Search company, investor...


Founded Year



Loan | Alive

Total Raised


Last Raised

$1.5M | 3 yrs ago

About Kappler

Kappler is a Consumer Products & Services/Clothing & Accessories company based in Guntersville, Alabama . Kappler's investors include Enterprise Ventures and Catapult.

Headquarters Location

55 Grimes Drive

Guntersville, Alabama, 35976,

United States


Missing: Kappler's Product Demo & Case Studies

Promote your product offering to tech buyers.

Reach 1000s of buyers who use CB Insights to identify vendors, demo products, and make purchasing decisions.

Missing: Kappler's Product & Differentiators

Don’t let your products get skipped. Buyers use our vendor rankings to shortlist companies and drive requests for proposals (RFPs).

Kappler Patents

Kappler has filed 1 patent.

The 3 most popular patent topics include:

  • Handedness
  • High schools in Saskatoon
  • Mirrors
patents chart

Application Date

Grant Date


Related Topics




Occupational safety and health, Protective gear, Safety clothing, Polymers, Mirrors


Application Date


Grant Date



Related Topics

Occupational safety and health, Protective gear, Safety clothing, Polymers, Mirrors



Latest Kappler News

An evolving view on biogeochemical cycling of iron

Feb 1, 2021

Abstract Biogeochemical cycling of iron is crucial to many environmental processes, such as ocean productivity, carbon storage, greenhouse gas emissions and the fate of nutrients, toxic metals and metalloids. Knowledge of the underlying processes involved in iron cycling has accelerated in recent years along with appreciation of the complex network of biotic and abiotic reactions dictating the speciation, mobility and reactivity of iron in the environment. Recent studies have provided insights into novel processes in the biogeochemical iron cycle such as microbial ammonium oxidation and methane oxidation coupled to Fe(iii) reduction. They have also revealed that processes in the biogeochemical iron cycle spatially overlap and may compete with each other, and that oxidation and reduction of iron occur cyclically or simultaneously in many environments. This Review discusses these advances with particular focus on their environmental consequences, including the formation of greenhouse gases and the fate of nutrients and contaminants. $259.00 VAT will be added later in the checkout. Rent or Buy article from$8.99 Additional access options: Fig. 2: Redox potentials of diverse Fe(ii)–Fe(iii) redox couples. Fig. 3: Electron transfer mechanisms from microorganisms to Fe(iii) minerals. Fig. 4: Overview of processes that can overlap and lead to cryptic iron cycling. References 1. Ehrenberg, C. Vorläufige Mitteilungen über das wirkliche Vorkommen fossiler Infusorien und ihre große Verbreitung. Poggendorff Ann. 38, 213–227 (1836). 2. Chan, C. S. et al. The architecture of iron microbial mats reflects the adaptation of chemolithotrophic iron oxidation in freshwater and marine environments. Front. Microbiol. (2016). Microscopic analysis indicates how the morphology of iron-oxidizing bacteria in microbial mats responds to environmental conditions. 3. Melton, E. D., Swanner, E. D., Behrens, S., Schmidt, C. & Kappler, A. The interplay of microbially mediated and abiotic reactions in the biogeochemical Fe cycle. Nat. Rev. Microbiol. 12, 797–808 (2014). 5. Byrne, J. M. et al. Redox cycling of Fe(II) and Fe(III) in magnetite by Fe-metabolizing bacteria. Science 347, 1473–1476 (2015). First article to demonstrate magnetite could support complete microbial iron cycling; that is, Fe(ii) in magnetite can be used as an electron source by Fe(ii) oxidizers and Fe(iii) can be used by Fe(iii) reducers as an electron acceptor in a cycling fashion. 6. Berg, J. S. et al. Intensive cryptic microbial iron cycling in the low iron water column of the meromictic Lake Cadagno. Environ. Microbiol. 18, 5288–5302 (2016). Kappler, A. & Bryce, C. Cryptic biogeochemical cycles: unravelling hidden redox reactions. Environ. Microbiol. 19, 842–846 (2017). 8. Wang, M., Hu, R., Zhao, J., Kuzyakov, Y. & Liu, S. Iron oxidation affects nitrous oxide emissions via donating electrons to denitrification in paddy soils. Geoderma 271, 173–180 (2016). 9. Beal, E. J., House, C. H. & Orphan, V. J. Manganese- and iron-dependent marine methane oxidation. Science 325, 184–187 (2009). First demonstration that methane oxidation can be coupled to reduction of iron(iii) oxides and manganese(iv) oxides. 10. Orihel, D. M. et al. The “nutrient pump:” iron-poor sediments fuel low nitrogen-to-phosphorus ratios and cyanobacterial blooms in polymictic lakes. Limnol. Oceanogr. 60, 856–871 (2015). 11. Lalonde, K., Mucci, A., Ouellet, A. & Gélinas, Y. Preservation of organic matter in sediments promoted by iron. Nature 483, 198–200 (2012). 12. Muehe, E. M. et al. Fate of Cd during microbial Fe(III) mineral reduction by a novel and Cd-tolerant Geobacter species. Environ. Sci. Technol. 47, 14099–14109 (2013). 13. Glodowska, M. et al. Role of in situ natural organic matter in mobilizing As during microbial reduction of FeIII-mineral-bearing aquifer sediments from Hanoi (Vietnam). Environ. Sci. Technol. 54, 4149–4159 (2020). 14. Cutting, R. S., Coker, V. S., Fellowes, J. W., Lloyd, J. R. & Vaughan, D. J. Mineralogical and morphological constraints on the reduction of Fe(III) minerals by Geobacter sulfurreducens. Geochim. Cosmochim. Acta 73, 4004–4022 (2009). 15. Wu, T. et al. Interactions between Fe(III)-oxides and Fe(III)-phyllosilicates during microbial reduction 2: natural subsurface sediments. Geomicrobiol. J. 34, 231–241 (2017). 16. Jaisi, D. P., Dong, H. & Liu, C. Influence of biogenic Fe(II) on the extent of microbial reduction of Fe(III) in clay minerals nontronite, illite, and chlorite. Geochim. Cosmochim. Acta 71, 1145–1158 (2007). 17. Bosch, J., Heister, K., Hofmann, T. & Meckenstock, R. U. Nanosized iron oxide colloids strongly enhance microbial iron reduction. Appl. Environ. Microbiol. 76, 184–189 (2010). 18. Aeppli, M. et al. Decreases in iron oxide reducibility during microbial reductive dissolution and transformation of ferrihydrite. Environ. Sci. Technol. 53, 8736–8746 (2019). 19. Levar, C. E., Hoffman, C. L., Dunshee, A. J., Toner, B. M. & Bond, D. R. Redox potential as a master variable controlling pathways of metal reduction by Geobacter sulfurreducens. ISME J. 11, 741–752 (2017). 20. Wang, Z. et al. Kinetics of reduction of Fe(III) complexes by outer membrane cytochromes MtrC and OmcA of Shewanella oneidensis MR-1. Appl. Environ. Microbiol. 74, 6746–6755 (2008). 21. Kügler, S. et al. Iron-organic matter complexes accelerate microbial iron cycling in an iron-rich Fen. Sci. Total. Environ. 646, 972–988 (2019). 22. Daugherty, E. E., Gilbert, B., Nico, P. S. & Borch, T. Complexation and redox buffering of iron(II) by dissolved organic matter. Environ. Sci. Technol. 51, 11096–11104 (2017). 23. von der Heyden, B., Roychoudhury, A. & Myneni, S. Iron-rich nanoparticles in natural aquatic environments. Minerals 9, 287 (2019). Thorough review of the nature and impact of iron nanoparticles in the environment. 24. Hassellöv, M. & von der Kammer, F. Iron oxides as geochemical nanovectors for metal transport in soil-river systems. Elements 4, 401–406 (2008). 25. Liu, J. et al. Particle size effect and the mechanism of hematite reduction by the outer membrane cytochrome OmcA of Shewanella oneidensis MR-1. Geochim. Cosmochim. Acta 193, 160–175 (2016). 26. Druschel, G. K., Emerson, D., Sutka, R., Suchecki, P. & Luther, G. W. Low-oxygen and chemical kinetic constraints on the geochemical niche of neutrophilic iron(II) oxidizing microorganisms. Geochim. Cosmochim. Acta 72, 3358–3370 (2008). Landmark study using voltammetric electrodes to elucidate the optimum geochemical conditions of microaerophilic Fe(ii) oxidizers. 27. Barnes, A., Sapsford, D. J., Dey, M. & Williams, K. P. Heterogeneous Fe(II) oxidation and zeta potential. J. Geochem. Explor. 100, 192–198 (2009). 28. González-Davila, M., Santana-Casiano, J. M. & Millero, F. J. Oxidation of iron (II) nanomolar with H2O2 in seawater. Geochim. Cosmochim. Acta 69, 83–93 (2005). 29. Kanzaki, Y. & Murakami, T. Rate law of Fe(II) oxidation under low O2 conditions. Geochim. Cosmochim. Acta 123, 338–350 (2013). 30. King, D. W., Lounsbury, H. A. & Millero, F. J. Rates and mechanism of Fe(II) oxidation at nanomolar total iron concentrations. Environ. Sci. Technol. 29, 818–824 (1995). 31. Emerson, D., Fleming, E. J. & McBeth, J. M. Iron-oxidizing bacteria: an environmental and genomic perspective. Annu. Rev. Microbiol. 64, 561–583 (2010). 32. Chan, C. S., Emerson, D. & Luther, G. W. III The role of microaerophilic Fe-oxidizing micro-organisms in producing banded iron formations. Geobiology 14, 509–528 (2016). 33. Mori, J. F. et al. Physiological and ecological implications of an iron- or hydrogen-oxidizing member of the Zetaproteobacteria, Ghiorsea bivora, gen. nov., sp. nov. ISME J. 11, 2624–2636 (2017). 34. Emerson, D. & De Vet, W. The role of FeOB in engineered water ecosystems: a review. J. AWWA 107, E47–E57 (2015). 35. MacDonald, D. J. et al. Using in situ voltammetry as a tool to identify and characterize habitats of iron-oxidizing bacteria: from fresh water wetlands to hydrothermal vent sites. Environ. Sci. Process. Impacts 16, 2117–2126 (2014). 36. Emerson, D., Weiss, J. V. & Megonigal, J. P. Iron-oxidizing bacteria are associated with ferric hydroxide precipitates (Fe-plaque) on the roots of wetland plants. Appl. Environ. Microbiol. 65, 2758–2761 (1999). 37. Laufer, K. et al. Microaerophilic Fe(II)-oxidizing Zetaproteobacteria isolated from low-Fe marine coastal sediments: physiology and composition of their twisted stalks. Appl. Environ. Microbiol. 83, e03118–03116 (2017). 38. Orcutt, B. N. et al. Colonization of subsurface microbial observatories deployed in young ocean crust. ISME J. 5, 692–703 (2011). 39. Field, E. K. et al. Planktonic marine iron oxidizers drive iron mineralization under low-oxygen conditions. Geobiology 14, 499–508 (2016). 40. Maisch, M. et al. Contribution of microaerophilic iron(II)-oxidizers to iron(III) mineral formation. Environ. Sci. Technol. 53, 8197–8204 (2019). 41. Chiu, B. K., Kato, S., McAllister, S. M., Field, E. K. & Chan, C. S. Novel pelagic iron-oxidizing Zetaproteobacteria from the Chesapeake Bay oxic–anoxic transition zone. Front. Microbiol. 8, 1280 (2017). 42. McAllister, S. M. et al. The Fe(II)-oxidizing Zetaproteobacteria: historical, ecological and genomic perspectives. FEMS Microbiol. Ecol. (2019). 43. Barco, R. A. et al. New insight into microbial iron oxidation as revealed by the proteomic profile of an obligate iron-oxidizing chemolithoautotroph. Appl. Environ. Microbiol. 81, 5927–5937 (2015). 44. McAllister, S. M. et al. Validating the Cyc2 neutrophilic iron oxidation pathway using meta-omics of Zetaproteobacteria iron mats at marine hydrothermal vents. mSystems 5, e00553–00519 (2020). Support for Cyc2 as the iron oxidase in microaerophilic Fe(ii) oxidizers. 45. Jeans, C. et al. Cytochrome 572 is a conspicuous membrane protein with iron oxidation activity purified directly from a natural acidophilic microbial community. ISME J. 2, 542–550 (2008). 46. Edwards, B. A. & Ferris, F. G. Influence of water flow on in situ rates of bacterial Fe(II) oxidation. Geomicrobiol. J. 37, 67–75 (2020). 47. Liu, J. et al. Identification and characterization of MtoA: a decaheme c-type cytochrome of the neutrophilic Fe(II)-oxidizing bacterium Sideroxydans lithotrophicus ES-1. Front. Microbiol. 3, 37 (2012). 48. Chan, C. S., McAllister, S. M., Garber, A., Hallahan, B. J. & Rozovsky, S. Fe oxidation by a fused cytochrome-porin common to diverse Fe-oxidizing bacteria. bioRxiv (2018). 49. Byrne, J. M., Schmidt, M., Gauger, T., Bryce, C. & Kappler, A. Imaging organic–mineral aggregates formed by Fe(II)-oxidizing bacteria using helium ion microscopy. Environ. Sci. Technol. Lett. 5, 209–213 (2018). 50. Krepski, S. T., Emerson, D., Hredzak-Showalter, P. L., Luther, G. W. III & Chan, C. S. Morphology of biogenic iron oxides records microbial physiology and environmental conditions: toward interpreting iron microfossils. Geobiology 11, 457–471 (2013). 51. Sowers, T. D., Holden, K. L., Coward, E. K. & Sparks, D. L. Dissolved organic matter sorption and molecular fractionation by naturally occurring bacteriogenic iron (oxyhydr)oxides. Environ. Sci. Technol. 53, 4295–4304 (2019). 52. Lueder, U., Druschel, G., Emerson, D., Kappler, A. & Schmidt, C. Quantitative analysis of O2 and Fe2+ profiles in gradient tubes for cultivation of microaerophilic iron(II)-oxidizing bacteria. FEMS Microbiol. Ecol. (2017). 53. van der Grift, B., Rozemeijer, J. C., Griffioen, J. & van der Velde, Y. Iron oxidation kinetics and phosphate immobilization along the flow-path from groundwater into surface water. Hydrol. Earth Syst. Sci. 18, 4687–4702 (2014). 54. Enright, A. M. L. & Ferris, F. G. Bacterial Fe(II) oxidation distinguished by long-range correlation in redox potential. J. Geophys. Res. Biogeosci. 121, 1249–1257 (2016). 55. Lueder, U., Jørgensen, B. B., Kappler, A. & Schmidt, C. Photochemistry of iron in aquatic environments. Environ. Sci. Process. Impacts 22, 12–24 (2020). 57. Hartman, H. The Evolution of Photosynthesis and Microbial Mats: A Speculation on the Banded Iron Formations. (Alan R. Liss, Inc., 1984). 58. Ozaki, K., Tajika, E., Hong, P. K., Nakagawa, Y. & Reinhard, C. T. Effects of primitive photosynthesis on Earth’s early climate system. Nat. Geosci. 11, 55–59 (2018). 59. Croal, L. R., Jiao, Y. & Newman, D. K. The fox operon from Rhodobacter strain SW2 promotes phototrophic Fe(II) oxidation in Rhodobacter capsulatus SB1003. J. Bacteriol. 189, 1774–1782 (2007). 60. Ehrenreich, A. & Widdel, F. Anaerobic oxidation of ferrous iron by purple bacteria, a new type of phototrophic metabolism. Appl. Environ. Microbiol. 60, 4517–4526 (1994). 61. Jiao, Y., Kappler, A., Croal, L. R. & Newman, D. K. Isolation and characterization of a genetically tractable photoautotrophic Fe(II)-oxidizing bacterium, Rhodopseudomonas palustris strain TIE-1. Appl. Environ. Microbiol. 71, 4487–4496 (2005). 62. Straub, K. L., Rainey, F. A. & Widdel, F. Rhodovulum iodosum sp. nov. and Rhodovulum robiginosum sp. nov., two new marine phototrophic ferrous-iron-oxidizing purple bacteria. Int. J. Syst. Evol. Microbiol. 49, 729–735 (1999). 63. Heising, S., Richter, L., Ludwig, W. & Schink, B. Chlorobium ferrooxidans sp. nov., a phototrophic green sulfur bacterium that oxidizes ferrous iron in coculture with a Geospirillum sp. strain. Arch. Microbiol. 172, 116–124 (1999). 64. Llirós, M. et al. Pelagic photoferrotrophy and iron cycling in a modern ferruginous basin. Sci. Rep. 5, 13803 (2015). 65. Laufer, K. et al. Physiological characterization of a halotolerant anoxygenic phototrophic Fe(II)-oxidizing green-sulfur bacterium isolated from a marine sediment. FEMS Microbiol. Ecol. (2017). 66. Jiao, Y. & Newman, D. K. The pio operon is essential for phototrophic Fe(II) oxidation in Rhodopseudomonas palustris TIE-1. J. Bacteriol. 189, 1765–1773 (2007). 67. Gupta, D. et al. Photoferrotrophs produce a PioAB electron conduit for extracellular electron uptake. mBio 10, e02668–02619 (2019). 68. Gledhill, M. & Buck, K. The organic complexation of iron in the marine environment: A review. Front. Microbiol. 3, 69 (2012). 69. Saraiva, I. H., Newman, D. K. & Louro, R. O. Functional characterization of the FoxE iron oxidoreductase from the photoferrotroph Rhodobacter ferrooxidans SW2. J. Biol. Chem. 287, 25541–25548 (2012). 70. Crowe, S. A. et al. Draft genome sequence of the pelagic photoferrotroph Chlorobium phaeoferrooxidans. Genome Announc. 5, e01584–01516 (2017). 71. Bryce, C., Blackwell, N., Straub, D., Kleindienst, S. & Kappler, A. Draft genome sequence of Chlorobium sp. strain N1, a marine Fe(II)-oxidizing green sulfur bacterium. Microbiol. Resour. Announc. 8, e00080–00019 (2019). 72. Miot, J. et al. Iron biomineralization by anaerobic neutrophilic iron-oxidizing bacteria. Geochim. Cosmochim. Acta 73, 696–711 (2009). 73. Schaedler, S. et al. Formation of cell-iron-mineral aggregates by phototrophic and nitrate-reducing anaerobic Fe(II)-oxidizing bacteria. Geomicrobiol. J. 26, 93–103 (2009). 74. Hegler, F., Schmidt, C., Schwarz, H. & Kappler, A. Does a low-pH microenvironment around phototrophic FeII-oxidizing bacteria prevent cell encrustation by FeIII minerals? FEMS Microbiol. Ecol. 74, 592–600 (2010). 75. Swanner, E. D. et al. Fractionation of Fe isotopes during Fe(II) oxidation by a marine photoferrotroph is controlled by the formation of organic Fe-complexes and colloidal Fe fractions. Geochim. Cosmochim. Acta 165, 44–61 (2015). 76. Boyd, P. W. & Ellwood, M. J. The biogeochemical cycle of iron in the ocean. Nat. Geosci. 3, 675–682 (2010). A comprehensive review of the many dynamic processes which influence iron cycling in the oceans. 77. Faust, B. C. & Zepp, R. G. Photochemistry of aqueous iron(III)-polycarboxylate complexes: roles in the chemistry of atmospheric and surface waters. Environ. Sci. Technol. 27, 2517–2522 (1993). 78. Rose, A. L. & Waite, T. D. Reduction of organically complexed ferric iron by superoxide in a simulated natural water. Environ. Sci. Technol. 39, 2645–2650 (2005). 79. Voelker, B. M., Morel, F. M. M. & Sulzberger, B. Iron redox cycling in surface waters: Effects of humic substances and light. Environ. Sci. Technol. 31, 1004–1011 (1997). 80. Barbeau, K., Zhang, G., Live, D. H. & Butler, A. Petrobactin, a photoreactive siderophore produced by the oil-degrading marine bacterium Marinobacter hydrocarbonoclasticus. J. Am. Chem. Soc. 124, 378–379 (2002). 81. Waite, T. D. & Morel, F. M. M. Photoreductive dissolution of colloidal iron oxides in natural waters. Environ. Sci. Technol. 18, 860–868 (1984). 82. Sulzberger, B. Light-induced redox cycling of iron: roles for CO2 uptake and release by aquatic ecosystems. Aquat. Geochem. 21, 65–80 (2015). 83. Garg, S., Rose, A. L. & Waite, T. D. Photochemical production of superoxide and hydrogen peroxide from natural organic matter. Geochim. Cosmochim. Acta 75, 4310–4320 (2011). 84. Xing, G., Garg, S. & Waite, T. D. Is superoxide-mediated Fe(III) reduction important in sunlit surface waters? Environ. Sci. Technol. 53, 13179–13190 (2019). 85. Sutherland, K. M., Wankel, S. D. & Hansel, C. M. Dark biological superoxide production as a significant flux and sink of marine dissolved oxygen. Proc. Natl. Acad. Sci. USA 117, 3433–3439 (2020). 86. Diaz, J. M. et al. Widespread production of extracellular superoxide by heterotrophic bacteria. Science 340, 1223–1226 (2013). 87. Lis, H., Kranzler, C., Keren, N. & Shaked, Y. A comparative study of iron uptake rates and mechanisms amongst marine and fresh water cyanobacteria: prevalence of reductive iron uptake. Life 5, 841–860 (2015). 88. Swanner, E. D., Maisch, M., Wu, W. & Kappler, A. Oxic Fe(III) reduction could have generated Fe(II) in the photic zone of Precambrian seawater. Sci. Rep. 8, 4238 (2018). 89. Emmenegger, L., Schönenberger, R., Sigg, L. & Sulzberger, B. Light-induced redox cycling of iron in circumneutral lakes. Limnol. Oceanogr. 46, 49–61 (2001). 90. Lueder, U., Jørgensen, B. B., Kappler, A. & Schmidt, C. Fe(III) photoreduction producing Feaq2+ in oxic freshwater sediment. Environ. Sci. Technol. 54, 862–869 (2020). 91. Lueder, U. et al. Influence of physical perturbation on Fe(II) supply in coastal marine sediments. Environ. Sci. Technol. 54, 3209–3218 (2020). 92. Peng, C., Bryce, C., Sundman, A. & Kappler, A. Cryptic cycling of complexes containing Fe(III) and organic matter by phototrophic Fe(II)-oxidizing bacteria. Appl. Environ. Microbiol. 85, e02826–02818 (2019). 93. Schmidt, C., Behrens, S. & Kappler, A. Ecosystem functioning from a geomicrobiological perspective a conceptual framework for biogeochemical iron cycling. Environ. Chem. 7, 399–405 (2010). 94. Raven, J. A., Kübler, J. E. & Beardall, J. Put out the light, and then put out the light. J. Mar. Biol. Assoc. U.K. 80, 1–25 (2000). 95. Camacho, A., Walter, X. A., Picazo, A. & Zopfi, J. Photoferrotrophy: Remains of an ancient photosynthesis in modern environments. Front. Microbiol. 8 (2017). A review on the physiology of anoxygenic phototrophic Fe(ii) oxidizers and their role in modern and ancient redox-stratified systems. 96. Crowe, S. A. et al. Deep-water anoxygenic photosythesis in a ferruginous chemocline. Geobiology 12, 322–339 (2014). 97. Straub, K. L., Benz, M., Schink, B. & Widdel, F. Anaerobic, nitrate-dependent microbial oxidation of ferrous iron. Appl. Environ. Microbiol. 62, 1458–1460 (1996). 98. Bryce, C. et al. Microbial anaerobic Fe(II) oxidation – Ecology, mechanisms and environmental implications. Environ. Microbiol. 20, 3462–3483 (2018). 99. Blöthe, M. & Roden, E. E. Composition and activity of an autotrophic Fe(II)-oxidizing, nitrate-reducing enrichment culture. Appl. Environ. Microbiol. 75, 6937–6940 (2009). Article describing the composition of the only confirmed autotrophic nitrate-dependent, Fe(ii)-oxidizing enrichment culture. 100. Laufer, K., Røy, H., Jørgensen, B. B. & Kappler, A. Evidence for the existence of autotrophic nitrate-reducing Fe(II)-oxidizing bacteria in marine coastal sediment. Appl. Environ. Microbiol. 82, 6120–6131 (2016). 101. Liu, T., Chen, D., Luo, X., Li, X. & Li, F. Microbially mediated nitrate-reducing Fe(II) oxidation: quantification of chemodenitrification and biological reactions. Geochim. Cosmochim. Acta 256, 97–115 (2019). 102. Otte, J. M. et al. N2O formation by nitrite-induced (chemo)denitrification in coastal marine sediment. Sci. Rep. 9, 10691 (2019). 103. Wang, M., Hu, R., Ruser, R., Schmidt, C. & Kappler, A. Role of chemodenitrification for N2O emissions from nitrate reduction in rice paddy soils. ACS Earth Space Chem. 4, 122–132 (2020). 104. He, S., Tominski, C., Kappler, A., Behrens, S. & Roden, E. E. Metagenomic analyses of the autotrophic Fe(II)-oxidizing, nitrate-reducing enrichment culture KS. Appl. Environ. Microbiol. 82, 2656–2668 (2016). 105. Buchwald, C., Grabb, K., Hansel, C. M. & Wankel, S. D. Constraining the role of iron in environmental nitrogen transformations: Dual stable isotope systematics of abiotic NO2− reduction by Fe(II) and its production of N2O. Geochim. Cosmochim. Acta 186, 1–12 (2016). 106. Haaijer, S. C. M., Lamers, L. P. M., Smolders, A. J. P., Jetten, M. S. M. & Op den Camp, H. J. M. Iron sulfide and pyrite as potential electron donors for microbial nitrate reduction in freshwater wetlands. Geomicrobiol. J. 24, 391–401 (2007). 107. Edwards, K. J., Rogers, D. R., Wirsen, C. O. & McCollom, T. M. Isolation and characterization of novel psychrophilic, neutrophilic, Fe-oxidizing, chemolithoautotrophic α- and γ-Proteobacteria from the deep sea. Appl. Environ. Microbiol. 69, 2906–2913 (2003). 108. Yan, R. et al. Effect of reduced sulfur species on chemolithoautotrophic pyrite oxidation with nitrate. Geomicrobiol. J. 36, 19–29 (2019). 109. Holmes, P. R. & Crundwell, F. K. The kinetics of the oxidation of pyrite by ferric ions and dissolved oxygen: an electrochemical study. Geochim. Cosmochim. Acta 64, 263–274 (2000). 110. Zhao, L., Dong, H., Edelmann, R. E., Zeng, Q. & Agrawal, A. Coupling of Fe(II) oxidation in illite with nitrate reduction and its role in clay mineral transformation. Geochim. Cosmochim. Acta 200, 353–366 (2017). 111. Zhang, L., Dong, H., Kukkadapu, R. K., Jin, Q. & Kovarik, L. Electron transfer between sorbed Fe(II) and structural Fe(III) in smectites and its effect on nitrate-dependent iron oxidation by Pseudogulbenkiania sp. strain 2002. Geochim. Cosmochim. Acta 265, 132–147 (2019). 112. Shelobolina, E. S., VanPraagh, C. G. & Lovley, D. R. Use of ferric and ferrous iron containing minerals for respiration by Desulfitobacterium frappieri. Geomicrobiol. J. 20, 143–156 (2003). 113. Larese-Casanova, P., Haderlein, S. B. & Kappler, A. Biomineralization of lepidocrocite and goethite by nitrate-reducing Fe(II)-oxidizing bacteria: effect of pH, bicarbonate, phosphate, and humic acids. Geochim. Cosmochim. Acta 74, 3721–3734 (2010). 114. Pantke, C. et al. Green rust formation during Fe(II) oxidation by the nitrate-reducing Acidovorax sp. strain BoFeN1. Environ. Sci. Technol. 46, 1439–1446 (2012). 115. Nordhoff, M. et al. Insights into nitrate-reducing Fe(II) oxidation mechanisms through analysis of cell-mineral associations, cell encrustation, and mineralogy in the chemolithoautotrophic enrichment culture KS. Appl. Environ. Microbiol. 83, e00752–00717 (2017). 116. Smith, R. L., Kent, D. B., Repert, D. A. & Böhlke, J. K. Anoxic nitrate reduction coupled with iron oxidation and attenuation of dissolved arsenic and phosphate in a sand and gravel aquifer. Geochim. Cosmochim. Acta 196, 102–120 (2017). 117. Madison, A. S., Tebo, B. M., Mucci, A., Sundby, B. & Luther, G. W. Abundant porewater Mn(III) is a major component of the sedimentary redox system. Science 341, 875–878 (2013). 118. Gillispie, E. C., Taylor, S. E., Qafoku, N. P. & Hochella, M. F. Jr. Impact of iron and manganese nano-metal-oxides on contaminant interaction and fortification potential in agricultural systems – a review. Environ. Chem. 16, 377–390 (2019). 119. Siebecker, M., Madison, A. S. & Luther, G. W. Reduction kinetics of polymeric (soluble) manganese (IV) oxide (MnO2) by ferrous iron (Fe2+). Aquat. Geochem. 21, 143–158 (2015). 120. Herndon, E. M., Havig, J. R., Singer, D. M., McCormick, M. L. & Kump, L. R. Manganese and iron geochemistry in sediments underlying the redox-stratified Fayetteville Green Lake. Geochim. Cosmochim. Acta 231, 50–63 (2018). 121. Maguffin, S. C. et al. Influence of manganese abundances on iron and arsenic solubility in rice paddy soils. Geochim. Cosmochim. Acta 276, 50–69 (2020). 122. Lovley, D. R. & Phillips, E. J. P. Novel mode of microbial energy metabolism: Organic carbon oxidation coupled to dissimilatory reduction of iron or manganese. Appl. Environ. Microbiol. 54, 1472–1480 (1988). 123. Myers, C. R. & Nealson, K. H. Respiration-linked proton translocation coupled to anaerobic reduction of manganese(IV) and iron(III) in Shewanella putrefaciens MR-1. J. Bacteriol. 172, 6232–6238 (1990). 124. Lovley, D. R., Coates, J. D., Blunt-Harris, E. L., Phillips, E. J. P. & Woodward, J. C. Humic substances as electron acceptors for microbial respiration. Nature 382, 445–448 (1996). 125. Coates, J. D., Ellis, D. J., Gaw, C. V. & Lovley, D. R. Geothrix fermentans gen. nov., sp. nov., a novel Fe(III)-reducing bacterium from a hydrocarbon-contaminated aquifer. Int. J. Syst. Evol. Microbiol. 49, 1615–1622 (1999). 126. Tor, J. M. & Lovley, D. R. Anaerobic degradation of aromatic compounds coupled to Fe(III) reduction by Ferroglobus placidus. Environ. Microbiol. 3, 281–287 (2001). 127. Hansel, C. M., Benner, S. G. & Fendorf, S. Competing Fe(II)-induced mineralization pathways of ferrihydrite. Environ. Sci. Technol. 39, 7147–7153 (2005). 128. Shi, L. et al. The roles of outer membrane cytochromes of Shewanella and Geobacter in extracellular electron transfer. Environ. Microbiol. Rep. 1, 220–227 (2009). 129. Shi, L., Squier, T. C., Zachara, J. M. & Fredrickson, J. K. Respiration of metal (hydr)oxides by Shewanella and Geobacter: a key role for multihaem c-type cytochromes. Mol. Microbiol. 65, 12–20 (2007). 130. Butler, J. E., Young, N. D. & Lovley, D. R. Evolution of electron transfer out of the cell: comparative genomics of six Geobacter genomes. BMC Genomics 11, 40 (2010). 131. Reguera, G. et al. Biofilm and nanowire production leads to increased current in Geobacter sulfurreducens fuel cells. Appl. Environ. Microbiol. 72, 7345–7348 (2006). 132. Lovley, D. R. & Holmes, D. E. Protein nanowires: the electrification of the microbial world and maybe our own. J. Bacteriol. 202, e00331–00320 (2020). A comprehensive and recent review on extracellular electron transfer by bacteria. 134. Cologgi, D. L., Lampa-Pastirk, S., Speers, A. M., Kelly, S. D. & Reguera, G. Extracellular reduction of uranium via Geobacter conductive pili as a protective cellular mechanism. Proc. Natl. Acad. Sci. USA 108, 15248–15252 (2011). 135. Ueki, T. et al. Decorating the outer surface of microbially produced protein nanowires with peptides. ACS Synth. Biol. 8, 1809–1817 (2019). 136. Smith, J. A., Lovley, D. R. & Tremblay, P.-L. Outer cell surface components essential for Fe(III) oxide reduction by Geobacter metallireducens. Appl. Environ. Microbiol. 79, 901–907 (2013). 137. Pirbadian, S. et al. Shewanella oneidensis MR-1 nanowires are outer membrane and periplasmic extensions of the extracellular electron transport components. Proc. Natl. Acad. Sci. USA 111, 12883–12888 (2014). 138. El-Naggar, M. Y. et al. Electrical transport along bacterial nanowires from Shewanella oneidensis MR-1. Proc. Natl. Acad. Sci. USA 107, 18127–18131 (2010). 139. Roden, E. E. et al. Extracellular electron transfer through microbial reduction of solid-phase humic substances. Nat. Geosci. 3, 417–421 (2010). 140. Lohmayer, R., Kappler, A., Lösekann-Behrens, T. & Planer-Friedrich, B. Sulfur species as redox partners and electron shuttles for ferrihydrite reduction by Sulfurospirillum deleyianum. Appl. Environ. Microbiol. 80, 3141–3149 (2014). 141. Kappler, A., Benz, M., Schink, B. & Brune, A. Electron shuttling via humic acids in microbial iron(III) reduction in a freshwater sediment. FEMS Microbiol. Ecol. 47, 85–92 (2004). 142. Cervantes, F. J. et al. Reduction of humic substances by halorespiring, sulphate-reducing and methanogenic microorganisms. Environ. Microbiol. 4, 51–57 (2002). 143. Coates, J. D. et al. Recovery of humic-reducing bacteria from a diversity of environments. Appl. Environ. Microbiol. 64, 1504–1509 (1998). 144. Piepenbrock, A., Behrens, S. & Kappler, A. Comparison of humic substance- and Fe(III)-reducing microbial communities in anoxic aquifers. Geomicrobiol. J. 31, 917–928 (2014). 146. Marsili, E. et al. Shewanella secretes flavins that mediate extracellular electron transfer. Proc. Natl. Acad. Sci. USA 105, 3968–3973 (2008). 147. von Canstein, H., Ogawa, J., Shimizu, S. & Lloyd, J. R. Secretion of flavins by Shewanella species and their role in extracellular electron transfer. Appl. Environ. Microbiol. 74, 615–623 (2008). 148. Nevin, K. P. & Lovley, D. R. Mechanisms for Fe(III) oxide reduction in sedimentary environments. Geomicrobiol. J. 19, 141–159 (2002). 149. Markelova, E. et al. Deconstructing the redox cascade: what role do microbial exudates (flavins) play? Environ. Chem. 14, 515–524 (2017). 150. Bai, Y. et al. AQDS and redox-active NOM enables microbial Fe(III)-mineral reduction at cm-scales. Environ. Sci. Technol. 54, 4131–4139 (2020). The first article to demonstrate that microorganisms can transfer electrons to Fe(iii) over centimetre distances by electron shuttling. 151. Bai, Y., Sun, T., Angenent, L. T., Haderlein, S. B. & Kappler, A. Electron hopping enables rapid electron transfer between quinone-/hydroquinone-containing organic molecules in microbial iron(III) mineral reduction. Environ. Sci. Technol. 54, 10646–10653 (2020). 152. Liu, F. et al. Magnetite compensates for the lack of a pilin-associated c-type cytochrome in extracellular electron exchange. Environ. Microbiol. 17, 648–655 (2015). 153. Taillefert, M. et al. Shewanella putrefaciens produces an Fe(III)-solubilizing organic ligand during anaerobic respiration on insoluble Fe(III) oxides. J. Inorg. Biochem. 101, 1760–1767 (2007). 154. in ‘t Zandt, M. H., de Jong, A. E., Slomp, C. P. & Jetten, M. S. The hunt for the most-wanted chemolithoautotrophic spookmicrobes. FEMS Microbiol. Ecol. (2018). 155. Sivan, O. et al. Geochemical evidence for iron-mediated anaerobic oxidation of methane. Limnol. Oceanogr. 56, 1536–1544 (2011). 156. Miura, Y., Watanabe, A., Murase, J. & Kimura, M. Methane production and its fate in paddy fields. Soil Sci. Plant Nutr. 38, 673–679 (1992). 158. Amos, R. T. et al. Evidence for iron-mediated anaerobic methane oxidation in a crude oil-contaminated aquifer. Geobiology 10, 506–517 (2012). 159. Glodowska, M. et al. Arsenic mobilization by anaerobic iron-dependent methane oxidation. Commun. Earth Environ. 1, 42 (2020). First study providing evidence that anaerobic oxidation of methane coupled to reduction of arsenic-bearing Fe(iii) minerals can lead to arsenic mobilization in groundwater. 160. Scheller, S., Yu, H., Chadwick, G. L., McGlynn, S. E. & Orphan, V. J. Artificial electron acceptors decouple archaeal methane oxidation from sulfate reduction. Science 351, 703–707 (2016). 161. Wegener, G., Krukenberg, V., Riedel, D., Tegetmeyer, H. E. & Boetius, A. Intercellular wiring enables electron transfer between methanotrophic archaea and bacteria. Nature 526, 587–590 (2015). 162. Ettwig, K. F. et al. Archaea catalyze iron-dependent anaerobic oxidation of methane. Proc. Natl. Acad. Sci. USA 113, 12792–12796 (2016). 163. Cai, C. et al. A methanotrophic archaeon couples anaerobic oxidation of methane to Fe(III) reduction. ISME J. 12, 1929–1939 (2018). 164. Clément, J.-C., Shrestha, J., Ehrenfeld, J. G. & Jaffé, P. R. Ammonium oxidation coupled to dissimilatory reduction of iron under anaerobic conditions in wetland soils. Soil. Biol. Biochem. 37, 2323–2328 (2005). 165. Huang, S. & Jaffé, P. R. Characterization of incubation experiments and development of an enrichment culture capable of ammonium oxidation under iron-reducing conditions. Biogeosciences 12, 769–779 (2015). 166. Li, X. et al. Evidence of nitrogen loss from anaerobic ammonium oxidation coupled with ferric iron reduction in an intertidal wetland. Environ. Sci. Technol. 49, 11560–11568 (2015). 167. Zhou, G.-W. et al. Electron shuttles enhance anaerobic ammonium oxidation coupled to iron(III) reduction. Environ. Sci. Technol. 50, 9298–9307 (2016). 168. Yang, W. H., Weber, K. A. & Silver, W. L. Nitrogen loss from soil through anaerobic ammonium oxidation coupled to iron reduction. Nat. Geosci. 5, 538–541 (2012). 169. Li, X. et al. Simultaneous Fe(III) reduction and ammonia oxidation process in Anammox sludge. J. Environ. Sci. 64, 42–50 (2018). 170. Huang, S. & Jaffé, P. R. Isolation and characterization of an ammonium-oxidizing iron reducer: Acidimicrobiaceae sp. A6. PLoS ONE 13, e0194007 (2018). 172. Zhu, X., Burger, M., Doane, T. A. & Horwath, W. R. Ammonia oxidation pathways and nitrifier denitrification are significant sources of N2O and NO under low oxygen availability. Proc. Natl. Acad. Sci. USA 110, 6328–6333 (2013). 173. Ginn, B., Meile, C., Wilmoth, J., Tang, Y. & Thompson, A. Rapid iron reduction rates are stimulated by high-amplitude redox fluctuations in a tropical forest soil. Environ. Sci. Technol. 51, 3250–3259 (2017). A good example of the dynamic nature of iron cycling in the environment and its impact on the reducibility of minerals. 174. Mejia, J., Roden, E. E. & Ginder-Vogel, M. Influence of oxygen and nitrate on Fe (hydr)oxide mineral transformation and soil microbial communities during redox cycling. Environ. Sci. Technol. 50, 3580–3588 (2016). 175. Laufer, K. et al. Coexistence of microaerophilic, nitrate-reducing, and phototrophic Fe(II) oxidizers and Fe(III) reducers in coastal marine sediment. Appl. Environ. Microbiol. 82, 1433–1447 (2016). 176. Hansel, C. M., Ferdelman, T. G. & Tebo, B. M. Cryptic cross-linkages among biogeochemical cycles: novel insights from reactive intermediates. Elements 11, 409–414 (2015). A review on cryptic element cycling in the environment, including cryptic iron cycling. 177. Klueglein, N. & Kappler, A. Abiotic oxidation of Fe(II) by reactive nitrogen species in cultures of the nitrate-reducing Fe(II) oxidizer Acidovorax sp. BoFeN1 – questioning the existence of enzymatic Fe(II) oxidation. Geobiology 11, 180–190 (2013). 178. Matus, F. et al. Ferrous wheel hypothesis: Abiotic nitrate incorporation into dissolved organic matter. Geochim. Cosmochim. Acta 245, 514–524 (2019). Demonstration of the ‘ferrous wheel hypothesis’ with insights for the role of coupled iron and nitrogen cycling in the environment. 179. Chen, C., Hall, S. J., Coward, E. & Thompson, A. Iron-mediated organic matter decomposition in humid soils can counteract protection. Nat. Commun. 11, 2255 (2020). 180. Patzner, M. S. et al. Iron mineral dissolution releases iron and associated organic carbon during permafrost thaw. Nat. Commun. 11, 6329 (2020). 181. Beckwith, C. R. et al. Characterization of MtoD from Sideroxydans lithotrophicus: a cytochrome c electron shuttle used in lithoautotrophic growth. Front. Microbiol. 6, 332 (2015). 182. Bird, L. J., Bonnefoy, V. & Newman, D. K. Bioenergetic challenges of microbial iron metabolisms. Trends Microbiol. 19, 330–340 (2011). 183. Field, S. J. et al. Purification and magneto-optical spectroscopic characterization of cytoplasmic membrane and outer membrane multiheme c-type cytochromes from Shewanella frigidimarina NCIMB400. J. Biol. Chem. 275, 8515–8522 (2000). 185. Salmon, T. P., Rose, A. L., Neilan, B. A. & Waite, T. D. The FeL model of iron acquisition: nondissociative reduction of ferric complexes in the marine environment. Limnol. Oceanogr. 51, 1744–1754 (2006). 186. Navrotsky, A., Mazeina, L. & Majzlan, J. Size-driven structural and thermodynamic complexity in iron oxides. Science 319, 1635–1638 (2008). 187. Gorski, C. A., Edwards, R., Sander, M., Hofstetter, T. B. & Stewart, S. M. Thermodynamic characterization of iron oxide–aqueous Fe2+ redox couples. Environ. Sci. Technol. 50, 8538–8547 (2016). One of the first examples of using electrochemical methods to better understand the range of redox potentials present in different iron phases. 188. Robie, R. A. & Heminway, B. S. Thermodynamic properties of minerals and related substances at 298.15 K and 1 bar (105 pascals) pressure and at higher temperatures. (United States Printing Office, 1995). 189. Navrotsky, A., Ma, C., Lilova, K. & Birkner, N. Nanophase transition metal oxides show large thermodynamically driven shifts in oxidation-reduction equilibria. Science 330, 199–201 (2010). 190. Robie, R. A. & Bethke, P. Molar Volumes and Densities of Minerals. Report TEI-822 (United States Department of the Interior Geological Survey, 1962). 191. Gorski, C. A., Nurmi, J. T., Tratnyek, P. G., Hofstetter, T. B. & Scherer, M. M. Redox behavior of magnetite: implications for contaminant reduction. Environ. Sci. Technol. 44, 55–60 (2010). 192. Gorski, C. A., Klüpfel, L. E., Voegelin, A., Sander, M. & Hofstetter, T. B. Redox properties of structural Fe in clay minerals: 3. Relationships between smectite redox and structural properties. Environ. Sci. Technol. 47, 13477–13485 (2013). 193. Oswald, K. et al. Aerobic gammaproteobacterial methanotrophs mitigate methane emissions from oxic and anoxic lake waters. Limnol. Oceanogr. 61, S101–S118 (2016). 194. Braunschweig, J., Bosch, J. & Meckenstock, R. U. Iron oxide nanoparticles in geomicrobiology: from biogeochemistry to bioremediation. N. Biotechnol. 30, 793–802 (2013). 195. Villa, R. D., Trovó, A. G. & Nogueira, R. F. P. Environmental implications of soil remediation using the Fenton process. Chemosphere 71, 43–50 (2008). 196. Wagai, R. & Mayer, L. M. Sorptive stabilization of organic matter in soils by hydrous iron oxides. Geochim. Cosmochim. Acta 71, 25–35 (2007). 197. Nitzsche, K. S. et al. Arsenic removal from drinking water by a household sand filter in Vietnam — effect of filter usage practices on arsenic removal efficiency and microbiological water quality. Sci. Total. Environ. 502, 526–536 (2015). 198. Sipos, P., Németh, T., Kis, V. K. & Mohai, I. Sorption of copper, zinc and lead on soil mineral phases. Chemosphere 73, 461–469 (2008). 199. Poulton, S. W. & Canfield, D. E. Development of a sequential extraction procedure for iron: implications for iron partitioning in continentally derived particulates. Chem. Geol. 214, 209–221 (2005). 200. Schaedler, F., Kappler, A. & Schmidt, C. A revised iron extraction protocol for environmental samples rich in nitrite and carbonate. Geomicrobiol. J. 35, 23–30 (2018). 201. Porsch, K. & Kappler, A. FeII oxidation by molecular O2 during HCl extraction. Environ. Chem. 8, 190–197 (2011). 202. Roden, E. E. & Zachara, J. M. Microbial reduction of crystalline iron(III) oxides: Influence of oxide surface area and potential for cell growth. Environ. Sci. Technol. 30, 1618–1628 (1996). 203. Tessier, A., Campbell, P. G. C. & Bisson, M. Sequential extraction procedure for the speciation of particulate trace metals. Anal. Chem. 51, 844–851 (1979).

Kappler Web Traffic

Page Views per User (PVPU)
Page Views per Million (PVPM)
Reach per Million (RPM)
CBI Logo

Kappler Rank

Kappler Frequently Asked Questions (FAQ)

  • When was Kappler founded?

    Kappler was founded in 1976.

  • Where is Kappler's headquarters?

    Kappler's headquarters is located at 55 Grimes Drive, Guntersville.

  • What is Kappler's latest funding round?

    Kappler's latest funding round is Loan.

  • How much did Kappler raise?

    Kappler raised a total of $2.49M.

  • Who are the investors of Kappler?

    Investors of Kappler include Paycheck Protection Program, Catapult, Enterprise Partners Venture Capital, East Midlands Early Growth Fund and Coalfields Enterprise Fund.

Discover the right solution for your team

The CB Insights tech market intelligence platform analyzes millions of data points on vendors, products, partnerships, and patents to help your team find their next technology solution.

Request a demo

CBI websites generally use certain cookies to enable better interactions with our sites and services. Use of these cookies, which may be stored on your device, permits us to improve and customize your experience. You can read more about your cookie choices at our privacy policy here. By continuing to use this site you are consenting to these choices.