Desulfovibrio inopinatus (cat. no. 10711) was obtained from the German Collection of Microorganisms and Cell Cultures (DSMZ, Braunschweig, Germany: Catalogue of strains 1993). This microbe was isolated from marine sediments of Venice, Italy (Reichenbecher and Schink, 1997). Cultures were maintained anaerobically in glass serum bottles capped with butyl rubber septa under an atmosphere containing 80% N₂ and 20% CO₂. Culture medium was prepared according to DSMZ recipe #196-13871, with the exception of NaCl concentration because D. inopinatus grew better at 21 g/L than at the 7 g/L concentration suggested in the DSMZ recipe. The medium contains (per liter): NaHCO₃, 9.0 g; Na₂SO₄, 3.0 g; KH₂PO₄, 0.20 g; NH₄Cl, 0.30 g; NaCl, 21 g; KCl, 0.50 g, MgCl₂·6H₂O, 3.1 g; CaCl₂·2H₂O, 0.15 g; resazurin, 1 mg; a trace element solution (SL-10), 1 ml (Widdel and Pfennig, 1981; Imhoff-Stuckle and Pfennig, 1983), and a general vitamin mix (described in DSMZ recipe #141), 10 ml. Sodium ascorbate (1.5 g/L) was added to maintain anoxic conditions (Kligler and Guggenheim, 1938). The medium did not contain any yeast extract. The pH of the medium was adjusted to 7.5 by dropwise addition of 1 M NaOH or 1 M HCl prior to inoculation.

Culture medium was supplemented with lactate (28 mM), malate (28 mM), or fructose (14 mM) as organic substrate and sole electron donor. Initial concentrations of organic substrates were chosen such that sulfate would be in excess: sulfate reduction with stoichiometric conversion of any substrate to acetate would produce only ~14 mM of sulfide from the initial 21 mM sulfate, see Table 1. For each experiment, 7–15 bottles were inoculated with 5% (v/v) of a mid- to late-exponential phase pre-culture grown on the same organic substrate as that used in the experiment. Inoculums were pelleted via centrifugation and rinsed three times with clean medium in an anaerobic chamber (under an atmosphere of 80% N₂, 15% CO₂, and 5% H₂) to ensure the removal of all residual sulfide and organic substrate, and then transferred into 40 mL of fresh media. Bottles were incubated in the dark at room temperature. Microbial growth was monitored daily in one “master” bottle to minimize puncturing of septa of the experimental bottles. At each time point (Tables 2, 3), one experimental bottle was subsampled for cell counts and the colorimetric assay of sulfide concentration. The remaining culture volume was sacrificed by the addition of a 1 M zinc acetate solution (at a 10% v/v ratio) to terminate microbial activity and precipitate sulfide as ZnS for isotopic analyses.

To test the effect of phosphate limitation, additional batch cultures were grown with varying initial concentrations of phosphate in the medium. Serum bottles used for these cultures were autoclaved three times with deionized water (18.2 MΩ·cm, Barnstead Nanopure™ filtration system) to remove residual phosphate. Medium was prepared as described above, with lactate (28 mM) as organic substrate and without added phosphate. Phosphate (as KH₂PO₄) was then added from a concentrated stock solution, to final concentrations of 150, 15, and 3 μM. Another set of serum bottles did not receive any added phosphate. The concentration of phosphate in this nominally “0” phosphate medium was estimated to be <1 μM based on lot analyses provided with the chemicals used in the media (see also Experiments with Different Phosphate Concentrations). Figure A2 shows that ~3 to 5 μM of added phosphate is limiting, suggesting that the blank is likely <5 μM. Conclusions reached in this paper would be unaffected by a blank of 5 μM or less. Experimental bottles (containing 10 mL of fresh media) were inoculated with a 5% (v/v) sample of pre-cultures that were already conditioned to growth in the presence of lower or limiting concentrations of phosphate. Cell densities were low when phosphate was limiting, so inoculums were pelleted and rinsed only once to minimize the loss of biomass prior to inoculation. Cultures were monitored and sampled as described above.

Desulfovibrio fructosovorans strain JJ (cat. no. 3604) was obtained from DSMZ. This strain was isolated from estuarine sediment (Jones et al., 1984; Cord-Ruwisch et al., 1986), and it incompletely oxidizes pyruvate, lactate, or fructose to acetate (Ollivier et al., 1988). The mineral medium (modified from DSMZ recipe #63) contained (per liter): NaHCO₃, 2.6 g; Na₂SO₄, 3.0 g; KH₂PO₄, 0.30 g; NH₄Cl, 0.50 g; KCl, 0.20 g; CaCl₂, 0.10 g; MgCl₂, 2.0 g; resazurin, 1 mg; SL-10 trace element solution (see above), 1 ml; vitamin solution #141 (see above), 10 ml; and sodium ascorbate, 1.5 g (see above). Media were supplemented with limiting concentrations of one of the following organic substrates: lactate, 20 mM; pyruvate, 40 mM; or fructose, 10 mM. The starting concentrations of lactate, fructose, and pyruvate were chosen such that 10 mM of sulfide would be produced by stoichiometric conversion of the substrate to acetate (Table 1). The pH of the medium was adjusted to 7.0 before inoculation. Bottles for all experiments with D. fructosovorans were inoculated with 5% (v/v) of a late-exponential phase pre-culture grown on lactate. Media and cultures were otherwise prepared and maintained as described for Desulfovibrio inopinatus above.

Desulfovibrio sp. DMSS-1 (henceforth “DMSS-1”), isolated from a salt marsh on Cape Cod, Massachusetts, USA and characterized by Sim et al. (2011b), was grown in batch cultures in the presence of varying concentrations of phosphate. Serum bottles were cleaned and prepared in the same manner as described above for D. inopinatus. Medium for DMSS-1 was prepared following Sim et al. (2011b), with the exception of phosphate, and contained (per liter): NaHCO₃, 9 g; Na₂SO₄, 3 g; NH₄Cl, 0.3 g; NaCl, 21 g; KCl, 0.5 g; MgCl₂·6H₂O, 6 g; CaCl₂·2H₂O 0.3 g; resazurin, 1 mg; SL-10 trace element solution (see above), 1 ml; vitamin solution #141 (see above), 10 ml; 1 ml of selenium stock solution (0.4 mg of Na₂SeO₃ per 200 ml of 0.01 N NaOH); and sodium ascorbate, 1.5 g (see above). Phosphate was added to final concentrations of 360, 36, 5 μM, and “0” (<1 μM).

Growth was monitored using optical density (OD) measurements and microscopic cell counts. OD was measured at 630 or 660 nm using a spectrophotometer (Synergy 2 microplate reader, BioTek, Winooski, Vermont, USA). For cell counts, subsamples of experimental cultures were preserved in 2.5% glutaraldehyde at 4°C. Preserved cells were stained with SYBR Green I nucleic acid stain (Invitrogen Molecular Probes, Eugene, Oregon, USA), and filtered onto Whatman 0.2 μm Nuclepore polycarbonate filters. Stained cells were visualized and imaged by epifluorescence microscopy using a Zeiss Axio Imager M1 microscope (Carl Zeiss Microscopy, LLC), and cell densities were determined by manual counting (Noble and Fuhrman, 1998). Cell lengths and widths were measured in the epifluorescence micrographs using measuring tools in Zeiss AxioVision software.

Sulfide from subsamples was precipitated as ZnS in a 50 mM zinc acetate solution, and stored at 4°C until analysis. Sulfide concentrations (ΣH₂S = H₂S + HS⁻ + S²⁻) were determined by a modified methylene blue colorimetric method (Cline, 1969). Briefly, 200 μL samples of medium were reacted with 1 ml of 0.05 M zinc acetate and 10 μL of N,N-dimethyl-p-phenylenediamine sulfate solution. Optical density was read at 670 nm using a microplate reader (Sim et al., 2011b).

Sulfide was extracted for isotopic analysis by acidifying each ZnS sample with 3 to 6 M HCl, and gently boiling for 1 h under a stream of N₂ gas. Sulfate in samples were then converted to H₂S using a general reducing agent (HCl, HI, and H₃PO₂) (Thode et al., 1961; Forrest and Newman, 1977; Arnold et al., 2014). Volatiles were passed through a condenser and a distilled H₂O trap. H₂S produced in the reactions was precipitated as ZnS in zinc acetate, and then converted to Ag₂S via addition of silver nitrate, or precipitated as Ag₂S directly in a silver nitrate solution. The recovered Ag₂S was washed with deionized water, dried at 70°C, and converted to SF₆ by reaction with F₂ at 300°C overnight. The SF₆ product was purified by cryogenic trapping and preparative gas chromatography, and analyzed on a ThermoFinnigan MAT 253 isotope-ratio mass spectrometer operated in dual-inlet mode as described previously (Ono et al., 2006).

Sulfur isotope values are reported in the standard δ notation against Vienna Cañon Diablo Troilite (VCDT):

Following IUPAC recommendations (Coplen, 2011), we have omitted the factor of 1000‰ from the definition of δ in Equation 1.

Both D. fructosovorans and D. inopinatus discriminate against heavier isotopes of sulfur more when they grow on fructose (³⁴ε ~ 30‰) relative to the growth on lactate (10 to 15‰). Data from Sim et al. (2011b) for DMSS-1 growing on fructose (30 to 40‰) and lactate (~6‰) in batch cultures exhibited the same trend and similar ranges of ³⁴ε. Growth on pyruvate and malate yielded ³⁴ε values that were dissimilar among the studied species. While fractionations during the growth on pyruvate were larger than those during the growth on lactate and smaller than those during the growth on fructose for both D. fructosovorans and DMSS-1, ³⁴ε was ~27‰ for the former species, but only 8‰ for the latter. The relatively high fractionation by pyruvate-grown D. fructosovorans contrasts with previous studies of sulfate reducing bacteria that oxidize organic substrates completely to CO₂. These microbes generally produce values of ³⁴ε smaller than 19‰ when oxidizing pyruvate to acetate in batch culture (Brüchert, 2004; Sim et al., 2011b). An increased fractionation may be a consequence of simultaneous fermentation and respiration of pyruvate, an effect that has been inferred in cultures of D. vulgaris Hildenborough and other species (Sass et al., 2002; Sim et al., 2013), and to which higher fractionations could be attributed (Sim et al., 2013). Another unexplained difference in fractionation is evident during the growth on malate by D. inopinatus (up to ~40‰) and DMSS-1 (ca. 17‰). Such differences suggest that the magnitude of sulfur isotope fractionations is not a direct function of the organic substrate itself, but depends on the specific pathways by which organisms take up and oxidize organic compounds, transfer the reducing equivalents to sulfate (or instead to an organic compound via fermentation), and generate ATP.

The trend in ³⁴ε vs. csSRR for D. fructosovorans generally resembles the inverse correlations reported by several previous studies (Harrison and Thode, 1958; Kaplan and Rittenberg, 1964; Chambers et al., 1975; Kleikemper et al., 2004; Hoek et al., 2006; Sim et al., 2011a,b; Leavitt et al., 2013). Data from several of these studies are shown in Figure 5a. D. inopinatus data fall within the range observed for other Desulfovibrio species, but there is no clear trend between ³⁴ε and csSRR; this may be in part due to the high fractionations observed during growth on malate. In general, trends for different species are subparallel in ³⁴ε–log₁₀(csSRR) space, have different slopes and intercepts, and may be offset by several orders of magnitude in csSRR. These observations are evidence of the complexity that depends on species-specific responses to substrates and growth conditions.

The three Desulfovibrio species have very different cell sizes despite their grossly similar vibrioid morphology (Figure 1), and generally, higher sulfate reduction rates are observed in the larger cells. To test whether the surface areas or volumes of cells account for some of the observed differences in csSRR between species, we expressed ³⁴ε–log₁₀(sulfate reduction rate) relationships per unit of cell surface area or volume. To estimate the surface areas and volumes, we measured the cell dimensions of D. inopinatus, D. fructosovorans and DMSS-1 grown in this study and those of DvH previously grown in our laboratory (Sim et al., 2013; Table 4). The csSRR values were divided by either surface area or volume, and expressed as “cell area (or volume) normalized sulfate reduction rate” [canSRR or cvnSRR; units of fmol H₂S (μm)⁻² day⁻¹ or fmol H₂S (μm)⁻³ day⁻¹, respectively]. Figures 5b,c show the ³⁴ε data for these species plotted against sulfate reduction rates normalized in this manner. Non-linear least-squares regression of a modified Michaelis-Menten-type equation (see Figure 5 legend) to plots of ³⁴ε against canSRR or cvnSRR has higher coefficients of determination (R²) of 0.55 or 0.56 (respectively), compared to 0.43 for ³⁴ε against csSRR. Thus, normalizing sulfate reduction rates to either cell surface area or cell volume reduces the scatter of the data (Figure 5). The improved fits may arise from the distribution of membrane transport proteins across the cell surface, and from the constraints on the amounts of enzyme in a single cell by the cell volume (Pomeroy et al., 2007). This result underscores similarities in the isotopic signals produced per unit of membrane area or biomass of different cultured sulfate reducing microbes, in spite of their differing cell geometries, growth rates, and capacities to oxidize different electron donors. We hypothesize that the larger cells require more energy (electrons) per cell to synthesize cellular components, and therefore exhibit higher csSRR values. Normalizing the sulfate reduction rate to cell volume eliminates that inherent difference amongst microbial species. Alternatively or additionally, larger cells also have larger surface areas that allow larger fluxes of sulfate in to the cell through the cell membrane.

We recognize that cell sizes almost certainly cannot explain all of the variation between the subparallel ³⁴ε–log₁₀(csSRR) trends, and other factors such as the capacity for complete oxidation of organic substrates, growth temperature, and efficiency of sulfate transport may be important (Detmers et al., 2001; Canfield et al., 2006; Bradley et al., 2015). However, similarities amongst the normalized data for several Desulfovibrio species strongly indicate that biomass and cell sizes influence the observed ³⁴ε in experimental and environmental studies. Ideally, we would compare our results to more of the previously published ³⁴ε values reported for SRB and archaea. However, neither the original descriptions of the SRBs, nor previous isotopic studies of MSR provide detailed cell size measurements that are required for this type of analysis. Rather, they report cell size ranges and do not examine the dependence of the cell size on the growth condition. The broad variations in these ranges translate into a large range of cell volumes and do not allow meaningful comparisons with our data. We encourage future studies of MSR in pure cultures to report cell sizes or biomass as functions of growth condition.

Both D. inopinatus and DMSS-1 had slower growth rates (k) when limited by phosphate (Figure 4b and Table 3). The growth rate data for D. inopinatus are in good agreement with the Monod equation (Equation 7) (Monod, 1949) for a half-saturation constant (K_M) of ~10 μM phosphate (Figure 4b). Growth rates for DMSS-1 are more scattered but also decrease below ~10 μM phosphate.

Phosphate limitation at ~10 μM for these species is similar to observations of phosphate limited growth at <12 μM for D. desulfuricans (Okabe and Characklis, 1992) and ~10 μM for D. alaskensis G20 (Bosak et al., 2016), suggesting that many Desulfovibrio species may use similar mechanisms to acquire phosphate during growth. Cellular yields (Y) decreased by several fold at lower phosphate concentrations (Table 3). Lower cell yields were also observed by Okabe and Characklis (1992) when the phosphate concentration was about 10 μM for D. desulfuricans grown in a chemostat; the C:P ratio of cells started to increase before the cell yield began to decrease. Thus, cells first adapted to moderately low phosphorus concentrations by reducing the cellular requirement for P. When the phosphorus availability decreased even further, cell yields began to decrease as well. Morphological changes observed in the two Desulfovibrio spp. we tested (see below), as well as structural adaptations to phosphate limitation and starvation observed in D. desulfuricans (Weimer et al., 1988) and D. alaskensis G20 (Bosak et al., 2016) are consistent with the higher bulk C:P ratio in phosphate-limited D. desulfuricans (Okabe and Characklis, 1992).

D. inopinatus and DMSS-1 cells elongated up to several hundred percent in the lowest phosphate conditions (Table 4). Cell elongation also occurs in phosphate limited cultures of D. alaskensis G20 (Bosak et al., 2016), albeit the overall effect in this microbe is smaller. Microscopic images of phosphate-limited D. inopinatus (Figure 2) revealed long chain-like cells as well as cells with more “typical” lengths. We attribute this cell morphology to incomplete cell division. Nucleic acids (particularly RNA, Elser et al., 2008; Yao et al., 2015) contain a very large fraction (up to tens of percent) of total cellular phosphorus; minimizing DNA and RNA synthesis and division may therefore enable survival in a competitive environment (see also Appendix). Growing phosphate-limited D. alaskensis G20 also replaces its membrane phospholipids by phosphorus-free lipids, synthesizes carbon-rich storage granules and metabolizes nucleic acids (Bosak et al., 2016). Cell elongation and the formation of intracellular carbon-rich granules may also aid in nutrient acquisition by increasing the surface area-to-volume ratio (Thingstad et al., 2005; Godwin and Cotner, 2015; Bosak et al., 2016). Also expected are decreases in the pools of phosphorus-containing metabolites dissolved in the cytoplasm: these metabolites should consist principally of inorganic phosphate, P_i. Below, we discuss how a diminished cytoplasmic phosphorus content can influence the kinetics of enzymatic processes and influence the correlation between ³⁴ε and csSRR.

Phosphorus-limited D. inopinatus and DMSS-1 cultures grown on lactate did not fractionate sulfur isotopes very much (~5 to 8‰). These ³⁴ε values were generally lower in phosphate-limited than in phosphorus-replete cultures (Figure 4a) and we did not observe a distinct correlation with csSRR. In contrast, ³⁴ε values in DMSS-1 cultures grown in iron- and ammonium-limited media were ~5 and ~1‰ higher relative to the cultures grown in nutrient-replete media (Sim et al., 2012). The lower selectivity of the two phosphate-limited SRB species for sulfur isotopes may be related to any number of physiological changes experienced by cells at low phosphate concentrations: the composition and appearance of cell membranes and envelopes, the accumulation of storage polymers, transport of metals and others (Bosak et al., 2016). Some possibilities that directly influence the sulfate reducing pathway are discussed below.

The fitted half-saturation constant (K_M = 10 μM, Figure 4) appears to coincide with the phosphate concentration at which the isotopic fractionation changes from <10‰ to >10‰ for D. inopinatus. Phosphate-limited DMSS-1 cultures exhibit ³⁴ε–log₁₀(csSRR) values that fall off and plot to the right (higher csSRR) of the trend observed by Sim et al. (2011b) (Figure 6). The large heterogeneity in cell lengths of D. inopinatus at low phosphate concentrations (Table 4 and Figure 2) complicates interpretations of csSRR, but assuming that per-cell enzyme activities scale linearly with either cell volume or surface area, the increases in cell size (up to threefold for DMSS-1, Table 4) related to phosphate depletion cannot explain up to tenfold higher csSRRs. The elevated rates may additionally reflect changes in biochemical kinetics within the cell.

Rees (1973) introduced the reversibility of steps in the respiratory chain as a control on overall sulfur isotope fractionation, and this concept has been invoked or extended to explain features of many experimental datasets (Trudinger and Chambers, 1973; Eckert et al., 2011; Sim et al., 2011a,b; Brunner et al., 2012; Antler et al., 2017). More recently, Wing and Halevy (2014) proposed a quantitative model of dissimilatory sulfate reduction that uses enzyme kinetic and thermodynamic data to calculate ³⁴ε values as a function of csSRR. Their model explicitly calculates reversibilities, and thereby isotopic fractionations, for each enzymatic step in MSR. This approach offers insights into links between phosphorus, the sulfate reducing pathway and sulfur isotope fractionations. Phosphorus-containing metabolites (including the adenosine phosphates and inorganic pyrophosphate (PP_i)) influence the thermodynamic drive of reactions in the respiratory chain of sulfate reducers (Equations A1–A4), and their cytoplasmic concentrations are either assumed or explicitly predicted in their model (Figure 7b). The major conclusion of Wing and Halevy's study is that concentrations of intracellular metabolites dictate the energetic favorability and reversibility (ratio of backwards to forwards fluxes) at each step, and influence the overall sulfate reduction rate and the expression of kinetic and equilibrium isotope effects.

We reconstructed the numerical simulations of Wing and Halevy (2014), and extended them to simulate the changes in intracellular concentrations of orthophosphate (P_i) and other phosphorus-containing metabolites as a function of csSRR. The most salient features of the model are described in the Appendix. Our model results suggest that high csSRRs are associated with lower phosphorus contents within the cytoplasm (Figure 7a). This is a direct consequence of Wing and Halevy's finding that reduction of adenosine-5′-phosphosulfate (APS) to sulfite (Equation A3) is the primary rate-limiting step in MSR under conditions probed by most experiments. Because [PP_i] depends on [P_i]² (according to Equation A6), and because P_i accounts for most phosphorus that is dissolved in the cytoplasm (see discussion in Appendix), a small decrease in total cytoplasmic phosphorus during phosphorus-limited growth and starvation induces a large decrease in the PP_i concentration. Lowered PP_i levels are counterbalanced by an increase in the concentration of APS (because Equation A2, sulfate activation to APS, is always near equilibrium; Sat in Figure A3a), as shown in Figure 7b. The buildup of [APS] associated with lowered [P_i] increases the thermodynamic drive for APS reduction (Equation A3). Intuitively, APS reduction becomes less reversible (Apr in Figure A3a), and therefore isotope fractionation between APS and the instantaneous sulfite product approaches the intrinsic kinetic isotope effect for this enzymatic step (22‰ in the default model of Wing and Halevy). Because the remaining step (sulfite reduction, Equation A4) is downstream of APS reduction and is almost fully reversible (dSiR in Figure A3a), the δ³⁴S value of H₂S is not sensitive to the kinetic isotope effect assumed for dSiR (in agreement with some experimental results, Leavitt et al., 2015, 2016) and is ~22‰ lower than that of APS when csSRR is high (Figure A3b).

Modeled ³⁴ε values asymptote to a limit of ~5 ± 2‰ at high csSRRs, in good agreement with our data from DMSS-1 cultures (Figure A4). Several assumptions built into the model require further examination to determine if they are applicable under nutrient-limited conditions. In particular, this includes the parameterization of the scaling factor u_vivo−vitro (see Appendix). Although the agreement with our data may be circumstantial, the above treatment suggests a plausible mechanistic link between low extracellular phosphate concentrations and small ³⁴ε both in the laboratory and in the environment. How phosphate limitation affects the fractionation of sulfur isotopes during growth on substrates that typically produce large fractionations (e.g., fructose) is an open question. The above analysis predicts a decrease in ³⁴ε and increase in csSRR; a hypothesis that can be experimentally tested.

Studies of the sulfur cycle and the sulfur isotope record may benefit from consideration of the possible effects of phosphate limitation. While phosphorus is unlikely to limit growth of sulfate reducing bacteria in modern anoxic sediments where porewater phosphate concentrations are tens of micromolar or greater (Sundby et al., 1992), some studies hypothesize a more limited delivery flux of phosphate to Precambrian sediments (Bjerrum and Canfield, 2002; Reinhard et al., 2017). The very low sulfur isotope fractionation in sedimentary sulfates and sulfides during most of the Precambrian (Canfield, 1998) are consistent with this hypothesis. Overall, our data emphasize potential contributions of organic substrates and phosphate limitation to the persistently low sulfur isotope fractionations in the Archean and most of the Proterozoic.