Characterization of Methanogenic Communities and Nickel
Total Page：16
File Type：pdf, Size：1020Kb
Microbial Diversity 2011 Characterization of methanogenic communities and nickel requirements for methane production from Wood Hole marshes and isolation of a novel methanogen of the order Methanomicrobiales from Eel Pond mud Jennifer Glass California Institute of Technology [email protected] July 28, 2011 Microbial Diversity Course Marine Biological Laboratory 1 Highlights of Mini-Project • Isolation of anaerobic Citrobacter sp. (F1_G2) from School Street Marsh that seems to be producing acetate (further testing required). Glycerol stock made. • Isolation of methanogen with 85% similarity to Methanoplanus petrolearius from Eel Pond mud (S1_G3). Glycerol stock made. • Construction of mcrA clone libraries and community analysis for Cedar Swamp (165 clones) and School Street Marsh (62 clones). • 454 pyrosequencing and community analysis of sediment samples from Cedar Swamp (JG4: 6166 total OTUs and 224 methanogen OTUs), School Street Marsh (JG2: 5279 total OTUs and1 methanogen OTU), Great Sippewissett (JG3: 4131 total OTUs and 3 methanogen OTUs) and marine mud from the south coast of Martha’s Vineyard (JG1: 4192 total OTUs and 0 methanogen OTUs). • CARD-FISH analyses of complex populations of Methanosarcinales and other archaea and bacteria. Acquisition of two new probes (MSMX860 and MS1414) for the course. • Attempt at nickel limitation experiment was not useful for testing hypothesis likely due to nickel carryover and contamination. Introduction Methane is a very potent greenhouse gas and the only organisms known to produce it in significant quantities are methanogenic archaea. The marshes and salt ponds around Woods Hole offer excellent environments for sampling and enrichment of methanogens, as exemplified by the brilliant pyrotechnic displays every summer at Cedar Swamp during the Volta experiment. Although methanogens play a vital role in carbon cycling and require high concentrations of micronutrients such as iron, molybdenum, cobalt and nickel for growth (Schönheit et al., 1979; Sowers and Ferry, 1985), little is known about the environmental availability of such trace metals in the ecosystems when methanogens live or how methanogens acquire such metals. Furthermore, little is known about how differences in the primary substrate that methanogens use for growth (H2 CO2, acetate or methyl-compounds) affects their micronutrient/trace metal requirements. Nickel is a particularly important metal for methanogens compared to other microbes because methanogenesis involves four Ni enzymes: [NiFe] hydrogenase, Ni-dependent 2 carbon monoxide dehydrogenase, acetyl SCoA synthase and most importantly methyl-SCoM (Mcr) which catalyzes the last step of methanogenesis. During this project, I developed a hypothesis that methanogens might have higher nickel requirements when grown on H2 CO2 than when grown on acetate or methyl-compounds because additional Ni-Fe hydrogenases are required for hydrogenotrophic growth via aceticlastic or methylotrophic methanogenesis (Thauer et al., 2008). The goals of this mini-project were as follows: (1) isolate and characterize a methanogen using the group enrichments started at the beginning of the course; (2) test the nickel hypothesis by measuring methane production of pure cultures of methanogens grown on H2 CO2 vs. methylated compounds; (3) use CARD-FISH to analyze the spatial structure of methanogens in mixed culture and (4) perform community analysis of methanogenic diversity in a range of natural environments around the Woods Hole/Falmouth area. Methods and Materials Sample sites Samples were collected from the following sites around Woods Hole, Massachusetts: Great and Little Sippewissett Marsh, Eel Pond, Cedar Swamp, School Street Marsh and marine sediment (kindly provided by Erik Zettler) from the south coast of Martha’s Vineyard (Fig. 1). The first round of sampling occurred on June 15, 2011 at School Street Marsh and Little Sippewissett Marsh as part of the class field trip. Each of the four student groups collected sediment from the two sampling localities. In addition, group 3 collected a sample on June 13, 2011 from a patch of black, smelly, fine mud from Eel Pond directly across from the Loeb building near the parking lot and boats. Anaerobic chemotrophic group enrichments from these sites later became classified as the “freshwater” (School Street Marsh) and “seawater” (Little Sippewissett for groups 1, 2 and 4; Eel Pond for group 3) cultures (see below). The second round of sampling occurred on July 3-4, 2011 at Cedar Swamp, School Street Marsh and Great Sippewissett Marsh. Samples of sediment collected at Cedar Swamp, School Street Marsh and Great Sippewissett were frozen at -80°C immediately upon return to the lab. Along with a sample of marine sediment (C235AA-002-SG) collected on June 20, 2011 by Erik Zettler from 41°14’94009N/70°53’93055W, these samples were used for constructing mcrA clone libraries and for 454 pyrosequencing. At Great Sippewissett, anaerobic enrichment cultures 3 were set up within several hours after sampling (see below) whereas at School Street Marsh and Cedar Swamp, enrichment samples were taken with an N2-flushed syringe in the field and immediately inoculated into sterile anaerobic freshwater media in 160-mL serum bottles. The marine sediment enrichment culture was inoculated ~2 weeks after collection. Figure 1. Map of sampling localities in the Woods Hole and Falmouth area, Cape Cod, Massachusetts Anaerobic chemotrophic media Anaerobic basal modular medium was used throughout this project. This media was made by mixing 1 mL 0.1% resazurin with 1L of 1x seawater (342.2 mM NaCl; 14.8 mM MgCl22H2O; 1.0 mM CaCl22H2O; 6.71 mM KCl) or freshwater (17.1 mM NaCl; 1.97 mM MgCl22H2O; 0.68 mM CaCl22H2O; 6.71 mM KCl) basal medium in 1-L Pyrex bottle. The following additions were made: 20 mL 1 M MOPS (pH 7.2) to maintain the pH, 1 mL 0.1M potassium phosphate (pH 7.2), 1 mL 1000x trace element solution (made without Ni for the nickel experiments) and 0.7 g of sodium acetate (for a final concentration of 5 mM acetate). This media was purged for 1 hour under a stream of anaerobic 80% N2: 20%CO2 gas and then brought 4 into the anaerobic chamber. The following anaerobic stock solutions were then added: 70 mL 1 M NaHCO3 to adjust the pH to 7.0 under 20% CO2, 10 mL of 0.5 M NH4Cl, 2.0 mL of 0.2 M cysteine-HCl and 2.0 mL of Na2S to reduce the medium and serve as a source of sulfur. Once the medium turned from pink to clear in color, indicating complete reduction, 30 mL of media was distributed into 160-mL serum bottles. Bottles intended for nickel experiments were acid-washed in 20% hydrochloric acid and then rinsed 3x in MilliQ water. An anaerobic stock solution of nickel (30 mM NiCl26H2O) was added to final concentrations of 0 (no added Ni), 10, 100, 500 and 1000 nM Ni. For experiments with trimethylamine and methanol, anaerobic stock solutions were added to serum bottles a final concentration of 10 and 62 mM, respectively. Bottles were stoppered with blue butyl stoppers (HCl-washed for nickel experiments) and crimped inside the anaerobic chamber. Bottles were then autoclaved and allowed to cool before inoculation. Plates were made using the same media recipe described above, but with Gelrite added to the seawater media and agar added to the freshwater media. Inoculation for anaerobic chemotrophic enrichments For group enrichments, bottles containing 30-mL of anaerobic basal modular media containing either no added or 62 mM methanol were decrimped, purged with N2/CO2 gas (80%:20%) for 10 minutes and inoculated with 5-mL of a prepared soil slurry (2-3 g of soil or sediment into 15 mL of media). Bottles without added methanol were then overpressured with H2/CO2 (80%:20%) gas to 7 psi (in later transfers, higher overpressures were used, up to 30 psi). The inventory of inoculated cultures included 4 freshwater (from School Street Marsh) cultures grown on H2 CO2, 4 freshwater cultures grown on methanol, 4 seawater (from Little Sippewissett) cultures grown on H2 CO2 and 4 seawater cultures grown on methanol, totaling to 16 bottles. The annotation used for labeling cultures in this mini-project was “F” or “S” for freshwater or seawater, “1” for H2 CO2 growth, “4” for methanol growth and “G1, G2, G3 or G4” for group 1, 2, 3, or 4. Enrichment and isolation of methanogens All sixteen group cultures were transferred after 6 days of growth by withdrawing 1-mL of culture with an N2-purged syringe and injecting it into 30 mL of fresh sterile anaerobic basal modular medium. Cultures were monitored for growth by visually checking for turbidity, by 5 observing F420 autofluorescence under the microscope and by measurement of methane production by gas chromatography. The culture was grown for another 2 days and plated onto agar plates for freshwater samples and Gelrite plates for seawater samples, otherwise of the same composition as the liquid media. Plates were grown inside the anaerobic chamber in H2S incubators with the same headspace previously used for incubation. Small colonies appeared after several days of growth in the H2S incubators. These colonies were checked for F420 autofluorescence and colony PCR with universal and archeal primers (see below). I restreaked single colonies from all plates onto new plates after approximately 1 week of growth. After another week, single colonies were picked and transferred into sterile anaerobic basal modular media. Figure 2. Protocol for isolation of methanogens used during the group enrichments (first three steps) and the steps conducted in this mini-project (last three steps). Numbers indicate the number of different cultures at each step. DNA extraction DNA was extracted from four sediment samples (Great Sippewissett, Cedar Swamp, School Street Marsh and marine sediment) and from pure liquid cultures using the Mo-Bio 6 PowerSoil® DNA Isolation Kit following manufacturer’s directions. DNA concentration and quality was checked with a NanoDrop.