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Research Article
13 January 2015

Rapid Redox Signal Transmission by “Cable Bacteria” beneath a Photosynthetic Biofilm

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ABSTRACT

Recently, long filamentous bacteria, belonging to the family Desulfobulbaceae, were shown to induce electrical currents over long distances in the surface layer of marine sediments. These “cable bacteria” are capable of harvesting electrons from free sulfide in deeper sediment horizons and transferring these electrons along their longitudinal axes to oxygen present near the sediment-water interface. In the present work, we investigated the relationship between cable bacteria and a photosynthetic algal biofilm. In a first experiment, we investigated sediment that hosted both cable bacteria and a photosynthetic biofilm and tested the effect of an imposed diel light-dark cycle by continuously monitoring sulfide at depth. Changes in photosynthesis at the sediment surface had an immediate and repeatable effect on sulfide concentrations at depth, indicating that cable bacteria can rapidly transmit a geochemical effect to centimeters of depth in response to changing conditions at the sediment surface. We also observed a secondary response of the free sulfide at depth manifest on the time scale of hours, suggesting that cable bacteria adjust to a moving oxygen front with a regulatory or a behavioral response, such as motility. Finally, we show that on the time scale of days, the presence of an oxygenic biofilm results in a deeper and more acidic suboxic zone, indicating that a greater oxygen supply can enable cable bacteria to harvest a greater quantity of electrons from marine sediments. Rapid acclimation strategies and highly efficient electron harvesting are likely key advantages of cable bacteria, enabling their success in high sulfide generating coastal sediments.

INTRODUCTION

During the degradation of organic matter, the microbial competition for terminal electron acceptors results in a characteristic vertical redox zonation in marine sediments (1, 2). Typically, oxygen and nitrate are consumed near the sediment surface, whereas sulfate, being a less energetically favorable electron acceptor, becomes utilized only in deeper sediment horizons. In the absence of porewater flushing by waves, currents, and fauna (3) or the downmixing of metal oxy/hydroxides by bioturbation (4), sulfide produced in deeper sediment layers is transported by molecular diffusion toward the sediment surface where it can be reoxidized. Molecular diffusion is a slow transport process that links centimeter-scale distances over time scales of days.
Recently, however, a direct electrical connection between sulfidic and oxic sediment horizons was observed in a marine sediment, whereby electrons harvested from sulfide at depth were shunted directly to oxygen over centimeter distances (5). This novel microbial lifestyle enabling sulfur oxidation by long-distance electron transport can be distinguished by a distinct geochemical fingerprint of microsensor depth profiles (5). Close microscopic examination of sediments exhibiting this geochemical signature revealed that long filamentous bacteria belonging to the family Desulfobulbaceae were associated with the long-distance electron transport (6, 7). These so-called “cable bacteria” have been observed to exceed 8 mm in length and are thought to span the entire suboxic zone (6, 7). Perturbation experiments further suggested that cable bacteria were directly implicated in the long-distance electron transport (6). The occurrence of such long-distance electron transport by cable bacteria challenges conventional ideas about electron transport across redox gradients and thus could have profound implications for our understanding of the microbial ecology and biogeochemistry of marine sediments.
Electrogenic sulfur oxidation by cable bacteria appears to be a highly efficient mechanism for sulfide removal. In sediments with high rates of sulfate reduction, all detectable free sulfide can be removed by this process from a suboxic zone up to about 30 mm (5, 7, 8). Moreover, electrogenic sulfur oxidation appears to be competitively successful over other microbial sulfur oxidation strategies in laboratory experiments when organic-rich coastal sediments are incubated with oxygenated overlying water (5, 9). The geochemical signature of long-distance electron transport and the associated cable bacteria has also been found under natural conditions in a range of coastal habitats with organic-rich sediments (8). A literature survey combined with data mining from 16S rRNA gene sequence archives suggested that electrogenic sulfur oxidation has a cosmopolitan distribution, occurring in a range of organic rich areas, including aquaculture-impacted areas, coastal mud plains, salt marshes, seasonally hypoxic basins, mangrove swamps, and cold seeps. The primary niche for cable bacteria appears to be sediments with high rates of sulfate reduction, together with an oxygenated overlying water column and limited sediment disturbance.
In a previous field study (8), we observed the geochemical fingerprint of electrogenic sulfur oxidation together with high densities of cable bacteria in a drainage channel of an intertidal salt marsh (Rattekaai salt marsh, The Netherlands). The drainage channels are exposed to sunlight and consequently also host diatom-dominated photosynthetic biofilms at the sediment surface. Given that electrogenic sulfur oxidation can co-occur with oxygenic photosynthesis, we hypothesized that benthic photosynthesis could stimulate electrogenic sulfur oxidation by increasing the availability of oxygen, the terminal electron acceptor for electrogenic sulfur oxidation (5). If so, this would imply a novel synergistic interaction in biofilms. In the present work, we performed two experiments to investigate the effects of a photosynthetic biofilm on electrogenic sulfide oxidation by cable bacteria across a range of time scales. We first tested whether deep sediment horizons respond to photosynthetic oxygen production at the sediment-water interface on rapid time-scales (i.e., minutes to hours) and then tested whether the presence of a photosynthetic biofilm affected the depth of sulfide removal over a time scale of days.

MATERIALS AND METHODS

Study site and sediment collection.

Sediment was obtained from Rattekaai, a salt marsh located within the Oosterschelde tidal inlet (51.4391°N, 4.1697°E, The Netherlands) in June 2012 and again in September 2013. Rapid accretion of organic-rich detrital material leads to high rates of sulfate reduction and hence high sulfide generation within the Rattekaai salt marsh sediments (10). Sediment was collected by manual core insertion from a drainage channel within an area dominated by tussocks of Common Cordgrass (Spartina anglica). At low tide, sulfide-rich porewater was observed to seep out from the sediments of the drainage channel (Fig. 1a). These sediments were previously shown to support high rates of sulfide oxidation by long-distance electron transport and were demonstrated to harbor high densities of cable bacteria (8). Repeated visits to the field site between spring and early autumn revealed that the creek sediment typically supported a surface biofilm dominated by diatoms, indicated by a yellow to gold-brown color at the surface and confirmed by microscopic examination.
FIG 1
FIG 1 (a) Photograph of the drainage channel at Rattekaai salt marsh, where sediment with sulfide seepage and a surficial diatom mat are observable. (Courtesy of Jack van de Vossenberg. Reproduced with permission.) (b) Isolated filament of cable bacterial filament, targeted with the CARD-FISH probe DSB706. Scale bar, 10 μm.
In the laboratory, the top 8 cm of sediment was sieved (500-μm mesh), homogenized, and repacked into Plexiglas core liners (4-cm diameter). These cores were subsequently used in two sediment incubation experiments, as described below. The sediment surface was adjusted to be level with the top of the core liner to create continuous water flow over the sediment surface during microsensor profiling. In experiments involving illumination of the sediment surface, the sides of the core liners were covered with opaque tape to ensure that only the sediment surface was illuminated.

Measurements of sediment characteristics.

Sediment porosity was calculated from the water content and density of the solid-phase components. Water content was measured as the difference in weight before and after drying a sediment sample in an oven at 60°C for 48 h, and the solid-phase density was assumed to be 2.65 g cm−3. Loss on ignition was measured as the decrease in dry mass following combustion for 4 h at 450°C. Sediment grain size was measured by laser diffraction on a Malvern Mastersizer 2000.
Anoxygenic phototrophic sulfur oxidizing bacteria have pigments that absorb light in the red and far-red light spectrum (i.e., 650 to 1,100 nm). To assess whether anoxygenic sulfur-oxidizing bacteria (i.e., rather than cable bacteria) could be responsible for sulfide removal in the light, pigments were extracted from sediments in cold 90% acetone with mechanical disruption. A sediment core was cut into successive 0.5-cm slices to a depth of 3 cm, and pigment extraction was performed on each sediment slice. The extract was analyzed with a scanning spectrophotometer (Analytik Jena Specord 210) between 350 and 1,100 nm.
To confirm the presence of the cable bacteria, catalyzed reporter deposition-fluorescence in situ hybridization (CARD-FISH) was performed, using the oligonucleotide probe DSB706 (7). Detailed methods are given in Malkin et al. (8), with the modification that preservation of the sediment samples was accomplished by addition of ethanol to a final concentration of 50%.

Background theory on electrogenic sulfur oxidation.

Electrogenic sulfur oxidation by long-distance electron transport creates a unique geochemical signature in marine sediments which can be assayed using a combination of oxygen, pH, and sulfide microsensor profiling (5). In the absence of photosynthesis, a pH maximum in the oxic zone is generated by the cathodic redox half-reaction of electrogenic sulfur oxidation: O2 + 4e + 4H+ → 2H2O. In the suboxic zone, a pH minimum is generated by an anodic half-reaction of electrogenic sulfur oxidation, such as 0.5H2S + 2H2O → 0.5SO42− + 5H+ + 4e. Thus, electrogenic sulfur oxidation can be identified by the combination of the presence of a pH maximum in the oxic zone, a suboxic space (i.e., where sulfide and oxygen are not detectable), and a pH minimum in proximity to the deeper sulfide horizon. Carbon assimilation associated with photosynthesis also elevates porewater pH within the oxic zone, and this pH elevation can persist for several hours after photosynthesis has stopped (11). Accordingly, it was important to distinguish the surface pH increase due to photosynthesis from that generated by electrogenic sulfur oxidation. To determine the time scales over which photosynthesis affects porewater pH, sediment cores were transferred from the light to the dark, and a series of O2 and pH microsensor depth profiles were recorded, until steady-state concentration profiles were observed.

Microsensor profiling and geochemical rate estimation.

We used high-resolution microsensor profiling to measure changes in O2, pH, and H2S induced by oxygenic photosynthesis and electrogenic sulfur oxidation. For experiment 1, free sulfide (H2S) is reported. For experiment 2, total sulfide (ΣH2S = H2S + HS) was calculated from the H2S measurements and pH values measured at the same depth. Microsensor profiling, including all calibration procedures, was as previously described (8). Calculations for diffusive oxygen uptake, cathodic proton consumption, and current density were likewise conducted as previously described (8). The oxygen penetration depth (OPD) was operationally defined as the sediment depth where the O2 concentration drops below 1 μM, and the sulfide appearance depth was defined as the depth where the H2S concentration first exceeds 1 μM.

Experiment 1: continuous sulfide microprofiling under a diel light/dark cycle.

In a first experiment, we investigated the influence of photosynthetic oxygen production on sulfide concentrations deep in the sediment over the time scale of minutes to hours. The experiment involved two separate pre-experimental steps before measurements were started. In a first pre-experimental step, sediment cores were incubated in the dark in an aquarium containing aerated filtered seawater from the Oosterschelde (salinity 28‰ ± 1‰) in a temperature-controlled room (16 ± ∼1°C). After 20 days of incubation, O2, pH, and H2S microsensor profiling confirmed that the geochemical signal of electrogenic sulfur oxidation was present in the sediment. Examination of the sediment by microscopy also confirmed the presence of a large abundance of long filamentous bacteria, identified with CARD-FISH (probe DSB706) as Desulfobulbaceae filaments (also know as “cable bacteria”; Fig. 1b). In a second pre-experimental step, the sediment cores were exposed to indirect sunlight in an aerated aquarium for 7 days following a natural dark/light regime. During this week, a dense photosynthetic biofilm, dominated by diatoms, grew at the sediment surface, and small gas bubbles were observed during the period when the cores were exposed to light. Oxygen and pH microsensor profiles were recorded during illumination to confirm that photosynthesis was occurring at the sediment surface.
The two pre-experimental steps ensured that the sediment cores at the start of the experiment supported a subsurface filament network of cable bacteria, which was able to perform electrogenic sulfur oxidation, and a surficial diatom-dominated biofilm, which was able to perform oxygenic photosynthesis in the light. The experiment itself consisted of exposing the sediment surface to a 12-h light/dark cycle with an artificial light source for 4 days, while continuously recording changes in pH and free sulfide concentrations within the sediment. Light was provided by cool white fluorescent lighting (∼300 μmol of photons m−2 s−1) mounted above the aquarium on a programmed timer. Free sulfide was monitored continuously by immersing a sulfide microsensor in the suboxic zone of the sediment and programming it to profile the depth interval from 15 to 32 mm at 200-μm-depth increments for the duration of the experiment. Keeping the sensor submerged in the sediment throughout the experiment served to prevent flushing the suboxic zone with oxygenated overlying water during the profiling. pH was continuously monitored near the sediment surface in order to monitor both the cathodic response of electrogenic sulfur oxidation and the photosynthetic activity. To accomplish this, the pH microsensor was mounted on the same micromanipulator as the sulfide microsensor but was adjusted to record profiles in the depth interval from above the sediment water interface down to 8 mm. pH depth profiles were thereby recorded continuously for 4 days at 200-μm-depth increments concurrently with the sulfide depth profiles.

Experiment 2: comparison between sediment cores incubated with or without a photosynthetic biofilm.

In a second experiment, we investigated the influence of a photosynthetic biofilm on the depth of the suboxic zone over a time scale of days, during a period of cable bacteria filament growth. As in the first experiment, a pre-experimental step was performed. Sieved sediment was incubated in the dark, using the same aquarium setup described above. For this second experiment, the preincubation period was shortened to 1 week, after which time electrogenic sulfur oxidation was detected by microsensor profiling (O2, pH, and H2S). Microsensor depth profiles were measured in triplicate cores on this date and are referred to as the “baseline” data.
The experiment began by exposing a subset of the baseline cores (n = 2) to a cool white light-emitting diode (LED) light source (∼300 μE m−2 s−1) programmed on a 12-h light/dark cycle, which served to stimulate the growth of a photosynthetic biofilm. A second subset of the baseline control cores (n = 2) were kept in the dark as controls. All sediment cores were incubated with the same recirculating water, pumped at 20 liters h−1. After a period of 7 days, microsensor profiles (O2, pH, H2S) of the light/dark treatment cores were compared to the dark control cores. All microsensor profiling measurements were made after at least 4 h of darkness to exclude transient effects of photosynthesis on the pH profiles.

RESULTS

Sediment characteristics.

The sieved and homogenized sediment retrieved from Rattekaai salt marsh was a muddy sand composed of 3.8% clay, 35.5% silt, and 61.1% sand (φ = 3.7) and had a sediment porosity of 0.74 in the top centimeters. The sediment was characterized by high organic carbon content, with a mean loss on ignition of 7.5% ± 0.05% (standard deviation [SD]).

Experiment 1: continuous profiles of sulfide under a diel light/dark cycle.

After the pre-experimental incubations, a high-resolution time series of O2 and pH depth profiles confirmed that the sediment used in this experiment supported electrogenic sulfur oxidation, together with benthic photosynthesis (Fig. 2). When the light was turned on, the oxygen tension rapidly increased, reaching more than 250% saturation within 1 h (Fig. 2a). The oxygen penetration depth (OPD) increased from 1.2 mm in the dark to 2.7 mm during illumination. When the light was subsequently switched off, the oxygen penetration depth returned to previous dark values, and the oxygen concentration returned to 100% saturation just above the sediment surface. This transition occurred within 60 min, and the oxygen depth profile did not detectably change after that time (Fig. 2b). Depth profiles of pH were recorded alongside the O2 profiles over the same dark-to-light and light-to-dark transitions (Fig. 2c and d). In the dark, there was a near-surface pH maximum of 8.5, which is attributable to proton consumption associated with the cathodic half-reaction of electrogenic sulfur oxidation (Fig. 2c). During illumination, the near-surface pH maximum increased to >9.0 over a period of about 2 h, attributable to CO2 uptake during oxygenic photosynthesis (128 min; Fig. 2c). When the light was switched off again, the pH decreased back to initial conditions and did not change substantially after about 4 h (267 min; Fig. 2d). Together, these O2 and pH profiles confirmed that during the pre-experimental period, intense electrogenic sulfur oxidation and strong photosynthetic activity were co-occurring within the sediment cores.
FIG 2
FIG 2 Microsensor profiles of oxygen (a and b) and pH (c and d) from the sediment core used in experiment 1. A time series was made for oxygen and pH after a transition from dark to light (a and c), and after a transition from light to dark (b and d). The time after the light transition is indicated in minutes. Note that there is a persistent pH maximum in the oxic zone under dark conditions, which is indicative of electrogenic sulfur oxidation by cable bacteria.
During the course of the light/dark experiment, the depth distributions of pH and H2S were continuously monitored over four light/dark cycles (Fig. 3 and 4). Near-surface sediment pH responded strongly to the consecutive light/dark cycles, and pH profiles indicated that photosynthesis was stimulated in the light and halted in the dark, as expected (Fig. 3a). To aid in visualization, pH at a constant depth (0.4 mm) was plotted in Fig. 4a. During each period of darkness, a shallow subsurface pH maximum remained (Fig. 3), suggesting that the cathodic reactions of electrogenic sulfur oxidation were maintained through subsequent cycles.
FIG 3
FIG 3 Continuous microsensor profiling of an incubated sediment core during 12:12-h light/dark cycles in experiment 1. Black bands at the top of panels indicate periods of darkness. pH at the sediment surface, as a proxy for photosynthesis (a) and concurrent measurements of sulfide concentrations at a depth of 15 to 32 mm (b) are shown.
FIG 4
FIG 4 pH at 0.4 mm (a) and sulfide at 20 mm (b) extracted from the continuous microsensor profiling shown in Fig. 2. There are four repeated phases in sulfide concentrations. (c) One full light/dark cycle, expanded from the dashed box in panel b, with tick marks indicated at 6-h intervals. Identified phases: I, stable sulfide concentrations during illumination; II, a rapid increase in sulfide concentration immediately after illumination is halted, lasting between 1.5 and 2.5 h; III, a decrease in sulfide concentration lasting for the remainder of the dark period; IV, a rapid decrease in sulfide concentration immediately upon illumination, lasting <1 h.
The porewater H2S concentrations between 15 and 32 mm revealed a strong and highly repeatable response to the imposed light/dark cycles (Fig. 3b). Sulfide levels rapidly built up in deeper sediment layers after the light to dark transitions and then gradually diminished over the remainder of the dark period, eventually dropping to low values during illumination. To further illustrate this, the H2S concentration at one depth (20 mm) is plotted in Fig. 4b. Maximum sulfide concentrations were always observed during the dark period and minimum concentrations were observed during the light period.
One light/dark cycle is further magnified to illustrate the four different phases in the concentration change of the sulfide at the fixed depth of 20 mm (Fig. 4c). In the light, sulfide concentrations were initially low (phase I; Fig. 4c). The transition from light to dark resulted in an immediate increase in free sulfide concentration (phase II; Fig. 4c). The time to reach maximum sulfide concentrations was between 1.5 and 2.5 h (mean, 2.1 ± 0.5 h) for the four consecutive light-to-dark transitions. The associated jump in free sulfide concentration at a 20-mm depth (ΔC20 mm) was between 2 and 4 μmol liter−1 and increased slightly over each consecutive cycle. Deeper down in the sediment, the jumps in concentration were progressively smaller, with concentration changes becoming barely detectable below a depth of 30 mm (Fig. 3). After the maximum H2S concentration was reached, a steady decrease occurred over the remaining period of darkness (phase III; Fig. 4c). This gradual concentration decrease eventually compensated ca. 80% of the initial jump in concentration. Upon the subsequent transition from dark to light, there was an immediate and rapid decrease in sulfide concentrations (phase IV; Fig. 4c). Over the four light-to-dark transitions, this sudden decrease occurred within 45 and 52 min. After minimum sulfide concentrations were reached, sulfide concentrations remained stable at all depths for the remaining part of the 12-h light period (i.e., a return to phase I).

Experiment 2: comparison between sediment cores incubated with or without a photosynthetic biofilm.

At the start of experiment 2, the sediment cores exhibited the geochemical signature of electrogenic sulfur oxidation (“baseline” conditions, Fig. 5a). Microscopic examination confirmed that cable bacteria were abundant. Under baseline conditions, the oxygen penetration depth (OPD) was 1.12 ± 0.08 (SD) mm, the diffusive oxygen uptake (DOU) was 0.78 ± 0.05 mmol m−2 h−1, and the current density was 44.5 ± 6.0 mA m−2 (Table 1). The sulfide appearance depth began at 7.7 ± 0.6 mm, and the pH minimum was 6.66 ± 0.04.
FIG 5
FIG 5 Microsensor profiles of oxygen (red), pH (black), and ΣH2S (blue) from sediment used in experiment 2, showing the “baseline” at the start of the experiment (a), the control cores incubated in the dark for 1 week (b), and the cores treated with a diel light/dark cycle to stimulate growth of a photosynthetic film for 1 week (c).
TABLE 1
TABLE 1 Comparison of variables measured in sediment cores at an initial time (“baseline”), after incubation in the dark (control), and in a 12:12-h light/dark cycle that stimulated the growth of a photosynthetic biofilm (treatment)
TreatmentAvg ± SDa
Sulfide depth (mm)pH minimumOPD (mm)DOU (mmol m−2 h−1)Current density (mA m−2)
Baseline7.7 ± 0.66.66 ± 0.041.12 ± 0.080.78 ± 0.0544.5 ± 6.0
Dark (control)9.6 ± 0.9*6.46 ± 0.00*1.08 ± 0.110.87 ± 0.06*27.2 ± 0.0
Light/dark cycle (treatment)14.6 ± 2.0*6.36 ± 0.01*0.78 ± 0.111.26 ± 0.11*52.3 ± 28.8
a
OPD, oxygen penetration depth; DOU, diffusive oxygen uptake. Differences were statistically tested between treatment and control cores with a two-tailed Student t test, and significance at α = 0.05 is indicated with an asterisk (*). Data represent averages of results from two replicate cores for treatments and controls and from three replicate cores for baseline measurements.
Subsets of sediment cores were subsequently incubated for 7 days either in a light/dark cycle (treatment) or in the dark (control). After 7 days of incubation, treatment cores were different from control cores in terms of oxygen consumption, sulfide appearance depth, and pH minimum in the suboxic zone (Table 1; Fig. 5b and c). Diffusive oxygen uptake in the dark was significantly higher in the treatment cores (treatment, 1.27 ± 0.11 mmol m−2 h−1; control, 0.87 ± 0.06; two-tailed Student t test = −4.647, P = 0.043). The oxygen penetration depth in the dark was concomitantly shallower in the treatment cores over the control cores, although the difference was not statistically significant (treatment, 0.78 ± 0.11 mm; control, 1.08 ± 0.11; two-tailed Student t test = 2.828, P = 0.1056). The pH at depth was significantly more acidic in the treatment cores (treatment, 6.36 ± 0.01; control, 6.46 ± 0.00; two-tailed Student t test = 21, P = 0.0023), and the sulfide horizon was deeper in the treatment cores (treatment, 14.6 ± 2.0 mm; control, 9.6 ± 0.9; 2-tailed Student t test = −3.28, P = 0.082). There were no significant differences between treatments in terms of the cathodic proton consumption rate or the current density (a quantity which directly scales with the cathodic proton consumption rate).

DISCUSSION

Rapid response of deep porewater sulfide to photosynthesis mediated by cable bacteria.

Our results demonstrate rapid changes (i.e., on the time scale of minutes) in the pore-water sulfide concentrations of deeper sediment horizons in response to light/dark oscillations. We attribute this deep sulfide response to changes in near-surface oxygen distribution induced by oxygenic photosynthesis. The onset of illumination caused a rapid (<10-min) stimulation of photosynthesis and a concomitant rapid decrease in sulfide concentrations at all measured depths (i.e., between 15 and 32 mm). When the light was switched off, photosynthesis stopped, and the sulfide concentrations at depth rapidly increased. These changes in sulfide were detectable within the first sequence of microprofiling, which took place within 8 min of the light-dark transition. The sulfide response was also depth dependent, with the greatest response in sulfide occurring toward the shallowest depths measured (∼20 mm). Within the measured zone, the sulfide appears suddenly and synchronously at all depths, as indicated by the isoclines of equal sulfide concentrations. This suggests that upon the transition from light to dark, sulfide is locally accumulating throughout the zone from depths of 15 and 30 mm, rather than being transported from below by diffusion. Ruling out two alternative explanations, we interpret these rapid changes in deep sulfide concentration as evidence that electrogenic sulfur oxidation by cable bacteria is stimulated by increased oxygen availability.
First, the possibility that the observed changes in sulfide concentrations were due to sulfide consumption resulting from anoxygenic photosynthesis was excluded. Although in cyanobacterium-dominated microbial mats, anoxygenic photosynthesis by purple and green sulfur bacteria can drive sulfide oxidation using deep penetrating red and infrared light energy, this is not a plausible explanation for the observed changes in sulfide below a depth of 20 mm in our experiment. Light penetration in cohesive sediments, such as those used in our study, is typically <0.5 mm (12). However, even if we consider an extremely deep light penetration depth (e.g., 8.9 mm as documented for far-red and infrared irradiance in a coarse grained sandy sediment [13]), this is still not deep enough to account for the observed sulfide removal between 15 and 32 mm. Furthermore, we did not detect bacterial chlorophyll pigments capable of light absorption in the red to infrared wavelengths in our sediment slices (data not shown), reinforcing that anoxygenic photosynthesis was not responsible for sulfide removal in our experiments. Hence, we conclude that the response of sulfide at depth was not a direct response to the light conditions per se but to a changing oxygen availability within the surface sediment.
Second, we ruled out that the observed changes in sulfide concentrations at depth could be explained by diffusion of redox active compounds linking the production of O2 by photosynthesis to an oxidative consumption of H2S. The diffusion time scale in the vertical dimension can be approximated as t = δ2/2D, where δ is the distance to the oxic zone, and D is the molecular diffusion coefficient of a soluble redox-active compound. Assuming a conservative value of 2 × 10−5 cm2 s−1 for D, diffusion across the suboxic depth to the sediment surface would be on the order of 15 to 63 h. In contrast, the observed changes in sulfide concentration after a transition from dark to light or back occurred on the order of minutes. This very rapid response affirms that a direct biogeochemical connection existed between the distant sediment layers.
A similar rapid response of deep pore water sulfide to changes in oxygen at the sediment surface has previously been documented in a laboratory experiment and gave rise to the hypothesis that surface and deep sediment horizons are connected by electrical currents (5). In this experiment, sediment from Aarhus Harbor was incubated under similar conditions as in first preincubation step of experiment 1 (i.e., incubation in the dark with oxygenated overlying water). This incubation was continued until the geochemical fingerprint of electrogenic sulfur oxidation became established and, subsequently, the sediment water interface was exposed to alternating oxygen levels. Oxygen supply to the overlying water was manipulated by alternately switching between bubbling with oxygenated air and bubbling with a nitrogen-carbon dioxide gas mix. Nielsen et al. (5) observed rapid and repeatable changes in the sulfide concentrations at depth in response to changes in oxygenation of the overlying water. The sulfide appearance depth, also defined by Nielsen et al. (5) as the 1 μM isocline, oscillated up and down. During anoxia, this sulfide horizon migrated ∼4 mm upward in ∼4 h, while during oxic conditions this horizon migrated back downward over the same distance and in about the same time (5). The sulfide oscillations that were observed here were similar, and we likewise attribute them to long-distance electron transfer mediated by cable bacteria (5, 6). Overall, our results substantially strengthen the idea that the biogeochemistry of deeper sediment horizons can rapidly respond to changes in the composition of the overlying water. Moreover, we found that such rapid responses can be induced naturally by the process of photosynthesis and, hence, could be widespread in illuminated coastal environments. Whether cable bacteria provide a reciprocal benefit to diatom biofilms by removing toxic sulfide from the surficial sediment layer remains to be seen.

Response of sulfide to changing oxygen availability: time scale of minutes.

The changes in the sulfide concentration at depth showed a regular cyclical pattern in response to the light/dark cycle (Fig. 4). In this section, we focus on the response of the deep porewater sulfide that occurred on rapid time scales (i.e., minutes). When the light was switched off, the oxic zone rapidly narrowed, and within 8 min the sulfide concentration at depth began to increase (phase II; Fig. 4c). During this phase, we calculated the sulfide production rate as H2SProd = φ·ΔCt, where Δt (∼2 h) is the time interval over which the sulfide increases, ΔC (∼7 μmol liter−1) is the increase in estimated total sulfide (ΣH2S = H2S + HS) concentration, and φ = 0.74 is the sediment porosity. Using ΣH2S estimates for a 20-mm depth, the resulting sulfide generation rate was 2.6 ± 1.2 (SD) nmol cm−3 h−1, where the mean and SD were calculated from the four transitions from light to dark. This value is comparable to the deep sulfate reduction rates previously made at the field site (∼3.0 nmol cm−3 h−1 [10]) and falls within the 2.5 to 7.9 nmol cm−3 h−1 reported from other coastal sediments (14). Together, these findings suggest that sulfide consumption in the deeper part of the suboxic zone is strongly decreased, or even completely arrested, upon transition from light to dark.
We suggest that these rapid changes in the spatial distribution of sulfide in response to changes in light can be best understood by considering the positioning of the cable bacteria in the sediment matrix, relative to a moving oxygen front, and we advance a simple conceptual model to explain these observations (Fig. 6). Invoking a simple electrical analogy, we imagine the long-distance electron transport to occur by the transfer of electrons over a series of redox active sites along the longitudinal axis of the cable bacteria (Fig. 6; discussed in detail in the supplemental material). These redox sites attain an elevated redox potential—Ehcath—in the oxic zone of the sediment, where cells perform the cathodic half-reaction (O2 + 4e + 4H+ → 2H2O) and electrons are removed from the redox sites (i.e., the site occupancy is low). Similarly, the redox sites attain a negative redox potential—Ehan—in the sulfidic zone where cells are performing an anodic half-reaction (0.5H2S + 2H2O → 0.5SO42− + 4e + 5H+) and electrons begin to occupy the redox sites (i.e., the site occupancy is high). If the chain of redox sites has an Ohmic conductivity σ, the electron flux can be expressed as:
FIG 6
FIG 6 Conceptual model explaining the temporal response of sulfide concentrations to photosynthetic oxygen production at the sediment surface. (a) Electrogenic sulfur oxidation during the illuminated phase with associated photosynthetic oxygen production. (b) Hypothetical depth profile of the redox potential of redox sites along the longitudinal axis of the filaments during the illuminated phase. (c) Electrogenic sulfur oxidation during the dark phase. The oxic zone becomes smaller, while the sulfide horizon moves upward. (d) The change in the depth profile of redox site potential as a result of a change in the oxygen penetration depth. (e) The change in the depth profile of redox site potential as a result of a decrease in the cathodic redox site potential in the oxic zone.
Je=σF(ΔEhΔL)
(1)
where ΔL is the width of the suboxic zone, ΔEh = EhcathEhan is the potential difference along the chain of redox sites that spans the length of the cable bacterium, and F is the Faraday constant. Assuming the electron flux Je remains constant between light and dark conditions (the justification of this assumption is discussed in the supplemental material), equation 1 allows us to investigate the processes that modulate ΔL. A change in the oxygen penetration depth shifts the redox depth profile upward, without an effect on ΔEh and hence leading to no effect on ΔL. However, in our measurements, ΔL decreased from 25.5 to 15.8 mm, indicating that a change in the oxygen penetration depth alone was not responsible for the observed changes in the spatial extent of the suboxic zone. Alternatively, if we allow Ehcath to be modulated by changing O2 levels, then equation 1 predicts that ΔL will change in such a way that the gradient in the redox potential (i.e., ΔEhL) remains constant between light and dark. Hence, the observed decrease in ΔL from 25.5 to 15.8 mm (over the transition from light to dark) would also represent a 40% decrease in ΔEh.
To interpret these calculations in terms of microbial physiology and metabolism, we considered the positioning of the cable bacterial cells relative to the fluctuating oxygen front. In the light, a given number of cells are in the oxic zone and perform cathodic oxygen consumption. When the light is switched off, the oxic zone shrinks, decreasing the number of cells in contact with oxygen. Because fewer cells are available to transfer the same quantity of electrons that arrive from the anodic reactions, the number of redox sites charged with electrons of the remaining cathodic cells increases (i.e., the site occupancy of the cathodic cells increases), resulting in a decreased Ehcath. The associated decrease in ΔEh could explain the observed contraction of the suboxic zone and observed H2S accumulation at depth.
Similarly, when the light is switched on, the oxic zone extends downward, rapidly increasing the number of cells in contact with oxygen. If this increase in the number of cells available to perform cathodic reactions is linked to an increase in Ehcath, this could explain our observation that sulfide concentrations rapidly decreased upon the start of photosynthesis. The important implication in terms of cable bacterium physiology is that the cells in the oxic and the suboxic zones are not differentiated but rather are instantly responsive to changes in the redox field.

Response of sulfide to changing oxygen availability: time scale of hours.

In this section, we consider the conspicuous decline in sulfide concentrations that occurs in the later stage of the dark period (phase III; Fig. 4c). After the transition to darkness, the sulfide concentrations first increased, as discussed above. Based on the conceptual model described above, however, we should expect the sulfide concentrations to remain at this elevated level. Intriguingly, however, after about 2 h of darkness, sulfide concentrations started to decrease and steadily declined over the remainder of the dark period (phase III; Fig. 4c), suggesting that sulfide oxidation was reinitiated. We propose two plausible mechanisms to explain these observations.
Our first proposed mechanism is that cable bacteria are stationary in the sediment (i.e., the filaments retain a fixed position), and individual cells regulate their efficiency of electron transfer to oxygen in response to changes in redox gradients. By this proposed mechanism, when fewer cells are in contact with oxygen, the remaining cells that are operating cathodically must be capable of upregulating their efficiency in transferring electrons to oxygen. In terms of the above conceptual model, this would decrease the site occupancy and increase Ehcath, thus explaining the observed expansion of the suboxic zone and the observed decline in sulfide at depth. The delay in response on the scale of ∼2 h suggests that this could be achieved at a transcriptional level of regulation, although the genetic underpinnings for how such a regulatory control could operate is not clear.
Our second proposed mechanism is that cable bacteria are motile and capable of physically reorienting their position in response to the changing redox gradient. By this proposed mechanism, the cable filaments would move upward in response to the upward movement of the oxic zone, thereby increasing the number of cells performing the cathodic half-reaction. During the transition from light to dark, the oxic zone moved upward by ∼1.2 mm. The time to reach maximum sulfide concentration during the first transition was 1.45 h, increasing to 2.5 h on the fourth transition, from which we derive a first-order approximation of (population) movement between 0.13 and 0.23 μm s−1 (0.48 to 0.83 mm h−1). Members of the family Beggiatoaceae are considered fast gliding filamentous bacteria. At the same temperature as our experiments (16°C), Beggiatoa collected from a temperate habitat were observed to glide at a rate of ∼3.0 μm s−1 (9), or about an order of magnitude faster than our tentative estimate for cable bacteria. Our estimated rate of movement for cable bacteria would therefore be difficult to detect by traditional microscopy but could readily explain the changes in sulfide concentrations observed on time scales of hours.
Motility is a well-known adaptive strategy throughout the bacterial kingdom. As a particularly relevant example, the majority of players that form microbial mats are motile (1518). These microorganisms constantly readjust their depth distribution in response to diel oscillations in light and redox conditions in order to maintain optimal positions for their growth (17). Gliding motility in the family Beggiatoaceae was acquired by horizontal gene transfer from cyanobacteria, and it has been proposed that the close physical proximity of these organisms within microbial mats over evolutionary time scales favored the gene transfer (19, 20). This highlights that strong selective pressures exist across widely divergent taxa for acquiring motility in environments where there are steep and fluctuating chemical gradients (21). In subtidal coastal sediments without photosynthesis, rapid oxygen fluctuations can be induced in the surface sediment by changes in bottom currents and the associated adjustment of the diffusive boundary layer (22). In intertidal sediments, rapid oxygen fluctuations are induced by photosynthesis, as in the experiments described here. We hypothesize that motility in cable bacteria could be an adaptive strategy to adjust the distribution of their cathodic and anodic cells to optimize their energy capture. Note that such an energy taxis would require an energy sensing mechanism coordinated across the length of the entire multicellular cable bacterium.

Response of suboxic zone to a photosynthetic biofilm: time scale of days.

In addition to the changes in sulfide concentration observed on time scales of minutes and hours, the presence of a photosynthetic biofilm also affected the sediment geochemistry on a time scale of days. After 7 days of incubation, sediment exposed to a light/dark cycle (treatment) had a significantly deeper suboxic depth and a significantly more acidic pH in the suboxic zone than sediments grown in the dark (control). Photosynthetic biofilms not only increase oxygen availability in the light, but they also increase respiration in the dark. We found significantly greater oxygen consumption (measured in the dark) in our treatment cores versus control cores, which we attribute predominantly to greater oxygen consumption by the biofilm community (i.e., phototrophs, cable bacteria, and other microbes). However, the overall net effect of the biofilm was enhanced electron harvesting from the suboxic zone. These results suggest that greater oxygen availability confers greater electron harvesting capacity from the sediments by the cable bacteria. In sediments that harbor cable bacteria, the sulfide that fuels the electrogenic sulfur oxidation appears to come from two sources (23). Sulfide is generated by microbial sulfate reduction, giving rise to a source of free sulfide throughout the suboxic zone and deeper sediment layers. In addition, free sulfide may arise from FeS dissolution, which only occurs in the suboxic zone, where the acidity generated by the anodic reaction of electrogenic sulfur oxidation strongly enhances FeS dissolution (23). In this experiment, the additional electrons were likely harvested from both sources. First, free sulfide, arising from sulfate reduction, was harvested to a greater depth, as indicated by the greater suboxic depth. Second, the greater acidity in the suboxic zone confers greater FeS dissolution, providing additional H2S release and hence additional electron harvesting from these depths (23). An ability to harvest electrons in both these ways may be important for the success of cable bacteria growing in highly dynamic environments, such as the intertidal salt marshes investigated here.

Conclusions.

We report here that cable bacteria can affect sulfide concentration below a depth of 15 mm within minutes in response to changes in oxygen supply at the sediment surface. The speed of this response implies an absence of differentiation between the cable bacterial cells positioned in the oxic versus suboxic zones (i.e., between cells operating cathodically versus anodically). Second, we report an additional change in sulfide concentration at depth on the scale of hours, indicating a behavioral or regulatory response of cable bacteria to changes in oxygen supply at the sediment surface. Our results add to the growing body of knowledge about cable bacteria metabolism by showing that these bacteria are capable of withstanding rapid changes in the availability of their electron acceptor (oxygen) and identifying that they may be capable of modulating their electron harvesting capacity based on the availability of oxygen.

ACKNOWLEDGMENTS

We gratefully acknowledge Francesco Cozzoli whose observations inspired this study. We also thank Anton Tramper for assistance in the field and lab, Laurine Burdorf for scientific input, and Jack van de Vossenberg for providing the photograph of the study site in Fig. 1. We thank three anonymous reviewers for their thoughtful comments and suggestions, which considerably improved the manuscript.
This research was financially supported by the Netherlands Organization for Scientific Research (NWO-VIDI grant 864.08.004 to F.J.R.M.), Research Foundation Flanders (FWO-Odysseus grant G.0929.08 to F.J.R.M.), and the European Research Council (ERC starting grant to F.J.R.M.).

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cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 81Number 31 February 2015
Pages: 948 - 956
Editor: G. Voordouw
PubMed: 25416774

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Received: 28 August 2014
Accepted: 17 November 2014
Published online: 13 January 2015

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S. Y. Malkin
Department of Analytical, Environmental, and Geo-chemistry, Vrije Universiteit Brussel, Brussels, Belgium, and Royal Netherlands Institute for Sea Research (NIOZ), Yerseke, The Netherlands
Present address: S. Y. Malkin, Department of Marine Sciences, University of Georgia, Athens, Georgia, USA.
F. J. R. Meysman
Department of Analytical, Environmental, and Geo-chemistry, Vrije Universiteit Brussel, Brussels, Belgium, and Royal Netherlands Institute for Sea Research (NIOZ), Yerseke, The Netherlands

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G. Voordouw
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Address correspondence to S. Y. Malkin, [email protected].

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