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Research Article
2 January 2014

Polysulfides as Intermediates in the Oxidation of Sulfide to Sulfate by Beggiatoa spp

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Zero-valent sulfur is a key intermediate in the microbial oxidation of sulfide to sulfate. Many sulfide-oxidizing bacteria produce and store large amounts of sulfur intra- or extracellularly. It is still not understood how the stored sulfur is metabolized, as the most stable form of S0 under standard biological conditions, orthorhombic α-sulfur, is most likely inaccessible to bacterial enzymes. Here we analyzed the speciation of sulfur in single cells of living sulfide-oxidizing bacteria via Raman spectroscopy. Our results showed that under various ecological and physiological conditions, all three investigated Beggiatoa strains stored sulfur as a combination of cyclooctasulfur (S8) and inorganic polysulfides (S n 2−). Linear sulfur chains were detected during both the oxidation and reduction of stored sulfur, suggesting that S n 2− species represent a universal pool of bioavailable sulfur. Formation of polysulfides due to the cleavage of sulfur rings could occur biologically by thiol-containing enzymes or chemically by the strong nucleophile HS as Beggiatoa migrates vertically between oxic and sulfidic zones in the environment. Most Beggiatoa spp. thus far studied can oxidize sulfur further to sulfate. Our results suggest that the ratio of produced sulfur and sulfate varies depending on the sulfide flux. Almost all of the sulfide was oxidized directly to sulfate under low-sulfide-flux conditions, whereas only 50% was oxidized to sulfate under high-sulfide-flux conditions leading to S0 deposition. With Raman spectroscopy we could show that sulfate accumulated in Beggiatoa filaments, reaching intracellular concentrations of 0.72 to 1.73 M.


Sulfur storage in bacteria was first documented in the 19th century when Winogradsky (1) proposed the concept of chemolithotrophy from observations of sulfur globules in Beggiatoa filaments. Since then, taxonomically diverse microorganisms have been reported to store zero-valent sulfur either intracellularly (e.g., Beggiatoaceae and Chromatiaceae) or extracellularly (e.g., Ectothiorhodospiraceae, Chlorobiaceae, and Thiobacillus) as an intermediate in the oxidation of reduced sulfur compounds (2). Internal sulfur, which is contained within invaginations of the periplasm, is often surrounded by a protein envelope thought to be purely structural in function (3, 4). External sulfur globules are not enclosed in a membrane; rather, they feature a hydrophilic surface of, e.g., polythionates (5, 6). It has been noted that biological sulfur is characterized by a lower density and different crystalline structure than orthorhombic sulfur (7, 8). However, our understanding of the chemical nature of stored sulfur is still limited by conventional methods of extraction and analysis that can induce artificial changes in the chemistry of biogenic sulfur compounds. In transmission electron micrographs, for example, the presence of sulfur in cells must be inferred from empty vesicles resulting from the dissolution of sulfur in ethanol during the dehydration step in resin embedding (see, e.g., reference 9). Only recently has the novel application of spectroscopic methods to biological samples indicated that the chemical species of stored sulfur may differ across groups of ecologically and physiologically distinct bacteria (see, e.g., references 6, 10, and 11). This has implications for the metabolism of stored sulfur, as enzyme-S0 interactions may be highly specific for the sulfur species utilized.
Sulfur atoms readily catenate to form linear or cyclic molecules and can form bonds with both organic and inorganic end groups. The resulting sulfur compounds are often extremely pH and redox sensitive and thus difficult to measure. The most thermodynamically stable form of elemental sulfur under standard biological conditions is orthorhombic α-sulfur, which consists of puckered S8 rings (12), but more than 180 allotropes have been described (13). Thus far, a variety of species other than cyclooctasulfur have been identified in sulfur-storing bacteria, including inorganic polysulfides (Sn2−), polythionates (O3S—Sn—SO3), and long-chain organosulfanes (R—Sn—R) (6, 11, 14). The solubilities of these S0 species in water are very different, ranging from the nearly insoluble S8 rings to extremely soluble inorganic polysulfides. Sulfur-oxidizing and sulfur-reducing bacteria growing on solid sulfur preferentially take up the charged and soluble fraction, leaving behind the insoluble rings (1517). In Beggiatoa spp., oxidation of elemental sulfur to sulfate is mediated by the cytoplasmic enzyme reverse dissimilatory sulfite reductase (rDSR) (18). It is not yet understood how extracellularly and periplasmically stored S8 rings are accessed by the Dsr system, as this implies transmembrane transport. It has been hypothesized that cyclooctasulfur is either reduced to H2S or taken up via an unknown persulfide (R—S—SH) carrier molecule across the cytoplasmic membrane (19). Furthermore, enzyme active sites probably do not accommodate large molecules of cyclooctasulfur, and recent studies indicate that at least the archaeal sulfur oxygenase reductase from Acidianus ambivalens contains an elongated active-site pocket which can accommodate only linear polysulfides of up to 8 atoms in length (20). The enzymes involved in sulfur utilization in bacteria are still poorly characterized, and therefore mechanisms of sulfur activation and transport remain speculative.
Published studies on the chemistry of stored sulfur frequently report contradictory findings: the same organism appears to contain different sulfur species depending on the culture conditions, sample preparation, and analytical techniques used. Moreover, many of these investigations were performed on dead bacteria at the risk of altering the sulfur chemistry. We tried to resolve some of the conflicting results while investigating the link between sulfur species and enzymatic pathways of sulfur utilization by comparing three strains of Beggiatoa under various growth conditions. In this study, sulfur compounds in single cells of living bacteria were characterized, and their distribution was mapped using confocal Raman microscopy, a technique which relies on the inelastic scattering of light to provide a unique molecular-structural fingerprint of a compound, allowing for its identification. The noninvasive, nondestructive nature of Raman spectroscopy makes it ideal for biological applications and especially for measuring pH- and redox-sensitive sulfur species.



Beggiatoa sp. strain 35Flor was cultivated in agar-stabilized sulfide gradient medium prepared in glass test tubes (1.4 by 14.5 cm) as described by Schwedt et al. (21). Each tube contained an ∼2-cm-high bottom layer of solid agar (1.5% [wt/vol] agar) and an ∼5-cm-high top layer of semisolid agar (0.25% [wt/vol] agar), which allowed Beggiatoa to glide through the medium. Agar layers were prepared by mixing separately autoclaved sulfate-free artificial seawater (24.34 g NaCl, 8.34 g MgCl2 · 6H2O, 0.66 g CaCl2 · 2H2O, and 1.02 g KCl per 1 liter Milli-Q water) and triple-washed agar solutions. The bottom agar was amended with a sterile 1 M sulfide solution to a final concentration of 4, 12, or 16 mM, referred to as low-, medium-, and high-sulfide-flux conditions, respectively. The top agar received additional trace elements, vitamins, minerals (555 mg K2HPO4, 28.72 mg Na2MoO4, 750 mg Na2S2O5, and 29 mg FeCl3 · 6H2O per liter), and HCO3 (2 mM final concentration) as specified by Schwedt et al. (21). Culture tubes were loosely capped, allowing gas exchange with the atmosphere. Oxygen gradients were allowed to develop overnight before inoculation with 100 μl filament suspension approximately 1 cm below the agar surface. Beggiatoa alba strains B15LD and B18LD were grown in freshwater-based agar-stabilized gradient medium (22) with a bottom agar sulfide concentration of 8 mM.
For Raman spectroscopy, 10 to 50 μl of each live bacterial culture was pipetted onto a 20- by 20- by 1-mm glass coverslip coated with 0.1% poly-l-lysine solution (Sigma-Aldrich). Samples were protected with a second coverslip and then sealed with electrical tape to minimize oxygen diffusion and prevent drying. At least 15 different filaments from each time point and culture condition were analyzed, and representative point spectra are shown in Results.

Reference compounds.

The following analytical grade (purity, >99%) compounds were used as references: elemental sulfur (Roth GmbH), sodium sulfate (Sigma-Aldrich), potassium sulfate (Merck), magnesium sulfate (Merck), and calcium sulfate (Applichem). A polysulfide solution was prepared from 5.06 g Na2S · 9H2O and 5.8 g elemental sulfur per 100 ml H2O, with a final pH of 9.5 and sulfide concentration of 210 mM. Spectra for aqueous polysulfides were recorded under anoxic to oxic conditions by transferring a 100-μl drop to a glass coverslip and immediately measuring in live mode, with a point spectrum taken every second. A subsample of the polysulfide solution was dried under a 90:10 N2-CO2 atmosphere for comparison. Crystalline standards, such as sodium sulfate, were measured directly on glass coverslips.

Raman spectroscopy.

All measurements were carried out with an NTEGRA Spectra confocal spectrometer (NT-MDT, Eindhoven, Netherlands) coupled to an inverted Olympus IX71 microscope. The excitation light from a 532-nm solid-state laser was focused on the sample through an Olympus 100× (numerical aperture [NA], 1.3) oil immersion objective. The pinhole aperture was maintained at 55 μm, corresponding to a spatial resolution of 250 to 300 μm. Raman scattered light was dispersed with a 150-line · mm−1 grating and collected by an electron-multiplying charge-coupled device (EMCCD) camera (Andor Technology, Belfast, Northern Ireland) cooled to −70°C. To verify resonance Raman spectra, a 785 nm diode laser coupled to an IR-CCD camera (Andor Technology, Belfast, Northern Ireland) was used, and the pinhole aperture was set at 80 μm. Exposure times varied between 0.2 and 6 s, and the Raman spectra were recorded between 0 and ∼4,500 cm−1 with a spectral resolution of 0.2 cm−1. The laser power at the sample was checked with a laser power meter and did not exceed 1 mW. The instrument was also equipped with a motorized piezostage for x-y scanning. Full-spectrum maps were taken with a maximum 1.2-s dwelling time per point and 0.3-mW laser power.

Data processing.

The software Nova_Px (NT-MDT, Eindhoven, Netherlands) was used for all spectral analyses. Spectra were normalized to an arbitrary unit of intensity on the y axis, maintaining relative peak heights, for easier comparison. Noisy spectra were smoothed using a Gaussian function with a tau value of 3. In order to map specific compounds, relevant peak areas were first background corrected, and images were then overlaid and brightness/contrast adjusted in Adobe Photoshop CS2.
For the Raman quantification of sulfate, spectra of 8 different Beggiatoa sp. 35Flor filaments grown on sulfate-free medium were baseline corrected using a modified average fit curve with a period of 20 (an example is provided in Fig. S1A in the supplemental material). Although peak area is generally used for Raman quantification, peak height can also be used as an accurate measure of intensity (see, e.g., reference 23). In our study, standard curves were generated from the height of the S=O peak at 997 cm−1 in spectra from a 0.2 to 2.0 M dilution series of Na2SO4 in Milli-Q water and resulted in a good fit with an r2 value of ≥0.9309 for the three different measurement conditions used (see Fig. S1B in the supplemental material). The detection limit for sulfate under the applied conditions on the instrument used was ∼25 mM.

Chromatographic sulfate measurements.

Sulfate was analyzed on a 761 Compact ion chromatograph (Metrohm) with a Metrosep A SUPP 5 column. A carbonate buffer, prepared with 3.2 mmol liter−1 Na2CO3 and 1 mmol liter−1 Na2CO3, was used as the eluent. The duration of a run was 14 min, with sulfate eluting at ∼11.5 min. Sodium sulfate standards of 1 to 400 μM and IAPSO standard seawater were used as references.
The intracellular sulfate content of Beggiatoa sp. 35Flor was determined by picking individual filaments with a glass needle and transferring them to 3 ml of 10 mM HCl. Triplicates of samples containing 25, 50, or 100 filaments were freeze-thawed twice to ensure cell lysis. The resulting solution was passed through a 0.45-μm syringe filter into ion chromatography (IC) vials before analysis. Because some liquid adhering to filaments is inevitably transferred with each sample, single liquid droplets from the inoculated medium were also measured in 10 mM HCl. Sulfate concentrations in these control samples were so close to the detection limits of our instrument that single-droplet effects were considered negligible here.
Extracellular sulfate concentrations were determined by extracting pore water from the agar medium with 2-cm-long microrhizones (Rhizosphere Research Products, Wageningen, Netherlands) inserted vertically into the culture tubes. It is possible that differential pressures occurred along the rhizone, resulting in unequal sampling of the upper 2 cm of medium. Parallel cultures with different bottom agar sulfide concentrations (4 mM, 12 mM, or 16 mM) were inoculated at the same time and sampled over a period of 14 days. Uninoculated cultures containing 4 mM sulfide were monitored for the chemical formation of sulfate. For this, one tube from each culture condition was sacrificed at every time point. All liquid samples were diluted in 10 mM HCl and filtered through a 0.2-μm filter before analysis.
For sulfate depth profiles, gradient media with a sulfide concentration in the bottom agar of 4 mM were prepared in triplicate in open-bottom glass tubes (1.4 by 14.5 cm) plugged from below with a tapered rubber stopper. After approximately 2 weeks of incubation, the lower end of each tube was immersed in liquid nitrogen up to a depth of 2 cm, leaving the semisolid top agar nonfrozen, and the rubber stopper was removed. The agar core was then mounted on a micromanipulator, which allowed for sectioning of the top agar in 2-mm steps. Each section of semisolid agar was gently passed through a 0.45-μm syringe filter and diluted in 3 ml of 10 mM HCl for analysis by ion chromatography. Sulfate production was calculated from the change in the measured sulfate concentrations over time and adjusted to the 2-mm-high Beggiatoa mat.

Microsensor measurements.

Gradients of pH and H2S were recorded in triplicate, and the culture tubes were subsequently sectioned for sulfate profiling as described above. The profiles were measured in 250-μm intervals from the agar surface to a depth of 1 cm. Microsensors for pH (pH-10) and H2S (H2S-10) were purchased from Unisense A/S (Aarhus, Denmark) and calibrated as described by Schwedt et al. (21). Total sulfide concentrations were calculated as described by Kühl et al. (24) using a pk1 value of 6.569. Sulfide fluxes were calculated using Fick's first law of diffusion with a diffusion coefficient D of 1.52 × 10−9 m2 s−1 for H2S/HS at 20°C (25). All microsensors were calibrated immediately before use and then after the final measurement to check for a possible drift.


Sulfur species during sulfur oxidation and reduction in a marine Beggiatoa strain.

Beggiatoa was grown in sulfide gradient tubes simulating the environmental conditions in which Beggiatoa can migrate vertically between oxic and anoxic sediment zones. In gradient medium with a low sulfide flux, Beggiatoa sp. 35Flor filaments were present in a dense mat ca. 0.5 cm below the air-agar interface at 16 days after inoculation. Within this mat, the Beggiatoa completely oxidized upward-diffusing sulfide. A peak in the sulfate concentration profile indicated that sulfate was formed as an end product (Fig. 1). The Beggiatoa mat was whitish in color due to intracellular deposition of sulfur globules (Fig. 1 and 2A), as demonstrated by Schwedt et al. using high-performance liquid chromatography of methanol extracts from filaments (21). Using Raman spectroscopy, we could confirm presence of zero-valent sulfur in these inclusions of living Beggiatoa. In Fig. 2B, Raman spectra of these sulfur inclusions are shown together with spectra of two sulfur standards for comparison. The spectrum of the cyclooctasulfur standard was characterized by two strong peaks at 152 and 216 cm−1 corresponding to the bending and stretching modes of the 8-fold ring and a third strong peak at 473 cm−1 corresponding to the vibration of the S—S bond (26). It is important to note that not only the absolute positions of peaks but also their relative heights are significant in a Raman spectrum and that the three peaks of S8 are similar in height. The spectrum of a globule from 5-day-old culture matched that of S8, but spectra from 8- and 16-day cultures exhibited a weakening of the 152- and 216-cm−1 peaks relative to the 473-cm−1 S—S peak, a ratio which is typical of aqueous polysulfides as visible in the Sn2− standard spectrum (see also references 27 and 28). Thus, globules in filaments from a later growth phase were composed of a mixture of polysulfides and cyclooctasulfur, with the ratio of Sn2− to S8 apparently increasing over time.
FIG 1 Example of H2S (○), total sulfide (●), and SO42− (□) profiles in a Beggiatoa sp. 35Flor culture after 16 days of incubation with a low sulfide flux. The Beggiatoa mat was located at a depth of 0.5 to 0.7 cm.
FIG 2 (A) Optical micrograph of a Beggiatoa sp. 35Flor filament with a low sulfide flux. (B) Point spectra of sulfur inclusions measured 5, 8, and 16 days after inoculation using constant 180-μW laser power and 1-s exposure time, compared to sulfur standards (Std). A spectrum of cytochrome c (cyt c)identified in the Beggiatoa membrane region is also shown. (C) Raman map of a filament, showing the 473-cm−1 sulfur peak in yellow and autofluorescence of the cell in green.
The distribution of sulfur within the cells was mapped using the characteristic S—S vibration peak at 473 cm−1 (Fig. 2C). A peak at 757 cm−1, which is depicted in the uppermost spectrum in Fig. 2B, was tentatively attributed to cytochrome c and used in the spectral map as a proxy for the Beggiatoa cell membrane. The 757-cm−1 peak cooccurred with peaks at 979, 1,136, 1,321, and 1,584 cm−1 which are the characteristic resonance Raman bands for cytochrome c at an excitation wavelength of 532 nm (29). The phenomenon of resonance Raman scattering occurs when the excitation frequency is close to an electronic transition of a molecule and thus is specific to the wavelength of the laser used. The fact that the peaks at 757, 979, 1,136, 1,321, and 1,584 cm−1 were not observed under excitation with a 785-nm diode laser (see Fig. S2 in the supplemental material) provides further support for the assignment of these peaks to cytochrome c.
In cultures grown with a high sulfide flux, filaments at the sulfide-oxygen interface accumulated so much sulfur (Fig. 3A and B) that some eventually burst. Presumably to avoid this fate, a subpopulation of filaments migrated downwards and established a second bacterial mat in the anoxic, sulfidic zone. In this subpopulation, a depletion of internal sulfur was observed coinciding with the production of sulfide (21). It has been hypothesized that the anoxic subpopulation of Beggiatoa sp. 35Flor reduces intracellular sulfur with stored polyhydroxyalkanoates (PHAs), which were identified using the nonspecific lipid stain Nile Red (21). Although PHA is a strong Raman scatterer, we were not able to detect this particular carbon storage compound in Beggiatoa sp. 35Flor filaments under any of the investigated growth conditions. The remaining sulfur globules in filaments of this lower Beggiatoa mat were composed of both S8 rings and linear Sn2− species (Fig. 3C), as indicated by the dominant S—S peak at 473 cm−1. Additionally, two weak S8 ring vibration peaks and a 272-cm−1 peak occurred in the S—S—S bending region. The 272-cm−1 peak is characteristic of long-chain (n ≥ 8) polymeric sulfur (30).
FIG 3 (A) Optical micrograph of Beggiatoa sp. 35Flor from the upper mat of a culture grown with a high sulfide flux. (B) Corresponding Raman map showing the occurrence of the 473-cm−1 sulfur peak in yellow/red and autofluorescence of the cell in green. (C) Representative point spectra of sulfur inclusions from upper and lower mat filaments together with polysulfide and cyclooctasulfur standards (Std).
Sulfate, a strong Raman scatterer with a signature peak belonging to the S=O stretch at 997 cm−1 (Fig. 4A), was also identified in Beggiatoa sp. 35Flor filaments but not in the surrounding medium. Porewater sulfate concentrations as measured by ion chromatography were below the ∼25 mM detection limit of the Raman microscope. In Beggiatoa filaments, sulfate appeared to be concentrated in particular regions of the cell, in some cases in association with the sulfur globules (Fig. 4B). The intracellular sulfate was further quantified by Raman spectroscopy using the height of the S=O peak as an indicator of sulfate concentration (see Fig. S1 in the supplemental material). Assuming that sulfate alone contributed to the 997-cm−1 peak, the estimated concentrations of intracellular sulfate were between 0.72 and 1.73 M, with an average of 1.14 M. Ion chromatography of individual, hand-picked filaments confirmed the presence of molar concentrations of sulfate in this marine Beggiatoa strain. We then monitored the production of sulfate from biological sulfide oxidation in the pore water of cultures grown in medium prepared with sulfate-free seawater. The pore water sulfate concentrations increased linearly during the first 2 weeks of incubation in all cultures (Fig. 5). The sulfate production increased proportionally to the sulfide flux, resulting in concentrations of 3.2, 5.1, and 9.9 mM in the 2-cm-thick layer the around the Beggiatoa mat in 14-day-old cultures with initial bottom agar sulfide concentrations of 4, 12, and 16 mM, respectively. No abiotic sulfate production was detected in the uninoculated controls (Fig. 5).
FIG 4 (A) From top to bottom: (i) a background-corrected spectrum of live Beggiatoa sp. 35Flor grown on sulfate-free medium exhibiting a sulfate peak at 997 cm−1 and (ii) sulfate detected in a dried filament in comparison to (iii) an Na2SO4 standard. (B) Raman map of a dried Beggiatoa sp. 35Flor filament, with the 473-cm−1 S—S peak shown in yellow/red, the 997-cm−1 S=O peak of sulfate in blue, and autofluorescence of the cell in green.
FIG 5 Porewater sulfate concentrations in the upper 2 cm of agar medium from cultures grown with different sulfide concentrations in the bottom agar: uninoculated control (■), 4 mM (□), 12 mM (▲), and 16 mM (○).
In order to relate the observed sulfate production to sulfide consumption, we estimated sulfide consumption rates by integrating the sulfide fluxes in Beggiatoa sp. 35Flor cultures measured by Schwedt et al. (21) over the height of the Beggiatoa mat. The resulting sulfide oxidation rates (± the standard deviation) for cultures with 16 mM sulfide were 13.65 ± 2.5 and 8.55 ± 1.8 mol · m−3 · day−1, after 7 and 13 days, respectively (Table 1). These rates were 30 to 40% higher than the corresponding sulfate production rates of 7.9 and 5.8 mol · m−3 · day−1. The sulfide flux in cultures with 4 mM sulfide did not fluctuate significantly over the 13-day period, which can be explained by a continuous downward migration of the Beggiatoa mat, adjusting the oxygen-sulfide interface to the decreasing flux. The average sulfide oxidation rate was 2.88 ± 0.6 mol · m−3 · day−1, and the sulfate production rate was 2.14 mol · m−3 · day−1, showing that at a low sulfide flux, sulfide may be directly oxidized to sulfate.
TABLE 1 Sulfide consumption and sulfate production rates for marine Beggiatoa sp. 35Flor cultures
Sulfide in bottom agar (mM)Time (days)Avg rate (mol · m−3 · day−1) ± SD
Sulfide consumptionSulfate production
16 (high flux)713.65 ± 2.5a7.9
 138.55 ± 1.8a5.8
4 (low flux)72.88 ± 0.6a2.14
 132.88 ± 0.6a2.14
 160.8 ± 0.252.84 ± 0.02
Value calculated from fluxes measured by Schwedt et al. (21).
Although the sulfate production rate remained relatively constant within the first 2 weeks after inoculation, sulfide consumption decreased after this period, implying that the ratio of H2S and S0 oxidation rates varied over time. To better compare the ratio of sulfide oxidation products, sulfate and sulfide fluxes were modeled from the respective chemical profiles (Fig. 1) measured in low-sulfide-flux cultures after 16 days of incubation. The calculated average sulfate production rate was 2.84 ± 0.02 mol · m−3 · day−1, more than four times higher than the sulfide consumption rate of 0.8 ± 0.25 mol · m−3 · day−1. Visual inspection of the filaments at this time point confirmed that most of refractive sulfur globules had disappeared.

Sulfur species in freshwater Beggiatoa strains oxidizing sulfide incompletely to sulfur.

Beggiatoa alba B18LD is a heterotrophic, freshwater strain capable of oxidizing sulfide to elemental sulfur but not to sulfate (31). Instead, the strain can reduce internally stored sulfur to sulfide, presumably by degrading carbon-rich compounds such as PHAs during periods of anoxia (31). In contrast to the case for the autotrophic Beggiatoa sp. 35Flor, polyhydroxybutyrate (PHB) inclusions (Fig. 6A) were easily identified in B. alba B18LD based on their characteristic Raman bands at 906, 1,357, 1,431, 1,733 and ∼2,940 cm−1 (32). After 4 days of growth in gradient cultures with a medium sulfide flux, highly refractive intracellular inclusions were visible under the optical microscope (Fig. 6B). These were identified as sulfur globules composed of a mixture of S8 rings and polysulfides by their characteristic Raman spectra (Fig. 6A). The wave numbers of the S—S vibration varied in different point spectra from 466 to 486 cm−1 (data not shown). Because the vibration frequency of the S—S bond depends on the number of sulfur atoms in a chain (28), the shift in peak position may reflect Sn2− species of different chain lengths (n) coexisting in a polysulfide “pool” within the sulfur globules. The characteristic sharp S=O peak of sulfate at 997 cm−1 was not detected in this freshwater Beggiatoa strain.
FIG 6 (A) From top to bottom: (i) representative point spectra of sulfur inclusions from B. alba B18LD, (ii) extracellular sulfur from B. alba B15LD, (iii) a polysulfide standard, (iv) cytochrome c, and (v) PHB. (B and C) Optical micrographs of strain B18LD (B) and strain B15LD (C) grown with 8 mM sulfide in the bottom agar. (D and E) Raman map of a B15LD filament, with the 2,900-cm−1 C—H peak in pink and the 757-cm−1 cytochrome c resonance peak in green (D), and the corresponding optical image (E). The red square outlines the Raman map.
Beggiatoa alba B15LD appears to be a special case in the family Beggiatoaceae, members of which have thus far all been reported to deposit sulfur intracellularly (33). Surprisingly, no sulfur could be detected inside B. alba B15LD filaments, but instead abundant, refractive spherules, composed of a mixture of S8 rings and polysulfides (Fig. 6A), were found in the medium surrounding the cells (Fig. 6C). Previously, this strain was observed to deposit intracellular sulfur (34), and therefore it is possible that a mutation causing the sulfur globules to be formed extracellularly occurred. In uninoculated controls the abiotically formed sulfur precipitated as crystals rather than globules, suggesting that the extracellular sulfur globules associated with the Beggiatoa alba filaments might be of biological origin. Indeed, these sulfur globules behaved differently from chemically produced crystalline sulfur; for example, exposure to strong laser radiation caused burning of the sulfur globules, suggesting that globules were more amorphous in structure. Full-spectrum Raman mapping of a B15LD filament after 4 days growth in sulfide gradient medium revealed PHB stored as inclusions within the cells and c-type cytochromes associated with the cell membrane (Fig. 6D and E). After 11 days, PHB could no longer be detected in Beggiatoa alba B15LD filaments.


All of the Beggiatoa strains that we studied deposited sulfur globules as a mixture of S8 rings and linear polysulfides, with the relative abundances of the different sulfur species depending on the growth conditions. Although only cyclooctasulfur was detected in Beggiatoa sp. 35Flor cultures at the earliest growth stage, it is possible that minor amounts of polysulfides were also present but could not be distinguished due to the overlap with the strong Raman signal of predominant S8 rings. This observation is consistent with X-ray absorption near edge structure (XANES) spectroscopy of Allochromatium vinosum, which demonstrated a correlation between intracellular concentrations of polysulfides (among other sulfur species) and the bacterial growth phase (17). In cultures of Beggiatoa sp. 35Flor, an increase of the polysulfide fraction under both sulfur-oxidizing and -reducing conditions suggests that Sn2− species represent a universal pool of activated sulfur utilized in both the oxidative and reductive parts of the sulfur cycle. Polysulfides can be derived from cyclic sulfur species by enzymatic cleavage of S—S bonds with membrane-bound thiol groups or glutathione, as postulated for, e.g., Thiobacillus thiooxidans, which oxidizes solid elemental sulfur (35). Alternatively, sulfur rings can be opened chemically by strong nucleophiles such as HS according to the following reaction: n/8 S8 + HS ↔ Sn+12− + H+ (36). Polysulfide anions are also strong nucleophiles, and therefore their formation could have an autocatalytic effect on S—S bond cleavage. It has been shown that the addition of sodium sulfide to cultures of sulfur-reducing bacteria enhances the solubilization and reduction of crystalline sulfur (15, 37). Thus, the cleavage of sulfur rings by HS or Sn2− could provide a fortuitous pool of linear sulfur species, which, in contrast to large sulfur ring structures, are accessible to enzymes involved in sulfur oxidation and sulfur reduction.
In the case of the Beggiatoa strains that we investigated, it could not be determined whether the activation of cyclooctasulfur to polysulfide was biologically or chemically regulated. It is, nonetheless, tempting to speculate that bacteria may harness the chemical reaction instead of expending energy to cleave sulfur rings. In the presence of free sulfide, bacteria might not be able to prevent the decomposition of elemental sulfur to polysulfides. This could explain the presence of polysulfides in both intra- and extracellular sulfur globules of two different Beggiatoa alba strains that cannot further oxidize zero-valent sulfur to sulfate. In the environment, Beggiatoa can migrate vertically between oxic and anoxic sediment zones, thus experiencing dramatic fluctuations in the sulfide supply. Periodic sulfide starvation would require the sulfur-oxidizing Beggiatoa to enzymatically activate S8 rings in order to utilize stored sulfur. Thus, a combination of environmental and physiological factors may influence the species of stored sulfur compounds in these sulfide-oxidizing bacteria.
It is important to note that the speciation of sulfur as a combination of S8 rings and Sn2− described here is discussed only for inclusions in neutrophilic, microaerophilic bacteria. Sufficient concentrations of polysulfides (∼10% of total dissolved HS) exist in solution to support bacterial growth at circumneutral to alkaline pH, but polysulfides undergo hydrolysis in acid (35). The acidophile Acidithiobacillus ferrooxidans produces extracellular globules thought to be composed of a cyclooctasulfur core and surrounded by a hydrophilic layer of polythionates, which are stable only at extremely low pH (6, 38). Although the ultrastructure of sulfur globules in Beggiatoa spp. could not yet be resolved, it is possible that polysulfides contribute to the hydrophilic behavior of the biogenic sulfur, analogous to the function of polythionates in Acidithiobacillus sp. sulfur globules. A proteinaceous membrane which encloses intracellular sulfur of Beggiatoa spp. (39) may protect other cellular components from the highly reactive Sn2− ions. With Raman spectroscopy, we were unable to resolve the exact localization of the sulfur globules within the cell. However, we could observe associations of the sulfur inclusions with the cell membrane, identified by the presence of c-type cytochromes. These observations further confirm previous reports of unusually high cytochrome c contents in many Beggiatoa strains (39, 40), which in some cases causes pink or orange coloration of the bacteria (41, 42).
Our data suggest that elemental sulfur is temporarily stored as an electron reserve, and the activated polysulfide fraction is further oxidized to sulfate or reduced to sulfide by Beggiatoa. Our study of sulfate production by Beggiatoa sp. 35Flor provides further insight into the complete biological pathway of sulfide oxidation. Unlike other sulfur bacteria such as Thiobacillus denitrificans, which oxidize stored sulfur only after other reduced sulfur compounds such as thiosulfate have been depleted (43), Beggiatoa spp. oxidize sulfide and sulfur simultaneously. Our data suggest that sulfide is completely oxidized to sulfate by Beggiatoa when these strains are grown with a low sulfide flux. However, it appears that under high sulfide fluxes, the rate of sulfur oxidation cannot keep up with sulfide oxidation, resulting in up to half of the oxidized sulfide being deposited as sulfur and in some cases causing cells to rupture (21). Conversely, when the sulfide supply has been depleted, the sulfur oxidation rate substantially exceeds the sulfide oxidation rate. This excess sulfate production can be attributed to the oxidation of stored sulfur. To date it has been shown that Beggiatoa spp. store phosphate, nitrate, carbon compounds (such as PHA), and elemental sulfur intracellularly. Our results show that Beggiatoa filaments can also contain molar concentrations of sulfate. Similarly, sulfate has been found inside the related gammaproteobacterium Thioploca in concentrations up to 100 to 1,000 times higher than that in its freshwater environment (44). Storing sulfate offers no apparent advantage to marine bacteria, and it is therefore likely that sulfate is not stored per se but accumulates inside the cell as a product of the sulfate-evolving enzyme complex APS reductase/ATP sulfurylase or another sulfite-oxidizing enzyme, depending on the organism (40, 42, 45). In fact, sulfate appears to be localized in regions around degraded sulfur globules (Fig. 4B) where it is produced. Not much is known about the mechanism of sulfate export in Beggiatoa. A gene for the H+-driven sulfate permease sulP was identified in the genome of Beggiatoa sp. 35Flor (M. Winkel, S. Verena, T. Woyke, H. Schulz-Vogt, M. Richter, and M. Mußmann, unpublished data). The activity of this symporter may be limited by the energy required for ion export or by the low surface-to-volume ratio of these large sulfur bacteria, which restricts substrate-product exchange with the environment.
With our in vivo Raman analyses presented here, we attempted to resolve some of the discrepancies surrounding the discussions of the chemical species composition of bacterial sulfur globules. In the literature there are conflicting reports of sulfur rings (10, 46), polysulfides (11), or polythionates (5, 6) in these inclusions. Based on our results, we hypothesize that even within the same species of bacteria, the speciation of stored sulfur varies under different ecophysiological conditions.


We thank Martina Meyer, Kirsten Imhoff, and Daniela Franzke for technical support and Sandra Havemeyer for kindly providing Beggiatoa alba strains B15LD and B18LD. We are grateful to Heide Schulz-Vogt for helpful discussions and for providing cultures of Beggiatoa sp. 35Flor.
Funding was provided by the International Max Planck Research School of Marine Microbiology, the Max Planck Society, and the Deutsche Forschungsgemeinschaft (through the MARUM Center for Marine Environmental Sciences).

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 80Number 215 January 2014
Pages: 629 - 636
PubMed: 24212585


Received: 23 August 2013
Accepted: 4 November 2013
Published online: 2 January 2014


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Jasmine S. Berg
Department of Biogeochemistry, Max Planck Institute for Marine Microbiology, Bremen, Germany
Anne Schwedt
Department of Microbiology, Max Planck Institute for Marine Microbiology, Bremen, Germany
Present address: Anne Schwedt, Department of Biogeochemistry, Max Planck Institute for Marine Microbiology, Bremen, Germany.
Anne-Christin Kreutzmann
Department of Microbiology, Max Planck Institute for Marine Microbiology, Bremen, Germany
Marcel M. M. Kuypers
Department of Biogeochemistry, Max Planck Institute for Marine Microbiology, Bremen, Germany
Jana Milucka
Department of Biogeochemistry, Max Planck Institute for Marine Microbiology, Bremen, Germany


Address correspondence to Jasmine S. Berg, [email protected].

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