Microsensor measurements of hydrogen gas dynamics in cyanobacterial microbial mats

We used a novel amperometric microsensor for measuring hydrogen gas production and consumption at high spatio-temporal resolution in cyanobacterial biofilms and mats dominated by non-heterocystous filamentous cyanobacteria (Microcoleus chtonoplastes and Oscillatoria sp.). The new microsensor is based on the use of an organic electrolyte and a stable internal reference system and can be equipped with a chemical sulfide trap in the measuring tip; it exhibits very stable and sulfide-insensitive measuring signals and a high sensitivity (1.5–5 pA per μmol L-1 H2). Hydrogen gas measurements were done in combination with microsensor measurements of scalar irradiance, O2, pH, and H2S and showed a pronounced H2 accumulation (of up to 8–10% H2 saturation) within the upper mm of cyanobacterial mats after onset of darkness and O2 depletion. The peak concentration of H2 increased with the irradiance level prior to darkening. After an initial build-up over the first 1–2 h in darkness, H2 was depleted over several hours due to eﬄux to the overlaying water, and due to biogeochemical processes in the uppermost oxic layers and the anoxic layers of the mats. Depletion could be prevented by addition of molybdate pointing to sulfate reduction as a major sink for H2. Immediately after onset of illumination, a short burst of presumably photo-produced H2 due to direct biophotolysis was observed in the illuminated but anoxic mat layers. As soon as O2 from photosynthesis started to accumulate, the H2 was consumed rapidly and production ceased. Our data give detailed insights into the microscale distribution and dynamics of H2 in cyanobacterial biofilms and mats, and further support that cyanobacterial H2 production can play a significant role in fueling anaerobic processes like e.g., sulfate reduction or anoxygenic photosynthesis in microbial mats.


Introduction
Molecular hydrogen (H 2 ) is produced during fermentative anaerobic degradation of organic matter (Conrad, 1988). Formation of H 2 as a by-product of nitrogenase activity is also described as a major H 2 producing process (Bothe et al., 2010). In excess of energy and reducing power, photosynthetic bacteria can also photo-produce H 2 (Gest and Kamen, 1949;Warthmann et al., 1992). Hydrogen is a very good energy source that is readily reacting with O 2 (chemically or catalyzed by "Knallgas"-bacteria) or is consumed by anaerobic mineralization processes, e.g., as an electron donor in sulfate reduction and in methanogenesis (Hoehler et al., 2002). Anoxygenic photosynthetic bacteria can also utilize hydrogen as an electron donor (Overmann and Garcia-Pichel, 2000).
Efficient inter-species hydrogen transfer in consortia of microorganisms allows for syntrophic processes, which separately would otherwise be energetically unfavorable (Wolin, 1982;Hoehler et al., 2001). Consequently, only very low H 2 concentrations are detected in most natural environments. Higher H 2 levels can, however, be found in special environments like the digestive tracts of termites (Ebert and Brune, 1997) or in legume nodules harboring N 2 -fixing bacteria (Witty, 1991). Geothermal features can also exhibit high levels of H 2 , and hydrogenotrophs are widespread in many hot springs, where H 2 metabolism can be predominant (Spear et al., 2005).
Hydrogen production in cyanobacteria has been known for a long time (Jackson and Ellms, 1896;Benemann and Weare, 1974;Oschchepkov et al., 1974) and has been studied for a large number of strains and in different environments (Lambert and Smith, 1981;Houchins, 1984;Kothari et al., 2012;Otaki et al., 2012). In recent years, cyanobacterial H 2 production has also become a major research topic in connection with the search for new clean energy generating processes (e.g., Lee et al., 2010;Hallenbeck, 2012). Production of H 2 in cyanobacteria is primarily associated with N 2 fixation, where H 2 is a major by-product (Bothe et al., 2010), or due to dark fermentation of storage products accumulated during daytime photosynthesis (Moezelaar et al., 1996;Stal and Moezelaar, 1997). In a survey of different cyanobacterial strains, Kothari et al. (2014) demonstrated that fermentative pathways and bidirectional NADH-linked [Ni-Fe] hydrogenases are of prime importance for H 2 production under dark anoxic conditions in the filamentous non-heterocystous cyanobacteria Microcoleus chtonoplastes and Lyngbya aestuarii that form dense microbial mats in coastal and hypersaline environments; these species exhibited higher production rates and reached much higher steady state H 2 concentrations than many other cyanobacteria. Bidirectional hydrogenases are also involved in light-driven H 2 formation under anoxia via direct biophotolysis, i.e., the light-driven splitting of water (Appel et al., 2000).
Earlier reports of significant H 2 production in cyanobacterial mats were based on gas chromatographic analysis of intact mats (Skyring et al., 1988(Skyring et al., , 1989 and of gas bubbles carefully sampled from the surface of intact hypersaline cyanobacterial mats (Hoehler et al., 2001). The latter study, and more recently Burow et al. (2012) studying a coastal microbial mat also obtained a coarse depth distribution of H 2 production by incubating 2 mm thick slices of mats from different depth horizons below the surface, showing maximal H 2 production in the top 2 mm of the mat during night time. These findings were used to hypothesize that H 2 production by ancient microbial mats and subsequent escape of the H 2 to space was a major mechanism for facilitating the oxidation of the primitive Earth (Hoehler et al., 2001;Jørgensen, 2001). A series of elegant follow up studies combining biogeochemical process measurements with modern molecular tools, have (i) identified filamentous non-heterocystous cyanobacteria as the major H 2 producers in such mats Marschall et al., 2012), (ii) demonstrated cyanobacterial fermentation as the major H 2 producing process Lee et al., 2014), and (iii) demonstrated that sulfate reducing bacteria (SRB) are predominant hydrogenotrophs in cyanobacterial mats . There is thus increasing evidence that fermentative H 2 and organic acid production is a key component in microbial mat biogeochemistry facilitating close interactions between cyanobacteria, anoxygenic phototrophs and heterotrophic bacteria (Otaki et al., 2012;Lee et al., 2014).
Despite an increasing interest in understanding the production and consumption of H 2 in the environment, very few studies have described the fine scale distribution and dynamics of H 2 (Witty, 1991;Ebert and Brune, 1997). Part of the reason has been lack of suitable technology (Hübert et al., 2011). Conventional Clark-type electrochemical H 2 microsensors, which are based on the oxidation of H 2 at a positively charged platinum electrode in an acidic KCl containing electrolyte (Wang et al., 1971;Witty, 1991), often suffer from unstable signals and calibration drift when used in natural systems. This has limited their applicability, especially in environments like sediments and microbial mats, where H 2 measurements were hampered by sulfide interference on the measuring signal.
A sulfide-sensitive amperometric H 2 microsensor based on the use of non-aqueous electrolyte is commercially available (Unisense A/S, Denmark), and a robust version of the sensor has proven useful for quantifying H 2 production in vials with cyanobacterial cultures (Kothari et al., 2012(Kothari et al., , 2014. This sensor has very recently been employed for first microscale H 2 measurement in intertidal microbial mats (Hoffmann et al., 2015) showing pronounced accumulation and efflux of H 2 in darkness driven by cyanobacterial fermentation in the upper mm's of the mat. The microenvironmental dynamics of H 2 in hypersaline water covered mats remain to be studied in more detail, as these mat types often exhibit high sulfide levels causing interference on the commercial H 2 microsensor The sensor design has now been further improved and a sulfide-insensitive H 2 microsensor was recently developed (Nielsen et al., 2015). In the present study we use these microsensors for studying the H 2 microenvironment and its relation to light, O 2 , pH and H 2 S micro gradients in coastal and hypersaline microbial mats. We demonstrate pronounced H 2 dynamics during experimental light-dark shifts, and discuss the role of H 2 for biogeochemical processes in microbial mats.

Experimental Setup
We studied H 2 dynamics in two different microbial mats, both harboring a 1-3 mm thick dark-green surface layer with dense populations of filamentous non-heterocystous cyanobacteria, and anoxygenic Chloroflexi-like phototrophs.

Hypersaline Mat
Dense biofilms of filamentous cyanobacteria were retrieved from the top layer of a hypersaline microbial mat sampled in a salt evaporation pond of Saline de Giraud, Camargue, France. The mat locality and a detailed description of the biogeochemistry and microbial composition of the mat are presented elsewhere (Fourcans et al., 2004;Wieland et al., 2005). Microbial mat samples were transported to our laboratory and were kept in trays with aerated brine at in situ salinity (∼80-100 ppt) under a 12 h light-12 h dark period in a thermostated room at 16 • C. The mat was covered by a 2-3 mm thick deep-green biofilm of motile filamentous cyanobacteria. Microscopic investigations of the biofilm showed a dominance of morphotypes similar to M. chtonoplastes mixed with other motile filaments of Oscillatoria sp. and Spirulina sp. The upper millimeters of the mat remained non-sulfidic due to a conspicuous layer of oxidized iron below the cyanobacterial layer that buffered against sulfide formation in the uppermost mat layer during night-time (see details in Wieland et al., 2005).
Prior to experiments, a small piece of the surface biofilm was transferred to a small 3-4 mm high and 8 mm wide glass beaker with a thin layer of semisolid agar at ∼38-40 • C. During subsequent cooling the biofilm bottom and side adhered to the solidifying agar leaving the upper biofilm surface uncovered. The small beaker was mounted in a flow chamber (Lorenzen et al., 1995) with the biofilm surface flush with a larger agar slab, and aerated brine (90 ppt, 25 • C, pH 8) was constantly circulated over the biofilm surface. The air-saturated brine contained 154 µmol O 2 l −1 according to a table compiled from published empirical solubility equations 1 . Illumination was provided with a fiberoptic halogen lamp equipped with a collimating lens (KL-2500, Schott, Germany), where the irradiance was regulated with a built-in neutral density screen. During long-term incubations the lamp was turned on and off at defined times by an electrical switch with a timer. Irradiance levels at defined lamp settings were determined with a quantum irradiance meter (LI-250, LiCor, USA) equipped with a small spherical irradiance sensor (Walz GmbH, Germany).

Coastal Mat
Coastal microbial mat samples were collected in small acrylate coring tubes from the upper air-exposed yet moist part of a sandbar in Limfjorden near Aggersund, Denmark (57 • 00 02.15N; 9 • 17 12.89E). The mat undergoes irregular cycles of inundation and air exposure depending on prevailing wind directions and consisted of well sorted fine grained sand bound together by a dense 2-3 mm top layer of motile filamentous cyanobacteria (M. chtonoplastes and Oscillatoria sp.) and some green filamentous anoxygenic phototrophs Chloroflexus sp. and exopolymers on top of a black sulfidic layer, which also contained filamentous sulfide oxidizing bacteria (Beggiatoa sp.; Lassen et al., 1992). The mat samples were incubated for 1-2 days in aerated seawater under moderate illumination by halogen lamps (∼100-200 µmol photons m −2 s −1 ) prior to experiments.
During this time, the surface became densely covered by a dense layer of filamentous cyanobacteria (Supplementary Figure S1A). For comparison, we also sampled and investigated permanently submerged sediment samples from the same locality that were predominated by a dense benthic diatom film (Supplementary Figure S1B); these samples were obtained at the same location but from a sandier and less sulfidic sediment that was permanently water covered.
Experiments were conducted with the core samples mounted in an aquarium 1 cm below the surface of continuously aerated seawater (25 ppt, 21-22 • C), which was circulated over the mat by a gentle airstream from a Pasteur pipette. The airsaturated seawater contained 240 µmol O 2 l −1 according to a table compiled from published empirical solubility equations 1 . Illumination was provided by a halogen lamp bulb, where the irradiance was regulated by varying the distance to the mat surface. During long-term incubations, the lamp was turned on and off at defined times by an electrical switch with a timer. Irradiance levels at defined lamp distance were determined with a quantum irradiance meter (LI-250, LiCor, USA) equipped with a small spherical irradiance sensor (Walz GmbH, Germany).
Inhibition of sulfate reduction in the mat was done by incubating a mat sample in aerated and stirred sea water with 2.5 mM sodium molybdate for 6 h prior to measurements. Using published diffusion coefficients (D) of molybdate in water (9.91·10 −6 cm 2 s −1 ; Li and Gregory, 1974) and gel (6.48·10 −6 cm 2 s −1 ; Mason et al., 2005), we estimated the penetration depth of molybdate after t = 6 h of incubation as L = √ 2Dt assuming a one-dimensional diffusion geometry (Berg, 1983). This showed that after 6 h of incubation, molybdate penetrated about 5.3-6.5 mm into the microbial mat ensuring sufficient exposure of SRB's in the complete photic zone as well as several mm's of the underlaying aphotic zone of the mat.
The H 2 microsensor is constructed like a Clark-type O 2 microsensor (Revsbech, 1989) and consists of an outer casing sealed by a thin silicon rubber membrane, and an internal measuring microanode polarized at +0.6 to +1.0 V relative to an internal reference electrode. The casing is filled with an organic electrolyte and this configuration facilitates a stable measurement of H 2 via its oxidation at the measuring anode. The H 2 microsensor is commercially available and further details on the sensor and its calibrations can be obtained from the manufacturer's website 2 . We tested the interference of several compounds, which can pass the silicon membrane of the H 2 microsensor and react at the measuring anode. Compounds like dimethyl sulfide (DMS) and methyl mercaptan can strongly affect the microsensor performance and seem to have a poisoning effect, but levels of these potential interfering agents in microbial mats have been found to be in the lower nM range (Visscher et al., 2003) and should thus not affect our measurements significantly. We found no interference from carbon monoxide, which has been shown to be present in hypersaline microbial mats during daytime (Hoehler et al., 2001) and is a known interfering agent of amperometric H 2 sensors with aquatic electrolytes (Hübert et al., 2011). No sensitivity to light was observed. In the absence of a H 2 S shield, dissolved H 2 S is a major interfering substance and for a given concentration gives rise to a signal of ∼20-30% of the signal measured for the same concentration of H 2 (Nielsen et al., 2015). However, by mounting a thin outer capillary containing ZnCl 2 in propylene carbonate and sealed with a thin silicone membrane, sulfide insensitive H 2 microsensors can be constructed (Nielsen et al., 2015). We did not repeat tests of CO, DMS, and methyl mercaptan interference on the sulfide insensitive H 2 sensors, but such interference will not be larger than on the unshielded sensors.
The new H 2 S insensitive H 2 microsensor (see Nielsen et al., 2015 for details on construction and sensor design) exhibits a linear response from 0 to 100% H 2 . By varying the thickness and diameter of the silicone membrane sealing the microsensor tip as well as the distance from the membrane to the internal measuring anode, we could manufacture H 2 microsensors with various measuring characteristics. Sensors without a H 2 S shield could be constructed with a very fast response time of <0.2 s, but they also exhibited a relatively large stirring sensitivity, which can cause severe measuring artifacts, especially when measuring concentration gradients within a gradient of flow, e.g., in the diffusive boundary layer above sediments and biofilms (Revsbech, 1989;Klimant et al., 1995). Hydrogen gas microsensors with a lower stirring sensitivity can be constructed by using smaller silicone membrane diameters and a longer internal diffusion path, and here the presence of a H 2 S shield contributes to the latter. The sulfide insensitive H 2 microsensors used in this study exhibited a sensitivity of 1.5-5 pA µM −1 H 2 , with a negligible stirring sensitivity and a 90% response time of ∼20-40 s. In all cases, the new H 2 microsensors exhibited low and stable zero currents (1-10 pA) and a temperature sensitivity similar to other amperometric microsensors, i.e., an increase in sensor signal of 2-3% per • C.
We calibrated the sensor in salt water flushed with various defined amounts of H 2 , either by help of a gas-mixing unit or by using commercially available defined mixtures of H 2 and N 2 . Hydrogen data were either expressed as partial pressures (%H 2 saturation) or in molar concentration units. The H 2 concentration in saturated water at experimental salinity and temperature was calculated from tabulated values of H 2 solubility according to Wiesenburg and Guinasso (1979).
The amperometric microsensors were used in connection with a pA-meter (PA2000 and Microsensor Multimeter, Unisense A/S, Denmark), while the pH microsensors were used with a standard calomel reference electrode both connected to a high impedance mV meter (Microsensor Multimeter, Unisense A/S, Denmark). Measuring signals were either recorded on a stripchart recorder (BD-25, Kipp&Zonen, Netherlands) or via an A/D converter (Unisense A/S, Denmark) connected to a PC. Microsensors were mounted in a motorized micromanipulator that was mounted on a heavy stand and was remotely controlled by a PC-interfaced motor controller (Unisense A/S, Denmark). Automated data acquisition and positioning of microsensors was done with commercial software (Profix and Sensor TracePro, Unisense A/S, Denmark). The microsensors were inserted into the biofilm vertically from above in defined steps of 100-200 µm.
The efflux of H 2 from the microbial mats, J(H 2 ) quantified net H 2 production and was calculated from measured concentration gradients using Fick's firs law, J(H 2 ) = D * (dC/dz), where D is the molecular diffusion coefficient of H 2 at experimental temperature and salinity and dC/dz is the linear H 2 concentration gradient in the diffusive boundary layer above the mat. Similar flux calculations, J(O 2 ) were done with O 2 concentration profiles to quantify net photosynthesis in the light and dark O 2 uptake rates, using the molecular diffusion coefficient of O 2 at experimental temperature and salinity. Diffusion coefficients were taken from Broecker and Peng (1974) and corrected for temperature and salinity according to Li and Gregory (1974): D(O 2 ) = 2.05·10 −5 cm 2 s −1 and D(H 2 ) = 3.93·10 −5 cm 2 s −1 at 21 • C and 25 ppt; D(O 2 ) = 2.04·10 −5 cm 2 s −1 and D(H 2 ) = 3.90·10 −5 cm 2 s −1 at 25 • C and 90 ppt.

Microsensor Measurements of Scalar Irradiance
Light penetration in the coastal microbial mat was measured with a scalar irradiance microsensor (Lassen et al., 1992;Kühl et al., 1997;Kühl, 2005) connected to a sensitive fiber-optic spectrometer (QE65000, Ocean Optics, USA) that was interfaced to a PC running dedicated spectral acquisition software (Spectrasuite, Ocean Optics, USA). Mat samples were illuminated vertically from above with a fiber-optic halogen lamp equipped with a collimating lens (KL-2500, Schott, Germany), where the downwelling photon irradiance was regulated with a built-in neutral density screen to 500 µmol photons m −2 s −1 . A scalar irradiance microsensor was mounted in a manually operated micromanipulator (MM33, Märtzhäuser GmbH, Germany) and inserted into the mat at a 45 • angle relative to the vertically incident light. Measurements were corrected for the measuring angle, and depths are given as vertical depth below the mat surface. Data were normalized to the incident downwelling irradiance as measured with the scalar irradiance microsensor positioned in the light path at similar distance as the mat surface but over a black light absorbing well.

Results and Discussion
We measured H 2 dynamics in two different cyanobacterial mats: (i) a hypersaline mat with a pronounced layer of oxidized iron buffering the cyanobacterial top layer against sulfide exposure (Wieland et al., 2005), and (ii) a highly sulfidic coastal cyanobacterial mat (Lassen et al., 1992). Data from the hypersaline mat were measured with H 2 microsensors without a sulfide trap and in the absence of H 2 S, as checked with a H 2 S microsensor (data not shown). Data in the highly sulfidic coastal mat were measured with H 2 microsensors equipped with a chemical sulfide trap in front of the measuring tip (Nielsen et al., 2015).

Hydrogen Production in the Hypersaline Cyanobacterial Mat
When incubated under an irradiance of 800 µmol photons m −2 s −1 for 2.5 h, intense photosynthesis in the dense 2-3 mm thick hypersaline cyanobacterial biofilm lead to hyperoxic conditions reaching 4-5 times air saturation in the upper mm and supersaturating O 2 levels throughout the whole sample, which was contained in a small glass container ( Figure 1A). Upon darkening, O 2 was most rapidly depleted in the region showing highest O 2 production activity in light, and H 2 was first detected in this zone after 15 min. As O 2 became further depleted, H 2 accumulated to higher concentrations and over a wider zone in the biofilm reaching a maximum of 8 µmol H 2 L −1 (∼1.6% H 2 ) at 1 mm depth after 2 h in the dark. Hydrogen was consumed in the lowermost parts of the biofilm sample, which was constrained by the bottom of the small glass incubation container. The apparent migration of the H 2 peak into slightly deeper layers probably reflects a shift in the relative balance between H 2 production, consumption and transport, especially as the mat sample was confined in a small glass vial presenting a diffusion barrier ∼4 mm below the mat surface.
We note that the absolute amount of H 2 produced in the mats after darkening apparently depended on the irradiance level during the previous light incubation. When we increased the irradiance to 1800 µmol photons m −2 s −1 for 2.5 h we again saw a very strong O 2 accumulation in the sample but observed a much higher H 2 production reaching ∼2% H 2 15 min after onset of darkness. Maximal levels of 40-50 µM (8-9% H 2 ) were reached in the upper millimeters of the mat within 30 min after darkening (Figure 2A). These H 2 levels are higher than most other findings  in more permanently submerged hypersaline mats that generally exhibit lower H 2 accumulation than intertidal mats ( Table 1).
We speculate that the apparent enhancement in H 2 production with irradiance reflects a higher accumulation of storage products in the cyanobacteria enhancing subsequent dark fermentation. A similar explanation was proposed by Hoffmann et al. (2015), who measured sustained H 2 production during darkness in intertidal microbial mats kept in a greenhouse for 1.5 years, and with an apparent positive correlation between the solar radiative flux during daytime and the night-time H 2 production in the mats. A rigorous test of this hypothesis would require measurements of cyanobacterial photosynthate accumulation as well as H 2 and fermentation products as a function of light incubation time and may be complicated by the cross-feeding of cyanobacterial fermentation products to other mat members such as SRB and Chloroflexi Lee et al., 2014). Measurements on cyanobacterial cultures using methodology described by Kothari et al. (2014) may thus be more straightforward.
Alignment of H 2 and O 2 microsensor measurements showed that H 2 diffused out of the mat in the dark ( Figure 2B) with an estimated maximal H 2 efflux of ∼23.8 nmol H 2 cm −2 h −1 amounting to about 13% of the diffusive O 2 uptake of 182.2 nmol O 2 cm −2 h −1 in the dark and 2% of the net photosynthetic O 2 production in light of 1420 nmol O 2 cm −2 h −1 . Hoffmann et al. (2015) did similar measurements in different intertidal mats and found that H 2 production in the dark amounted to 0.2-5% of net photosynthesis and 0.4-28% of O 2 respiration.
Long-term experiments with simultaneous O 2 and H 2 measurements over several days (14 h dark: 10 h light) with the microsensor tips positioned 0.8 mm below the mat surface, i.e., in the zone of maximal O 2 production in the light, showed recurrent H 2 production that persisted in the anoxic mat throughout the 14 h dark incubation period (Figure 3).  M. chtonoplastes cultures showed similar long term persistence for >24 h (Kothari et al., 2014). The build-up of H 2 was highest immediately after darkening and then leveled off after 1-2 h. Strong H 2 depletion occurred rapidly after onset of the illumination leading to O 2 accumulation from photosynthesis. Such depletion can be explained by several mechanisms such as (i) O 2 inhibition of H 2 production coupled with diffusive losses, (ii) H 2 consumption with O 2, by e.g., Knallgas bacteria, and/or intermittent anoxygenic photosynthesis. However, our limited experimental data do not allow us to discriminate between the relative importance of these H 2 consuming processes.

Quantification of H 2 production in Lyngbya aestuarii and
In the second light period, the apparently constant O 2 level at this particular depth over many hours indicated formation of a gas bubble in the mat that acted as an O 2 and H 2 reservoir slowing the O 2 depletion and the build-up of H 2 after the subsequent light-dark shift. However, the slower O 2 and H 2 dynamic in subsequent light-dark shifts could also indicate an increasing substrate limitation as the light period may not have been sufficient to restock cyanobacterial storage products.
Overall, the observed H 2 dynamics in the hypersaline mat is very similar to patterns recently observed in cyanobacterial cultures (Kothari et al., 2014) and intertidal microbial mats (Hoffmann et al., 2015). In comparison to other studies of H 2 in hypersaline mats (Table 1), we found much higher H 2 levels after darkening. The reasons for such high H 2 accumulation remain to be studied in detail, but we speculate that the high content of oxidized iron in the upper layers of the Saline des Giraud mat (Wieland et al., 2005) may lead to less sulfate reduction and thus less consumption of H 2 , in contrast to most other hypersaline mat systems that often become highly sulfidic in darkness due to intense sulfate reduction in the top layers. However, the higher H 2 levels may also simply reflect that the small glass incubation vial confined the sample to only a 3-4 mm thick top layer and thus did not allow a diffusive exchange and consumption in deeper more sulfidic mat layers. There is thus clearly a need for more detailed H 2 and H 2 S measurements on deeper mat cores from Saline des Giraud.

Hydrogen Dynamics in Coastal Cyanobacterial Mats
More detailed microenvironmental analyses of H 2 dynamics were done in a sulfidic coastal cyanobacterial mat (Supplementary Figure S1A), whereas measurements in coastal sediment with a surface biofilm of diatoms (Supplementary Figure S1B) showed no accumulation of H 2 (data not shown). Spectral scalar irradiance measurements showed strong light attenuation with depth in the dense 1-2 mm thick top layer of the coastal mat, where distinct throughs in the transmission spectra indicated a high density of cyanobacteria with Chl a and phycobilins, as well as anoxygenic phototrophs with Bchl a and Bchl c (Supplementary Figure S2), with the latter being indicative of the presence of Chloroflexi. The euphotic zone for oxygenic photosynthesis was limited to the uppermost mm of the mat, wherein visible light (PAR, 400-700 nm) was attenuated to ∼0.1 µmol photons m −2 s −1 (Figure 4A). Similar optical characteristics were found in samples from the same site by Lassen et al. (1992) >20 years ago.

Chemical Microenvironment in Light
The chemical conditions in the mat exhibited steep concentration gradients (Figure 4). Under high irradiance of PAR (500 µmol photons m −2 s −1 ), intense photosynthesis led to peak O 2 concentrations of ∼4.5 times air-saturation 0.5 mm below the mat surface. However, O 2 only penetrated to ∼1.2 mm in the light due to intense respiration and re-oxidation of reduced chemical species. Sulfide was produced by SRB in deeper mat layers, where H 2 S levels reached about 0.5 mM at 3 mm depth. Sulfide was re-oxidized by O 2 in a thin zone around 1.2-1.4 mm depth. Strong photosynthesis caused a strong pH increase in the photic zone peaking at pH 10 around 0.5-0.7 mm below the mat surface, i.e., >2 pH units above the overlaying water pH of 7.9. With increasing depth, pH dropped by ∼3 units reaching pH <7 in the sulfide oxidation zone before stabilizing around pH 7 in deeper mat layers. No H 2 was detected in the upper millimeters of the illuminated mat. Measurements in another mat sample from the same habitat showed the same chemical zonations and extremes, albeit with a somewhat deeper O 2 penetration depth in light of ∼1.5 mm and a more heterogeneous distribution of H 2 S in deeper mat layers (Supplementary Figure S3).

Hydrogen Gas Production and Chemical Dynamics after Darkening
The chemical conditions in the coastal mat changed dramatically after darkening (Figure 5). Within 5 min after darkening, O 2 became strongly depleted and only penetrated ∼0.3 mm into the mat, with a further decrease in the O 2 penetration depth to 0.2 mm over the following 85 min. The O 2 and H 2 S concentration profiles were initially separated by a ∼0.7 mm wide zone, wherein H 2 accumulated rapidly after darkening. Peak concentrations were measured 0.5-0.7 mm below the mat surface increasing from ∼13 µM H 2 after 5 min to ∼22 µM H 2 after 45 min dark incubation. Over this time interval, pH in the H 2 production zone decreased to a stable value of pH 7.2-7.8. Produced H 2 diffused both toward the mat surface and toward the sulfidic zone, where it was consumed. After 45 min, H 2 levels in the mat started to decrease, while sulfide levels continued to increase in the upper mat layers. Sulfide started to overlap with the O 2 concentration profile after 90 min. Hydrogen levels in the mat continued to decrease slowly and in a second experiment complete H 2 depletion was only found after about 7 h dark incubation as seen in Figure 6, where data from continuous measurements at 0.6 mm depth are shown. The observed pattern of rapid build-up followed by a slow decline in H 2 concentration follows a similar pattern observed in studies of H 2 evolution in cyanobacterial isolates (Kothari et al., 2012).

Hydrogen Accumulation in the Presence of Molybdate
Additional measurements of H 2 production after onset of darkness were done in another coastal mat sample from the same habitat (Figure 7) that was incubated 8 h under a photon irradiance of 500 µmol photons m −2 s −1 prior to darkening. When incubated in normal seawater, the mat reached maximal concentrations of ∼18 µM H 2 at 0.5-0.7 mm depth within FIGURE 5 | Chemical gradients in a coastal cyanobacterial mat (same sample as in Figure 4) as a function of time after darkening. Prior to darkening, the mat was incubated in aerated seawater under a photon irradiance of 500 µmol photons m −2 s −1 .
FIGURE 6 | Accumulation and depletion of H 2 in a coastal cyanobacterial mat (same sample as in Figure 4) as a function of time after darkening. Measurements were performed 0.6 mm below the mat surface in the zone of maximal H 2 production (see Figure 5). 60 min after darkening, where after H 2 levels in the mat declined gradually to ∼1.5 µM H 2 after 720 min in darkness (Figures 7A,B). Thereafter, the same mat was incubated 6 h under a photon irradiance of 500 µmol photons m −2 s −1 in seawater with 2.5 mM molybdate, an inhibitor of sulfate reduction. Measurements in light at the end of this incubation showed a similar O 2 and pH distribution, whereas H 2 S levels in the mat were much lower below the photic zone than in the absence of molybdate ( Supplementary Figures S3 and S5); interestingly, a slight accumulation of H 2 (reaching 1-1.5 µM) was also observed in deeper mat layers around 3 mm depth after the molybdate treatment, i.e., in the aphotic zone that exhibited high H 2 S levels in the absence of molybdate.
In presence of molybdate, the H 2 accumulation in the mat after darkening was much stronger and H 2 penetrated deeper into the mat (Figure 7C). Within 30 min after darkening, H 2 concentrations reached peak values of >60 µM H 2 0.6-0.8 mm below the mat surface. The H 2 concentrations remained high in the dark incubated mat for about 5 h and showed FIGURE 7 | Accumulation and depletion of H 2 in a coastal cyanobacterial mat as a function of time after darkening (see numbers and color legends in the graph). Prior to darkening, the mat was incubated under a photon irradiance of 500 µmol photons m −2 s −1 in aerated seawater for 8 h without (A,B) and for 6 h with 2.5 mM molybdate (C). Note the use of different concentration ranges in the different panels. much slower H 2 depletion than in the absence of molybdate. Without molybdate, the produced H 2 penetrated to a depth of 1.1-1.3 mm where it became fully depleted in a relatively narrow zone (Figures 7A,B). In the presence of molybdate, H 2 penetrated to a depth of almost 2 mm and the H 2 concentration profile showed a much more gradual depletion with depth ( Figure 7B). Both with and without molybdate, there was no indication of strong H 2 depletion in the uppermost mat layer and the microprofiles showed an efflux of H 2 into the overlaying water. The H 2 efflux was strongly stimulated in the presence of molybdate (Table 1). In the absence of molybdate, the maximal H 2 efflux 60 min after onset of darkness reached 16.3 nmol H 2 cm −2 h −1 amounting to 5.5% of the dark O 2 consumption (296.2 nmol O 2 cm −2 h −1 ) and 1% of the net photosynthetic O 2 production prior to darkening (1500 nmol O 2 cm −2 h −1 ). In the presence of molybdate, the maximal H 2 efflux reached 86.5 nmol H 2 cm −2 h −1 amounting to 29% of the dark O 2 consumption and 6% of the net photosynthesis. Hoffmann et al. (2015) did a similar experiment in intertidal microbial mats showing stimulated H 2 production reaching 4 times higher peak concentrations of H 2 in the presence of molybdate reaching up to ∼20 µM H 2 in the upper mm of the mat (Table 1). However, their measurements with a sulfide sensitive H 2 microsensor showed a H 2 concentration peak on top of an apparent gradually increasing H 2 concentration with depth, which can be interpreted as an sulfide interference on the microsensor signal (see below).
Bulk measurements of the H 2 production of coastal and hypersaline mats (Skyring et al., 1989) and in the upper 2 mm of a coastal cyanobacterial mat  and two hypersaline cyanobacterial mats (Lee et al., 2014) all showed a strong stimulation of H 2 production upon inhibition of sulfate reduction activity by addition of molybdate to the incubation vials. Molecular analyses of the microbial diversity and gene expression in such mats identified SRB as major hydrogenotrophs in the mat along with filamentous anoxygenic phototrophs belonging to the Chloroflexi Lee et al., 2014). In the present study, we did not investigate the distribution and identity of SRB or Chloroflexi or their hydrogenase gene expression in the mat samples, but our H 2 microsensor data strongly support the findings in other cyanobacterial mats identifying SRB as primary hydrogenotrophs in the upper mat layers.
A close spatial co-occurrence of SRB and cyanobacteria has been demonstrated in the upper millimeters of several microbial mat environments (e.g., Baumgartner et al., 2006;Fike et al., 2008), including observation of migratory behavior of motile SRB toward the photic zone (Krekeler et al., 1998). This has been ascribed to aerobic sulfate reduction (Canfield and Des Marais, 1991) and/or a possible aerotaxis combined with aggregation and high O 2 respiration as a survival mechanism for SRB in the photic zone (Cypionka, 2000;Baumgartner et al., 2006). But the presence of SRB within the photic zone also reflects the easy access of SRB to both electron donor and acceptor immediately after onset of darkness and onset of fermentative H 2 production (Lee et al., 2014), and we note that some SRB can also catalyze the oxidation of H 2 , organic substrates and inorganic sulfur species with O 2 as an electron acceptor (Dannenberg et al., 1992). Metabolic flexibility including a versatile H 2 metabolism and chemotaxis of SRB may thus enable them to thrive in the highly variable chemical microenvironment of the photic zone in microbial mats.

Photo-Stimulation of H 2 Production
Simultaneous measurements of O 2 and H 2 concentration in the coastal mat at a depth of 0.6 mm, i.e., within the zone of maximal H 2 production in the dark and photosynthetic O 2 production in the light, showed pronounced dynamics (Figure 8). In the dark, O 2 was depleted completely and H 2 concentrations reached levels of 20-25 µM H 2 after 15-20 min. However, immediately after onset of illumination we observed a burst in H 2 production driving local concentrations up to 30-40 µM H 2 . This burst only lasted for 20-30 s, where after H 2 became rapidly depleted as O 2 from photosynthesis accumulated to super saturating concentration levels in the mat. Similar measurements (data not shown) in depth horizons closer to the mat surface showed a shorter and less intense burst due to more rapid O 2 accumulation and thus faster H 2 depletion, whereas measurement in deeper mat layers showed a less intense build-up of H 2 upon onset of illumination due to strong light limitation; at 0.8 mm depth we observed no photo stimulation of H 2 production. Such intermittent pulses of H 2 upon illumination have been ascribed to direct biophotolysis in cyanobacteria involving a bidirectional Ni-Fe hydrogenase (Appel et al., 2000). While our data give first evidence that such biophotolysis can occur in the uppermost parts of cyanobacterial mats, the process is limited to <1 min after onset of illumination and thus plays a very minor role for the total H 2 production.

Sulfide Interference on H 2 Microsensor Measurements
Our measurements in the hypersaline mat were done under absence of sulfide (checked by H 2 S microsensor measurements) due to a pronounced layer of oxidized iron buffering against accumulation of free sulfide in the photic zone during darkness (Wieland et al., 2005). Under such conditions, the commercially available H 2 microsensor from Unisense performs well and gives accurate quantifications of H 2 concentrations. However, this sensor is also sensitive to hydrogen sulfide giving rise to about 20-30% of the signal for a given H 2 S concentration as compared to the same H 2 concentration, and exposure to high H 2 S levels can also affect sensor calibration (cf. H 2 microsensor manual available at: www.unisense.com/manuals/). Typical H 2 S concentrations in the upper millimeters of hypersaline and coastal mats can reach up to 500 µM just below the photic zone in light or in the uppermost mat layers in darkness (Wieland and Kühl, 2000). A strong gradient of increasing H 2 S concentration with depth would thus be detected in H 2 microsensor measurements giving rise to false H 2 signals of up to >50 µM and typically showing a continuously increasing H 2 level with depth in the mat following the increasing H 2 S concentration. Hoffmann et al. (2015) measured with the sulfide-sensitive H 2 microsensor in three types of intertidal mats. In the upper tidal apparently less sulfidic mat they found a clear peak of H 2 production overlapping with the photic zone, while measurements in the other mats showed that such a H 2 production peak was overlayed by a strong continuously increasing H 2 concentration with depth (cf. Figure 3 in Hoffmann et al., 2015). While the authors did not report on H 2 S measurements in their mats, we suggest that their measurements in deeper mat layers may include H 2 S interference. We have observed similar patterns when using sulfide sensitive H 2 microsensors in the coastal mat from Aggersund (data not shown). Such interference would also affect subsequent rate calculations on measured H 2 concentration profiles, especially in deeper zones. With the new sulfide-insensitive microsensor it is now possible to measure in strongly sulfidic microbial mats without such potential artifacts.

Conclusion
Our measurements demonstrated distinct microscale dynamics of H 2 in hypersaline and coastal microbial mats that are densely populated by filamentous cyanobacteria. We found a pronounced build-up of H 2 in the upper millimeters of such mats upon darkening, while more oxidized coastal sediments with a surface biofilm of diatoms did not show any H 2 accumulation. In general, our results support other recent demonstrations of strong H 2 production in mat-forming cyanobacteria (Kothari et al., 2012(Kothari et al., , 2014 and intact microbial mats (Hoehler et al., 2001;Burow et al., 2012;Hoffmann et al., 2015), where cyanobacterial fermentation of photosynthate in darkness is the major H 2 source. Such H 2 formation upon light-dark shifts in the upper photic zone thus seems inherent in many coastal and hypersaline microbial mats and presents an important, yet intermittent energy source that together with photosynthate fermentation products can fuel anaerobic respiration processes such as sulfate reduction (Lee et al., 2014).
In conclusion, this first application of a new H 2 microsensor (Nielsen et al., 2015) in concert with microsensors for O 2 , pH, H 2 S, and scalar irradiance demonstrated a pronounced potential for H 2 production in the photic zone of microbial mats. Strong intermittent H 2 accumulation (up to 30-40 µM H 2 ) and efflux of H 2 to the overlaying water originated in the uppermost cyanobacterial layers, with the most intense H 2 formation in the depth horizons exhibiting maximal photosynthesis in the light. In the dark, H 2 first accumulates in the uppermost mm of the mat and is then released to the overlaying water and consumed over 6-7 h by anaerobic respiration. Depletion of H 2 within the mat was strongly inhibited by molybdate addition pointing to SRB as major hydrogenotrophs in the mat. Strong H 2 production by biophotolysis was observed in the uppermost anoxic mat layers (0.2-0.8 mm) immediately (for <1 min) after onset of illumination but was quickly inhibited by oxygenic photosynthesis and did not contribute significantly to the H 2 production, which was primarily observed in the dark. The new microsensors allow detailed studies of H 2 dynamics in sulfidic environments at high spatio-temporal resolution. Such studies have until recently been limited to mm-scale measurements of net H 2 production from incubated mat samples using gas chromatography or other bulk phase measurements. Here we have focused on costal and hypersaline cyanobacterial mats, but the new H 2 sensor is also suitable for measurements at higher temperatures (up to 60 • C; Nielsen et al., 2015) and we are currently investigating H 2 dynamics in hot spring microbial mats, where H 2 metabolism often plays a major role (Spear et al., 2005) although the relative importance of H 2 and H 2 S as an energy source remains debated (D'Imperio et al., 2008).