AEM Accepts, published online ahead of print on 16 May 2014 Appl. Environ. Microbiol. doi:10.1128/AEM.01259-14 Copyright © 2014, American Society for Microbiology. All Rights Reserved.
1
5-May-2014
2
Response of a paddy soil methanogen to syntrophic growth as revealed by
3
transcriptional analyses
4
5
6 7 8 9
Pengfei Liu1, Yanxiang Yang1, Zhe Lü1 and Yahai Lu1,2 1
College of Resources and Environmental Sciences, China Agricultural University,
Beijing 100193, China 2
College of Urban and Environmental Sciences, Peking University, Beijing 100871,
China
10 11
*Corresponding author:
12
Yahai Lu
13
Urban and Environmental Sciences,
14
Peking University,
15
Beijing 100871, China.
16
Phone and fax: + 86 10 62755683.
17
E-mail:
[email protected].
18 19 20
Running title: Response to syntrophic growth of a paddy soil methanogen.
21
Key words: Methanogens; the Wolfe cycle; syntrophy; propionate; butyrate.
22
1
23
ABSTRACT
24
Members of Methanocellales are widespread in paddy field soils and play the key role in methane
25
production. These methanogens feature largely in their adaptation to low H2 and syntrophic
26
growth with anaerobic fatty acids oxidizers. The adaptive mechanisms, however, remain unknown.
27
In the present study, we determined the transcripts of 21 genes involved in the key steps of
28
methanogenesis and acetate assimilation of Methanocella conradii HZ254, a strain recently
29
isolated from paddy field soil. M. conradii was grown in monoculture and syntrophically with
30
Pelotomaculum thermopropionicum (propionate syntroph) or Syntrophothermus lipocalidus
31
(butyrate syntroph). Comparison of the relative transcript abundances showed that three
32
hydrogenase encoding genes and all methanogenesis related genes tested were up-regulated in
33
cocultures relative to monoculture. The genes encoding for formylmethanofuran dehydrogenase
34
(Fwd), heterodisulfide reductase (Hdr) and the membrane-bound energy converting hydrogenase
35
(Ech) were the most up-regulated among the evaluated genes. The expression of formate
36
dehydrogenase (Fdh) encoding gene was also significantly up-regulated. By contrast, an acetate
37
assimilation gene was down-regulated in cocultures. The genes coding for Fwd, Hdr and the D
38
subunit of F420-nonreducing hydrogenase (Mvh) form a large predicted transcription unit and
39
therefore the Mvh/Hdr/Fwd complex capable of mediating the electron bifurcation and connecting
40
the first and last steps of methanogenesis was predicted to be formed in M. conradii. We propose
41
that Methanocella methanogens cope with low H2 and syntrophic growth by: (1) stabilizing of
42
Mvh/Hdr/Fwd complex; and (2) activating the formate-dependent methanogenesis.
43
2
44
INTRODUCTION
45
The 16S rRNA gene of a new type methanogen Methanocellales was discovered in 1998
46
from the surface of rice root (1). The 16S rRNA gene sequences were found phylogenetically
47
branching off between Methanosarcinaceae and Methanomicrobiales. An enrichment culture at
48
50°C revealed that these organisms contained the methyl-coenzyme M reductase encoding genes,
49
thus confirming their nature as methanogenic archaea (2, 3). They were found widespread in rice
50
field soils and the environments including the desert soil (4-7). Due to the difficulty of cultivation,
51
many molecular studies were conducted to characterize their ecological functions prior to the
52
isolation into pure culture. The application of stable isotope probing technology showed that these
53
organisms outcompeted other methanogens under low H2 condition (8). This result suggested for
54
the first time that these methanogens might be intrinsically adaptive to low H2. Moreover, they
55
were found to play the key role in CH4 production from root-derived material in the rice
56
rhizosphere in situ, illustrating their ecological significance and niche specificity (9).
57
More evidences for the adaptation of Methanocellales to low H2 came from the investigations
58
of syntrophic oxidation of short-chain fatty acids (10-14). In the investigations of syntrophic
59
oxidations of acetate, propionate and butyrate in Italian and Chinese rice soils, it was revealed that
60
Methanocellales played the key role in H2 consumption for the syntrophic interactions (10-14).
61
These ecological studies reconfirm that Methanocellales are adapted to low H2 conditions. The
62
mechanism, however, remains unknown.
63
H2 is the major electron donor for CH4 production and retains the central role in the
64
interspecies electron transfer for syntrophic fatty acids oxidation (15). Several studies have aimed
65
to understand the mechanisms for the low H2 adaptation of methanogens. A study on 3
66
Methanothermobacter thermautotrophicus revealed that one of two methyl-coenzyme M
67
reductases (MCRI) dominated under syntrophic conditions while both MCRI and MCRII were
68
expressed in pure culture (16). MCRI has a higher substrate affinity than MCRII and hence may
69
play an important role in low H2 adaptation. The study on Methanococcus maripaludis (17)
70
showed that the genes coding for enzymes that reduce or oxidize the deazaflavin coenzyme F420
71
were significantly up-regulated under H2 limitation. Similar results were observed in several other
72
studies (16, 18-23). A recent study on M. maripaludis grown syntrophically with Desulfovibrio
73
vulgaris revealed that most of genes for methanogenesis were up-regulated while those for
74
biosynthetic functions declined compared with monoculture (24). In addition, this methanogen
75
appears capable of activating the functions of paralogous genes to cope with the changing
76
substrate availability (24). The transcript level of paralogs that were previously found upregulated
77
with hydrogen limitation in monoculture (17) was significantly down regulated in the syntrophic
78
coculture. Therefore, it appears that different paralogs are activated under pure low H2 and
79
syntrophic growth conditions. Methanospirillum hungatei, on the other hand, showed that the gene
80
expression levels of hydrogenase and formate dehydrogenase did not vary with culture conditions
81
(25). Given that Methanocella are environmentally important, elucidating the mechanisms for
82
their low H2 adaptation is of great interest.
83
The first Methanocella strain, M. paludicola SANAE, was isolated from a Japanese rice field
84
soil in 2007 (26). Thereafter, two more isolates, M. arvoryzae MRE50 and M. conradii HZ254,
85
were obtained from the Italian and Chinese rice field soils, respectively (27, 28). The whole
86
genome sequences of all three strains are now available (29-31). Of the three strains, M. conradii
87
HZ254 shows the fastest growth with a doubling time of 6.5-7.8 hours under optimum condition
4
88
(27). In the present study, we constructed two cocultures by growing M. conradii syntrophically
89
with Pelotomaculum thermopropionicum or Syntrophothermus lipocalidus. The transcript levels
90
of key genes associated with methanogenesis and acetate assimilation in M. conradii were
91
determined during the syntrophic growth in comparison with the growth in monoculture. The
92
genome of M. conradii lacks carbon monoxide dehydrogenase and hence acetate is essential for
93
growth (27). Therefore, the expression of acetate assimilation-related genes is related to the first
94
step of M. conradii biosynthesis. Formate may serve as an alternative shuttle for the interspecies
95
electron transfer in the syntrophic fatty acids oxidations (32, 33). Physiological tests showed no
96
growth of M. conradii on formate alone (27), but the genes coding for formate dehydrogenase
97
(Fdh) are present and were expressed under the experimental conditions. We therefore reevaluated
98
the growth of M. conradii on formate in the presence of H2.
99
100
MATERIALS AND METHODS
101
Microorganisms and cultivation. Methanocella conradii strain HZ254T was isolated in our lab
102
and has been described previously (27). Pelotomaculum thermopropionicum strain SI (DSM13744)
103
and Syntrophothermus lipocalidus strain TGB-C1 (DSM12680) were purchased from Deutsche
104
Sammlung von Mikroorganismen und Zellkulturen GmbH (Braunschweig, Germany).
105
M. conradii monoculture was grown in 550 ml serum bottles containing 100 ml of minimal
106
salts medium (with 10% volume of inoculation) under H2: CO2 (80:20 [v/v]) at 1.7 atm and
107
incubated for about 24 days to allow its full utilization of H2 in the headspace (Fig. 1A), as
108
described previously (27). The production of CH4 and the consumption of H2 were determined.
5
109
For the test of formate utilization, 4 mM H13COONa (99 atom% 13C, Sigma-Aldrich) was added
110
to a 125 ml serum bottle containing 50 ml medium. All other conditions were same as H2 only
111
incubation.
112
Syntrophic cocultures were obtained by adding ~5% volume of P. thermopropionicum or S.
113
lipocalidus to a vigorously growing culture of M. conradii, flushing the headspace with N2 and
114
finally adjusting the headspace with N2: CO2 (80:20 [v/v]) to a final pressure of 1.7 atm. The
115
substrate sodium propionate or sodium butyrate was added to a final concentration of 20 mM. The
116
production of CH4 and acetate and the consumption of propionate and butyrate were monitored to
117
verify growth. After fourteen and four successive transfers were conducted (each time with 10%
118
volume of inoculation) for propionate and butyrate cocultures, respectively, 550 ml serum bottles
119
containing 100 ml of minimal salts medium plus 25 mM butyrate or 20 mM proprionate were
120
inoculated and incubated for cell collections. Cultivation was carried out at 55 °C in dark without
121
agitation.
122
The cell density of HZ254 at the early stage of logarithmic growth was low and it was
123
technically problematic to harvest sufficient amount of cells for RNA extraction. Therefore, cells
124
were harvested only once for M. conradii monoculture at the middle point of logarithmic growth,
125
which was about 4-5 days after inoculation (Fig. 1A, arrows). For two syntrophic cocultures, the
126
time window of logarithmic growth was sufficient for collecting the cells twice at the early and
127
middle of logarithmic phases, respectively (sampling was carried out about 5, 7 days and 6, 8 days
128
after inoculation for syntrophic propionate and butyrate cocultures, respectively, Fig. 1B and C,
129
arrows). This would allow us to track the possible change in gene expression during the
130
logarithmic growth. For all sampling points, three replicates were carried out except for MP2, 6
131
which had only two replicates. The cell suspensions were centrifuged at 16, 000 × g at 4 °C for 10
132
min (Avanti J-26XP, Beckman Coulter, USA), and the pellets were stored immediately at -80 °C
133
until RNA extraction.
134
Chemical analyses. Gasses were sampled with a pressure-lock syringe (VICI, USA) and the
135
concentration of methane and hydrogen were determined using a gas chromatograph (7890A,
136
Agilent, USA) equipped with 80/100-mesh, PORAPAK Q column (SUPELCO, Sigma-Aldrich,
137
USA) and a thermal conductivity detector. Nitrogen was used as a carrier gas and as reference for
138
thermal conductivity detector (TCD). The column and the detector temperatures were 45 °C and
139
250 °C, respectively. The concentrations of propionate, butyrate and acetate in the medium were
140
measured with a high-performance liquid chromatograph (1200, Agilent, USA) equipped with 4.6
141
mm ID × 250 mm (5μm), ZORBAX SB-Aq C18 column (Agilent, USA) and a UV detector. The
142
column was operated at 25 °C, and 20 mM NaH2PO4 in 0.5% acetonitrile (adjust pH to 2.0 with
143
H3PO4) was used as carrier at a flow rate of 0.8 ml min-1. The liquid samples were filtrated with a
144
PES membrane filter of 0.22-μm pore size (ANPEL, Shanghai, China) before the
145
measurements.
146
For the measurements of 13CH4 and 13CO2, 0.5 ml of gas sample was taken from the
147
headspace with a pressure-lock syringe (VICI, USA) and diluted into 15 ml serum tube containing
148
pure N2. Stable isotope ratios of 13C/12C of CH4 and CO2 were determined using gas
149
chromatograph-combustion-isotope ratio mass spectrometry (GC-C-IRMS) system (Thermo
150
Finnigan, Germany) following the procedures described previously (34).
151
Nucleic acids extraction, purification and cDNA synthesis. For each samples collected above,
152
RNA was extracted using TRIzol reagent (Invitrogen) as described previously (35) with 7
153
modifications. Cell pellets with 1.2 ml of TRIzol reagent were homogenized at 5.0 m s-1 for 20 s
154
in a FastPrep-24 (MP Biomedicals, USA). Tubes were incubated at 25 °C for 3 min, and 250 μl
155
chloroform was added followed by 15 s shaking by hand. After incubation at 25 °C for 10 min, the
156
suspension was centrifuged at 16, 000 × g at 4 °C for 15 min. The supernatants 750 - 800 μl was
157
transferred to a new tube, and 800 μl of 2-propanol was added, mixed and incubated at 25 °C for
158
15 min. The mixtures were centrifuged at 16,000 × g for 15 min at 4 °C. The supernatants were
159
discarded and the pellets were washed with 1000 μl of 70% ethanol. The pellets were air-dried and
160
the RNA samples were finally dissolved in 50 μl RNase-free water. DNA digestion was performed
161
by incubation at 37 °C for 1 h in a total volume of 50 μl containing 41 μl of RNA extracts, 5 μl of
162
RQ1 RNase-Free DNase 10× reaction buffer, 3 μl of RQ1 RNase-free DNase, and 1 μl of
163
recombinant RNasin RNase inhibitor (Promega, Madison, WI, USA). RNA was further purified
164
using an RNeasy mini kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions.
165
The quality and purity of RNA were checked by 1% agarose gel electrophoresis and
166
NanoDrop-2000 spectrophotometry (NanoDrop Technologies, Wilmington, DE). Both indicated a
167
good quality of RNA extracts. The ImProm-II reverse transcription system (Promega, Madison,
168
WI, USA) was used to synthesize the complete cDNA according to the manufacturer’s instructions.
169
To verify the absence of DNA, a control reaction was performed with nuclease-free water instead
170
of reverse transcriptase.
171
For the quantification of 16S rRNA gene and transcripts, DNA and RNA were co-extracted
172
by using AllPrep DNA/RNA Mini Kit (Qiagen, Germany) according to the manufacturer manual.
173
DNA in co-extracts was directly used for quantification. For RNA quantification the co-extracts
174
were subjected to DNA digestion and purification and then RNA was converted to cDNA as
8
175
described above.
176
Primer design, qPCR and data analysis. Primers for the relative quantification of functional
177
genes were designed with Primer Premier 6 (Premier, Canada) and synthesized by Life
178
Technologies (Shanghai, China) (Table S1). The optimal melting temperature (58 °C) of each
179
primer set was experimentally verified with genomic DNA from M. conardii as template. qPCR
180
was performed with the 7500 real-time PCR system (Applied Biosystems, USA).Each 25 μl
181
reaction mixture contained 5 μl 25× diluted cDNA, 12.5 μl 2× SYBR Green PCR mix (Applied
182
Biosystems, USA), 0.5 μl of bovine serum albumin and 0.2 μM each primer. Thermocycling
183
conditions were as follows: 50 °C for 2 min, 95 °C for 10 min, 45 cycles of 95 °C for 15 s, 58 °C
184
for 30 s and 72 °C for 1 min, with fluorescence detection at the end of each extension step.
185
Amplification was followed by a melting program consisting of 95 °C for 1 min, 60 °C for 1 min
186
and a step-wise temperature increase of 0.5 °C per 10 s with fluorescence detection at each
187
temperature transition. Single peak of the melting curve and the presence of only one band in the
188
electrophoresis gel indicated that single amplicon was generated from each primer pair. The
189
optimal baseline and threshold cycle (Ct) values were calculated by using the ABI 7500 real time
190
PCR system sequence detection software version 1.3.1 according to the user’s manual.
191
Amplification efficiency for each individual reaction was calculated from the amplification profile
192
of each sample (36). All transcript abundances were normalized to 16S rRNA levels of M.
193
conradii to obtain the relative expression levels. It is known that rRNA constitute the major part of
194
total RNA (over 90%), and the ratio of 16S rRNA to total RNA is relatively constant (37).
195
Therefore, we assumed that the ratio of a given mRNA target to 16S rRNA transcripts represented
196
the relative expression level of that gene against the total RNA. Three biological replicates (two
9
197
198
for MP2) and three technical replicates were performed for measurement. For the quantification of 16S rRNA gene and transcripts, primer pair Ar364f/Ar915r pair (38)
199
was used. Each 25 μl reaction mixture contained 12.5 μl 2× Go Taq® qPCR Master mix (Promega,
200
Madison, WI, USA) and 0.2 μM Ar364f/Ar915r pair, 0.5 μl of bovine serum albumin (10mg/ml),
201
and 5 μl of DNA (1/103 dilution) or cDNA (1/104 dilution). Thermocycling conditions were as
202
follows: 95 °C for 2 min, 45 cycles of 95 °C for 30 s, 65 °C for 30 s, 72 °C for 1 min and 82 °C
203
for 1 min, with fluorescence detection at the end of each extension step. Amplification was
204
followed by a melting program consisting of 95 °C for 1 min, 60 °C for 1 min and a step-wise
205
temperature increase of 0.5 °C per 10 s with fluorescence detection at each temperature transition.
206
The DNA standard and transcript standard were prepared as described earlier (39), excepted that
207
we used the purified PCR product of a 16S rRNA clone amplified by T7/M13R primer pairs as the
208
DNA standard. The concentration of DNA and transcript standard ranged from 1.0×102 to 1.0×108
209
copies μl-1. Each measurement was performed in three replicates. Since the genome of M. conradii
210
contains two copies of 16S rRNA gene, the cell number of M. conradii per ml was estimated by
211
dividing the total quantity of 16S rRNA gene per ml by 2.
212
The mean ± SD of all replicates were calculated and the analysis of variance was performed
213
to test significant differences between treatments using DUNCAN test in the SAS program (SAS
214
Institute, Cary, IN).
215
216
RESULTS
217
M. conradii monoculture showed a lag phase of 3-4 days before the exponential growth (Fig. 1A).
10
218
CH4 was stoichiometrically produced from H2 and CO2. The production of CH4 showed a lag
219
phase of 3-4 days in propionate coculture and 6 days in butyrate coculture (Fig. 1B and C). H2 in
220
the headspace reached a maximal partial pressure of 30-40 Pa in propionate and 80 Pa in butyrate
221
cocultures, respectively. CH4 and acetate were produced stoichiometrically during the syntrophic
222
oxidations of propionate and butyrate. The doubling time of M. conradii was estimated to be 7-8
223
hours in monoculture based on CH4 production, consistent with the maximal growth rate under
224
optimum condition (27). The doubling time during the syntrophic growth increased to 14-16 hours
225
in butyrate and 40-45 hours in propionate cocultures, respectively.
226
RNA sample in monoculture was collected in the middle exponential phase (Fig. 1A, arrows).
227
RNA sampling in syntrophic cocultures was performed twice, in the early and mid exponential
228
phases, respectively (Fig. 1B and C, arrows). Quantification of rRNA transcripts indicated that the
229
rRNA expressions of M. conradii in syntrophic cultures decreased by a factor of 1.8 to 2.4
230
compared with monoculture (Table 1). Twenty-one genes involved in methanogenesis and acetate
231
assimilation were selected for the relative expression analyses (Table S1). Primers targeting each
232
gene were designed according to the genome sequence. If a predicted transcription unit contains
233
multiple genes, one gene was selected for the measurement (Table S1).
234
Hydrogenases and methanogenesis. M. conradii contains two types of membrane-bound
235
hydrogenases, the energy-converting hydrogenase (Ech) and the methanophenazine-reducing
236
hydrogenase (Vht), and two types of cytoplasmic hydrogenase, the F420-reducing hydrogenase
237
(Frh) and the methyl viologen-reducing hydrogenase (Mvh). An Ech-like hydrogenase was
238
predicted, which was similar to Ech but lacking EchA, the subunit coding for proton transporter.
239
Frh has two predicted transcription units, fhrA1G1B1 and fhrA2DG2B2, and a third single frhB3. 11
240
Mvh also has two predicted transcription units, mvhG1A1 and mvhG2A2, with the genes mvhD1
241
and mvhD2 located at other loci (Fig. S1).
242
The transcript level of echE was significantly up-regulated 2 to 3-fold in cocultures compared
243
with pure culture (Table 2). The transcript level of vhtG did not change in response to coculture
244
except for an increase in propionate at the first sampling. The transcript level of frhA2 was
245
up-regulated 1.6 to 1.9-fold in coculture. The transcript abundance of frhA1 was approximately
246
two orders of magnitude lower than frhA2 and showed no response to syntrophic growth. Similarly,
247
the transcript level of mvhA2 increased 1.7 to 1.9-fold in cocultures except in butyrate coculture at
248
the early sampling. The transcript abundance of mvhA1 was 20 to 33-fold lower than mvhA2 and
249
showed no response to syntrophic growth.
250
The enzymes involved in methanogenesis of M. conradii comprise formylmethanofuran
251
dehydrogenase (Fmd and Fwd, containing Mo and W, respectively), formylmethanofuran: H4MPT
252
formyltransferase (Ftr), methenyl-H4MPT cyclohydrolase (Mch), methylene-H4MPT
253
dehydrogense (Mtd), methylene-H4MPT reductase (Mer), methyl- H4MPT: coenzyme M
254
methyltransferase (Mtr), methyl-coenzyme M reductase (Mcr), and CoB-CoM heterodisulfide
255
reductase (Hdr) (Fig. S1). Transcript analysis was conducted except for Ftr and Mch enzymes.
256
The transcript abundance of fwdB was significantly up-regulated 3.4 to 4.6-fold in cocultures
257
(Table 2). The transcript level of fmdB (encoding the B subunit of Fmd) was four orders of
258
magnitude lower than fwdB and showed no response to syntrophic growth. The transcript levels of
259
mer, mtrH and mcrA were moderately up-regulated (1.6- to 2.3-fold) in cocultures, while mtd
260
showed an increase only in propionate coculture at the first sampling. The gene hdrB has three
261
homologous copies. The hdrB2 is located in the large predicted transcription unit 12
262
fwdGF/hdrC2B2A2/mvhD2/fwdBAC and its expression was significantly up-regulated 2.4 to
263
3.7-fold in cocultures (Table 2). The hdrB1 is located in the predicted transcription unit of
264
hdrA1B1C1 and hdrB3 is linked to the predicted transcription unit coding for Ech-like
265
hydrogenase (Fig. S1). The transcripts of hdrB1 and hdrB3 were also up-regulated, but
266
significantly only in propionate coculture. The transcript abundance of hdrB1 was two orders of
267
magnitude lower than hdrB2 and hdrB3 (Table 2).
268
The gene coding for the A subunit of F420-reducing formate dehydrogenase (fdhA) was
269
transcribed to a similar level as hdrB2 and mtrH and was markedly up-regulated (3 to 5-fold) in
270
cocultures relative to monoculture (Table 2).
271
Acetate assimilation. M. conradii employs the AMP-forming acetyl-coenzyme A (CoA)
272
synthetase (ACS) for acetate assimilation, similarly as most obligate hydrogenotrophic
273
methanogens (40). This organism lacks genes encoding carbon monoxide dehydrogenase for the
274
acetyl-CoA biosynthesis from CO2 (30, 41). Two putative pathways for the conversion between
275
acetyl-CoA and pyruvate involved the pyruvate-ferredoxin oxidoreductase (Por) and the pyruvate
276
dehydrogenase (Pdh) complex; the latter was presumably expressed under oxic conditions (29). M.
277
conradii contains two acs copies. The transcript level was comparable between the two genes in
278
the monoculture (Table 2). The transcript level of acs2, however, was significantly
279
down-regulated 8- to10-fold in cocultures, while acs1 remained unaffected. The transcript
280
abundance of a gene coding for the membrane-bound inorganic pyrophosphatase was also
281
analyzed and showed no significant change among cultures (Table 2). Two por copies showed
282
similar transcript levels (Table 2) and did not change significantly in cocultures except an increase
283
of porA1 in butyrate coculture. 13
284
Growth on formate. To determine whether the strain could grow on formate in the presence of H2,
285
we performed a carbon isotope labelling experiment. [13C]-formate (99% 13C) was added to a final
286
concentration of 4 mM in the pure culture under H2/CO2 (4:1, 170 kPa). All of formate was
287
consumed in 6 days in parallel with the consumption of H2 (Fig. 2A). Meanwhile, the atomic 13C
288
ratio of CO2 increased rapidly in the first 4-5 days, and thereafter leveled off around 9% (Fig. 2B).
289
These data indicated that formate dehydrogenase catalyzed the conversion of formate to CO2 as:
290
Formate + F420 F420H2 + CO2. However, it had to be indicated that the rapid increase of 13CO2
291
could be resulted from the vigorous isotope exchange. The 13CO2 produced in this process was
292
then converted to 13CH4. Nevertheless, in total 38.5 μmol of 13CH4 were recovered in the
293
headspace at the end of incubation. If one molecule of CH4 was produced from 4 molecules of
294
formate according to the reaction stoichiometry (4 HCOOH CH4 + 3 CO2 + 2 H2O), the carbon
295
mass balance indicated that at least 77% of formate added was utilized for CH4 production. These
296
results evidenced that Fdh was functionally active.
297
298
DISCUSSION
299
Environmental studies have demonstrated that Methanocellales methanogens are adapted to low
300
H2 and syntrophic growth with fatty acids-oxidizing bacteria. To understand the adaptive
301
mechanisms, we determined the transcripts of 21 genes for methanogenesis and acetate
302
assimilation in M. conradii monoculture and syntrophic cocultures with P. thermopropionicum or
303
S. lipocalidus. We found that the expressions of three hydrogenase encoding genes (ech, frh and
304
mvh) and six methanogenesis genes (fwd, mtd, mer, mtr, mcr, hdr) were up-regulated (summarized
305
in Fig. 3), while the acetate assimilation gene (acs2) was down-regulated in cocultures. In addition, 14
306
the expression of formate dehydrogenase encoding gene (fdhA) was up-regulated in cocultures.
307
Apparently, M. conradii displayed a metabolic change, repressing the energy-consuming
308
biosynthesis while maintaining the energy-generating methanogenesis during syntrophic growth.
309
The methanogen possibly also activated the utilization of formate for the syntrophic interaction.
310
Among the genes up-regulated, the expressions of four genes fwd, hdr, ech and fdh showed
311
substantial increases (larger than 3-fold on average), while mer, mtr, mcr, mvh, and frh displayed
312
moderate increases (Fig. 3). Since the transcript level of 16S rRNA in syntrophic cocultures
313
decreased by a factor of about two compared with monoculture (Table 1), 2 to 3-fold increases of
314
the relative expression levels of the above genes indicated that M. conradii in cocultures
315
maintained nearly the same absolute expression levels of methanogenesis genes as in monoculture
316
per cell, though their growth rate significantly decreased. Genes coding for Fwd, Hdr and MvhD
317
in M. conradii are lined in a same large predicted transcription unit
318
(fwdGF/hdrC2B2A2/mvhD2/fwdCBAD). In hydrogenotrophic methanogens without cytochromes,
319
MvhADG and HdrABC form a complex (42, 43) that reduces heterodisulfide CoM-S-S-CoB in
320
conjunction with the reduction of the oxidized ferredoxin via the flavin-based electron bifurcation
321
(43, 44). The reduced ferredoxin is then used to reduce CO2 to formylmethanofuran, the first step
322
of methanogenesis. This coupling saves the Na+ motive force generated by Mtr available for ATP
323
synthesis. Costa and colleagues (45) showed that in M. maripaludis Hdr, Vhu, Fwd and Fdh
324
formed a supercomplex, proving that the first step and last step were physically connected. This
325
pointed to a full cycle of the methanogenic pathway, now referred to as the Wolfe cycle (46). The
326
presence of the large predicted transcription unit in M. conradii indicates that a complex
327
consisting of Mvh, Hdr and Fwd is likely to form. The marked up-regulation of hdr and fwd
15
328
transcript levels indicates the response to energy limitation imposed by low H2 or syntrophic
329
growth and suggesting that M. conradii tend to stabilize the Fwd-Hdr interaction under these
330
conditions. Given that electron bifurcation is crucial in the energy conservation of
331
hydrogenotrophic methanogens, the increase of the related gene expressions, and hence the
332
enzyme abundances will support an increased flux through this step.
333
The membrane-bound energy-converting hydrogenase is responsible for the reversible
334
reduction of ferredoxin with H2, driven by a proton or sodium ion motive force. There are two
335
types of energy-converting hydrogenases, namely Eha and Ehb, in M. maripaludis. Ehb is
336
responsible for anabolic ferredoxin synthesis (47, 48), while the function of Eha is probably
337
related to anaplerotically replenishing the intermediates of methanogenic pathway that is required
338
to sustain the Wolfe cycle (46, 49). M. conradii has only one Ech hydrogenase (encoded by
339
echABCDEF). Hence, its function is probably more close to methanogens with cytochromes like
340
Methanosarcina barkeri, generating ferredoxin for both biosynthesis and methanogenesis (50).
341
Since the growth rate of M. conradii was slower in the cocultures, it was unlikely that the
342
up-regulated Ech was due to the requirement of biosynthesis. Instead, the increased Ech very
343
possibly functioned like Eha in M. maripaludis to generate ferredoxin for methanogenesis and
344
hence stabilizing the Wolfe cycle (Fig. 3). The expressions of other methanogenesis genes
345
including frh, mer, mtr and mcr were moderately up-regulated during the syntrophic growth.
346
These enzymes would be all required for strengthening the central pathway of methanogenesis.
347
The formate dehydrogenase encoding genes were actively transcribed and markedly
348
up-regulated in cocultures. Formate is needed for the biosynthesis of purine (51). However, it was
349
unlikely that the significant up-regulation of fdh in M. conradii cocultures was for the need of 16
350
biosynthesis, as the growth was slower. By using carbon isotope labeling, we showed that M.
351
conradii could in fact convert formate to CH4 in the presence of H2, indicating that formate
352
dehydrogenase was functionally active for methanogenesis. The genomes of P.
353
thermopropionicum and S. lipocalidus encode both hydrogenases and formate dehydrogenases
354
(52-54). Thus, formate was possibly produced by the syntrophs and served as an alternative shuttle
355
for interspecies electron transfer between syntrophic partners. The up-regulation of Fdh in
356
methanogen therefore could facilitate the use of formate for methanogenesis and growth. Similar
357
up-regulation of Fdh under H2-limitation or syntrophic conditions was detected in M. maripaludis
358
(24). However, M. hungatei monoculture (on either formate or H2) and syntrophic coculture with S.
359
fumaroxidans showed no difference in the transcript levels of hydrogenase and formate
360
dehydrogenase encoding genes (25). The function of Fdh in hydrogenotrophic methanogens
361
therefore deserves further investigations.
362
Among acetate assimilation genes, one gene (acs2) coding for the synthesis of acetyl-CoA
363
from acetate was significantly down-regulated in cocultures while the other genes including acs1
364
remained unaffected. This result indicates that acs2 is more dynamic and possibly functionally
365
more important than acs1. Significant down regulation of acs genes was also observed in M.
366
thermautotrophicus and M. maripaludis (18, 24). Apparently, the lower growth rate of M. conradii
367
in cocultures was correlated with the down regulation of ACS that metabolized the first step of
368
acetate assimilation. The expression of acs might be also influenced by the acetate concentration
369
in the culture medium. Acetate was produced in the syntrophic cocultures to a much higher
370
concentration (Fig. 1B and C) than the amount added in the monoculture, which might exceed the
371
Km of ACS and elevate the efficiency of ACS. Consequently, less amount of ACS was needed,
17
372
373
resulting in the down-regulation of ACS. We have shown that M. conradii exhibited substantial changes of gene expressions associated
374
with the methanogenesis, acetate assimilation and formate utilization during the syntrophic growth.
375
We assumed that the major factor causing the shift of gene expressions was the H2 limitation. But
376
other factors could not be ruled out. Most obviously, the up-regulation of fdh and hence very likely
377
the activation of formate utilization have suggested that more complicated effect than H2
378
limitation alone has occurred during the syntrophic growth. Furthermore, it is also probable that
379
the interspecies transfers of other metabolites may happen that can influence the gene expressions.
380
The transfer of alanine has been identified in the syntrophic coculture of M. maripaludis with D.
381
vulgaris (24). Such kinds of metabolite transfers may diminish the pressure for biosynthesis in
382
methanogen and hence the down-regulation of the genes like acs. As with the effect of H2
383
limitation alone, previous studies on M. thermoformicicum (55), M. thermautotrophicus (19, 20,
384
22), M. maripaludis (17) and M. jannaschii (21) indicated that mainly the genes (frh, mtd, mer)
385
coding for enzymes catalyzing the oxidation and reduction of coenzyme F420 showed the
386
significant elevation of mRNA levels under H2 limitation. By comparison, the transcript levels of
387
these genes were only moderately up-regulated in the present study.
388
It has to be noted that batch cultivation was employed in the present study. The growth rate of
389
M. conradii decreased in cocultures and to a greater extent in propionate than butyrate coculture.
390
The difference between two syntrophic cocultures might be related to the thermodynamic
391
conditions. The higher H2 partial pressure in butyrate coculture probably supported a greater
392
growth rate for M. conradii in comparison with the propionate coculture (56-58). Growth rate was
393
believed to influence the gene expressions in both bacteria and archaea. The 16S rRNA expression 18
394
per cell in M. conradii monoculture was 1.8 and 2.4 times higher than in butyrate and propionate
395
cocultures respectively, consistent with the growth rate differences (Table 1).
396
The function of a few genes in M. conradii remains difficult to predict. For instance, we
397
detected a high transcript level of vhtG (Table 2 and Table S2) though no difference between
398
coculture and monoculture. M. conradii lacks methanophenazine (30). Thus, the function of Vht
399
remains unclear. We did not analyze the gene transcripts for the putative Ech-like hydrogenase.
400
We found, however, that the gene hdrB3 that is located in the predicted transcription unit of
401
ech-like genes (Fig. S1) was significantly up-regulated in cocultures (propionate coculture in
402
particular, Table 2). This implies that the Ech-like genes are possibly transcribed. Its function,
403
however, remains unknown. In addition, several enzymes like Frh, Mvh, and Hdr show multiple
404
copies. While one of copies was expressed usually to a relatively high level, others showed very
405
low expressions (orders of magnitude lower than the average). We found that the highly expressed
406
genes responded to the growth conditions substantially, while the lower expression copies did not
407
show changes. Apparently, these results imply the functioning differences of homologous genes in
408
M. conradii. The function and activity of multiple-copy genes deserve further investigations.
409
In conclusion, by determining the gene expression of M. conradii grown in monoculture and
410
syntrophically with P. thermopropionicum or S. lipocalidus, we illustrated that Methanocella
411
methanogens used different strategies to cope with low H2 and syntrophic growth conditions.
412
Firstly, they tend to strengthen the Wolfe cycle under the syntrophic conditions. M. conradii is
413
likely to form the Mvh/Hdr/Fwd complex. The significant up-regulation of Hdr and Fwd encoding
414
genes was in favor to maintain this complex. In addition, Ech that probably contributed to
415
replenish the methanogenesis intermediates was significantly up-regulated during the syntrophic 19
416
growth. The expression of other genes within the Wolfe cycle was also moderately upregulated.
417
All these results point to a strategy of M. conradii to reinforce the Wolfe cycle during the
418
syntrophic growth. Secondly, M. conradii appeared to down-regulate the energy-demanding
419
biosynthesis by repressing the acetate assimilation. Thirdly, M. conradii probably activated the
420
formate-dependent methanogenesis during the syntrophic growth. In conclusion, we propose that
421
maintaining the integration of the Wolfe cycle might be crucial for the methanogen to cope with
422
low H2 and syntrophic growth conditions that prevail in nature. It remains, however, unknown
423
how methanogens initiate these metabolic shifts in response to environmental changes.
424
425
426
ACKNOWLEDGMENTS
427
This study was financially supported by the National Basic Research Program of China
428
(2011CB100505) and the National Natural Science Foundation of China (41130527, 40625003).
429 430
20
431
REFERENCES
432
1. Groβkopf R, Stubner S and Liesack W. 1998. Novel euryarchaeotal lineages detected on rice roots
433 434 435
and in the anoxic bulk soil of flooded rice microcosms. Appl. Environ. Microb. 64:4983-4989. 2. Fey A, Chin K and Conrad R. 2001. Thermophilic methanogens in rice field soil. Environ. Microbiol. 3:295-303.
436
3. Lueders T, Chin K, Conrad R and Friedrich M. 2001. Molecular analyses of methyl-coenzyme M
437
reductase α-subunit (mcrA) genes in rice field soil and enrichment cultures reveal the methanogenic
438
phenotype of a novel archaeal lineage. Environ. Microbiol. 3:194-204.
439 440 441 442 443 444 445
4. Angel R, Claus P and Conrad R. 2012. Methanogenic archaea are globally ubiquitous in aerated soils and become active under wet anoxic conditions. ISME J. 6:847-862. 5. Angel R, Matthies D and Conrad R. 2011. Activation of methanogenesis in arid biological soil crusts despite the presence of oxygen. Plos One 6:e20453. 6. Conrad R, Erkel C and Liesack W. 2006. Rice Cluster I methanogens, an important group of Archaea producing greenhouse gas in soil. Curr. Opin. Biotech. 17:262-267. 7. Krüger M, Frenzel P, Kemnitz D and Conrad R. 2005. Activity, structure and dynamics of the
446
methanogenic archaeal community in a flooded Italian rice field. FEMS Microbiol. Ecol.
447
51:323-331.
448 449 450 451 452
8. Lu YH, Lueders T, Friedrich MW and Conrad R. 2005. Detecting active methanogenic populations on rice roots using stable isotope probing. Environ. Microbiol. 7:326-336. 9. Lu YH and Conrad R. 2005. In situ stable isotope probing of methanogenic archaea in the rice rhizosphere. Science 309:1088-1090. 10. Gan Y, Qiu Q, Liu P, Rui J and Lu Y. 2012. Syntrophic oxidation of propionate in rice field soil
21
453 454 455 456 457 458 459
at 15 and 30 °C under methanogenic conditions. Appl. Environ. Microb. 78:4923-4932. 11. Liu FH and Conrad R. 2010. Thermoanaerobacteriaceae oxidize acetate in methanogenic rice field soil at 50 ºC. Environ. Microbiol. 12:2341-2354. 12. Liu P, Qiu Q and Lu Y. 2011. Syntrophomonadaceae-affiliated species as active butyrate-utilizing syntrophs in paddy field soil. Appl. Environ. Microb. 77:3884-3887. 13. Rui J, Qiu Q and Lu Y. 2011. Syntrophic acetate oxidation under thermophilic methanogenic condition in Chinese paddy field soil. FEMS Microbiol. Ecol. 77:264-273.
460
14. Lueders T, Pommerenke B and Friedrich MW. 2004. Stable-isotope probing of microorganisms
461
thriving at thermodynamic limits: syntrophic propionate oxidation in flooded soil. Appl. Environ.
462
Microb. 70:5778-5786.
463
15. McInerney MJ, Struchtemeyer CG, Sieber J, Mouttaki H, Stams AJ, Schink B, Rohlin L and
464
Gunsalus RP. 2008. Physiology, ecology, phylogeny, and genomics of microorganisms capable of
465
syntrophic metabolism. Ann. Ny. Acad. Sci. 1125:58-72.
466
16. Luo H, Zhang H, Suzuki T, Hattori S and Kamagata Y. 2002. Differential expression of
467
methanogenesis genes of Methanothermobacter thermoautotrophicus (formerly Methanobacterium
468
thermoautotrophicum) in pure culture and in cocultures with fatty acid-oxidizing syntrophs. Appl.
469
Environ. Microb. 68:1173-1179.
470
17. Hendrickson E, Haydock A, Moore B, Whitman W and Leigh J. 2007. Functionally distinct
471
genes regulated by hydrogen limitation and growth rate in methanogenic Archaea. P. Natl. Acad. Sci.
472
USA 104:8930-8934.
473
18. Enoki M, Shinzato N, Sato H, Nakamura K and Kamagata Y. 2011. Comparative proteomic
474
analysis of Methanothermobacter themautotrophicus ΔH in pure culture and in co-culture with a
22
475
butyrate-oxidizing bacterium. Plos One 6:e24309.
476
19. Kato S, Kosaka T and Watanabe K. 2008. Comparative transcriptome analysis of responses of
477
Methanothermobacter thermautotrophicus to different environmental stimuli. Environ. Microbiol.
478
10:893-905.
479
20. Morgan R, Pihl T, Nolling J and Reeve J. 1997. Hydrogen regulation of growth, growth yields,
480
and methane gene transcription in Methanobacterium thermoautotrophicum deltaH. J. Bacteriol.
481
179:889-898.
482
21. Mukhopadhyay B, Johnson EF and Wolfe RS. 2000. A novel pH2 control on the expression of
483
flagella in the hyperthermophilic strictly hydrogenotrophic methanarchaeaon Methanococcus
484
jannaschii. P. Natl. Acad. Sci. USA 97:11522-11527.
485 486
22. Reeve JN, Nolling J, Morgan RM and Smith DR. 1997. Methanogenesis: genes, genomes, and who's on first? J. Bacteriol. 179:5975-5986.
487
23. Shinzato N, Enoki M, Sato H, Nakamura K, Matsui T and Kamagata Y. 2008. Specific DNA
488
binding of a potential transcriptional regulator, inosine 5′-monophosphate dehydrogenase-related
489
protein VII, to the promoter region of a methyl coenzyme M reductase I-encoding operon retrieved
490
from Methanothermobacter thermautotrophicus strain ΔH. Appl. Environ. Microb. 74:6239-6247.
491
24. Walker CB, Redding-Johanson AM, Baidoo EE, Rajeev L, He Z, Hendrickson EL,
492
Joachimiak MP, Stolyar S, Arkin AP, Leigh JA, Zhou J, Keasling JD, Mukhopadhyay A and
493
Stahl DA. 2012. Functional responses of methanogenic archaea to syntrophic growth. ISME J.
494
6:2045-2055.
495
25. Worm P, Stams AJ, Cheng X and Plugge CM. 2011. Growth- and substrate-dependent
496
transcription of formate dehydrogenase and hydrogenase coding genes in Syntrophobacter
23
497
fumaroxidans and Methanospirillum hungatei. Microbiology 157:280-289.
498
26. Sakai S, Imachi H, Sekiguchi Y, Ohashi A, Harada H and Kamagata Y. 2007. Isolation of key
499
methanogens for global methane emission from rice paddy fields: a novel isolate affiliated with the
500
clone cluster rice cluster I. Appl. Environ. Microb. 73:4326-4331.
501 502 503
27. Lü Z and Lu Y. 2012. Methanocella conradii sp. nov., a thermophilic, obligate hydrogenotrophic methanogen, isolated from Chinese rice field soil. Plos One 7:e35279. 28. Sakai S, Conrad R, Liesack W and Imachi H. 2010. Methanocella arvoryzae sp. nov., a
504
hydrogenotrophic methanogen isolated from rice field soil. Int. J. Syst. Evol. Micr. 60:2918-2923.
505
29. Erkel C, Kube M, Reinhardt R and Liesack W. 2006. Genome of Rice Cluster I archaea--the key
506 507 508
methane producers in the rice rhizosphere. Science 313:370-372. 30. Lü Z and Lu Y. 2012. Complete genome sequence of a thermophilic methanogen, Methanocella conradii HZ254, isolated from Chinese rice field soil. J. Bacteriol. 194:2398-2399.
509
31. Sakai S, Takaki Y, Shimamura S, Sekine M, Tajima T, Kosugi H, Ichikawa N, Tasumi E,
510
Hiraki AT, Shimizu A, Kato Y, Nishiko R, Mori K, Fujita N, Imachi H and Takai K. 2011.
511
Genome sequence of a mesophilic hydrogenotrophic methanogen Methanocella paludicola, the first
512
cultivated representative of the order Methanocellales. Plos One 6:e22898.
513
32. Stams AJM and Dong XZ. 1995. Role of formate and hydrogen in the degradation of propionate
514
and butyrate by defined suspended cocultures of acetogenic and methanogenic bacteria. Anton.
515
Leeuw. Int. J. G. 68:281-284.
516
33. Thiele JH and Zeikus JG. 1988. Control of interspecies electron flow during anaerobic digestion:
517
significance of formate transfer versus hydrogen transfer during syntrophic methanogenesis in flocs.
518
Appl. Environ. Microb. 54:20-29.
24
519 520 521
34. Yuan QA and Lu YH. 2009. Response of methanogenic archaeal community to nitrate addition in rice field soil. Environ. Microbiol. Rep. 1:362-369. 35. Culley D, Kovacikjr W, Brockman F and Zhang W. 2006. Optimization of RNA isolation from
522
the archaebacterium Methanosarcina barkeri and validation for oligonucleotide microarray analysis.
523
J. Microbiol. Meth. 67:36-43.
524 525
36. Peirson SN, Butler JN and Foster RG. 2003. Experimental validation of novel and conventional approaches to quantitative real-time PCR data analysis. Nucleic. Acids. Res. 31:e73.
526
37. Hendrickson EL, Liu Y, Rosas-Sandoval G, Porat I, Söll D, Whitman WB and Leigh JA. 2008.
527
Global responses of Methanococcus Maripaludis to specific nutrient limitations and growth rate. J.
528
Bacteriol. 190:2198-2205.
529
38. Kemnitz D, Kolb S and Conrad R. 2005. Phenotypic characterization of Rice Cluster III archaea
530
without prior isolation by applying quantitative polymerase chain reaction to an enrichment culture.
531
Environ. Microbiol. 7:553-565.
532 533 534 535 536 537 538
39. Ma K, Conrad R and Lu Y. 2012. Responses of methanogen mcrA genes and their transcripts to an alternate dry/wet cycle of paddy field soil. Appl. Environ. Microb. 78:445-454. 40. Oberlies G, Fuchs G and Thauer R. 1980. Acetate thiokinase and the assimilation of acetate in Methanobacterium thermoautotrophicum. Arch. Microbiol. 128:248-252. 41. Shieh J and Whitman WB. 1988. Autotrophic acetyl coenzyme A biosynthesis in Methanococcus Maripaludis. J. Bacteriol. 170:3072-3079. 42. Buckel W and Thauer RK. 2013. Energy conservation via electron bifurcating ferredoxin
539
reduction and proton/Na+ translocating ferredoxin oxidation. Biochim. Biophys. Acta 1827:94-113.
540
43. Thauer RK, Kaster AK, Seedorf H, Buckel W and Hedderich R. 2008. Methanogenic archaea:
25
541
ecologically relevant differences in energy conservation. Nat. Rev. Microbiol. 6:579-591.
542
44. Kaster AK, Moll J, Parey K and Thauer RK. 2011. Coupling of ferredoxin and heterodisulfide
543
reduction via electron bifurcation in hydrogenotrophic methanogenic archaea. P. Natl. Acad. Sci.
544
USA 108:2981-2986.
545
45. Costa KC, Wong PM, Wang T, Lie TJ, Dodsworth JA, Swanson I, Burn JA, Hackett M and
546
Leigh JA. 2010. Protein complexing in a methanogen suggests electron bifurcation and electron
547
delivery from formate to heterodisulfide reductase. P. Natl. Acad. Sci. USA 107:11050-11055.
548
46. Thauer RK. 2012. The Wolfe cycle comes full circle. P. Natl. Acad. Sci. USA 109:15084-15085.
549
47. Major TA, Liu YC and Whitman WB. 2010. Characterization of Energy-Conserving
550
Hydrogenase B in Methanococcus maripaludis. J. Bacteriol. 192:4022-4030.
551
48. Porat I, Kim W, Hendrickson EL, Xia Q, Zhang Y, Wang T, Taub F, Moore BC, Anderson IJ
552
and Hackett M. 2006. Disruption of the operon encoding Ehb hydrogenase limits anabolic CO2
553
assimilation in the archaeon Methanococcus maripaludis. J. Bacteriol. 188:1373-1380.
554
49. Lie TJ, Costa KC, Lupa B, Korpole S, Whitman WB and Leigh JA. 2012. Essential anaplerotic
555
role for the energy-converting hydrogenase Eha in hydrogenotrophic methanogenesis. P. Natl. Acad.
556
Sci. USA 109:15473-15478.
557
50. Meuer J, Kuettner HC, Zhang JK, Hedderich R and Metcalf WW. 2002. Genetic analysis of
558
the archaeon Methanosarcina barkeri Fusaro reveals a central role for Ech hydrogenase and
559
ferredoxin in methanogenesis and carbon fixation. P. Natl. Acad. Sci. USA 99:5632-5637.
560 561 562
51. White RH. 1997. Purine biosynthesis in the domain Archaea without folates or modified folates. J. Bacteriol. 179:3374-3377. 52. Müller N, Worm P, Schink B, Stams AJM and Plugge CM. 2010. Syntrophic butyrate and
26
563
propionate oxidation processes: from genomes to reaction mechanisms. Environ. Microbiol. Rep.
564
2:489-499.
565 566
53. Sieber JR, McInerney MJ and Gunsalus RP. 2012. Genomic insights into syntrophy: The paradigm for anaerobic metabolic cooperation. Annu. Rev. Microbiol. 66:429-452.
567
54. Sieber JR, Sims DR, Han C, Kim E, Lykidis A, Lapidus AL, McDonnald E, Rohlin L, Culley
568
DE, Gunsalus R and McInerney MJ. 2010. The genome of Syntrophomonas wolfei: new insights
569
into syntrophic metabolism and biohydrogen production. Environ. Microbiol. 12:2289-2301.
570
55. Nolling J and Reeve JN. 1997. Growth- and substrate-dependent transcription of the formate
571
dehydrogenase (fdhCAB) operon in Methanobacterium thermoformicicum Z-245. J. Bacteriol.
572
179:899-908.
573
56. Ishii S, Kosaka T, Hori K, Hotta Y and Watanabe K. 2005. Coaggregation facilitates
574
interspecies hydrogen transfer between Pelotomaculum thermopropionicum and
575
Methanothermobacter thermautotrophicus. Appl. Environ. Microb. 71:7838-7845.
576
57. Kato S, Kosaka T and Watanabe K. 2009. Substrate-dependent transcriptomic shifts in
577
Pelotomaculum thermopropionicum grown in syntrophic co-culture with Methanothermobacter
578
thermautotrophicus. Microbial Biotech. 2:575-584.
579 580
58. Schink B. 1997. Energetics of syntrophic cooperation in methanogenic degradation. Microbiol. Mol. Biol. R. 61:262-280.
581 582
27
583
TABLE 1 Numbers of 16S rRNA transcripts per cell in M. conradii growing in monoculture and
584
syntrophic cocultures with P. thermopropionicum or S. lipocalidus. Culture
Transcript number per cell
M MB1 MB2 MP1 MP2
509.5±87.6 a 294.2±54.7 b 252.7±94.3 b 195.1±106 b 216.3±35.1 b
585
Note:
586
M, monoculture; MB and MP, syntrophic butyrate and propionate cocultures, respectively; the
587
number after culture (1 and 2) indicates the early- and mid-exponential phases, respectively.
588
Values are means of four biological replicates (three for MP1 and MP2) for each sample,
589
means±SD are given. Different letters after the transcript number indicate significant differences
590
at the level of p
