRNA gene (Fig. 2-1), the sMMO gene (Fig. 2-2a),
the methanol dehydrogenase gene (Fig. 2-2b), and the pMMO gene (Fig. 2-2c) supports
placement of Strain CSC1 within the closely related genera 2ethylocystis and
Given the suspected placement in the genera M~ethylocystis, DNA-DNA
hybridization was performed with M~ethylocystis echinoides. Based on the DNA-DNA
hybridization results showing only 3.8% similarity, Strain CSC1 does not belong to the
species M~ethylocystis echinoides, following the threshold value recommendation of
Wayne et al. (1987). Phenotypic results in Table 2-1 show that Strain CSC1
differentiates from M~ethylosinus trichosporium, M~ethylocystis echinoides, and
M\~ethylocystis parvus, three closely matching cultured strains in the phylogenetic
analysis, in various characteristics. All of the strains shown in Table 2-1 have been
reported to be oxidase- and catalase-positive, possess colonies that are of opaque
transparency, smooth edge, convex elevation, form poly-P-hydroxybutyrate, grow on
methanol, and capable of fixing atmospheric nitrogen. Unlike the known strains, Strain
CSC1 was not capable of growth at 370C; however, all of the methanotrophs grow
optimally near 300C. It is important to note that Gal'chenko et al. (1977) and Bowman et
al. (1993a) report conflicting information concerning the ability ofM~ echinoides to
accumulate poly-P-hydroxybutyrate and grow at 370C, the former reporting positive
results for each and the latter reporting negative results. Our TEM and growth studies
with this strain agreed with Gal'chenko et al. (1977) (data not shown).
The elongated dumbbell shape of Strain CSC1, lack of motility, and ability to form
polyphosphate separate it from the M~ethylosinus trichosporium. Other distinguishing
characteristics between Strain CSC1 and M~ trichosporium include smaller cell size, S-
layer morphology, and lack of heat resistance. Also, unlike reported observations ofM~
trichosporium, Strain CSC1 can use L-asparagine, L-aspartate and L-glutamine and
cannot use L-lysine as nitrogen sources. Both strains share the ability to use L-ornithine
Most similarities, however, are shared with the two M~ethylocystis strains, including
lack of motility, heat resistance, and rosette formation. Strain CSC1's cell shape, ability
to form polyphosphate, and colony color of yellow-white differ from M~ethylocystis
echinoides and M\~ethylocystis parvus (Table 2-1). Unlike M~ echinoides, Strain CSC1 is
capable of using L-ornithine and putrescine as nitrogen sources, whereas, unlike M~
parvus, Strain CSC1 is not capable of using L-lysine. In addition, as reported for M
trichosporium, M~ echinoides, and M~ parvus, Strain CSC1 is not lysed by a 2% solution
(w/v) of SDS.
While Strain CSC1 has previously been shown by TEM to contain characteristic
Type II membranes (Henry and Grbic-Galic, 1990; Hrliak and Begonja, 1998), it was
revealed here to accumulate both polyphosphate bodies (Fig. 2-2) and poly-P-
hydroxybutyrate storage granules, consistent with M\~ethylocystisparvus (Bowman et al.,
1993). No study has reported the structure of Strain CSC1's cell envelope in comparison
to that of other well-characterized methanotrophs. Of special interest are the surface- (S-)
layers, regular crystalline surface layers in Archaebacteria and Eubacteria, composed of
protein or glycoprotein subunits (Sleytr et al., 1993; Sidhu and Olsen, 1997).
It is not known why S-layers develop in some strains of closely related bacteria and
not in others. However, one hypothesis is that formation of these structures reflects
adaptation to an ecological niche (Easterbrook and Alexander, 1983; Easterbrook, 1989)
or a response to exposure to harsh environments (Minsky et al., 2002). Others suggest
that S-layers may provide microorganisms with a selective advantage by serving as a
protective coating or as molecular porins or sieves and traps for substrates, in maintaining
the rigidity of the cell envelope, or providing a means of cell adhesion and surface
recognition (Sara and Sleytr, 1987; Sleytr and Messner, 1988; Sara et al., 1992; Sidhu
and Olsen, 1997). Easterbrook and Sperker (1982) hypothesized that spinae may
simultaneously fulfill many fortuitous roles, analogous to "arms" with multipotential
activities, including attachment, distance-keeping, and protection. However, why some
species are prone to spine formation and others not, why S-layers exist in a variety of
shapes and symmetries, and why these structures develop among species of
methanotrophs is not clearly understood.
M~ethylocystis echinoides strain IlVET 10491 was isolated from lake mud in Russia
(Gal'chenko et al., 1977), possibly a more nutrient-rich environment than the sediments
of the uncontaminated groundwater aquifer in California where Strain CSC1 was
isolated. Echinoides is the Latinized adjective derived from the Greek word echinos,
meaning "hedgehog," named for the hedgehog-like appearance of this bacterium.
However, as reported by Gal'chenko et al. (1977), and verified in this study (Fig. 2-3c,d),
the spines on this methanotroph appear to be tubular and less dense in comparison to the
spikes observed on Strain CSC1, which would be more aptly named for a hedgehog.
Despite the different originating environments of these two strains, proximity in the
grouping of the M~ethylocystis genus, as strongly suggested by the results of this study,
adds credence to the hypothesis that phylogeny and ecology may both play a role in S-
layer formation. Similar clustering of S-layer-producing strains of Bacillus cereus has
been observed, and, similar to these results, strains in this cluster do not possess S-layers,
while others do (Mignot et al., 2001). These authors concluded that ecological pressure
is associated with the acquisition and maintenance of S-layers in hosts that fall into a
Phylogenetically, Strain CSC1 is most closely related to 2ethylocystis sp. Its cell
size, rosette formation, and presence of surface layers are most similar to M~ethylocystis
echinoides. However, Strain CSC1 showed only 3.8% similarity with2~ethylocystis
echinoides by DNA-DNA hybridization, and these two strains showed differences in
surface-layer morphology, cell shape, colony color, formation of polyphosphate, and
ability to use L-ornithine or putrescine as a nitrogen source. Characteristics of cell shape
and the presence of surface layers, genes encoding for slV1VO expression, and ability to
use L-lysine as a nitrogen source are divergent from those of 2ethylocystis parvus. The
lack of polar flagella, smaller cell size, different cell shape, lack of heat resistance,
presence of polyphosphate, ability to use L-asparagine, L-aspartate, or L-glutamine and
inability to use L-lysine as a nitrogen source differentiate Strain CSC1 from 2ethylosinus
trichosporium. Accepting these differences, Strain CSC1 could be described as a new
species in the M~ethylocystis genus. We proposed this species be name M~ethylocystis
aldrichii sp. nov.
Description of Methylocystis aldrichii sp. nov.
M~ethylocystis aldrichii sp. nov. (al.drich'i.i ML gen. N. aldrichii of Aldrich;
named after H. C. Aldrich, an American microbiologist, deceased August 9, 2005).
Cells are aerobic, gram-negative, 0.3-0.6 x 0.7-1 Clm in size that occur singly or in
clusters. Reproduces by normal cell division. Budding division does not occur. Cells
are not motile but possess a spiney surface layer composed of polysaccharide. Produces
oxidase and catalase. Forms lipid cysts. Poly-P-hydroxybutyrate accumulates. Contains
Type II intracytoplasmic membranes which are aligned parallel to the cell wall. Type II
methanotroph. Methane and methanol are the sole sources of carbon and energy.
Capable of using KNO3, L-asparagine, L-aspartate, L-glutamine, L-ornithine, and
putrescine as nitrogen sources. Capable of fixing atmospheric nitrogen. Expresses
sMMO under low copper concentrations. Capable of cometabolically oxidizing a variety
of aliphatic and aromatic compounds. Not resistant to pasteurization. Is not lysed by 2%
(w/v) SDS. Is lysed by 10% (w/v) SDS. Colonies are white/yellow, slow-growing and
0.8-1.5 mm in diameter after 17-18 days at 300C on NMS agar plates, incubated in the
presence of 20% by volume methane in the headspace of a sealed desiccator. No growth
on complex organic media. Optimal pH for growth is 7.0; does not grow at pH 4.0 or
9.0. Is not capable of growth at 370C. Optimal temperature for growth is approximately
300C. The type strain, Strain CSC1 (ATCC BAA-1344), was isolated from an
uncontaminated groundwater aquifer in the mid-1980s from Moffett Naval Air Station in
Mountain View, CA, USA.
Methylocystis sp. KS33
Methylocystis sp. 39
Methylocystis sp. SC2
Methylocystis sp. L32
Uncultured bacterium L013.7
Methylocystis sp. 5FB2
Methylocystis sp. 50/54
Methylocystis sp. SV97
Methylocystis sp. DWT
58 Methylosinus parvus (Y18945)
55 Methylocystis echinoides str.
IMET 10491 (AJ458473)
Methylosinus sporium SK13
Figure 2-1. 16S rRNA phylogeny of Strain CSC1 and related M~ethylosinus and
M~ethylocystis species. Numbers at branch points represent bootstrap values
based on 100 replicates.
Methylorystis sp M
Uncultured clone LOPB13
51 1 Strain CSC1
100 Methylocystis sp. 50/54
100 1 Methylocystissp. B2/7
Methylocystis parvus OBBP
99 Mehlsnssporium str. KS16
Methylocystis sp. M
Methylosinus sp. LW4
10 Methylosinus sp. LW3
M~et 1702ins ssp.LW8
100 Methylocystis sp. 45/7a
S rain CS 1
Uncultured bacterium clone W9
56rMethylocystis sp. 51
Methylocystis sp. 41
53U~n Itr~ed bacterium clone 9
Methylocystis sp. IM ET 10486
~IMET 10486 (AJ459077)
(Met86y cystis sp. 5FB2
(AJ8h6y ytisp L
~Methylocystis sp. DWT
Methylocystis sp. SK28
100 IMET 10489 (AF488302)
Figure 2-2. Functional genes phylogenies of Strain CSC1. (a) Phylogenetic tree of Strain
CSC1 soluble methane monooxygenase (sMMO) gene sequence. (b)
Phylogenetic tree of Strain CSC1 methanol dehydrogenase (MDH) gene
sequence. (c) Phylogenetic tree of Strain CSC1 particulate methane
monooxygenase (pMMO) gene sequence. Numbers at branch points represent
bootstrap values based on 100 replicates.
Table 2-1. Phenotypic characteristics differentiating Strain CSC1 from M~ethylosinus
trichosporiurm, Me~cthyloc:ystis echinoidesy, and Miethylocy:~ stis palrvus.'
Characteristic Strain CSC1
trichosporium echmnoldes parvus
Yellow/ White/pale White/pale
Color White/buff .
white pink pink/tan
Width (Cim) 0.3-0.6 0.5-1.5 0.6-0.8 0.3-0.5
Length (Cim) 0.7-1 2-3 0.8-1.2 0.5-1.5
Sha e Dumbbell Rods, Pear-shape, Pear-shape,
Pyriform Ovoid Ovoid
Sharp, solid Bead-like/ Tubular
S-layersspinestf filamentous spines
Polyphosphate +f +
Poy--+ + +TI +
Motility -Polar flagella
Rosettes -Jf +
Heat resistance -Jf +
2%(w/v) SDS lysed -Jf
Growth at 37oC -Jf + +TI +
N,-source and use:
L-asparagine +t + +#
L-aspartate +f + +#
L-glutamine +t + +#
L-lysine -"f + -+#
L-ornithine +"r + -+#
Putrescine +"r + -+#
NMS no N2-SOUTCe "
C-source and use:
Methylamine -Jf -f
Methanol +"r + +"r +
Glucose -Jf -Jf
'References: Whittenbury, 1970; Gal'chenko et al., 1977; Fassel et al., 1990, Fassel et al. 1992;
Henry and Grbic-Galic, 1991; Bowman et al., 1993a; Hanson and Hanson, 1996; Hrliak and
Begonja, 1998. This study. Gal'chenko et al. (1977) reported that M echinoides forms lipid
cysts and does grow to a limited extent at 37oC. Bowman et al. (1993a), however, reported that
this strain does not accumulate poly- B-hydroxybutyrate, and only 0-10% of the strains tested
grew at 37oC. #Reported by Bowman et al. (1993a) as 75-87% of the strains were positive.
Figure 2-3. Transmission electron microscopy photographs of Strain CSC1 and
M~ethylocystis echinoides. Panels (a) and (b) show the morphology of cells of
Strain CSC1 grown with 10 ELM Cu. In panel (b), numerous lamellae (La) are
present. Lipid inclusions (Li) and polyphosphate (P) storage inclusions are
also present. S indicates a surface view of the spiny surface of a cell. Cells of
M~ethylocystis echinoides viewed with negative stain (panels (c) and (d)) are
elliptical in profile and have numerous tubular projections from the surface.
Some may be seen in circular end-on profile at the upper right of panel (c).
The high magnification view in panel (d) shows that the tubes are striated.
Markers in panels (a) and (b) indicate 1 Epm; in panel (c), 0.5 Epm; in panel (d),
Figure 2-4. Electron microscope cytochemistry of the S-layer of Strain CSC1. Panel (a)
shows the surface of a cell fixed initially with glutaraldehyde alone. Spiny
layer is indistinct and lightly stained. Panel (b) shows the surface of a cell
fixed initially with a glutaraldehyde/Alcian blue mixture to selectively stain
polysaccharide. Compared to panel (a), the spiny layer stains darker and is
thicker and is more distinct. Panels (c) and (d) show cells after pronase
digestion. In panel (c), a cross section, a light layer around the cell has been
left where the pronase removed the protein in the S-layer. In panel (d), a
grazing section of the spines at the cell surface, numerous light spots in the
plastic show where the pronase removed the spikes. Markers represent 0.2
EFFECTS OF ALPHA-PINENE AND TRICHLOROETHYLENE ON OXIDATION
POTENTIALS OF IVETHANOTROPHIC BACTERIA
Note: Published manuscript (Pacheco and Lindner, 2005)
Pacheco, A., and Lindner, A.S. (2005) Effects of alpha-pinene and trichloroethylene on
oxidation potentials of methanotrophic bacteria. Bulletin ofEnvironmental
Conttttttttttttttttttamnt io and Toxicology 74:133-140.
Trichloroethylene (TCE), a widely used solvent notable for its degreasing
properties, is a common environmental contaminant that poses significant risk to public
health ((ATSDR), 1999). TCE has been shown to be effectively removed from soil and
water by phytoremediation, often favored over other methods because of its
effectiveness, low cost, and aesthetic benefits. More rapid TCE removal has been
observed in the root zone of plants (rhizosphere) used in phytoremediation (Walton and
Anderson, 1990; Anderson and Walton, 1995; Brigmon et al., 1999), and methanotrophs,
methane-oxidizing bacteria that thrive on oxygen and methane and are capable of co-
oxidizing TCE (Wilson and Wilson, 1985; Little et al., 1988), have been implicated in
this increased activity (Brigmon et al., 1999).
Loblolly pines (Pinus taeda), shown to support large rhizosphere populations of
methanotrophs. (Brigmon et al., 1999), have been considered for TCE remediation. These
trees produce and release significant quantities of monoterpenes, the most predominant
being (R)-a-pinene, composing over 65% of the total oleoresin composition in different
plant tissues (Phillips et al., 1999). Since concentrations of (R)-a-pinene have been
observed to be as high as 1.4 mg gl in fresh litter layers of pine forest soils (White,
1994), the probability that soil microorganisms encounter these compounds in nature is
high. Previous studies have shown that (R)-a-pinene has a concentrate on-dependent
inhibitory effect on methane oxidation by methanotrophs (Amaral and Knowles, 1997;
Amaral et al., 1998; Amaral and Knowles, 1998), and, thus, may impact not only the
growth of these bacteria in the rhizosphere but also their ability to co-oxidize TCE.
While methanotrophs. were shown to regain methane oxidation activity one to three days
after exposure to (R)-a-pinene (Amaral et al., 1998), the implications of the long-term
presence of this monoterpene on methanotrophic activity in the rhizosphere, in particular
concentration effects of this chemical and its influence on TCE removal potentials, are
To this end, this study sought to first assess the ability of representative Type I, II,
and X methanotrophs, grouped by their differences in carbon assimilation pathways,
intracytoplasmic membrane structures, fatty acid carbon lengths, and phylogeny
(Bowman et al., 1993a), to oxidize (R)-a-pinene over a range of concentrations using
oxygen uptake analysis. Secondly, this study sought to gain a better preliminary
understanding of the variation in oxygen uptake responses to mixtures of (R)-a-pinene
and TCE by representative methanotrophs, thus ultimately providing insight into the
effect of (R)-a-pinene on TCE oxidation potentials of these bacteria and guidance for the
phytoremediation practitioner to more accurately predict the extent of TCE
rhizodegradation when using monoterpene-releasing plants. We report herein
observations of the potential of methanotrophs. to oxidize (R)-a-pinene over a broad
range of concentrations and (R)-a-pinene/TCE mixture effects on methanotrophic
oxygen uptake activity.
Materials and Methods
Methanotroph strains used in this study included Type I2~ethylomicrobium album
BG8 (ATCC 33003) and Type II 2ethylosinus trichosporiunt OB3b (ATCC 35070),
obtained from Dr. Jeremy Semrau (University of Michigan, Ann Arbor, MI, USA), and
Type X M~ethylococcus capsulatus (Bath) (ATCC 33009), purchased from the American
Type Culture Collection (Manassas, VA, USA). Cultures were grown in nitrate mineral
salts (NMS) medium (Whittenbury et al., 1970), with or without 10 CtM Cu(NO3)2 to
provide conditions for expression of pMMO or sMMO, respectively. With the exception
of M capsulatus (Bath), incubated at 45 oC with 50% methane (99.99% pure, Strate
Welding, Jacksonville, FL, USA) in the headspace, all organisms were routinely
subcultured in sealed erlenmeyer flasks containing 20% methane in the headspace and
incubated at 30oC in a rotary shaker at 250 rpm, as previously described (Lindner et al.,
2000). Purity of the cultures was verified by routine streaking on 2% (w/v) nutrient agar
plates (Difco, Sparks, MD, USA). Expression of sMMO was qualitatively verified by a
naphthalene assay modified from Brusseau et al. (1990) and described by Lindner et al.
Oxygen uptake analysis was performed in this study, as it has been shown to be a
rapid, effective means of assessing oxidative potential of whole cells (Lindner et al.,
2000; Lindner et al., 2003). (R)-a-pinene was chosen to represent monoterpenes because
it is a major component of loblolly pine oleoresin (Phillips et al., 1999). (R)-a-pinene
and TCE were obtained in the highest purity available from Aldrich Chemical Co.
(Milwaukee, WI, USA). Standard solutions of 10 Eomol mlF were prepared in 1,4-
dioxane (Fisher Scientifie, Pittsburgh, PA, USA), used as the carrier solvent because it
easily solubilized the substrates, was not oxidized by any of the cultures studied, and
caused no probe effects during oxygen uptake analysis (Lindner et al., 2000). Resting-
cell suspensions were prepared from 500 ml cultures harvested at %/-log phase by
centrifugation in a J2-HS Beckman floor model centrifuge (Beckman-Coulter, Fullerton,
CA, USA) at 2460 x g, 4oC, for 20 min. To ensure removal of all methane, the cells were
washed with NMS medium, recentrifuged, and resuspended in the NMS medium to a wet
cell concentration of 0.2 g ml l. The oxygen uptake system was composed of a 1.9 ml,
well-stirred, enclosed reactor held at room temperature, as described by Lindner et al.
(2000). After assessing the ability of methanotrophs to oxidize TCE and (R)-a-pinene
alone, the study proceeded to investigate the effect of (R)-a-pinene on TCE oxidation by
adding both substrates simultaneously into the oxygen uptake system before addition of
the resting cells.
Despite storage of the resuspended cells on ice throughout the oxygen uptake
experiments, loss of cell activity over time was observed. To ensure comparability of
measurements throughout the 2-3-day testing period, all rates of oxygen uptake were
normalized to the rates observed with 4 ml of methane gas, measured just prior to a
change to a new substrate concentration. Details of this normalization procedure are
presented in Lindner et al. (2000). The electrode was calibrated at least daily with a
saturated sodium sulfite solution, and "live" runs were performed at least in triplicate for
each concentration tested. All runs were corrected for endogenous metabolism.
Controls without cells and with 4 ml of acetylene gas, a known inhibitor of MMO (Prior
and Dalton, 1985), were routinely run to verify that depletion of oxygen, hence, oxidation
activity, was a result of MMO activity. Initial rates of oxygen uptake were calculated by
linear or polynomial fits to the data points using Microsoft Excel software (Microsoft
Corp., Redmond, WA, USA).
Results and Discussion
M. trichosporium OB3b and M~ capsulatus (Bath), when cultured with no copper,
expressed positive sMMO activity, as evidenced by a bright pink-to-purplish color in the
assay. All of the strains tested negative for sMMO activity (no color change observed)
when cultured with 10 CIM Cu(NO3)2. SMMO and pMMO expression under culturing
conditions without and with copper, respectively, was thus assumed, a reasonable
conclusion given that enzyme expression in these methanotrophs under these conditions
is well characterized. Active resting cells of all three representative methanotrophs
consumed oxygen over a range of TCE and (R)-a-pinene concentrations, regardless of the
type of MMO expressed (Fig. 3-1, A-E). No oxygen uptake was observed after addition
of acetylene or without cells present, verifying MMO activity in all cases. As shown in
Figure 3-1, regardless of the methanotroph or substrate tested, a maximum rate of oxygen
uptake was observed, followed by a rapid decrease in rates, suggesting toxic effects of
either the substrate itself or of oxidation products formed. This oxygen uptake behavior
has been reported previously for methanotrophs with aromatic substrates (Lindner et al.,
2000), and, while both substrates have been shown to have toxic effects on
methanotrophic activity (Fox et al., 1990; Alvarez-Cohen and McCarty, 1991a; Henry
and Grbic-Galic, 1991; Oldenhuis et al., 1991; White, 1994; Amaral and Knowles, 1997;
Amaral et al., 1998; Amaral and Knowles, 1998), there have been no previous reports on
the effects of a range of substrate concentrations on relative activities.
As shown in Figure 3-1, methanotrophs expressing sMMO (plots A, C) oxidized
TCE at higher maximum rates than those expressing pMMO (plots B, D, E), as
previously reported (Little et al., 1988; DiSpirito et al., 1992; Lontoh and Semrau, 1998).
The maximum normalized rates of oxidation by M~ trichosporium OB3b and M~
capsulatus (Bath) expressing sMMO or pMMO were 0. 11 + 0.01 and 0.03 + 0.00 and
0.05 + 0.01 and 0.03 + 0.01, respectively (Fig. 3-1, A-D), while the maximum rate
expressed by M~ album BG8, capable of pMMO expression only, was 0.05 + 0.01 (Fig. 3-
1, E). The TCE concentrations where the observed normalized oxygen uptake rate was
the highest ranged from 20 to 35 ppm for the tested strains. M~ trichosporium OB3b and
M~ capsulatus (Bath) expressing sMMO exhibited oxygen uptake maxima at higher TCE
concentrations (35 ppm) than when expressing pMMO (20-25 ppm), and M~ album BG8
expressing pMMO showed a maximum observed rate at 35 ppm TCE. These results do
suggest differing sensitivity levels to TCE, depending on the methanotroph and type of
As observed with TCE, both sMMO-expressing methanotrophs were also capable
of oxidizing (R)-a-pinene at higher rates than their pMMO-expressing counterparts (Fig.
3-1, A-D). The maximum normalized rate of oxygen uptake by M~ trichosporium OB3b
expressing sMMO was almost 10 times the rate observed with pMMO-expressing cells
(0.28 + 0.04 and 0.02 + 0.01, respectively); however, both rate maxima occurred at 20
ppm (R)-a-pinene (Fig. 3-1, A, B). The maximum normalized oxygen uptake rate with
M~ capsulatus (Bath), expressing sMMO, was 0. 10 + 0.02 at 20 ppm (R)-a-pinene,
compared to 0.08 + 0.01 at 50 ppm (R)-a-pinene under pMMO expression (Fig. 3-1, C,
D). The observed maximum normalized rate of oxygen uptake by M. album BG8 was
0.04 + 0.00, between the values observed for the other two strains under pMMO
expression (Fig. 3-1E). Previous studies have reported higher TCE oxidation rates by
pure methanotrophs under sMMO expression (Wilson and Wilson, 1985; Little et al.,
1988; DiSpirito et al., 1992; Lontoh and Semrau, 1998); however, this is the first report
of such a trend with (R)-a-pinene. These results bring direct relevance to the
environment, as sMMO expression in methanotrophs occurs only at very low copper
concentrations (Lontoh and Semrau, 1998). Measurement of bioavailable copper is
essential, therefore, for effective prediction of methanotrophic activity potential.
The response of each methanotroph in the presence of 20 ppm TCE over a range of
(R)-a-pinene concentrations is shown in Figure 3-2, A-E. This plot presents the change
in normalized oxygen uptake rate with 20 ppm TCE alone caused by the presence of
different concentrations of (R)-a-pinene and thus represents the influence of (R)-a-
pinene on TCE oxidation and provides insight into mixture effects on methanotroph
activity. The concentration of 20 ppm TCE was chosen because it was not observed to be
toxic to any of the methanotrophs tested previously (Fig. 3-1).
The responses to (R)-a-pinene were highly dependent on the type of methanotroph
and MMO expression, with M~ trichosporium OB3b showing decreased rates relative to
20 ppm TCE alone regardless of (R)-a-pinene concentration (Fig. 3-2, A, B) and M~
capsulatus (Bath) and M~ album BG8 showing mostly increased rates (Fig. 3-2, C, D, E).
With the exception ofM~ capsulatus (Bath) under pMMO expression, the highest
observed rates in the presence of the mixture were lower than those observed with (R)-a-
pinene alone. M~ trichosporium OB3b expressing pMMO showed consistently small
decreases in oxygen uptake activity in the presence of the mixture compared to 20 ppm
TCE alone; however, the activity of this strain when expressing sMMO appeared to be
inhibited to a greater extent in the presence of all tested concentrations (2 to 20 ppm) of
(R)-a-pinene in the mixtures (Fig. 3-2, A, B). Regardless of 1VMO expression, M~
capsulatus (Bath) yielded increased normalized oxygen uptake rates in the presence of
the mixture above approximately 20 ppm (R)-a-pinene relative to its observed rate at 20
ppm TCE alone. The greatest rate increase shown by M~ capsulatus (Bath) expressing
slV1VO in the presence of the mixture was observed at 40 ppm (R)-a-pinene. This
maximum rate observed with the mixture was 1.8 times the rate with 20 ppm TCE alone,
suggesting a lessoning of toxicity effects on the cells. The maximum increase with this
strain under plV1VO expression was observed at 30 ppm (R)-a-pinene and was
approximately 3.5 times higher than with 20 ppm TCE alone and 1.5 times higher than
observed at 50 ppm (R)-a-pinene alone. Increase in oxidation potential ofM~ album BG8
was also observed when (R)-a-pinene was in the presence of 20 ppm TCE (Fig. 3-2, E).
At the highest concentration of (R)-a-pinene tested (30 ppm) with this strain, the increase
in normalized oxygen uptake rate was 1.8 times the rate observed with TCE alone.
In conclusion, all of the tested methanotrophs expressing either slVMVO or plV1VO
were capable of oxidizing (R)-a-pinene over a range of environmentally relevant
concentrations. However, toxicity effects of this monoterpene, similar to those shown
with TCE, were observed. When both (R)-a-pinene and TCE were introduced to the
representative methanotrophs, varying responses in the rates--decreases with the Type II
methanotroph and increases with the Types I and X methanotrophs-were observed in
comparison to those observed in the TCE-only experiments. Whether TCE and/or (R)-a-
pinene were oxidized in the mixture is not known, given the indirect measurement
method of oxygen uptake analysis; however, it is suggested here that the total oxidation
potential of methanotrophs is affected, either antagonistically or synergistically, in the
presence of TCE and (R)-a-pinene mixtures. These results emphasize the importance of
not only assessing the concentration levels of both contaminants and monoterpenes and
but also of measuring the oxidation potentials and diversity of rhizosphere methanotrophs
at phytoremediation sites where plants that release large amounts of monoterpenes are
being contemplated for use.
0 20 40 60 0 20 40 60 0 20 40 60 0 20 40 60 0 20 40 60
TCE and (R)-co-pinene concentration (ppm)
Figure 3-1. Normalized rate of oxygen uptake by the representative methanotrophs in the
presence of varying concentrations of TCE (*) and (R)-a-pinene (0). (A),
(B): M~ trichosporium OB3b cultured without and with copper, respectively.
(C), (D): M~ capsulatus (Bath) cultured without and with Cu, respectively.
(E): M~ album BG8 cultured with Cu. Error bars represent the standard
deviation for triplicate samples.
-A -- B -- C- -- D -
0 20 4060 0 20 4060 0 20 400 0~ 20 4060 0 20 4080
(R)-a.-pinene concentration (ppm)
Figure 3-2. Change in the normalized oxygen uptake rate by representative
methanotrophs observed in the presence of 20 ppm TCE at varying
concentrations of (R)-a-pinene. (A), (B): M. trichosporium OB3b cultured
without and with Cu, respectively. (C), (D): M~ capsulatus (Bath) cultured
without and with Cu, respectively. (E): M. album BG8 cultured with Cu.
Error bars represented the standard deviation for triplicate samples.
STABLE ISOTOPE PROBING FOR CHARACTERIZATION OF
IVETHANOTROPHIC BACTERIA INT THE RHIZOSPHERE OF
PHYTORE1VEDIATING PLANT S
Note: Manuscript to be submitted to Biology Letters
Phytoremediation, the use of plants to remove a variety of contaminants from soil
and aqueous environments, has been shown to be more economical and aesthetically
pleasing than traditional remediation methods, such as pump-and-treat approaches
(McCutcheon and Schnoor, 2003). Despite its observed effectiveness in removal of
contaminants, including trichloroethylene (TCE) and tetrachloroethylene (PCE), two
widely distributed chlorinated solvents that cause concern because of their potential
health effects (ATSDR, 2006), phytoremediation is still limited by a lack of
understanding of the primary removal processes involved. In particular, the potential
roles that root-zone (rhizosphere) bacteria can assume in the overall removal of
contaminants is not fully appreciated (Walton and Anderson, 1990; Anderson and
Walton, 1995; Brigmon et al., 1998; Brigmon et al., 1999).
One reason for the lack of specific information on the degradation potentials of
root-zone bacteria is that traditional culture-dependent methods are not capable of
directly assessing the activity and diversity of microorganisms in situ. Furthermore, these
methods provide limited information because of their associated inherent cultivation bias
(Fry, 2004; Smalla, 2004). The development of culture-independent molecular methods
such as stable isotope probing (SIP), has enabled scientists to study in situ conditions
more effectively and, more importantly, to characterize the active microbial populations.
The promising SIP technique relies on the incorporation of a labeled substrate with a less
naturally frequent isotope, into the active microbial community of a sample that later can
be separated from the unlabeled biomass (Radaj ewski et al., 2000).
Recent studies of methanotrophic bacteria have shown successful results using the
SIP approach in environments, such as peat soils, acidic forest soils, cave water, and soda
lake sediments (Morris et al., 2002; Radajewski et al., 2002; Hutchens et al., 2004; Lin et
al., 2004). Methanotrophs. are among the aerobic bacteria that are known to reside in the
rhizosphere of plants and that are capable of oxidizing chlorinated contaminants such as
TCE (Wilson and Wilson, 1985; Hanson and Hanson, 1996; Brigmon et al., 1999;
Doronina et al., 2004; Pilon-Smits, 2005). Because methane serves as the sole source of
carbon and energy for methanotrophs, it is often used as the measure of methanotroph
activity in the environment, and is a natural substrate for SIP testing. Labeled methane
(13C-CH4) has been successfully incorporated into the DNA of growing cells of
methanotrophs. (13C-DNA) and separated from the naturally occurring 12C-DNA by
density gradient centrifugation (Radajewski et al., 2000; Morris et al., 2002; Radajewski
et al., 2002; McDonald et al., 2005). Molecular fingerprinting techniques, such as
denaturing gradient gel electrophoresis (DGGE) with DNA fragments of specific
methanotroph enzymes, such as particulate methane monoxygenase (pMMO), can be
subsequently used to identify and assess the relative abundance of the active populations
(Muyzer et al., 1993).
With the advent of this sophisticated molecular biology method that links identity
with function, development of a protocol using SIP methods that is specific to the
rhizosphere holds promise in better understanding the rhizodegradation process of
phytoremediation systems. This study provides a first-basis in method development and
analysis of the SIP technique for the measurement of potential in situ activity and
diversity of methanotrophic bacteria in the root-zone of trees used for remediation of
TCE from contaminated groundwater and soil.
Materials and Methods
This study was located at a Superfund site, the former LaSalle Electrical Utilities in
LaSalle, IL (USA). The company manufactured capacitors from 1943 to 1982, resulting
in soil and groundwater contamination of mostly polychlorinated biphenyls and the
chlorinated solvents, TCE and PCE. Currently, in the final stages of the cleanup process
at the site, two phytoremediation plots have been implemented to enhance chlorinated
solvent removal (Lange, 2004). The first plot (0.25 ha), contaminated with TCE (0-254
ppb), was installed in September 2002 (labeled as "TCE Site"). Poplar (18 clones) and
willow (24 clones) genotypes were planted by lowering 1.8 m rooted cuttings to the
bottom of boreholes (0.6 m diameter) lined with high-density polyethylene pipe and filled
with an equal mix of sand, soil, bark, and peat (pH 7.8). The second phytoremediation
plot (0.21 ha), contaminated with PCE (0-838 ppb), was established in March 2002
(labeled as "PCE Site"). At this plot, poplar trees were planted directly into the improved
soil (pH 7.3) with mulch composed of tree chips on the top 0.5 m of the soil surface.
All rhizosphere soil samples were collected using a small diameter (1.9 cm) hand
soil auger to minimize disturbance in the pots. Samples were taken at three time periods,
July 2003, July 2004, and November 2004, in order to compare summer and fall
conditions and time effects on the activity and population diversity of methanotrophs.
Each sampling period, mid-summer (July) and early fall (November), fell in the "wet
season" of April to December when 642 mm of precipitation were collected at the site
during 2004. The average daily air and soil temperatures were 210C and 110C,
respectively, in the 2004 summer sampling, and 200C and 90C, respectively, in the 2004
fall sampling. Groundwater levels below the planted plots fluctuated in 2004 from 2.1 to
3.1 m and 1.8 to 3.3 m from the soil surface at the TCE and PCE Site, respectively.
Root growth of the trees at the TCE and PCE Site was observed to extend from the
surface to 90-120 cm below surface. A composite sample, in regions of high contaminant
concentration, was taken from two opposite locations around the tree base at a depth of
30-60 cm. This soil layer was chosen as a potential zone of intermediacy rhizosphere
activity between surface and deeper soil layers. At the TCE Site, samples were also
removed from non-planted pots in the contaminated area to serve as a control when
compare to the planted pots. Also different tree clones, showing the greatest vigor, were
sampled at the TCE Site. They were one poplar clone 145/51 (Populus deltoides x P.
nigra; origin, North America x Europe) and three willow clones, SX61 (Salix
sachalinensis; origin, Japan; exotic), S365 (S. discolor 18; origin, University of Toronto),
and 94014 (S. purpurea; origin, State University of New York; exotic).
While it is well known that methanotrophs. are not capable of oxidizing PCE,
samples were removed from the PCE Site to serve as a mean of comparing methanotroph
activity and diversity with the TCE Site samples. One poplar clone, 145/51, was sampled
at the PCE Site, along with a non-planted sample removed from outside the plot in an
uncontaminated region that was eventually converted to an irrigated, fertilized soccer
Hield after the first sampling in July 2003.
To prevent cross-contamination, the sampling auger was washed with sterile water,
rinsed with 95% ethanol, and washed again before each sample was taken. Samples were
immediately placed in sterile bags (Nasco Whirl-Pak, Fort Atkinson, WI, USA), placed
on ice, and transported to the UF laboratory where they were subsequently stored at 4oC
until testing. In the laboratory, samples were gently homogenized using a sterile spatula,
and Eine roots of less than 2 mm diameter were separated from the soil for separate
Stable Isotope Probing (SIP) Soil Microcosms
Experimental conditions. In order to assess activity and diversity of the active
methanotroph populations in poplar and willow tree rhizospheres, during TCE
remediation, soil microcosms were prepared from samples collected over time.
Microcosms consisted of 10 g wet soil (plant material removed) normalized to 16% water
content with sterile water. This water content represented approximately 40% of the Hield
capacity of the TCE and PCE Site soils, where the greatest extent of CH4 Oxidation was
observed in preliminary experiments, as previously described by Reay et al. (2001). The
"wetted" soil was placed in sterile 160 ml serum vials, which were subsequently sealed
with gray butyl rubber stoppers and crimp tops. Ten ml (0.4 mmol; ~7%, v/v) of filter-
sterilized 13CH4 (99.9%; Isotec, Miamisburg, OH, USA) or 12CH4 (99.9%; Airco-BOC,
Murray Hill, NJ, USA), used in preliminary experiments to optimize conditions and
assess any effects of the labeled substrate by comparing to the 13CH4 miCTOCOSms rates,
was then added as previously described (Morris et al., 2002; Radajewski et al., 2002), and
each vial was wrapped with aluminum foil for subsequent incubation in the dark at room
temperature (~250C). Headspace CH4 depletion was monitored every 2 to 5 days by
removal of 25 ul of CH4 with a gastight syringe and analysis using a gas chromatograph
(Model HP5809A GC/TCD; Hewlett Packard, PaloAlto, CA, USA) equipped with a GS-
Carbon plot column (Agilent Technology, PaloAlto, CA, USA). The gas chromatograph
was maintained at a head pressure of 5 psi and programmed in a 4 min run with
temperatures of 25, 120 and 2000C in the oven, injector and detector, respectively.
When more than 90% of the CH4 WAS COnSumed, vials were opened, gently flushed
with fi1ter-sterilized air for 5 s to remove any accumulated 13CO2 and to maintain aerobic
conditions, resealed, and the same initial amount of 13CH4 added. The procedure was
repeated Hyve times until a total of 2.0 mmol of CH4 WAS COnSumed (Radaj ewski et al.,
2002). A positive control with pure methanotroph, M~ethylocystis trichosporium OB3b,
and three negative controls with no CH4 added, with twice-autoclaved (killed) soil, and
with 20% (v/v) each of CH4 and acetylene (a known inhibitor of methane
monooxygenase, Prior and Dalton (1985)) in the headspace, were also included.
Additionally, some of the microcosms were set in replicates to assure reproducible
results. Initial CH4 depletion rates were calculated from data taken during incubation
after the first CH4 addition by linear regression analysis of the consumption curve.
DNA extraction and ultracentrifugation. The content of the microcosms (10 g
soil) was processed using a PowerMax Soil DNA Extraction Kit (Mo-Bio, Carlsbad, CA,
USA). DNA extracts were resolved by CsCl density gradient centrifugation. Briefly, 1 g
mll of CsCl (Fisher Scientifie, Pittsburgh, PA, USA) was dissolved in the DNA solution,
and 100 El1 of ethidium bromide (10 mg ml l; Bio-Rad, Hercules, CA, USA) was added
before loading the solution into 5.1 ml quick-seal polyallomer ultracentrifuge tubes
(Beckman Coulter, Fullerton, CA, USA). Ultracentrifugation was performed using a
VTi65 vertical rotor in a Model L8-80 ultracentrifuge (Beckman Instruments, Fullerton,
CA, USA) at 265,000 x g for 16 h at 200C. After ultracentrifugation, fractions were
visualized with UV light at 365 nm (Sambrook et al., 1989; Radajewski et al., 2002).
Three DNA bands were generally observed and collected: (1) a light-DNA upper band
(12C-DNA); (2) a middle band, smear of 12C- and 13C-DNA; and (3) a heavy-DNA lower
band (13C-DNA). DNA fractions were collected and purified as described by Sambrook
et al. (1989). Ethidium bromide was extracted from the DNA with 1-butanol (Fisher
Scientific, Pittsburgh, PA, USA) saturated with water. Following fiye extractions the
DNA solution was diluted in water and precipitated with ethanol overnight at -200C and
dissolved in 100 ul TE buffer (Sambrook et al., 1989). A second ultracentrifugation step
was not necessary after confirming that the protocol of DNA band extraction was exact.
Re-runs verified the presence of only one distinct band in the new columns.
Polymerase chain reaction (PCR) amplification. The purified DNA fractions
(12C- and 13C-DNA) were used as a template for PCR analysis. The phylogenetic
analysis was performed with the functional pmoA gene, targeted using the primer set
Al89f (5' -GGNGACTGGGACTTCTGG-3 ') and mb661 (5'-
CCGGMGCAACGTCYTTACC-3 ') (Integrated DNA Technologies, Coralville, IA,
USA), specific to the pMMO active site (Costello and Lidstrom, 1999). A GC-clamp (5'-
cccccccccccccgccccccgccccccgcccccgccgccc-3') was attached to the Al 89f primer as
described by Henckel et al. (1999).
PCR amplification was performed according to the procedure described by Knief et
al. (2003). PCR reactions consisted of the MasterAmp 2X PCR premixture F (Epicentre
Technologies, Madison, WI, USA) containing 100 mM Tris-HCI (pH 8.3), 100 mM KC1,
400 ELM each of dNTP, 3-7 mM MgCl2, and the enhancer betaine (0-8 X), combined with
0.5 ELM each primer, lU Taq polymerase, and sterile water to a total reaction volume of
50 El1 (Knief et al., 2003). All reactions were assembled on ice, and the cooled tubes
were placed in a preheated (940C) thermal block for PCR initiation (Henckel et al.,
The PCR protocol consisted of a touchdown program using a thermocycler
(Mastercycler Personal 5332; Eppendorf, Westbury, NY, USA) with the following
parameters: initial denaturation of 5 min at 940C, followed by 35 cycles of 1 min at 940C
for denaturation, 1.5 min at 62 to 550C in -0.50C increments for annealing, 1 min at 720C
for elongation, with a final extension step of 7 min at 720C (Knief et al., 2003). PCR
product size (540 bp) was examined by horizontal agarose electrophoresis. PCR positive
controls included representatives of all methanotrophs types (type X M~ethylococcus
capsulatus (Bath) (ATCC 33009), type II strains, M~ethylosinus trichosporium OB3b
(ATCC 35070), Strain CSC1, M~ethylocystis echinoides (IMET 10491), M~ethylocystis
parvus OBBP (NCIMB 11129), and type I2~ethylomicrobium album BG8 (ATCC
Denaturing Gradient Gel Electrophoresis Analysis (DGGE), Sequencing, and
DGGE. PCR products were separated by DGGE in the DCode System (Bio-Rad,
Hercules, CA, USA) as described by Henckel et al. (1999). Briefly, 1 mm thick 6.5%
(w/v) polyacrilamide gels (37.5:1 acrylamide-bisacrylamide) (Fisher Scientific,
Pittsburgh, PA, USA) were prepared and electrophoresed in lX TAE buffer at 610C and
180 V for 5 h in a 35-65% linear denaturant gradient (65% is 4.5 M urea and 26% (v/v)
deionized formamide). Gels were loaded with 25-45 El1 of PCR product, according to
band intensity in agarose gels, and '/ volume of loading buffer. Gels were stained with
ethidium bromide according to the manufacture' s instructions (Bio-Rad, Hercules, CA,
USA), visualized at 312 nm on a UV transilluminator (Model 88A, Fisher Scientifie,
Pittsburgh, PA, USA), and photo-documented with the system DigiDoc-IT TM (Daigger,
Vernon Hill, IL, USA) using the Doc-It software v. 2.4 (UVP, Upland, CA, USA).
DGGE bands were excised from the middle part of the band with a sterile scalpel and the
DNA eluted according to the protocol described by Chory and Pollard (1999). The eluted
DNA was reamplified and reanalyzed on DGGE to verify sample purity. Band
reamplifieation was performed by modifying the PCR protocol to 30 cycles of 30 s at
940C for denaturation, 45 s at 660C for annealing (to avoid sequence ambiguity as
reported by Dunfield et al. (2002)) and 30 s at 720C for elongation, with the same initial
and Einal steps. Several bands with the same mobility were excised from different lanes
to check for sequence identity.
Sequencing. Reamplified PCR products of excised DGGE bands were purified
with a PCR purification kit (Mo-Bio, Carlsbad, CA, USA) before sequencing. PCR
product concentration and purity was determined by UV absorption spectrophotometry
(1:20 dilution). PCR products were sequenced by the Interdisciplinary Center for
Biotechnology Research (ICBR), University of Florida (Gainesville, Florida, USA).
Phylogenetic analysis. Sequences were compared in the National Center for
Biotechnology Information (NCBI) database using BLAST (Altschul et al., 1990).
Related sequences identified in BLAST, as well as sequences of extant methanotrophs,
were aligned and adjusted manually with CLUSTALX v. 1.8 (Thompson et al., 1997).
Phylogenetic trees were generated by the neighbor j oining (NJ) method with
CLUSTALX and displayed in TreeView v. 1.6.6 (Page, 1996). Nucleotide accession
numbers of all obtained sequences were placed in the GenBank for future access
CH4 depletion rates observed in the SIP microcosms were analyzed by comparing
the initial slopes of the linear regression curves fitted to the consumption curve during
incubation after the first CH4 addition. When a set of samples showed no significant
differences in rates, an average depletion rate (common regression coefficient) was
calculated as an estimate of the CH4 depletion rate underlying all rates of a particular set
of samples (for example, rates among the same plant type in each sampling period).
Additionally, differences among sample means and between samples and the control
were analyzed by Tukey's and Dunnett' s test, respectively (Zar, 1984). SAS software v.
7 (SAS Institute, Cary, NC, USA) was used for all the analysis.
SIP Protocol Implementation
The SIP technique was successfully applied to the rhizosphere soils. The rates
observed in the 13CH4 and 12CH4 miCTOCOSms were comparable, and variability among
the replicates was low. The M. trichosporium OB3b control was effectively labeled by
the 13CH4 and no 13CH4 COnSumption was observed in the negative controls.
DNA extracts from the microcosms were effectively separated by CsCl density
gradients (Fig. 4-1A). When a smear was present between the unlabeled (12C-DNA) and
labeled (13C-DNA) fractions, it was collected as a 12-13C-DNA combined fraction and was
not included in the study. Correct recovery of these fractions was verified by a second
ultracentrifugation under the same protocol, even though band position varied because of
changes in the density of the new solution (Fig. 4-1, B-C). The extracted 13C-DNA
fractions produced pmoA gene fragments of the expected size (540 bp). The
methanotroph positive control cultures revealed multiple DGGE bands (Fig. Al,
Appendix), in keeping with earlier reports of the high probability of encountering
multiple copies of the pmoA gene in methanotrophs (Semrau et al., 1995; Dunfield et al.,
2002). Because DGGE band purification was difficult in most samples yielding
ambiguous positions after sequencing, the annealing temperature was increased from 62
to 660C and only the reverse primer was used for sequencing (Dunfield et al., 2002).
Furthermore, 16S rDNA-DGGE profies revealed complex band patterns and smears that
were difficult to examine (data not shown).
In general, pmoA-DGGE profies of the 13C-DNA fractions of the different
rhizosphere soil microcosms did not differ greatly among sites, plant type or sampling
period. Consequently, it was useful to set a reference profie for the analysis (Fig. 4-3A,
lane 1). Reference bands 1, 3, 9, 10 and 11 were not possible to sequence.
Methanotroph Activity and Composition in the TCE Site
13CH4 added in Hyve additions to a total of approximately 2.0 mmol, was consumed
within 31-37 days in all TCE Site soil microcosms. Initial CH4 depletion rates (Fig. 4-
2A), calculated as a measure of the oxidative potential at the time of sampling, showed
no significant differences within plant type over the 16-month sampling period. A
comparison of average rates per plant type, which represents plant type's overall activity
throughout the study, shows also no significant difference in CH4 depletion rates among
plant types and with the non-planted sample (P<0.05). The overall average CH4
depletion rate at the TCE Site was 0. 11 Epmol h-l g-l dry weight soil (+ 0.01, SE).
As shown in Fig. 4-3, TCE Site (panel A to C) and PCE Site (panel D) pnzoA-
DGGE profies were mostly described by the same group of bands numbered from 1 to
11 in the reference profie (Fig. 4-3A, lane 1). Concentric circles with letters denoted
bands different from the reference profie. Profies were labeled starting with the plot
location (TCE or PCE) followed by the tree type (poplar or willow), soil compartment
analyzed (rhizosphere, rhizoplane, or non-planted soil), and sampling period. Willow
clones SX61 and 94019 were not sampled in the November 2004 sampling period,
consequently, they only showed two profies each (Fig. 4-3C, lanes 1-4). The pnoA-
amplified sequences from the 13C-DNA fractions of the TCE Site microcosms revealed
DGGE profies composed by 2 to 11 bands (Fig. 4-3, B-C). These profies were mainly
described by three groups of highly similar sequences according to band position and
BLAST alignments, indicating that some bands may represent copies of multiple pnzoA
genes. However, some profies that exhibited these groupings did not reveal all of the
The TCE Site DGGE profies throughout the study did not vary to a great extent
among plant type (Fig. 4-3, B-C), but they were distinct from the non-planted soil (Fig. 4-
3B, lane 4 to 6). Rhizosphere DGGE profies from poplar (Fig. 4-3B, lane 1-2) and all
willow clones (Fig. 4-3, panel B, lanes 7-8, and, panel C, lanes 1 to 4) collected in July
2003 and 2004 revealed the same community of organisms. These profiles were
composed of two groupings of bands represented by reference bands 4 and 6 to 8, and
bands 2 and 5 (Fig. 4-3A, lane 1). Within each grouping, bands shared 100% sequence
similarity and aligned in BLAST (>99% similarity) with two clones of uncultured
M~ethylocaldunt sp. isolated from a landfill cover soil. The phylogenetic tree clustered
these sequences as the described groupings (Fig. 4-4, Group 1 and 2) and within the
M~ethylocaldunt branch (type X methanotroph). These groupings are closely related to
the cultured organism, M~ gracile. Also, a third group of bands, designated by reference
bands 9 to 11 (Fig. 4-3A, lane 1), was detected in all profiles. However, because
reamplifieation of this group was not possible, and since they were the only bands present
in the July 2003 TCE Site non-planted profie (Fig. 4-3B, lane 4), it is possible they
represent another set of similar pnoA genes.
From the planted microcosms at the TCE Site, only the polar and willow clone
94014 exhibited changes in their methanotroph community throughout the study. The
poplar tree, in the November 2004 sampling (Fig. 4-3B, lane 3), showed less than half of
the bands in the July samplings (Fig. 4-3B, lane 1-2). It only revealed three bands, two at
very low intensity and the same maj or band of previous samplings (reference band 5).
Willow clone 94014, in the July 2004 sampling (Fig. 4-3C, lane 4), did not exhibit this
maj or band and showed an extra band of an uncultured methanotroph that closely related
(88% similarity) to another type X methanotroph, M~ethylococcus capsulatus (Fig. 4-4).
The TCE Site non-planted microcosms, over the three sampling times, exhibited
variable profies at very low intensity (Fig. 4-3B, lane 4 to 6). In the July 2003 sampling,
only the 9 to 11 grouping was retrieved. However, in the July and November 2004
samplings, the 9 to 11 grouping was observed along with some of the bands that
described the M~ethylocaldunt clones found in the planted samples and uncultured bacteria
from rice Hields and upland soils (>87% similarity). The uncultured bacteria (Fig. 4-3B,
lane 5-6) were placed within the M~ethylocaldunt and M~ethylococcus branch (Fig. 4-4),