Seasonal methanotrophy across a hydrological gradient in a freshwater wetland

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Seasonal methanotrophy across a hydrological gradient in a freshwater wetland Taniya Roy Chowdhury a,1 , William J. Mitsch b,2 , Richard P. Dick a,∗ a Soil Microbial Ecology Laboratory, School of Environment & Natural Resources, The Ohio State University, 210 Kottman Hall, 2021 Coffey Road, Columbus, OH 43210, USA b Wilma H. Schiermeier Olentangy River Wetland Research Park, School of Environment & Natural Resources, The Ohio State University, 352 W Dodridge Street, Columbus, OH 43202 USA

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Article history: Received 28 April 2014 Received in revised form 10 August 2014 Accepted 20 August 2014 Available online xxx Keywords: Methane oxidation Wetland Pulsing hydrology Methanotroph PLFA Climate change

a b s t r a c t Wetlands provide significant ecosystem services but are also the largest natural source of methane (CH4 ), a critical greenhouse gas. Oxidation of CH4 in soils/sediments, driven by methanotrophs, offsets CH4 losses to the atmosphere. Manipulation of flooding regimes to optimize methanotrophy is a potential management strategy to reduce CH4 emissions from wetlands and offset global climate change. Therefore, the objectives of this study were to determine rates of potential CH4 oxidation (PMO) and shifts in methanotrophs over hydrological and seasonal gradients. Surface and subsurface soil samples (0–8 or 8–16 cm depths) were analyzed for PMO and profiled for methanotroph community structure using phospholipid fatty acid (PLFA) analysis over four seasons (winter, spring, summer and fall) and 3 landscape positions (upland, intermittently flooded, and permanently flooded sites). PMO rates were highest in the winter. The permanently flooded sites had higher PMO rates than the intermittently flooded sites (p < 0.05). Significantly higher PMO rates were observed in the 0–8 cm compared to the 8–16 cm soil depths (p < 0.05). PLFA profiling of methanotrophs showed that both Type I and Type II methanotrophs were dominant in winter. Concentrations of the Type II methanotroph PLFA (18:␻9c) was significantly higher (p < 0.05) than those for Type I in all seasons and landscape positions. PMO and methanotroph biomass were highest in the winter and in the PFS which suggested substrate (CH4 ) concentration was more important in regulating methanotrophy than redox potential or seasonal shifts in temperature under flooded conditions. PFS had the lowest redox potential (which would not favor aerobic CH4 oxidation), yet it had the highest PMO rates, suggesting anaerobic methane oxidizers may be important in flooded soils. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Wetlands provide significant ecosystem services like flood control, remediation of wastewater and agricultural runoff, and carbon sequestration. These benefits are, however, offset by the fact that wetlands are the largest natural source of methane. Methane (CH4 ) is a critical greenhouse gas and management strategies are necessary not only to limit CH4 emission, but also to facilitate its

∗ Corresponding author. Tel.: +1 614 247 7605; fax: +1 614 292 7432. E-mail addresses: [email protected], [email protected] (R.P. Dick). 1 Current address: Oak Ridge National Laboratory, One Bethel Valley Road, Oak Ridge, TN 37831, USA. 2 Current address: Everglades Wetland Research Park, Florida Gulf Coast University, 110 Kapnick Center, 4940 Bayshore Drive, Naples, FL 34112, USA.

biological sequestration into the soil. Manipulation of the flooding regimes can be a potential management strategy to mitigate CH4 emission especially from constructed wetlands. There is an interest in particularly promoting wetlands with fluctuating or “pulsing” hydrology because of the additional benefits they provide over static wetlands (Mitsch et al., 2005; Roslev and King, 1996). The fringe or pulsing zones of these wetlands provide favorable conditions like higher supply of oxygen for CH4 consumption through oxidation by CH4 oxidizing bacteria or methanotrophs in soil. However, most studies in wetlands have been limited to only quantify the surface CH4 fluxes, and not explored the environmental or hydrological controls on bacterial CH4 consumption in the soil. The ecophysiology of soil microbial communities and their feedback effects on global C fluxes under changing climate is yet unclear (Bardgett et al., 2008), and particularly the metabolic potential of methanotrophs is under-investigated.

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The rate of CH4 emission from an ecosystem is controlled by the rates of two opposing processes: (i) CH4 production in the soil/sediments by methanogenic archaea; and (ii) oxidation of CH4 in the soil/sediments and the water column by methanotrophic bacteria. But the relationship of potential methane oxidation (PMO) to the methanotroph biomass and community structure is not fully explored in wetlands (Siljanen et al., 2011). Methanotrophs are ubiquitous in soil (Liu et al., 2013; Roy Chowdhury and Dick, 2013; Sundh et al., 2000). The biological oxidation of CH4 by methanotrophs, which belong to the ␣-proteobacteria (Type II methanotrophs) and ␥-proteobacteria (Type I methanotrophs), represent the major terrestrial sinks for CH4 . The methanotrophs contain specific phospholipid fatty acids (PLFAs) of 16 (Type I methanotrophs) or 18 (Type II) carbon (C) atoms in chain length (Bowman et al., 1991). A number of such methanotrophic biomarker PLFA have been identified using stable isotope probing in numerous studies in a diversity of study sites and previously listed (Roy Chowdhury and Dick, 2013). These PLFAs have also been used to estimate the relative abundance of Type I and II methanotrophs in environments well supplied with CH4 (Borjesson et al., 1998; Nichols et al., 1987; Sundh et al., 1995b). In this study, we studied the dynamics of the soil methanotroph community structure using the biomarker PLFAs and their relationship to CH4 oxidation potential from wetlands in response to “pulsing” hydrology. Our objectives were (i) to estimate the rates of CH4 oxidation in response to seasonal fluctuations in temperature and water-levels in two freshwater wetlands; (ii) study the relative shifts in the methanotrophic community structure along seasonally induced hydrologic gradients; and (iii) understand the relative response within the methanotrophic communities. 2. Materials and methods 2.1. Description of field site Two experimental wetlands located at the Wilma H. Schiermier Olentangy River Wetland Research Park (ORW) at The Ohio State University, Columbus, Ohio (Mitsch, 2005) were chosen as the site for this study. Details about the ORW can be found in several studies (Mitsch et al., 1998, 2005, 2012). The two wetlands are field-scale replicates with the same basin morphology; the western basin (W1) was planted with 13 native species of macrophytes in May 1994 while the eastern basin (W2) was allowed to naturally colonize. Water from the Olentangy River, a fourth-order river that drains the central part of an agricultural/urban watershed in Ohio, is controlled by pumps into both the experimental wetlands (Mitsch et al., 1998; Nahlik and Mitsch, 2010). During this study period of 2008–2010, the average annual inflow rates ranged from 975 to 1550 m3 d−1 (36–57 m yr−1 ), total nitrogen inflow concentrations averaged 2.85 mg-N L−1 and total phosphorus concentrations averaged 160 ␮g-P L−1 (Mitsch et al., 2012; Nahlik and Mitsch, 2010, 2011). 2.2. Experimental design and soil sampling The experimental design was a randomized block design with the following treatments: three hydrologically distinct landscape positions viz., Upland Site (UPS), Intermittently Flooded Site (IFS) and the Permanently Flooded Site (PFS) in two replicate wetlands W1 and W2, four seasons (winter, spring, summer and fall) and two soil depths viz., 0–8 and 8–16 cm. The IFS were located at 0–15 cm closer to the edge of the wetlands and within the wetland boundaries. This landscape position was subjected to pulsed flooding with water levels ranging from 10 to 20 cm during spring and fall, and

<1-cm during summer. The soil surface in the UPS and IFS was nearly frozen to an average depth of ∼5-cm during winter sampling. The PFS had water levels >20 cm throughout the year and were located near the middle of the wetlands. Soil samples were collected over a two-year period, from October 2008 to October 2010. During the entire study period, pumping rates were adjusted daily according to a predetermined calculation based on the flow of the Olentangy River. This was done to ensure that the wetlands experienced conditions similar to naturally occurring riverine wetlands and the same amount of water was introduced to each wetland (Nahlik and Mitsch, 2010). From November to April, during typically cold (mean air temperature of 4 ◦ C) and wet conditions in Ohio, the wetlands generally experience more frequent flooding and higher water levels. During the warm (mean air temperature of 21 ◦ C) and dry seasons from May to October, low water levels prevail in the wetlands due to reduced river flow. The seasonal sampling times were: winter (December–January–February), spring (March–April–May), summer (June–July–August) and fall (September–October–November). Soil samples were collected twice every season to capture any effects of short-term flooding within a season (Table 1). The withinseason samplings were separated by 3–4 weeks contingent on flooding events and water levels. Soil samples were collected from the inflow and the outflow of both wetlands W1 and W2. A minimum of 20 soil cores were collected to obtain a representative soil sample for each landscape position and depth. Soil cores were collected using a stainless-steel soil probe and placed in zip-lock bags. Soil samples were immediately sieved and homogenized by passing through a 2-mm mesh screen and stored at appropriate temperatures until analysis. Samples for analysis of PMO rates were stored at 4 ◦ C, and those for PLFA analysis were stored at −20 ◦ C. Potential methane oxidation rates were measured within a day of soil sampling. Phospholipid fatty acids were extracted within a week after soil collection. 2.3. Potential methane oxidation (PMO) rates measurement Potential methane oxidation rates were measured according to modified methods after Crossman et al. (2004) and Sundh et al. (1995a). 10 mL of standard 99.99% CH4 gas was injected into a polytetrafluoroethylene (PTFE) gas sampling bag to prepare standard calibration curves. Appropriate volumes of the standard CH4 gas were taken to prepare four calibration standards of concentrations 0.2, 0.25, 0.3 and 0.35% prepared in 125 mL Wheaton serum bottles with butyl rubber stoppers equipped with crimps and purged with N2 gas. The standards were run in a staggered manner at the beginning and end of the run of each set of samples by gas chromatography and concentrations calculation based on linear regression of five-point standard curves. Potential methane oxidation rates were measured in triplicates for each soil sample. Fifteen grams of homogenized soil was transferred to 125 mL Wheaton serum bottles and 15 mL of deionized water was added and shaken vigorously; the bottles were evacuated and refilled with air in three cycles. 0.32 mL of the standard CH4 was added to create a final concentration of 0.35% (v/v) CH4 . The flasks were incubated at 25 ◦ C, shaken horizontally at 150 rpm, and the CH4 concentration in the gas phase was monitored every hour for 8 h. Gas samples were removed (0.75 mL to optimize detection and sensitivity) from the incubation bottles using a syringe and analyzed by gas chromatography as described below. PMO rate was subsequently obtained from a linear regression fitted to all measurements above 0.1%. The PMO rates are reported as nmol CH4 g−1 dry weight soil h−1 . A gas chromatograph (Shimadzu GC-2010, Kyoto, Tokyo) with a RT-QPLOT column was used to measure CH4 . The column was

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Table 1 Physico-chemical characteristics of soil. Season

Sampling site description

Depth (cm)

pHwater (1:1)

Total carbon (g C kg−1 )

Redox potential (mV)

Gravimetric water content (g H2 O g−1 soil)

Winter

Intermittently Flooded Site (IFS) Permanently Flooded Site (PFS) Upland (UP)

0–8 8–16 0–8 8–16 0–8 8–16

6.3 6.5 6.6 6.6 6.5 6.2

6.67 7.12 6.97 6.8 5.63 5.43

−52 −88 −275 −400 45 60

0.58 0.46 0.52 0.27 0.36 0.35

Spring

Intermittently Flooded Site (IFS) Permanently Flooded Site (PFS) Upland (UP)

0–8 8–16 0–8 8–16 0–8 8–16

6.2 6.4 6.7 6.7 6.5 6.1

6.12 8.32 7.21 7.62 4.94 5.33

−80 −110 −310 −348 33 45

0.62 0.33 0.50 0.29 0.33 0.42

Summer

Intermittently Flooded Site (IFS) Permanently Flooded Site (PFS) Upland (UP)

0–8 8–16 0–8 8–16 0–8 8–16

6.2 6.1 6.6 6.6 6.4 6.3

5.95 5.76 4.52 4.66 3.95 3.62

49 −120 −220 −310 48 52

0.45 0.45 0.58 0.49 0.51 0.47

Fall

Intermittently Flooded Site (IFS) Permanently Flooded Site (PFS) Upland (UP)

0–8 8–16 0–8 8–16 0–8 8–16

6.3 6.3 6.6 6.6 6.5 6.5

6.22 7.29 6.98 7.23 5.50 5.76

−115 −130 −330 −350 72 58

0.59 0.57 0.34 0.37 0.44 0.49

isothermally maintained at 40 ◦ C. The carrier gas was helium. The injector and the detector (Flame Ionization Detector) were kept isothermally at 150 ◦ C. Hydrogen flow was maintained at 45 mL min−1 and that of synthetic air at 450 mL min−1 . Both hydrogen and helium used were of Ultra-High purity grade and hydrocarbon-free grade air was used. The supporting program for data analysis used was EZ-Start.

Standard nomenclature for the FAMEs include the number of C atoms counted from the omega (␻) end (i.e., opposite the carboxyl end), followed by the number of double bonds after the colon; cis conformations are designated with the suffix c, and the prefixes i and a are given for iso- and anteiso-branched FAMEs, respectively.

2.5. Statistical analysis 2.4. Phospholipid fatty acid (PLFA) analysis A modified version of the PLFA method of Bligh and Dyer (1959) and Roy Chowdhury and Dick (2012) was used. Briefly, lipids were extracted from the soil (4 g) by shaking in 35 mL glass vials equipped with Teflon-lined screw caps for 2 h in a solvent phase consisting of up to 1.5 mL of aqueous citrate buffer (0.15 M, pH 4.0), 1.9 mL of chloroform (CHCl3 ), and 3.8 mL of methanol (MeOH) and 2 mL Bligh and Dyer reagent (chloroform:methanol:aqueous citrate buffer mixed in the proportions 1:2:0.8 (v/v/v)). The phospholipid fraction (methanol fraction) was dried down under mild stream of N2 and re-dissolved in 1 mL of 1:1 (v/v) methanol/toluene. This fraction was used for the PLFA profiling. The PLFAs were redissolved in 1 mL of toluene and subjected to a mild alkaline transmethylation with 1 mL addition of methanolic-KOH at 37 ◦ C for 15 min, followed by cooling to room temperature. The PLFAs were quantified by gas chromatograph (HewlettPackard model 5890 Series II) equipped with a HP Ultra 2 capillary column and a flame ionization detector. Fatty acids were identified by the Microbial Identification Index System, Version 5.0 (MIDI Inc., Newark, DE). The GC temperature program ramped from 170 to 270 ◦ C at 5 ◦ C min−1 . Following the analysis, a ballistic increase to 300 ◦ C allows cleaning of the column during a hold of 2 min. Hydrogen was the carrier gas, nitrogen the “makeup” gas, and air is used to support the flame. Peak areas were used to quantify each PLFA relative to the internal standard (19:0). Method blanks were extracted with each set of samples and were assumed to be free of contamination if chromatograms of the blanks contained no peaks.

Data were tested for normality assumptions using Shapiro–Wilke tests, confirming which post hoc tests (Tukey’s Honestly Significant Difference) were done to determine significant difference of means. Analyses of variance to determine significant differences between means were performed with Tukey’s post hoc HSD test. In most cases the distribution was non-normal, and so the data were subjected to non-parametric tests. Kruskal–Wallis tests (alpha = 0.05) were used to examine the treatment effects on the PMO rates for the following treatments: landscape position, considering the three levels (Intermittently Flooded Site, Permanently Flooded Site and Upland Site); four Seasons and the methanotroph type (Type I and Type II). All statistical analyses of data were performed using SAS ver. 9.1 (SAS Institute Inc., USA).

3. Results 3.1. Soil physicochemical properties Representative samples from each site and soil depth were analyzed for gravimetric water content, pH, soil texture and total carbon. The gravimetric water content of soils was determined by weighing subsamples before and after oven drying at 105 ◦ C to constant weight (n = 3). pH was measured using a Accumet Basic AB15 bench pH meter (Fisher Scientific Inc.) using the slurry method (mixing 1:1, w:v ratio of soil to de-ionized water, n = 3). Soil physicochemical properties are reported in Table 2. Results for the seasonal patterns of soil temperature (◦ C), water level (cm) and

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Table 2 Pearson’s correlation coefficients between PMO and the relative abundance of Type II methanotrophs to Type I methanotrophs (Type II/Type I). Soil depth (cm)

Winter

Spring

Summer

Fall

Inflow

0–8 8–16

0.72 0.87

0.84 0.66

−0.96 0.6

−0.84 0.51

Outflow

0–8 8–16

0.93 0.89

0.89 0.62

−0.99 0.48

−0.89 0.73

Overall, both Type I and II methanotroph biomarker concentrations exhibited distinct seasonal dynamics and positively co-varied with PMO rates at sites where they both could be measured (Fig. 3a–f). It is noteworthy that negative or poor correlation between PMO and concentrations of methanotroph PLFAs was observed at those sites where the methanotroph Type II biomarkers were absent (Table 2). This was especially true in summer and fall. A pattern in the distribution of Type I and Type II methanotrophs and season was observed showing an increase in Type II methanotroph biomass with apparent and relative dissolved oxygen availability, in turn a function of inundation vs. exposed surface (Fig. 3e and f).

3.3. Potential methane oxidation (PMO) rates The rates of potential CH4 oxidation over the four seasons ranged from 20.6 to 11.8 at 0–8 cm and 8.5 to 5.6 at 8–16 cm depths for the PFS, 18.7 to 6.8 at 0–8 cm and 17.8 to 4.8 at 8–16 cm depths for the IFS, and 14.2 to 5.7 at 0–8 cm and 13.7 to 3 at 8–16 cm depths for the UPS. The highest PMO rate at all landscape position was observed in winter and was statistically higher in the PFS. A seasonal difference in PMO rates was observed at all landscape positions but were more obvious in the PFS. At 0–8 cm depth, the PFS had significantly lower PMO rates in summer (Fig. 2). At 8–16 cm depth, the PMO rates in the PFS were significantly lower compared to the 0–8 cm depth in IFS in all seasons. The UPS had more consistent PMO rates between the two soil depths in all seasons except in winter. The IFS site showed considerable depth effects on PMO rates. The greatest difference in PMO rates between depths was seen for the PFS.

4. Discussion 4.1. Landscape position and hydrology controls on methanotrophy

Fig. 1. Seasonal variation in (a) Type I and (b) Type II methanotroph phospholipid fatty acid concentration across landscape position.

CH4 emissions (mg CH4 -C m−2 h−1 ) for the two wetlands (W1 and W2) can be found in Nahlik and Mitsch (2010). 3.2. Distribution of Type I and Type II methanotrophs Distribution of phospholipid fatty acids (PLFA) characteristic of Type I (16:1C; Fig. 1a) and Type II (18:1C; Fig. 1b) methanotrophs varied between soil depths and landscape position. Temporal variation of the methanotrophic biomass controlled by seasonal changes of temperature and water levels was remarkable. Both Type I and Type II methanotroph PLFA concentrations were significantly higher (p < 0.001) at 0–8 than at 8–16 cm soil depths at the permanently flooded site in winter. Interestingly, Type II methanotroph PLFA concentrations were significantly higher (p < 0.007) in the surface layer (0–8 cm) at permanently flooded site in winter (Fig. 3e and f) and at upland at the proximity of the wetland inflow. However, no statistically significant differences were observed between the distributions of Type I and Type II methanotroph PLFA concentrations at the other sites in winter. In contrast, in summer both Type I and II methanotrophs were significantly higher in the intermittently flooded site at 0–8 cm in both inflow and outflow of the wetlands.

Seasonal fluctuations in water levels typical of the IFS resulting in short-term saturated (in fall, winter and spring) and unsaturated conditions (in summer) have previously been reported to increase CH4 flux (Roslev and King, 1995) and oxidation (Kettunen et al., 1999). In a peat wetland, Hughes et al. (1999) found that an imposed drought caused by draining of the surface layer and lowering of the water table initially resulted in an increase in CH4 flux with no net change in CH4 oxidation (Freeman et al., 2002). Our results of PMO did not follow similar trends as the IFS site had lower PMO levels than the permanently flooded sites and indeed it would be expected that PMO rates should be higher in the aerobic zones. In contrast, our results suggest substrate concentration (CH4 ) was the major controller of both PMO and the biomass of Type I and II methanotroph communities. Indeed, the redox potential measurements (Table 1) became more negative from UPS to IFS (albeit with levels fluctuating, seasonally) to PFS locations where the prolonged reduced conditions of PFS (−200 to 230 mV) would be the most conducive for methanogenesis. These observations are strongly supported by the reports of Altor and Mitsch (2006) and Nahlik and Mitsch (2010) who at the same wetlands and sampling sites as our study, reported the highest CH4 fluxes at the PFS followed by IFS and UPS locations. Another factor unique to the IFS location which exhibited reduced subsurface PMO rates in summer could be attributed to physical changes of soil in this fluctuating hydrologic zone. Rapid degassing of CH4 trapped in the subsurface (8–16 cm) of the IFS as a result of draw-down and de-saturation in summer may bypass the CH4 oxidizing bacteria. Small decreases in water levels (<5 cm)

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Fig. 2. Seasonal variation in potential methane oxidation. † Bars within a landscape position and soil depth are not significantly different at p < 0.05 when indicated by same letters.

decrease oxygen availability and thereby methanotrophy as previously shown by Roslev and King (1995). Additionally, as water recedes and the soil profile drains, combined with evaporation, soil compaction may occur. This is because macropores shrink and the soil consolidates with the loss of water. Such compression is another possible mechanism that would increase the oxygen consumption per volume of soil (or peat), decreasing oxygen diffusivity, increasing soil matric potential and resulting in anoxic microsites (Teh et al., 2005; Verchot et al., 2000). Methane oxidation in these rapidly created microsites would thus be limited by the transport of CH4 from source to sink and by mass transfer (Bogner et al., 1997; Whalen and Reeburgh, 1990) in the seasonally saturated and pulsed IFS soils. Overall, CH4 oxidation rates were more sensitive to CH4 concentration rather than O2 availability, suggesting that methanotrophs may respond strongly to in situ fluctuations in CH4 production. Aquatic vegetation is an important parameter that affects CH4 dynamics in wetlands. Methanotrophic activity in submerged freshwater marshes is predominantly found in the oxic rhizosphere of macrophytes (Gerard and Chanton, 1993; Hirota et al., 2004; Murase and Frenzel, 2007) and as such is an important factor that might control CH4 oxidation in situ. However, Gerard and Chanton (1993) argued that enhanced CH4 transport via the aerenchyma structure of aquatic plants enables CH4 to bypass rhizosphere methanotrophs and is of greater significance than enhanced CH4 oxidation in the rhizosphere. Furthermore, there can be competition for oxygen transported to the rhizosphere by root respiration, aerobic microbial metabolism, and ferrous iron oxidation (Chanton et al., 1992). However, previous studies (Roura-Carol and Freeman, 1999) have suggested that this competition between CH4 transport to the atmosphere vs. oxygen transport to support CH4 oxidation varies with aquatic plant species and the process that dominates and ultimately affects net CH4 flux depends on the composition of aquatic vegetation species. The effect of aquatic plants and their associated root exoderms and aerenchyma transport system would only be relevant at the

IFS soils (dominated by Schoenoplectus tabernaemontani, Typha latifolia L., T. augustifolia L., Juncus effusus to name a few) in this study, as the UPS plant composition do not perform this process and the PFS sampling location had open water and no macrophytes. It is important to note here that previous studies at ORW did not detect significant CH4 emission due to presence of emergent macrophytes in the intermittently flooded site while reporting 70% less CH4 in these exposed soils when compared to the permanently flooded conditions (Altor and Mitsch, 2006). Thus, for the IFS sites the lower CH4 emission reported by Nahlik and Mitsch (2010) and our lower PMO rates compared to the PFS could be in part affected by the presence of aquatic plants. By transporting oxygen to the rhizosphere, this could suppress methanogenesis which in turn limited substrate (CH4 ) availability to stimulate methanotrophy. This observation in juxtaposition to the PFS for the 8–16 cm depth further highlights that CH4 oxidation can flourish under microaerophilic (van Bodegom et al., 2001) and oxygen-limiting conditions (Roslev and King, 1995). 4.2. Seasonal and temperature relationships Potential methane oxidation (PMO) rates measured under laboratory conditions were highest in the winter samples from all landscape positions (mean air temperature < 4 ◦ C). Methane oxidation rates were significantly higher at 0–8 cm in the PFS followed by the IFS. These results are particularly interesting because at the time of sampling the IFS in winter, the ∼0–4 cm depth had been under frozen conditions for ∼45 days compared to the 8–16 cm depth where the soils were not frozen. At the PFS, the water surface is frozen but not the sediment and/or the water column below it. The high PMO rates measured for PFS-0–8 cm (Fig. 3f) suggest absence of any diffusion barrier and favorable availability of dissolved oxygen for aerobic CH4 oxidation (King, 1990; Megonigal and Schlesinger, 2002). The low redox potentials at 8–16 cm depth also confirm that conditions are conducive to methanogenesis. Methane fluxes from freshwater environments

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Fig. 3. Dynamics of methane oxidation and PLFA concentration with soil depth.

are reportedly dominated by ebullition (Bastviken et al., 2011) in all seasons except winter (Segarra et al., 2013) when the amount of dissolved CH4 would be higher than summer due to increased solubility (Chanton et al., 1989). In other words, seasonally driven lower winter temperatures create conditions of higher substrate availability for methanotrophs in the dissolved phase/absorbed in sediment pores in contrast to accelerated loss from wetland water column by ebullition in the summer. In fall (Fig. 3c and d) and spring, PMO patterns across the landscape positions were similar. The active methanotrophic community under the temperature and moisture conditions of spring and late fall may be a mix of mesophiles and psychrotolerants (Liebner and Wagner, 2007). The methanotrophic community would be expected to shift toward the mesophiles in summer with mean air temperatures ranging between 30 and 35 ◦ C, and

shallow water column at the PFS (≤10 cm) and IFS (≤2 cm when wet following rainfall). The PLFA biomarkers of methanotrophs gradually declined in spring to summer and fall (Fig. 1). These results suggest that the seasonal lowering of water levels in the PFS reduces CH4 fluxes. Conversely, seasonal pulsing enhances greater diffusion/advection of O2 into the oxic–anoxic interface thereby lowering methanogenesis and promoting CH4 oxidation. The positive correlation between the PMO rates and the ratio of Type II/Type I methanotroph PLFAs concentrations (Table 2) provide indirect evidence for this phenomenon as further discussed below. The lack of a positive relationship between warmer seasonal temperatures with methanotrophic activity reported here is consistent with previous research directly focused on methanotrophic responses to temperature. Although recent studies report higher temperature conditions to generally enhance both bacterial CH4

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oxidation and archaeal methanogenesis (Kip et al., 2010; van Winden et al., 2012), studies in freshwater wetlands (Kelley et al., 1995; Neubauer et al., 2000) showed only weak reliance of CH4 fluxes on temperature (Segarra et al., 2013). Furthermore, the effect of soil temperature on CH4 oxidation estimated at Q10 values between 1.4 and 1.9 was shown to be negligible (Crill et al., 1994; Segers, 1998) and increases in CH4 oxidation rates in summer are attributed to drier soil conditions and temperature has only a secondary effect (Brumme and Borken, 1999). In a recent study, van Winden et al. (2012) showed reduced consumption by methanotrophs at temperature optimum of 20 ◦ C due to reduced solubility of CH4 with increasing temperature. While the apparent warm-season decline in CH4 oxidation rates suggests that the process is O2 -limited (King, 1996), the linear relationship between CH4 oxidation and gross CH4 emission suggests substrate availability (CH4 ) is a major controller of methanotrophy (Megonigal and Schlesinger, 2002). Our results uniquely show that seasonal-responses of CH4 oxidation approximately follow that of CH4 emission as reported by Nahlik and Mitsch (2010), and are likely controlled by seasonally induced redox fluctuations and methanotroph community dynamics. 4.3. Seasonal dynamics of methanotroph community Biomass estimation using biomarker PLFAs for Type I and II methanotrophs for the 0–8 cm depth were significantly higher in the PFS than the IFS sites in winter. It is important to note here that the detection and estimation of active methanotrophs in these environments using PLFA analyses are limited by known methanotrophic PLFA profiles and are often rich in the 18:1␻7c PLFA biomarker, but lack the fingerprint 18:1␻8c or 16:1␻8c fatty acids (Dedysh et al., 2001, 2004). As much as 65% of the cells specifically profiled using oligonucleotide probes for methanotrophs are reported as unclassified bacteria (Kobabe et al., 2004). These unidentifiable groups of methanotrophs (not detectable by the known Type I and II methanotrophic-PLFA biomarkers) might represent the recently discovered denitrifying ammonia oxidizers (Wu et al., 2013) and anaerobic CH4 oxidizers (Smemo and Yavitt, 2007) which our PLFA methods did not detect. These organisms have not been cultured and only been identified phylogenetically in freshwater environments, canals, lake sediments, wastewater treatment plants, and peatlands (Callaghan, 2013; Deutzmann and Schink, 2011; Gupta et al., 2013; He et al., 2012; Zhu et al., 2010). These specialized organisms may explain the high PMO rates we measured at the PFS, which is the most anoxic site. Examining the differences in 16:1C and 18:1C PLFA profiles at the two depths and three landscape positions in winter (Fig. 1) showed that either all methanotrophic species are not equally adaptive to the cold temperatures, or biochemical changes in their membrane lipids occur under temperature influences. We suggest that the psychrotolerant methanotrophs predominate at the IFS in winter and are sparse in the PFS where the water column temperature is above freezing-point, but future studies need to be carried out to test this. The strong activity of these specialized groups of methanotrophs is reflected in the highest PMO rates at 0–8 cm in the PFS. Our results are consistent with the previous observations of Liebner and Wagner (2007), a phenomenon that could be due to adaptation or a shift in methanotroph species abundance to lower sediment temperatures during the winter. The seasonal shifts of methanotrophy are more closely related to CH4 production, meaning methanotrophs are stimulated by the substrate (CH4 ) that results in a net increase in methanotrophic biomass. Regulation of aerobic CH4 oxidation by bacteria is controlled by the physiological requirement for O2 and CH4 . Type I methanotrophs were reported to be more active in the surface layers

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(Evershed et al., 2006), where O2 concentrations were the highest and CH4 concentrations were low. In contrast, the Type II methanotrophs dominated the deeper layers where CH4 concentrations were higher and oxygen availability was low. Our observation is further supported by changes in the PLFA profiles at the depths of 0–8 cm and 8–16 cm, respectively. We observed increased amounts of unsaturated and short-chain PLFAs in the deeper (8–16 cm) compared with that in the active layer above (0–8 cm) (data not shown). One of the most important results of this study is the strong positive correlation between the potential CH4 oxidation rates and the relative abundance of methanotrophic community (Type II/Type I) as reflected by the biomarker PLFA concentrations (Table 2). The Type II over Type I methanotrophs profiled with PLFA markers in our study, appear to be the “drivers” of CH4 oxidation under the favorable conditions of high CH4 concentration. Evidence for this is that the PMO rates and the ratio of Type II/Type I PLFA concentrations had similar seasonal and depth distributions. This is further supported by the strong positive correlation of PMO rates and Type II PLFAs (Table 2). Bogner et al. (1997) speculated that methanotrophs first deplete the dissolved CH4 in pore waters that in turn stimulate methanotrophic activity. During the winter at PFS, the water column over the sediment surface is frozen for nearly 2 months. This limits gas diffusional exchange with the atmosphere. In a situation of continued methanogenesis and diffusion of CH4 from the 8–16 cm to the 0–8 cm sediment depth, oxygen availability in the watersediment interface is limited and likely very rapidly consumed by the oxidizing microorganism. These conditions would result in methanotrophs oxidizing a larger fraction of the dissolved CH4 pool at the 0–8 cm as reflected in the PLFA concentrations, particularly by the Type II methanotrophs (Fig. 1). On the other hand, methanotrophic uptake of the gas may have exceeded mass transfer rates in the micro-environments where CH4 availability was low (assuming constrained diffusion of CH4 from the 8–16 cm to the 0–8 cm), resulting in more complete oxidation of the dissolved CH4 pool. The depth-dependent differences in methanotroph biomass clearly observed at the IFS locations could also be due to the differences in the C metabolism efficiencies of the Type I and Type II methanotrophs. The Type I methanotrophs (RuMP pathway) have greater C assimilation efficiency than Type II (Serine pathway) methanotrophs (Hanson and Hanson, 1996). Also, Type II (or the low-affinity) methanotroph populations are known to be strongly related to soil porewater CH4 concentrations (Costello et al., 2002). Our results show the dominance of the high-affinity Type I methanotrophs in the 0–8 cm depth and of the Type II in the 8–16 cm depth, observations similar to that reported in highly reduced and cold soil condition (Wagner et al., 2005). Therefore, each soil depth represents dynamic changes in biogeochemical conditions and establishes links between specialized microbial population and functional acclimation (Hughes et al., 1999; Kim et al., 2008).

5. Implications and perspectives We demonstrated that within the saturated zones (the pulsing and the permanently flooded zones) the abundance and distribution of CH4 oxidizing bacteria are closely related to seasonally controlled sediment saturation and redox conditions and negatively to seasonal temperature. This temporal pattern of methane oxidation and methanotroph biomass corresponded to CH4 emission fluxes reported by Altor and Mitsch (2006) with the permanently flooded zones being highest in the non-growing season. This provides indirect evidence that substrate (CH4 ) concentration is the dominant factor in controlling methanotrophy on a seasonal. This does not mean necessarily that temperature is

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not a controlling factor in methanotrophy (although the literature is inconsistent at this point), but rather that under field conditions, substrate concentration (CH4 ) has a much greater influence than temperature. We found a strong relationship of PMO rates with methanotroph biomarker PLFA concentrations over seasons and soil depth. This showed that the Type I and II PLFA biomarkers are good indicators of aerobic methanotrophs in wetland systems. Methane dynamics is very complex in soils because of the heterogeneity in spatial distribution of anoxic and oxic microsites that provide niche for the counteracting processes of methanogenesis and methanotrophy. Our results are promising to support the concept of “pulsing” wetlands as a management strategy to facilitate biological CH4 consumption. Finally, to improve estimates of net CH4 emission from freshwater wetland systems our results suggest that it is important to include the dynamic effects of depth-dependent methane oxidation potential linked to methanotroph biomass. Under changing climate scenarios it is urgent that process-based models include the nuanced effects of environmental factors and landscape position in controlling methanotrophy and ultimately net CH4 losses from wetlands. References Altor, A.E., Mitsch, W.J., 2006. Methane flux from created riparian marshes: relationship to intermittent versus continuous inundation and emergent macrophytes. Ecol. Eng. 28, 224–234. Bardgett, R.D., Freeman, C., Ostle, N.J., 2008. Microbial contributions to climate change through carbon cycle feedbacks. ISME J. 2, 805–814. Bastviken, D., Tranvik, L.J., Downing, J.A., Crill, P.M., Enrich-Prast, A., 2011. Freshwater methane emissions offset the continental carbon sink. Science 331, 50. Bligh, E.G., Dyer, W.J., 1959. A rapid method of total lipid extraction and purification. Can. J. Biochem. Phys. 37, 911–917. Bogner, J.E., Spokas, K.A., Burton, E.A., 1997. Kinetics of methane oxidation in a landfill cover soil: temporal variations, a whole landfill oxidation experiment, and modeling of net CH4 emissions. Environ. Sci. Technol. 31, 2504–2514. Borjesson, G., Sundh, I., Tunlid, A., Svensson, B.H., 1998. Methane oxidation in landfill cover soils, as revealed by potential oxidation measurements and phospholipid fatty acid analyses. Soil Biol. Biochem. 30, 1423–1433. Bowman, J.P., Skerratt, J.H., Nichols, P.D., Sly, L.I., 1991. Phospholipid fatty-acid and lipopolysaccharide fatty-acid signature lipids in methane-utilizing bacteria. FEMS Microbiol. Ecol. 85, 15–22. Brumme, R., Borken, W., 1999. Site variation in methane oxidation as affected by atmospheric deposition and type of temperate forest ecosystem. Glob. Biogeochem. Cycle 13, 493–501. Callaghan, A.V., 2013. Enzymes involved in the anaerobic oxidation of n-alkanes: from methane to long-chain paraffins. Front. Microbiol. 4. Chanton, J.P., Martens, C.S., Kelley, C.A., 1989. Gas-transport from methanesaturated, tidal fresh-water and wetland sediments. Limnol. Oceanogr. 34, 807–819. Chanton, J.P., Martens, C.S., Kelley, C.A., Crill, P.M., Showers, W.J., 1992. Methane transport mechanisms and isotopic fractionation in emergent macrophytes of an Alaskan Tundra Lake. J. Geophys. Res. Atmos. 97, 16681–16688. Costello, A.M., Auman, A.J., Macalady, J.L., Scow, K.M., Lidstrom, M.E., 2002. Estimation of methanotroph abundance in a freshwater lake sediment. Environ. Microbiol. 4, 443–450. Crill, P.M., Martikainen, P.J., Nykanen, H., Silvola, J., 1994. Temperature and Nfertilization effects on methane oxidation in a drained peatland soil. Soil Biol. Biochem. 26, 1331–1339. Crossman, Z.M., Abraham, F., Evershed, R.P., 2004. Stable isotope pulse-chasing and compound specific stable carbon isotope analysis of phospholipid fatty acids to assess methane oxidizing bacterial populations in landfill cover soils. Environ. Sci. Technol. 38, 1359–1367. Dedysh, S.N., Horz, H.P., Dunfield, P.F., Liesack, W., 2001. A novel pmoA lineage represented by the acidophilic methanotrophic bacterium Methylocapsa acidophila B2. Arch. Microbiol. 177, 117–121. Dedysh, S.N., Berestovskaya, Y.Y., Vasylieva, L.V., Belova, S.E., Khmelenina, V.N., Suzina, N.E., et al., 2004. Methylocella tundrae sp nov., a novel methanotrophic bacterium from acidic tundra peatlands. Int. J. Syst. Evol. Microbiol. 54, 151–156. Deutzmann, J.S., Schink, B., 2011. Anaerobic oxidation of methane in sediments of lake Constance, an oligotrophic freshwater lake. Appl. Environ. Microbiol. 77, 4429–4436. Evershed, R.P., Crossman, Z.M., Bull, I.D., Mottram, H., Dungait, J.A.J., Maxfield, P.J., et al., 2006. C-13-labelling of lipids to investigate microbial communities in the environment. Curr. Opin. Biotechnol. 17, 72–82. Freeman, C., Nevison, G.B., Kang, H., Hughes, S., Reynolds, B., Hudson, J.A., 2002. Contrasted effects of simulated drought on the production and oxidation of methane in a mid-Wales wetland. Soil Biol. Biochem. 34, 61–67.

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