Microbial gene expression in anoxic subseafloor sediment was recently explored in the Baltic Sea and the Peru Margin. Our analysis of these data reveals diverse transcripts encoding proteins associated with neutralization of reactive oxygen species, including catalase, which may provide an in situ source of oxygen. We also detect transcripts associated with oxidation of iron and sulfur, and with reduction of arsenate, selenate and nitrate. Given limited input of electron acceptors from outside the system, these results suggest that the microbial communities use an unexpectedly diverse variety of electron acceptors. Products of water radiolysis and their interactions with sediment continuously provide diverse electron acceptors and hydrogen. Cryptic microbial utilization of these oxidized substrates and H2 may be an important mechanism for multi-million-year survival under the extreme energy limitation in subseafloor sediment.
Chloroflexi are widespread in subsurface environments, and recent studies indicate that they represent a major fraction of the communities in subseafloor sediment. Here, we compare the abundance, diversity, metabolic potential and gene expression of Chloroflexi from three abyssal sediment cores from the western North Atlantic Gyre (water depth >5400 m) covering up to 15 million years of sediment deposition, where Chloroflexi were found to represent major components of the community at all sites. Chloroflexi communities die off in oxic red clay over 10–15 million years, and gene expression was below detection. In contrast, Chloroflexi abundance and gene expression at the anoxic abyssal clay site increase below the seafloor and peak in 2–3 million-year-old sediment, indicating a comparably higher activity. Metatranscriptomes from the anoxic site reveal increased expression of Chloroflexi genes involved in cell wall biogenesis, protein turnover, inorganic ion transport, defense mechanisms and prophages. Phylogenetic analysis shows that these Chloroflexi are closely related to homoacetogenic subseafloor clades and actively transcribe genes involved in sugar fermentations, gluconeogenesis and Wood–Ljungdahl pathway in the subseafloor. Concomitant expression of cell division genes indicates that these putative homoacetogenic Chloroflexi are actively growing in these million-year-old anoxic abyssal sediments.
How microbial metabolism is translated into cellular reproduction under energy-limited settings below the seafloor over long timescales is poorly understood. Here, we show that microbial abundance increases an order of magnitude over a 5 million-year-long sequence in anoxic subseafloor clay of the abyssal North Atlantic Ocean. This increase in biomass correlated with an increased number of transcribed protein-encoding genes that included those involved in cytokinesis, demonstrating that active microbial reproduction outpaces cell death in these ancient sediments. Metagenomes, metatranscriptomes, and 16S rRNA gene sequencing all show that the actively reproducing community was dominated by the candidate phylum “Candidatus Atribacteria,” which exhibited patterns of gene expression consistent with fermentative, and potentially acetogenic, metabolism. “Ca. Atribacteria” dominated throughout the 8 million-year-old cored sequence, despite the detection limit for gene expression being reached in 5 million-year-old sediments. The subseafloor reproducing “Ca. Atribacteria” also expressed genes encoding a bacterial microcompartment that has potential to assist in secondary fermentation by recycling aldehydes and, thereby, harness additional power to reduce ferredoxin and NAD+. Expression of genes encoding the Rnf complex for generation of chemiosmotic ATP synthesis were also detected from the subseafloor “Ca. Atribacteria,” as well as the Wood-Ljungdahl pathway that could potentially have an anabolic or catabolic function. The correlation of this metabolism with cytokinesis gene expression and a net increase in biomass over the million-year-old sampled interval indicates that the “Ca. Atribacteria” can perform the necessary catabolic and anabolic functions necessary for cellular reproduction, even under energy limitation in millions-of-years-old anoxic sediments.
Organic carbon in marine sediments is a critical component of the global carbon cycle, and its degradation influences a wide range of phenomena, including the magnitude of carbon sequestration over geologic timescales, the recycling of inorganic carbon and nutrients, the dissolution and precipitation of carbonates, the production of methane and the nature of the seafloor biosphere. Although much has been learned about the factors that promote and hinder rates of organic carbon degradation in natural systems, the controls on the distribution of organic carbon in modern and ancient sediments are still not fully understood. In this review, we summarize how recent findings are changing entrenched perspectives on organic matter degradation in marine sediments: a shift from a structurally-based chemical reactivity viewpoint towards an emerging acceptance of the role of the ecosystem in organic matter degradation rates. That is, organic carbon has a range of reactivities determined by not only the nature of the organic compounds, but by the biological, geochemical, and physical attributes of its environment. This shift in mindset has gradually come about due to a greater diversity of sample sites, the molecular revolution in biology, discoveries concerning the extent and limits of life, advances in quantitative modeling, investigations of ocean carbon cycling under a variety of extreme paleo-conditions (e.g. greenhouse environments, euxinic/anoxic oceans), the application of novel analytical techniques and interdisciplinary efforts. Adopting this view across scientific disciplines will enable additional progress in understanding how marine sediments influence the global carbon cycle.
| Project Title | Microbial activity in oxygenated subseafloor sediment |
|---|---|
| Acronym | Microbial activity subseafloor sediment |
| URL | https://www.bco-dmo.org/project/788507 |
| Created | January 27, 2020 |
| Modified | February 28, 2020 |
The subseafloor sedimentary biosphere is the largest ecosystem on Earth, where microbes subsist under energy-limited conditions over long timescales. It is poorly understood how activity is converted into microbial reproduction and survival under these conditions. Here, we examine this question in deep-sea subseafloor communities subsisting in oxic and anoxic abyssal sediments for over multimillion year timescales. Ammonia-oxidizing Archaea (AOA) dominate oxic abyssal microbial communities by up to an order of magnitude for 15 million years in the oxic subseafloor of the North Atlantic Ocean. Rates of nitrification correlated with the abundance of these dominant AOA populations, whose metabolism is characterized by ammonia oxidation, mixotrophic utilization of organic nitrogen, deamination, and the energetically efficient chemolithoautotrophic hydroxypropionate/ hydroxybutyrate carbon fixation cycle. These AOA thus have the potential to couple mixotrophic and chemolithoautotrophic metabolism via mixotrophic deamination of organic nitrogen, followed by oxidation of the regenerated ammonia for additional energy to fuel carbon fixation. This metabolic feature likely reduces energy loss and improves AOA fitness under energy starved, oxic conditions, thereby allowing them to outcompete other taxa for millions of years. In abyssal anoxic sites, a single population affiliated with the candidate Phylum “Candidatus Atribacteria” dominates the subseafloor community for up to 8 million years. Expression of genes encoding proteins for cell division were detected only in the upper 10 mbsf, indicating increased abundances of “Ca. Atribacteria” were due to actively reproducing microbes. Mean net sulfate reduction rate is relatively high over the upper 10-meter interval. At greater depths, the ecosystem is subject to net death, with mean net sulfate reduction rate below detection, microbial abundance steadily declining, and no detectable expression of cell division genes. Even in this net-death interval, “Ca. Atribacteria” dominates the subseafloor community. The transcriptomes indicate that “Ca. Atribacteria” is homoacetogenic, utilizing electron bifurcation to couple fermentative H2 production from sugars with the Wood-Ljungdahl carbon fixation pathway. Additional reducing power for ATP synthesis appears to be gained by secondary fermentations via a bacterial micro-compartment. This energy-efficient metabolism apparently improves the fitness of “Ca. Atribacteria” in this energy-limited setting, allowing this group to dominate communities over multimillion-year timescales.
Additional project information is available from C-DEBI: https://www.darkenergybiosphere.org/award/microbial-activity-in-oxygenated-subseafloor-sediment/
| Name | Affiliation | Role |
|---|---|---|
| William D. Orsi | University of Munich | Lead Principal Investigator |
| URL | https://www.bco-dmo.org/dataset/798144 |
|---|---|
| Download URL | https://www.bco-dmo.org/dataset/798144/data/download |
| Media Type | application/octet-stream |
| Created | February 28, 2020 |
| Modified | March 3, 2020 |
| State | Final no updates expected |
| Brief Description | 16S rRNA gene datasets, metagenomes, and metatranscriptomes |
Methodology: We optimized a DNA extraction protocol for ultra-organic lean sediments that provided increased yields of DNA, enabling sequencing of 16S rRNA and ammonia monooxygenase [amoA] genes, as well as metagenomes, from two deep oxic subseafloor sediment cores reaching up to ~15 million years old. In addition, we demonstrate the viability of uncultivated microbial populations via ¹⁸O-labeling in long-term (18-month) incubations in the presence of H₂¹⁸O, a method for identifying actively growing microbes.
Sampling and analytical procedures: All samples were taken by Cruise KN223 of the R/V Knorr, in the North Atlantic, from October 26th 2014 – December 2014 (Woods Hole, MA – Woods Hole, MA). At both Site 11 (22° 47.0’N, 56° 31.0’ W, water depth ~5,600 m) and Site 12 (29° 40.6’ N, 58° 19.7’ W, water depth ~5,400 m), successively deeper subseafloor cores were taken with a multicorer [to ~0.4 meters below seafloor (mbsf)], a gravity corer (to ~3 mbsf), and the 45-m WHOI “long corer” piston-coring device (https://www2.whoi.edu/site/longcore/) (to ~28 mbsf). Subsamples from the core sections for DNA extraction were sampled on board the ship immediately after retrieval with sterile 60 mL syringes with the Luer taper end cut off and frozen immediately at -80 °C prior to DNA extraction. Subsamples for the 18O-labeling experiment were sampled from the core sections in the same manner, but were stored at +4 °C prior to incubation set up.
DNA extraction: Subcores sampled aseptically with sterile syringes were sampled aseptically in a UV-sterilized DNA/RNA clean HEPA filtered laminar flow hood. To reduce contamination, the paraffin was removed and the outer 3 cm of sediment pushed out of the syringe, which was then sliced off with a red-hot, sterile spatula. A second unused, sterile, spatula was used to carefully sample the uncontaminated center of the remaining core sample inside the syringe. In brief, 10 g of sediment was transferred into 50 mL Lysing Matrix E tubes (MP Biomedicals) containing silica glass beads and homogenized for 40 sec at 6 m/s using a FastPrep 24 5-G homogenizer (MP Biomedicals) in the presence of 15 mL preheated (65 C) sterile filtered extraction buffer (76 vol% vol 1M NaPO₄ pH 8, 15 vol% 200 proof ethanol, 8 vol% MoBio’s lysis buffer solution C1, and 1 vol% SDS). The samples were incubated at 99°C for 2 minutes and frozen overnight at -20°C, thawed, and frozen again at -20°C overnight, followed by additional incubation at 99°C for 2 minutes and a second homogenization using the settings described above. The additional freeze thaw steps, particularly freezing overnight, was found to increase DNA yield 2–10 fold. After the second homogenization, the samples were centrifuged for 15 minutes, and the supernatants concentrated to a volume of 100 uL using 50 KDa Amicon centrifugal filters (Millipore). Co-extracted PCR-inhibiting humic acids and other compounds were removed from the concentrated extract using the PowerClean Pro DNA Clean-up Kit (MoBio). Extraction blanks were performed alongside the samples to assess laboratory contamination during the extraction process.
qPCR: DNA was quantified fluorometrically using a Qubit with a dsDNA high-sensititivity kit (Life Technologies). Quantitative PCR (qPCR) was performed using the custom primer dual indexed approach that targets the V4 hypervariable region of the 16S rRNA gene using updated 16S rRNA gene primers 515F/806R (515F: 5′ – GTGYCAGCMGCCGCGGTAA– 3′, 806R: GGACTACNVGGGTWTCTAAT) that increase coverage of ammonia oxidizing Thaumarchaea and other marine strains(59). To measure the abundance of amoA genes from archaea, the primers Arch amoA-1F (STAATGGTCTGGCTTAGACG) and Arch amoA-2R (GCGGCCATCCATCTGTATGT) were used. qPCR reactions were prepared using an automated liquid handler (pipetting robot), the EpMotion 5070 (Eppendorf), was used to set up all qPCR reactions and standard curves. The efficiency values of the qPCR was <90% and R² values >0.95. qPCR was performed using white 96-well plates as this was found to increase the signal to noise in the SYBR green assay 2-fold compared to clear plates. The technical variability of 16S rRNA gene qPCR measurements was determined to be consistently <5% under the the EpMotion 5070.
16S rRNA and amoA gene sequencing: Barcoded V4 hypervariable regions of amplified 16S rRNA genes were sequenced on an Illumina MiniSeq. This yielded a total of >20,000,000 raw sequencing reads that were then subjected to quality control. In order to quality control the OTU picking algorithm for the data, we also sequenced a “mock community” alongside our environmental samples. The mock communities contained a defined number of species (n=18) all containing 16S rRNA genes >3% difference. USEARCH version 10.0.240 was used for quality control and OTU picking (61), OTUs were clustered at 97% sequence identity. The taxonomic relationship of OTU representative sequences were identified by BLASTn searches against SILVA database (www.arb-silva.de) version 128. To identify contaminants, 16S rRNA genes from extraction blanks and dust samples from the lab were also sequenced. These 16S rRNA gene sequences from contaminants were used to identify any contaminating bacteria in our oxic abyssal clay samples. All OTUs containing sequences from these ‘contaminant’ samples were removed prior to downstream analysis.
qPCR of 16S rRNA genes in DNA extraction blanks were consistently <10² copies per extraction, and thus we used 10² copies to define our detection limit for the abyssal clay samples. Consistent with this, high-throughput sequencing of amplicons with qPCR values <10² copies per g sediment had up to 50% sequence representation from contaminant taxa, whereas samples with values >10² copies per g sediment had <5% representation from contaminant taxa. This further supported our definition of <10² as a realistic detection limit. Using samples that had 16S rRNA gene copies >10² copies per g sediment, we were able to analyze microbial communities down to ca. 15 mbsf at Site 11 and ca. 8 mbsf at Site 12.
Thaumarchaeal amoA genes amplified via qPCR using the method described above were cloned using the TOPO TA cloning kit, and Sanger sequenced at the LMU Munich Sequencing Service at the Faculty of Biology (http://www.gi.bio.lmu.de/sequencing). Prior to phylogenetic analysis, the reads were quality trimmed in CLC Genomics using the default settings for quality control.
Experimental setups: DNA-SIP experiments with H₂¹⁸O were set up at two North Atlantic coring sites: Site 11 and Site 12, from depths 2.8 and 3.5 mbsf, respectively. Prior to setting up the incubations, the subcores were sampled with sterile syringes using the sample aseptic technique used for the DNA extraction. For each sample depth, seven grams of abyssal clay was placed into sterile 20-mL glass flasks and incubated with 4 mL of sterile artificial seawater composed of either H₂¹⁸O (97% atomic enrichment) or unlabeled artificial seawater. Vials were crimp sealed, with an oxygenated headspace of approximately 10 mL, and incubated at 8 C. The water content of the clay was measured to be approximately 40% (+/- 5%) of the total weight. This initial water content diluted the final concentration of added H₂¹⁸O to be ca. 60% of the total water content of the sample. The artificial seawater was different from the porewater at depth because there was no added nitrate, but there was also no added ammonia which should be similar to the in situ conditions where ammonia is generally below detection. Oxygen was measured continuously throughout the incubations using non-invasive fiberoptic measurements as described previously. Small fluctuations in the oxygen measurements in the killed control, and experimental incubations (Fig S3), were likely due to temperature fluctuations of the incubator itself (1°C), since the non-invasive fiber optic oxygen sensor spots are temperature sensitive.
To assess the preservation potential of extracellular DNA (eDNA), and its ability to bias our study that is based on DNA from living organisms, we monitored microbial growth in the presence and absence of added eDNA over a 210-day incubation experiment. eDNA extracted from a culture of Rhodococcus erythropolis was added to sediment slurries from 2.8 mbsf at Site 11 at a concentration of 5 ng g⁻¹. Microbial growth was measured over time with 16S rRNA gene qPCR, and also in a control that did not receive the eDNA. As a second control, we added eDNA to autoclaved (dead) sediment. DNA was extracted from each timepoint and measure with qPCR using the methods described above.
Density gradient fractionation, qPCR: We used Tag-SIP to measure the atom % ¹⁸O-enrichment of actively growing microbial taxa. In brief, after 7 and 18 months incubations DNA was extracted and subjected to Cesium Chloride (CsCl) density gradient centrifugation as described previously. The same 16S 515F/806R primers (described above) were used in qPCR (described above) to determine density shifts in the peak DNA of buoyant density (BD) for each incubation. 16S rRNA gene amplicons from each fractions resulting from the density gradient fractionation were Illumina sequenced as described above. To identify contaminants that may have entered during the fractionation process, we also included in the sequencing run extraction blanks from the SIP fractionation. OTUs containing sequences from extraction blanks were removed.
Metagenome library preparation, sequencing and bioinformatics analysis: Prior to library preparation, whole genome amplifications were performed on DNA extracts through a multiple displacement amplification (MDA) step of 6 to 7 hours, using the REPLI-g Midi Kit (QIAGEN) and following the manufacturer’s instructions. We monitored amplification using SYBR green I (Invitrogen) on a CFX-Connect qPCR machine, stopping amplifications once the exponential phase was reached. Metagenomic libraries were prepared using the Nextera XT DNA Library Prep Kit (Illumina). Quality control and quantification of the libraries were obtained on an Agilent 2100 Bioanalyzer System, using the High Sensitivity DNA reagents and DNA chips (Agilent Genomics). was used to normalize library DNA concentrations. Metagenomic libraries were diluted to 1 nM using the Select-a-Size DNA Clean and Concentrator MagBead Kit (Zymo Research), and pooled for further sequencing on the Illumina MiniSeq platform.
Data processing: Contigs were assembled on CLC Genomics Workbench v. 9.5.4 (QIAGEN), using a word size = 20, bubble size = 50, and a minimum contig length of 300 nucleotides. Reads were then mapped to the contigs using the following parameters (mismatch penalty = 3, insertion penalty = 3, deletion penalty = 3, minimum alignment length = 50% of read length, minimum percent identity = 95%). We then performed even further stringency controls, by removing any contig that had less than 5x coverage, e.g. reads per kilobase mapped (RPKM). The final resulting dataset of contigs was then used for open reading frame (ORF) searches and BLAST analysis. Protein encoding genes and open reading frames (ORFs) were extracted using FragGeneScan v. 1.30. Cut-off values for assigning hits to specific taxa were performed at a minimum bit score of 50, minimum amino acid similarity of 60, and an alignment length of 50 residues.
For phylogenetic analyses, OTUs of AOAs were aligned with SINA online v.1.2.11 and plotted in ARB against the SILVA 16S rRNA SSU NR99 reference database release 132. Closest environmental sequences with nearly full-length sequences (>1400 bp) were selected as taxonomic references and used to calculate trees using the Maximum Likelihood algorithm RAxML implemented with the archaeal filter and advanced bootstrap refinement selecting the best tree among 100 replicates. Partial OTU sequences were added to the tree using the maximum parsimony algorithm without allowing changes of tree topology. Statistical analyses of beta-diversity were performed using R.Studio Version 3.3.0 with the vegan package.
A homogenizer is a piece of laboratory equipment used for the homogenization of various types of material, such as tissue, plant, food, soil, and many others.
An instrument for quantitative polymerase chain reaction (qPCR), also known as real-time polymerase chain reaction (Real-Time PCR).
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.
| Name | Affiliation | Contact |
|---|---|---|
| William D. Orsi | University of Munich | ✓ |
| Shannon Rauch | University of Munich | ✓ |
| Project Title | Microbial activity in oxygenated subseafloor sediment |
|---|---|
| Acronym | Microbial activity subseafloor sediment |
| URL | https://www.bco-dmo.org/project/788507 |
| Created | January 27, 2020 |
| Modified | February 28, 2020 |
The subseafloor sedimentary biosphere is the largest ecosystem on Earth, where microbes subsist under energy-limited conditions over long timescales. It is poorly understood how activity is converted into microbial reproduction and survival under these conditions. Here, we examine this question in deep-sea subseafloor communities subsisting in oxic and anoxic abyssal sediments for over multimillion year timescales. Ammonia-oxidizing Archaea (AOA) dominate oxic abyssal microbial communities by up to an order of magnitude for 15 million years in the oxic subseafloor of the North Atlantic Ocean. Rates of nitrification correlated with the abundance of these dominant AOA populations, whose metabolism is characterized by ammonia oxidation, mixotrophic utilization of organic nitrogen, deamination, and the energetically efficient chemolithoautotrophic hydroxypropionate/ hydroxybutyrate carbon fixation cycle. These AOA thus have the potential to couple mixotrophic and chemolithoautotrophic metabolism via mixotrophic deamination of organic nitrogen, followed by oxidation of the regenerated ammonia for additional energy to fuel carbon fixation. This metabolic feature likely reduces energy loss and improves AOA fitness under energy starved, oxic conditions, thereby allowing them to outcompete other taxa for millions of years. In abyssal anoxic sites, a single population affiliated with the candidate Phylum “Candidatus Atribacteria” dominates the subseafloor community for up to 8 million years. Expression of genes encoding proteins for cell division were detected only in the upper 10 mbsf, indicating increased abundances of “Ca. Atribacteria” were due to actively reproducing microbes. Mean net sulfate reduction rate is relatively high over the upper 10-meter interval. At greater depths, the ecosystem is subject to net death, with mean net sulfate reduction rate below detection, microbial abundance steadily declining, and no detectable expression of cell division genes. Even in this net-death interval, “Ca. Atribacteria” dominates the subseafloor community. The transcriptomes indicate that “Ca. Atribacteria” is homoacetogenic, utilizing electron bifurcation to couple fermentative H2 production from sugars with the Wood-Ljungdahl carbon fixation pathway. Additional reducing power for ATP synthesis appears to be gained by secondary fermentations via a bacterial micro-compartment. This energy-efficient metabolism apparently improves the fitness of “Ca. Atribacteria” in this energy-limited setting, allowing this group to dominate communities over multimillion-year timescales.
Additional project information is available from C-DEBI: https://www.darkenergybiosphere.org/award/microbial-activity-in-oxygenated-subseafloor-sediment/
| Name | Affiliation | Role |
|---|---|---|
| William D. Orsi | University of Munich | Lead Principal Investigator |
Ammonia-oxidizing archaea (AOA) dominate microbial communities throughout oxic subseafloor sediment deposited over millions of years in the North Atlantic Ocean. Rates of nitrification correlated with the abundance of these dominant AOA populations, whose metabolism is characterized by ammonia oxidation, mixotrophic utilization of organic nitrogen, deamination, and the energetically efficient chemolithoautotrophic hydroxypropionate/hydroxybutyrate carbon fixation cycle. These AOA thus have the potential to couple mixotrophic and chemolithoautotrophic metabolism via mixotrophic deamination of organic nitrogen, followed by oxidation of the regenerated ammonia for additional energy to fuel carbon fixation. This metabolic feature likely reduces energy loss and improves AOA fitness under energy-starved, oxic conditions, thereby allowing them to outcompete other taxa for millions of years.
Selection of microorganisms in marine sediment is shaped by energy-yielding electron acceptors for respiration that are depleted in vertical succession. However, some taxa have been reported to reflect past depositional conditions suggesting they have experienced weak selection after burial. In sediments underlying the Arabian Sea oxygen minimum zone (OMZ), we performed the first metagenomic profiling of sedimentary DNA at centennial-scale resolution in the context of a multi-proxy paleoclimate reconstruction. While vertical distributions of sulfate reducing bacteria and methanogens indicate energy-based selection typical of anoxic marine sediments, 5–15% of taxa per sample exhibit depth-independent stratigraphies indicative of paleoenvironmental selection over relatively short geological timescales. Despite being vertically separated, indicator taxa deposited under OMZ conditions were more similar to one another than those deposited in bioturbated intervals under intervening higher oxygen. The genomic potential for denitrification also correlated with palaeo-OMZ proxies, independent of sediment depth and available nitrate and nitrite. However, metagenomes revealed mixed acid and Entner-Dourdoroff fermentation pathways encoded by many of the same denitrifier groups. Fermentation thus may explain the subsistence of these facultatively anaerobic microbes whose stratigraphy follows changing paleoceanographic conditions. At least for certain taxa, our analysis provides evidence of their paleoenvironmental selection over the last glacial-interglacial cycle.
| URL | https://www.bco-dmo.org/dataset/626184 |
|---|---|
| Download URL | https://www.bco-dmo.org/dataset/626184/data/download |
| Media Type | text/tab-separated-values |
| Created | November 6, 2015 |
| Modified | September 30, 2016 |
| State | Final no updates expected |
| Brief Description | Sub-seafloor sediment eukaryotic rRNA collected on JOIDES Resolution Legs 201 and 204, R/V Maria Merian at North Pond, and R/V Meteor at the Benguela Upwelling System. |
Sample collection and storage: Subsurface sediment samples from Hydrate Ridge (IODP Leg 204 Site 1244a; 44° 35′ 17″ N, 125° 07′ 19″ W), Peru Margin (IODP Leg 201 Site 1227a; 79° 57′ 349″ W, 08° 59′ 463″ S), and Eastern Equatorial Pacific (IODP Leg 201 Site 1225a; 110° 43′ 289″ W, 02° 46′ 247″ N) were obtained from the Gulf Coast Core Repository (University of Texas A&M). Gravity core subsurface samples from North Pond near the Mid-Atlantic Ridge (22° 48′ 04″ E, 46° 06′ 30″ N) and Benguela Upwelling System (14° 15′ 04″ E, 27° 44′ 40″ S) were collected on March 3rd 2009 onboard the R/V Maria Merian and April 21st 2008 onboard the R/V Meteor, respectively and were provided by Andreas Teske (University of North Carolina, Chapel Hill, NC). Careful precautions were taken during sampling to avoid contamination during the sampling process. For IODP cores, contamination tests were performed using Perfluorocarbon tracers and fluorescent microspheres. Sediment samples were immediately frozen at −80 degrees C after sampling and stored at −80 degrees C until RNA was extracted. Sediment samples at a sediment depth of 0.01 and 0.08 mbsf from Little Sippewissett Salt Marsh were taken November 13th 2011 using a sterile syringe. Sulfide was detectable in both samples and thus samples were presumed anoxic. No specific permits were required for the described field studies. The locations sampled are not privately owned or protected and field studies did not involve endangered or protected species.
RNA extraction and purification: RNA was extracted from 25 grams of sediment using the FastRNA Pro Soil-Direct Kit in a laminar flow hood to reduce contamination from aerosols. Extractions were performed at Woods Hole Oceanographic Institution. Several modifications were made to the protocol provided with the kit to increase RNA yield from low biomass subseafloor samples. It was necessary to scale up the volume of sediment that is typically extracted with the kit (~0.5 grams) due to the expected low biomass of subsurface eukaryotes. Four 15 ml Lysing Matrix E tubes (MP Biomedicals, Solon, OH) were filled with 5 g sediment and 5 ml of Soil Lysis Solution. Tubes were vortexed to suspend the sediment and Soil Lysis Solution was added to the tube leaving 1 ml of headspace. Tubes were then homogenized for 60 seconds at a setting of 4.5 on the FastPrep-24 homogenizer. Contents of the 15 ml tubes were combined into two RNAse free 50 ml falcon tubes and centrifuged for 30 minutes at 4000 RPM. The supernatants were combined in a new 50 ml RNAse-free falcon tube and 1/10 volume of 2M Sodium Acetate (pH 4.0) was added. An equal volume of phenol-chloroform (pH 6.5) was added and vortexed for 30 seconds, incubated for 5 minutes at room temperature, and centrifuged at 4000 RPM for 20 minutes at 4 degrees C. The top phase was carefully transferred to a new 50 ml falcon tube and 2.5x volumes 100% ethanol and 1/10 volume 3M Sodium Acetate were added and incubated overnight at −80 degrees C. After incubation, tubes were centrifuged at 4000 RPM for 60 minutes at 4 degrees C and the supernatant removed. Pellets were washed with 70% ethanol, centrifuged for 15 minutes at 4 degrees C, and air-dried. Dried pellets were resuspended with 0.25 ml RNAse-free sterile water and combined into a new 1.5 ml RNAse-free tube. 1/10 volume of 2M Sodium Acetate (pH 4.0) and an equal volume of phenol:chloroform (pH 6.5) were added, the tube was vortexed for 1 minute, and incubated for 5 minutes at room temperature. The tube was then centrifuged for 10 minutes at 4 degrees C, the top phase removed into a new RNAse free 1.5 ml tube, and 0.7 volumes of 100% isopropanol was added and incubated for 1hour at −20 degrees C. After incubation tubes were centrifuged for 20 minutes at 14000 RPM at 4 degrees C and the supernatant was removed. Pellets were washed with 70% ethanol and centrifuged at 14000 RPM for 5 minutes at 4 degrees C. Ethanol was removed and the pellets air-dried. Pellets were resuspended with 200 ul of RNAse free sterile water and DNA was removed using the Turbo DNA-free kit (Life Technologies, Grand Island, NY). DNAse incubation times were increased to 1 hour to ensure removal of contaminating DNA. Samples were then taken through the protocol supplied with the FastRNA Pro Soil-Direct kit to the end (starting at the RNA Matrix and RNA Slurry addition step), including the optional column purification step to remove residual humic acids. To further purify the RNA, we used the MEGA-Clear RNA Purification Kit. Extraction blanks were performed (adding sterile water instead of sample) to identify aerosolized contaminants that may have entered sample and reagent tubes during the extraction process. To reduce contamination, all RNA extractions were performed in a laminar flow hood.
RT-PCR amplification of eukaryotic rRNA: To amplify the V4 hypervariable region of eukaryotic rRNA, we used PCR primers targeting this region: EukV4F (5′ – CGTATCGCCTCCCTCGCGCCATCAGxxxxxxxxxxCCAGCASCYGCGGTAATTCC – 3′) and EukV4R (5′ – CTATGCGCCTTGCCAGCCCGCTCAGACTTTCGTTCTTGATYRA – 3′), where the x region represents the unique MID barcode used for each sample, the linker primer sequence is underlined, and the 18S rRNA eukaryotic primer is bold. These primers were chosen because they target a wide range of eukaryotic taxa. RT-PCR was performed using the SuperScript One-Step RT-PCR with Platinum Taq kit. Individual reactions consisted of 2 ul RNA template, 25 ul buffer, 1 ul of forward Primer, 1 ul of reverse primer, 2 ul of the Platinum RT-Taq enzyme mix, and 18 ul RNAse free sterile water. The cDNA step was performed at 55 degrees C and cDNA was amplified in 40 cycles of PCR with an annealing temperature of 65 degrees C (55 degrees C for 30 minutes, 95 degrees C for 5 minutes, [95 degrees C for 15 seconds, 65 degrees C for 30 seconds, 68 degrees C for 1 minute]x40, 68 degrees C for 5 minutes). To check for DNA carryover during the RNA extraction protocol, a separate PCR reaction (at the same number of cycles) was included in which Taq polymerase was substituted for the reverse-transcriptase/platinum Taq enzyme mix. For each sample, 5–10 RT-PCR reactions were performed and extracted using the Zymo Research Gel Extraction Kit. A gel volume of 100% isopropanol was added to each dissolved gel slice before addition to the DNA collection column. Dissolved gel slices from each sample were pooled by centrifuging them all through the same DNA collection column. cDNA was quantified fluorometrically prior to 454 sequencing using the Qubit 2.0. To identify contaminants we performed additional RT-PCR amplifications at 55 cycles using RNAse free sterile water and RNA extraction blanks (resulting from RNA extractions in which no sample was added) as template. Contaminants were amplified with primers containing a unique MID in 55 cycles of PCR.
Quality control, clustering, and taxonomic assignment of 454 data: cDNA amplicons were sequenced on a GS-FLX Titanium 454 sequencer at EnGenCore (University of South Carolina, Columbia, SC), which resulted in ~37000 reads. To reduce homopolymer errors inherent to 454 sequencing, the dataset was put through the denoise protocol as described in the QIIME software package using the denoise_wrapper.py command. After denoising, chimeric sequences were identified and removed using ChimeraSlayer with the blast_fragments method in QIIME. The data were subjected to quality score filtering using the split_libraries.py command and clustered at various levels of sequence identity (80%, 85%, 90%, 93%, 95%, 97%) in QIIME using the uclust method of all-to-all pair-wise comparisons via the pick_otus.py command.
The QIIME taxonomy classification pipeline was not able to accurately classify the majority of eukaryotic OTUs. Thus, we used Jaguc, a program developed specifically for classification of eukaryotic rRNA sequence data, to classify our sequence reads. 90% of eukaryotic OTUs were classified to genus using this approach. OTU tables were created using the make_otu_table.py command in QIIME and the Jaguc taxonomy for each OTU was amended onto this table using a custom perl script developed by the authors for this purpose. This perl script is available from the authors upon request.
Terminal Restriction Fragment Length Polymorphism (TRFLP) analysis of fungal rRNA: To further investigate the fungal diversity in our samples, we used a TRFLP approach using PCR primers specific to fungal 18S rRNA. The fungal primers used were EF3 (5′ – TCCTCTAAATGACCAAGTTTG – 3′) and Fung5 (5′ – GTAAAAGTCCTGGTTCCCC – 3′). The forward primer, EF3, was labeled with the phosphoramidite dye 6-Carboxyfluorescein (6-FAM) at the 5′-end (Integrated DNA Technologies, Coralville, Iowa). Fungal rRNA was amplified using a cDNA incubation step at 50 degrees C followed by 40 cycles of PCR with an annealing temperature of 53 degrees C. Three RT-PCR reactions were performed for each sample, gel extracted, and pooled using the same protocol as above. Fungal rRNA amplicons were digested with three different restriction enzymes: MspI, RsaI, and HhaI (New England Biolabs, Ipswich, MA), for 1 hr at 37 degrees C. These restriction enzymes were chosen because they have been shown to provide statistically significant TRFLP data for interpreting fungal community structure across different samples. Digests were mixed with the Applied Biosystems size marker GS600LIZ and HiDi Formamide in the ratio 1:1:9 and run on an Applied Biosystems 3730 DNA analyzer. Electropherograms were analyzed using the PeakScanner software package (Applied Biosystems, Carlsbad, CA) to identify the size, height, and peak area of each T-RF. T-REX was used to filter out noise from true peaks and to align peaks.
Related references:
The manuscript is at http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0056335
Statistical Analyses: Canonical Correspondence Analysis (CCA) was used to elucidate relationships between eukaryotic community structure and concentrations of dissolved oxygen (O2), nitrate (NO3−) dissolved inorganic carbon (DIC), total organic carbon (TOC), and sulfide. Multi-response Permutation Procedure (MRPP) was used to test for a statistically significant influence of sediment depth, DIC, sulfide, TOC, and oxygen on the observed OTU distributions. All ordination and multivariate statistical analyses were performed on the TRFLP and pyrosequenced datasets as a whole, as well as the five major eukaryotic subgroups that dominated our 454 dataset: Metazoa, Viridiplantae, Diatoms, Alveolates, and Fungi. Analyses were performed on sequences affiliated with these groups clustered at 80, 85, 90, 93, and 97% sequence identity thresholds as well as the fungal TRFLP dataset. MRPP and CCA were implemented using the PC-ORD software package (MjM Software Design). Weighted UniFrac analysis was performed in QIIME. Prior to UniFrac and alpha-diversity comparisons, the number of sequences per sample were normalized to the sample with the least number of sequences by randomly selecting a subset of sequences from each sample using the multiple_rarefactions.py script in QIIME.
Quality-filtered reads and raw reads are publicly available through the NCBI SRA at http://www.ncbi.nlm.nih.gov/sra?term= SRA052670
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)
cDNA amplicons were sequenced on a GS-FLX Titanium 454 sequencer at EnGenCore (University of South Carolina, Columbia, SC), which resulted in ~37000 reads.
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.
NCBI SRA accession number.
Brief description of the sequence.
brief description, open ended, specific to the data set in which it appears
Hyperlink to NCBI SRA accession.
| Name | Affiliation | Contact |
|---|---|---|
| William D. Orsi | Woods Hole Oceanographic Institution (WHOI) | |
| Virginia P. Edgcomb | Woods Hole Oceanographic Institution (WHOI) | |
| Jennifer F. Biddle | Woods Hole Oceanographic Institution (WHOI) | ✓ |
| William D. Orsi | University of Delaware | |
| Shannon Rauch | University of Delaware | |
| William D. Orsi | ||
| William D. Orsi | ||
| Shannon Rauch | Woods Hole Oceanographic Institution (WHOI BCO-DMO) |
| Project Title | World-wide exploration of microbial eukaryote diversity and activity in the marine subsurface |
|---|---|
| Acronym | Microbial Euk Div Mar Subsurface |
| URL | https://www.bco-dmo.org/project/626119 |
| Created | November 5, 2015 |
| Modified | November 10, 2015 |
Project description obtained from C-DEBI:
Practically nothing is known about microbial eukaryotes (mEuks) in the marine subsurface. mEuks are pivotal members of microbial communities because they regenerate nutrients and modify or remineralize organic matter through grazing on prokaryotic and other eukaryotic prey. Thus, mEuks help determine metabolic potentials of microbial communities and influence elemental cycling. Only one study has addressed mEuk diversity in the marine subsurface (Edgcomb et al. 2010), which suggested Fungi dominate the eukaryotic subsurface community and are active in sediments 35 mbsf at the Peru Margin. Thus, some mEuks may be specifically adapted to the deep subsurface and may play significant roles in the utilization and regeneration of organic matter and nutrients in deep-sea sediments.
One objective of this study will be to further investigate whether Fungi are consistently the dominant group of mEuks in the marine subsurface by examining mEuk diversity in a broad range of subsurface samples from ODP expeditions spanning the world’s oceans. Deep sequencing of SSU rRNA in these samples will provide a proxy for mEuk diversity and activity in the marine subsurface. A second objective will be to ‘ground truth’ an mRNA isolation protocol for mEuks in marine subsurface sediments. Once established, this protocol will enable the third objective, which is the creation of a eukaryotic metatranscriptome from ODP site 1229. This metatranscriptome will provide insights into the functional role of mEuks in the marine subsurface and perhaps new insights into microbial evolution.
This project was funded by a C-DEBI Postdoctoral Fellowship.
| Name | Affiliation | Role |
|---|---|---|
| William D. Orsi | University of Munich | Principal Investigator |
| Glenn D. Christman | Woods Hole Oceanographic Institution (WHOI) | Co-Principal Investigator |
| Virginia P. Edgcomb | University of Delaware | Co-Principal Investigator |
| Jennifer F. Biddle | University of Delaware | Co-Principal Investigator |
| URL | https://www.bco-dmo.org/dataset/626150 |
|---|---|
| Download URL | https://www.bco-dmo.org/dataset/626150/data/download |
| Media Type | text/tab-separated-values |
| Created | November 5, 2015 |
| Modified | September 30, 2016 |
| State | Final no updates expected |
| Brief Description | Sub-seafloor metatranscriptomes from anaerobic Peru Margin sediments collected on JOIDES Resolution, Leg 201. |
Sample collection and storage: Subsurface sediment samples from the continental shelf of Peru, Ocean Drilling Program (ODP) Site 1229D (77° 57.4590′ W, 10° 58.5721′ S), were obtained during ODP Leg 201 on 6 March 2002. Careful precautions were taken to avoid contamination during the sampling process. For Integrated Ocean Drilling Program (IODP) cores, contamination tests were performed using perfluorocarbon tracers and fluorescent microspheres. Sediment samples were immediately frozen at −80 degrees C after sampling and stored at −80 degrees C until used for mRNA extractions in this study (10-year storage time at −80 degrees C).
RNA extraction and purification: Extraction of sub-seafloor RNA was performed according to the protocol described previously. In brief, RNA was extracted from 25 g of sediment using the FastRNA Pro Soil-Direct Kit (MP Biomedicals). It was necessary to scale up the volume of sediment that is typically extracted with the kit (~0.5 g) owing to the low biomass inherent to marine subsurface samples. All tubes, tips and disposables used were certified RNase free and all extraction procedures were performed in a laminar flow hood to reduce aerosol contamination by bacterial and fungal cells/spores. Five 15-ml Lysing Matrix E tubes (MP Biomedicals) were filled with 5 g sediment and 5 ml of Soil Lysis Solution (MP Biomedicals). Tubes were vortexed to suspend the sediment and Soil Lysis Solution was added to the tube leaving 1 ml of headspace. Tubes were then homogenized for 60 s on the FastPrep-24 homogenizer (MP Biomedicals) with a setting of 4.5. Contents were pooled into two 50-ml tubes and centrifuged for 30 min at 4000 r.p.m. (3220g) at room temperature (25 degrees C). Supernatants were combined in a new 50-ml tube and 1/10 volume of 2 M sodium acetate (pH 4.0) was added. An equal volume of phenol-chloroform (pH 6.5) was added and vortexed for 30 s, incubated for 5 min at room temperature, and spun at 4000 r.p.m. (3220g) for 20 min at 4 degrees C. The aqueous phase was transferred to a new 50-ml tube. Nucleic acids were precipitated by adding 2.5 and 1/10 volumes 100% ethanol and 3 M sodium acetate, respectively, and incubating overnight at −80 degrees C. The next day, tubes were spun at 4000 r.p.m. (3220g) for 60 min at 4 degrees C and the supernatant removed. Pellets were washed with 70% ethanol, spun for 15 min at 4 degrees C and air-dried. Dried pellets were resuspended with 0.25 ml RNase-free sterile water and combined into a new 1.5-ml tube. 1/10 volume of 2 M sodium acetate (pH 4.0) and an equal volume of phenol-chloroform (pH 6.5) were added, vortexed for 1 min and incubated for 5 min at room temperature. This was necessary to remove residual organic material (that is, humic acids) resulting from the rather large pellet/precipitate. After centrifuging at 14000 r.p.m. (20817g) for 10 min at 4 degrees C, the top phase was removed into a new 1.5-ml tube. 0.7 volumes of 100% isopropanol was added and incubated for 1 h at −20 degrees C (to precipitate nucleic acids). Tubes were then centrifuged for 20 min at 14000 r.p.m. (20,817g) at 4 degrees C and the supernatant removed. Pellets were washed with 70% ethanol and centrifuged at 14000 r.p.m. (20817g) for 5 min at 4 degrees C. After removing ethanol and air-drying, pellets were re-suspended in 0.2 ml of RNase free sterile water. DNA was removed using the Turbo DNA-free kit (Life Technologies), increasing the incubation time to 1 h to ensure rigorous DNA removal. After this step, samples were taken through the protocol supplied with the FastRNA Pro Soil-Direct kit to the end (starting at the RNA Matrix and RNA Slurry addition step), including the column purification step to remove residual humic acids (see FastRNA Pro Soil-Direct Kit manual). Extraction blanks were performed (adding sterile water instead of sample) to ensure that aerosolized contaminants did not enter sample and reagent tubes during the extraction process. Absence of DNA and RNA contamination was confirmed by no visible amplification of small subunit (SSU) ribosomal RNA (rRNA) and rRNA genes from extraction blanks after 35 cycles of PCR and RT–PCR.
After RNA extraction, the MEGA-Clear RNA Purification Kit (Life Technologies) was used to purify the RNA. This kit removes short RNA fragments (mostly produced during the extraction protocol) and residual inhibitors (that is, humics). We followed the protocol all the way through the optional precipitation/concentration step, re-suspending the RNA pellet in 10 ul of RNase-free sterile water. Before cDNA amplification, the removal of contaminating DNA in RNA extracts was confirmed by the absence of visible amplification of SSU rRNA genes after 35 cycles of PCR using the RNA extracts as template.
cDNA amplification and Illumina sequencing: Five microlitres of purified RNA was used as template for whole-cDNA amplification using the Ovation RNA-Seq v2 System (NuGEN technologies, http://www.nugeninc.com/nugen/index.cfm/products/cs/ngs/rna-seq-v2/). We followed the manufacturer’s instructions for cDNA amplification, and the resulting quantity of cDNA was checked on a Nanodrop (Thermo Scientific) and Fluorometer (Qubit 2.0, Life Technologies). Quality of the amplified cDNA was checked on a Bioanalyzer (Agilent Biotechnologies) before Illumina sequencing. Illumina library preparation and paired-end sequencing was performed at the University of Delaware Sequencing and Genotyping Center (Delaware Biotechnology Institute).
Related references:
For more sampling information see www-odp.tamu.edu/publications/prelim/201_prel/201toc.html
The manuscript is at http://www.nature.com/nature/journal/v499/n7457/full/nature12230.html
Quality control of the data set was performed using FastQC (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/), with a quality score cutoff of 28. Approximately 1 billion paired-end reads that passed quality control were imported into CLC Genomics Workbench 5.0 (CLC Bio) and assembled using the paired-end Illumina assembler. Contigs were assembled over a range of k-mer sizes (20, 50, 60, 64) with a minimum contig size cutoff of 300 nucleotides. The k-mer size of 50 resulted in the highest number of contigs and these contigs were chosen for use in downstream analyses. To reduce the formation of chimaeric assemblies, we used a paired-end sequencing approach and performed assemblies without scaffolding. Reads were mapped onto the contigs using the read mapping option in CLC Genomics Workbench to retain information on relative abundance of contigs. Quality-filtered reads and raw reads are publicly available through the NCBI SRA at http://www.ncbi.nlm.nih.gov/sra?term=SRA058813
Absence of DNA and RNA contamination was confirmed by no visible amplification of small subunit (SSU) ribosomal RNA (rRNA) and rRNA genes from extraction blanks after 35 cycles of PCR and RT–PCR.
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)
We followed the manufacturer’s instructions for cDNA amplification, and the resulting quantity of cDNA was checked on a Nanodrop (Thermo Scientific) and Fluorometer (Qubit 2.0, Life Technologies).
An instrument used to measure the relative absorption of electromagnetic radiation of different wavelengths in the near infra-red, visible and ultraviolet wavebands by samples.
We followed the manufacturer’s instructions for cDNA amplification, and the resulting quantity of cDNA was checked on a Nanodrop (Thermo Scientific) and Fluorometer (Qubit 2.0, Life Technologies).
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.
Quality of the amplified cDNA was checked on a Bioanalyzer (Agilent Biotechnologies) before Illumina sequencing.
A Bioanalyzer is a laboratory instrument that provides the sizing and quantification of DNA, RNA, and proteins. One example is the Agilent Bioanalyzer 2100.
NCBI accession number.
NCBI SRA accession number.
Brief description of the sequence.
brief description, open ended, specific to the data set in which it appears
Hyperlink to NCBI SRA accession.
Hyperlink to NCBI accession.
| Name | Affiliation | Contact |
|---|---|---|
| William D. Orsi | Woods Hole Oceanographic Institution (WHOI) | |
| Glenn D. Christman | Woods Hole Oceanographic Institution (WHOI) | |
| Virginia P. Edgcomb | Woods Hole Oceanographic Institution (WHOI) | ✓ |
| Jennifer F. Biddle | Woods Hole Oceanographic Institution (WHOI) | ✓ |
| William D. Orsi | University of Delaware | |
| Shannon Rauch | University of Delaware | |
| Shannon Rauch | University of Delaware | |
| William D. Orsi | ||
| William D. Orsi | ||
| Shannon Rauch | Woods Hole Oceanographic Institution (WHOI BCO-DMO) |
| Project Title | World-wide exploration of microbial eukaryote diversity and activity in the marine subsurface |
|---|---|
| Acronym | Microbial Euk Div Mar Subsurface |
| URL | https://www.bco-dmo.org/project/626119 |
| Created | November 5, 2015 |
| Modified | November 10, 2015 |
Project description obtained from C-DEBI:
Practically nothing is known about microbial eukaryotes (mEuks) in the marine subsurface. mEuks are pivotal members of microbial communities because they regenerate nutrients and modify or remineralize organic matter through grazing on prokaryotic and other eukaryotic prey. Thus, mEuks help determine metabolic potentials of microbial communities and influence elemental cycling. Only one study has addressed mEuk diversity in the marine subsurface (Edgcomb et al. 2010), which suggested Fungi dominate the eukaryotic subsurface community and are active in sediments 35 mbsf at the Peru Margin. Thus, some mEuks may be specifically adapted to the deep subsurface and may play significant roles in the utilization and regeneration of organic matter and nutrients in deep-sea sediments.
One objective of this study will be to further investigate whether Fungi are consistently the dominant group of mEuks in the marine subsurface by examining mEuk diversity in a broad range of subsurface samples from ODP expeditions spanning the world’s oceans. Deep sequencing of SSU rRNA in these samples will provide a proxy for mEuk diversity and activity in the marine subsurface. A second objective will be to ‘ground truth’ an mRNA isolation protocol for mEuks in marine subsurface sediments. Once established, this protocol will enable the third objective, which is the creation of a eukaryotic metatranscriptome from ODP site 1229. This metatranscriptome will provide insights into the functional role of mEuks in the marine subsurface and perhaps new insights into microbial evolution.
This project was funded by a C-DEBI Postdoctoral Fellowship.
| Name | Affiliation | Role |
|---|---|---|
| William D. Orsi | University of Munich | Principal Investigator |
| Glenn D. Christman | Woods Hole Oceanographic Institution (WHOI) | Co-Principal Investigator |
| Virginia P. Edgcomb | University of Delaware | Co-Principal Investigator |
| Jennifer F. Biddle | University of Delaware | Co-Principal Investigator |
| Project Title | World-wide exploration of microbial eukaryote diversity and activity in the marine subsurface |
|---|---|
| Acronym | Microbial Euk Div Mar Subsurface |
| URL | https://www.bco-dmo.org/project/626119 |
| Created | November 5, 2015 |
| Modified | November 10, 2015 |
Project description obtained from C-DEBI:
Practically nothing is known about microbial eukaryotes (mEuks) in the marine subsurface. mEuks are pivotal members of microbial communities because they regenerate nutrients and modify or remineralize organic matter through grazing on prokaryotic and other eukaryotic prey. Thus, mEuks help determine metabolic potentials of microbial communities and influence elemental cycling. Only one study has addressed mEuk diversity in the marine subsurface (Edgcomb et al. 2010), which suggested Fungi dominate the eukaryotic subsurface community and are active in sediments 35 mbsf at the Peru Margin. Thus, some mEuks may be specifically adapted to the deep subsurface and may play significant roles in the utilization and regeneration of organic matter and nutrients in deep-sea sediments.
One objective of this study will be to further investigate whether Fungi are consistently the dominant group of mEuks in the marine subsurface by examining mEuk diversity in a broad range of subsurface samples from ODP expeditions spanning the world’s oceans. Deep sequencing of SSU rRNA in these samples will provide a proxy for mEuk diversity and activity in the marine subsurface. A second objective will be to ‘ground truth’ an mRNA isolation protocol for mEuks in marine subsurface sediments. Once established, this protocol will enable the third objective, which is the creation of a eukaryotic metatranscriptome from ODP site 1229. This metatranscriptome will provide insights into the functional role of mEuks in the marine subsurface and perhaps new insights into microbial evolution.
This project was funded by a C-DEBI Postdoctoral Fellowship.
| Name | Affiliation | Role |
|---|---|---|
| William D. Orsi | University of Munich | Principal Investigator |
| Glenn D. Christman | Woods Hole Oceanographic Institution (WHOI) | Co-Principal Investigator |
| Virginia P. Edgcomb | University of Delaware | Co-Principal Investigator |
| Jennifer F. Biddle | University of Delaware | Co-Principal Investigator |
Sulfate reducing bacteria (SRB) oxidize a significant proportion of subseafloor organic carbon, but their metabolic activities and subsistence mechanisms are poorly understood. Here, we report in depth phylogenetic and metabolic analyses of SRB transcripts in the Peru Margin subseafloor and interpret these results in the context of sulfate reduction activity in the sediment. Relative abundance of overall SRB gene transcripts declines strongly whereas relative abundance of ribosomal protein transcripts from sulfate reducing δ‐Proteobacteria peak at 90 m below seafloor (mbsf) within a deep sulfate methane transition zone. This coincides with isotopically heavy δ34S values of pore water sulfate (70‰), indicating active subseafloor microbial sulfate reduction. Within the shallow sulfate reduction zone (0–5 mbsf), a transcript encoding the beta subunit of dissimilatory sulfite reductase (dsrB) was related to Desulfitobacterium dehalogenans and environmental sequences from Aarhus Bay (Denmark). At 159 mbsf we discovered a transcript encoding the reversely operating dissimilatory sulfite reductase α‐subunit (rdsrA), with basal phylogenetic relation to the chemolithoautotrophic SUP05 Group II clade. A diversity of SRB transcripts involved in cellular maintenance point toward potential subsistence mechanisms under low‐energy over long time periods, and provide a detailed new picture of SRB activities underlying sulfur cycling in the deep subseafloor.
Subseafloor mixing of reduced hydrothermal fluids with seawater is believed to provide the energy and substrates needed to support deep chemolithoautotrophic life in the hydrated oceanic mantle (i.e., serpentinite). However, geosphere-biosphere interactions in serpentinite-hosted subseafloor mixing zones remain poorly constrained. Here we examine fossil microbial communities and fluid mixing processes in the subseafloor of a Cretaceous Lost City-type hydrothermal system at the magma-poor passive Iberia Margin (Ocean Drilling Program Leg 149, Hole 897D). Brucite−calcite mineral assemblages precipitated from mixed fluids ca. 65 m below the Cretaceous paleo-seafloor at temperatures of 31.7 ± 4.3 °C within steep chemical gradients between weathered, carbonate-rich serpentinite breccia and serpentinite. Mixing of oxidized seawater and strongly reducing hydrothermal fluid at moderate temperatures created conditions capable of supporting microbial activity. Dense microbial colonies are fossilized in brucite−calcite veins that are strongly enriched in organic carbon (up to 0.5 wt.% of the total carbon) but depleted in 13C (δ13CTOC = −19.4‰). We detected a combination of bacterial diether lipid biomarkers, archaeol, and archaeal tetraethers analogous to those found in carbonate chimneys at the active Lost City hydrothermal field. The exposure of mantle rocks to seawater during the breakup of Pangaea fueled chemolithoautotrophic microbial communities at the Iberia Margin, possibly before the onset of seafloor spreading. Lost City-type serpentinization systems have been discovered at midocean ridges, in forearc settings of subduction zones, and at continental margins. It appears that, wherever they occur, they can support microbial life, even in deep subseafloor environments.
Viruses are highly abundant in marine subsurface sediments and can even exceed the number of prokaryotes. However, their activity and quantitative impact on microbial populations are still poorly understood. Here, we use gene expression data from published continental margin subseafloor metatranscriptomes to qualitatively assess viral diversity and activity in sediments up to 159 metres below seafloor (mbsf). Mining of the metatranscriptomic data revealed 4651 representative viral homologues (RVHs), representing 2.2% of all metatranscriptome sequence reads, which have close translated homology (average 77%, range 60–97% amino acid identity) to viral proteins. Archaea‐infecting RVHs are exclusively detected in the upper 30 mbsf, whereas RVHs for filamentous inoviruses predominate in the deepest sediment layers. RVHs indicative of lysogenic phage–host interactions and lytic activity, notably cell lysis, are detected at all analysed depths and suggest a dynamic virus–host association in the marine deep biosphere studied here. Ongoing lytic viral activity is further indicated by the expression of clustered, regularly interspaced, short palindromic repeat‐associated cascade genes involved in cellular defence against viral attacks. The data indicate the activity of viruses in subsurface sediment of the Peruvian margin and suggest that viruses indeed cause cell mortality and may play an important role in the turnover of subseafloor microbial biomass.
The subsurface environment of Earth contains a diverse and complex community comprised of >1028 microbial cells globally, with a significant portion of this biomass being present in continental margin sediment. The cellular functions, and the genes that encode them, enabling subsurface microbial survival over geological timescales are unknown. To better understand the cellular functions that enable microbial survival over geological timescales in subseafloor sediment, metatranscriptomes from subseafloor continental margin sediment (5–159 m below the seafloor) were compared to soil metatranscriptomes using fungal genomes from the genera Cryptococcus and Aspergillus, taxa known to occupy representative samples of each environment, as reference. Soil metatranscriptomes contain a relatively higher number of overexpressed representative fungal homologous genes involved in catabolism of labile substrates, reflecting the increased bioavailability of the soil substrate pool relative to subseafloor sediment. In contrast, many fungal homologs with overexpression in subseafloor samples encode proteasomes and autophagosomes that likely help conserve and recycle amino acids under reduced availability of labile organic matter. Genes associated with stationary phase were significantly overexpressed in the subseafloor suggesting a relatively higher investment into cellular maintenance energy. Such differences indicate that 1.) subseafloor fungal transcripts are not contaminants and 2.) subseafloor fungi are relatively dormant compared to soil fungi, and likely persist for long periods in stationary phase. Our findings provide insights into biochemical mechanisms enabling subseafloor survival for long periods, under extreme pressure, with relatively recalcitrant carbon sources.
Thawing of permafrost soils is expected to stimulate microbial decomposition and respiration of sequestered carbon. This could, in turn, increase atmospheric concentrations of greenhouse gasses, such as carbon dioxide and methane, and create a positive feedback to climate warming. Recent metagenomic studies suggest that permafrost has a large metabolic potential for carbon processing, including pathways for fermentation and methanogenesis. Here, we performed a pilot study using ultrahigh throughput Illumina HiSeq sequencing of reverse transcribed messenger RNA to obtain a detailed overview of active metabolic pathways and responsible organisms in up to 70 cm deep permafrost soils at a moist acidic tundra location in Arctic Alaska. The transcriptional response of the permafrost microbial community was compared before and after 11 days of thaw. In general, the transcriptional profile under frozen conditions suggests a dominance of stress responses, survival strategies, and maintenance processes, whereas upon thaw a rapid enzymatic response to decomposing soil organic matter (SOM) was observed. Bacteroidetes, Firmicutes, ascomycete fungi, and methanogens were responsible for largest transcriptional response upon thaw. Transcripts indicative of heterotrophic methanogenic pathways utilizing acetate, methanol, and methylamine were found predominantly in the permafrost table after thaw. Furthermore, transcripts involved in acetogenesis were expressed exclusively after thaw suggesting that acetogenic bacteria are a potential source of acetate for acetoclastic methanogenesis in freshly thawed permafrost. Metatranscriptomics is shown here to be a useful approach for inferring the activity of permafrost microbes that has potential to improve our understanding of permafrost SOM bioavailability and biogeochemical mechanisms contributing to greenhouse gas emissions as a result of permafrost thaw.
During the past decade, the IODP (International Ocean Discovery Program) has fostered a significant increase in deep biosphere investigations in the marine sedimentary and crustal environments, and scientists are well-poised to continue this momentum into the next phase of the IODP. The goals of this workshop were to evaluate recent findings in a global context, synthesize available biogeochemical data to foster thermodynamic and metabolic activity modeling and measurements, identify regional targets for future targeted sampling and dedicated expeditions, foster collaborations, and highlight the accomplishments of deep biosphere research within IODP. Twenty-four scientists from around the world participated in this one-day workshop sponsored by IODP-MI and held in Florence, Italy, immediately prior to the Goldschmidt 2013 conference. A major topic of discussion at the workshop was the continued need for standard biological sampling and measurements across IODP platforms. Workshop participants renew the call to IODP operators to implement recommended protocols.
The complex interplay of climate shifts over Eurasia and global sea level changes modulates freshwater and saltwater inputs to the Black Sea. The dynamics of the hydrologic changes from the Late Glacial into the Holocene remain a matter of debate, and information on how these changes affected the ecology of the Black Sea is sparse. Here we used Roche 454 next-generation pyrosequencing of sedimentary 18S rRNA genes to reconstruct the plankton community structure in the Black Sea over the last ca. 11,400 y. We found that 150 of 2,710 species showed a statistically significant response to four environmental stages. Freshwater chlorophytes were the best indicator species for lacustrine conditions (>9.0 ka B.P.), although the copresence of previously unidentified marine taxa indicated that the Black Sea might have been influenced to some extent by the Marmara Sea since at least 9.6 ka calendar (cal) B.P. Dinoflagellates, cercozoa, eustigmatophytes, and haptophytes responded most dramatically to the gradual increase in salinity after the latest marine reconnection and during the warm and moist mid-Holocene climatic optimum. According to paired analysis of deuterium/hydrogen (D/H) isotope ratios in fossil alkenones, salinity increased rapidly with the onset of the dry Subboreal after ∼5.2 ka B.P., leading to an increase in marine fungi and the first occurrence of marine copepods. A gradual succession of dinoflagellates, diatoms, and chrysophytes occurred during the refreshening after ∼2.5 ka cal B.P. with the onset of the cool and wet Subatlantic climate and recent anthropogenic perturbations.
Scientific ocean drilling has revealed a deep biosphere of widespread microbial life in sub-seafloor sediment. Microbial metabolism in the marine subsurface probably has an important role in global biogeochemical cycles, but deep biosphere activities are not well understood. Here we describe and analyse the first sub-seafloor metatranscriptomes from anaerobic Peru Margin sediment up to 159 metres below the sea floor, represented by over 1 billion complementary DNA (cDNA) sequence reads. Anaerobic metabolism of amino acids, carbohydrates and lipids seem to be the dominant metabolic processes, and profiles of dissimilatory sulfite reductase (dsr) transcripts are consistent with pore-water sulphate concentration profiles. Moreover, transcripts involved in cell division increase as a function of microbial cell concentration, indicating that increases in sub-seafloor microbial abundance are a function of cell division across all three domains of life. These data support calculations and models of sub-seafloor microbial metabolism and represent the first holistic picture of deep biosphere activities.
The deep marine subsurface is a vast habitat for microbial life where cells may live on geologic timescales. Because DNA in sediments may be preserved on long timescales, ribosomal RNA (rRNA) is suggested to be a proxy for the active fraction of a microbial community in the subsurface. During an investigation of eukaryotic 18S rRNA by amplicon pyrosequencing, unique profiles of Fungi were found across a range of marine subsurface provinces including ridge flanks, continental margins, and abyssal plains. Subseafloor fungal populations exhibit statistically significant correlations with total organic carbon (TOC), nitrate, sulfide, and dissolved inorganic carbon (DIC). These correlations are supported by terminal restriction length polymorphism (TRFLP) analyses of fungal rRNA. Geochemical correlations with fungal pyrosequencing and TRFLP data from this geographically broad sample set suggests environmental selection of active Fungi in the marine subsurface. Within the same dataset, ancient rRNA signatures were recovered from plants and diatoms in marine sediments ranging from 0.03 to 2.7 million years old, suggesting that rRNA from some eukaryotic taxa may be much more stable than previously considered in the marine subsurface.
My experience at the 2014 Goldschmidt Conference in Sacramento was very positive. I presented the Keynote lecture in the session 18f: “Life and Death: Molecular Biomarkers to Study Current and Past Ecosystems”, which was well received by many colleagues whom I spoke with after the talk who clearly had strong interest in the work. The presentation that I gave was also highlighted by the Goldschmidt conference in a press release, and was highlighted by several news agencies. Following the publication of the Goldschmidt press release, I was interviewed by a science reporter from Science Now at the LA Times, who published another research highlight on my work in the LA Times. Since being published in the LA Times, this story has been published on numerous other science websites including Geochemical News, ScienceDaily, and Science World Report. By taking part in the session, I was also able to network with several other presenters and organizers of the session and discuss ideas for future collaborative work and proposals. Establishing these connections will hopefully lead to successful proposals that I will write together with new collaborators. I am very grateful for the support from C-DEBI that allowed me to attend the Goldschmidt Conference, and I am excited about new products that will result from future C-DEBI supported research.
Practically nothing is known about microbial eukaryotes (mEuks) in the marine subsurface. mEuks are pivotal members of microbial communities because they regenerate nutrients and modify or remineralize organic matter through grazing on prokaryotic and other eukaryotic prey. Thus, mEuks help determine metabolic potentials of microbial communities and influence elemental cycling. Only one study has addressed mEuk diversity in the marine subsurface (Edgcomb et al. 2010), which suggested Fungi dominate the eukaryotic subsurface community and are active in sediments 35 mbsf at the Peru Margin. Thus, some mEuks may be specifically adapted to the deep subsurface and may play significant roles in the utilization and regeneration of organic matter and nutrients in deep-sea sediments. One objective of this study will be to further investigate whether Fungi are consistently the dominant group of mEuks in the marine subsurface by examining mEuk diversity in a broad range of subsurface samples from ODP expeditions spanning the world’s oceans. Deep sequencing of SSU rRNA in these samples will provide a proxy for mEuk diversity and activity in the marine subsurface. A second objective will be to ‘ground truth’ an mRNA isolation protocol for mEuks in marine subsurface sediments. Once established, this protocol will enable the third objective, which is the creation of a eukaryotic metatranscriptome from ODP site 1229. This metatranscriptome will provide insights into the functional role of mEuks in the marine subsurface and perhaps new insights into microbial evolution.
The purpose of this project was to apply metatranscriptomic methods to North Atlantic pelagic clay sediment, in order to better understand the microbial activities in this habitat. Deep-sea pelagic clays contain extremely low organic matter and oxygen penetrates deeply into the sediment, which should be the major high-energy electron acceptor supporting microbial life. However, cell counts in pelagic clays are relatively low suggesting they are limited by availability of electron donors (e.g., organic matter). In contrast, organic rich yet oxidant limited continental margin sediment has much higher cell abundances. This project has produced new metatranscriptomes in electron donor limited pelagic clays from two sites in the North Atlantic, which are being compared to existing metatranscriptomic data from oxidant limited Peru Margin sediment. Metatranscriptomes from one hole have been sequenced and preliminary analysis indicates active DNA repair, energy production, and organic matter degradation by the in situ microbial communities. These data are currently being analyzed in greater detail, and funds are being sought to sequence the metatranscriptome libraries from a second hole, which will strengthen the conclusions. The results will be compared to metatranscriptomes from oxidant limited Peru Margin, in order to better understand subseafloor microbial adaptations and activities under electron donor versus oxidant limitation.
Little is known about which microorganisms have the most impact on biogeochemical cycles in the subsurface, and practically nothing is known about microbial eukaryotes (mEuks) in these ecosystems. During the first year of the C-DEBI funded postdoctoral fellowship project “World-wide Exploration of Microbial Eukaryote Diversity and Activity in the Marine Subsurface” we produced the first survey of active subsurface mEuks across a globally distributed sample collection of sediments from up to 48 meters below seafloor (mbsf). The data reveal a dramatic increase in fungal diversity with increasing sediment depth. Unique communities of fungi inhabit different locations and are selected for as a result of geographic isolation and differential responses to in situ geochemical conditions. These findings support the hypotheses that the diversity of subsurface fungi increases substantially with sediment depth, and that fungi may play an important role in large scale elemental cycling and organic substrate turnover in the marine subsurface. We propose to test these hypotheses by 1) surveying fungal diversity across five depths from Iberian Margin sediments spanning 10-120 mbsf, and 2) sequencing metatranscriptomes for analysis of fungal derived message RNA coding for functional proteins in Peru Margin (IODP site 1229) sediments from 5 and 50-mbsf.
