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APPLIED AND ENVIRONMENTAL MICROBIOLOGY,
0099-2240/98/$04.0010
Mar. 1998, p. 1099–1105 Vol. 64, No. 3
Copyright © 1998, American Society for Microbiology
Characterization of Root-Associated Methanotrophs from
Three Freshwater Macrophytes: Pontederia cordata,
Sparganium eurycarpum, and Sagittaria latifolia†
A. CALHOUN AND G. M. KING*
Darling Marine Center, University of Maine, Walpole, Maine 04573
Received 15 September 1997/Accepted 8 December 1997
Root-associated methanotrophic bacteria were enriched from three common aquatic macrophytes: Ponte-
deria cordata, Sparganium eurycarpum, and Sagittaria latifolia. At least seven distinct taxa belonging to groups
I and II were identified and presumptively assigned to the genera Methylosinus, Methylocystis, Methylomonas,
and Methylococcus. Four of these strains appeared to be novel on the basis of partial 16S ribosomal DNA
sequence analysis. The root-methanotroph association did not appear to be highly specific, since multiple
methanotrophs were isolated from each of the three plant species. Group II methanotrophs were isolated most
frequently; though less common, group I isolates accounted for three of the seven distinct methanotrophs.
Apparent K
m
values for methane uptake by representative cultures ranged from 3 to >17 mM; for five of the
eight cultures examined, apparent K
m
values agreed well with apparent K
m
estimates for plant roots, suggesting
that these strains may be representative of those active in situ.
So¨hngen (33) isolated and characterized the first methane-
oxidizing bacterium by using enrichments from the leaves of
submerged aquatic macrophytes. Subsequently, methanotrophs
have been isolated from soils, sediments, and the water column
of freshwater and marine systems (18, 20, 24, 35, 36). Meth-
anotrophs and methanotrophic activity have also been de-
scribed for mytilid mussels harboring bacterial endosymbionts
(10, 11, 13). In addition, both in vitro and in situ methane
oxidation rates have been documented for various aquatic
plant roots (16, 25, 26, 31).
Although root-associated methanotrophy limits methane
emission from wetlands to the atmosphere and thus plays an
important role in the global methane budget, little is known
about the bacteria responsible for this activity. To date, there
are no published studies of methanotrophic enrichments or
isolates specifically derived from the roots or rhizospheres of
aquatic plants. As a result, the similarity of root meth-
anotrophs to isolates from other systems remains unclear; like-
wise, the host plant specificity of root methanotrophs is un-
known. King (25) and Hanson and Hanson (18) report that
group II methanotrophs dominate root populations on the
basis of signature deoxyribonucleotide hybridization patterns.
However, these studies provide no information on the diversity
or characteristics of root methanotrophs, nor do they address
the extent to which such organisms can be routinely isolated.
We describe here characteristics of root-associated meth-
anotrophs and their distribution among three common aquatic
plant species. The populations of root-associated meth-
anotrophs include at least seven distinct taxa, three and four
each from the phylogenetically coherent groups I and II, re-
spectively; all of the latter four appear novel, based on partial
16S ribosomal DNA (rDNA) sequence analysis. The isolates
most frequently obtained were assigned to group II; group I
isolates were rarer, an observation consistent with previous
results (25). The various isolates from both groups are similar
to extant cultures, based on morphology, colony characteristics
on solid media, and physiological attributes. One-half-satura-
tion constants for methane (apparent K
m
) are also consistent
with previously reported apparent K
m
values for sediment-free
roots of various freshwater plants and extant cultures (25).
Ranges in apparent K
m
and V
max
suggest the possibility that
some methanotrophs may be adapted for colonization of the
root surface while others may be better adapted for coloniza-
tion of the root interior.
MATERIALS AND METHODS
Root enrichments. Pontederia cordata, Sagittaria latifolia, and Sparganium eu-
rycarpum were collected from marshes in Bristol and Orono, Maine (9, 26).
Methanotrophs on or in approximately 4 g (fresh weight) of sediment-free
excised roots were enriched in each of the following nutrient solutions: (i)
Higgins nitrate mineral salts medium (NMS) (10.0 mM KNO
3
, 6.1 mM
Na
2
HPO
4
, 3.9 mM KH
2
PO
4
, 0.8 mM Na
2
SO
4
, 0.2 mM MgSO
4
z 7H
2
O, 0.1 mM
CaCl
2
z 2H
2
O), (ii) NMS without added copper (NMS [2Cu]), (iii) Higgins
ammonium mineral salts (AMS) (same composition as NMS but with 10.0 mM
NH
4
Cl and no nitrate, and (iv) mineral salts medium with no added nitrogen
(2N).
For each of the four media, trace elements were added to give the following
final concentrations: 2 mM ZnCl
2
,2mM CuCl
2
,1mM NaBr, 0.5 mMNa
2
MoO
2
,
2 mM MnCl
2
,1mM KI, 2 mMH
3
BO
3
,1mM CoCl
2
, and 1 mM NiCl
2
. Iron was
added to autoclaved media as FeSO
4
in 1 M HCl to produce a final concentration
of 50 mM and a pH between 6.8 and 7.0. NMS (2Cu) and nitrogen-free basic
mineral salts media were used to select for group II methanotrophs that fix
nitrogen and express soluble methane monooxygenase (sMMO) under copper-
limited conditions.
Roots were agitated at 100 rpm in 300 ml of medium in 1-liter flasks with a 30
to 70% methane-air headspace at 32°C. Roots from each plant were incubated in
triplicate in each of the four media; this procedure was conducted twice for each
plant species, once in September 1994 and once in July 1995.
Culture isolation. Root enrichments were incubated until the medium was
turbid. One milliliter from each of the enrichments was transferred to 10 ml of
nutrient medium (each of the four described above) in 160-ml culture bottles
(incubated at 32°C, with shaking at 100 rpm and a 30 to 70% methane-air
headspace). The enrichments were subcultured weekly for approximately 6
months until there were #5 distinct morphotypes per culture as determined from
phase-contrast microscopy. Subsequent efforts to isolate pure cultures included
a series of serial dilutions in liquid media and on mineral salts agar plates with
washed Bacto agar (Difco, Inc.). The plates were incubated at 32°C in sealed jars
with a 30 to 40% methane headspace. Colonies from the plates were subcultured
every 2 to 3 weeks and transferred to liquid and solid media. Cultures containing
#3 discrete morphotypes as determined by microscopic examination were used
for further characterization.
* Corresponding author. Phone: (207) 563-3146 ext. 207. Fax: (207)
563-3119. E-mail: GKing@Maine.Maine.Edu.
† Contribution 311 from the Darling Marine Center.
1099
Culture characterization. BacLight viability and Gram stains (Molecular
Probes, Inc.) were used for morphological and Gram reaction analysis. One
milliliter of exponentially growing broth culture was transferred to 1.5-ml micro-
centrifuge tubes. BacLight stain was added to the tubes, which were then incu-
bated in darkness for 20 min according to the manufacturer’s instructions; sub-
samples of the stained cultures were transferred to agar-coated slides and
examined with a Zeiss Axioscope with epifluorescence illumination and 4003
Achrostigmat and 1,0003 Plan-neofluar phase-contrast objectives. A small vol-
ume of culture was heat fixed on slides for poly-b-hydroxybutyrate staining with
0.03% (wt/vol) Sudan black B and a 0.5% safranin counterstain (15). Loeffler
methylene blue was used for staining polyphosphate inclusions. Broth cultures
for these stains were 2 to 3 weeks old. A Difco Gram stain kit was also used on
fresh cultures. Bacterial cysts were stained with neutral red and light green S.F.
yellowish dyes (15) with broth cultures at least 3 weeks old. Capsules were
stained with India ink according to the Duguid method (15).
Exospore formation was determined with 2-week-old broth cultures grown in
10 ml of Higgins NMS at 32°C in 160-ml culture bottles with a 30% methane
headspace. A volume of culture (0.5 ml) was transferred to fresh medium to
produce a set of controls; a second set was pasteurized at 80°C for 20 min. Initial
cell density was determined by measuring absorbance at 600 nm on a Beckman
DU 640 spectrophotometer; absorbance was assayed periodically for an addi-
tional 3 weeks. Exospore formation was also determined by microscopy.
Lysis in 0.2 and 2% sodium dodecyl sulfate (SDS) was determined with 1 ml
of unwashed culture. Microcuvettes containing culture and SDS were vortexed
and assayed spectrophotometrically (A
600
). Colony morphology and pigmenta-
tion were determined by examining 1-week-old NMS agar plate cultures. Plates
also were incubated in air and compared to plates incubated in methane to
ensure positive identification of methanotrophic colonies.
Physiological assays. sMMO production was determined by using a modifica-
tion of the naphthalene oxidation assay of Brusseau et al. (6). Cell suspensions
were diluted to an A
600
of 0.2. One milliliter of culture was transferred to a 10-ml
screw-cap tube, and 1 ml of saturated naphthalene solution (about 234 mMat
25°C) was added. The solution was incubated at 25°C and shaken at 200 rpm for
1 h. After incubation, 100 ml of fresh 0.2% tetrazotized o-dianisidine was added.
The absorbance of the resulting solution was read at 525 nm. The intensity of
diazo dye formation was proportional to the oxidation of naphthalene (6). Cul-
tures were considered positive for sMMO when samples appeared blue.
Temperature and pH response measurements were conducted with fresh,
washed cultures. Cells were grown to an A
600
of 0.2 to 0.4 and harvested in
exponential phase by centrifugation for 10 min at 4°C and 9,500 3 g. The pellets
were washed twice in Higgins neutral phosphate buffer (NPB) (6.1 mM
Na
2
HPO
4
, 3.9 mM KH
2
PO
4
[pH 7]) and resuspended in NPB. Nine milliliters of
Higgins nutrient medium was inoculated with 1 ml of washed culture and incu-
bated with a 30% methane headspace at the desired temperature or pH in 60-ml
culture bottles stoppered with green neoprene stoppers. Cultures were agitated
at 100 rpm. Absorbances were assayed periodically and compared to the initial
readings. For the pH assays, pH was adjusted after the addition of Fe, trace
metals, and salts. For pHs of #7.0, a citrate-phosphate buffer (5 mM citric acid,
10 mM Na
2
HPO
4
) was used; for pHs of $7.0, a phosphate buffer solution (10
mM Na
2
HPO
4
,10mMNaH
2
PO
4
) was used.
DNA analysis. Selected cultures (isolates 2, 7, 8, 12, 13, 19, 20, 22, and 23 and
Methylomonas albus BG8 and Methylosinus trichosporium OB3b) were grown in
Higgins medium with nitrate and a 20% methane headspace. The cultures were
harvested by centrifugation and washed twice with 10 mM phosphate buffer (pH
7). The final pellets were resuspended in 13 Tris-EDTA buffer (TE) and stored
frozen (220°C) prior to further analysis. A subsample of thawed cell suspension
was subjected to three cycles of freezing (270°C) and thawing (65°C) and then
incubated with lysis buffer. DNA from cell lysates was extracted with phenol-
chloroform-isoamyl alcohol and purified by standard methods (2). DNA was
stored frozen (220°C) in TE prior to PCR assays.
PCR of methanotroph DNA was based on the use of signature oligonucleo-
tides as specific primers. The specificity of the two primers has been described
previously by Brusseau et al. (7); briefly, 1034-SER (59-CCA-TAC-CGG-ACA-
TGT-CAA-AAG-C-39) hybridizes with the group II methanotrophs, and 1035-
RuMP (59-GAT-TCT-CTG-GAT-GTC-AAG-GG-39) hybridizes with group I
methanotrophs other than those in the genus Methylococcus. These oligonucle-
otides were used as reverse primers in combination with the forward primer, 530f
(59-GTG-CCA-GCM-GCC-GCG-G-39), which hybridizes with eubacteria in
general (34). A typical PCR was based on a 100-ml volume with 1 to2UofTaq
polymerase (Promega, Inc.), 15 mM MgCl
2
, 100 ng of template, and approxi-
mately 100 mM of one of the pairs of forward-reverse primers. Amplification
conditions included the following (after a hot start): 5 min at 95°C (1 cycle); 94°C
denaturation, 55°C annealing, 72°C polymerization (30 cycles); and 72°C final
extension (10 min). PCR products were electrophoresed in 2% NuSieve 3:1
agarose (FMC Corporation) with 13 tris-borate-EDTA buffer (TBE) and visu-
alized with UV illumination after staining with ethidium bromide (27a). All
products were of the expected size (about 500 bp) as determined by comparison
with the electrophoresis of a set of PCR markers (Bio-Rad Laboratories, Inc.).
Deionized water controls were always negative, as were samples of M. albus BG8
and M. trichosporium OB3b DNA when amplified with the group II and group I
primers, respectively.
PCR products were picked from the agarose gels and extracted after treatment
with agarase according to the manufacturer’s (FMC Corp.) instructions. A sub-
sample of the amplified DNA was sequenced at the University of Maine Se-
quencing Laboratory with an ABI 373A sequencer and a Perkin-Elmer ABI
prism dye terminator cycle sequencing kit with Amplitaq DNA polymerase. The
resulting sequences, along with various sequences from GenBank, were aligned
with CLUSTAL V; alignments were also compared with those previously used by
Brusseau et al. (7). Phylogenetic relationships were determined from 432 bp of
sequence (corresponding to Escherichia coli positions 732 to 1194) with programs
from the PHYLIP version 3.5 package (DNAML, DNAPARS, DNADIST,
SEQBOOT, and CONSENSE [12]).
Kinetic analyses. Kinetic assays were conducted with M. trichosporium OB3b
and root isolates dominated by a single morphotype (2, 4, 8, 10, 12, 13, 19, 20).
Cultures were grown at 32°C in 1-liter flasks with 200 ml of Higgins NMS and
were shaken at 150 rpm. Cells were grown to an A
600
of 0.2 to 0.4 and harvested
by centrifugation for 10 min at 10°C and 9,500 3 g. The pellets were washed with
Higgins NPB (10 mM, pH 7) and recentrifuged. The final pellets were resus-
pended in 20 ml of 10% NMS (A
600
, 0.02 to 0.04), and a volume was transferred
to 160-ml cultures bottles to yield final bacterial concentrations in four ranges of
approximately 0.02 to 0.04, 0.04 to 0.06, 0.08 to 0.1, and 0.1 to 0.15 mg ml
21
in
a total volume of 5 ml. These bacterial concentrations were used for incubations
with dissolved methane concentrations of 0.15 to 1, 4 to 8, 8 to 16, and .32 mM,
respectively. The cultures were incubated horizontally in triplicate with vigorous
shaking at 32°C. Headspace samples of 0.2 cm
3
were collected with 1-ml dispos-
able syringes and needles for methane analysis with a Shimadzu 14A gas chro-
matograph and a flame ionization detector operated at 150°C. Methane was
separated with a Porapak Q column in series with a wide-bore capillary column
(DB-1; 30 m by 0.53 mm [outside diameter]) (J&W Scientific, Inc.). Samples
were collected at 10- to 20-min intervals during a 2- to 4-h incubation. Kinetic
parameters (V
max
and apparent K
m
) were determined by fitting data to the
Michaelis-Menten model by using the nonlinear curve-fitting algorithm of Ka-
leidagraph version 3.0.3 (Adelbeck Software). Cell densities were measured at
the beginning and termination of the experiment to determine if the cultures had
grown significantly during the sampling period.
RESULTS
Root enrichments. Enrichments for methanotrophs were
successful in each of 24 attempts, from which 13 cultures were
selected for characterization. The remaining 11 cultures were
morphologically indistinguishable from the vibrioid exospore
formers described below and were not further characterized.
Although pure cultures were not obtained from any of the
enrichments, the majority existed in stable consortia, with only
one consort present at very low densities. In some cases, the
consorts were not evident by microscopy but grew on various
solid media incubated without methane. The consorts used
diverse organic compounds as carbon and energy sources,
including organic acids, amino acids, sugars, and methanol.
Because of its distinctive morphology, it was evident that a
Hyphomicrobium sp. occurred as a consort in some of the
cultures.
Methanotroph characterization. All methanotrophs were
gram negative and mesophilic, growing best at temperatures of
.20°C and at pH values between 6 and 7 (Table 1). Six cul-
tures were dominated by encapsulated rosette-forming vibrios
(strains 7, 10, 13, 20, 23, and 24) 2 to 3 mm long and 1 to 1.5 mm
in width; two of the 6 (strains 13 and 20) were motile. During
stationary phase, all of the rosette-forming vibrios elongated to
a comma shape and produced encapsulated exospores rela-
tively quickly (after 4 to 5 days) in liquid cultures. Four of the
six rosette-forming vibrio cultures grew after pasteurization at
80°C for 20 min. Colonies of the rosette-forming vibrios on
NMS agar plates were low convex with entire edges, of buty-
rous consistency, and buff colored. All of the rosette-forming
vibrios produced sMMO (based on naphthalene oxidation) in
a copper-limited medium (Table 1).
Strains 2, 4, 8, and 12 were rods (0.6 to 1.25 mm by 2.0 to 3.0
mm) that did not produce sMMO in copper-limited medium.
Strain 8 was distinguished from the others by the absence of
cysts, poly-b-hydroxybutyrate, and polyphosphate inclusions.
None of these strains formed rosettes. Capsules were present
for strains 4, 8, and 12. Most colonies on NMS agar were
1100 CALHOUN AND KING APPL.ENVIRON.MICROBIOL.
opaque, low convex, of butyrous consistency, and buff colored.
However, strain 2 differed by forming a high-convex, mucoid
colony with irregular edges.
Strain 19 consisted of encapsulated, nonmotile paired cocci
(0.6 to 1 mm by 0.5 to 1.0 mm) which often formed tetrads.
Cysts were formed in older cultures. Colonies were buff col-
ored and low convex with entire edges. This strain did not
produce sMMO in copper-limited medium.
Strains 21 and 22 were encapsulated, small, motile rods that
formed Azotobacter-like cysts and were sMMO negative in
copper-limited medium. Cells lysed in 0.2% SDS. Strain 21
formed bright pink, low-convex colonies with entire edges.
Strain 22 formed salmon-to-orange colonies of the same gen-
eral description. Both strains formed pellicles in static and
agitated cultures.
Strains 2, 4, 7, 8, 10, 12, 13, 20, 23, and 24 yielded a PCR
product with the 1034-SER but not the 1035-RuMP primer;
on this basis, these strains are assigned to the group II
methanotrophs. Strain 22 yielded a product only with the
1035-RuMP primer and is therefore assigned to the group I
methanotrophs. No amplification product was obtained
from strain 19, but its similarity to the genus Methylococcus
indicates placement in group I. Strain 21 was not assayed by
PCR, but its similarity to the genus Methylomonas also in-
dicates that it is probably in group I. The remaining strains
(1, 3, 5, 6, 9, 11, and 14 to 18) were indistinguishable mor-
phologically from the vibrioid exospore formers (e.g., strains
7, 10, 13, 20, 23, and 24) and are presumably group II
methanotrophs.
Comparison of partial 16S rDNA sequences from selected
strains indicated that many were equivalent (e.g., strain 7 was
equivalent to strain 10; strain 13 was equivalent to strains 20,
23, and 24; and strain 8 was equivalent to strain 12). Maximum-
likelihood phylogenetic analysis based on a 100-iteration boot-
strap data set of aligned sequences for the root strains and
other methanotrophs (Fig. 1) indicated that strains 2 and 12
were distinct but related to previously reported group II se-
quences from a peat bog and that strains 10 and 13 were
TABLE 1. Phenotypic characteristics of methanotrophic isolates from the roots of P. cordata, S. eurycarpum, and S. latifolia
Parameter
Characteristic for isolate:
2 4 812 7 1013202324212219
Cell morphology
Cocci, paired 1
Bacilli 1111 1 1
Vibrioid 1 11111
Length ,1–,2.0 mm 111
Length 2.0–3.0 mm 1111 1 11111
Width 0.6–1.25 mm 1111 1 11111111
Poly-b-hydroxybutyrate 2122 2 22222211
Polyphosphate 1221 2 22222121
Cyst formation 1121 2 22222111
Exospore formation 2111 1 11111222
Group, presumptive II II II II II II II II II II I I I
Gram stain 2222 2 22222222
Capsule 2111 1 11111111
Motility 1222 2 21122112
Rosette formation 2222 1 11111121
Chain formation 1222 2 22222121
Colony morphology
Translucent
Opaque 1111 1 11111111
Low convex 111 1 11111111
High convex 1
Lobate or irregular edges 1
Entire edges 111 1 11111111
Butyrous consistency 111 1 11111111
Mucoid consistency 1
Pigmentation
White or buff 1111 1 11111 1
Pink or rose 1
Salmon to orange 1
Growth in static culture as evenly dispersed
clumps or pellicles
1111 1 11111111
Physiological characteristics
sMMO 2222 1 11111222
Lysis by 0.2% SDS 2222 2 22222112
Lysis by 2% SDS 1111 1 11111111
Growth at 10°C 1221 ND 22112122
Growth at 20°C 1111 ND 11111111
Growth at 30°C 1111 ND 11111111
Strain no. 2 4 8 12 7 10 13 20 23 24 21 22 19
a
ND, no data.
VOL. 64, 1998 ROOT-ASSOCIATED METHANE-OXIDIZING BACTERIA 1101
related but distinct from other root methanotrophs and group
II isolates.
Of the seven distinct strains identified from the various char-
acterizations, four were obtained from P. cordata (Table 2),
including two vibrios (strains 12 and 13), a paired coccus
(strain 19), and a rod (strain 22). Three strains each were
obtained from S. eurycarpum (a vibrio [strain 24] and two
distinct rods [strains 2 and 21]) and S. latifolia (two vibrios
[strains 10 and 20] and one rod [strain 4]). Note that some of
the strain numbers on different plant taxa represented equiv-
alent methanotrophs (e.g., strains 13 and 20). Both group I and
group II methanotrophs were obtained from P. cordata and S.
eurycarpum, while only group II isolates were cultured from S.
latifolia. Group I and group II methanotrophs were also ob-
tained from copper-sufficient NMS, while only group II isolates
were enriched from copper-limited NMS. AMS and nitrogen-
free media supported one group I methanotroph, with group II
strains otherwise dominant.
Kinetic analyses. The methanotrophs used for the kinetic
analyses did not grow measurably during the assays. Methane
oxidation kinetics conformed to a Michaelis-Menten model
(Fig. 2). V
max
values ranged from 42.2 to 118 mmol of methane
mg (dry weight)
21
h
21
; apparent K
m
values ranged from 3.0 to
17.0 mM. The V
max
and apparent K
m
for M. trichosporium
OB3b were 24 6 1.5 mmol mg (dry weight)
21
h
21
and 1.0 6 0.3
mM, respectively. Strains 2, 8, 13, and 20 had similar V
max
and
apparent K
m
values, while cultures 10, 12, and 19 had the
highest apparent K
m
s (Table 3). V
max
values and apparent K
m
s
were significantly correlated when they were plotted as pairs
for all of the cultures (r 5 0.72; P 5 0.03) (Fig. 3).
DISCUSSION
In spite of the global importance of aquatic plants for the
production, transport, and oxidation of methane, little is
known about the specific groups of root-associated bacteria
that directly affect methane dynamics. The results presented
here provide new insights into the taxonomic diversity of root-
associated methanotrophs and some of the important charac-
teristics of these organisms (e.g., methane uptake kinetics).
Although the relative importance in situ of the various meth-
anotrophs obtained during this study is as yet unknown, the
FIG. 1. Unrooted consensus tree derived from a maximum-likelihood anal-
ysis of a 100-iteration bootstrap data set of partial 16S rDNA sequences for
selected group II methanotrophs and root isolates; analysis was performed with
the SEQBOOT, DNAML, and CONSENSE packages of PHYLIP version 3.5.
MPH14 and MPH17 represent sequences obtained from group-specific PCR of
a genomic extract from bog peats (28). Strains I2 and I12 are presumed Methy-
locystis spp., and I10 and I13 are assigned to the genus Methylosinus. Parvus,
Methylocystis parvus; Echinoides, Methylocystis echinoides; Minimus, Methylocystis
minimus; Sporium, Methylosinus sporium; lac, Methylosinus sp. strain lac; ER,
Methylosinus sp. strain ER2.
FIG. 2. Methane uptake rate versus dissolved methane concentration for two
presumed Methylocystis spp. (strain 4 [F] and strain 8 [E]); values represent
means 6 1 standard error. Plots are representative of the seven additional strains
assayed.
TABLE 2. Distribution of presumed methanotrophic strains from the roots of three aquatic macrophytes under four enrichment regimes
Enrichment
Distribution of strains
a
from:
P. cordata S. eurycarpum S. latifolia
NMS Methylococcus sp. (19),
Methylocystis sp. (12)
Methylocystis sp. (8),
Methylomonas sp. (21)
Methylocystis sp. (4),
Methylosinus sp. (20)
NMS, Cu2 Methylosinus sp. (6, 13) Methylosinus sp. (5, 15) Methylosinus sp. (1, 14)
AMS Methylosinus sp. (16) Methylocystis sp. (2),
Methylosinus sp. (18)
Methylosinus sp. (7, 17)
2N Methylosinus sp. (9),
Methylomonas sp. (22)
Methylosinus sp. (11, 24) Methylosinus sp. (10, 23)
a
Genus designations are presumptive and based on strain descriptions and assignments in the text; strain numbers are given in parentheses.
1102 CALHOUN AND KING APPL.ENVIRON.MICROBIOL.
dominance of group II forms in the enrichments is consistent
with earlier studies that showed a greater abundance of group
II than of group I based on signature oligonucleotide hybrid-
ization to 16S rRNA in genomic root extracts.
Most of the methanotrophs obtained in this study (e.g.,
strains 7, 10, 13, 20, 23, and 24) have been assigned to the
genus Methylosinus on the basis of various morphological and
physiological characteristics. Characteristics shared by these
strains and the genus Methylosinus include Gram reaction
(negative); lack of motility; vibrioid morphology; size (2 to 3
mm by 1.0 to 1.5 mm); formation of rosettes and exospores;
expression of sMMO in copper-limited media; absence of
polyphosphate, poly-b-hydroxybutyrate, and cysts; and colony
morphology and color (low convex and buff colored).
Although the morphology of the vegetative cells is most
similar to descriptions of Methylosinus sporium (5, 14, 19, 36),
the exospores of M. sporium lack capsules (36), in contrast to
consistent encapsulation for the isolates described here. Anal-
ysis of partial 16S rDNA sequences also supports the assign-
ment of these strains to the genus Methylosinus, as indicated by
their relationship to M. sporium and Methylosinus strain LAC.
However, the two distinct but closely related root meth-
anotrophs represented by strains 10 and 13 clearly differed
from other Methylosinus strains, including two apparently
novel sequences recently reported from a peat bog (Fig. 1)
(28).
A second taxonomic grouping encompasses strains 2, 4, 8,
and 12. Size, morphology, lack of motility, and response to
0.2% SDS suggest an affinity to the genus Methylocystis (5). The
lack of sMMO expression is consistent with Methylocystis par-
vus. In addition, polyphosphate, poly-b-hydroxybutyrate, mo-
tility, and capsule formation are variable in the genus Methy-
locystis (5) as they are in strains 2, 4, 8, and 12. Results of a
phylogenetic analysis indicate that strains 2, 4, 8, and 12 (strain
4 was not sequenced but is otherwise identical to 8 and 12)
form a distinct group more closely related to sequences from a
peat bog in the United Kingdom (MPH14 and MPH17) than to
the Methylosinus-like strains from roots or other known meth-
anotrophs (Fig. 1). In addition, strain 2 differs from strains 8
and 12, which is consistent with its morphological and physio-
logical characteristics (e.g., lack of spores and capsules [Table
1]). Thus, this second grouping of strains differs from the first
and likely includes at least two distinct taxa as well (e.g., strain
2 versus strains 4, 8, and 12).
A third grouping is suggested by the characteristics of strains
21 and 22. Morphology, size, encapsulation, motility, pigmen-
tation, and Azotobacter-like cysts strongly suggest affinities with
the genus Methylomonas. Pigmentation and polyphosphate in-
clusions in strain 21 and poly-b-hydroxybutyrate inclusions in
strain 22 indicate their assignment to Methylomonas methanica
and the closely related Methylomonas aurantiaca and Methyl-
omonas fodinarum, respectively (4). The latter assignment is
supported by 16S rDNA sequence analyses which show a high
degree of identity between strain 22 and M. aurantiaca and M.
fodinarum.
Strain 19 is most similar to the genus Methylococcus. Encap-
sulation, lack of motility, the presence of paired cocci (0.6 to 1
mm by 0.5 to 1.0 mm) that often formed tetrads, and the
presence of polyphosphate and poly-b-hydroxybutyrate are all
consistent with the characteristics of this genus (5). Colony
characteristics and the lack of lysis in 0.2% SDS for strain 19
are also similar to characteristics of this genus. However, in
contrast to the well-known production of sMMO by Methylo-
coccus capsulatus Bath, strain 19 is sMMO negative. The in-
ability of the 1035-RuMP primer to amplify DNA from strain
19 is consistent with previous reports of the response of Methy-
lococcus spp. (7) to this primer and supports the generic as-
signment, albeit indirectly.
Distribution and diversity of root-associated methanotrophs.
Although at least seven distinct taxa were enriched from three
plant species in this study, the true level of methanotroph
diversity is likely higher, since enrichments generally select
against some strains. Several of the methanotrophs obtained
from the enrichments were cosmopolitan, appearing in all nu-
trient regimens for all plants. These may be representative of
the most common taxa, or at least the taxa that are generally
distributed and most competitive in enrichments. In contrast,
several strains appeared more restricted in their distribution,
occurring only in a specific medium or on a single plant species.
These taxa may be representative of the least-abundant meth-
anotrophs.
The cosmopolitan distribution of several of the taxa suggests
that at least some root-methanotroph associations may be op-
portunistic. This differs from other microbe-wetland root as-
sociations that involve much more specific host interactions
(e.g., associations based on nitrogen-fixing actinomycetes [32]).
Whether the few, more specific methanotroph associations
known for animals (10, 11) represent exceptions among a
larger number of opportunistic associations is unclear.
Several lines of evidence, including results from PCR assays
with group I- and group II-specific primers, indicate that most
of the root methanotrophs belong to group II (four of the
seven distinct taxa and 21 of 24 total isolates). Group I and
FIG. 3. V
max
versus K
m app
for selected methanotrophic strains (r 5 0.72; P 5
0.03); values are summarized in Table 3.
TABLE 3. Kinetic parameters for methane oxidation by selected
methanotrophic strains from sediment-free roots of S. eurycarpum,
P. cordata, and S. latifolia and by Methylosinus trichosporium OB3b
Strain
V
max
a
(mean 6 1 SE)
K
m app
(mM)
(mean 6 1 SE)
V
max
K
m app
21
2 55.0 6 7.0 7.2 6 2.3 7.6
4 102.0 6 5.0 4.3 6 0.7 23.7
8 66.2 6 8.0 3.0 6 1.0 22.1
12 118.0 6 11.0 13.2 6 3.8 8.9
19 60.0 6 8.0 13.0 6 3.7 4.6
10 133.0 6 25.0 17.0 6 10.0 7.8
13 42.2 6 6.4 3.4 6 1.6 12.4
20 71.2 6 3.0 6.8 6 1.0 10.5
OB3b 24.0 6 1.5 1.0 6 0.3 24.5
a
Expressed in micromoles per milligram (dry weight) per hour.
VOL. 64, 1998 ROOT-ASSOCIATED METHANE-OXIDIZING BACTERIA 1103
group II methanotrophs occur on P. cordata and S. eurycarpum
roots, but only group II was isolated from S. latifolia. However,
this probably does not accurately reflect methanotroph diver-
sity in situ, since PCR of genomic DNA extracts from S. lati-
folia reveals both groups (37).
The availability of methane, oxygen, nitrogen, and copper
probably plays a major role in determining methanotrophic
population structure (1). Group II methanotrophs may domi-
nate in environments where growth rates are restricted peri-
odically by deprivation of nutrients, particularly nitrogen (17).
Under such conditions, nitrogenase expression presumably
provides a selective advantage for group II methanotrophs.
Group II methanotrophs might also be expected to dominate
in systems with an abundance of methane, such as wetlands,
since they grow more efficiently than group I methanotrophs at
high substrate concentrations (1, 7, 25, 30).
Kinetic analyses. The V
max
and apparent K
m
reported here
(Table 3) for M. trichosporium OB3b (24 6 1.5 mmol of meth-
ane mg [dry weight]
21
h
21
and 1.0 6 0.3 mM, respectively)
agree well with results reported by others, especially the values
of Joergensen and Degn (22) that were based on membrane
inlet mass spectroscopy, which eliminates phase transfer limi-
tations. Root methanotroph V
max
values (42 to 133 mmol mg
[dry weight]
21
h
21
) significantly exceed those of other meth-
anotrophs characterized to date (3, 18, 22). Using root meth-
anotroph V
max
values and the observed maximal methane ox-
idation rates of washed roots in vitro (1 to 10 mmol g [dry
weight]
21
h
21
[25]), one can estimate the population size nec-
essary to account for root activity. The values thus obtained,
3 3 10
7
to 9.5 3 10
8
cells g (dry weight)
21
of root, are clearly
speculative, but they indicate that methanotrophs likely repre-
sent a significant fraction of the root microbiota.
Apparent K
m
values (about 3 to 7 mM [Table 3]) for 5 of the
8 strains assayed are comparable to estimates obtained for the
washed roots of a variety of wetland macrophytes (3 to 6 mM)
(25), which suggests that at least some of the root enrichments
may be representative of the dominant methanotrophs in situ.
Several strains with somewhat higher values (.10 mM) may
represent less important populations. The apparent K
m
softhe
root strains are also comparable to values obtained for other
aquatic systems, including lake and wetland sediments (about
2to10mM) (8, 23, 27, 29) and peats (about 4 mM) (38).
In general, neither V
max
, apparent K
m
, nor V
max
/apparent
(K
m app
) K
m
varied consistently among the strains (Table 3).
For instance, strains 4, 8, and 12 are very similar taxonomically;
however, although V
max
is comparable for 4 and 12, the ap-
parent K
m
s for these strains differ. Likewise, the apparent K
m
s
for strains 4 and 8 are comparable, but the V
max
s differ. The
most consistent trend among the kinetic results is the positive
correlation (r 5 0.72; P 5 0.03) between V
max
and K
m app
,
which suggests that a lower uptake affinity accompanies in-
creased uptake capacity for methane. This relationship may
reflect methanotrophic ecological strategies. Strains with a rel-
atively low K
m app
may have a competitive advantage in the
root interior, where methane concentrations are low, while
strains with a higher K
m app
may have an advantage in the
rhizoplane. Future efforts based on strain-specific fluorescent
probes could prove useful in determining the relationship be-
tween kinetic characteristics and methanotroph microzona-
tion.
In summary, representatives of four methanotrophic genera
and both of the phylogenetically coherent groups (I and II)
have been enriched from the roots of aquatic macrophytes.
Group II taxa (e.g., Methylocystis and Methylosinus spp.) were
most abundant and included four potentially novel strains,
based on partial 16S rDNA analysis. The isolation and char-
acterization of root methanotrophs provide a basis for under-
standing the physiological limitations of methane oxidation in
situ, as well as material for the development of specific nucleic
acid and immunological probes for assessing in situ distribu-
tion and abundance.
ACKNOWLEDGMENTS
This work was funded by NASA grant NAGW-3346.
We thank K. Hardy for technical support and S. Schnell and H. G.
Williams for helpful input.
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