ABSTRACT Oxygen minimum zones (OMZs) contain the largest pools of oceanic methane but its origin and fate are poorly understood. High-resolution (<15 m) water column profiles revealed a

300 m thick layer of elevated methane (20–105 nM) in the anoxic core of the largest OMZ, the Eastern Tropical North Pacific. Sediment core incubations identified a clear benthic methane

source where the OMZ meets the continental shelf, between 350 and 650 m, with the flux reflecting the concentration of methane in the overlying anoxic water. Further incubations

characterised a methanogenic potential in the presence of both porewater sulphate and nitrate of up to 88 nmol g−1day−1 in the sediment surface layer. In these methane-producing sediments,

anaerobic methane oxidation is corroborated by the presence of particulate methane monooxygenase (_pmoA_) gene sequences, related to type I methanotrophs and the lineage of _Candidatus_

Methylomirabilis oxyfera, known to perform nitrite-dependent anaerobic methane oxidation (N-DAMO), respectively. SIMILAR CONTENT BEING VIEWED BY OTHERS THERMOCHEMICAL OXIDATION OF METHANE BY

MANGANESE OXIDES IN HYDROTHERMAL SEDIMENTS Article Open access 24 June 2023 SEAWATER SULPHATE HERITAGE GOVERNED EARLY LATE MIOCENE METHANE CONSUMPTION IN THE LONG-LIVED LAKE PANNON Article

Open access 13 June 2023 METHANE LEAKAGE THROUGH THE SULFATE–METHANE TRANSITION ZONE OF THE BALTIC SEABED Article 06 December 2024 INTRODUCTION Methane is the most abundant hydrocarbon in

the atmosphere and a potent greenhouse gas, which has contributed ~20% to the Earth’s warming since pre-industrial times (Lashof and Ahuja, 1990; Reeburgh, 2007; Kirschke et al., 2013). The

marine environment encompasses large reservoirs of methane (Zhang et al., 2011; Pack et al., 2015), particularly in oxygen minimum zones (OMZs). Here, oxygen is consumed faster than it is

resupplied, resulting in a layer of hypoxic waters surrounding a functionally anoxic core (Thamdrup et al., 2012), where methane accumulates (Wright et al., 2012). Under a warming climate,

the dissolution of oxygen in seawater will decrease, whereas its consumption through respiration will likely increase (Vázquez-Domínguez et al., 2007) and thermal stratification could become

more intense. Together, these biotic and abiotic changes will thicken OMZs moving these large methane pools closer to the zone of atmospheric exchange (Stramma et al., 2008; Keeling et al.,

2010; Helm et al., 2011). Marine methanogenesis, which produces 0.7–1.4Tg each year (Krüger et al., 2005), forms an essential link in the carbon cycle, preventing the long-term burial of

carbon in the sediments by mineralising it and returning it to the water in gaseous form (Ferry and Lessner, 2008). The thermodynamics of organic matter oxidation dictate that sulphate

reduction and methanogenesis should be mutually exclusive reactions. Although the clear spatial partitioning of these two microbial processes has been widely observed in marine sediments

(Martens and Berner, 1977; Whiticar, 2002; Reeburgh, 2007), non-competitive methanogenesis can co-occur with other anaerobic processes (Valentine, 2011). Non-competitive methanogenisis

disproportionates methylated substrates (for example, methyl amine, methane thiols and methanol) to yield methane and carbon dioxide, and since its discovery in the early 1980s (Oremland et

al., 1982), it has been found to occur in all major oceans (D’Hondt et al., 2002; Mitterer, 2010; Valentine, 2011). However, relatively little scientific attention has focused on microbial

methanogenesis compared with that around gas hydrates and cold seeps (Shakhova et al., 2005; Valentine, 2011; Boetius and Wenzhofer, 2013). The majority of methane produced in marine

sediments is thought to be oxidised anaerobically, limiting its flux to the overlying water (Knittel and Boetius, 2009). Any methane leaking into the water column may still be oxidised, by

pelagic aerobic or anaerobic bacteria, which form a final barrier preventing its escape to the atmosphere (Reeburgh et al., 1991; Blumenberg et al., 2007; Kessler et al., 2011; Heintz et

al., 2012). We focused on locating the origin of methane in the Eastern Tropical North Pacific (ETNP), between 70 and 720 km off the Guatemalan coast. The ETNP OMZ is both the world’s

largest OMZ (Paulmier and Ruiz-Pino, 2009) and the largest reservoir of oceanic methane (Sansone et al., 2001, 2004; Reeburgh, 2007; Naqvi et al., 2010). Here, the methane is thought to be

formed by a combination of decomposing sinking organic matter and coastal or benthic sources but neither have been directly measured (Sansone et al., 2001, 2004). Porewater and bottom-water

methane concentrations along with stable isotope ratio data suggested the sediments were the source of the pelagic methane and the flux was greatest where the anoxic core of the OMZ touched

the sediment (western Mexican margin, Sansone et al. (2004)). Although these studies offer useful insights, there are no direct measurements of sediment methanogenesis or methane efflux in a

marine OMZ. Pelagic methane oxidation in marine environments is a rarely quantified process but on the margins of an OMZ, where methane intersects traces of oxygen, it could be a

significant process. Published rates span ~0.001–10 nmol l−1day−1 and all studies used either 3H-CH4 or LL-14C as a tracer (Mau et al., 2013). The only study to have successfully measured

methane oxidation in the ETNP OMZ (Pack et al., 2015) found exceptionally slow rates (0.000034–4 nmol l−1day−1), which could explain how the methane, if of benthic origin, can be sustained

hundreds of kilometres offshore. We used high-resolution water column profiles to show that methane concentrations peak in the anoxic core of the ETNP OMZ. We then used a combination of

water and sediment incubations, along with stable isotope tracers and molecular analyses, to quantify sediment methane flux, methanogenic potential and pelagic methane oxidation potentials.

We hypothesised that all sediments would contain active methanogens, but that their activity would be controlled by the oxygen concentration in the bottom-water. Further, the methane

released from the sediments is then oxidised by aerobic and/or anaerobic methanotrophs in the water column as it moves towards the OMZ margins. To the best of our knowledge, this is the

first study to combine biogeochemical with molecular microbial data, in order to better understand the origins and fate of methane in the ocean’s largest OMZ. MATERIALS AND METHODS SAMPLE

SITES This study comprised two cruises in the ETNP: the first (D373, 11 December 2011–13 January 2012), which focused on the water column (0–4000 m), was structured around 6 ‘offshore’ sites

located along 92.5oW, between 8 and 13°N (Supplementary Figure S1). The second (JC097, 28 December 2013–10 February 2014), concentrated on the continental shelf and slope, 70–150 km off the

Guatemalan coast (Supplementary Figure S1) and here, both sediments and water column samples were collected. A standard conductivity–temperature–depth rosette, comprising 24 Niskin (20

litre) bottles and a Sea-Bird 24 electronics system (fluorimeter, altimeter, photosynthetically active radiation and oxygen sensors, and so on) was used to collect water and a multi-corer

(Mega Corer, OSIL, Havant, UK) was used to recover intact cores of sediment and overlying water. WATER COLUMN GAS AND NUTRIENT PROFILES High-resolution (5–15 m) water column profiles

(_n_=21) were constructed to define the OMZ and locate the methane. To minimise atmospheric contamination, water for methane analysis was discharged from the Niskin bottles into 12.5 ml

gas-tight vials (Labco, Lampeter, UK) via Tygon tubing and allowed to overflow three times before capping, temperature equilibration and head-spacing (2 ml helium (BOC, Guildford, UK)).

Methane was measured on-board using a gas chromatograph fitted with a flame ionisation detector (gas chromatography/flame ionization detector Agilent Technologies (Santa Clara, CA, USA), see

Sanders et al. (2007) for details). Oxygen concentrations were measured by the Sea-Bird sensor (Bellevue, WA, USA) (with a limit of detection (LOD) ~1.4 μmol l−1) and nitrite was measured

using a segmented flow auto-analyser (Skalar, Breda, Netherlands; LOD=0.05 μmol l−1, Nicholls et al., 2007). SEDIMENTS AS A METHANE SOURCE Sediment-water flux was determined using intact

cores and the methanogenic potential of discrete layers was quantified using slurries. As the conductivity–temperature–depth could not sample closer than ~10 m from the seabed, the water

overlying the sediment (_n_=3 from the least disturbed core) was sampled, as above, to measure the methane concentration as close to the seabed as possible (<15 cm). Next, six sediment

mini-cores were subsampled from three of the large cores (using Perspex tubes, 3.4 × 25 cm), sealed with rubber bungs and transferred to a temperature controlled (10 °C) tank. This was

repeated at 16 locations ranging in seabed depth from 100 to 900 m. Methane flux was quantified by measuring methane in the overlying water before and after a sealed 24-h incubation. First,

the overlying water was degassed by bubbling (2 min) with oxygen-free nitrogen (BOC), to ensure all cores were incubated under the same hypoxic conditions (precise concentration verified

using an oxygen micro-sensor, Unisense, Aarhus, Denmark) and that the majority of ambient methane was removed (previous experiment had demonstrated that 2 min was sufficient to remove

>90% methane). Water samples were taken from each mini-core after degassing (T0), they were then sealed with bungs with inbuilt magnetic stirrers, and left for 24 h in the dark until a

second water sample (Tfinal) was taken for methane analysis. The daily flux of methane was calculated as the increase between T0 and Tfinal. To identify the sediment layer with the greatest

methanogenic potential, additional large sediment cores (six locations, Table 1) were carefully extruded and ~4 ml of sediment and 3 ml of bottom water (overlying the cores) was transferred

to gas-tight vials using a truncated 1 ml syringe (to minimise air contamination) to create a slurry. The headspace and water was purged with helium for 2 min to deoxygenate the vials and

optimise conditions for methanogenesis. The methane concentration in the headspace was measured by gas chromatography/ flame ionization detector 4–8 times over the following 4–12 days and

between measurements vials were kept at 12 °C in the dark. Following the first two experiments (550 m and 650 m), only the top 5 cm was used for further sites. The concentration of sulphate,

nitrite and nitrate in the sediment porewater was measured in eight large cores from four different locations (150, 350, 550 and 750 m seabed depth) by ion chromatography (Dionex,

Sunnyvale, CA, USA; for sulphate) and segmented flow auto-analyser (Skalar for nitrite and nitrate), after separating the porewater from the sediment by centrifugation. Finally, hydrogen

sulphide was measured in the cores by inserting a calibrated, miniaturised amperometric sensor (Unisense) into an extruded portion of the core, from the side, at 2 cm intervals. AEROBIC AND

ANAEROBIC WATER COLUMN METHANE OXIDATION We set up four experiments using 13C-labelled methane to quantify the potential for aerobic and anaerobic methane oxidation in the water column

(Supplementary Figure S2). First, we set up short time experiments with water from the upper margin of the OMZ, where oxygen is at the LOD (200 and 226 m). Seawater saturated with 99.9%

13C-CH4 was used to spike the samples with 3.3 nmol 13C-CH4 (264 nmol L−1) to avoid the need for a headspace and, thus, maintain ambient oxygen conditions. Samples were fixed (100 μl of 12.2

 m HCl) at 3–5 time points over 10–15 days to track the accumulation of dissolved inorganic carbon (13C-DIC). Second, we set up dose-response experiments (65 and 200 m), whereby we varied

the injection volume to give a range of methane concentrations (44–790 nmol l−1) to assess the extent to which the methanotrophic community was substrate limited. These were left for the

duration that the samples took to get back to the UK (5 months), without a headspace, before fixing. Third, we started more widespread long-term incubations (eight locations) at one methane

concentration (264 nmol l−1) and after fixing a sample within 2 min (control), we incubated the remaining samples for 5 months at 12 °C in the dark. Finally, to test for the potential for

nitrite-dependent anaerobic methane oxidation (N-DAMO) we incubated water from five depths spanning the upper boundary and into the core of the OMZ, where nitrite and methane were both

present (235–412 m, Supplementary Figure S2 and Table 2) with 13C-CH4 (3.4 μmol l−1) and 15N-NO2 (11.4 μmol l−1), just 13C-CH4 or no spike. These samples were taken from depths with no

detectable oxygen but, to ensure complete anoxia in the water, we introduced a 2 ml helium headspace before adding the gas spikes. For all four types of incubations, we included a reference

sample (no spike, fixed at same time as samples), a spiked-control (spiked with 13C-CH4 and/or 15NO2 and killed at the beginning of the experiment) and three technical replicates of each

treatment. Microbial activity was stopped, and any resulting 13C-DIC converted to 13C-CO2 for analysis, by injecting HCl through the septa (as above). See Supplementary Figure S2 for sample

location details. Upon return to the UK, all unfixed samples were fixed and a 2 ml helium headspace was introduced into those incubated without one. To confirm the initial CH4 concentration,

spiked-control samples were analysed on a gas chromatography/flame ionization detector and then the 13C-DIC (and 15N-N2 where necessary) was quantified using an elemental analyser

interfaced with a continuous flow isotope ratio mass spectrometer (Sercon 20–22, Sercon Group, Crewe, UK), calibrated against sodium bicarbonate (0–4 mM for DIC) or air (for N2). MOLECULAR

ANALYSIS Water (see Supplementary Table S1 for details) was filtered either through stand alone pumps or Nalgene filtration units (Supor, Pall, Port Washington, NY, USA; ø 293 mm for stand

alone pumps or ø 47 mm for Nalgene units, 0.2 μm pore size filters). Filters were immediately frozen in liquid nitrogen, placed in –80 °C freezer and transferred to the UK for DNA

extraction. Sediments (seabed depth 222 , 342, 550, 650 and 657 m) were collected from the top 2 cm of the sediment cores into 2 ml cryovials and frozen (as above) until DNA extraction.

Details of the extraction process and downstream analysis are given in Supplementary Information. ACCESSION NUMBERS The DNA sequences reported in this study were deposited in the EMBL

database under the accession numbers LT575999–LT576028. RESULTS WATER COLUMN PROFILES Over the two cruises, we constructed 21 water column profiles covering seabed depths ranging from 55 to

5320 m. Oxygen declined steeply from an average of 193 μmol l−1 in the top 20 m, to 6.4 μmol l−1 at 80 m and then slowly until it went below our LOD (1.4 μmol l−1) at 230 m (Figure 1). Below

800 m, oxygen returned to detectable concentrations and reached 100 μmol l−1 (30% saturation) at 2500 m where it remained stable until the seabed (>5000 m). The baseline nitrite

concentrations were 0.05 (LOD) to 0.2 μmol l−1 but within the OMZ, at 275–600 m, there was a large, secondary nitrite maximum of up to 1.8 μmol l−1 at 345 m and in the epipelagic waters, a

smaller, primary maximum at around 50 m (max 1.37 μmol l−1) in most profiles (Figure 1). The true anoxic core of the OMZ was where oxygen was below the LOD and a clear secondary nitrite

maximum was present (230–600 m). Methane was supersaturated relative to the atmosphere throughout the water column and there was a clear peak around 250–600 m (Figure 1). The maximum

concentration (102 nmol l−1) was measured at 368 m on the continental shelf where the seabed depth was 506 m, although very close to the sediments (<15 cm) 254 nmol l−1 was measured.

Outside the core of the OMZ, the methane was consistently above atmospheric equilibration at 3–5 nmol l−1, except for a small epipelagic methane peak (Figure 1), which was only found in some

of the profiles (maximum concentration 25 nmol l−1 at 65 m). METHANE FLUX, METHANOGENESIS AND THE METHANOGEN COMMUNITY In the intact, anoxic core incubations, methane flux averaged 262±65.5

 nmol m−2day−1, peaking at 1007 nmol m−2day−1 at 550 m and with the slowest of 162 nmol m−2day−1 measured at 300 m (Figure 2a). Although all sediment cores were degassed to remove oxygen

(that is, optimal conditions for methanogenesis), methane efflux was greater in sediments from locations where the OMZ intersected the shelf (indicated by shaded area on Figure 2a) compared

with those with oxygenated (2–4.6 μmol l−1) bottom-water (_X_2(1)=13.261, _P_<0.0001). The efflux of methane from the sediments was positively correlated with the concentration of methane

in the bottom-water (_X_2(1)=23.233, _P_<0.0001), which ranged from 6 to 254 nmol l−1 (Figure 2b). Further, there was a strong, non-linear inverse relationship between the concentration

of methane and oxygen in the bottom-water (Figure 2b, inset, LOD~1.4 μM O2). Incubating anoxic sediment slurries from discrete depth intervals from two locations revealed that the bulk of

the methanogenic potential was in the surface sediments (Figure 2c) and so all further experiments were focused on this layer (Table 1). Methanogenesis was also detected in the overlying

water (0–5 cm above sediment) and in sediments down to 25 cm, but in the uppermost layer the rate was at least an order of magnitude higher than any other depth (Figure 2c). The greatest

potential was measured in sediment from 750 m (88 nmol g−1day−1, Table 1). Hydrogen sulphide was detected in two out of the six cores in which it was measured; at 550 m the concentration

reached 59 μmol l−1 at 25 cm (Figure 2c) and in a core from 350 m it reached 219 μmol l−1 at 23 cm. The porewater profiles (Supplementary Figure S3) revealed that in the top 2 cm of

sediment, where methane production was most active, sulphate (23 mmol l−1), nitrite (2.8 μmol l−1) and nitrate (86 μmol l−1) were similar to, or above, bottom-water concentrations. A total

of 126 303 _mcrA_ gene sequences from the top 2 cm were retrieved and clustered into 16 operational taxonomic units (OTUs), hereafter named as ETNP_MG. The majority of sequences (50% of

sequences represented by ETNP_MG2 and 35% represented by ETNP_MG1) were 96–97% similar to _Methanococcoides_ sp. (Supplementary Table S2). Most of the OTUs (13 out of 16) clustered within

the order Methanosarcinales, whereas two OTUs (ETNP_MG3 and ETNP_MG5) clustered within the order Methanomicrobiales and one (ETNP_MG14) within the order Methanocellales (Figure 3a). The

_Methanococcoides_-like species dominated all five seabed samples, which displayed a similar level of intra-sample diversity (assessed by Shannon and Simpson indices, Supplementary Table

S2). Principal coordinate analysis indicates that most of the variation (91.2%) in the methanogen community is explained by the first two principal coordinates (58.5% of the variation

explained by axis 1 and 32.7% by axis 2, Figure 3b). The most separated communities, by the first coordinate, are those from 650 to 222 m, because of unique Methanomicrobiales (ETNP_MG5) and

Methanocellales (ETNP_MG14) sequences at 650 m and Methanosarcinales at 222 m. The community at 342 m is also somewhat separated from the others, as it is the only one with ETNP_MG3

Methanomicrobiales-like sequences. METHANE OXIDATION AND METHANOTROPHS Methane oxidation was measured, through the accumulation of 13C-DIC, in short-term (10–15 days) time series incubations

with water from the uppermost margins of the OMZ (Figure 4a and Supplementary Figure S2); 13C-DIC was produced at a rate of 3.0–5.9 nmol l−1 day−1, and, after 15 days, 26% of the 13C-CH4

had been oxidised to 13C-DIC. In the epipelagic zone, we could not measure any methane oxidation at 47 m but we did at 65 m (Table 2). Our dose-response experiments indicate that the

methanotrophs can oxidise methane at concentrations much higher than the ambient concentration (Figure 4b). Water from both the epipelagic (65 m) and mesopelagic waters (200 m, where oxygen

was below detection) oxidised increasing amounts of 13C-CH4 to 13DIC with increasing initial methane spike (Figure 4b) and there was a good correlation between methane oxidised and 13C-DIC

produced (R2=0.96). This relationship between 13C-DIC produced and starting methane concentration was linear (_R_2(200m)=0.94 and _R_2(65m)=0.54) for the range of concentrations tested

(85–760 nmol l−1) and the slope (b1(200m)=0.0055 and b1(65m)=0.0057) of the relationship was similar for the two different water samples (Figure 4b). Long-term (5 months) incubations from

eight different locations (47–228 m), yielded mixed results, with methane oxidation being undetectable in some vials (Table 2). The greatest amount of 13C-DIC produced was at 200 m where,

following a 3.3 nmol spike of 13C-CH4, 0.6 nmol 13C-DIC was measured in the water after 5 months. In the shorter incubations, water from the same location, produced a similar amount in only

10 days (Figure 4a), which indicates methane oxidation did not continue linearly during the 5 months. For comparison, if a total of 0.6 nmol 13C-DIC in the vial accumulated linearly, after

150 days, it would equate to 0.42 nmol l−1 day−1, which is 14 times slower than that measured in the short-term incubations. Water incubated with 13CH4 and 15NO2− did produce both 13DIC

(0.9–15.4 nmol) and 29+30N2 (8.9–80.5 nmol), and the relative proportions produced varied across depths with high rates of 29+30N2 production at 235 and 264 m, indicative of nitrite

reduction alongside methanotrophy (Table 2 and Supplementary Figure S2). Methanotrophic bacteria were targeted in waters offshore (30–1250 m depth) and closer to the coast (200 and 228 m).

Analysed aerobic _pmoA_ sequences in the offshore samples (6202 in total) were clustered into six OTUs, hereafter called ETNP_Offshore_MO, and in the inshore samples (363 816 in total) were

clustered into six OTUs, hereafter called ETNP_Inshore_MO. The sequences from both the offshore and inshore samples were highly similar (97–100% BLAST similarity) to uncultured bacteria from

marine environments (Supplementary Table S3, Figure 5a). The vast majority of the offshore sequences are represented by two OTUs, that is, ETNP_Offshore_MO1 (44.16% of sequences) and

ETNP_Offshore_MO2 (47.19% of sequences). Similarly, ETNP_Inshore_MO1 represents the majority (89.43%) of the analysed inshore sequences. Phylogenetic analysis shows that all the OTUs cluster

within known type I methanotrophs (Figure 5a). Among them, three OTUs of the offshore samples (ETNP_Offshore_MO1/MO3/MO5) sit within a sub-cluster of the family Methylococcaceae including

_Methylococcus_ and _Methylomonas_ species. The diversity (based on Shannon and Simpson indices) within all the analysed samples and particularly of the inshore ones is small, with the most

diverse sample being that of 290 m offshore (Shannon=1.19, Simpson=0.64; Supplementary Table S3). The principal coordinate analysis plot also shows a very close proximity of all the inshore

samples (that is, green circles on Figure 5b practically overlap), whereas there is some variance among the offshore samples, as indicated by their good separation along the second principal

coordinate, that is, axis 2, explaining 26.9% of the observed variance (triangles in Figure 5b). However, most of the principal coordinate analysis variance is explained by the first

principal coordinate (axis 1, explaining 70.8% of the variance), which is mainly driven by the divergence of the anaerobic methanotroph community (overlapping purple circles on Figure 5b)

and, to a lesser extent, by the divergence of two offshore aerobic methanotroph samples (30 and 645 m; blue triangles on Figure 5b). The diversity within the two OTUs of the anaerobic

methanotrophs (ETNP_NDAMO1 and ETNP_NDAMO2) is minimal (see Shannon and Simpson indices, Supplementary Table S3). Indeed, phylogenetic analysis placed both of these OTUs into a separate and

well-defined cluster, related to the _Candidatus_ Methylomirabilis oxyfera anaerobic methanotroph (Figure 5a). DISCUSSION Here we have shown that biological methanogenesis, in the surface

layer of the seabed sediments, is a major source of methane to the ETNP OMZ. These are the first direct measurements of methane production in sediments from this region. The reactivity of

these sulphate and nitrate-rich surface sediments highlights the potential importance of non-competitive methanogenesis to the marine methane pool. Our oxygen profiles agree with previously

published data for the ETNP OMZ (Burke et al., 1983; Sansone et al., 2001, 2004; Pack et al., 2015) and show oxygen to be ⩽1.4 μmol l−1 between 200 and 800 m. However, as a secondary nitrite

maximum occurs when oxygen is below 0.05 μmol l−1 (Thamdrup et al., 2012), we used this profile to define the true core of the OMZ (230–600 m). In addition, we have presented evidence for

microbial methane oxidation, which can be sustained under a wide range of oxygen (<1.4–65 μmol l−1) and methane (44–790 nmol l−1) concentrations, potentially controlling the release of

methane emissions from the OMZ. The highest potential for methanogenesis is in the top 2 cm of seabed, in the presence of ample sulphate, nitrate and nitrite (as alternative electron

acceptors) and >20 cm above the hydrogen sulphide peak (Figure 2c, Table 1). The co-occurrence of the greatest potential for methanogenesis and highest concentration of sulphate indicates

that this is likely to be non-competitive methanogenesis and this is supported by the methanogen community findings. The majority of methanogens in all the analysed samples (97.31% of total

sequences) clustered within the family Methanosarcinaceae and the dominant OTUs were similar to _Methanococcoides_ sp. deriving from sub-seafloor sediments (Imachi et al., 2011) or

estuarine mudflats (Watkins et al., 2012). _Methanococcoides_ sp. have often been isolated from marine sediments (for example, Singh et al., 2005; Lazar et al., 2011; Webster et al., 2015)

and they are obligatory methylotrophic methanogens, that is, utilising only non-competitive substrates, such as methanol or methylamines (Garcia et al., 2000; Ferry, 2010). Although we could

not find other direct measurements of methanogenesis in the ETNP OMZ, there are data reported from other locations. Krüger et al. (2005) reported rates of methanogenesis in sediment surface

slurries from eight different marine sites in the Atlantic, Pacific and Arctic Oceans and the North and Baltic Seas, and their results (0.01–0.1 μmol g−1 day−1) agree well with our

measurements using the same technique (0.001–0.09 μmol g−1 day−1, Table 1). They noted that the highest methanogenic potentials were measured in regions with high input of organic matter

from the water column (Krüger et al., 2005). The only intact core experiment (Crill and Martens, 1983) to report marine methane flux was performed on coastal sediments from Cape Lookout

Bight, and showed a similar range (0.18–1.56 μmol m−2 day−1) to those found in our ETNP sediments (0.16–1.01 μmol m−2 day−1). The potential for methanogenesis was markedly reduced below the

top 2 cm and we propose that this is linked to organic carbon supply raining down from above, which the surface methanogens can preferentially access. Continental shelves are known for high

productivity and therefore, the delivery of carbon to the seafloor is high relative to less productive areas of the ocean (Ramaswamy et al., 2008; Fennel, 2010). We propose that this benthic

methanogenesis supplies the water column with methane, which persists far offshore. The location of the methane peak (250–700 m), agrees well with other ETNP studies but the magnitude in

our study (102 nmol l−1 using the conductivity–temperature–depth and 254 nmol l−1 very close to the sediment surface using Mega-Cores) was considerably higher than previously reported

(maximum 5–80 nmol l−1, Burke et al., 1983; Sansone et al., 2001, 2004 and Pack et al., 2015). Variation in maximum concentrations found across the ETNP is likely due to proximity to the

source of methane, dilution and slow microbial oxidation. The flux was greatest when oxygen concentration in the bottom-water was below the LOD (Figure 2a) and a clear plume, originating in

the continental shelf slope and extending offshore, can be seen in our profiles (Supplementary Figure S1), both of which support our theory. Even when there was a wedge of oxygenated water

between the OMZ and the seafloor, methane was supersaturated in the OMZ and the maximum concentration of methane decreased with distance offshore. Indeed, methane was only found to be over

35 nmol l−1 when the maximum water depth was between 350 and 650 m, and in the deeper water (seabed >750 m) the methane did not exceed 25 nmol l−1 even when oxygen and nitrite indicated

true OMZ conditions. The close agreement between our potential methanogenesis rates and the flux data show that benthic methanogenesis could be responsible for all the methane measured in

the bottom-water without the need to invoke additional methane sources, for example, seeps or dissociation of hydrates. Further, to the best of our knowledge, there are no reports of methane

seeps in this OMZ. The spatial alignment of the methane and nitrite peaks suggests that methane could be oxidised, in the presence of nitrite and the absence of measureable oxygen, that is,

anaerobically. Our attempts to measure the potential for N-DAMO were inconclusive, and others (Padilla et al., 2016) using a similar dual-isotope incubation technique, recently tried and

failed to fully quantify this process in the ETNP OMZ. However, in our experiments, water incubated with 13C-CH4 and 15N-NO2 did produce 13C-DIC and 29+30N-N2 but the stoichiometry (Table 2)

was not indicative of pure N-DAMO (3CH4 and 8NO2 produce 3CO2 and 4N2, Ettwig et al., 2010), nor were the rates of 13C-DIC production consistently stimulated by addition of nitrite.

Nonetheless, sequences from N-DAMO-like bacteria were detected in all the targeted water depths. The sequences belonged to just two closely related phylotypes (ETNP_NDAMO_1 and ETNP_NDAMO_2;

Figure 5,Supplementary Table S3) affiliated with uncultured anaerobic methanotrophs from South China Sea sediments (Chen et al., 2014, 2015). They clustered within the _Candidatus_

Methylomirabilis oxyfera lineage, which is known to couple anaerobic methane oxidation to the reduction of nitrite (Ettwig et al., 2010; Haroon et al., 2013) and, although they are well

described in lakes (Deutzmann and Schink, 2011; Kojima et al., 2012), paddy soils (Wang et al., 2012) and peatlands (Zhu et al., 2012), the ecological role of these phylotypes in marine

environments has only recently been addressed (Chen et al., 2014; Li-Dong et al., 2014). More recently, Padilla et al. (2016) reported transcriptionally active Methylomirabilis-like NC10

phylotypes in all their ETNP sites, off the North Mexican coast, with the abundance of 16S rRNA transcripts peaking in the core of the OMZ, thereby confirming marine OMZs as a niche for such

phylotypes. In agreement with recent findings in the South China Sea (Chen et al., 2015), we show that these marine phylotypes form a separate cluster from their equivalent freshwater

phylotypes. We were able to confirm the potential for aerobic methane oxidation in the OMZ of ETNP by measuring the conversion of 13C-CH4 to 13C-DIC over relatively short timescales (<2

weeks). We artificially raised the methane concentration, to ensure that 13C-CH4 (rather than 12C-CH4) constituted the overwhelming majority of the methane available for oxidation. However,

we can use our dose-response experiment to approximate ambient rates of methane oxidation. For example, at 200 m the average methane concentration was 3.3 nmol l−1 so although we measured

4.5 nmol l−1 day−1 (incubated with 300 nmol l−1 CH4) _in situ_ we would expect 0.0495 nmol l−1 day−1 with a turnover time of 67 days. Even in anoxic incubations (LOD for oxygen), 13C-DIC was

produced following a spike of 13C-CH4 and so it is reasonable to predict that methanotrophs are oxidising methane right at the margin of the OMZ core and our measurements fall within the

range recently reported for methane oxidation in the ETNP (0.000034–4 nmol l−1 day−1, Pack et al., 2015). Molecular analysis confirmed the presence of aerobic methane oxidisers at a wide

range of depths (ranging from 30 to 1250 m) in both offshore (ETNP_Offsshore_MO) and coastal (ETNP_Inshore_MO) waters. The majority of methanotrophs from inshore waters (99.96% of sequences)

were phylogenetically related (>97% similarity, Figure 5,Supplementary Table S3) to uncultured bacteria detected in the ETNP (Hayashi et al., 2007). The methanotrophs in the offshore

samples were somewhat more diverse, with some similar to those in the inshore samples and others forming a separate sub-cluster with known Methylococcaceae species (for example,

_Methylomonas methanica_ and _Methylococcus capsulatus_ str. Bath, Figure 5). This could be partly attributed to the difference in the range of depths from which the samples were obtained,

that is, offshore samples were collected from depths between 30 and 1250 m, whereas inshore samples were collected from a much narrower range of depths (200–228 m). Depth-related differences

in the aerobic methanotroph community along vertical water horizons have been reported elsewhere (for example, Tavormina et al., 2010, 2013). Such differences may be related to the physical

transport of waters, harbouring distinct microbial communities, which, along with environmental selection and spatial separation, has been shown to shape the distribution of marine microbes

(Wilkins et al., 2013; Steinle et al., 2015). The diversity of methanotroph phylotypes in the water column is likely controlled by environmental factors rather than geographical proximity,

and the same phylotypes may be adapted to a range of methane concentrations (Tavormina et al., 2008). Indeed, a later study of Cu-MMO phylotypes from the Costa-Rican OMZ showed that methane

concentration did not predict the occurrence, abundance or distribution of any phylotypes; instead environmental factors such as depth, salinity, temperature and dissolved oxygen

concentrations accounted for most of the observed phylotype variance (Tavormina et al., 2013). Aerobic methanotrophs in both offshore and inshore samples clustered within type I, whereas no

sequences were affiliated to type II methanotrophs, which is in accordance with the findings in other marine environments (Schubert et al., 2006; Tavormina et al., 2008; Wasmund et al.,

2009; Schmale et al., 2012). The lack of close affiliation of marine phylotypes with established methanotroph lineages has been reported previously and it has been linked to specialisation

of these phylotypes in marine environments and to the rather small representation of marine methanotroph sequences in public databases (Tavormina et al., 2008; Wasmund et al., 2009). Here we

present clear evidence for microbial methanogenesis in the continental shelf sediments fuelling the ETNP OMZ methane plume, which is sustained several 100 km offshore, despite biological

oxidation. Molecular analyses support the methanogenesis and methanotrophy potentials presented, however, more studies are needed to fully unravel the diversity of pelagic methanotrophs and

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8657–8665. Article  CAS  Google Scholar  Download references ACKNOWLEDGEMENTS This work was funded by the Natural Environment Research Council (grant NE/E01559X/1). We thank Dr Ian Sanders

for general technical assistance, John Green for support with the Dionex and Nele Thijs for DNA extraction of the ETNP_Offshore_MO samples. We appreciate the support of the Captain and crew

of RRS Discovery (cruise D373) and RRS James Cook (cruise JC097). AUTHOR INFORMATION Author notes * Panagiota-Myrsini Chronopoulou Present address: 3Current address: Department of

Environmental Sciences, University of Helsinki, PO Box (Viikinkaari 1), 00014, Helsinki, Finland., * William J Pritchard Present address: 4Current address: Faculty of Life Sciences,

University of Manchester, A4051 Michael Smith Building, Dover Street, Manchester, M13 9PT, UK., * Susanna T Maanoja Present address: 5Current address: Department of Chemistry and

Bioengineering, Tampere University of Technology, PO Box 541, FI-33101, Tampere, Finland., * Panagiota-Myrsini Chronopoulou and Felicity Shelley: These authors contributed equally to this

work. AUTHORS AND AFFILIATIONS * School of Biological and Chemical Sciences, Queen Mary University of London, London, UK Panagiota-Myrsini Chronopoulou, Felicity Shelley, William J

Pritchard, Susanna T Maanoja & Mark Trimmer Authors * Panagiota-Myrsini Chronopoulou View author publications You can also search for this author inPubMed Google Scholar * Felicity

Shelley View author publications You can also search for this author inPubMed Google Scholar * William J Pritchard View author publications You can also search for this author inPubMed 

Google Scholar * Susanna T Maanoja View author publications You can also search for this author inPubMed Google Scholar * Mark Trimmer View author publications You can also search for this

author inPubMed Google Scholar CORRESPONDING AUTHOR Correspondence to Mark Trimmer. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no conflict of interest. ADDITIONAL

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the Eastern Tropical North Pacific oxygen minimum zone. _ISME J_ 11, 1386–1399 (2017). https://doi.org/10.1038/ismej.2017.6 Download citation * Received: 22 June 2016 * Revised: 06 December

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