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Applied and Environmental Microbiology, February 1999, p. 534-539, Vol. 65, No. 2
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Influence of Light Intensity on Methanotrophic
Bacterial Activity in Petit Saut Reservoir, French Guiana
J. F.
Dumestre,1,*
J.
Guézennec,2
C.
Galy-Lacaux,3
R.
Delmas,3
S.
Richard,4 and
L.
Labroue1
Centre d'Ecologie des Systèmes
Aquatiques Continentaux, UMR CNRS/UPS 5576, Université Paul
Sabatier, 31062 Toulouse Cedex,1
Laboratoire de Biotechnologie des Microorganismes
Hydrothermaux, DRV/VP/BMH, Ifremer Centre de Brest, 29280 Plouzané,2 and
Laboratoire d'
Aérologie, UMR CNRS/UPS 5560, OMP, 31400 Toulouse
Cedex,3 France, and
Laboratoire
Environnement de Petit Saut, Hydreco, Aménagement de Petit
Saut, 97388 Kourou Cedex, French Guiana4
Received 7 April 1998/Accepted 17 November 1998
 |
ABSTRACT |
One year after impoundment in January 1994, methanotrophic bacteria
in Petit Saut Reservoir (French Guiana) were active at the oxic-anoxic
interface. This activity was revealed by the sudden extinction of
diffusive methane emission (600 metric tons of CH4 · day
1 for the whole lake surface area, i.e., 360 km2). Lifting of inhibition was suspected. After reviewing
the potential inhibitors of this physiological guild (O2,
NH4+, sulfides) and considering the
similarities with nitrifiers, we suggest that sunlight influenced the
methanotrophic bacteria. On the basis of phospholipid analysis, only a
type II methanotrophic community was identified in the lake. Both
growth and methanotrophic activity of an enriched culture, obtained in
the laboratory, were largely inhibited by illumination over 150 microeinsteins · m2 · s1.
These results were confirmed on a pure culture of Methylosinus trichosporium OB3B. In situ conditions showed that water
transparency was quite stable in 1994 and 1995 and that the oxycline
moved steadily deeper until January 1995. Considering the mean
illumination profile during this period, we showed that removal of
methanotrophic growth inhibition could only occur below a 2-m depth.
The oxycline reached this level in October 1994, allowing
methanotrophic bacteria to develop and to consume the entire methane
emission 4 months later.
| |
INTRODUCTION |
Within the three past decades,
methanotrophic bacterial guilds have been studied in the laboratory for
industrial applications, and the major factors controlling their
activity have been examined in detail in pure and mixed cultures
(8, 20, 26, 32). These experiments contributed to the
understanding of how methanotrophs operate in natural environments,
such as termite mounds, lake water, sediments, and soils (7, 28,
35, 39). Most of the data set obtained in tropical or equatorial
lakes concerned only measurements of methane emissions or
concentrations (21, 22, 36, 38). Only a few works reported
experimental determination of bacterial methane oxidation in such
environments (19, 31). So we chose to develop such an
approach for a new equatorial reservoir.
Petit Saut Dam was built on the Sinnamary River, by Electricité
de France Company, in order to sustain the economic development of
French Guiana. More than 360 km2 of equatorial rain forest
was flooded. The anaerobic degradation of submerged organic matter
rapidly induced strong emission of methane into the atmosphere
(12). Because of the huge emission of reduced elements and
the very little mixing by the wind, the oxygenated layer of the lake
remained thin during the two first years.
The aim of this study was to determine the role played by
methanotrophic bacteria in regulating methane emission and the factors controlling their growth in this equatorial medium.
| |
MATERIALS AND METHODS |
In situ measurements.
Since the dam has been closed, water
analyses have been regularly performed at a floating station (Barge
Petit Saut [BPS]) located 300 m upstream of the dam (35-m
maximum depth). Sampling was performed and physical-chemical profiles
were determined by means of a peristaltic pump fitted with silicone
rubber tubing (7.9-mm inside diameter; Masterflex Ltd.). A septum
located just before the pump head allowed the water to be sampled
without gas stripping. The water was pumped at 800 ml · min1, and the line was flushed for 3 min before any
samples were taken.
Methane emission fluxes were measured at the surface of the lake in a
floating stainless steel chamber (50 by 50 by 18 cm). The concentration
of methane gas was determined in the chamber by gas chromatography
analyses at time zero and 15, 30, and 60 min later. Before the samples
were taken, the atmosphere in the chamber was mixed by connecting the
chamber to the peristaltic pump in a closed circuit. The linear
regression determined from the experimental points allowed the methane
flux to be calculated. During each trial, fluxes were measured at four
different sites (two were in the flooded forest, one was the BPS site,
and one was above the former river bed) distributed over the whole
lake, in order to evaluate an average total methane emission, expressed as moles of CH4 per square meter per day. The method used
did, however, lead to an underestimation of the methane flux since the
chamber reduced wind effects on surface roughness and transfer velocity.
Methane concentrations were determined by the headspace method. A 20-ml
water sample was injected into a 57-ml glass flask which had been
sealed with a Teflon septum and previously emptied with a vacuum pump.
The methane concentration in the water was calculated after analyzing
the headspace with a Hewlett Packard HP 5890 A gas chromatograph fitted
with a flame ionization detector and a Poraplot Q semicapillary column.
Details concerning the analysis conditions are described elsewhere
(11).
The concentration of dissolved O
2 was determined in situ
with an OXY 196 Wissenschaftlich Technische Werkstatten (Weilheim,
Germany)-specific probe. Ammonium ion was analyzed immediately
after
sampling by spectrophotometry with the Nessler reagent.
Total sulfides
were determined by the colorimetric method of Cline
(
5) in
samples preserved with zinc
acetate.
Illumination in the water column was measured with a LI-COR 1000 luxmeter (wavelengths integrated from 400 to 700
nm).
Transparency was assessed with a 33-cm-diameter white disk (Secchi
disk). The average depth at which the disk disappeared
from view was
noted after three
readings.
The actual methane consumption rate was measured to document the
methanotrophic activity in the water column. For each depth,
four 26-ml
glass flasks, closed with a Teflon septum, were filled
with lake water.
Twenty-milliliter volumes of the water contained
in two flasks were
rapidly sampled to measure the initial methane
concentration as
described above. The two other glass flasks were
incubated under in
situ conditions for 1 h and the final methane
concentration was
measured. The difference between the two average
measurements gives an
estimation of the methane consumption rate
in milligrams of
CH
4 per liter per hour. This determination was
repeated
three times at the BPS station in December 1995. The
values were very
similar over the whole lake. No abiotic methane
disappearance was
observed in samples preserved with thimerosal
(BDH Chemicals Ltd.,
Poole, England), a chemical compound blocking
bacterial enzyme activity
in the same way mercuric chloride
does.
Laboratory experiments.
Water samples were taken at the BPS
site from around the oxycline in September 1995 for enrichment of their
methanotrophic bacterial population. Ten-milliliter aliquots of water
complemented with 10% glycerol were frozen at 
18°C until required
for enrichment. The samples were thawed and centrifuged three times to
eliminate the glycerol carbon source. The resulting pellet was
resuspended in 20 ml of nitrate mineral salt medium (41),
buffered at pH 6.8, and incubated in a 250-ml glass Erlenmeyer flask at
30°C in darkness, with magnetic stirring (130 rpm). The headspace of the flask was filled with a methane-air atmosphere (50/50, vol/vol) complemented with 4 ml of CO2, to enhance the start of
bacterial growth (17). The atmosphere was replaced every day
and the bacteria were subcultured to obtain a large active bacterial
biomass. A freeze-dried pure culture of Methylosinus
trichosporium OB3B (purchased at the National Collections of
Industrial and Marine Bacteria Ltd., Aberdeen, Scotland) was also
resuscitated by the same method in order to perform control experiments.
Bacterial growth was monitored by spectrophotometry (Anthelie; Secomam,
Domont, France) in a 1-cm cell (optical density [OD]
measured at 540
nm).
Replicates of the bacterial enrichment were placed in a 30°C
regulated incubator with different illumination intensities (from
10 to
1,100 microeinsteins · m
2 · s
1). The white light was delivered for 12 h a day
(the day length
in French Guiana) with two 400-W lamps (Phytoclaude;
Claude, Paris,
France). The emission spectrum is given in Fig.
1. The incident
light was integrated from
400 to 700 nm by a LI-COR 189 luxmeter.
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FIG. 1.
Emission spectrum of the Phytoclaude lamp used to
demonstrate the inhibitory effect of light on methanotrophic bacterial
growth and activity.
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|
The total methane concentration was monitored during the incubations by
analyzing the flask's headspace with a gas chromatograph
fitted as
described in the previous
paragraph.
Phospholipid fatty acid (PLFA) analysis was performed on the enriched
culture and on water taken directly from the lake, which
had been
freeze-dried after filtration on a Nuclepore polycarbonate
membrane
(0.22-µm pores, 47-mm diameter). Briefly, total lipids
were extracted
by a modified Bligh and Dyer extraction procedure
(
40). The
extracted lipids were fractionated by silica column
chromatography with
dichloromethane, acetone, and methanol as
eluants. The methanol
fraction (polar lipids) was dried under
a stream of nitrogen and
submitted to acid methanolysis. Nonadecanoic
acid was added before
methanolysis as an internal standard. The
resulting fatty acid methyl
esters were resuspended in 50 µl of
dichloromethane and analyzed by
gas chromatography. The double
bond positions in the monounsaturated
fatty acids were determined
by dimethyl disulfide derivatization
(
25). The double bond positions
in the polyunsaturated fatty
acids were determined after 4,4-dimethyloxazoline
derivatization
(
10). Detailed procedures are reported elsewhere
(
16).
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RESULTS AND DISCUSSION |
In situ measurements.
Since the beginning of reservoir
impounding, the anaerobic degradation of submerged organic matter
caused the production of large quantities of reduced compounds. The
bottom methane concentration increased until the filling was finished,
i.e., in July 1995 (Fig. 2). After that
time the bottom methane concentration evolved in a cyclic manner, with
a maximum during the dry season. This phenomenon could be due to a
variation in methane production or dilution by the huge amounts of rain
in the wet season.
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FIG. 2.
Variation of total methane emission and methane
concentrations in the water column (at depths of 0, 1, and 2 m and
at the bottom), since the beginning of filling (day zero), in January
1994. d, day.
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|
Average diffusive methane emission reached a maximum of 0.103 mol of
CH
4 · m
2 · d
1
(600 metric tons of CH
4 · day
1 for the
whole lake surface area) in February 1995 (Fig.
2) and
then decreased
suddenly, falling to a level close to zero. This
phenomenon could be
attributed not to a drop in production, since
the bottom concentration
was still increasing, but rather to the
activation of methanotrophic
populations present in the upper
layers of the water column. The
biological methane-oxidizing activity
was directly demonstrated by
measuring the actual methane consumption
rate at the BPS site (Fig.
3). This vertical profile was obtained
in
December 1995 and showed that the methanotrophic activity was
localized
at the oxycline level, where methane and oxygen were
both available in
low but sufficient quantities. Sharp stratifications
of this bacterial
activity have already been shown to occur in
aquatic environments
(
19,
30) and in sediments (
6). The
methanotrophic
activity was assumed to act in a 1-m-thick stratum
with a maximum
activity of 0.275 mg of CH
4 · liter
1 · h
1 in the middle of the
stratum (4.5 m from the surface) and over
a surface area of about 250 km
2, at the oxycline (
34). Considering that the
methanotrophic
activity was equally distributed across the lake, we
obtained
a total methane consumption of 825 metric tons of
CH
4 · day
1, which could explain the
sudden drop to zero of the methane emission
recorded in February 1995.
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FIG. 3.
Dissolved oxygen and methane profiles and methane
consumption measured at the BPS site in December 1995. Values are in
tenths.
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|
Hypothesis of methanotrophic bacterial inhibition.
This sharp
decrease in methane emission occurred more than 1 year after the
beginning of filling, suggesting a long lag phase or inhibition by
environmental factors. During preliminary measures realized in 1994, Gosse and Grégoire (13) demonstrated downstream of the
dam a strong oxygen depletion due to methane oxidation. This phenomenon
occurred only when bottom water masses (anaerobic with methane) and
upper water masses (aerobic with methanotrophic bacteria) were
evacuated simultaneously. When bottom water was discharged alone into
the Sinnamary River, oxygen consumption was reduced. So, we
hypothesized that methanotrophic bacteria were present and inhibited in
the upper water layers of the lake during the first year of filling.
A few inhibitory factors have been well documented in the literature.
In Petit Saut Lake, a marked gradient in oxygen concentrations
has been
observed a few centimeters or meters below the lake surface
since the
beginning of the filling phase (Fig.
3). Oxygen concentrations
were
sufficiently low to enable methanotroph development. So,
the hypothesis
of inhibition by excessive concentrations of dissolved
oxygen was
improbable (
29).
The ammonium concentrations observed around the oxycline, at the BPS
site (Table
1), were very low compared
with the reported
inhibitory concentrations, which range from 1,000 to
10000 µmol
of NH
4+ · liter
1 (
1,
4). So,
NH
4+ could only have had only a trivial
inhibitory effect in Petit
Saut Lake in 1994.
Great amounts of sulfides were produced, and a maximum of 188 µg of
sulfides S · liter
1 was measured at the bottom of
the water column in May 1995 (Table
1). However, the sulfides could not
attain the oxycline level
in sufficiently high quantities, because they
were totally consumed
by phototrophic sulfur-oxidizing bacteria,
located in the anaerobic
zone, 1 or 2 m below the oxycline
(
9). So, the inhibitory effect
of sulfur-containing
compounds on methanotroph growth (
1) can
be also
excluded.
Laboratory experiments.
Methanotrophic bacteria are
genetically related to nitrifying bacteria (18). This second
physiological guild has an ammonia monooxygenase (AMO) which is
inactivated by light (15, 33), specifically in the near-UV
region (300 to 375 nm) and in the blue region of the spectrum (400 to
475 nm). AMO and methane monooxygenase (MMO) are closely related by
their substrate specificities, the structures of their active sites,
and their sensitivities to inhibitors (1). As a consequence,
we suspected light as a possible environmental factor able to influence
methanotrophic guilds in the upper layers of the water column. So, the
effect of light on methanotrophic growth was tested in the laboratory
on enriched samples.
Methane-oxidizing bacteria contain unusual PLFAs in their membranes
(
2,
14,
23,
24,
37). On the basis of the PLFA
analyses and
the presence of 18:1
8 acid, a biomarker for methanotrophs,
only type
II was present in our enrichments (Table
2). Moreover,
it seemed to be widespread
in the whole lake and in the Sinnamary
River.
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TABLE 2.
PLFA profiles determined on methanotrophic cultures and
on a water sample from around the BPS oxycline in December 1996.
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|
Our mixed methane-oxidizing bacterial culture was capable of stable and
predictable growth in batch culture. It exhibited
a fatty acid profile
with 18:1
8 and 18:1
7 acids accounting for
57.6 and 19.63% of the
total PLFAs, respectively (Table
2). This
profile is very close to
those of
Methylosinus or
Methylocystis strains
previously described (
3), with 18:1
8 acid (from 52.9
to
73.6%) and 18:1
7 acid (from 14.8 to 37.7%) as major fatty
acids.
However, we can assume that the exact composition of the
bacterial
community remained unknown and that our enrichment conditions
probably
gave not a pure culture but a mixed culture composed
essentially of
methanotrophic
bacteria.
During our enrichment procedure and for every assay submitted to
illumination, the exponential growth phase stopped at the
end of the
first day of culture. So, we calculated for that time
the percent
inhibition versus the bacterial growth measured in
the reference
culture, after subtracting the OD value measured
at the initial time.
Figure
4 shows the results of an
experiment
run with an illumination of 168 microeinsteins · m
2 · s
1. The change in the methane
quantity (in water-air) in the different
culture flasks clearly showed
that the growth inhibition was related
to a strong decrease of actual
methane consumption. We did not
observe any abiotic methane oxidation
with noninoculated flasks
submitted to illumination. The experiments
were performed with
two separately enriched cultures showing almost
identical PLFA
compositions. Considering all the data obtained for both
enrichments,
the percent inhibition increased rapidly to reach high
values
(over 90% inhibition, taking into account the standard error)
from an illumination about 150 microeinsteins · m
2 · s
1. This inhibition profile was
confirmed with experiments using
a pure
M. trichosporium
OB3B culture that exhibited about the
same sensitivity to light (Fig.
5).
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FIG. 4.
Bacterial growth determined by OD at 540 nm and by total
methane measurements in the incubated flasks. Replicates were incubated
in darkness or under illumination at about 168 microeinsteins · m2 · s1 for 12 h a day. Abiotic
experiments were run without an inoculum. Percent growth inhibition was
calculated after 1 day of incubation. Q, quantity.
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FIG. 5.
Percent growth inhibition versus illumination delivered
during bacterial growth. Experiments were run on two identical enriched
cultures (A and B) obtained from water sampled at the oxycline in
September 1995 and on a pure culture of M. trichosporium
OB3B. µE, microeinsteins.
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|
Relation between laboratory experiments and environmental
conditions.
So, under our experimental conditions, light inhibited
the growth and the activity of the type II methanotrophic bacteria sampled in Petit Saut Lake. In order to relate this observation to
environmental data, we monitored the oxycline level (where methanotrophic activity is located) and the Secchi disk depth since the
beginning of reservoir filling. The average Secchi disk depth was about
1.5 m during the first 2 years of filling, revealing quite steady
illumination of the water column during this period (Fig.
6). Small variations corresponding to
periods of low sunshine during the rainy season were observed. On the
other hand, the oxycline level sank deeper until January 1995. In
consequence, the illumination that would have been measured at the
oxycline decreased regularly. If we consider an average illumination
profile in the water column obtained at the end of 1994 (Table
3), and the levels of inhibition versus
illumination previously measured, we can see that the percent
inhibition significantly dropped only for depths below 2 m (Table
3).
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FIG. 6.
Variation of the total methane emission, the Secchi disk
depth, and the oxycline depth in Petit Saut reservoir since the
beginning of the filling in January 1994. d, day.
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The average depth of the oxycline fell below 2 m in October 1994 (Fig.
6). Considering our laboratory results, the inhibition
of
methanotrophic guild activity could have been largely lifted
during
this period. The extinction of methane emissions occurred
only 4 months
later, a duration in agreement with the well-known
low growth rate of
methanotrophs in natural environments. Since
this time, we have not
detected any diffusive methane emission
from the lake surface into the
atmosphere. Currently, methane
production at the bottom is still high,
but most of the methane
is evacuated through the hydroelectric plant
(
12) or rises as
bubbles in the shallow flooded
forest.
In conclusion, this work allowed us to formulate a plausible
explanation for the sudden decrease in diffusive methane emissions
from
Petit Saut Lake in French Guiana. Environmental observations
were
closely paralleled by laboratory experiments which showed
that light
intensity has a serious inhibitory effect on the growth
of the
methanotrophic guild present in the water column around
the oxycline
and also on pure methanotroph cultures. So, the light
intensity in the
upper water column is thought to have seriously
inhibited the
methanotrophic activity during the year 1994. However,
other unknown
inhibitory factors might have been active during
this first year of
filling. Our experiments allowed us to conclude
that the lifting of
sunlight inhibition on this physiological
guild occurred about 4 months
before the methanotrophs had consumed
large amounts of methane.
Unfortunately, this study does not explain
the actual mechanism by
which light inhibits MMO. However, there
is a great similarity of
structure with the AMO of nitrifying
bacteria, which is also
photosensitive. Shears and Wood (
33)
speculated on the
existence of a particular configuration of the
active site of AMO
having two copper atoms which become photosensitive
after binding to
oxygen (O
2). Taking into account our experimental
illumination conditions, one hypothesis is that the soluble MMO
is
inhibited by the UV or blue region of the spectrum, as Guerrero
and
Jones showed for nitrifying bacteria (
15). We suggest that
immunochemical and structural analyses after isolation of soluble
MMO
from methanotroph cultures may contribute, in further works,
to
explaining the inactivation by high
illumination.
| |
ACKNOWLEDGMENTS |
We are grateful to the Laboratoire Environnement de Petit Saut
for its technical and human support and to Peter Winterton of the
Université Paul Sabatier for his help in translating the manuscript.
We thank Electricité de France (EDF/CNEH) for its financial
support (Convention GP 7573).
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FOOTNOTES |
*
Corresponding author. Mailing address: Centre
d'Ecologie des Systèmes Aquatiques Continentaux, UMR CNRS/UPS
5576, Université Paul Sabatier, 118 Route de Narbonne, 31062 Toulouse Cedex, France. Phone: (33) 5 61 55 67 26. Fax: (33) 5 61 55 60 96. E-mail: dumestre{at}cict.fr.
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REFERENCES |
| 1.
|
Bédard, C., and R. Knowles.
1989.
Physiology, biochemistry, and specific inhibitors of CH4, NH4+, and CO oxidation by methanotrophs and nitrifiers.
Microbiol. Rev.
53:68-84[Abstract/Free Full Text].
|
| 2.
|
Bowman, J. P.,
J. H. Skerratt,
P. D. Nichols, and L. I. Sly.
1991.
Phospholipid fatty acid and lipopolysaccharide fatty acid signature lipids in methane-utilizing bacteria.
FEMS Microbiol. Ecol.
85:15-22.
|
| 3.
|
Bowman, J. P.,
L. I. Sly,
P. D. Nichols, and A. C. Hayward.
1993.
Revised taxonomy of the methanotrophs: description of Methylobacter gen. nov., emendation of Methylococcus, validation of Methylosinus and Methylocystis species, and a proposal that the family Methylococcaceae includes only the group I methanotrophs.
Int. J. Syst. Bacteriol.
43:735-753[Abstract/Free Full Text].
|
| 4.
|
Carlsen, H. N.,
L. Joergensen, and H. Degn.
1991.
Inhibition by ammonia of methane utilization in Methylococcus capsulatus (Bath).
Appl. Microbiol. Biotechnol.
35:124-127.
|
| 5.
|
Cline, J. D.
1969.
Spectrophotometric determination of hydrogen sulfides in natural waters.
Limnol. Oceanogr.
14:454-458.
|
| 6.
|
Conrad, R., and F. Rothfuss.
1991.
Methane oxidation in the soil surface layer of a flooded rice field and the effect of ammonium.
Biol. Fertil. Soils
12:28-32.
|
| 7.
|
Conrad, R.
1996.
Soil microorganisms as controllers of atmospheric trace gases (H2, CO, CH4, OCS, N2O, and NO).
Microbiol. Rev.
60:609-640[Abstract/Free Full Text].
|
| 8.
|
Dalton, H.,
S. D. Prior,
D. J. Leak, and S. H. Stanley.
1984.
Regulation and control of methane monooxygenase, p. 75-82.
In
R. L. Crawford, and R. S. Hanson (ed.), Microbial growth on C1 compounds. American Society for Microbiology, Washington, D.C.
|
| 9.
|
Dumestre, J.-F.,
L. Labroue,
C. Galy-Lacaux,
C. Reynouard, and S. Richard.
1997.
Biomasses et activités bactériennes dans la retenue et à l'aval du barrage de Petit Saut (Guyane): influence sur les émissions de méthane et la consommation d'oxygène.
Hydroécol. Appliquée
9:139-167.
|
| 10.
|
Fay, L., and U. Richli.
1991.
Location of double bonds in polyunsaturated fatty acids by gas chromatography-mass spectrometry after 4,4-dimethyloxazoline derivatization.
J. Chromatogr.
541:89-98.
|
| 11.
|
Galy-Lacaux, C.,
C. Jambert,
R. Delmas,
J.-F. Dumestre,
L. Labroue,
P. Cerdan, and S. Richard.
1996.
Emission de méthane et consommation d'oxygène dans la retenue hydroélectrique de Petit Saut en Guyane.
C. R. Acad. Sci. Series IIa
322:1013-1019.
|
| 12.
|
Galy-Lacaux, C.,
R. Delmas,
C. Jambert,
J.-F. Dumestre,
L. Labroue, and S. Richard.
1997.
Gaseous emissions and oxygen consumption in hydroelectric dams. A case study in French Guiana.
Global Biogeochem. Cycles
11:471-483.
|
| 13.
|
Gosse, P., and A. Grégoire.
1997.
Dispositif de réoxygénation artificielle du Sinnamary à l'aval du barrage de Petit Saut (Guyane).
Hydroécol. Appliquée
9:23-56.
|
| 14.
|
Guckert, J. B.,
D. B. Ringelberg,
D. C. White,
R. S. Hanson, and B. J. Bratina.
1991.
Membrane fatty acids as phenotypic markers in the polyphasic taxonomy of methylotrophs within the Proteobacteria.
J. Gen. Microbiol.
137:2631-2641[Abstract/Free Full Text].
|
| 15.
|
Guerrero, M. A., and R. D. Jones.
1996.
Photoinhibition of marine nitrifying bacteria. 1. Wavelength-dependent response.
Mar. Ecol. Prog. Ser.
141:183-192.
|
| 16.
|
Guézennec, J., and A. Fiala-Medioni.
1996.
Bacterial abundance and diversity in the Barbados Trench determined by phospholipid analysis.
FEMS Microbiol. Ecol.
19:83-93.
|
| 17.
|
Hanson, R. S.,
A. I. Netrusov, and K. Tsuji.
1991.
The obligate methanotrophic bacteria Methylococcus, Methylomonas and Methylosinus, p. 2350-2363.
In
A. Balows, and H. G. Truper (ed.), The prokaryotes. Springer-Verlag, New York, N.Y.
|
| 18.
|
Holmes, A. J.,
A. Costello,
M. E. Lidstrom, and J. C. Murrell.
1995.
Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related.
FEMS Microbiol. Lett.
132:203-208[Medline].
|
| 19.
|
Jannasch, H. W.
1975.
Methane oxidation in Lake Kivu (Central Africa).
Limnol. Oceanogr.
80:860-864.
|
| 20.
|
Joergensen, L., and H. Degn.
1987.
Growth rate and methane affinity of a turbidostatic and oxystatic continuous culture of Methylococcus capsulatus (Bath).
Biotechnol. Lett.
9:71-76.
|
| 21.
|
Keller, M.,
W. A. Kaplan, and S. C. Wofsy.
1986.
Emissions of N2O, CH4 and CO2 from tropical forest soils.
J. Geophys. Res.
91:791-802.
|
| 22.
|
Keller, M.,
M. E. Mitre, and R. F. Stallar.
1990.
Consumption of atmospheric methane in soils of Central Panama: effects of agricultural development.
Global Biogeochem. Cycles
4:21-27.
|
| 23.
|
Makula, R. A.
1978.
Phospholipid composition of methane-utilizing bacteria.
J. Bacteriol.
134:771-777[Abstract/Free Full Text].
|
| 24.
|
Nichols, P. D.,
G. A. Smith,
C. P. Antworth,
R. S. Hanson, and D. C. White.
1985.
Phospholipid and lipopolysaccharide normal and hydroxy fatty acid as potential signatures for methane-oxidising bacteria.
FEMS Microbiol. Ecol.
31:327-335.
|
| 25.
|
Nichols, P. D.,
J. B. Guckert, and D. C. White.
1986.
Determination of monounsaturated fatty acid double bond position and geometry for microbial monocultures and complex consortia by capillary GC-MS of their dimethyldisulfur adducts.
J. Microbiol. Methods
5:49-55.
|
| 26.
|
Patel, R. N.,
C. T. Hou,
A. I. Laskin, and A. Felix.
1982.
Microbial oxidation of hydrocarbons: properties of a soluble methane monooxygenase from a facultative methane-utilizing organism, Methylobacterium sp. strain CRL-26.
Appl. Environ. Microbiol.
44:1130-1137[Abstract/Free Full Text].
|
| 27.
|
Porter, K. G., and Y. S. Feig.
1980.
The use of DAPI for identifying and counting aquatic microflora.
Limnol. Oceanogr.
25:943-948.
|
| 28.
|
Rudd, J. W.,
R. D. Hamilton, and N. E. R. Campbell.
1974.
Measurement of microbial oxidation of methane in lake water.
Limnol. Oceanogr.
19:519-524.
|
| 29.
|
Rudd, J. W.,
A. Furutani,
R. J. Flett, and R. D. Hamilton.
1976.
Factors controlling methane oxidation in shield lakes: the role of nitrogen fixation and oxygen concentration.
Limnol. Oceanogr.
21:357-364.
|
| 30.
|
Rudd, J. W. M., and R. D. Hamilton.
1975.
Methane oxidation in a eutrophic Canadian shield lake.
Ver. Int. Ver. Limnol.
19:2669-2673.
|
| 31.
|
Rudd, J. W. M., and C. D. Taylor.
1980.
Methane cycling in aquatic environments.
Adv. Aquat. Microbiol.
2:77-150.
|
| 32.
|
Scott, D.,
J. Brannan, and I. J. Higgins.
1981.
The effect of growth conditions on intracytoplasmic membrane and MMO activities in Methylosinus trichosporium OB3b.
J. Microbiol.
125:105-119.
|
| 33.
|
Shears, J. H., and P. M. Wood.
1985.
Spectroscopic evidence for a photosensitive state of ammonia mono-oxygenase.
Biochem. J.
226:499-507[Medline].
|
| 34.
|
Sissakian, C.
1992.
Présentation de la retenue de Petit Saut en Guyane Française: cartographie partition de la retenuevolumes et surfacesintégration paysagère.
Hydroécol. Appliquée
4:121-132.
|
| 35.
|
Sundh, I.,
P. Borga,
M. Nilsson, and B. H. Svensson.
1995.
Estimation of cell numbers of methanotrophic bacteria in boreal peatlands based on analysis of specific phospholipid fatty acids.
FEMS Microbiol. Ecol.
18:103-112.
|
| 36.
|
Tathy, J. P.,
R. A. Delmas,
A. Marenco,
B. Cros,
M. Labat, and J. Servant.
1992.
Methane emission from flooded forest in Central Africa.
J. Geophys. Res.
97:6159-6168.
|
| 37.
|
Urakami, T., and K. Komagata.
1984.
Cellular fatty acid composition and quinone system in methane-utilizing bacteria and methylamine-utilizing bacteria, p. 123-133.
In
R. L. Crawford, and R. S. Hanson (ed.), Microbial growth on C1 compounds. American Society for Microbiology, Washington, D.C.
|
| 38.
|
Wassman, R.,
U. G. Thein,
H. Whiticar,
H. Rennenberg,
W. Seiler, and W. J. Junk.
1992.
Methane emission from the Amazon floodplain: characterization of production and transport.
Global Biogeochem. Cycles
6:3-13.
|
| 39.
|
Whalen, S. C.,
W. S. Reeburgh, and C. E. Reimers.
1996.
Control of tundra methane emission by microbial oxidation, p. 257-274.
In
J. F. Reynolds, and J. D. Tenhunen (ed.), Landscape function: implications for ecosystem response to disturbance, a case study in arctic tundra. Springer-Verlag, Berlin, Germany.
|
| 40.
|
White, D. C.,
R. J. Bobbie,
J. S. Herron,
J. D. King, and S. J. Morrison.
1979.
Biochemical measurements of microbial mass and activity from environmental samples. Native aquatic bacteria: enumeration, activity and ecology.
ASTM Spec. Tech. Publ.
695:69-81.
|
| 41.
|
Whittenbury, R.,
K. C. Phillips, and J. F. Wilkinson.
1970.
Enrichment, isolation and some properties of methane-utilizing bacteria.
J. Gen. Microbiol.
61:205-218[Abstract/Free Full Text].
|
Applied and Environmental Microbiology, February 1999, p. 534-539, Vol. 65, No. 2
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
