Global marine sediments harbor a large and highly diverse microbial biosphere, but the mechanism by which this biosphere is established during sediment burial is largely unknown. During burial in marine sediments, concentrations of easily-metabolized organic compounds and total microbial cell abundance decrease. However, it is unknown whether some microbial clades increase with depth. We show total population increases in 38 microbial families over 3 cm of sediment depth in the upper 7.5 cm of White Oak River (WOR) estuary sediments. Clades that increased with depth were more often associated with one or more of the following: anaerobes, uncultured, or common in deep marine sediments relative to those that decreased. Maximum doubling times (in situ steady state growth rates could be faster to balance cell decay) were estimated as 2-25 years by combining sedimentation rate with either quantitative PCR (qPCR) or the product of the Fraction Read Abundance of 16S rRNA genes and total Cell counts (FRAxC). Doubling times were within an order of magnitude of each other in two adjacent cores, as well as in two laboratory enrichments of Cape Lookout Bight (CLB), NC, sediments (average difference of 28 ± 19%). qPCR and FRAxC in sediment cores and laboratory enrichments produced similar doubling times for key deep subsurface uncultured clades Bathyarchaeota (8.7 ± 1.9 years) and Thermoprofundales/MBG-D (4.1 ± 0.7 years). We conclude that common deep subsurface microbial clades experience a narrow zone of growth in shallow sediments, offering an opportunity for selection of long-term subsistence traits after resuspension events.
Anoxic subsurface sediments contain communities of heterotrophic microorganisms that metabolize organic carbon at extraordinarily slow rates. In order to assess the mechanisms by which subsurface microorganisms access detrital sedimentary organic matter, we measured kinetics of a range of extracellular peptidases in anoxic sediments of the White Oak River estuary, NC. Nine distinct peptidase substrates were enzymatically hydrolyzed at all depths. Potential peptidase activities (Vmax) decreased with increasing sediment depth, although Vmaxexpressed on a per cell basis was approximately the same at all depths. Half-saturation constants (Km) decreased with depth, indicating peptidases that functioned more efficiently at low substrate concentrations. Potential activities of extracellular peptidases acting on molecules that are enriched in degraded organic matter (D-phenylalanine and L-ornithine) increased relative to enzymes that act on L-phenylalanine, further suggesting microbial community adaptation to access degraded organic matter. Nineteen classes of predicted, exported peptidases were identified in genomic data from the same site, of which genes for class C25 (gingipain-like) peptidases represented more than 40% at each depth. Methionine aminopeptidases, zinc carboxypeptidases, and class S24-like peptidases, which are involved in single-stranded DNA repair, were also abundant. These results suggest a subsurface heterotrophic microbial community that primarily accesses low-quality detrital organic matter via a diverse suite of well-adapted extracellular enzymes.
Difficulty quantifying biogeochemically significant microbes in marine sediments limits our ability to assess interspecific interactions, population turnover times, and niches of uncultured taxa. We incubated surface sediments from Cape Lookout Bight, North Carolina USA, anoxically at 21°C for 122 days. Sulfate decreased until day 68, after which methane increased, with hydrogen concentration consistent with predicted values of an electron donor exerting thermodynamic control. We measured turnover times using two relative quantification methods, quantitative PCR (qPCR) and the product of 16S gene read abundance and total cell abundance (FRAxC, for fraction of read abundance times cells), to estimate population turnover rates of uncultured clades. Most 16S rRNA reads were from deeply-branching uncultured groups and ∼ 98% of 16S rRNA genes did not abruptly shift in relative abundance when sulfate reduction gave way to methanogenesis. Uncultured Methanomicrobiales and Methanosarcinales increased at the onset of methanogenesis with population turnover times estimated from quantitative PCR (qPCR) at 9.7 ± 3.9 and 12.6 ± 4.1 days, respectively. These were consistent with FRAxC turnover times of 9.4 ± 5.8 and 9.2 ± 3.5 days, respectively. Uncultured Syntrophaceae, which are possibly fermentative syntrophs of methanogens, and uncultured Kazan-3A-21 archaea also increased at the onset of methanogenesis with FRAxC turnover times of 14.7 ± 6.9 and 10.6 ± 3.6 days. Kazan-3A-21 may therefore either perform methanogenesis or form a fermentative syntrophy with methanogens. Three genera of sulfate reducing bacteria, Desulfovibrio sp., Desulfobacter sp., and Desulfobacterium sp. increased in the first 19 days before declining rapidly during sulfate reduction. We conclude that population turnover times on the order of days can be measured robustly in organic-rich marine sediment, and the transition from sulfate-reducing to methanogenic conditions only stimulates growth in a few clades directly involved in methanogenesis, rather than the whole microbial community.
| URL | https://www.bco-dmo.org/dataset/649807 |
|---|---|
| Download URL | https://www.bco-dmo.org/dataset/649807/data/download |
| Media Type | text/tab-separated-values |
| Created | June 20, 2016 |
| Modified | August 2, 2016 |
| State | Final no updates expected |
| Brief Description | Microbial diversity and geochemistry of marine sediment mesocosm, Cape lookout bight, North Carolina |
Samples were collected with SCUBA divers at Cape Lookout Bight, North Carolina (34.6205°N, 76.5500°W), October 2, 2013. Thirty 20 cm PVC push cores were collected, capped, refrigerated, and then returned to the lab on ice in Tennessee within 48 hrs. Bubbles of methane were released from the sediments as each core was taken, indicating the presence of methane at the surface of the core. Using a plunger inserted from the bottom the first three centimeters of sediment taken from each core was placed in a 2L Erlenmeyer flask by way of a funnel. About ten core tubes were needed to fill each of the three flasks to 1.5 liters of sediment. About 100 ml of slurried sediment were autoclaved and incubated alongside the experiments under anoxic conditions as a negative control. Cell counts, hydrogen, sulfate, and methane were measured for the negative control on day 260. All measurements described below were performed for 18 weekly timepoints for all three incubations (0-122 days). An additional timepoint at 802 days was taken for 16S rRNA gene sequence analysis only.
Geochemistry
Each of the three flasks was fitted with a custom butyl rubber stopper with a hole drilled through the center to accommodate a wide bore (6 mm) glass and Teflon stopcock for the removal of samples. Two 18-gauge needles with stainless steel stopcocks were inserted into the stopper as well. Using the luer-lock fitting on the needles, ultra high purity nitrogen gas (99.999%) that had been scrubbed of oxygen using heated copper fillings was flowed through the bottles using the second needle for the outflow to make the headspace anoxic. Incubation 3 was then flushed with ≥ 99.0% methane gas (Sigma-Aldrich, St. Louis, MO). Then all the ports were closed and the flasks inverted so that sediment covered the gas ports, stopper, and stopcock, and placed in a ring stand at constant room temperature (21.4°C) in the dark.
The incubations were turned over once every seven days just before sampling. Prior to gas sampling, 2 ml of anoxic N2 gas (99.999%) was used to blow the needle clear of sediment. Hydrogen and methane gas samples were collected in glass gastight Hamilton syringes using the steel needle ports in the custom stopper. About 32 ml of sediment was removed through the glass and Teflon stopcock using a sterile 60 ml plastic catheter tip syringe. From this, two 15ml conical centrifuge tubes were filled and capped, one used for porewater analysis and the other frozen at -80°C for later molecular analysis. One ml of sediment was placed in a 2 ml screw cap tube to be fixed and used for cell counts as described below. After sampling, 30ml of oxygen- and hydrogen-scrubbed N2 was injected into the bottle to replace the lost volume. The 15 ml tube destined for porewater analysis was centrifuged at 5000 xG for 5 minutes. A syringe was used to remove the supernatant not in contact with the air. The porewater was then filtered using a 0.2 μm syringe filter into 100 μl of 10% HCl to a final volume of 1 ml. Porewater sulfate content was determined by ion chromatography (Dionex, Sunnyvale, CA), with the remaining porewater used for determining the pH.
500µl of headspace gas was injected into a Peak Performer 1 Reducing Compound Photometer (Peak Laboratories, Mountain View, CA). Premixed hydrogen ppm lab bottles (Sigma-Aldrich, St. Louis, MO) were used as standards. Hydrogen was assumed to be equilibrated between headspace and porewater, since the equilibration time for Cape Lookout Bight sediments is < 2 days (Hoehler et al. 1998), which is less than the 7 days between timepoints in our study. Therefore, gas phase partial pressures were converted to aqueous hydrogen concentrations using the solubility coefficient of hydrogen corrected for salinity of 35 ppt and temperature of 21.4°C (Crozier & Yamamoto 1980). Methane was determined by using injected 500 µl of gas from the headspace into an evacuated glass bottle to be later analyzed on a gas chromatograph with a flame ionized detector (Agilent, Santa Clara, CA). Methane concentrations were not assumed to be equilibrated with the aqueous phase, therefore concentrations are presented as partial pressures.
Cell quantification
Total cell counts were determined by direct epifluorescence microscopy SYBRGold DNA stain (Invitrogen, Carlsbad, CA). Sediments were sonicated at 20% power for 40 seconds to disaggregates cells from sediments and diluted 40-fold into PBS prior to filtration onto a 0.2 μm polycarbonate filter (Fisher Scientific, Waltham, MA) and mounted onto a slide. An autoclaved sediment sample was used as a negative control.
Quantitative PCR
DNA was extracted from sediment samples frozen at -80°C using the Fast DNA kit for Soil (MP Bio, Santa Ana, CA). Negative controls of sterilized sediment and a blank water extract were used as well. Quantitative PCR was used to determine 16S rRNA gene copy numbers of several taxa with Quantifast SYBRGreen kit (Qiagen) on a BioRad IQ5 machine. DNA standards were prepared from either existing stocks (Lloyd et al., 2011) or from TOPO plasmids (Invitrogen, Carlsbad, CA) containing PCR amplified 16S rRNA gene products of closely related relatives in the clades Methanomicrobiales and Methanosarcinales synthesized by Invitrogen (accession # AB236118 and AB679168 respectively). Standards were quantified using Hoechst dye in a flourimeter (Hoefer, Holliston, MA). 16s rRNA specific primers Methanomicrobiales and Methanosarcinales primers were selected to have good coverage of the taxa (Narihiro and Sekiguchi, 2011).
16S Ribosomal RNA Gene Amplicons
Extracted DNA was used for 16S rRNA gene amplicon analysis. The V4 region of each DNA extraction was amplified using primers 806r and 515f (Caporaso et al., 2012), as a universal primer pair for Bacteria and Archaea. Library preparations via Nexterra kit and sequencing using an Illumina MiSeq were performed at the Center for Environmental Biotechnology at the University of Tennessee in Knoxville. The Mothur MiSeq Standard Operating Procedure was used to make contigs of bidirectional sequences, cluster operational taxonomic units (OTUs) at 97% similarity, and classify them with the Silva reference set 119 (Schloss et al., 2009, Pruesse et al., 2007). 26.4% of unique sequences were removed as chimeric and then approximately 5% of total sequences were removed for failing to classify at the domain level. Reads were normalized against the sum of reads classified as bacteria and archaea.
Analyses of time course patterns of various microbial taxa were considered only for those with more than 20 reads when summed from the 18 timepoints from each of the three incubations, leaving between 593 and 669 genus-level clades of bacteria and archaea across the three incubations. Total reads ranged from 20,922 to 329,380 for the 54 libraries. 16S rRNA sequences were normalized to total classifiable bacterial and archaeal reads. ]
Raw data has been converted to environmentally relevant units as calculated from appropriate standards. Data that is below detection limits is flagged as no data (nd).
BCO-DMO Processing:
– added conventional header with dataset name, PI name, version date
– renamed parameters to BCO-DMO and BODC standards
– reformatted date from m/d/yyyy to yyyy-mm-dd
– changed Incbuation to Incubation
– added lat and lon
Dionex, Sunnyvale, CA
Ion chromatography is a form of liquid chromatography that measures concentrations of ionic species by separating them based on their interaction with a resin. Ionic species separate differently depending on species type and size. Ion chromatographs are able to measure concentrations of major anions, such as fluoride, chloride, nitrate, nitrite, and sulfate, as well as major cations such as lithium, sodium, ammonium, potassium, calcium, and magnesium in the parts-per-billion (ppb) range. (from http://serc.carleton.edu/microbelife/research_methods/biogeochemical/ic.html)
gas chromatograph with a flame ionized detector (Agilent, Santa Clara, CA)
Instruments that generate enlarged images of samples using the phenomena of fluorescence and phosphorescence instead of, or in addition to, reflection and absorption of visible light. Includes conventional and inverted instruments.
General term for a laboratory apparatus commonly used for performing polymerase chain reaction (PCR). The device has a thermal block with holes where tubes with the PCR reaction mixtures can be inserted. The cycler then raises and lowers the temperature of the block in discrete, pre-programmed steps.
(adapted from http://serc.carleton.edu/microbelife/research_methods/genomics/pcr.html)
flourimeter Hoefer, Holliston, MA
A fluorometer or fluorimeter is a device used to measure parameters of fluorescence: its intensity and wavelength distribution of emission spectrum after excitation by a certain spectrum of light. The instrument is designed to measure the amount of stimulated electromagnetic radiation produced by pulses of electromagnetic radiation emitted into a water sample or in situ.
Illumina MiSeq at the Center for Environmental Biotechnology at the University of Tennessee in Knoxville
General term for a laboratory instrument used for deciphering the order of bases in a strand of DNA. Sanger sequencers detect fluorescence from different dyes that are used to identify the A, C, G, and T extension reactions. Contemporary or Pyrosequencer methods are based on detecting the activity of DNA polymerase (a DNA synthesizing enzyme) with another chemoluminescent enzyme. Essentially, the method allows sequencing of a single strand of DNA by synthesizing the complementary strand along it, one base pair at a time, and detecting which base was actually added at each step.
unique sample identification or number; any combination of alpha numeric characters; precise definition is file dependent
date sampled; formatted as YYYY-MM-DD
date; generally reported in GMT as YYYYMMDD (year; month; day); also as MMDD (month; day); EqPac dates are local Hawaii time. ISO_Date format is YYYY-MM-DD (http://www.iso.org/iso/home/standards/iso8601.htm)
Days as in days spent sampling or days post treatment for sampling
Total cell counts determined by direct epifluorescence microscopy SYBRGold DNA stain
Concentration of cells; often determined by spectrophotometry, flow cytometry, or using a microscope.
pH: The measure of the acidity or basicity of an aqueous solution
| Name | Affiliation | Contact |
|---|---|---|
| Karen G. Lloyd | University of Tennessee Knoxville (UTK) | |
| Richard T. Kevorkian | University of Tennessee Knoxville (UTK) | |
| Nancy Copley | University of Tennessee Knoxville (UTK) | ✓ |
| Project Title | Quantifying the contribution of the deep biosphere in the marine sediment carbon cycle using deep-sea sediment cores from the Baltic Sea |
|---|---|
| Acronym | IODP-347 Microbial Quantification |
| URL | https://www.bco-dmo.org/project/639417 |
| Created | February 29, 2016 |
| Modified | December 6, 2017 |
Marine sediments contain a microbial population large enough to rival that of Earth’s oceans, but much about this vast community is unknown. Innovations in total cell counting methods have refined estimates of cell concentrations, but tell us nothing about specific taxa. Isotopic data provides evidence that a majority of subsurface microorganisms survive by breaking down organic matter, yet measurable links between specific microbial taxa and their organic matter substrates are untested. The proposed work overcomes these limitations, with a particular focus on the degradation of proteins and carbohydrates, which comprise the bulk of classifiable sedimentary organic matter. The project will link specific taxa to potential extracellular enzyme activity in the genomes of single microbial cells, apply newly-identified, optimal methods for counting viable cells belonging to specific taxa using catalyzed reporter deposition fluorescent in situ hybridization (CARD-FISH), and measure the potential activity of their enzymes in situ. The resulting data will provide key evidence about the strategies subsurface life uses to overcome extreme energy limitation and contribute to the long-term carbon cycle.
The Principal Investigators are employing novel,improved methods to quantify cells of specific taxa in the marine subsurface and to determine the biogeochemical functions of those uncultured taxa, including:
1) Determine the pathway of organic carbon degradation in single cell genomes of uncultured, numerically dominant subsurface microorganisms.
2) Quantify viable bacteria and archaea in the deep subsurface using an improvement on the existing technology of CARD-FISH.
3 )Measure the potential activities (Vmax values) of enzymes in deep Baltic Sea sediments, and use the abundances of enzyme-producing microorganisms to calculate depth profiles of cell-specific Vmax values.
The project combines these methods in order to identify and quantify the cells capable of degrading organic matter in deep sediments of the Baltic Sea, obtained from Integrated Ocean Drilling Program (IODP) expedition 347. These results will greatly expand our knowledge of the function and activity of uncultured microorganisms in the deep subsurface.
This project is associated with C-DEBI account number 157595.
| Name | Affiliation | Role |
|---|---|---|
| Karen G. Lloyd | University of Tennessee Knoxville (UTK) | Principal Investigator |
| Andrew D. Steen | University of Tennessee Knoxville (UTK) | Co-Principal Investigator |
| Project Title | Discovering pathways for organic matter breakdown in deep subsurface sediments using detailed gene homologue analysis of metagenomes |
|---|---|
| Acronym | Subsurface Organic Breakdown |
| URL | https://www.bco-dmo.org/project/649845 |
| Created | June 21, 2016 |
| Modified | June 21, 2016 |
Heterotrophy supports much of the energy-starved life in the deep biosphere, yet we know almost nothing about the pathways by which organic matter (OM) is remineralized in the deep subseafloor. The microbes responsible for OM breakdown in the subsurface are taxonomically diverse, but their lack of resemblance to well-characterized lab cultures makes it impossible to use taxonomy alone to predict the nature of their interactions with OM. Even when metagenomes or metatranscriptomes are available to provide functional information, common analysis tools only categorize annotated genes into general classes that do not distinguish between degradative, synthetic, or housekeeping processes. We propose to develop detailed gene homologue analyses to unlock hidden details about OM degradation pathways in subsurface metagenomes from a range of OM types and amounts in the Peru Margin deep biosphere. This will allow us to describe the intricate landscape of biological remineralization of OM in the marine subsurface. Finally, we will develop an open-source software tool that replicates our analysis methods (freely available to the community via web-interface or as source code from a public repository such as GitHub) to allow other researchers to perform this analysis automatically.
Reference:
Lloyd, K.G., Schreiber, L., Petersen, D.G., Kjeldsen, K.U., Lever, M.A., Steen, A.D., Stepanauskas, R., Richter, M., Kleindienst, S., Lenk, S., Schramm, A., Jørgensen, B.B. 2013. Predominant archaea in marine sediments degrade detrital proteins. Nature 496, 215–218. C-DEBI Contribution 177. Publisher’s Link
| Name | Affiliation | Role |
|---|---|---|
| Karen G. Lloyd | University of Tennessee Knoxville (UTK) | Principal Investigator |
| Andrew D. Steen | University of Tennessee Knoxville (UTK) | Co-Principal Investigator |
There is no universally accepted method to quantify bacteria and archaea in seawater and marine sediments, and different methods have produced conflicting results with the same samples. To identify best practices, we compiled data from 65 studies, plus our own measurements, in which bacteria and archaea were quantified with fluorescent in situ hybridization (FISH), catalyzed reporter deposition FISH (CARD-FISH), polyribonucleotide FISH, or quantitative PCR (qPCR). To estimate efficiency, we defined “yield” to be the sum of bacteria and archaea counted by these techniques divided by the total number of cells. In seawater, the yield was high (median, 71%) and was similar for FISH, CARD-FISH, and polyribonucleotide FISH. In sediments, only measurements by CARD-FISH in which archaeal cells were permeabilized with proteinase K showed high yields (median, 84%). Therefore, the majority of cells in both environments appear to be alive, since they contain intact ribosomes. In sediments, the sum of bacterial and archaeal 16S rRNA gene qPCR counts was not closely related to cell counts, even after accounting for variations in copy numbers per genome. However, qPCR measurements were precise relative to other qPCR measurements made on the same samples. qPCR is therefore a reliable relative quantification method. Inconsistent results for the relative abundance of bacteria versus archaea in deep subsurface sediments were resolved by the removal of CARD-FISH measurements in which lysozyme was used to permeabilize archaeal cells and qPCR measurements which used ARCH516 as an archaeal primer or TaqMan probe. Data from best-practice methods showed that archaea and bacteria decreased as the depth in seawater and marine sediments increased, although archaea decreased more slowly.
