Anaerobic Methane Oxidation Driven by Microbial Reduction of Natural Organic Matter in a Tropical Wetland

Abstract

Wetlands constitute the main natural source of methane on Earth due to their high content of natural organic matter (NOM), but key drivers, such as electron acceptors, supporting methanotrophic activities in these habitats are poorly understood. We performed anoxic incubations using freshly collected sediment, along with water samples harvested from a tropical wetland, amended with ¹³C-methane (0.67 atm) to test the capacity of its microbial community to perform anaerobic oxidation of methane (AOM) linked to the reduction of the humic fraction of its NOM. Collected evidence demonstrates that electron-accepting functional groups (e.g., quinones) present in NOM fueled AOM by serving as a terminal electron acceptor. Indeed, while sulfate reduction was the predominant process, accounting for up to 42.5% of the AOM activities, the microbial reduction of NOM concomitantly occurred. Furthermore, enrichment of wetland sediment with external NOM provided a complementary electron-accepting capacity, of which reduction accounted for ~100 nmol ¹³CH4 oxidized · cm⁻³ · day⁻¹. Spectroscopic evidence showed that quinone moieties were heterogeneously distributed in the wetland sediment, and their reduction occurred during the course of AOM. Moreover, an enrichment derived from wetland sediments performing AOM linked to NOM reduction stoichiometrically oxidized methane coupled to the reduction of the humic analogue anthraquinone-2,6- disulfonate. Microbial populations potentially involved in AOM coupled to microbial reduction of NOM were dominated by divergent biota from putative AOMassociated archaea. We estimate that this microbial process potentially contributes to the suppression of up to 114 teragrams (Tg) of CH4 · year⁻¹ in coastal wetlands and more than 1,300 Tg · year⁻¹, considering the global wetland area.

Full text

Anaerobic Methane Oxidation Driven by
Microbial Reduction of Natural Organic
Matter in a Tropical Wetland
Edgardo I. Valenzuela,
a
Alejandra Prieto-Davó,
b
Nguyen E. López-Lozano,
a
Alberto Hernández-Eligio,
c
Leticia Vega-Alvarado,
d
Katy Juárez,
c
Ana Sarahí García-González,
e
Mercedes G. López,
e
Francisco J. Cervantes
a
División de Ciencias Ambientales, Instituto Potosino de Investigación Científica y Tecnológica, San Luis Potosí,
México
a
; Facultad de Química, Unidad Sisal, Universidad Nacional Autónoma de México, Sisal, Yucatán,
México
b
; Departamento de Ingeniería Celular y Biocatálisis, Instituto de Biotecnología, Universidad Nacional
Autónoma de México, Campus Morelos, Cuernavaca, Morelos, México
c
; Centro de Ciencias Aplicadas y
Desarrollo Tecnológico, Universidad Nacional Autónoma de México, Ciudad Universitaria, Coyoacán, Distrito
Federal, México
d
; Departamento de Biotecnología y Bioquímica, Centro de Investigación y de Estudios
Avanzados del IPN, Unidad Irapuato, Irapuato, México
e
ABSTRACT Wetlands constitute the main natural source of methane on Earth due
to their high content of natural organic matter (NOM), but key drivers, such as elec-
tron acceptors, supporting methanotrophic activities in these habitats are poorly un-
derstood. We performed anoxic incubations using freshly collected sediment, along
with water samples harvested from a tropical wetland, amended with
13
C-methane
(0.67 atm) to test the capacity of its microbial community to perform anaerobic oxi-
dation of methane (AOM) linked to the reduction of the humic fraction of its NOM.
Collected evidence demonstrates that electron-accepting functional groups (e.g., qui-
nones) present in NOM fueled AOM by serving as a terminal electron acceptor. In-
deed, while sulfate reduction was the predominant process, accounting for up to
42.5% of the AOM activities, the microbial reduction of NOM concomitantly oc-
curred. Furthermore, enrichment of wetland sediment with external NOM provided a
complementary electron-accepting capacity, of which reduction accounted for ⬃100
nmol
13
CH
4
oxidized · cm
⫺3
· day
⫺1
. Spectroscopic evidence showed that quinone
moieties were heterogeneously distributed in the wetland sediment, and their re-
duction occurred during the course of AOM. Moreover, an enrichment derived from
wetland sediments performing AOM linked to NOM reduction stoichiometrically oxi-
dized methane coupled to the reduction of the humic analogue anthraquinone-2,6-
disulfonate. Microbial populations potentially involved in AOM coupled to microbial
reduction of NOM were dominated by divergent biota from putative AOM-
associated archaea. We estimate that this microbial process potentially contributes
to the suppression of up to 114 teragrams (Tg) of CH
4
· year
⫺1
in coastal wetlands
and more than 1,300 Tg · year
⫺1
, considering the global wetland area.
IMPORTANCE The identification of key processes governing methane emissions
from natural systems is of major importance considering the global warming effects
triggered by this greenhouse gas. Anaerobic oxidation of methane (AOM) coupled to
the microbial reduction of distinct electron acceptors plays a pivotal role in mitigat-
ing methane emissions from ecosystems. Given their high organic content, wetlands
constitute the largest natural source of atmospheric methane. Nevertheless, pro-
cesses controlling methane emissions in these environments are poorly understood.
Here, we provide tracer analysis with
13
CH
4
and spectroscopic evidence revealing
that AOM linked to the microbial reduction of redox functional groups in natural or-
ganic matter (NOM) prevails in a tropical wetland. We suggest that microbial reduc-
tion of NOM may largely contribute to the suppression of methane emissions from
Received 16 March 2017 Accepted 18 March
2017
Accepted manuscript posted online 24
March 2017
Citation Valenzuela EI, Prieto-Davó A, López-
Lozano NE, Hernández-Eligio A, Vega-Alvarado
L, Juárez K, García-González AS, López MG,
Cervantes FJ. 2017. Anaerobic methane
oxidation driven by microbial reduction of
natural organic matter in a tropical wetland.
Appl Environ Microbiol 83:e00645-17. https://
doi.org/10.1128/AEM.00645-17.
Editor Frank E. Löffler, University of Tennessee
and Oak Ridge National Laboratory
Copyright © 2017 American Society for
Microbiology. All Rights Reserved.
Address correspondence to Francisco J.
Cervantes, fjcervantes@ipicyt.edu.mx.
ENVIRONMENTAL MICROBIOLOGY
crossm
June 2017 Volume 83 Issue 11 e00645-17 aem.asm.org 1Applied and Environmental Microbiology

tropical wetlands. This is a novel avenue within the carbon cycle in which slowly de-
caying NOM (e.g., humic fraction) in organotrophic environments fuels AOM by serv-
ing as a terminal electron acceptor.
KEYWORDS anaerobic methane oxidation, humus, methanotrophy, wetlands
Microbial processes produce and consume methane (CH
4
) in anoxic sediments,
playing a crucial role in regulating Earth’s climate. Virtually 90% of the CH
4
produced from marine environments is oxidized by microorganisms, preventing its
release into the atmosphere (1). Anaerobic oxidation of methane (AOM) associated with
sulfate reduction was first discovered in marine environments (2). More recently, AOM
has also been linked to the microbial reduction of nitrate (3, 4) and nitrite (5), as well
as Fe(III) and Mn(IV) oxides (6–8), in freshwater and marine environments. Wetlands are
the largest natural source of CH
4
(9), contributing to about one-third of global emis-
sions (10), but key drivers, such as electron acceptors fueling methanotrophic activities
in these habitats, are poorly understood. CH
4
emissions from wetlands have been
strongly responsive to climate in the past and will likely continue to be responsive to
anthropogenic-driven climate change in the future, predicting a large impact on global
atmospheric CH
4
concentration (10). The traditional assumption is that aerobic metha-
notrophy dominates CH
4
cycling in wetlands by oxidizing an estimated 40 to 70% of
the gross CH
4
production in these ecosystems (11). Recent findings (12) challenged this
conjecture by providing evidence that AOM may consume up to 200 Tg of CH
4
· year
⫺1
,
decreasing their potential CH
4
emission by 50% in these habitats. Most AOM activities
observed in wetlands have been related to sulfate reduction (12, 13), but other electron
acceptors remain feasible. Natural organic matter (NOM), circumscribed to humic
substances (HS) in many studies (14), occurs at high concentrations in wetlands in both
soluble and solid phases (15). Recent evidence indicates that HS suppress methane
production in different ecosystems (16, 17), yet the mechanisms involved are still
enigmatic. HS can theoretically promote AOM, as they can serve as terminal electron
acceptors for microbial respiration (18, 19) and have a higher redox potential than
sulfate (20). However, compelling evidence demonstrating AOM driven by the microbial
reduction of NOM present in anoxic environments remains elusive (21, 22).
We aimed to document
13
CH
4
anaerobic oxidation and the ongoing reduction of
intrinsic electron acceptors, including the electron-accepting fraction of NOM, by the
biota of freshly sampled sediment from a coastal tropical wetland. We provide
13
CH
4
tracer studies and spectroscopic evidence demonstrating that AOM is linked to the
microbial reduction of redox functional groups present in the NOM of this tropical
marsh. Furthermore, we found evidence, based on 16S rRNA gene sequences, indicat-
ing that microbial populations potentially involved in AOM coupled to microbial
reduction of NOM were dominated by divergent biota from putative AOM-associated
microorganisms.
RESULTS
Kinetics of
13
C-methane oxidation and electron balances. The exponential phase
of AOM was observed in microcosms over the first 15 days of incubation in cases with
unamended sediment (i.e., sediment without external NOM addition). The methanotrophic
rate in this experimental treatment was ⬃1.34
␮
mol
13
C-methane oxidized · cm
⫺3
·
day
⫺1
(Fig. 1). At the end of the exponential phase, sulfate and Fe(III) reduction
accounted for 42.5% and 0.5% of the
13
C-methane oxidized, respectively, while the role
of nitrate was marginal (Fig. 2 and Table S2). These unamended sediment microcosms
exhibited a reduction in intrinsic NOM during the course of AOM, which was expected
due to the high concentration of organic carbon in the tropical wetland, with the
capacity to accept electrons (Table S1 in the supplemental material and Fig. 2).
Nevertheless, large perturbation caused by endogenous NOM reduction in experimen-
tal controls lacking
13
C-methane obstructed an accurate assessment of AOM driven by
this microbial process (Fig. 2). The large endogenous NOM reduction observed in these
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control experiments may be explained by the concomitant methane production (and
subsequent consumption) observed (Fig. S2) and by oxidation of labile organic matter
present in the sediment (Table S1). Supplementary incubations spiked with the sulfate
reduction inhibitor molybdate (25 mmol liter
⫺1
) showed decreased sulfate-reducing
activity (⬃50%, Fig. 2), while AOM rates remained high compared against their non-
inhibited counterparts (Fig. 1). Remarkably, when sulfate reduction was inhibited, the
reduction of intrinsic NOM was doubled (from 1.6 ⫾0.11 to 3.4 ⫾0.19 millielectron
equivalents [meq] · liter
⫺1
), implying that the reduction of redox functional groups in
NOM was promoted when the utilization of sulfate was impeded.
Further enrichment of wetland sediment with external NOM, in the form of HS
derived from Pahokee peat (Florida Everglades, 2.5 g · liter
⫺1
), provided complementary
electron-accepting capacity, which significantly elicited AOM up to ⬃1.88
␮
mol
13
C-
methane oxidized · cm
⫺3
· day
⫺1
and extended the exponential phase to 20 days (Fig.
1). In this experimental treatment, electron balances revealed methanotrophic activity
responsible for ⬃100 nmol
13
CH
4
oxidized · cm
⫺3
· day
⫺1
linked to microbial reduction
of NOM (including both intrinsic and externally added as Pahokee peat HS). As
hypothesized before, the consumption of intrinsically produced methane was con-
firmed by experimental controls enriched with HS from Pahokee peat and incubated in
FIG 1 Anaerobic methane oxidation measured as
13
CO
2
production in microcosms’ headspace and
13
C enrichment (inset) calculated
as
13
F
CO2
⫽(
13
CO
2
/[
13
CO
2
⫹
12
CO
2
]). (a) Kinetics for incubations performed with unamended sediment. (b) Kinetics for incubations
performed with sediment enriched with 2.5 g · liter
⫺1
of external NOM in the form of Pahokee peat humic substances. Error bars
represent the standard error among replicates (n⫽4, or 3*). SR-INH stands for sediment incubations performed with molybdate (25
mmol liter
⫺1
) in order to inhibit sulfate reduction.
13
CO
2
production rates were based on the maximum slope observed on linear
regressions considering at least three sampling points.
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the absence of
13
C-methane, which showed significant consumption of
12
CH
4
(Fig. S2).
This was also confirmed by an increase in quantified
12
CO
2
production, which was
reflected in the 2- to 4-fold-lower enrichment of
13
CO
2
in HS-enriched incubations
compared to unamended controls (see
13
F
CO2
values in Fig. 1). Reports (23, 24) indicate
that methanotrophic microorganisms prefer to oxidize
12
CH
4
compared to
13
CH
4
, which
may partly explain our findings.
The role of sulfate reduction on AOM when wetland sediment was enriched with HS
was not possible to assess (Table S2) due to the large degree of endogenous sulfate
reduction elicited by the degradation of the labile fraction of externally added NOM
(Fig. 2), which also triggered methanogenesis in these microcosms. Since no significant
differences in iron reduction were detected between microcosms with or without
added
13
CH
4
, the only microbial process clearly identified as driving AOM in Pahokee
peat-enriched sediments was the microbial reduction of HS (Table S2).
Spectroscopic evidence on the presence and reduction of redox functional
groups in NOM. Initial exploration of the solid-phase NOM present in wetland sedi-
ment by microattenuated total reflection-Fourier transform infrared (micro-ATR-FTIR)
spectra revealed the presence of electron-accepting moieties in both unamended and
HS-enriched wetland sediments. By mapping of acquisition points at 1,650 to 1,620 · cm
⫺1
,
the presence and heterogeneous distribution of quinone functional groups were evi-
denced in sediments, confirming the presence of nonsoluble electron-accepting moi-
eties classically attributed to humic-like materials (Fig. 3a and b). To further confirm this,
FIG 2 Production of
13
CO
2
and reduction of intrinsic or added electron acceptors at the end of the exponential phase (20
days of incubation) in the absence (a and b) and in the presence (c and d) of external NOM as HS from Pahokee peat.
SR-INH stands for controls amended with the sulfate reduction inhibitor molybdate (25 mmol liter
⫺1
). Error bars represent
the standard error among replicates.
13
CO
2
produced was measured as described for Fig. 1. Quantification of sulfate and
nitrate reduction implies a decrease in their concentration at this sampling time, whereas Fe(III) reduction was quantified
in terms of the ferrous iron produced. Reduction of NOM and HS was determined by the ferrozine technique.
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we looked for double-bonded carbon and oxygen (C⫽O) by use of X-ray photoelectron
spectra (XPS); this technique supported the existence of quinone-like functional groups
in unamended sediment and furthermore provided evidence of the reduction of these
moieties by showing the disappearance of the C⫽O signal from C1s and O1s high-
resolution spectra in a comparison of signals from sediment analyzed before and after
incubation with
13
CH
4
in the absence of external HS (Fig. 3c to f). Another missing
signal after the AOM process was that which corresponds to metallic oxides, evidenced
by an analysis of the O1s high-resolution spectra (Fig. 3d and f), which may imply the
reduction of intrinsic iron oxides that supported ⬃0.5% of methanotrophy, according
to electron balances (Table S2). Further analysis of the liquid phase of pristine sediment
microcosms also revealed the reduction of quinone-like moieties during the course of
AOM (Fig. 4). Initial samples exhibited a well-defined and strong peak at 1,690 · cm
⫺1
associated with quinone moieties, while reduced samples, at the end of the incubation
FIG 3 Spectroscopic evidence of the presence of quinone moieties and their reduction in wetland sediment
samples. (a and b) Micro-ATR-FTIR representative spectra taken from imaged areas generated after processing
quinone functional groups (1,650 to 1,620 · cm
⫺1
) of sediment samples before incubation in the absence (a) and
in the presence (b) of external NOM in the form of Pahokee peat HS. (c and e) XPS high-resolution profiles of C1s.
(d and f) O1s signal. (c to f) Sediment samples prior to incubation (c and d) and sediment samples after incubation
(e and f) with
13
C-methane. Regions and components were corrected at 284.8 eV for the COC adventitious carbon
A, B and G components belong to COO bond (⬃286.6 and ⬃532 eV, respectively), C and H correspond to C⫽O
functional group (⬃288.9 and ⬃533.3 eV, respectively), D belongs to –COOH (⬃289.6 eV), E is typical of the
presence of carbonate (⬃291 eV), and F suggests the occurrence of a metallic oxide (⬃530 eV). a.u., absorbance
units.
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period, showed an increase in the signal related to phenolic groups (1,660 · cm
⫺1
).
Additional signals of phenolic groups were detected after incubation with
13
CH
4
and
Pahokee peat by spectral signals detected around 2,260 to 2,500 · cm
⫺1
(25).
Microbial communities performing AOM. According to 16S rRNA gene sequences
from wetland sediment samples performing AOM, anaerobic methanotrophic archaea
(ANME), which are traditionally linked to anaerobic methanotrophy under sulfate-
reducing (2, 26), Fe(III)-reducing (6, 8), and artificial electron acceptor-reducing condi-
tions (27), were barely detected in our experiments, with ANME-1b and ANME-3
representing less than 0.5% and 0.2%, respectively, from the archaeal community in all
experimental treatments (Fig. 5). The only abundant Euryarchaeota members detected
were affiliated with an unclassified genus of the marine benthic group D (MBG-D) family
(deep hydrothermal vent euryarchaeotal group 1 [DHVEG-1]), which accounted for 18
to 23% of the archaeal biota in all treatments. Outside the Euryarchaeota phylum,
members from the newly named Bathyarchaeota lineage (formerly known as Miscella-
neous Crenarchaeotic Group) were another cluster of microorganisms that remained in
high percentages (from 8 to 14%) in all treatments. Two genera from the Thaumar-
chaeota phylum, one belonging to the pMC2A209 class, and the other from marine
benthic group B (MBG-B), were also consistently present in all sediment samples
showing AOM, with the MBG-B genus increasing its proportion up to 12% when sulfate
reduction was inhibited (Fig. 5). From the bacterial counterpart, the most abundant
bacteria in two of the treatments were from the genus Oceanimonas from the Aero-
monadaceae family (Gammaproteobacteria), whose presence was diminished when
sulfate reduction was inhibited and when
13
CH
4
was absent (Fig. S3), suggesting that
this microorganism might have been involved in sulfate-dependent AOM. Other evi-
dent changes in the bacterial community included the increase in Clostridia and Bacilli
members when external NOM was supplied (Fig. S3), which agrees with their capacity
to reduce HS (28).
FIG 4 High-performance UV-visible-near infrared spectra obtained from liquid samples before and after
incubation with
13
CH
4
. (a and c) Spectra obtained before incubation with
13
CH
4
. (b and d) Spectra
obtained after incubation with
13
CH
4
.
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AOM linked to AQDS reduction. In order to confirm the capacity of the sediment
biota to channel
13
C-methane-derived electrons to quinone groups, the humic analogue
anthraquinone-2,6-disulfonate (AQDS) was added as an electron acceptor to the artificial
basal medium for sediment enrichments. AQDS reduction and methane consumption
were observed since the first enrichment cycle, although no clear relationship between
net methane consumption and anthrahydroquinone-2,6-disulfonate (AH
2
QDS) produc-
tion was observed due to high concentrations of intrinsic electron donors and accep-
tors (data not shown). Nevertheless, during the third incubation cycle, net AOM was
observed within 11 days, which corresponded to a final ratio of oxidized methane to
reduced AQDS of 1:4.7, corrected for endogenous controls, which is very close to the
stoichiometric 1:4 ratio, according to the following equation (Fig. 6a):
CH4⫹4AQDS ⫹2H2O→CO2⫹4AH2QDS
Gibbs free energy (⌬G°')⫽⫺43.2 kJ mol⫺1
Analysis of 16S rRNA gene sequences from enriched sediment sampled at the end
of the third cycle of AQDS-dependent AOM activity (Fig. S1) displayed a significant
decrease in the diversity of the microbial community, evidenced by a decrease in
Shannon index, from 5.52 in freshly sampled sediment to 3.56 after enrichment with
CH
4
and AQDS. Significant increments and decreases in specific groups of archaea and
bacteria did occur in this enrichment (Fig. 6b and c). From the archaeal fraction, the
pMC2A209 class from the Thaumarchaeota and the Methanosaeta genus were archaeal
FIG 5 Archaeal composition in wetland sediment samples performing AOM. Shown are the most abundant archaeal genera detected,
based on 16S rRNA amplicon gene libraries, on selected experimental treatments shown in Fig. 1 at the end of the incubation period (30
days).
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clusters that significantly increased their presence in the AQDS enrichment (34% and
23%, respectively). Also in the AQDS enrichment, the Bathyarchaeota phylum previously
detected in wetland sediments, both in the presence and in the absence of external
NOM, significantly increased its proportion in the archaeal community (by around 10%
with respect to the original composition), suggesting potential metabolic arrangements
that thrive under AQDS-dependent AOM conditions (Fig. 6b). Humus-reducing bacteria
FIG 6 AOM with AQDS as electron acceptor by an enrichment derived from wetland sediment. (a) Kinetics of methane consumption linked to AQDS reduction
(to AH
2
QDS) observed during the last 11 days of the entire enrichment process lasting 151 days: filled squares () represent microcosms with CH
4
as an electron
donor and AQDS as an electron acceptor (complete experiments, n⫽3), open squares (e) represent controls without an electron acceptor provided (without
AQDS control, n⫽3), filled circles () represent CH
4
-free microcosms (endogenous controls, n⫽3), and crosses (⫻) represent heat-killed controls (sterile
controls, n⫽2). Error bars represent the standard error among replicates. (b and c) Microbial community changes at the end of the enrichment (151 days of
incubation) at the phylum level based on Illumina sequencing of 16S rRNA V3 to V4 region. Fresh sediment composition was used as a reference.
Valenzuela et al. Applied and Environmental Microbiology
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that proliferated throughout the 5 months of enrichment included genera from the
Desulfuromonadales (29, 30), Clostridiales (14, 28), and Propionibacteriales (31) orders,
which increased by 27%, 7%, and 12%, respectively, with respect to the original
composition (Fig. 6c).
DISCUSSION
NOM as terminal electron acceptor fueling AOM in wetland sediment. Although
the complex composition of the studied wetland sediment challenged efforts to
elucidate the microbial processes responsible for the high methanotrophic activities
quantified, the present study provides multiple lines of evidence demonstrating that
electron-accepting functional groups present in its NOM fueled AOM by serving as a
terminal electron acceptor. Indeed, while sulfate reduction was the predominant
process, accounting for up to 42.5% of AOM activities, microbial reduction of NOM
concomitantly occurred. Furthermore, enrichment of wetland sediment with external
NOM, as Pahokee peat HS, significantly promoted AOM, with ⬃100 nmol
13
CH
4
oxidized · cm
⫺3
· day
⫺1
attributed to this microbial process. Spectroscopic evidence
also demonstrated that quinone moieties, which are main redox functional groups in
HS (19), were heterogeneously distributed in the studied wetland sediment and that
their reduction occurred during the course of AOM. Moreover, an enrichment derived
from wetland sediments performing AOM linked to NOM reduction stoichiometrically
oxidized methane coupled to AQDS. Sediment incubations performed in the presence
of the sulfate reduction inhibitor molybdate further confirmed the role of HS in AOM.
Certainly, even though sulfate-reducing activities significantly decreased in the pres-
ence of molybdate, AOM activities remained high, while microbial reduction of NOM
was doubled under these conditions. These interesting findings suggest that metha-
notrophic microorganisms performing sulfate-dependent AOM might have directed
electrons derived from AOM toward NOM when sulfate reduction became blocked, as
has been suggested based on experiments performed under artificial conditions (27).
Microbial communities in wetland sediments performing AOM. Archaeal clus-
ters consistently found in wetland sediment incubations performing AOM included
members from the MBG-D family, which have already been proposed as players in
metal-dependent AOM (6); thus, their presence agrees with evidence indicating that
AOM is linked to iron reduction observed in some experimental controls (Table S2).
Additionally, these microorganisms were not found in the AQDS enrichment, probably
due to the depletion of intrinsic ferric iron throughout the incubation cycles. Archaea
constantly present among fresh sediment incubation and AQDS enrichment were those
from the pMC2A209 class and the Bathyarchaeota phylum. To our knowledge, the
pMC2A209 class of archaea has not been related to AOM, but its close partners from the
MBG-B class have been consistently found in environments in which AOM occurs
(32–35). In fact, Thaumarchaeota members, including the MBG-B, have been found in
consortia performing AOM in the absence of ANME clades (36). Interestingly, the
pMC2A209 cluster seemed to duplicate its proportion up to 12% when sulfate reduc-
tion was inhibited (by molybdate), which might suggest that the impediment of sulfate
reduction enhanced its activity promoting AOM coupled to NOM reduction. With
respect to the Bathyarchaeota phylum, increasing evidence suggests that this lineage
might be involved in the methane cycle. Recently, it has been demonstrated that this
cluster possesses the necessary genetic elements to express the enzymatic machinery
required for methane production, and potentially methane consumption (37). Addi-
tionally, Saxton and colleagues have found abundant Bathyarchaeota representation in
a fulvic acid-rich deep sediment that oxidizes methane uncoupled from sulfate reduc-
tion (22). Unexpectedly, a very low percentage within the archaeal population was
identified as members from the ANME type archaea, even though it would be expected
to find ANME-2 members, since it is the only ANME subgroup with a proven capability
to derive electrons extracellularly toward humus and its analogues under artificial
conditions (27). Our microcosms, both in fresh sediment as well as in the AQDS
long-term enrichment, showed a barely detectable number of copies of ANME-1b and
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ANME-3 sequences retrieved by the methodology employed, suggesting a low pres-
ence of ANME microorganisms in the ecosystem studied.
Regarding the bacterial composition, while Clostridia,Bacilli, and Gammaproteobac-
teria were significantly represented within the fresh sediment performing AOM (Fig. S3),
the AQDS enrichment (Fig. 6) exhibited the most significant increase in Deltaproteo-
bacteria of the Desulfuromonadales order, which includes several humus-reducing
microorganisms (14). Since a wide diversity of microorganisms have been proven to
reduce humus analogues or HS, we do not rule out that diverse bacterial clusters could
have participated in partnership with detected archaea to jointly perform AOM coupled
to NOM reduction. Nevertheless, humus-reducing bacteria possess metabolic versatility
and capability to reduce miscellaneous electron acceptors, which makes it difficult to
come to conclusions about their participation in our experiments. Further investigation
must be done to unravel the potential involvement of humus-reducing bacteria in
AOM.
Ecological significance. Here, we report AOM coupled to microbial reduction of
NOM, which constitutes a missing link within the carbon cycle. HS frequently contribute
up to 80% of soil NOM and up to 50% of dissolved NOM in aquatic environments. While
the labile fraction of NOM promotes methanogenesis in anaerobic environments, the
slowly decomposing humic portion may serve as an important barricade to prevent
methane emissions in organotrophic ecosystems by serving as a terminal electron
acceptor driving AOM (Fig. 7). As an example, considering the maximum AOM driven
by the microbial reduction of NOM measured in humus-enriched sediments and the
global area of coastal wetlands (38, 39), we approximate that this microbial process
consumes up to 114 Tg of CH
4
· year
⫺1
. Considering the global wetland area (10), we
anticipate suppression of more than 1,300 Tg of CH
4
· year
⫺1
(see supplemental
material for details). Accordingly, NOM-driven AOM may be more prominent in or-
ganotrophic sites with poor sulfate content, such as peatlands, swamps, and or-
ganotrophic lakes. This premise is supported by the suppression of methanogenesis by
HS observed in different ecosystems (16, 17) and by the widespread AOM activity
reported across many peatland types (40–42). The potential role of HS is further
emphasized, because their electron-accepting capacity is fully recycled in recurrently
anoxic environments. Thus, the suppression of methanogenesis by HS may be much
greater than previously considered (it had been estimated to be on the order of 190,000
mol CH
4
·km
⫺2
· year
⫺1
[43]).
MATERIALS AND METHODS
Sediment sampling and characterization. Sediment cores were collected from the tropical marsh
Sisal, located in the Yucatán Peninsula, southeastern Mexico (21°09=26⬙N, 90°03=09⬙W) in January 2016.
Sediment cores with a depth of 15 cm were collected under a water column of approximately 70 cm.
Water samples were also collected from the area of sediment sampling points to be used as liquid
medium in anaerobic incubations. All sediment and water samples were sealed in hermetic flasks and
were maintained in ice until arrival at the laboratory. Upon arrival, all sampled materials were stored at
4°C in a dark room until analysis and incubation. Sediment cores were opened and homogenized within
an anaerobic chamber (atmosphere composed of 95%/5% [vol/vol] N
2
/H
2
) before characterization and
incubation. No amendments (addition of chemicals, washing, or exposure to air) were allowed on the
sediment and water samples in order to reflect the actual conditions prevailing in situ as closely as
possible. The characterization of water and sediment samples is described in Table S1.
Sediment incubations. Water samples collected from sediment sampling points were thoroughly
mixed before amendment with HS (2.5 g · liter
⫺1
) by magnetic stirring. Pahokee peat (Florida Everglades)
HS, purchased from the International Humic Substances Society, was employed as external NOM in
sediment incubations. Humus-enriched water was flushed with N
2
to blow away any dissolved oxygen.
Portions of 15 ml were then distributed in 25-ml serological flasks. Sediment containers were opened
inside an anaerobic chamber. Portions of 2.5 ml of previously homogenized wet sediment were then
inoculated into each serological bottle. After sealing all bottles with rubber stoppers and aluminum rings
inside the anaerobic chamber, they were flushed with N
2
. Once anaerobic conditions were established,
5mlof
13
C-methane was injected into each vial to reach a
13
CH
4
partial pressure of 0.67 atm in a
headspace of 7.5 ml. Controls incubated in the absence of external HS were also prepared according to
an identical protocol. Killed controls included chloroform at a concentration of 10% (vol/vol) to annihilate
any microbial activity. Additional incubations were executed in the presence of the sulfate reduction
inhibitor molybdate (25 mmol liter
⫺1
) in the presence and absence of external NOM. All incubation
Valenzuela et al. Applied and Environmental Microbiology
June 2017 Volume 83 Issue 11 e00645-17 aem.asm.org 10

bottles were statically placed in a dark room at 28°C (the temperature prevailing at the Sisal wetland at
the sampling time). The pH remained at 7.5 ⫾0.05 throughout all incubations.
Enrichment incubations with AQDS. Incubations were commenced by inoculating 120-ml serolog-
ical bottles with 10 g of volatile suspended solids (VSS) per liter of Sisal sediment. Prior to inoculation,
portions of 60 ml of artificial medium were distributed into the incubation bottles and flushed for 15 min
with a mixture of N
2
and CO
2
(80%/20% [vol/vol]) for stripping any dissolved oxygen from the medium.
AQDS (⬎98.0% purity; TCI America Chemicals) was added at a concentration of 10 mmol liter
⫺1
as a
terminal electron acceptor, along with the following basal medium components (in grams per liter):
NaHCO
3
(5), NH
4
Cl (0.3), K
2
HPO
4
(0.2), MgCl
2
·6H
2
O (0.03), and CaCl
2
(0.1). Trace elements were included
in the medium by adding 1 ml · liter
⫺1
of a solution with the following composition (in milligrams per
liter): FeCl
2
·4H
2
O (2,000), H
2
BO
3
(50), ZnCl
2
(50), CuCl
2
·6H
2
O (90), MnCl
2
·4H
2
O (500), AlCl
3
·6H
2
O (90),
CoCl
2
·6H
2
O (2,000), NiCl·6H
2
O (920), Na
2
SeO·5H
2
O (162), (NH
4
)
6
Mo
7
O
24
(500), EDTA (1,000), Na
2
WO
4
·H
2
O
(100), and 1 ml · liter
⫺1
HCl at 36%. The final pH of the medium was 7.2, and no changes were observed
throughout the incubation time. Once inoculation took place, microcosms were sealed with rubber
stoppers and aluminum rings and then flushed with the same N
2
-CO
2
mixture. After anoxic conditions
were established, 1 ml of sodium sulfide stock solution was injected into each vial to reach a sulfide
concentration of 0.1 g · liter
⫺1
in order to consume any traces of dissolved oxygen. Methane was
provided into the microcosms by injecting 30 ml of CH
4
(99.9% purity; Praxair), reaching a partial pressure
of methane of 0.54 atm. Subsequent incubations were performed after AQDS was reduced (i.e.,
converted to AH
2
QDS), coupled to anaerobic oxidation of methane (AOM). A new set of bottles
containing basal medium with AQDS (10 mmol liter
⫺1
) were inoculated within an anaerobic chamber by
transferring 10 ml of slurry (sediment and medium) taken from previous incubations (Fig. S1). The
following incubations were completed under the same experimental conditions.
Analytical techniques. (i) Isotopic carbon dioxide and methane measurements. Ions 16 (
12
CH
4
),
17 (
13
CH
4
), 44 (
12
CO
2
), and 45 (
13
CO
2
) were detected and quantified in an Agilent Technologies 7890A gas
FIG 7 Schematic representation of methane generation and consumption by wetland sediment biota. While a fraction of NOM may serve
as an electron acceptor to support AOM (NOM-AOM) and decouple sulfate reduction-dependent AOM (SR-AOM), depending on its
chemical properties, a labile fraction of NOM could also be degraded following the methanogenesis pathway by a fermenting and
methanogenic fraction of the consortia. Equilibrium between these three phenomena must be tightly dependent on thermodynamic
conditions, concentration of chemical species, and composition of microbial community. *, anaerobic methanotrophic archaea are
considered in a broader perspective than ANME clades from the Euryarchaeota phylum.
Anaerobic Methane Oxidation Linked to Humus Reduction Applied and Environmental Microbiology
June 2017 Volume 83 Issue 11 e00645-17 aem.asm.org 11

chromatograph (GC) coupled to an Agilent Technologies 5975C mass spectrometer (detector); ionization
was achieved by electronic impact and a quadrupole analyzer. For the analysis, an Agilent Technologies
HP-PLOT/Q capillary column with a stationary phase of poly(styrene-divinylbenzene) (30 m by 0.320 mm
by 20
␮
m) was employed as stationary phase using helium as a carrier gas. The chromatographic method
was as follows: the starting temperature was 70°C, which was held for 3 min, and then a ramp with an
increase of 20°C per min was implemented until 250°C was reached and maintained for 1 min. The total
time of the run was 13 min. The temperature of the injection port was 250°C. The injection volume was
20
␮
l, and there was only one replicate of injection per bottle. The gas injected into the GC was taken
directly from the headspace of the incubations and immediately injected into the GC port. Methane
calibration curves were made by injection of different methane (99.9% purity) volumes into serological
bottles under the same experimental conditions (atmosphere composition, pressure, temperature, and
liquid volume).
12
CO
2
and
13
CO
2
curves were made using different dried sodium bicarbonate (99% purity;
Sigma-Aldrich) and sodium
13
C-labeled carbonate (99 atom %
13
C; Sigma-Aldrich) concentrations, respec-
tively, in serological bottles which contained the same volume of wetland sediment and water used in
incubations. Standards were incubated at room temperature for 12 h until equilibrium with the gaseous
phase was reached. The linear regression analysis of obtained measurements had a correlation coefficient
higher than 0.97.
13
CO
2
production rates were based on the maximum slope observed on linear
regressions considering at least three sampling points.
(ii) Methane quantification in AQDS enrichment. Net methane consumption was assessed in terms
of methane concentration measurements in the headspace of microcosms. These measurements were
carried out by injecting 100
␮
l of gas samples from the headspace of incubation bottles into a gas
chromatograph (Agilent Technologies 6890M) equipped with a thermal conductivity detector and a
HayeSep D column (Alltech, Deerfield, IL, USA) with the dimensions 3.048 m by 3.185 m by 2.16 mm.
Helium was employed as a carrier gas at a flux of 12 ml · min
⫺1
. The temperatures of the injection port,
oven, and detector were 250, 60, and 250°C, respectively. Calibration curves were made for each reaction
volume used by injecting different methane concentrations into serological bottles under the same
experimental conditions at which microcosms were studied (atmosphere composition, pressure, tem-
perature, and liquid volume).
(iii) Determination of electron-accepting functional groups in solid phase by XPS. Sediment
samples (solid fraction of microcosms) were dried under a constant nitrogen flow after incubation with
methane. Once sediments became dried, bottles were open inside an anaerobic chamber with an
atmosphere composed of 95%/5% (vol/vol) N
2
/H
2
and were triturated on an agate mortar. Samples were
then kept under anaerobic conditions until analysis in a PHI VersaProbe II X-ray photoelectron spectros-
copy analyzer (Physical Electronics, ULVAC-PHI). Two representative spectra were recorded per scanned
sample.
(iv) Determination of electron-accepting functional groups in solid phase by micro-ATR-FTIR
imaging. Micro-ATR-FTIR images were collected from each sample with a continuous-scan spectrometer,
the Agilent 660 FTIR interfaced to a 620 infrared microscope with a 32 by 32 focal plane array (FPA)
detector and Ge ATR objective for micro-ATR. Each pixel obtains a full IR spectrum or a total of 1,024
spectra. Background spectra were collected from a clean ATR crystal (i.e., without sample). The Ge crystal
of the ATR microscope was lowered onto the surface of each sample for a contact area of approximately
100 by 100
␮
m. Spectra were collected by coaddition of 256 scans over a spectral range of 4,000 to 900 ·
cm
⫺1
, at a spectral resolution of4·cm
⫺1
. In all images, a color scale bar is set within the software to reflect
the relative concentration range, from low to high. Agilent Resolutions Pro was used for data acquisition
and analysis.
(v) Determination of electron-accepting functional groups in liquid phase by high-resolution
UV-Vis–near-infrared spectroscopy. After each incubation cycle, liquid samples (1.5 ml) were taken in
an anaerobic chamber with a disposable syringe and put into a quartz cell, which was sealed with plastic
film in order to maintain anoxic conditions during spectrometric analysis. Spectra were obtained in a
Varian Cary 5000 UV-Vis (diffuse reflectance) spectrophotometer equipped with an integrating sphere.
(vi) Nitrite and nitrate determinations. Nitrite and nitrate concentrations were measured according
to spectrometric techniques established by Standard Methods (44). Nitrate measurement is taken under
acidic conditions at a wavelength of 275 nm, and the value obtained is corrected for dissolved organic
matter, which has its maximum absorbance at 220 nm. Nitrite forms a purple complex through a reaction
with sulfanilamide and N-(1-naphthyl) ethylene diamine, which presents its maximum absorbance at a
wavelength of 543 nm. Samples were taken with a disposable syringe directly from the microcosms,
injected into sealed quartz cuvettes or glass tubes (depending on the required lecture wavelength), and
immediately taken to the spectrophotometer to avoid any reaction of the sample with atmospheric
oxygen.
(vii) Sulfate and sulfide determinations. Samples were extracted from microcosms and immedi-
ately filtered through 0.22-
␮
m-pore-size nitrocellulose membranes. Filtered samples were then diluted
(1:10) with deionized water and processed in an Agilent capillary electrophoresis system (Agilent
Technologies), according to the methodology proposed by Soga and Ross (45). Dissolved sulfide was
measured by the spectrometric method proposed by Cord-Ruwisch (46). Briefly, 100
␮
l of sample
was taken and immediately vortexed with 4 ml of an acidic CuSO
4
solution. Absorbance at 480 nm was
immediately registered in a UV-Vis spectrophotometer (Thermo Spectronic) to avoid sulfide oxidation
before measurements.
(viii) Humic substance reduction and ferrous iron measurements. Quantification of the reduction
of electron-accepting functional groups in HS was performed according to Lovley et al. (18). Slurry
samples (⬃500
␮
l) were taken from microcosms with a disposable syringe while bottles were being
Valenzuela et al. Applied and Environmental Microbiology
June 2017 Volume 83 Issue 11 e00645-17 aem.asm.org 12

manually shaken inside an anaerobic chamber. A portion of each sample (200
␮
l) was mixed with an
equal volume of an acidic solution (0.5 mol liter
⫺1
HCl) and allowed to stand for 30 min, while the same
volume of sample was reacted with ferric citrate (20 mmol liter
⫺1
) for 3 h. After reaction with ferric citrate,
samples were mildly resuspended in a vortex, and 200
␮
l was left repose with the same volume of HCl
solution for 30 min. Afterwards, each sample was centrifuged for 10 min at 10,000 ⫻gin a Spectrafuge
16M centrifuge, and 200
␮
l of supernatant was then recovered and reacted with a 0.2 g · liter
⫺1
solution
of 2,4,6-Tris(2-pyridyl)-1,3,5-triazine (ferrozine reagent). Ferrous iron produced due to the chemical
reduction of ferric citrate by reduced functional groups in HS forms a purple complex along with
ferrozine reagent, which has its maximum absorbance at 562 nm. The ferrozine solution was buffered
with HEPES (50 mmol liter
⫺1
). Once centrifuged samples were mixed with ferrozine solution, they were
left reacting for 10 min before their measurement in a Thermo Scientific Genesis 10 UV spectrometer
located inside an anaerobic chamber. All solutions employed in this determination were bubbled with
N
2
for 30 min to ensure the absence of dissolved oxygen.
(ix) Total carbon, TOC, and total inorganic carbon measurements. Water samples were filtered
through 0.22-
␮
m-pore-size nitrocellulose membranes and diluted with deionized water, while sediment
samples were dried until constant weight. Both liquid and solid samples were analyzed in a Shimadzu
TOCVCS/TNM-1 total organic carbon (TOC) analyzer equipped with a solids sampling port (SSM-5000A).
The solid-sample processing time was 6 min at 900°C using O
2
(500 ml · min
⫺1
, 99.9% purity) as a carrier
gas; all samples were analyzed in triplicate.
(x) Total, volatile, and fixed solids. Total, fixed, and volatile solids were measured in triplicate
according to the Standard Methods procedure (44).
(xi) Elemental composition. The elemental composition of the sediments was assessed by analyzing
acid extracts from2gofdrysediment. In the case of iron and manganese measurements in microcosms,
supernatant samples were taken with disposable syringes, filtered, and acidified prior to analysis.
Samples were then analyzed by inductively coupled plasma-optical emission spectrometry (ICP-OES) in
an equipment Varian 730-ES. The operational conditions were: 1 kW potency, 1.5 liters · min
⫺1
auxiliary
flow, 0.75 liters · min
⫺1
net flow, 30-s sample-taking delay, and 3 measured replicates by sample. Argon
was employed as a carrier gas.
(xii) DNA extraction, PCR amplification, and sequencing. One microcosm for each selected
treatment was randomly chosen at the end of the incubation period (30 days for experiments presented
in Fig. 1 and 151 days for experiments depicted in Fig. 6). Before DNA extraction, liquid medium was
decanted and extracted from the serological bottles. The total sediment was homogenized afterwards,
and a subsample of 0.5 g was taken to proceed with DNA extraction. The remaining sediment and the
other microcosms were used for material characterization. The total DNA was extracted from sediment
samples using the PowerSoil DNA extraction kit (Mo Bio Laboratories, Carlsbad, CA, USA), according to
the protocol described by the manufacturer. DNA isolated from each sample was amplified using primers
341F and 785R, targeting the V3 and V4 regions of the 16S rRNA gene fused with Illumina adapter
overhang nucleotide sequences (47). The PCRs were performed in 50-
␮
l reaction mixtures using Phusion
Taq polymerase (Thermo Scientific, USA) under the following conditions: denaturation at 98°C for 60 s,
followed by 5 cycles of amplification at 98°C for 60 s, 50°C for 30 s, and 72°C for 30 s, followed by 25
cycles of amplification at 98°C for 60 s, 55°C for 30 s, and 72°C for 30 s, and a final extension of 72°C for
5 min. Two independent PCRs were performed for each sample. The products were indexed using
Illumina’s 16S metagenomic sequencing library preparation protocol and Nextera XT index kit version 2
(Illumina, San Diego, CA). Libraries were deep sequenced with the Illumina MiSeq sequencer.
(xiii) Bioinformatics analysis. An analysis of 16S rRNA gene libraries was carried out using the
mothur open source software package (version 1.34.4) (48). The high-quality sequence data were
analyzed for potential chimeric reads using the UCHIME algorithm. Sequences containing homopolymer
runs of 9 or more bases, those with more than one mismatch to the sequencing primer, and those with
a Q-value average below 25 were eliminated. Group membership was determined prior to the trimming
of the barcode and primer sequence. Sequences were aligned against the SILVA 123 16S/18S rRNA gene
template using the Nearest Alignment Space Termination (NAST) algorithm and trimmed for the optimal
alignment region. A pairwise distance matrix was calculated across the nonredundant sequence set, and
reads were clustered into operational taxonomic units (OTUs) at 3% distance using the farthest neighbor
method. The sequences and OTUs were categorized taxonomically using mothur’s Bayesian classifier and
the SILVA 123 reference set. The sequences obtained have been submitted to the NCBI GenBank
database.
Accession number(s). The accession numbers of the sequences in this work were deposited in the
GenBank Sequence Read Archive under BioProject number SRP094593.
SUPPLEMENTAL MATERIAL
Supplemental material for this article may be found at https://doi.org/10.1128/AEM
.00645-17.
SUPPLEMENTAL FILE 1, PDF file, 0.3 MB.
ACKNOWLEDGMENTS
We thank Derek R. Lovley, Jim A. Field, Alfons Stams, Frederic Thalasso, Cesar Nieto,
and Sonia Arriaga for discussions. We are also grateful to Matthew Tippett for proof-
reading the manuscript. We thank Dulce Partida, Ma. Carmen Rocha-Medina, Guadalupe
Anaerobic Methane Oxidation Linked to Humus Reduction Applied and Environmental Microbiology
June 2017 Volume 83 Issue 11 e00645-17 aem.asm.org 13

Ortega-Salazar, Mariela Bravo-Sánchez, Roberto Camposeco, Elizabeth Cortez and
Guillermo Vidriales, for technical support, as well as Lluvia Korynthia López-Aguiar and
Yessica Parera-Valadez for their help during sediment collection expeditions. We thank
Vicente Rodríguez for access to the Cary 5000 UV-Vis spectrophotometer. We are also
grateful for the use of infrastructure of the National laboratories LINAN and LANBAMA
at IPICYT, as well as USMB at UNAM.
This work was financially supported by grants from the Council of Science and
Technology of Mexico (Program Frontiers in Science, grant 1289) and the Marcos
Moshinsky Foundation to F.J.C.
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