Hydrogen Dynamics in Trichodesmium Colonies and Their Potential Role in Mineral Iron Acquisition

N₂-fixing cyanobacteria mediate H₂ fluxes through the opposing processes of H₂ evolution, which is a by-product of the N₂ fixation reaction, and H₂ uptake, which is driven by uptake hydrogenases. Here, we used microelectrodes to characterize H₂ and O₂ dynamics in single natural colonies of the globally important N₂ fixer Trichodesmium collected from the Gulf of Eilat. We observed gradually changing H₂ dynamics over the course of the day, including both net H₂ evolution and net H₂ uptake, as well as large differences in H₂ fluxes between individual colonies. Net H₂ uptake was observed in colonies amended with H₂ in both light and dark. Net H₂ evolution was recorded in the light only, reflecting light-dependent N₂ fixation coupled to H₂ evolution. Both net H₂ evolution and H₂ uptake rates were higher before 2 pm than later in the day. These pronounced H₂ dynamics in the morning coincided with strong net O₂ uptake and the previously reported diel peak in N₂ fixation. Later in the afternoon, when photosynthesis rates determined by O₂ measurements were highest, and N₂ fixation rates decrease according to previous studies, the H₂ dynamics were also less pronounced. Thus, the observed diel variations in H₂ dynamics reflect diel changes in the rates of O₂ consumption and N₂ fixation. Remarkably, the presence of H₂ strongly stimulated the uptake of mineral iron by natural colonies. The magnitude of this effect was dependent on the time of day, with the strongest response in incubations that started before 2 pm, i.e., the period that covered the time of highest uptake hydrogenase activity. Based on these findings, we propose that by providing an electron source for mineral iron reduction in N₂-fixing cells, H₂ may contribute to iron uptake in Trichodesmium colonies.

Introduction

Marine primary productivity is often limited by the availability of dissolved organic or inorganic nitrogen (Moore et al., 2013). Diazotrophic cyanobacteria can access an additional nitrogen source, dinitrogen (N₂) gas, and reduce it to ammonia, thereby making it available for other phytoplankton. As part of the N₂ fixation reaction catalyzed by nitrogenase, an equimolar amount of hydrogen (H₂) is produced:

$${\text{N}}_2+8{\text{e}}^{-}+8{\text{H}}^{+}+16\text{ATP}\to 2{\text{NH}}_3+{\text{H}}_2+16(\text{ADP}+{\text{P}}_i)\left(1\right)$$

Since the reduction of protons (H⁺) consumes both reducing equivalents and ATP, H₂ evolution contributes to the high energy costs related to N₂ fixation. However, all N₂-fixing cyanobacteria analyzed so far have either uptake hydrogenases or bidirectional hydrogenases that allow them to recycle some of this H₂ (Tamagnini et al., 2002). As these hydrogenases can feed electrons from H₂ into the respiratory electron transport chain, the uptake of H₂ provides a mechanism for recycling reducing equivalents.

The cyanobacterium Trichodesmium is one of the major marine N₂ fixers and has been estimated to contribute up to 50% to total nitrogen fixation in some areas (e.g., Karl et al., 1997; Dore et al., 2002). A range of different protective mechanisms has been proposed to shield the O₂-sensitive nitrogenase from photosynthetically evolved O₂ in Trichodesmium. Firstly, spatial separation of N₂ fixation and photosynthesis was suggested for the colony-level (Paerl and Bebout, 1988) as well as the single-cell level (N₂-fixing cells termed diazocytes; Bergman and Carpenter, 1991; Berman-Frank et al., 2001). Additionally, nitrogenase activity was shown to be strictly regulated over the diel cycle by daily synthesis of the nitrogenase pool in the morning and inactivation by post-translational modification in the afternoon, allowing for a relatively narrow peak of activity around midday (Zehr et al., 1993; Berman-Frank et al., 2001). A down-regulation of photosynthesis during this midday peak in N₂ fixation was suggested as a mechanism to protect nitrogenase from O₂ (Berman-Frank et al., 2001). Regarding the H₂ metabolism of Trichodesmium, the genome of Trichodesmium erythraeum IMS101 encodes for the uptake hydrogenase hupSL¹ and previous studies on its H₂ metabolism showed that it can recapture ca. 70% of the evolved H₂ (Wilson et al., 2010, 2012).

In natural systems, Trichodesmium often forms colonies that host a wide range of epibiotic bacteria with diverse functions (Hewson et al., 2009; Hmelo et al., 2012; Lee et al., 2017; Frischkorn et al., 2018). Apart from providing a substrate for bacteria, colonies can induce formation of distinct chemical microenvironments. Early studies suggested anoxic microenvironments in Trichodesmium colonies to facilitate N₂ fixation in colonies (Paerl and Bebout, 1988), potentially including the Knallgas reaction (i.e., the reaction of H₂ with O₂ to form H₂O) as a mechanism to reduce O₂ concentrations (Saino and Hattori, 1982). However, more recent studies showed that during day-time in the light, when photosynthesis is active, O₂ concentrations in colonies can increase to as much as 200% of air saturation, which would impose a considerable challenge to the O₂-sensitive nitrogenase in Trichodesmium (Eichner et al., 2017, 2019). In view of this colony-specific potential impediment to N₂ fixation, it becomes all the more relevant to identify the role of colony formation.

Recent studies have suggested colony formation to facilitate iron acquisition from dust, as Trichodesmium colonies were shown to efficiently capture dust particles, move them to the colony center and enhance iron dissolution from the dust (Rubin et al., 2011). The physiological and biochemical mechanisms of dust particle translocation to the colony center and iron dissolution from dust by Trichodesmium are still under investigation. In a parallel study (Eichner et al., submitted), we examined whether chemical gradients within colonies provide beneficial conditions for iron dissolution in the colony center. Measuring O₂ and pH gradients within Trichodesmium colonies with microsensors, we found that the respiration-induced decreases in O₂ and pH in colonies are not large enough to significantly enhance iron dissolution. Hence, the active centering of dust particles in the colony must have a different reason.

The release of soluble iron from minerals, where it is generally present in the form of Fe(III) oxide or oxyhydroxide, can be facilitated by ligand (L) binding (Eq. 2) and/or the reduction of Fe³⁺ to Fe²⁺ (Eq. 3):

$$\text{Fe}{(\text{OH})}_{3(\text{solid})}+{\text{L}}^{-}+3{\text{H}}^{+}\to {[\text{FeL}]}_{(\text{soluble})}+3{\text{H}}_2\text{O}\left(2\right)$$

$$\text{Fe}{(\text{OH})}_{3(\text{solid})}+{\text{e}}^{-}+3{\text{H}}^{+}\to {\text{Fe}}^{2+}{}_{(\text{soluble})}+3{\text{H}}_2\text{O}\left(3\right)$$

Mineral iron dissolution by Trichodesmium has been suggested to involve ligand-promoted dissolution (Basu et al., in press) and/or reductive dissolution (Rubin et al., 2011). While Trichodesmium has been demonstrated to reduce dissolved iron (Roe and Barbeau, 2014; Lis et al., 2015), direct measurements of mineral iron reduction by this organism are still lacking. Reductive dissolution of mineral iron most likely occurs in the vicinity of the cells in the colony center, where the minerals are concentrated. One possibility is electron transfer to Fe(III) directly on the cell surface, using electrons derived from photosynthesis or respiration, by a yet unidentified mechanism. Alternatively, accumulation of a reductant in the colony microenvironment might enable reductive dissolution. Here, we propose that the uptake of H₂, which is produced during N₂ fixation, might play a previously unrecognized role in supplying reducing power for iron acquisition. Specifically, we hypothesized that H₂ can act as a reductant for mineral-bound Fe(III), which is then liberated from the dust matrix for uptake and assimilation.

To address this hypothesis, we firstly used microelectrodes to characterize H₂ fluxes and their dynamics in relation to O₂ fluxes over the diel cycle in the microenvironment of single colonies collected in the Gulf of Eilat. In a second step, we used this data to analyze the kinetics of H₂ uptake under different light conditions and stages of the diel cycle. Finally, to evaluate the potential of H₂ as a reductant for mineral iron dissolution and acquisition, we investigated whether addition of H₂ affects the uptake of mineral iron by Trichodesmium colonies.

Materials and Methods

Colony Sampling

Trichodesmium colonies were sampled from the Gulf of Eilat/Aqaba in the Red Sea over a period of 2 months from March to May 2018. Colonies were collected with a 200 μm net, either placed statically at ca. 1–2 m depth on a pole extended from the pier (3–4 m bottom depth) for approx. 2 h, or by vertical net tows from 20 m to the surface carried out from a boat at ca. 300 m bottom depth in the Gulf. Puff-shaped colonies were then hand-picked with Pasteur pipettes and washed with trace metal free filtered sea water (cleaned using ion exchange resin Chelex 100). Colonies were collected multiple times during the day and kept in filtered seawater under stable light conditions (ca. 130 μmol photons m⁻² s⁻¹) at 25°C for ca. 1–5 h until measurements were started. Assuming that the diel rhythm of metabolic activity by the colonies is maintained under these conditions, rate measurements over the diel cycle are reported relative to the time of measurement rather than sample collection time.

Microsensor Measurements

Microsensor measurements were performed on single colonies. The colonies were placed in filtered seawater in Petri dishes of approx. 50 ml volume, and held in position with a thin glass needle above a nylon mesh to ensure unperturbed diffusion in all directions. The seawater was slowly mixed during measurements by an air flow over the water surface produced with a pipette connected to an aquarium air pump. Measurements were performed at approx. 25°C and 350–450 μmol photons m⁻² s⁻¹, unless specified otherwise.

Clark-type O₂ microelectrodes with ca. 10 μm tip diameter were made at Max Planck Institute for Marine Microbiology as described earlier (e.g., Kühl and Revsbech, 2000). O₂ sensors were calibrated with seawater bubbled with N₂ gas (0% reading) and with air-equilibrated seawater (21% reading; 212 μmol L⁻¹ O₂ at 23°C and salinity of 40) and corrected for changes in electrode performance over the course of the experiments by assuming that O₂ concentrations in the bulk seawater surrounding the colonies (i.e., outside of the microenvironment affected by cellular O₂ fluxes) was in equilibrium with the room air, which was facilitated by the large surface area of the Petri dish used for measurements. H₂ microelectrodes (Unisense) had a tip size of 50 μm and minimum detection limit below 50 nmol L⁻¹ and were calibrated with solutions of known H₂ concentrations prepared by dilution of H₂-saturated seawater (solubility of 652 μmol L⁻¹ at 26°C and salinity of 40) assuming a linear response of the electrode. Calibrations were performed daily to account for changes in electrode sensitivity over the duration of the study.

For recording depth profiles, microelectrodes were moved toward and through the colonies at 50–100 μm step size with a micromanipulator (VT-80, Micos) driven by a motor (Faulhaber minimotor, SA) while observing the colony at ca. 5× magnification in a stereomicroscope (SMZ1500, Nikon with DMC G5, Panasonic camera). For simultaneous measurements of O₂ and H₂ profiles on the same colony, the two sensors were mounted in a 90° angle (Figure 1A) on two separate micromanipulators. Each colony was photographed for determination of colony dimensions and calculation of volume and surface area assuming spherical geometry.

FIGURE 1

Figure 1. (A) Trichodesmium colony during a microsensor measurement. O₂ sensor and H₂ sensor as well as the glass needle and nylon mesh holding the colony in place are shown. The size of the colony core as used for flux calculations is indicated by the bracket. (B) Example of O₂ and H₂ depth profiles measured simultaneously on the same colony. Measurements were performed in the dark after H₂ addition. Gray shading indicates the approximate area taken by the colony core, the center of the colony is approximately at depth –70 μm. The gray line shows an example of a linear fit to the O₂ gradient at the colony surface. The slope of this line gives ΔC/Δr which is used to calculate the flux of O₂ across the colony surface according to Eq. 4.

Net O₂ and H₂ fluxes in and out of the colonies were calculated from the steady state gradients at the colony surface (Figure 1B) according to Fick’s first law of diffusion:

$$\text{J}=-\text{D}(\Delta \text{C}/\Delta \text{r})\left(4\right)$$

where J represents the interfacial O₂ or H₂ flux, D the diffusion coefficient (2.2 × 10⁻⁹ m² s⁻¹ for O₂ and 4.3 × 10⁻⁹ m² s⁻¹ for H₂ at 25°C and salinity 40; Broecker and Peng, 1974), and ΔC the concentration difference measured over the respective distance, Δr, at the colony surface. Interfacial flux was converted to the volume-normalized rate using estimates of surface area and volume for each colony. The surface of the colony was defined visually based on measured profiles as the depth where gradients in O₂ concentrations were steepest. Note that this was typically the boundary of the relatively dense colony core (as indicated in Figure 1), i.e., excluding single filaments protruding outward from the core area.

For detecting not only net H₂ evolution but also potential net H₂ uptake rates, H₂ profiles were recorded under artificially elevated H₂ concentrations. To this end, H₂ was added by pipetting H₂-saturated seawater into the measurement container, reaching final concentrations from below detection limit up to 13 μmol L⁻¹ (6 ± 12 μmol L⁻¹).

Iron Uptake Measurements

Iron uptake from radioactively labeled ⁵⁵ferrihydrite colloids by natural colonies was determined as described by Basu and Shaked (2018). Briefly, ca. 50 colonies were placed in 5 ml acid-cleaned glass vials and ⁵⁵ferrihydrite was added to a final concentration of 100 nmol L⁻¹. Vials were then amended with H₂-saturated seawater to reach a final H₂ concentration of 45 ± 34 μmol L⁻¹ (18–102 μmol L⁻¹), closed without headspace and incubated at ca. 300 μmol photons m⁻² s⁻¹ and 25°C for 3–12 h. Control samples without added H₂ were prepared for each experiment and H₂ concentrations in all vials were determined at the end of incubations with H₂ microelectrodes. At the end of the incubations, samples were transferred into Ti-EDTA-citrate solution (20 min) to ensure removal of absorbed ⁵⁵ferrihydrite and effective detachment of bacteria, and subsequently filtered on 8 μm polycarbonate filters to retain Trichodesmium biomass. ⁵⁵Fe internalized by Trichodesmium was determined by β-counting of the filters in a Tri-carb 1600 CA liquid scintillation counter (Packard).

Results

H₂ Dynamics Over the Diel Cycle – Relation to O₂ Fluxes

Microsensor measurements revealed a large variability in H₂ and O₂ fluxes between single colonies of Trichodesmium analyzed under similar conditions at the same time of the day (Figure 2). Both net evolution and net uptake of H₂ were observed in the presence of added H₂ (Figures 2A,B). The range of net H₂ and O₂ fluxes was clearly dependent on light and showed a distinct pattern over the diel cycle. Net H₂ uptake was observed in both light and dark, whereas net H₂ evolution was observed only in the light. Both net H₂ evolution and net H₂ uptake were highest during morning and midday. In the afternoon, there was no net H₂ evolution and H₂ uptake rates were relatively low. The peak in net H₂ evolution and uptake in the late morning/midday coincided with a peak in O₂ uptake (respiration) rates (Figures 2C,D) as well as the previously reported daily maximum in N₂ fixation rates in Trichodesmium, ca. 5 h after beginning of the light period (e.g., Berman-Frank et al., 2001; Levitan et al., 2007; Milligan et al., 2007; Kranz et al., 2010; Eichner et al., 2014). Following a transient decrease in O₂ fluxes (Figures 2C,D) after midday, respiration and photosynthesis increased again. Photosynthesis rates reached their daily maximum in the late afternoon, when there was no net H₂ evolution, moderate H₂ uptake and N₂ fixation rates were reported to be low (Figure 2C; Berman-Frank et al., 2001; Levitan et al., 2007; Milligan et al., 2007; Kranz et al., 2010; Eichner et al., 2014).

FIGURE 2

Figure 2. Diel cycle of potential H₂ evolution or uptake rates (A,B) and O₂ evolution or uptake rates (C,D) based on depth profiles measured on Trichodesmium colonies in the light (A,C) and dark (B,D). Data points represent single measurements of H₂ or O₂ profiles. Measurements were performed on a total of 130 and 23 colonies for O₂ and H₂, respectively. X-axis denotes time of measurement. Note that all data shown are net rates, with positive numbers reflecting net evolution of H₂ or O₂ by a specific colony, and negative numbers reflecting net uptake of H₂ or O₂, respectively. The green dotted line represents a typical diel cycle in N₂ fixation (arbitrary units), which was obtained by a third order polynomial fit to an average of N₂ fixation rates measured by Berman-Frank et al. (2001), Milligan et al. (2007), Kranz et al. (2010), and Eichner et al. (2014). N₂ fixation rates were normalized to the diel maximum in the respective study. For laboratory studies, start and end of the 12 h light period were aligned to the times of sunrise (6 am) and sunset (7 pm), respectively, in Eilat at the time of the experiments.

In a separate experiment, O₂ and H₂ concentrations were followed over time at the same position close to the center of a colony during consecutive light and dark phases (Figure 3A). An increase in H₂ concentrations by up to 200 nmol L⁻¹ was observed when the lights were switched on in four out of eight colonies tested (Figure 3B). One colony tested in the morning showed the opposite response to light, i.e., higher H₂ concentrations in the dark than in the light (colony b, Figure 3B). The response to light was strongest around midday, as observed for absolute fluxes of H₂ (Figure 2), and in concert with the previously reported peak in N₂ fixation. Two colonies tested in the afternoon, i.e., outside of the N₂ fixation period, showed no light response in H₂ fluxes, although the same colonies were clearly active as indicated by their strong response in O₂ concentrations to light (colonies g and h, Figures 3B,C). O₂ responses to light showed an opposite diel pattern compared to H₂ responses, with stronger responses in the afternoon than at morning/midday (Figure 3C).

FIGURE 3

Figure 3. (A) Example of time series measurement of H₂ performed on a single Trichodesmium colony. Gray shading indicates dark phases, arrow indicates light response (as plotted in B). (B,C) Light response in H₂ (B) and O₂ (C) concentrations (i.e., difference in concentration in light versus dark) measured at different times of the day. Data points labeled with the same letter in (B) and (C) were measured simultaneously using two microelectrodes (H₂ and O₂) on the same colony. Error bars indicate standard deviation of multiple measurements on the same colony (n ≥ 2, except for colonies j and k). The dotted line represents a typical diel cycle in N₂ fixation (arbitrary units), which was obtained by a third order polynomial fit to N₂ fixation rates measured in previous studies as specified in the legend of Figure 2.

Simultaneous measurements of H₂ and O₂ gradients performed by using two microelectrodes on the same colony showed a positive correlation (R² = 0.67) between net O₂ and H₂ uptake when measured in the afternoon in the dark, with a ratio of ca. 50 mol O₂ per mol H₂ taken up (Figure 4A). In light, when photosynthesis was active, O₂ fluxes were not strongly correlated with H₂ uptake (R² = 0.2; Figure 4B).

FIGURE 4

Figure 4. H₂ and O₂ evolution rates measured simultaneously by using two electrodes on the same Trichodesmium colonies in dark (A) and light (B). All measurements were conducted after 2 pm. Note that all data shown are net rates, with positive numbers reflecting net evolution of H₂ or O₂ by a specific colony, and negative numbers reflecting net uptake of H₂ or O₂, respectively. (A) shows a linear regression (solid line) and 95% confidence interval of the fit (dashed lines).

H₂ Dynamics Over the Diel Cycle – Reaction Kinetics

Aiming for a more systematic understanding of the kinetics of H₂ uptake and evolution in Trichodesmium, we analyzed the dependence of H₂ fluxes across the colony surface on the concentration of added H₂ under different conditions (Figure 5). This approach of measuring H₂ evolution under different H₂ concentrations as well as light conditions over the diel cycle allowed us to explore the regulation of uptake hydrogenase activity on different levels, separating the roles of substrate availability, instantaneous light intensity and the underlying diel rhythm. To evaluate the relative importance of each of these parameters for H₂ uptake, we firstly focused on H₂ fluxes under conditions when we expected no H₂ evolution, i.e., measurements carried out in the dark and in the afternoon, when nitrogenase is presumably inactive (Figures 5A–C). Under these conditions, the rate of H₂ uptake can be simply described as

FIGURE 5

Figure 5. Dependence of net H₂ evolution and uptake by Trichodesmium colonies on bulk H₂ concentration measured at different times of the day [before 2 pm (morning) and after 2 pm (afternoon)] and under different light conditions. Ticked boxes above the plots indicate the dominant H₂ and O₂ producing and consuming processes under each condition, as indicated by microsensor results from this study. (A–C) show linear regression lines (y-axis intercept forced to zero). (D) Solid line shows predicted net H₂ flux, dashed line shows gross H₂ evolution, predicted as indicated in the text. Note that all data shown are net rates with positive numbers reflecting net evolution of H₂ by a specific colony, and negative numbers reflecting net uptake of H₂.

$${\text{H}}_2\text{ uptake}=\text{k}\times [{\text{H}}_2],\left(5\right)$$

where k is a pseudo first order kinetic rate constant with units of h⁻¹, which appears as the slope of the regression lines in Figures 5A–C.

In the afternoon, H₂ uptake was not significantly different between light and dark (F-test, p > 0.05; Figures 5A,B), with k values of 175 and 259 h⁻¹ for dark and light, respectively. On the other hand, comparison of the dark k value determined in the afternoon to that determined in the morning shows that H₂ uptake was strongly affected by the time of the day, with a threefold stimulation in the morning (k = 586 h⁻¹; F-test, p < 0.05; Figure 5C).

To estimate the implications of our findings for H₂ fluxes under natural conditions, we furthermore used our experimental data to predict the average H₂ flux under typical bulk H₂ concentrations in natural systems. Under N₂-fixing conditions, when H₂ evolution (by nitrogenase) and H₂ uptake (by hydrogenase) co-occur (Figure 5D), the net H₂ flux can either be positive (net evolution) or negative (net uptake), depending on the bulk H₂ concentration. The rate of net H₂ evolution under these conditions can be described as the difference between gross H₂ evolution and H₂ uptake, which is, in turn, given by the product of k and [H₂] (cf. Eq. 5):

$$\mathrm{Net}{\text{H}}_2\mathrm{evolution}=\mathrm{Gross}{\text{H}}_2\mathrm{evolution}-\text{k}\times [{\text{H}}_2]\left(6\right)$$

Given that H₂ uptake across the colony surface cannot persist when there is no H₂ present in the bulk seawater, the rate of gross H₂ evolution was estimated from net H₂ evolution rates measured at the lowest bulk H₂ concentrations in our experiments (average net H₂ evolution below 0.5 μmol L⁻¹ bulk H₂). Please note that this concept of “gross H₂ evolution” refers to fluxes across the colony surface, whereas significantly larger fluxes may occur within the colony. Assuming that nitrogenase activity is not dependent on external H₂ concentrations, the gross H₂ evolution rate would be constant across all H₂ concentrations at 0.68 nmol mm⁻³ h⁻¹ (dashed line in Figure 5D). Since light had only a minor effect on k (Figure 5B), we assumed k determined in the morning in dark to be valid also under light conditions. The resulting predicted net H₂ flux is plotted in Figure 5D (solid line). Based on the x-axis intercept of this line, our data suggest net H₂ evolution to occur below a bulk H₂ concentration of ca. 1 μmol L⁻¹, whereas at higher H₂ concentrations, H₂ uptake is dominant.

Response of Mineral Iron Uptake to H₂ Addition

Mineral iron uptake by Trichodesmium colonies in incubations without added H₂ was similar across five experiments conducted at different times of the day (Figure 6), with average uptake rates of 14 ± 3 fmol Fe colony⁻¹ day⁻¹. In the presence of H₂, mineral iron uptake was strongly elevated in three out of five experiments, including both puff-shaped colonies (experiments #2 and #3; Figure 6) and tuft-shaped colonies (experiment #1; Figure 6). Over a whole day-night cycle, iron uptake was increased by more than a factor of 2 in the presence of H₂ (experiment #1, Figure 6). In shorter incubations, the response in uptake to H₂ was dependent on the time of the day. The strongest response was observed in incubations that were started at midday (experiments #2 and 3, Figure 6), and thus included the previously reported maximum in N₂ fixation rates. In incubations lasting from the end of the N₂ fixation period till the evening (experiments #4 and 5, Figure 6), there was no response to H₂.

FIGURE 6

Figure 6. Effect of H₂ on rates of iron uptake from ferrihydrite by natural Trichodesmium colonies measured at different times of the day. The time line below the plot indicates timing of the respective incubations as well as a typical diel cycle in N₂ fixation (dotted line), which was obtained by a third order polynomial fit to N₂ fixation rates measured in previous studies as specified in the legend of Figure 2. Error bars for incubation #4 show standard deviation of two replicate samples. Note incubation #1 continued through the night.

Discussion

H₂ Uptake and Evolution by Individual Colonies

Our microsensor measurements of H₂ and O₂ gradients in single, field-collected Trichodesmium colonies demonstrate significant variability in H₂ metabolism between individual colonies, which resulted in net H₂ fluxes directed either in or out of the colony. Single Trichodesmium colonies thus provide distinct and highly diverse microenvironments with regard to H₂ concentrations. Our experimental approach does not distinguish between the contribution of Trichodesmium and its associated bacteria to the H₂ fluxes. However, the fact that Trichodesmium colonies in our study were always oxic (>100 μmol L⁻¹ O₂) precludes activity of many of the known H₂-consuming taxa, such as anoxygenic phototrophs and sulfate reducing bacteria, and H₂-producers such as fermenting cyanobacteria. It should be noted that gross H₂ fluxes may be significantly larger than the net H₂ fluxes determined in this study, particularly in the confined space of a colony microenvironment with close spatial coupling of sources and sinks.

Previous studies have reported a wide range of H₂ evolution rates by Trichodesmium. Field studies in different locations across the North Atlantic and Caribbean showed H₂ evolution rates ranging between 0.002 and 0.05 nmol colony⁻¹ h⁻¹ (Scranton, 1983, 1984; Scranton et al., 1987). The maximum net H₂ evolution rate per colony in our study (0.08 nmol colony⁻¹ h⁻¹) strongly exceeds these previous field-based estimates. For laboratory cultures grown in single filaments under ca. 50 μmol photons m⁻² s⁻¹, net H₂ evolution rates in the order of 1 nmol (μg chl a)⁻¹h⁻¹ and up to 3 nmol (μg chl a)⁻¹h⁻¹ were reported (Wilson et al., 2010, 2012). Using a chlorophyll a content per colony determined previously in the North Pacific Subtropical Gyre (14 ng colony⁻¹; Eichner et al., 2017) to normalize H₂ fluxes, our maximum estimate also exceeds net H₂ evolution determined in the laboratory by approx. a factor of two (Wilson et al., 2010). Similarly, maximum net H₂ uptake in our study [9 nmol H₂ (μg chl a)⁻¹ h⁻¹] exceeds previous estimates of H₂ uptake for Trichodesmium (estimated as the effect of the hydrogenase inhibitor carbon monoxide on net H₂ evolution; Saino and Hattori, 1982) by approx. a factor of two. Aside from potential effects of experimental conditions among these studies, these comparisons demonstrate that H₂ fluxes in and out of the microenvironment of single colonies can be significantly larger (at their diel maximum) than average H₂ fluxes determined in incubations of several colonies or homogenous cultures.

In contrast to the previous reports of net H₂ evolution, the average across all our measurements in light was a net H₂ uptake rate of 0.01 ± 0.03 nmol colony⁻¹ h⁻¹. While experimental conditions in previous laboratory as well as field studies (such as different light intensities, colony formation and different species) most probably contribute to the differences between ours and previous results, possibly the most important difference was that our measurements were performed under micromolar H₂ concentrations, while in natural waters H₂ concentrations are typically in the low nanomolar range (e.g., Scranton, 1983; Wilson et al., 2013). Assuming a linear relation between net H₂ evolution/uptake and bulk H₂ concentration (Figure 5D), we extrapolated our data to lower H₂ concentrations, and predicted that under bulk H₂ concentrations below ca. 1 μmol L⁻¹, H₂ evolution would exceed H₂ uptake. Under natural H₂ conditions, the net H₂ flux would thus – on average – be directed out of the colony, implying net H₂ evolution in line with previous observations. However, our measurements also demonstrate that substantial variability in H₂ fluxes (as observed within and between previous studies) can even be manifested on the level of single colonies sampled over a short period of time at the same location.

Dependence of H₂ Fluxes on Light and the Diel Cycle

Measuring the steady-state H₂ gradients on the colony surface against a background of added H₂ enabled us to estimate the capacity of Trichodesmium colonies for H₂ uptake under various conditions. Since the H₂ uptake rates were measured under artificially elevated background concentrations of H₂, the rates we report should be considered as potential rates. The capacity for H₂ uptake was not uniform over the diel cycle and dependent on light (Figures 2, 3), indicating that uptake hydrogenase is not only regulated by the availability of its substrate H₂, in line with previous findings (Tamagnini et al., 2002). The fact that both net evolution and net uptake of H₂ were observed reflects the delicate balance between the two highly active, opposing processes of H₂ evolution by nitrogenase and H₂ uptake by uptake hydrogenase. However, considering only conditions where nitrogenase is presumably inactive (afternoon and/or dark) allowed us to separate these process and indicated that light itself had only a minor effect on uptake hydrogenase (Figures 5A–C). Consequently, the light effects on net H₂ fluxes in our study (Figures 2, 3) were most likely driven by the light response of nitrogenase activity, whereas uptake hydrogenase was not directly affected by light. The strong stimulation of dark H₂ uptake in the morning compared to the afternoon (Figure 5C), in turn, indicates that H₂ uptake is regulated over the diel cycle, potentially via an endogenous rhythm. The links between H₂ metabolism, respiratory and photosynthetic electron transport in cyanobacteria make for a complex regulatory system. O₂ fluxes measured at the same time and/or on the same colonies thus provided an important reference for gaining insights into the physiological mechanisms driving the H₂ dynamics we observed in Trichodesmium colonies. In the following paragraph, we discuss these potential physiological mechanisms.

Underlying Physiological Mechanisms

Processes Involved in the Dark

In the dark, O₂ uptake by respiration and H₂ uptake by hydrogenase were correlated (Figure 4A), in line with previously suggested links between these processes (e.g., Appel and Schulz, 1998). Uptake hydrogenases have been reported to be located on either the thylakoid membrane or the cytoplasmic membrane and to feed electrons into the plastoquinone (PQ) pool of the respiratory electron transport chain (ETC), potentially via a cytochrome-like anchoring unit that binds the H₂-oxidizing subunit to the membrane and facilitates electron transport to the respiratory ETC (Appel and Schulz, 1998; Khanna et al., 2016). Electrons originating from H₂ could then be transported from PQ to cytochrome b₆f, and further to a terminal oxidase where they are used to reduce O₂. The ratio of O₂ to H₂ uptake observed in the dark in the afternoon (Figure 4A) suggests that, under these conditions, 1 out of 50 electrons reducing O₂ comes from H₂, with the remainder originating from carbohydrates.

Processes Involved in the Light

In the light, O₂ and H₂ fluxes are determined by photosynthesis and N₂ fixation in addition to respiration and H₂ uptake. Net H₂ evolution was observed in light but not in dark (Figure 2), which we attribute to the light-dependence of nitrogenase activity observed in previous studies on Trichodesmium (e.g., Rabouille et al., 2006; Staal et al., 2007). Activation of nitrogenase by light was also reflected in the instantaneous increase in H₂ concentrations when the light was switched on during the time of highest N₂ fixation (Figure 3). These findings are in agreement with previous studies showing instantaneous decreases in net H₂ evolution in response to addition of NH₄⁺ (which inhibits nitrogenase), DCMU (an inhibitor of PSII), and darkness in Trichodesmium (Wilson et al., 2010, 2012), as well as a direct dependence of H₂ evolution on PSI activity in Cyanothece (Min and Sherman, 2010; Skizim et al., 2012). Notably, the interaction between O₂ evolution, nitrogenase activity and H₂ uptake can be modulated by the reduction state of the photosynthetic electron transport chain (Wilson et al., 2012), suggesting that the quantitative relationship between these processes is strongly dependent on the instantaneous light intensity. Due to the O₂-sensitivity of uptake hydrogenase (e.g., Arp and Burris, 1981; Houchins and Burris, 1981; Axelsson and Lindblad, 2002), the high levels of photosynthetic O₂ evolution in the afternoon (Figure 2C) might impede not only H₂ evolution by nitrogenase but also H₂ uptake, leading to lower levels of H₂ uptake in the afternoon than in the morning (Figure 2B), and causing a loss of the correlation between H₂ and O₂ fluxes in the light (Figure 4B).

Regulation Over the Diel Cycle

The timing of highest variability in net H₂ fluxes coincided with the time of highest net O₂ uptake as well as the midday peak in N₂ fixation reported previously (e.g., Berman-Frank et al., 2001). The down-regulation in net O₂ evolution during this part of the day is believed to act as a mechanism to protect nitrogenase from O₂ (e.g., Berman-Frank et al., 2001) and might additionally benefit uptake hydrogenase, which is also O₂-sensitive (e.g., Houchins and Burris, 1981; Axelsson and Lindblad, 2002; Tamagnini et al., 2007). H₂ evolution and N₂ fixation rates have been previously shown to be correlated over a range of conditions (Tamagnini et al., 2002 and references therein), yet most previous studies on H₂ evolution by Trichodesmium did not resolve H₂ fluxes over the diel cycle. A laboratory study on Trichodesmium cultures grown in single filaments under a 12:12 h light-dark cycle at significantly lower intensities (44 μmol photons m⁻² s⁻¹) did not show the same diel pattern in net H₂ evolution (Wilson et al., 2010), potentially due to feedbacks of colony formation and/or the range of light intensities in our field study.

During the morning, when N₂ fixation and thus H₂ evolution were presumably highest, elevated H₂ concentrations might elicit an increase in the activity of uptake hydrogenase. The fact that we observed high net H₂ evolution as well as uptake at this time of the day may reflect a mismatch in timing between the activation of nitrogenase and uptake hydrogenase – a slight offset in timing would result in strong net fluxes, while a balanced system with a constant fraction of H₂ recycling would result in minimal net fluxes. Interestingly, H₂ uptake was highest at morning/midday even in dark measurements (Figure 2B) where nitrogenase was presumably inactive, suggesting that uptake hydrogenase activity is regulated not only by H₂ availability (i.e., by nitrogenase activity) or by light, but potentially also via an endogenous rhythm, as shown for nitrogenase in Trichodesmium (Chen et al., 1998). Both, uptake hydrogenases and nitrogen metabolism, are regulated by the transcriptional regulator NtcA (e.g., Tamagnini et al., 2002; Khanna et al., 2016), which was suggested to be under the control of a circadian rhythm in Cyanothece (Bradley and Reddy, 1997). In the afternoon, H₂ fluxes were less pronounced (Figures 2A,B) and did not respond to light (Figure 3B) as strongly as in the morning, even though colonies were clearly active and generally responsive to light as indicated by the high O₂ evolution in the light (Figures 2C, 3C). H₂ evolution by nitrogenase can presumably not respond to light at this time of the day since nitrogenase becomes inactivated by post-translational modification in the afternoon (Zehr et al., 1993). The fact that also H₂ uptake did not respond strongly to light (Figure 3) and uptake rates were low despite addition of H₂ (Figure 2) at this time of the day suggests that uptake hydrogenase might be down-regulated in the afternoon in a similar way as nitrogenase.

The Potential Role of H₂ in Iron Acquisition

Using a recently optimized radiotracer method for measuring iron uptake from mineral iron by Trichodesmium colonies (Basu and Shaked, 2018), we were able to investigate the effects of H₂ on iron uptake under conditions that closely resembled the acquisition of iron from dust under natural conditions. Interestingly, these experiments revealed that the uptake of mineral iron was strongly stimulated by the presence of H₂ (Figure 6). This effect was observed across different colony morphologies, which are generally assumed to represent different species (e.g., Hynes et al., 2012). Also the timing of H₂ uptake and iron uptake was synchronized: Both H₂ uptake and evolution as well as the response of iron uptake to H₂ were strongest around midday, whereas in the afternoon, both H₂ uptake and iron uptake responded less strongly to H₂. Taken together, these findings suggest a potential mechanistic link between iron uptake and H₂ metabolism. While the exact mechanisms of mineral iron acquisition by Trichodesmium are yet to be elucidated, there are several indications for a reductive step involved in the process, such as the slow-down of iron uptake from ferrihydrite in the presence of the Fe(II) ligand ferrozine (Supplementary Figure 1). The electron source for this reduction has not been identified. Based on the large H₂ fluxes in colonies in combination with the stimulation of iron uptake by H₂ and indications for co-regulation of uptake hydrogenase and iron uptake over the diel cycle, we propose that H₂ may function as an electron source for mineral iron reduction. While the quantitative importance of this potential mechanism needs to be investigated in future studies under H₂ concentrations that more closely mimic natural conditions, we focus here on identifying a potential pathway of electron flow that might provide such a link between H₂ and mineral iron reduction on the cellular level (Figure 7).

FIGURE 7

Figure 7. Scheme of a potential cellular link between N₂ fixation, H₂ and O₂ fluxes and iron uptake in Trichodesmium. Dotted lines indicate electron flow. Please note that uptake hydrogenase may be restricted to non-photosynthetic diazocytes and/or located on the cytoplasmic membrane instead of the thylakoid membrane and thus coupled to respiratory electron transport but not photosynthesis (Tamagnini et al., 2002; Khanna et al., 2016). ARTO, alternative respiratory terminal oxidase; Cyt b₆f, cytochrome b₆f complex; Cyt c oxid, cytochrome c oxidase; Fd, ferredoxin; FeoB, ferrous iron transporter B; FeOH, iron oxide; FNR, ferredoxin-NADPH-oxidoreductase; hup, uptake hydrogenase; nif, nitrogenase; PQ, plastoquinone; PSI, photosystem I; PSII, photosystem II; SDH, succinate dehydrogenase.

The link between uptake hydrogenase and the respiratory electron transport chain has been reported previously (e.g., Appel and Schulz, 1998; Wilson et al., 2010), and was reflected in the correlation between H₂ and O₂ uptake in our study (Figure 4). In Synechocystis, mineral iron reduction was recently suggested to be facilitated by the alternative respiratory terminal oxidase ARTO (Kranzler et al., 2014). ARTO is also expressed and regulated in response to iron availability in Trichodesmium (Polyviou et al., 2018). As it accepts electrons from PQ (Lea-Smith et al., 2016), it could potentially serve as a link between the respiratory electron transport chain and iron reduction. Since on average, cellular iron uptake rates are several orders of magnitude lower than respiration rates (e.g., iron uptake of ca. 1–9 × 10⁻¹⁸ mol Fe cell⁻¹ d⁻¹, Basu and Shaked, 2018, as compared to respiration of ca. 3 × 10⁻¹² mol O₂ cell⁻¹ d⁻¹, Eichner et al., 2017), only a small fraction of electrons in the respiratory pathway would need to be channeled to iron reduction to meet cellular iron demands, while the remaining electrons would reduce O₂. Nevertheless, a coupling of H₂ uptake to iron acquisition via the respiratory ETC (i.e., electron flow from uptake hydrogenase via the PQ pool to ARTO) could explain the stimulation of iron uptake by H₂ observed in this study (Figure 6) during the time of the day when high rates of H₂ uptake were observed (Figures 2, 3). The reduced activity of uptake hydrogenase in the afternoon/evening (Figures 2, 3), potentially caused by a down-regulation/post-translational modification of the protein, in turn, could explain the lack of a response in iron uptake to H₂ at this time of the day (Figure 6).

Previous studies on heterocystous, filamentous cyanobacteria suggested that uptake hydrogenase is present only in heterocysts (e.g., Tamagnini et al., 2007). Such a restriction of uptake hydrogenase to N₂-fixing cells would provide not only a tight spatial coupling between H₂ evolution and uptake, but also the benefit of reduced O₂ concentrations that would protect both nitrogenase and uptake hydrogenase from oxidative damage. In Trichodesmium, it has been suggested that only 10–20% of cells in each filament, the diazocytes, express nitrogenase and actively fix N₂ (e.g., Bergman and Carpenter, 1991, Berman-Frank et al., 2001). Notably, those cells would have a significantly higher iron requirement associated to synthesis of the iron-rich nitrogenase than vegetative cells (19–53% of cellular iron was estimated to be bound in nitrogenase on average across all cell types; Kustka et al., 2003). The pathway of iron reduction coupled to H₂ uptake we propose here could provide a mechanism of surplus iron reduction to fuel the enhanced iron demand in these cells. The restriction of hydrogenase to diazocytes would also imply that H₂ uptake and photosynthesis are separated on a single-cell level, and thus do not share the same electron transport chain, potentially explaining why light did not have strong direct effects on H₂ uptake (at the time when nitrogenase was presumably inactive; Figure 5). Furthermore, assuming that the H₂ fluxes we detected as an average of all cells in a single colony were actually catalyzed by only 10–20% of the cells, it should be noted that the ratio of H₂:O₂ uptake in those cells would be 5–10× larger than the average ratio we measured (Figure 4). Particularly since photosynthetic water splitting is presumably not available as an electron source in diazocytes, H₂ as an alternative electron source might be relatively more important in these cells. In summary, although the contribution of H₂ to total electron flow in the ETC (as estimated from the ratio of H₂:O₂ uptake; Figure 4) was relatively low when averaged over all cells and determined outside of the reported peak in N₂ fixation, we suggest that in specialized N₂-fixing cells at the time of active N₂ fixation, a significant part of the electron flux in the respiratory electron transport chain might be channeled from H₂ to iron. Future studies should thus aim to confirm the physiological mechanism and quantitative importance of this pathway in Trichodesmium. Complementary molecular biological studies on the metabolic potential of associated bacteria in Trichodesmium colonies will bring further insights into the potential contribution of these bacteria to both hydrogen uptake and iron reduction within colonies.

Conclusion

By high-resolution measurements of H₂ and O₂ gradients on single colonies, we were able to demonstrate highly variable H₂ microenvironments in colonies of the important N₂ fixer Trichodesmium. We found that the net H₂ flux could be directed either in or out of the colony. H₂ uptake was regulated not only by H₂ availability and by light, but also over the diel cycle, potentially by an endogenous clock. Both H₂ uptake and evolution were generally most active in the morning at the time of highest N₂ fixation, reflecting the tight link between nitrogenase and uptake hydrogenase activity. The large variability in H₂ fluxes between individual colonies analyzed under similar conditions furthermore highlights that single colonies may provide diverse micro-niches differing strongly in H₂ metabolism, including systems characterized by either net H₂ evolution or net H₂ uptake. Given that the experimental approaches used in previous studies (measuring bulk H₂ evolution rates under natural H₂ concentrations) do not allow for detecting such potential hot-spots of net H₂ uptake, our findings highlight the need for more broadly applying (1) methods that allow for quantifying H₂ uptake rather than only net evolution (such as ³H₂ assays; Wilson et al., 2013), and (2) high-resolution measurements that can reveal small-scale environmental variability (such as microsensors).

Combining H₂ measurements with radiotracer experiments, we furthermore observed a strong stimulation of mineral iron uptake in the presence of H₂, indicating a previously unrecognized link between H₂ and acquisition of iron. We propose a potential pathway of electron flow linking N₂ fixation, H₂ uptake, respiration and iron reduction on the cellular level, which should be the subject of further physiological investigations. This link between H₂ uptake and mineral iron acquisition adds a new component to the complex network of nutrient acquisition mechanisms employed by Trichodesmium in the colony microenvironment.

Data Availability

The raw data supporting the conclusions of this manuscript will be made available by the authors, without undue reservation, to any qualified researcher.

Author Contributions

ME, YS, and DB designed the study. ME performed the microsensor measurements with help by DB and analyzed the data. SB performed the iron uptake experiments and analyzed the data. ME drafted the manuscript. All authors contributed to the data interpretation and writing the manuscript, and read and approved the final version of the manuscript for submission.

Funding

This work was supported in part by Israel Science Foundation (www.isf.org.il) Grant 458/15 and the German–Israeli Foundation for Scientific Research and Development (www.GIF.org.il) Grant 1349 awarded to YS and MG (and enabling visits by ME to IUI). Additional funding was provided by the Minerva Foundation (awarded to ME) and the Max Planck Society.

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We are grateful to Murielle Dray for technical assistance at IUI, to the technical assistants in the Microsensor Group at MPI for producing the microsensors, as well as to the staff of the Electronics Workshop at MPI for their technical support with the associated equipment. We also thank Ondrej Prasil and Nir Keren for providing valuable feedback on earlier versions of this manuscript.

Supplementary Material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2019.01565/full#supplementary-material

Footnotes

1. ^ http://genome.jgi-psf.org/finished_microbes/trier/trier.home.html

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