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An Insight Into the Diversity of Aerobic Methanotrophs in Different Habitats

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An Insight Into the Diversity of Aerobic Methanotrophs in Different Habitats AN INSIGHT INTO THE DIVERSITY OF AEROBIC METHANOTROPHS IN DIFFERENT HABITATS PRINCE MATHAI [email protected] MICROBIAL DIVERSITY 2013 ABSTRACT: Methane is an important greenhouse gas and atmospheric concentrations of methane have risen 2.6-fold since preindustrial times. Methane-oxidizing bacteria (aka methanotrophs) are a sink for methane and are found in different habitats such as natural wetlands, rice paddies and peat bogs. However, knowledge regarding the diversity and activity of these microorganisms is limited and thus, requires further investigation. In this study, a combination of culture-dependent and –independent techniques was employed to explore the diversity of aerobic methanotrophs in two distinct methanogenic environments in Woods Hole, Massachusetts: Cedar Swamp and School Street Marsh. Multiple microcosms were established and fed different concentrations (0.1%, 1% & 10%) of methane. This resulted in enrichment cultures which exhibited high degrees of methanotrophic activity, which was confirmed using gas chromatography and microscopy techniques (e.g. CARD-FISH). Stable isotope probing was performed to identify active methanotrophs. Furthermore, dilution to extinction and solid media plating experiments were performed to isolate these microorganisms. 1 INTRODUCTION: Methanotrophs are a unique group of obligately aerobic, gram-negative bacteria which utilize methane as the sole source of carbon and energy. This is feasible due to the presence of an enzyme ‘methane monooxygenase’ (MMO) which occurs, either in a particulate (membrane- bound; pMMO) or soluble (cytoplasmic; sMMO) form (Bowman et al, 2006). MMO oxidizes methane to methanol, and then methanol is further oxidized to formaldehyde, which methanotrophs use for cellular carbon. Due to their ability to oxidize methane, these bacteria can significantly reduce methane emissions to the atmosphere and play a crucial role in the global methane cycle (Martineau et al, 2010). The presence of pMMO has been reported in all methanotrophs except genus Methylocella (Theisen et al, 2005), whereas sMMO appears to occur only in certain methanotroph strains (Murrel et al, 2000). Based on their cell morphology, ultra-structure and metabolic pathways, most known methanotrophs can be classified into two families: Methylococcaceae (type I) and Methylocystaceae (type II). Type I methanotrophs (γ-proteobacteria) include the genera Methylobacter, Methylomicrobium, Methylomonas, Methylosphaera, Methylocaldum and Methylococcus; while type II methanotrophs (α-proteobacteria) include the genera Methylocystis, Methylosinus, Methylocella and Methylocapsa. Methanotrophs have been isolated from a wide variety of environments including soils, sediments, freshwater, and even more extreme environments such as Antarctic tundra and acid peat bogs (Hansen et al, 1996). This project focused on two main objectives: (1) to evaluate the methane oxidation capacity of biomass obtained from two different habitats: Cedar Swamp (CS) and School Street Marsh (SSM), and (2) to identify and characterize the diversity of active methanotrophs in these habitats by using a combination of cultivation-dependent and – independent techniques. MATERIALS & METHODS: Sample Collection & Analytical Measurements: Biomass samples were collected from Cedar Swamp (surface, -20cm, -40cm, -60cm, -80cm and sediments) and School Street Marsh (A- layer). Temperature and pH were recorded at each site. Dissolved oxygen and methane concentrations were measured for each sample using micro-electrodes and gas chromatography, respectively. Carbonate, sulfate, phosphate and chloride measurements were made using ion chromatography. 454 Sequencing: DNA was extracted from each biomass sample (CS: 6 and SSM: 1) using the MO-BIO PowerSoil DNA Isolation Kit. PCR was performed on the DNA extracts using universal 2 primers (515F; 907R) and sent to Penn State University for 454-sequencing. The QIIME software package was used to perform microbial community analysis (Caporaso et al, 2010). Establishment of Microcosms & Stable Isotope Probing: Twelve microcosms (CS: 6 and SSM: 6) were established in 160ml serum bottles with 25ml of biomass sample. 13-C labeled methane (0.1%, 1% and 10%) was added to the headspace of six microcosms (CS: 3 and SSM: 3) and 12-C methane to the other set. All microcosms were incubated in the dark on a shaker at 28ᴼC for two days. Headspace methane concentrations were determined using gas chromatography. DNA was extracted from each sample (CS: 10ml & SSM: 2ml) using the MO-BIO PowerSoil DNA Isolation Kit. DNA concentrations were measured using a Nanodrop and approximately 5µg DNA was added to CsCl gradient buffer. DNA fractions were resolved by centrifugation at 55,000 rpm for 60h at 20ᴼC. Density fractionation (24 fractions; 200uL each) was performed immediately after centrifugation using a syringe pump. The density of each fraction was measured and residual CsCl was removed from DNA by washing it with TE buffer. DNA was resuspended in TE buffer and quantified using Nanodrop. The 13C-DNA and 12C-DNA fractions were used as template for PCR. PCR was performed using methanotroph-specific primers targeting the pmoA gene: A189 (Homes et al, 1995) and mb661 (Costello et al, 1999). Enrichment media: Methanotrophic bacteria were enriched in nitrate mineral salts (NMS) medium with methane (Dedysh et al, 1998). For enrichment of nitrogen-fixing methanotrophs, the same medium without nitrate was used. The pH for each media was adjusted for 5.5 and 6.8. For solid media, agar was added to a concentration of 1.5 (w/v). Biomass from the microcosms was used as the inoculum for the liquid enrichments. Dilutions to extinction (up to 10-8) experiments were performed using liquid NMS media incubated with 10% methane. The enrichment cultures were grown in a shaker incubator at 28ᴼC. Methane consumption was monitored using gas chromatography. For methanotroph isolation, single colonies (obtained via spread-plating) were streaked on NMS media (with/without nitrate and pH 5.5/6.8) and maintained at 28ᴼC either in the presence of 10% methane (anaerobic jar) or air. CARD-FISH: Sample from the liquid enrichment culture (10-4 dilution: pH 5.5, +nitrate) obtained after four days of incubation and used for FISH analysis as described by Pernthaler et al (2004). The following probes were used: Eub I-III (all bacteria), Alf968 (α-proteobacteria) and Gam42a (γ-proteobacteria). RESULTS: Analytical Measurements: Dissolved oxygen concentrations dropped drastically within the first 20cm (from surface) and the water column was completely anaerobic below that depth (fig. 1). In addition, methane concentrations increased gradually with depth (fig.1). It was also observed 3 that there was a gradual increase in the concentrations of carbonates and sulfates till -60cm below surface (fig. 2) Fig. 1: Dissolved O2 and methane conc. Fig. 2: Anion concentrations 454 Sequencing: 454 sequencing of Cedar Swamp samples revealed that amongst all samples, sediments had the highest proportion of archaea (3.1%) (table 2). Proteobacteria was the most dominant group in all samples except the sediment (fig. 3). Within the sediment, the relative abundance of Bacteroidetes and Acidobacterium increased substantially, and the proportion of Proteobacteria declined fourfold (fig. 3). In the sediment sample, within the Proteobacteria, there was a substantial decline in the relative abundance of beta-proteobacteria and an increase in the relative abundance of alpha- and delta-proteobacteria (fig. 4). The relative abundance of the known aerobic methanotrophs (Methylococcaceae) declined seven-fold with increasing depth and Methylocystaceae increased two-fold (fig. 5). 454 sequencing of the soil sample obtained from School Street Marsh failed and as a result, no data could be generated. Table 1: 454 reads per sample Table 2: Relative abundance of Bacteria & Archaea 4 Fig. 3: Bacterial Community Structure at Phylum level Table 3: Phylogenetic groups (order-level) within Proteobacteria with substantial increase in representation from -80cm to sediments Fig 4: Relative abundance within Proteobacteria Fig 5: Relative abundance of key taxonomic groups (Methylococcaceae and Methylocystaceae) involved in aerobic methane oxidation 5 Methane uptake assays & Stable Isotope Probing: Based on dissolved oxygen and methane concentration profiles we selected biomass samples obtained from 40cm from surface water as inoculum for stable isotope probing. In addition, A-layer soil from School Street Marsh was selected for stable isotope analysis. Methane uptake (per day) was more than three times higher in samples obtained from School Street Marsh (~80%) when compared to Cedar Swamp (~25%). Methane concentration in headspace did not have an effect on methane consumption rates. However, biomass samples incubated at 10% methane concentration turned turbid within two days, followed by 1% (after three days). DNA extraction (after incubation with methane C-12/13) was more efficient in samples obtained from School Street Marsh when compared to Cedar Swamp. PCR analysis on C-12 and C-13 fractions (using pmoA primers) did not result in bands in the early fractions for C-13 DNA. However, the intensity of bands in the C- 13 bands were higher when compared to the corresponding C-12 fractions. Fig 6: Methane uptake in biomass samples obtained from Cedar Swamp & School Street Marsh when fed methane at different concentrations (0.1%, 1% & 10%) Liquid enrichments & Dilution to Extinction: Four different
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