Impacts of sulfonic acids on fungal manganese oxide production

Margaret A.G. Hinkle, Jeffrey E. Post, Javier Peralta, Cara M. Santelli

Research output: Contribution to journalArticlepeer-review

Abstract

Microbial manganese (Mn) oxidation plays a critical role in Mn(III/IV) oxide formation in modern day environmental systems. These oxidation processes and resulting biominerals are sensitive to aqueous conditions, such as pH and dissolved constituent concentrations. With Mn and sulfur (S) biogeochemical cycling closely linked in many environmental systems, and dissolved organic sulfur comprising a substantial pool of total sulfur in several environments, the impact of dissolved organic sulfur compounds on Mn redox processes is important to consider. Sulfonic acids, environmentally ubiquitous organosulfur compounds, play substantial roles in S cycling in many natural and contaminated systems. Research to assess the effects of these abundant sulfonic acids on Mn biogeochemical cycling, microbial Mn oxidation processes, and Mn biominerals is needed for understanding and predicting the impact of coupled S and Mn biogeochemical cycles, particularly in environments with dynamic redox gradients or in anthropogenically contaminated systems. Further, with research on microbial and abiotic Mn oxidation processes often using aminosulfonic acids to control pH, understanding the impact of such sulfonic acids on microbial Mn oxidation processes is critical. Several recent studies found that commonly used zwitterionic N substituted aminosulfonic acids, known as Good's buffers, such as HEPES and MES, can alter abiotic birnessite sheet structures. Here we investigate the impact of two sulfonic acids with broad applications to natural and contaminated sites as well as laboratory settings (HEPES and MES) on fungal Mn oxidation relative to a carbonate buffer and a buffer-free control by three Ascomycete fungi known to oxidize Mn(II): Stagonospora sp. SRC1lsM3a, Paraphaeosphaeria sporulosa AP3s5–JAC2a, and Plectosphaerella cucumerina DS2psM2a2. Structural analyses of the products show that sulfonic acids promote Mn oxidation by P. cucumerina, producing hexagonally symmetric phyllomanganates analogous to hexagonal birnessite or c–disordered H+ birnessite [(Ca,Na,K)(Mn4+,Mn3+,□)O2 nH2O], with solid–associated Mn(II) bound to vacancy sites and biomass, while in their absence almost all Mn remains as either aqueous Mn(II) or solid–associated Mn(II) bound to biomass. In contrast, sulfonic acids exert the opposite effect on Mn oxidation by P. sporulosa, with their presence suppressing Mn(II) oxidation to Mn(IV), likely leading to the formation of mycogenic bixbyite (Mn3+2O3) while the buffer–free control forms a poorly crystalline phyllo- or tectomanganate. Meanwhile, all treatments exert a minimal effect on Mn(II) uptake from solution and Mn oxidation with Stagonospora sp., with all experimental systems and controls forming poorly crystalline, hexagonally symmetric phyllomanganates. The fact that the sulfonic acids here studied exert similar effects on Mn oxidation, but substantially different effects for each fungus, suggests they affect Mn oxidation via mechanistically similar pathways that are likely dependent on interactions with fungal exudates (which vary from species to species) or specific fungal Mn oxidation processes. Interestingly, for all fungi, MES increases Mn(III) in the resulting biominerals, while the carbonate buffer consistently decreases Mn(III). These results clearly demonstrate that sulfonic acids not only alter Mn oxide structures, as has been previously noted in abiotic studies, but can interfere with Mn oxidation reactions themselves, highlighting the incredible sensitivity of both Mn oxide structures and the Mn oxidation process to the aqueous environment.

Original languageEnglish (US)
Pages (from-to)164-182
Number of pages19
JournalGeochimica et Cosmochimica Acta
Volume341
DOIs
StatePublished - Jan 15 2023

Bibliographical note

Funding Information:
M.A.G.H. was supported by the Peter Buck Postdoctoral Fellowship at the Smithsonian Institution's National Museum for Natural History (NMNH) and the Lenfest Grant Program at W&L. J.P. was supported by the Summer Research Scholars program at W&L as well as the W&L Geology Department through the Samuel J. Kozak-Odell S. McGuire-Edgar W. Spencer-Frederick L. Schwab Award. XAFS spectra collected at beamline 4–1 at SSRL (operated by SLAC National Accelerator Laboratory and supported by U.S. Department of Energy (DOE) Office of Science through Contract No. DE–AC02–76SF00515) and at beamline 12–BM–B at the APS (operated by Argonne National Laboratory and supported by the DOE Office of Science through Contract No. DE–AC02–06CH11357). XRD patterns were collected with a Rigaku D/MAX RAPID microdiffractometer supported by the Mineral Sciences Department at the NMNH and with the Bruker D8 Advanced XRD at Washington University in St. Louis (supported by the NSF through Award No. EAR–1161543). FTIR spectra were collected using the Thermo ScientificTM Nicolet 6700 Analytical FTIR spectrometer housed in the FTIR laboratory in the Mineral Sciences Department at the NMNH, which was started with support from Stephen Turner. The authors acknowledge the Geomicrobiology Laboratories at the Smithsonian Institution NMNH for the equipment, labware, and lab space to conduct this research. We especially thank Ryan Davis, Sungsik Lee, and Benjamin Reinhart for their help in XAFS spectra collection. We also thank Bri Mondesir for her help and enthusiasm in assisting Javier Peralta with his experiments and Alexandre Fowler and Dominique Chaput for helpful discussions and for advice on specific laboratory procedures in the early stages of the research. In addition, we would like to thank three anonymous reviewers for their input and guidance in improving the manuscript.

Funding Information:
M.A.G.H. was supported by the Peter Buck Postdoctoral Fellowship at the Smithsonian Institution’s National Museum for Natural History (NMNH) and the Lenfest Grant Program at W&L. J.P. was supported by the Summer Research Scholars program at W&L as well as the W&L Geology Department through the Samuel J. Kozak-Odell S. McGuire-Edgar W. Spencer-Frederick L. Schwab Award. XAFS spectra collected at beamline 4–1 at SSRL (operated by SLAC National Accelerator Laboratory and supported by U.S. Department of Energy (DOE) Office of Science through Contract No. DE–AC02–76SF00515) and at beamline 12–BM–B at the APS (operated by Argonne National Laboratory and supported by the DOE Office of Science through Contract No. DE–AC02–06CH11357). XRD patterns were collected with a Rigaku D/MAX RAPID microdiffractometer supported by the Mineral Sciences Department at the NMNH and with the Bruker D8 Advanced XRD at Washington University in St. Louis (supported by the NSF through Award No. EAR–1161543). FTIR spectra were collected using the Thermo Scientific TM Nicolet 6700 Analytical FTIR spectrometer housed in the FTIR laboratory in the Mineral Sciences Department at the NMNH, which was started with support from Stephen Turner. The authors acknowledge the Geomicrobiology Laboratories at the Smithsonian Institution NMNH for the equipment, labware, and lab space to conduct this research. We especially thank Ryan Davis, Sungsik Lee, and Benjamin Reinhart for their help in XAFS spectra collection. We also thank Bri Mondesir for her help and enthusiasm in assisting Javier Peralta with his experiments and Alexandre Fowler and Dominique Chaput for helpful discussions and for advice on specific laboratory procedures in the early stages of the research. In addition, we would like to thank three anonymous reviewers for their input and guidance in improving the manuscript.

Publisher Copyright:
© 2022

Keywords

  • Birnessite
  • Bixbyite
  • Fungi
  • Manganese biomineralization
  • Manganese oxidizing fungi
  • Mn oxidation
  • Mycogenic manganese oxides
  • Phyllomanganates
  • Redox reactions
  • Sulfonic acids

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