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Inactive hydrothermal vent microbial communities are important contributors to deep ocean primary productivity

Nature Microbiology volume 9, pages 657–668 (2024)Cite this article

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Abstract

Active hydrothermal vents are oases for productivity in the deep ocean, but the flow of dissolved substrates that fuel such abundant life ultimately ceases, leaving behind inactive mineral deposits. The rates of microbial activity on these deposits are largely unconstrained. Here we show primary production occurs on inactive hydrothermal deposits and quantify its contribution to new organic carbon production in the deep ocean. Measured incorporation of 14C-bicarbonate shows that microbial communities on inactive deposits fix inorganic carbon at rates comparable to those on actively venting deposits. Single-cell uptake experiments and nanoscale secondary ion mass spectrometry showed chemoautotrophs comprise a large fraction (>30%) of the active microbial cells. Metagenomic and lipidomic surveys of inactive deposits further revealed that the microbial communities are dominated by Alphaproteobacteria and Gammaproteobacteria using the Calvin–Benson–Bassham pathway for carbon fixation. These findings establish inactive vent deposits as important sites for microbial activity and organic carbon production on the seafloor.

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Fig. 1: Bathymetry of the East Pacific Rise 9–10° N region.
Fig. 2: Assimilation of 14C-bicarbonate by hydrothermal vent deposit hosted microbial communities.
Fig. 3: NanoSIMS analysis of 15N-ammonium and 13C-bicarbonate assimilation in individual microbial cells from active (Alvinellid Rock) and inactive (M Vent) hydrothermal vent deposits.
Fig. 4: Microbial community composition from metagenomes on inactive (Lucky’s Mound and Mosh Pit) and inactive-but-proximal (Bio9) hydrothermal vent deposits.
Fig. 5: Pathways of microbial carbon fixation on hydrothermal vent deposits.

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Data availability

Raw metagenomic data are available in the NCBI Sequence Read Archive (SRA) under BioProject PRJNA998752.

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Acknowledgements

We thank the captain and shipboard crew of the RV Atlantis during the AT42-09 and AT42-21 expeditions and of the RV Roger Revelle during the RR2102 expedition. We are also thankful for the input and hard work of the pilots and crew of the HOV Alvin, ROV Jason and AUV Sentry; the science done for this project would not be possible without the support of and collaboration with University–National Oceanographic Laboratory System operators and vessels and the National Deep Submergence Facility at Woods Hole Oceanographic Institution. This work was supported by grants from the US National Science Foundation to J.B.S. (OCE 1756339), B.M.T. (OCE 1756558), M.K.T. (OCE 1756419), D.J.F. (OCE 1949485) and A.E.D. (NSF CAREER award 2143035). J.B.S. acknowledges support from a Simons Foundation Early Career Investigator in Aquatic Microbial Ecology and Evolution Award. J.J. acknowledges financial support from the Canada Research Chairs program (CRC-2020-001165). S.M. acknowledges funding support from the Natural Sciences and Engineering Research Council of Canada Collaborative Research and Training Experience project in Marine Geodynamics and Georesources program. E.P.R. acknowledges funding support from the University of Bergen Meltzer Fund. We thank Emily Paris in the Dekas Laboratory for performing the ammonium quantification. The Stanford Nano Shared Facilities where the nanoSIMS analyses were conducted is supported in part by the National Science Foundation under award ECCS-2026822. F.S., J.B. and A.M. acknowledge funding from the German Research Foundation (DFG) under Germany’s Excellence Strategy—EXC 2077-390741603. Finally, thank you to T. Meek (Texas A&M University) for providing access to liquid scintillation counting equipment.

Author information

Authors and Affiliations

  1. Department of Oceanography, Texas A&M University, College Station, Texas, USA

    Amanda M. Achberger, Charles P. Holmes II & Jason B. Sylvan

  2. Department of Soil, Water and Climate, University of Minnesota-Twin Cities, St Paul, MN, USA

    Rose Jones & Brandy M. Toner

  3. Department of Earth Sciences, Memorial University of Newfoundland, St John’s, Newfoundland and Labrador, Canada

    John Jamieson & Sarah Moriarty

  4. MARUM Center for Marine Environmental Sciences, University of Bremen, Bremen, Germany

    Florence Schubotz, Alex Manthey & Jonas Brünjes

  5. Department of Earth System Science, Stanford University, Stanford, CA, USA

    Nicolette R. Meyer & Anne E. Dekas

  6. Department of Earth Science, Centre for Deep Sea Research, University of Bergen, Bergen, Norway

    Eoghan P. Reeves

  7. Department of Earth Sciences, University of Toronto, Toronto, Ontario, Canada

    Jonas Brünjes

  8. Department of Geology and Geophysics, Woods Hole Oceanographic Institution, Woods Hole, MA, USA

    Daniel J. Fornari

  9. Department of Marine Chemistry and Geochemistry, Woods Hole Oceanographic Institution, Woods Hole, MA, USA

    Margaret K. Tivey

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  1. Amanda M. Achberger
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Contributions

J.B.S., B.M.T., M.K.T. and A.M.A. conceived the study. A.M.A., J.B.S., J.J., A.E.D. and F.S. wrote the manuscript with input from all authors. D.J.F. contributed site survey information and provided support for field work. A.M.A., R.J., C.P.H., E.P.R. and J.B. performed sample collection. A.M.A. performed carbon fixation measurements and conducted bioinformatic analysis. R.J. contributed scanning electron microscopy images. J.J. and S.M. performed mineralogical analysis. A.E.D. and N.R.M. performed NanoSIMS analysis. F.S. and A.M. performed analysis of TOC and PLFA. C.P.H. performed cell counts. All authors reviewed and approved the final manuscript.

Corresponding authors

Correspondence to Amanda M. Achberger or Jason B. Sylvan.

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Extended data

Extended Data Table 1 Summary list of samples used to measure carbon fixation assimilation rates (14C-bicarbonate) including mineralogical and biological characteristics
Full size table
Extended Data Table 2 Key marker genes used to identify the presence of carbon fixation pathways in metagenomic datasets
Full size table
Extended Data Table 3 Stable carbon isotopic composition (δ13C) of the major polar lipid fatty acids (PLFA), analyzed as FAMEs (fatty acid methyl esters)
Full size table
Extended Data Table 4 Sample name reference table showing the original sample identifier assigned shipboard at the time of collection and the simplified sample identifier used in this manuscript
Full size table

Extended Data Fig. 1

a, Representative samples collected from inactive and active hydrothermal vent deposits including photographs of bulk samples (A1-C1) as well as thin section micrographs viewed with transmitted (A2-C2) and reflected light (A3-C3); Anh, anhydrite; Atc, atacamite; Cpy,chalcopyrite; Fe-oxy, iron oxyhydroxide; Mc, marcasite, Py, pyrite; Ps, pore space. b, Bulk minerology of samples. Black denotes iron oxyhydroxides, Yellow denotes sulfides, and Blue denotes sulfates. c, Photograph of representative subdivided inactive vent sample from Lucky’s Mound used in carbon assimilation assays showing the three regions of interest (inner conduit, middle region, and outer crust).

Extended Data Fig. 2

Assimilation of 14C−bicarbonate by microbial communities (shown in Fig. 1) from active (n = 8; biologically independent samples) and inactive (n = 8) hydrothermal vent deposits, normalized for cell abundance (Extended Data Table 1). Black line indicated mean value.

Extended Data Fig. 3

NanoSIMS comparison of uptake of 13C−bicarbonate a, and 15N−ammonium b, in individual cells (gray dots) from a single rock sample at each of AlvinellidRock (active vent deposit) and Mvent (inactive vent deposit) after 2 days of incubation. Box plots indicating the median, upper, and lower quartiles (but, appearing as lines due to mostly low activity/inactive cells) are overlaid in black. Numbers above the plots indicate the number of regions of interest (putative cells) analyzed, and asterisks indicate a statistically significant difference (p < 0.05) using a two-sided, Mann-Whitney (MW) test with a Bonferroni correction for multiple hypotheses.

Extended Data Fig. 4

Relative uptake of 13C-bicarbonate and 15N−ammonium in individual cells (black circles) measured by nanoSIMS from AlvinellidRock (active vent deposit) and Mvent (inactive vent deposit) after 2 days of incubation. Data is replotted from main text Fig. 3 with truncated axes to better display data points around the origin (lower activity cells). Based on their relative assimilation of each substrate, the regions below the 2:1 line include chemoautotrophic cells, the region above the 2:1 line includes heterotrophic cells, and the regions to the left (x-axis) or below (y-axis) the dashed lines include cells below the detection limit of uptake for that substrate. Regions are defined as in Dekas et al.41.

Extended Data Fig. 5

Abundance of key genes involved in total carbon fixation pathways and potential metabolic pathways for redox reactions annotated based on HMMs using MagicLamp72 (Supplementary Table 1).

Extended Data Fig. 6

Summary of published carbon fixation rates from active hydrothermal vent mineral deposits (black; right-axis) and hydrothermal vent fluids (grey; left-axis). The blue and orange dots denote the highest and lowest reported rates, respectively.

Supplementary information

Reporting Summary

Supplementary Tables 1–3

LithoGenie HMM list as implemented within MagicLamp (taken from https://github.com/Arkadiy-Garber/MagicLamp). List of metagenome-assembled genomes from inactive hydrothermal vent deposits showing the presence (indicated with a +) of key genes for the carbon fixation pathways examined (Extended Data Table 2). Cell-normalized carbon fixation rate mean and ranges from this work and other deep-sea environments.

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Achberger, A.M., Jones, R., Jamieson, J. et al. Inactive hydrothermal vent microbial communities are important contributors to deep ocean primary productivity. Nat Microbiol 9, 657–668 (2024). https://doi.org/10.1038/s41564-024-01599-9

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