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Formation and stabilization of elemental sulfur through
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organomineralization
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Julie Cosmidisa*1, Christine W. Nimsa1, David Diercksb, Alexis S. Templetona
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a
Department of Geological Sciences, University of Colorado, Boulder, CO 80309, USA
5
b
Department of Metallurgical and Materials Engineering, Colorado School of Mines, Golden, CO
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80401, USA
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*Corresponding author. Email address:
[email protected]
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Abstract
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Elemental sulfur (S(0)) is an important intermediate in the biogeochemical cycle of sulfur that is
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formed by chemical or biological oxidation of more reduced sulfur species. Given the restricted
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geochemical conditions under which S(0) should persist, the mechanisms whereby S(0) can be
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stabilized in the environment are not fully understood. Here we identify a process called “S(0)
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organomineralization”, by which S(0) minerals are produced and stabilized following the
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oxidation of hydrogen sulfide in the presence of numerous types of dissolved organics, including
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simple sugars and amino acids. The S(0) particles formed through this mechanism are closely
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associated with organics, which often form an envelope around the mineral. The organic envelopes
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are formed by self-assembly of the dissolved organic molecules in the presence of hydrogen sulfide
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and oxygen, and play in a role in the stabilization of S(0). Organic compound sulfurization
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probably plays an important role in the self-assembly mechanism, by causing the polymerization
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Present address: Department of Geosciences, Pennsylvania State University, University Park, PA 16802, USA
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of relatively small dissolved organic molecules into solid, macromolecular, polymeric organics.
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The organomineralized S(0) particles present unique and complex morphologies, which are
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controlled by the type of dissolved organic compound present in the experimental media.
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Depending on the organics present, organomineralized S(0) can exist as different combinations of
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several crystal structures, including the non-thermodynamically stable β- and γ-S8 allotropes,
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which are most likely stabilized by their close association with the organic phase. We propose that
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complex particle morphology combined with the presence of metastable S(0) allotropes could be
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used as a signature of S(0) organomineralization in natural settings. S(0) organomineralization was
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obtained in the laboratory under a wide range of experimental conditions that span across
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geochemical conditions which can be encountered in many sulfidic environments. It is possible
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that the reaction between reduced sulfur species and organics may significantly affect the
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production and preservation of S(0) in numerous natural systems.
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1. Introduction
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1.1.
Elemental sulfur formation in the environment
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Elemental sulfur (S(0)) is an important intermediate in the biogeochemical cycle of sulfur that is
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formed by the oxidation of more reduced species such as sulfide (Zopfi et al., 2004). S(0) is found
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in a variety of Earth surface environments such as marine sediments (Jørgensen and Nelson, 2004;
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Zopfi et al., 2004) and water columns (Jørgensen et al., 1991; Luther et al., 1991; Findlay et al.,
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2014), euxinic lakes (Zerkle et al., 2010; Kamyshny et al., 2011), caves (Galdenzi et al., 2008;
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Hamilton et al., 2015), aquifers (Einsiedl et al., 2015), hydrothermal vents (Taylor et al., 1999;
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Embley et al., 2007; Findlay et al., 2014; Gilhooly et al., 2014), as well as sub-glacial or hot springs
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(Boyd et al., 2007; Gleeson et al., 2011; Gleeson et al., 2012; Kamyshny et al., 2014). S(0) is
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consumed by a wide diversity of microorganisms that can gain energy from its oxidation (Schmidt
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et al., 1987; Suzuki et al., 1993; Franz et al., 2007; Marnocha et al., 2016), reduction (Boyd and
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Druschel, 2013) or disproportionation (Thamdrup et al., 1993; Canfield and Thamdrup, 1994;
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Finster et al., 1998).
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The chemical oxidation of sulfide to S(0) by molecular oxygen occurs at rates that are several
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orders of magnitude lower than the rate of microbial sulfide oxidation (Luther et al., 2011), and so
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it is assumed that in low-temperature environments, most S(0) formation results from microbial
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oxidation. Many different S-oxidizing bacteria are indeed known to biomineralize S(0) in the form
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of intracellular or extracellular S(0) globules (Kleinjan et al., 2003), or as extracellular S(0)
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filaments (Wirsen et al., 2002; Sievert et al., 2007). However, the microbial and enzymatic
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pathways for S(0) formation are still not completely understood. S(0) is considered a rather
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unstable and dynamic constituent of the sulfur pool of sediments (Troelsen and Jørgensen, 1982;
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Zopfi et al., 2004), and it is only thermodynamically stable under a very restricted range of Eh and
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pH conditions (Gleeson et al., 2010). S(0) can however persist and accumulate in some
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environments where it forms conspicuous deposits, although it is unclear what are the mechanisms
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responsible for its stabilization.
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1.2.
S(0) organomineralization: definition
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Recently, it was discovered that S(0) could be formed abiogenically through the interaction of
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hydrogen sulfide with dissolved organics in the presence of an oxygen gradient (Cosmidis and
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Templeton, 2016). The S(0) particles produced through this process presented filamentous and
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spherical morphologies and were encapsulated within an organic envelope. This reaction, i.e. the
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formation of S(0) where organics appear to play a necessary role, can be qualified as an
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organomineralization process.
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Organomineralization is a relatively recent concept in Earth Sciences, since its first occurrence in
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the literature of this field only dates back from about two decades ago (Défarge and Trichet, 1995;
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Trichet and Défarge, 1995). Contrary to microbial biomineralization, which has been studied since
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at least the late 19th century (Robinson, 1889), the importance of organomineralization in mineral
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formation in the environment has been widely under investigated. There has been some vigorous
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debate over the definition of this term (Perry et al., 2007; Altermann et al., 2009; Défarge et al.,
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2009; Perry and Sephton, 2009; Défarge et al., 2010), but a consensus seems to have been reached
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in
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“Organomineralization is the process of mineral formation mediated by organic matter,
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independent of the living organisms which the organic matter derives from”. The organic
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compounds may be by-products of biotic activity, relics of dead and decaying organisms, or non-
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biological
the definition provided by the
organic
compounds.
Encyclopedia
Importantly,
the
of
terms
Geobiology
(Défarge, 2011):
“organomineralization”
and 4
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“organominerals” should be used only when organic compounds have played an active role in the
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mineral precipitation, and not in the case when they have been passively entombed or complexed
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during crystal growth (Défarge et al., 2009).
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Some organomineralization processes have been thoroughly researched by chemists and material
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scientists, such as the synthesis of mesoporous organic-silica materials (Hoffmann et al., 2006) or
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organic–inorganic zeolites (Meng and Xiao, 2014), for applications such as heterogeneous
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catalysis or sensor technologies. Most studies published by geoscientists so far describing
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organomineralization processes have focused on the organically-mediated formation of calcium
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carbonates in modern and ancient microbialites (e.g,, Riding, 2000; Perry et al., 2007; Dupraz et
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al., 2009; Pacton et al., 2015; Bindschedler et al., 2016; Diaz et al., 2017), However,
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organomineralization reactions could also play a role in the formation of sulfides (Maclean et al.,
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2008), and sulfate minerals (Cámara et al., 2016) in a diversity of natural environments.
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The control exerted by organics on mineral nucleation and/or crystallization and growth has been
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investigated in many experimental studies with calcium carbonates (Meldrum and Hyde, 2001;
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Reitner, 2004; Braissant et al., 2007; Gallagher et al., 2013), calcium phosphates (Silverman and
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Boskey, 2004; Kniep and Simon, 2007; Gallagher et al., 2013; Crosby and Bailey, 2017), silica
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(Behrens et al., 2007; Delclos et al., 2008; Ahmed et al., 2010), iron oxides (Mirabello et al., 2016),
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iron sulfides (Grimes et al., 2001), sulfate minerals (Heywood and Mann, 1994; Borah et al., 2006),
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and metallic nanoparticles (Rashid et al., 2007; Alexandridis and Tsianou, 2011; Shah et al., 2015).
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Organomineralization often results in the formation of minerals with specific properties such as
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shape (Cody and Cody, 1991; Mann and Ozin, 1996; Meldrum and Hyde, 2001; Meldrum and
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Cölfen, 2008; Wu et al., 2010; Biacchi and Schaak, 2015), size (Ananikov et al., 2007; Kuwahara
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et al., 2014; Nakaya et al., 2014), crystal structure (Tong et al., 2004; Sand et al., 2012; Tobler et 5
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al., 2014; Lu et al., 2015; Moliner, 2015), chemical composition (Tobler et al., 2015) or even
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isotopic composition (Harouaka et al., 2017) that differ from minerals precipitated under purely
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inorganic conditions. Such special properties might be used as signatures of organomineralization
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processes in the rock record or in extraterrestrial samples (Reitner, 2004).
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The mechanism of S(0) organomineralization, as well as the potential contribution of this process
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to environmental S(0) formation, remains elusive. Most of what is known about the geochemical
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interactions of sulfur with organics has been derived from studies of the sulfurization of organic
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matter in sediments (e.g., Werne et al., 2004) or water columns (e.g., Raven et al., 2016). Some
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previous studies have shown that redox-active functional groups in organic matter such as
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quinones could oxidize sulfide to S(0) and thiosulfate (Heitmann and Blodau, 2006; Yu et al.,
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2015). However, the mechanism by which hydrogen sulfide reacts with non redox-active dissolved
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organics to produce S(0) needs to be elucidated. A better understanding of the mechanism of S(0)
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organomineralization, and of its potential role in the formation and/or preservation of S(0) in the
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environment, has the potential to reshape our knowledge of the sulfur biogeochemical cycle and
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its interactions with the carbon cycle. This requires better constraints on the geochemical
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conditions that are conducive to S(0) organomineralization. It also requires identifying the specific
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signatures of organomineralized S(0) that might be used to discriminate it from microbially- and
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inorganically-formed S(0) in the environment.
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In this study, we performed new experiments where several different types of dissolved organic
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compounds were reacted with hydrogen sulfide and oxygen, forming organomineralized S(0). In
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the original experiments by Cosmidis and Templeton (2016), S(0) organomineralization was
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performed using yeast extract and/or peptone – which are both complex mixtures of organic
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compounds – as the source of organics. Here we used well-defined organic compounds, such as 6
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simple sugars and amino acids, or more complex organics representative of natural organic matter
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(humic acids). The experiments were also performed under a wide range of geochemical
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conditions, including conditions representative of natural sulfidic environments. The combined
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results enable us to determine the geochemical parameters and specific types of organics conducive
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to S(0) organomineralization. We furthermore performed new characterizations of
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morphology, chemical speciation, and crystal structure of the organomineralization products, that
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allowed us to identify the specific signatures of organomineralized S(0). These experiments
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strongly demonstrate that dissolved organic compounds can directly influence the formation and/or
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stabilization of S(0) under our experimental conditions, and pave the way for future studies that
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will investigate the prevalence of S(0) organomineralization in the environment.
2. Methods: experimental
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the
2.1.
Gradient tubes experiments
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S(0) organomineralization experiments were performed in sulfide gradient tubes (Fig. EA-1A)
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prepared as described in Cosmidis and Templeton (2016). This setup was originally derived from
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a microbial culture experiment (Gleeson et al., 2011). In short, the gradient tubes consisted of a
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bottom layer containing an artificial mineral medium (modified EM medium), agar (1% w/v) and
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Na2S (5 mM), and a top layer containing the same mineral medium supplemented with vitamins
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and trace elements (see section 2.2 below) as well as different types of dissolved organic
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compounds at a 5 g.L-1 concentration: D(+)-glucose (Sigma-Aldrich), D(+)-cellobiose (Acros
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Organics), glycine (Acros Organics), casamino acids (Fisher Scientific), yeast extract (Fisher
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Scientific) or humic acids (humic acid sodium salt, Aldrich). In the case of humic acids, the top
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layer solution was filtered through 0.2 μm filters, to remove particulate material. The top layer
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solutions were de-oxygenated by sparging with N2:CO2 (80:20) before being added to the tubes.
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The use of a N2:CO2 gas mix instead of pure N2 was inherited from the original gradient tube
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protocol, which was designed for the growth of autotrophic bacteria (Gleeson et al., 2011). The
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top layer of each tube was oxygen-free when the experiment was started, but oxygen from the
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atmosphere was allowed to diffuse from the top, creating an oxygen gradient opposing the sulfide
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gradient. All the solutions, as well as the glass tubes themselves, were sterilized by autoclaving
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prior to the preparation of the gradient tubes, which was performed under aseptic conditions. The
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gradient tubes were kept in the dark (to exclude the possibility of photochemical reactions) and at
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room temperature during the duration of the experiments.
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2.2.
Batch experiments
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S(0) organomineralization was also performed in simplified “batch” experiments (Fig. EA-1B) in
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glass bottles (100 mL to 1 L) containing modified EM medium supplemented with dissolved
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organics at different concentrations ranging from 0.1 mg.L-1 to 15 g.L-1. Phosphate (K2HPO4) was
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omitted in order to avoid phosphate mineral precipitation. The organics tested were glucose,
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glycine, humic acids, and yeast extract. The bottles were autoclaved, allowed to cool down, and
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sparged with N2:CO2 (80:20) under aseptic conditions, before filter-sterilized solutions of vitamins
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(Wolfe’s vitamin solution; Atlas, 2010) and trace elements (0.52 g EDTA, 0.15 g FeCl2·4H2O, 7
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mg ZnCl, 10 mg MnCl·4H2O, 6.3 mg H3BO3, 19 mg CoCl2·6H2O, 1.7 mg CuCl2·2H2O, 24 mg
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NiCl2·6H2Oand 36 mg Na2MoO4·2H2O per 100mL deionized water) were added, both at 1 mL.L-
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1
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The solutions containing humic acids needed to be filtered through 0.2 μm filters (Millipore
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Steritop bottle top filter unit) to remove particulate material before sulfide was added in. The
. Filter-sterilized Na2S was finally added at final concentrations ranging from 50 μM to 5 mM.
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bottles were loosely capped, allowing for oxygen from the atmosphere to diffuse in. The
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experiments were stored in the dark at room temperature.
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Alterations to this protocol were made in order to test the effects of different geochemical
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parameters on S(0) organomineralization. In a set of experiments, the solutions were not
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autoclaved but were instead sterilized by vacuum filtration through 0.2 μm filters (Millipore
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Steritop bottle top filter units). An experiment was performed in which the bottles were stirred
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with sterile magnetic stirrers, in order to test the effect of agitation (i.e. preventing the formation
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of sulfide and oxygen gradients) on S(0) organomineralization. In another set of experiments, the
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N2:CO2 gas mix used to sparge the bottles was replaced with pure N2. Finally, experiments were
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performed in which the chemical composition of the medium was altered: trace elements and/or
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vitamins were omitted, or the EM medium was replaced with distilled water, or distilled water
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with 2.75 g.L-1 NaCl. These different experimental conditions are summarized in Table EA-1.
3. Methods: analytical
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3.1.
Light Microscopy
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Light microscopy was performed with a Zeiss Axio Imager Z1 on wet samples collected from the
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gradient tube and batch experiments. For each sample, a ~20 μL drop of liquid was mounted
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between a glass slide and a cover slip and imaged before it dried..
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3.2.
Scanning Electron Microscopy
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Samples from the gradient tubes and batch experiments were collected at different time intervals
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throughout the course of the experiments, deposited on polycarbonate filters (GTTP Isopore
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membrane filters, Merck Millipore, pore size 0.2 μm), and rinsed three times with distilled water.
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The filters were allowed to dry at ambient temperature and were coated with carbon or gold prior 9
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to analysis. The analyses were conducted on a JSM-7401F field emission Scanning Electron
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Microscope (SEM) at the Nanoscale Fabrication Laboratory at the University of Colorado at
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Boulder. Images were acquired in the secondary electron mode with the microscope operating at
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3 kV and a working distance (WD) of ~3 mm, and in the backscattered electron mode at 10 kV
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and WD ~8 mm. Energy-Dispersive X-ray Spectrometry (XEDS) analyses were performed at 20
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kV and WD ~8 mm.
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3.3.
Transmission Electron Microscopy
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Samples were collected by centrifugation from batch experiments, and rinsed three times with
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deionized water. They were then deposited on 200 mesh Cu grids covered with a lacy
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formvar/carbon film (Ted Pella), and allowed to air-dry at ambient temperature.
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A Philips CM200 transmission electron microscope operated at 200keV was used for imaging and
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selected area electron diffraction (SAED) of the sulfur particles. The particles proved to be fairly
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sensitive to electron beam exposure, with S(0) sublimating within a few minutes in the high-
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vacuum TEM chamber while being analyzed. The beam intensity was thus decreased through
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adjusting the spot size to allow more time for imaging and diffraction. A series of particle images
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and diffraction patterns were collected. The d-spacings, d-spacing ratios, and angles between
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planes observed in the diffraction patterns were compared to the known crystal structures of α-, β-
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, and γ-cyclooctasulfur from the American Mineralogist Crystal Structure Database. From this,
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diffraction patterns of several particles were indexed using the JEMS software package
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(http://www.jems-saas.ch/).
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3.4.
Scanning Transmission X-ray Microscopy
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Samples from the gradient tubes experiments were rinsed three times in deionized water and a
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small drop (~2 μL) was deposited on a Formvar-coated 200 mesh Cu TEM grid (Ted Pella) and
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allowed to air-dry at ambient temperature. Scanning Transmission X-ray Microscopy (STXM)
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analyses were performed on beamline 10ID-1 (SM) of the Canadian Light Source (Saskatoon,
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Canada) (Kaznatcheev et al., 2007) and beamline 11.0.2 of the Advanced Light Source (Berkeley,
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USA) (Bluhm et al., 2006). The X-ray beam was focused on the samples using a Fresnel zone plate
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objective and an order-sorting aperture yielding a focused X-ray beam spot of ~30 nm. After
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sample insertion in STXM, the chamber was evacuated to 100 mTorr and back-filled with He at
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~1 atm pressure. Energy calibration was achieved using the well-resolved 3p Rydberg peak of
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gaseous CO2 at 294.96 eV. Images, maps and image stacks were acquired in the 260-340 eV (C
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K-edge)
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(unicorn.mcmaster.ca/aXis2000.html) was used for data processing. A linear background
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correction was applied to the spectra at the C K and S L-edges, in the 260-280 eV region and 155-
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160 eV region, respectively, to eliminate the contribution of lower energy absorption edges. All
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spectra were normalized to the same maximum.
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Maps of organic C were obtained by subtracting an image obtained at 280 eV (i.e. below the C K-
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edge) and converted into optical density (OD) from an OD-converted image at 288.2 eV (1s →π*
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electronic transitions in peptides). Maps of S were obtained by subtracting an OD-converted image
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obtained at 160 eV (i.e. below the S L-edge) from an OD-converted image at 163.5 eV (energy of
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the S L3-edge). X-ray absorption near edge structure (XANES) spectra were extracted from image
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stacks as explained in Cosmidis and Benzerara (2014). The S L-edge spectra of a reference
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elemental sulfur compound (precipitated sulfur, Fisher Scientific) as well as of several
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organosulfur compounds (L-cystine, L-methionine, and L-cysteine) were also acquired using the
and
155-190
eV
(S
L-edge)
energy
ranges.
The
aXis2000
software
11
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same methods. Similarly, the C K-edge spectra of reference organic compound powders (glucose,
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cellobiose, glycine, casamino acids, and humic acids), were also obtained.
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3.5.
Fourier-Transform Infrared Spectroscopy
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Samples from the batch experiments were harvested by centrifugation, rinsed three times with
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deionized water, and air-dried. The powdered samples were then pressed into pellets in
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approximately 5% KBr. Fourier-Transform Infrared (FTIR) spectra were recorded between 4000
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and 400 cm−1 on a Thermo Nicolet NEXUS 670 FTIR spectrometer. Each spectrum was an
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integration of 150 spectral scans, with a wavenumber resolution of 1 cm−1. Background corrections
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were applied to the data following each measurement to compensate for instrumental noise and
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contributions from atmospheric CO2 and H2O by dividing the absorbance of the sample spectrum
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by the background spectrum at each data point. The spectra of reference organic compounds
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powders (glucose, glycine, yeast extract and humic acids) were acquired using the same method.
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For comparative purposes, all the spectra were normalized to a maximum absorbance of 1.
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3.6.
X-ray Diffraction
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Samples from the batch experiments were rinsed three times and concentrated in deionized water.
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The slurry was pipetted on a Si wafer to form a thin layer on the wafer surface. The wafer was
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gently heated on a slide warmer to dry the sample and create a thin film of sample on the wafer
251
surface. In order the check to stability of the sulfur phases in the atmosphere, some of the samples
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were “aged” by letting them air dry on the Si wafer surface for ten days before being analyzed.
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Samples were analyzed using a Siemens D500 X-ray diffractometer from 5 to 65 degrees two theta
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using Cu Kα X-ray radiation, with a step size of 0.02 degrees and a dwell time of 2 seconds per
12
255
step. Different S0 crystal structures were identified using Jade software (MDI, version 9) and the
256
International Centre for Diffraction Data (ICDD) 2003 database.
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3.7.
Geochemical profiles in gradient tube experiments
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The concentrations of dissolved sulfide and oxygen, as well as the pH, were measured at different
259
time intervals in gradient tube experiments performed either without added organics, or with added
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glucose, glycine, humic acids or yeast extract.
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The dissolved hydrogen sulfide (H2S) and oxygen profiles were measured using Unisense (Aarhus,
262
Denmark) H2S-100 and OX-100 microsensors, respectively. The oxygen microsensor was
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calibrated using freshly prepared de-oxygenated water (0% oxygen saturation) as well as a well-
264
aerated water (100% oxygen saturation). The H2S microsensor was calibrated using freshly
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prepared de-oxygenated solutions of Na2S in an acetate buffer (pH
