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Distribution Analysis of Hydrogenases in Surface Waters of Marine and Freshwater Environments

Distribution Analysis of Hydrogenases in Surface Waters of Marine and Freshwater Environments

  • Martin Barz, 
  • Christian Beimgraben, 
  • Torsten Staller, 
  • Frauke Germer, 
  • Friederike Opitz, 
  • Claudia Marquardt, 
  • Christoph Schwarz, 
  • Kirstin Gutekunst, 
  • Klaus Heinrich Vanselow, 
  • Ruth Schmitz



Surface waters of aquatic environments have been shown to both evolve and consume hydrogen and the ocean is estimated to be the principal natural source. In some marine habitats, H2 evolution and uptake are clearly due to biological activity, while contributions of abiotic sources must be considered in others. Until now the only known biological process involved in H2 metabolism in marine environments is nitrogen fixation.

Principal Findings

We analyzed marine and freshwater environments for the presence and distribution of genes of all known hydrogenases, the enzymes involved in biological hydrogen turnover. The total genomes and the available marine metagenome datasets were searched for hydrogenase sequences. Furthermore, we isolated DNA from samples from the North Atlantic, Mediterranean Sea, North Sea, Baltic Sea, and two fresh water lakes and amplified and sequenced part of the gene encoding the bidirectional NAD(P)-linked hydrogenase. In 21% of all marine heterotrophic bacterial genomes from surface waters, one or several hydrogenase genes were found, with the membrane-bound H2 uptake hydrogenase being the most widespread. A clear bias of hydrogenases to environments with terrestrial influence was found. This is exemplified by the cyanobacterial bidirectional NAD(P)-linked hydrogenase that was found in freshwater and coastal areas but not in the open ocean.


This study shows that hydrogenases are surprisingly abundant in marine environments. Due to its ecological distribution the primary function of the bidirectional NAD(P)-linked hydrogenase seems to be fermentative hydrogen evolution. Moreover, our data suggests that marine surface waters could be an interesting source of oxygen-resistant uptake hydrogenases. The respective genes occur in coastal as well as open ocean habitats and we presume that they are used as additional energy scavenging devices in otherwise nutrient limited environments. The membrane-bound H2-evolving hydrogenases might be useful as marker for bacteria living inside of marine snow particles.


The composition of earth's atmosphere is the result of a number of concurring processes and a matter of continuous change. Especially the amount of trace gases governs important aspects of the gas cover of our planet, such as its retention capacity of heat or the amount of ozone present. After methane, hydrogen is the second most abundant trace gas in the atmosphere, making up around 0.5 ppm to 0.6 ppm [1], [2].

Approximately 90% of hydrogen evolution is due to photochemical oxidation of hydrocarbons such as methane in the atmosphere, the combustion of fossil fuels and biomass burning. Natural evolution results from volcanic activity, the nitrogen fixation process in legumes and an uncharacterized source in the oceans. The latter comprises the majority with around 6% (6 Tg per year [3]).

The removal of hydrogen is either due to its reaction with hydroxyl radicals in the atmosphere or by its reaction with hydrogenases in the soil. In particular, hydrogen uptake into the soil is responsible for the largest term with an estimated 75% to 77% globally [1][5]. This is further corroborated by the lower average concentration of hydrogen found on the northern hemisphere, with its larger landmass [1]. Hydrogen uptake was attributed to aerobic hydrogen-oxidizing bacteria and extracellular enzymatic activity. Abiotic removal has been previously considered since hydrogen concentrations are below the threshold level found for cultures of aerobic hydrogen oxidizing bacteria that still maintains growth [6].

In contrast to soil, supersaturating concentrations of hydrogen have been measured in aquatic environments. In all cases, concentrations were highest at the surface and steeply decreased down to the thermocline while the deep ocean is undersaturated. Although a systematic analysis is not available it appears that surface waters of tropical and subtropical oceans are generally hydrogen sources [7][9]. In contrast, concentrations lower than the expected atmospheric equilibrium have been observed in higher latitudes and both hydrogen uptake and production vary depending on the season [10], [11]. In some fresh water lakes supersaturation has also been found [12], with a maximum at dawn [13]. The highest hydrogen concentrations were in the upper water column, which correlated with the maximum of primary production [13], [14].

Marine hydrogen uptake has been attributed to particulate fractions of 0.2 µm to 5 µm in size [11] and, like in freshwater lakes, most probably correlates with aerobic hydrogen-oxidizing bacteria [13]. Hydrogen production in the oceans was found to depend on solar radiation and clearly shows a diurnal variation with a maximum around noon [8], [9]. Since the nitrogenase inevitably produces at least one molecule of hydrogen per dinitrogen reduced to ammonia, cyanobacterial nitrogen fixation is thought to be the major source of hydrogen in these oceanic regions. Studies on heterocystous cyanobacteria demonstrated that hydrogen cycling by these strains is highly effective, although under CO2-limitation around 0.1 nmol H2 h−1 (mg chlorophyll)−1 escapes to the environment [15]. In contrast to this, in-situ measurements of Trichodesmium thiebautii (former Oscillatoria thiebautii), which is one of the major oceanic N-fixing strains, questioned whether its hydrogen evolution is actually sufficient to explain the concentrations found [16].

Recently it was shown that photochemical production of hydrogen from chromogenic dissolved organic matter can contribute, at least in part, to hydrogen production in fresh water lakes as well as coastal seawater [17]. Therefore, abiotic sources should be taken into account.

In the microbial world hydrogen is a valuable energy source that is exchanged efficiently between different prokaryotes and anaerobic eukaryotes. Some produce hydrogen while fermenting whereas others capture it to drive anaerobic or aerobic respiration and make use of its energy. A wealth of different enzymes called hydrogenases have been found in microorganisms that are able to split or form hydrogen [18], [19].

Hydrogenases are classified according to their metal content into the Fe-, FeFe-, and NiFe-varieties. Fe-hydrogenases are confined to the methanogenic archaea and FeFe-hydrogenases occur in bacteria and anaerobic eukaryotes. NiFe-hydrogenases are separated into 4 different groups and are widespread in archaea and bacteria [19], [20]. Most purified hydrogenases are only active under anoxic conditions, but there are some NiFe-hydrogenases from aerobic H2-oxidizing bacteria that are able to oxidize hydrogen at ambient oxygen concentrations [21].

Although hydrogenases have been investigated for a long time in a variety of different microorganisms it is rather difficult to deduce their physiological function on the basis of their classification alone. In Table 1 a tentative assignment of their metabolic roles is given. However, this assignment needs to be treated cautiously since several studies found surprising variations. Hydrogenase 2 of E. coli belongs to the group 1 H2-uptake hydrogenases and was originally described as H2-oxidizing enzyme [22]. In contrast, recent electrochemical data suggests that the hydrogenase 2 is working as a bidirectional enzyme [23]. Another interesting variance was found in case of the group 4 membrane-bound H2-evolving hydrogenase. In many cases these enzymes seem to be used under fermentative conditions to generate a proton gradient (e.g. [24]) but in other cases they might be used to oxidize H2 and reduce ferredoxin with the concomitant use of a proton gradient [25] or even for H2 uptake in N-fixing bacteria [26].

Systematic studies concerning the distribution of hydrogenases in different habitats to unravel their ecophysiological role are not yet available. Apart from the investigation of some specific soil hydrogenases [27], [28] only two studies attempted the amplification of FeFe-hydrogenase sequences from microbial mats [29], [30]. Although these works showed a surprising variety of these hydrogenases the short sequences amplified preclude any assignment of their function.

The hydrogen concentrations found in a variety of surface waters prompted us to investigate the presence and distribution of all known hydrogenases in marine and freshwater environments. Moreover, the ecological distribution of their genes was analyzed to collect valuable hints for their physiological functions and their oxygen tolerance.

To this end we analyzed the distribution of hydrogenases in cyanobacteria since they are one of the largest prokaryotic groups that occur in aquatic surface waters. The search was then expanded to the complete genomes of bacteria isolated from marine surface waters ( [31]) and the global ocean sampling metagenomic database ([32][34] for all the families of hydrogenases as classified by Vignais et al. [20] and Vignais and Billoud [19]. In parallel, we investigated DNA isolated from samples taken from the North Atlantic, Mediterranean Sea, North Sea, the Baltic Sea and the fresh water lakes Westensee and Selenter See in Northern Germany for the presence of the genes of the bidirectional NAD(P)-linked hydrogenase. Our results reveal that these enzymes are surprisingly widespread in surface waters and a clear bias toward waters with terrestrial influence is obvious.


Distribution of hydrogenases in cyanobacterial genomes

Cyanobacteria are known to harbor two different NiFe-hydrogenases. One is called bidirectional (group 3d) since it can produce or take up hydrogen, depending on the physiological conditions and the other is an uptake hydrogenase (group 2a) that is linked to the nitrogen fixation process and seems to be confined to diazotrophic strains [35], [36]. A phylogenetic analysis revealed a close ancestry of both hydrogenases to the filamentous anoxygenic photosynthetic bacteria (the former green non-sulfur bacteria)[37].

A search of genebank ( and cyanobase ( for all available cyanobacterial sequences revealed the presence of the bidirectional NAD(P)-linked hydrogenase (the large subunit HoxH was used in the BLAST search [38]) in all the freshwater strains and all the strains isolated from microbial mats, salt marshes, and the intertidal zone (Table 2). In contrast, only four out of the seven available coastal genomes harbor the gene for the bidirectional enzyme and it was completely absent in oceanic strains. Genomestreamlining and iron limitation [39] in the open ocean could be used as arguments for the absence of the bidirectional hydrogenase genes in the picoplanktonic Prochlorococcus and Synechococcus strains. But even the typical open ocean strains Crocosphaera watsonii and Trichodesmium erythraeum with genome sizes above 6 Mbp do not harbor this hydrogenase, although both have the uptake hydrogenase, which has an iron requirement similar to the bidirectional enzyme (Fig. 1 and Table 2). In addition the unicellular marine strain UCYN-A that lacks photosystem II shows an extremely reduced genome and still contains the hup-genes [40].

Figure 1. Comparison of cyanobacterial genome sizes and the distribution of the bidirectional NAD(P) linked hydrogenase gene hoxH.

Genomes without the bidirectional hydrogenase are depicted in black and those with it are red. The marine diazotrophic cyanobacteria containing the genes of the uptake hydrogenase hupL are shown in cyan. The cluster of black circles at the lower left end of the line represents the small genomes of the Prochlorococcus and Synechococcus strains.

Table 2. Occurrence of the bidirectional NAD(P)-linked hydrogenase (HoxH) and the membrane-bound uptake hydrogenase (HupL) in cyanobacteria.

All the completely sequenced cyanobacterial strains that harbor the bidirectional hydrogenase genes also harbor the gene of a pyruvate:flavodoxin/ferredoxin oxidoreductase (PFOR), nifJ. In two genomes (Synechococcus WH 5701 and Arthrospira maxima), this gene is either part of the hyp-gene cluster or in close proximity to the hox-genes, suggesting that the birdirectional hydrogenase is used to dispose of electrons during fermentation via a PFOR-like enzyme (Table 2).

The occurrence of the uptake hydrogenase (HupL, group 2a) in cyanobacteria does not correlate with a specific habitat but with the diazotrophy of the respective strains, as indicated by the presence of the nitrogenase genes (e.g. NifD)(Table 2). Of the completely sequenced genomes two Synechcococcus strains isolated from a hot spring and Cyanothece sp. PCC 7425 harbor the nitrogenase genes but no uptake hydrogenase. This confirms the previous finding of a marine nitrogen-fixing Synechococcus strain without an uptake hydrogenase [37].

Cyanothece sp. PCC 7425 is the only strain containing the genes of the bifunctional NAD(P) linked hydrogenase (group 3b)(Table 2) but expression and metabolic activity of this enzyme have not yet been demonstrated.

Distribution of hydrogenases in genomes of heterotrophic bacteria isolated from marine surface waters

Representatives of each of the hydrogenase classes were used to search the completely sequenced prokaryotic genomes in the genebank (Table 3). Of the approximately 1210 prokaryotic genomes (as of March 2010) 149 were isolated from marine surface waters and in 33 of these genomes, one or several hydrogenases occur, making up 22% of the total (Table 4, Table S1 supporting information). Since a number of the analyzed genomes is still not complete, this proportion is a minimum estimate. If divided into coastal and open ocean isolates, 25% of the coastal and 14% of the open ocean strains have hydrogenase genes.

Table 3. Hydrogenase and HypX sequences used for searches of the completely sequenced genomes and the GOS metagenomic database.

Table 4. Marine bacteria with FeFe-hydrogenases and NiFe-hydrogenases of the different classes.

The genomes of two Shewanella strains (ANA-3 and MR-4) have all the genes necessary for the expression of a FeFe-hydrogenase. Since this type of hydrogenase is extremely sensitive against and irreversibly inactivated by oxygen [41], this is a surprising finding. However, it should be noted that one strain (ANA-3) has been isolated from a wooden pier that might have been occupied by biofilms that could become anaerobic and the other strain (MR-4) was isolated from the Black Sea, which is the world largest anoxic basin [42]. Therefore, both are considered exceptions and will not be discussed any further.

Concerning the NiFe-hydrogenases, there are 24 genomes with a membrane-bound H2-uptake hydrogenase (group 1), two genomes with a cyanobacterial-type uptake hydrogenase (group 2a)(Sphingopyxis alaskensis RB2256 and Neptuniibacter caesariensis), six genomes with a sensor hydrogenase (group 2b), seven genomes with a bifunctional hydrogenase (group 3b), four genomes with a bidirectional NAD(P)-linked hydrogenase (group 3d), and three genomes with a membrane-bound H2-evolving hydrogenase (group 4) similar to hydrogenase 3 of E. coli.

The genomes of the Roseovarius group contain large gene clusters with the membrane-bound hydrogenase in conjunction with a sensor hydrogenase and the whole complement of the two-component system (Fig. S1, supporting information). The sensor enzyme is a receptor that enables the cells to detect hydrogen in the environment and to activate transcription of the hydrogenase structural genes [43][46]. The same gene clusters also contain a number of additional genes that encode for proteins such as HupK that have been shown to be necessary for the production of an oxygen tolerant hydrogenase in R. eutropha [47], [48].

The genomes of the Vibrionaceae harbor a membrane-bound H2-evolving hydrogenase (Fig. S2, supporting information) and a second membrane-bound hydrogenase. This is the necessary combination that can be used under anaerobic conditions to establish a proton gradient by hydrogen cycling in a single cell [49].

Additionally, the genome of N. caesariensis (former Oceanospirillum [50]) is worth mentioning. It contains a membrane-bound enzyme, a cyanobacterial like uptake hydrogenase, a sensor, and a bifunctional hydrogenase. A phylogenetic analysis confirmed that the HypX encoded in its genome belongs to the group of hydrogenase maturation factors (Fig. S3, supporting information). HypX was shown to render the soluble hydrogenase of the Knallgas bacterium Ralstonia eutropha oxygen insensitive [51]. The membrane-bound hydrogenase of N. caesariensis is a close relative of the same hydrogenase of R. eutropha (Fig. S4, supporting information), which is evidence that this bacterium and the Roseovarius strains are able to perform aerobic hydrogen oxidation in marine environments.

Distribution of hydrogenases in metagenomic databases

Single bacterial strains allow a detailed analysis of part of the genomes that occur in the specific environment they have been isolated from. However, isolated strains provide only a glimpse on the genetic diversity that might be present in the habitat from which they originate, given that most microbial strains are unculturable [52][54]. Therefore, we searched the global ocean sampling database (GOS)[32][34] with the same representative hydrogenases as given in Table 3 and the representatives of the small hydrogenase subunits (Fig. S5 to S7, supporting information).

This database contains millions of sequence reads that have been obtained mostly from biological samples with a particle size of 0.2 to 0.8 µm. Due to this size fractionation, the major proportion of the sequences belongs to Pelagibacter ubique and the Prochlorococcus/Synechococcus group of cyanobacteria [33]. Since the large number of sequences in the Sargasso Sea metagenome belonging to the Shewanellaceae and the Burkholderiaceae was discussed to be a contamination [55] Station 11 was not included in the analysis.

We could not detect any cyanobacterial bidirectional hydrogenase in the samples taken from the open ocean. All the cyanobacterial HoxH sequences that could be found in the database are from a single sample taken at Punta Comorant, a hypersaline pond with low oxygen levels [56] on the Galapagos Islands (Fig. 2). These sequences were most similar to the available bidirectional hydrogenases of Synechococcus strains (Fig. S8, supporting information). Thus, the GOS sampling and sequencing effort should have been able to capture any HoxH sequence present in the Prochlorococcus/Synechococcus group. Although it has to be taken into account that environmental sequencing does not capture 100% of the present DNA sequences it seems highly probable that this cyanobacterial hydrogenase is absent in these strains in these environments as already deduced from the whole genomes (Table 2, Table S1, supporting information).

Figure 2. Distribution of bidirectional NAD(P) linked hydrogenases found in the GOS database of the different prokaryotic groups.

The hoxH sequence of Synechocystis sp. PCC 6803 (Table 3) was used for the search and a total of 48 sequences has been found. On the right the number of sequences from the different sampling stations is shown.

These findings are also corroborated when looking at the hoxH sequences of the Burkholdericeae. Although these bacteria make up a major fraction of all the oceanic metagenome sequences, there are only representatives from Punta Cormorant with this hydrogenase (Fig. 2), whereas no sequences of this group have been retrieved from the open ocean. Altogether 48 hoxH sequences could be found but apart from three coastal stations (Mangrove on Isabella Island, Cape May and Dirty Rock), which accounted for 4 sequences all of the other 44 were exclusively from Punta Cormorant. This confirms the presence of hoxH in shallow coastal environments and ponds in a variety of different bacterial groups.

The largest group of sequences in the metagenome database were those of the membrane-bound NiFe-hydrogenases. Again most of the 51 sequences were found at Punta Cormorant, although 11 sequences were detected in the datasets of coastal stations (New Harbor, Dirty Rock, Yucatan Channel, Nags Head, a Mangrove on Isabella Island) and two were found in the open ocean (outside Seychelles and 250 miles of Panama) (Fig. 3).

Figure 3. Distribution of membrane-bound hydrogenases found in the GOS database of the different prokaryotic groups.

The hupL sequence of Desulfovibrio vulgaris (Table 3) was used for the search and a total of 51 sequences has been retrieved. On the right the number of sequences from the different sampling stations is shown.

Cyanobacterial-like uptake hydrogenases could also be found in the metagenomic dataset (Fig. 4). Because of the size fractionation (0.2–0.8 µm) most of the larger diazotrophic cyanobacteria have been excluded from this analysis. Therefore, although many of the samples have been taken in regions known to be inhabited by this cyanobacterial group only two sequences could be retrieved from the whole dataset. A total of 35 sequences could be found. Most of these sequences originate from coastal sites (28) but four sequences are from the open ocean (Sargasso Sea, Reunion Island and 250 miles off Panama City).

Figure 4. Distribution of cyanobacterial-like uptake hydrogenases found in the GOS database of the different prokaryotic groups.

The hupL sequence of Nostoc sp. PCC 7120 (Table 3) was used for the search and a total of 35 sequences has been retrieved. On the right the number of sequences from the different sampling stations is shown.

Searches for the small hydrogenase subunit genes retrieved 23 sequences of the bidirectional NAD(P)+-linked hydrogenases, 37 of the membrane bound H2 uptake hydrogenases and 18 of the cyanobacterial-like uptake hydrogenases. In all these cases the numbers are close to the expected number when comparing the gene sizes of the respective large and small hydrogenase genes (Fig. S5 to S7, supporting information).

Sequences of the oxygen sensitive FeFe-hydrogenases retrieved from the GOS database were from a Mangrove (Isabella Island) and the hypersaline pond at Punta Cormorant. In all other samples no FeFe-hydrogenase was found (Fig. 5) and none of the archaebacterial hydrogenases were found in the metagenome sequences.

Figure 5. Distribution of FeFe-hydrogenases found in the GOS database of the different prokaryotic groups.

The hydA sequence of Clostridium pasteurianum (Table 3) was used for the search and a total of 10 sequences have been found. On the right the number of sequences from the different sampling stations is shown.

Recently large amounts of metatranscriptomics data became available (e.g. [57]). A search of the respective dataset revealed the presence of three transcripts of membrane-bound H2-uptake hydrogenases. One transcript was most similar to a cyanobacterial uptake hydrogenase, one to the Flavobacteriaceae and one to the Bradyrhizobiaceae. In this dataset only samples from the open ocean are available.

Detection of sequences of the bidirectional NAD(P)-linked NiFe-hydrogenase in the North Atlantic, Mediterranean Sea, North Sea, Baltic Sea, and two freshwater lakes

Although all NiFe-hydrogenases share two characteristic motifs with altogether four cysteins at the N- and C-terminus for the binding of the NiFe active site, it is impossible to design degenerated primers that bind to the genes of all different classes of these enzymes. Therefore, we limited our effort to a single class and constructed degenerated primers specific for the bidirectional NAD(P)-linked hydrogenases of cyanobacteria, the Chloroflexaceae and some proteobacteria. In cyanobacteria this enzyme is known as the bidirectional hydrogenase. It is closely related to the soluble hydrogenase of Ralstonia eutropha and the respiratory complex I [58], [59].

We collected surface water from Stollergrundrinne outside the Kielfjord (Baltic Sea), in the Norderpiep west of Büsum (North Sea) and two freshwater lakes in northern Germany, Westensee and Selenter See. These samples were sequentially filtered on 10 µm and 0.2 µm filters and DNA isolated from the retained material. In samples from all these locations we could detect hoxH. In Fig. 6 the distribution of sequences on the different bacterial groups is shown for the different stations.

Figure 6. Distribution of bidirectional NAD(P) linked hydrogenases in samples taken from Norderpiep (North Sea), Stollergrundrinne (Baltic Sea) and the freshwater lakes Westensee and Selenter See.

From the Baltic Sea as well as the fresh water lakes we could amplify a large number of cyanobacterial hoxH that are most similar to the Chroococcales (most closely related to Cyanothece, Microcystis and Synechocystis) or the filamentous, heterocystous Nostocaceae. In the North Sea the α–proteobacterial group Rhodobacteraceae made up the same proportion as all the cyanobacterial sequences taken together. From the freshwater mesotrophic lakes Westensee and Selenter See we could only amplify cyanobacterial hoxH (Chlorococcales, Nostocaceae and Oscillatoriales) and in each case some sequences of methylotrophic bacteria and Dictyoglomaceae.

In contrast to this, all attempts to amplify sequences of the bidirectional NAD(P)-linked hydrogenases from the samples taken in the North Atlantic off the west African coast and the Ionian Sea (Mediterranean Sea) were negative. This corroborates that the open ocean and marine oligotrophic waters are devoid of this hydrogenase type.


Any conclusion concerning the activity of a gene from its environmental distribution is hampered by the fact that it is not necessarily expressed in a specific environment. Genomes might have genes in store that are not necessary to survive under the present-day conditions, but can be used to invade other niches or to prepare the organism for a drastic change. In the case of the distribution of hydrogenases found in this work, this scenario seems highly unlikely. For several reasons described in detail below, we think that biological hydrogen production and consumption, as depicted in Fig. 6, might be common in a large number of marine and freshwater habitats.

All strains from the open ocean were free of the bidirectional NAD(P) linked hydrogenase. Neither the cyanobacterial genomes nor all of the heterotrophic bacteria (Table 2 and Table S1, supporting information) or the metagenomic sequences harbor this hydrogenase. In addition, our efforts to amplify these hydrogenase genes from the North Atlantic or the Mediterranean Sea were unsuccessful. Since the diazotrophic cyanobacterial strains and the heterotrophic bacteria from the open ocean have other types of hydrogenases, there is no selection pressure against these enzymes per se. However, there is a clear bias of the bidirectional type to environments such as coastal marine waters, ponds, freshwater lakes and microbial mats (Table 2, Fig. 2, Fig. 6 and Table S1, supporting information), where cyanobacteria and heterotrophic bacteria might encounter micro-oxic or anaerobic conditions. In cyanobacteria this type of enzyme was shown to be activated under anaerobiosis and to be responsible for fermentative hydrogen production [60]. This is corroborated by the distribution of the PFOR gene, nifJ, in the same cyanobacteria (Table 2).

Starting from anaerobiosis, the bidirectional hydrogenase is known to be used as an electron valve, when cells switch from fermentation to photosynthesis [61][64]. These findings might explain the high hydrogen concentration found in the morning hours in a eutrophic lake that coincided with the phytoplankton maximum [13]. Oxygen depletion due to high respiratory activity during the night could have activated the hydrogenase in this zone and elicited a fermentative hydrogen production in the dark that continued at dawn until the next morning when photosynthesis resumed, thus causing supersaturating H2 concentrations. A similar diel variation of hydrogen concentrations has also been described for cyanobacterial mats (see e.g. [65]).

In both cases, hydrogen production is certainly not confined to the resident cyanobacteria but can also result from the activity of algae and other heterotrophic bacteria living in the same community.

The large number of genomes of marine bacteria from surface seawaters containing the membrane-bound H2-uptake hydrogenase is remarkable. A search of the current marine metatranscriptomics data [57] revealed the expression of these hydrogenases in cyanobacteria as well as other bacteria in the open ocean.

The membrane-bound hydrogenase gene clusters found in the Rhodobacteraceae (Fig. S1 supporting information) include all the accessory genes that are known from the membrane-bound hydrogenase of R. eutropha. One of the four hydrogenases of N. caesariensis and the hydrogenases of the Roseovarius strains are closely related to this hydrogenase as revealed by phylogenetic analysis (Fig. S1 and S4, supporting information). This type of enzyme is known to be oxygen insensitive and was shown to be active at ambient oxygen concentrations [66], [67]. Electrochemical investigations of this hydrogenase found measurable hydrogen uptake down to levels of 1 to 10 nM [67], which is well in the range of H2 concentrations in surface waters. One of these strains (Roseovarius sp. HTCC 2601) was isolated from the Sargasso Sea, but all of the others were from coastal areas. In these regions, this α-proteobacterial subclass makes up as much as 24% of the bacterioplankton [68] and therefore, their hydrogenases might be widespread in these environments.

Mycobacteria, known to colonize aquatic ecosystems, take up hydrogen in the same concentration range under aerobic conditions [69], supporting the notion that hydrogen consumption in these environments is a common microbial feature. Even though the supersaturating concentrations found in surface waters are below the threshold necessary to support growth exclusively on H2, hydrogen uptake could add to the ability to survive in a variety of these habitats. Similar suggestions have already been made for hydrogen uptake for long-term survival of bacteria [70] and for the ability to oxidize carbon monoxide in the coastal ocean [71], [72]. These suggestions coincide with the aerobic hydrogen uptake demonstrated for particle sizes between 0.2 and 5 µm in coastal waters [11]. This trait is especially important for litho- and heterotrophic bacteria that have to capitalize on as much of the available energy supply as possible, but can be disregarded by photoautotrophs like cyanobacteria.

Bacterial activity was found to be capable of depleting oxygen in marine organic aggregates. In particles as small as 1.5 mm, anoxic conditions emerged. In the same aggregates no methanogenic or sulfate-reducing bacteria could be detected [73]. Our results suggest that these anaerobic microniches might be specifically occupied by bacteria of the Vibrionaceae (Fig. S2, supporting information). Since their membrane-bound H2-evolving NiFe-hydrogenases are encoded in conjunction with subunits of the formate dehydrogenase it seems highly likely that it performs the formate:hydrogen lyase reaction. This reaction is well known from E. coli, where it detoxifies formate produced during fermentation, evolves hydrogen and might be involved in an additional energy-generating step [24]. The membrane-bound hydrogen uptake hydrogenase encoded in the same genomes would allow hydrogen cycling and might be used for additional net transport of protons across the cell membrane [49].

The Altermonadaceae are widespread in marine waters. Two different ecotypes have been sequenced, one is predominant in surface waters whereas the other is known from the deep Mediterranean Sea. The deep ecotype was originally found to harbor the genes of the membrane-bound H2 uptake hydrogenase but our analysis and that of others [74] also found the same sequences at the surface of the Sargasso Sea. It was speculated that these two strains are separated by either being associated with small aggregates (surface type) or large aggregates (deep ecotype)[75]. This might be further support for the use of hydrogenases in transiently anoxic microniches in the ocean.

The diel variation of the H2 concentration in marine surface waters [8], [9] that parallels solar radiation is still awaiting conclusive explanation. Nitrogen fixation is a major source of hydrogen in terrestrial ecosystems [4]. In-situ measurements of the diazotrophic cyanobacterium T. thiebautii suggest that it is a negligible source of hydrogen in the Sargasso Sea [15]. Therefore, nitrogen fixation by filamentous cyanobacteria is an insignificant source of H2 in aquatic ecosystems. Interestingly, a unicellular marine diazotrophic cyanobacterium has been shown to be devoid of the uptake hydrogenase [37] and to produce hydrogen while fixing nitrogen [76]. In general unicellular cyanobacteria perform a temporal separation of the oxygen sensitive energy consuming nitrogen fixation process and oxygenic energy generating photosynthesis between night and day, but some strains also fix nitrogen during the light phase [76], [77]. Unicellular strains are known to provide a considerable part of fixed nitrogen in marine waters [78], [79] and might therefore be responsible for part of the evolved H2. The newly discovered unicellular cyanobacteria without photosystem II [40], [80] harbor the genes of the cyanobacterial uptake hydrogenase (Table 2), which is most similar to those of the Cyanothece group (Fig. S4, supporting information) as expected. Therefore, these strains should be able to recycle the H2 evolved by the nitrogenase.

The distribution of cyanobacterial nitrogen fixers in the ocean and their seasonal abundance are poorly characterized although qPCR data has shown that all groups are widely distributed [81], [82]. One investigation suggests that their distribution is patchy and their rate of nitrogen fixation highly variable [79] and might therefore result in hydrogen evolution in some parts and very low or no evolution in other parts.

Although unicellular nitrogen-fixing cyanobacteria might be responsible for hydrogen evolution in some regions, part of the H2 produced during the day might be of photochemical origin, such as dissociation of organic matter by UV light [17].

Coastal waters are rich in hydrogenase sequences, as suggested by our analysis of complete genomes (Table 2, 4, Table S1, supporting information), and the number of sequences we could amplify of a single class of NiFe-hydrogenases from the North Sea and the Baltic Sea (Fig. 5). The apparent scarcity of sequences from coastal samples in the GOS database can be explained by the filtration procedure. Since mainly particle sizes between 0.2 and 0.8 µm have been used for DNA isolation many of the coastal bacteria and particle associated bacteria have been excluded from the analysis. We hypothezise that the membrane-bound H2 evolving hydrogenase in the genomes of the Vibrionaceae might be used as indicator for bacteria that colonize the inner parts of organic aggregates and thus, have not been sequenced yet in the GOS database.

Our analysis shows that the genetic repertoire of bacteria from surface waters of different environments enables them to produce hydrogen either by their nitrogenase, by hydrogenases linked to fermentative pathways (such as the bidirectional NAD(P) linked hydrogenase), or the membrane-bound H2-evolving hydrogenase. A number of bacteria could oxidize hydrogen as an energy source probably down to the lower nM range and might be responsible for biological hydrogen consumption in freshwater and marine systems.

This study intends to deliver a first key to the elucidation of the underlying biological processes of hydrogen turnover in aquatic ecosystems. Whether a specific body of water is a hydrogen sink or source will depend on a number of factors such as primary production, nitrogen fixation, the concentration of photodegradable organic compounds and organic particles, and the availability of electron acceptors. This is the first evidence that microorganisms can be an integral part of hydrogen turnover in marine waters, but much more remains to be learned. This is especially true when considering oxygen minimum zones [83] that have not been investigated for the presence of hydrogen or hydrogenases until now.

Materials and Methods

Sample collection

Samples were collected from the surface. In the North Sea water was collected in the Norderpiep (54°13′N/8°27′E), in the Baltic Sea it was collected in the Stollergundrinne (54°29′N/10°13′E) and from the freshwater lakes Selenter See (54°18′25N/10°28′53E) and Westensee (54°17′53N/9°57′09E) at least four times a year from every season. These samples were sequentially filtered on 10 µm and 0.2 µm filters with a peristaltic pump 620 S (Watson-Marlow Bredel).

Samples from the Mediterranean Sea were taken from the Ionian Sea at station 2 (36°41′N/21°39′E), station 3 (36°50′N/21°31′E), station 5.2 (36°37′N/21°17′E) and station 6 (36°42′N/21°04′E). In this case 5 l water from a depth of 5 m was filtered on 5 µm and then on 0.2 µm.

The samples from the North Atlantic were taken during the Poseidon 284 cruise at 18°N/30°W, 25°N/30°W and 29°/30°W in April 2002.

For DNA isolation the UltraClean™ Soil DNA Kit (Mo Bio, Carlsbad CA, USA) was used.

DNA amplification and sequence analysis

Sequences of the bidirectonal NAD(P)-linked hydrogenase were amplified with the primers HoxH-f GTATYTGYGGYATTTGTCCTGT and HoxH-r GGCATTTGTCCTRCTGYATGTGT were used. Prior to 40 cycles of the program the DNA was denatured for 5 min at 95°C. The temperature program was as follows: 30 sec at 95°C, 40 sec at 50°C, 2 min at 72°C. In a final step the temperature was kept at 72°C for 10 min. The reaction contained 0.5 µM of the two primers, 0.2 mM of dNTPs, 2.5 mM MgCl2, 0.025 U/µl Taq polymerase (MBI Fermentas, St. Leon-Roth, Germany) and 10x buffer as recommended by the manufacturer in a total volume of 50 µl. Of each sample different amounts of DNA between 2 and 100 ng were tested as template. If no PCR product was detected DNA concentrations were increased at least 10 times. Positive controls were run in parallel to prove the efficiency of the PCR. The approximate size of the product is around 1190 bp and covers close to 84% of the hoxH gene.

The resulting PCR products were ligated into the pCRII-topo (Invitrogen), sequenced with the Big-Dye Kit, and applied on a 96 capillary sequencer (3730 DNA Analyzer, Applied Biosystems).

If possible contigs were assembled from the obtained sequence data and the respective sequences deposited in the genebank (Accession numbers GQ454414 to GQ454443 and GU238237 to GU238258) including two additional cyanobacterial hoxH sequences of Aphanothece halophytica and Mastigocladus laminosus SAG 4.84 (Accession numbers GQ454444 and GQ454445).

Database searches

The genebank, cyanobase, and the GOS database were searched for hydrogenase specific sequences by using the hydrogenase sequences given in Table 3. Retrieved sequences were either run against the genebank by using the BLAST algorithm [38] to deduce the closest homolog or searched for the signature sequences as given by Vignais and Billoud [19] to unambiguously classify the respective hydrogenase. In case of the GOS database, the sequences found were aligned, and, if possible, larger contigs were formed from the same sampling station and used for all further analysis.

Phylogenetic analysis

In the case of critical candidates or unclear phylogenetic affiliation phylogenetic trees were used. Sequence alignments were made with ClustalW [84]. After manual optimization and removal of gaps from the alignments, parsimony, maximum likelihood, and distances were calculated with the 3.63 release of the PHYLIP package [85], using the Jones-Taylor-Thornton matrix and the algorithm of Fitch and Margoliash [86]. Maximum parsimony and distances were calculated for 1000 bootstraps and maximum likelihood for 100 bootstraps. The Unix-cluster at the computer center of the University of Kiel was used for most of the calculations. The resulting trees are given in Fig. S3 to S5 (supporting information).

Supporting Information

Table S1.

Complete list of all marine bacteria searched for hydrogenase genes

(0.05 MB XLS)

Figure S1.

Structure of the gene cluster of the membrane bound hydrogen uptake NiFe-hydrogenase of marine Rhodobacteraceae and the delta-proteobacterium Neptuniibacter caesariensis. The structural genes of the hydrogenase (hupS, hupL and hupZ the membrane bound cytochtrome) are shown in blue. Red genes (hoxAJBC) are involved in the regulation of the hydrogenase. HoxJ encodes a histidine kinase that is known to interact with a hydrogen sensor encoded by hoxB and hoxC and regulates the activity of the response regulator encoded by hoxA. HupK might encode a protein necessary to express an oxygen-tolerant hydrogenase. Accessory genes known to be necessary for this type of membrane hydrogenase are shown in grey, whereas grey patterned genes are general accessory genes for all NiFe-hydrogenases. Genes depicted in green are putative proteases that cleave the C-terminus of the hydrogenase. HypX of Ralstonia eutropha is known to render its soluble hydrogenase oxygen tolerant.

(0.06 MB DOC)

Figure S2.

Structure of three hydrogenase gene clusters of Vibrioanceae isolated from marine environments that are most similar to the energy converting H2-evolving NiFe-hydrogenases. The color code is the same as in Figure S1. Genes shown in plaid are part of the fromate dehydrogenase. FhlA is the transcriptional activator of the formate-hydrogen lyase. Those in black and grey-blue are additional subunits of the whole complex.

(0.05 MB DOC)

Figure S3.

Phylogenetic tree of HypX. Representatives of enoyl-CoA hydratase/crotonase have been used as outgroup. The abbreviations and the respective accession numbers are as follows: Aaeoli, Aquifex aeolicus VF5 NP_213788; Aehrli, Alkalilimnicola ehrlichei MLHE-1 YP_742845; Amarin, Acaryochloris marina MBIC11017 YP_001520946; BjapUSDA, Bradyrhizobium japonicum USDA 110 NP_773566; Cviola, Chromobacterium violaceum ATCC 12472 NP_903812; Daroma, Dechloromonas aromatica RCB YP_287160; Frankia Cc Frankia sp. CcI3 YP_482743; Frankia EA Frankia sp. EAN1pec YP_001505433; MmagAMB, Magnetospirillum magneticum AMB-1 YP_420998; MmagMS-1, Magnetospirillum magnetotacticum MS-1 ZP_00055441; Mmarina Microscilla marina ATCC 23134 ZP_01691397; Mpetro, Methylibium petroleiphilum PM1 YP_001021998; Ncaesar, Neptuniibacter caesariensis ZP_01166042; Nitrati, Nitratiruptor sp. SB155-2 YP_001356445; Pedobac Pedobacter sp. BAL39 ZP_01883353; Pnapht, Polaromonas naphthalenivorans CJ2 YP_982187, PfluPF-5, Pseudomonas fluorescens Pf-5 YP_260772; Pfluore, Pseudomonas fluorescens PfO-1 YP_348856; Reutro, Ralstonia eutropha H16 NP_942660; Rferri, Rhodoferax ferrireducens T118 YP_525330; Rmetalli, Ralstonia metallidurans CH34 YP_583693; Saverm, Streptomyces avermitilis MA-4680 NP_828541; Savermi Streptomyces avermitilis MA-4680 NP_823962; Scoelic Streptomyces coelicolor A3(2) NP_629596; Sdegra, Saccharophagus degradans 2-40 YP_526001; Smalto Stenotrophomonas maltophilia R551-3 YP_002027502; Ssedimi, Shewanella sediminis HAW-EB3 YP_001475080; Sulfuro, Sulfurovum sp. NBC37-1 YP_001358952; Xcamp Xanthomonas campestris pv. vesicatoria str. 85-10 YP_363011

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Figure S4.

Phylogenetic tree of HupL sequences. Representatives of the 49 kDa subunit of the complex I have been used as outgroup. The used abbreviations and their respective accession numbers are as follows: Abac345 Candidatus Koribacter versatilis Ellin345 YP_593314; Abut4018 Arcobacter butzleri RM4018 YP_001490358; Afer53993 Acidithiobacillus ferrooxidans ATCC 53993 YP_002219307; Ahyd7966 Aeromonas hydrophila subsp. hydrophila ATCC 7966 YP_857036; AmacDE Alteromonas macleodii ‘Deep ecotype’ YP_002124659; Aple4074 Actinobacillus pleuropneumoniae serovar 1 str. 4074 ZP_00134404; AsalA449 Aeromonas salmonicida subsp. salmonicida A449 YP_001141617; Asiam Anabaena siamensis TISTR 8012 AAN65266; Avar Anabaena variabilis ATCC 29413 YP_325087; Bac Ellin bacterium Ellin514 ZP_03626632; BBTAi1-2 Bradyrhizobium sp. BTAi1 YP_001220511; BBTAi1-3 Bradyrhizobium sp. BTAi1 YP_001236652; Bjap110 Bradyrhizobium japonicum USDA 110 NP_773581; Bphy815 Burkholderia phymatum STM815 YP_001863308; C.fer13031 Chlorobium ferrooxidans DSM 13031 ZP_01386726; C511412 Cyanothece sp. ATCC 51142 YP_001802481; C7424 Cyanothece sp. PCC 7424 YP_002377118; C7822 Cyanothece sp. PCC 7822 ZP_03153783; C8802 Cyanothece sp. PCC 8802 ZP_03142797; Cagg Chloroflexus aggregans DSM 9485 YP_002461848; Caur10-fl Chloroflexus aurantiacus J-10-fl YP_001636362; CCY0110 Cyanothece sp. CCY 0110 ZP_01728928; Chyd Carboxydothermus hydrogenoformans Z-2901 YP_360377; Cjej1221 Campylobacter jejuni RM1221 YP_179388; Ckos895 Citrobacter koseri ATCC BAA-895 YP_001455880; Clim245 Chlorobium limicola DSM 245 YP_001942914; CmedTB-2 Caminibacter mediatlanticus TB-2 ZP_01871651; Cpha Chlorobium phaeobacteroides DSM 266 YP_911445; CtepTLS Chlorobium tepidum TLS NP_661672; Cwat8501 Crocosphaera watsonii WH 8501 ZP_00519188; Dbac Desulfomicrobium baculatum 1CC1_L; DBAV1 Dehalococcoides sp. BAV1 YP_001213724; Deth Dehalococcoides ethenogenes 195 YP_180861; DvulDP4 Desulfovibrio vulgaris DP4 YP_966691; Ecar1043 Pectobacterium atrosepticum SCRI1043 YP_049334; EcolK12 Escherichia coli str. K-12 substr. MG1655 NP_415492; EcolNuoD Escherichia coli CAA48363; FACN14a Frankia alni ACN14a YP_712616; FACN14a-2 Frankia alni ACN14a YP_712064; Fbac Flavobacteria bacterium MS024-2A ZP_03702421; FCci3 Frankia sp. CcI3 YP_481046; FEAN Frankia sp. EAN1pec YP_001506830; FEAN2 Frankia sp. EAN1pec YP_001507712; Gaur Gemmatimonas aurantiaca T-27 YP_002759924; Gloeo Gloeothece sp. PCC 6909 AAP04005; GlovSZ Geobacter lovleyi SZ YP_001952291; GlovSZ-2 Geobacter lovleyi SZ YP_001950403; HpylJ99 Helicobacter pylori J99 NP_223293; L8106 Lyngbya sp. PCC 8106 ZP_01619041; Laes Lyngbya aestuarii ABD34838; Lint Lawsonia intracellularis PHE/MN1-00 YP_594816; Lmaj Lyngbya majuscula CCAP 1446/4 AAO66476; Mavi Mycobacterium avium 104 YP_881873; MJLS Mycobacterium sp. JLS YP_00107040; Mkan Mycobacterium kansasii ATCC 12478 ZP_04750138; Mmag-1-3 Magnetospirillum magneticum AMB-1 YP_421305; MmagMS-1 Magnetospirillum magnetotacticum MS-1 ZP_00052632; Mmar Mycobacterium marinum M YP_001850173; MMCS Mycobacterium sp. MCS YP_639307; Msil Methylocella silvestris BL2 YP_002364007; Msme Mycobacterium smegmatis str. MC2 155 YP_887053; N7120 Nostoc sp. PCC 7120 NP_484720; N7422 Nostoc sp. PCC 7422 BAE46791; Nazo ‘Nostoc azollae’ 0708 ZP_03768004; Neptuni2 Neptuniibacter caesariensis ZP_01167270; Neptuni 1 Neptuniibacter caesariensis ZP_01166595; Npun Nostoc punctiforme PCC 73102 AAC16277; Nspu Nodularia spumigena CCY 9414 ZP_01628406; Paes Prosthecochloris aestuarii DSM 271 YP_002015547; Pars Pyrobaculum arsenaticum DSM 13514 YP_001153513; Pdis8503 Parabacteroides distasonis ATCC 8503 YP_001303173; Photob34 Photobacterium sp. SKA34 ZP_01160131; Pisl Pyrobaculum islandicum DSM 4184 YP_929722; Plut Pelodictyon luteolum DSM 273 YP_375349; PMED4NdH Prochlorococcus marinus subsp. pastoris str. CCMP1986 NP_892293; Ppha Pelodictyon phaeoclathratiforme BU-1 YP_002018704; Rcap Rhodobacter capsulatus AAA69668; Rcas Roseiflexus castenholzii DSM 13941 YP_001433219; Rcas2 Roseiflexus castenholzii DSM 13941 YP_001433862; ReryPR4 Rhodococcus erythropolis PR4 YP_002766098; RerySK121 Rhodococcus erythropolis SK121 ZP_04384689; Reut Ralstonia eutropha H16 NP_942704; ReutC Ralstonia eutropha H16 NP_942663; ReutG Ralstonia eutropha H16 AAA16462; Rgel Methylibium petroleiphilum PM1 YP_001022015; RHTCC2501 Robiginitalea biformata HTCC2501 ZP_01119574; Rhtcc2601 Roseovarius sp. HTCC2601 ZP_01443057; RHTCC2601-Sens Roseovarius sp. HTCC2601 ZP_01443054; Rjos Rhodococcus jostii RHA1 YP_704548; Ropa Rhodococcus opacus B4 YP_002781742; Rpal009 Rhodopseudomonas palustris CGA009 NP_946314; RpalA53 Rhodopseudomonas palustris BisA53 YP_780164; RpalB5 Rhodopseudomonas palustris BisB5 YP_568300; RRS-1 Roseiflexus sp. RS-1 YP_001276649; Rrub Rhodospirillum rubrum ATCC 11170 YP_426250; Rsph17029 Rhodobacter sphaeroides ATCC 17029 YP_001044019; Rsph2.4.1 Rhodobacter sphaeroides 2.4.1 YP_353568; Rtm1035 Roseovarius sp. TM1035 ZP_01881109; Sag12614 Stappia aggregata IAM 12614 ZP_01550392; Sag12614-2 Stappia aggregata IAM 12614 ZP_01550270; Sala2256 Sphingopyxis alaskensis RB2256 YP_611130; Sama Shewanella amazonensis SB2B YP_927554; Save Streptomyces avermitilis MA-4680 NP_828543; SbalOS155 Shewanella baltica OS155 YP_001050263; Sdys197 Shigella dysenteriae Sd197 YP_402612; SentATCC Salmonella enterica subsp. enterica serovar Paratyphi A str. ATCC 9150 YP_152163; SentCT18 Salmonella enterica subsp. enterica serovar Typhi str. CT18 NP_456296; SfumMPOB Syntrophobacter fumaroxidans MPOB YP_847061; Slin Spirosoma linguale DSM 74 ZP_04492490; SoneMR-1 Shewanella oneidensis MR-1 NP_717701; SoneMR-4 Shewanella sp. MR-4 YP_733952; SoneMR-7 Shewanella sp. MR-7 YP_738201; Sros Streptosporangium roseum DSM 43021 ZP_04474993; Sste37 Sagittula stellata E-37 ZP_01748533; Ssvi Streptomyces sviceus ATCC 29083 YP_002204206; Susi Solibacter usitatus Ellin6076 YP_827763; Svir Saccharomonospora viridis DSM 43017 ZP_04507584; TcarNor1 Thermosinus carboxydivorans Nor1 ZP_01667576; Tden25259 Thiobacillus denitrificans ATCC 25259 YP_315133; Tden33889 Sulfurimonas denitrificans DSM 1251 YP_393947; Tery Trichodesmium erythraeum IMS101 YP_722943; TM1035-Sens Roseovarius sp. TM1035 ZP_01881113; Tros 5159 Thermomicrobium roseum DSM 5159 YP_002523076; Tros2 Thiocapsa roseopersicina AAA27410; Tros Thiocapsa roseopersicina AAC38282; Ucyn-A Cyanothece sp. CCY 0110 ZP_01728928; VangS14 Vibrio angustum S14 ZP_01234606; Wsuc1740 Wolinella succinogenes DSM 1740 NP_907813; Yent8081 Yersinia enterocolitica subsp. enterocolitica 8081 YP_001007767. The sequence of the marine unicellular group A cyanobacteria has been generated from the available short reads [70].

(0.67 MB DOC)

Figure S5.

Distribution of small subunits of the bidirectional NAD(P)+ linked hydrogenase found in the GOS database of the different prokaryotic groups. The small subunit gene, hoxY, of Synechocystis has been used for the search. All genes have been retrieved form Punta Comorant, a hypersaline pond on the Galapagos Islands.

(0.06 MB DOC)

Figure S6.

Distribution of small subunits of the membrane bound H2 uptake hydrogenasses found in the GOS database of the different prokaryotic groups. The hupS sequence of Desulfovibrio vulgaris was used for the search. On the right the number of sequences from the different sampling stations is shown.

(0.08 MB DOC)

Figure S7.

Distribution of small subunits of the cyanobacterial-like uptake hydrogenase found in the GOS database of the different prokaryotic groups. The small subunit gene, hupS, of Nostoc sp. PCC 7120 has been used for the search.

(0.05 MB DOC)

Figure S8.

Phylogenetic tree of HoxH sequences. Representatives of the 49 kDa subunit of the complex I have been used as outgroup. The used abbreviations and their respective accession numbers are as follows: Afla Acetomicrobium flavidum CAA56464; Ahalo Aphanothce halophytica GQ454444; Amar Acaryochloris marina MBIC11017 YP_001521996; Amax Arthrospira maxima FACHBSM AAQ63961; Apla1 Arthrospira platensis FACHB341 AAQ63964; Apla2 Arthrospira platensis FACHBOUQDS6 AAQ63959; Apla3 Arthrospira platensis FACHB439 AAQ63960; Apla4 Arthrospira platensis FACHB791 AAQ91344; Avar Anabaena variabilis ATCC 29413 YP_325153; Bxen Burkholderia xenovorans LB400 YP_555781; Cagg Chloroflexus aggregans DSM 9485 YP_002463784; CaggL Chlorobium chlorochromatii CaD3 YP_378564; Caur Chloroflexus aurantiacus J-10-fl YP_001634807; CCY0110 Cyanothece sp. CCY 0110 ZP_01727423; ClimL Chlorobium limicola DSM 245 YP_001944104; Cnec Ralstonia eutropha H16 NP_942730; CphaL Chlorobium phaeobacteroides DSM 266 YP_912598;CtepL Chlorobium tepidum TLS NP_662771; Daro Dechloromonas aromatica RCB YP_284208; DethV Dehalococcoides ethenogenes 195 YP_181357; Dpsy Desulfotalea psychrophila LSv54 YP_065948; DpsyV Desulfotalea psychrophila LSv54 YP_064749;Ecol Escherichia coli CAA48363; Galp Gloeocapsa alpicola str. CALU 743 AAO85440; Gmet1 Geobacter metallireducens GS-15 YP_384078; Gmet2 Geobacter metallireducens GS-15 YP_386258; GOS1 and GOS2 are the two consenus sequeces retrieved from the GOS database; Gsul1 Geobacter sulfurreducens PCA NP_953465; Gsul2 Geobacter sulfurreducens PCA NP_953763; Lyng Lyngbya majuscula CCAP 1446/4 AAT07678; Magneto Magnetococcus sp. MC-1 YP_864809; Mastigo Mastigocladus laminosus SAG 4.84 GQ454445; Mcap Methylococcus capsulatus str. Bath YP_112653; MferV Methanothermus fervidus Q49179; MjanV Methanocaldococcus jannaschii DSM 2661 NP_248187; Mkan Methanopyrus kandleri AV19 NP_613553; Mmag Magnetospirillum magnetotacticum MS-1 ZP_00053777; MmarV Methanococcus maripaludis S2 NP_987943;MvolV1 Methanococcus voltae Q00404; MvolV2 Methanococcus voltae Q00407;N7120 Nostoc sp. PCC 7120 NP_484809; N7422 Nostoc sp. PCC 7422 BAE46796; Neptuni Oceanospirillum sp. MED92 ZP_01164927; Nitrococcus Nitrococcus mobilis Nb-231 ZP_01126922; Nspu Nodularia spumigena CCY 9414 ZP_01629499; Nspu Nodularia spumigena CCY 9414 ZP_01629499; PaesL Prosthecochloris aestuarii DSM 271 YP_002016588; PfurL1 Pyrococcus furiosus DSM 3638 NP_578623; PfurL2 Pyrococcus furiosus DSM 3638 NP_579061; Phol Prochlorothrix hollandica AAB53705;Plancto Planctomyces maris DSM 8797 ZP_01852867; PMED4 Prochlorococcus marinus subsp. pastoris str. CCMP1986 NP_892293; PphaL Pelodictyon phaeoclathratiforme BU-1 YP_002019299; Rcas Roseiflexus castenholzii DSM 13941 YP_001431482; Rmet Ralstonia metallidurans CH34 YP_583677; Ropa Rhodococcus opacus AAB57892; RRS-1 Roseiflexus sp. RS-1 YP_001277847; S6301 Synechococcus elongatus PCC 6301 YP_172265; S6803 Synechocystis sp. PCC 6803 NP_441259; S6803 Synechocystis sp. PCC 6803 NP_441411;S7002 Synechococcus sp. PCC 7002 YP_001733469; S7942 Synechococcus elongatus PCC 7942 YP_401572; Spla Arthrospira platensis FACHB440 AAQ63963; Ssub Spirulina subsalsa FACHB351 AAQ63962; Susi Solibacter usitatus Ellin6076 YP_826256;Tros Thiocapsa roseopersicina AAP50523; WH5701 Synechococcus sp. WH 5701 ZP_01085930.

(0.13 MB DOC)


We are thankful for the help from the crews on the research vessels Polarfuchs (Institut für Meereskunde, Kiel) and the captain Uwe Becker of the Südfall and Andreas Ruser from FTZ (Büsum) and Christoph Keller and Conny Schmidt and Fischerei Reese while taking samples on Selenter See. Special thanks are to Jonathan Zehr and Jim Tripp for providing sequence reads of the unicellular N2-fixing cyanobacteria in "group A".

Author Contributions

Conceived and designed the experiments: RS JL RS JA. Performed the experiments: MB CB TS FG FO CM. Analyzed the data: MB CB TS FG FO CM CS KG JA. Contributed reagents/materials/analysis tools: KHV RS JL RS. Wrote the paper: JA.


  1. 1. Novelli PC, Lang PM, Masarie KA, Hurst DF, Myers R, Elkins JW (1999) Molecular hydrogen in the troposphere: global distribution and budget. J Geophys Res 104: 30427–30444.
  2. 2. Ehhalt DH, Rohrer F (2009) The tropospheric cycle of H2: a critical review. Tellus 61B: 500–535.
  3. 3. Price H, Jaegle L, Rice A, Quay P, Novelli PC, et al. (2007) Global budget of molecular hydrogen and its deuterium content: Constraints from ground station, cruise, and aircraft observations. J Geophys Res Atmospheres 112: D22108.
  4. 4. Conrad R, Seiler W (1980) Contribution of hydrogen production by biological nitrogen fixation to the global hydrogen budget. J Geophys. Res 85: 5493–5498.
  5. 5. Rhan T, Eiler JM, Boering KA, Wennberg PO, McCarthy MC, et al. (2003) Extreme deuterium enrichment in the stratospheric hydrogen and the global atmospheric budget of H2. Nature 424: 918–921.
  6. 6. Conrad R (1996) Soil microorganisms as controllers of atmospheric trace gases (H2, CO, CH4, OCS, N2O, and NO). Mol Microbiol Rev 60: 609–640.
  7. 7. Scranton MI, Jones MM, Herr FL (1982) Distribution and variability of dissolved hydrogen in the Mediterranean Sea. J Mar Res 40: 873–891.
  8. 8. Herr FL, Frank EC, Leone GM, Kennicutt MC (1984) Diurnal variability of dissolved molecular hydrogen in the tropical South Atlantic ocean. Deep Sea Res 31: 13–20.
  9. 9. Conrad R, Seiler W (1988) Methane and hydrogen in seawater (Atlantic Ocean). Deep Sea Res 35: 1903–1917.
  10. 10. Herr FL, Scranton MI, Barger WR (1981) Dissolved hydrogen in the Norvegian sea: mesoscale surface variability and deep water distribution. Deep Sea Res 28: 1001–1016.
  11. 11. Punshon S, Moore RM, Xie H (2007) Net loss rates and distribution of molecular hydrogen (H2) in mid-latitude coastal waters. Mar Chem 105: 129–139.
  12. 12. Schmidt U, Conrad R (1993) Hydrogen, carbon monoxide, and methane dynamics in Lake Constance. Limnol Oceanogr 38: 1214–1226.
  13. 13. Conrad R, Aragno M, Seiler W (1983) Production and consumption of hydrogen in a eutrophic lake. Appl Environ Microbiol 45: 502–510.
  14. 14. Schütz H, Conrad R, Goodwin S, Seiler W (1988) Emission of hydrogen from deep and shallow freshwater environments. Biogeochem 5: 295–311.
  15. 15. Lindberg P, Lindblad P, Cournac L (2004) Gas exchange in the filamentous cyanobacterium Nostoc punctiforme strain ATCC 29133 and its hydrogenase-deficient mutant strain NHM5. Appl Environ Microbiol 70: 2137–2145.
  16. 16. Scranton MI, Novelli PC, Michaels A, Horrigan SG, Carpenter EJ (1987) Hydrogen production and nitrogen fixation by Oscillatoria thiebautii during in situ incubations. Limnol Oceangr 32: 998–1006.
  17. 17. Punshon S, Moore RM (2008) Photochemical production of molecular hydrogen in lake water and coastal seawater. Mar Chem 108: 215–220.
  18. 18. Schwartz E, Friedrich B Dworkin M, Falkow S, editors. (2005) The H2-metabolising prokaryotes. 2: A Handbook of the Biology of Bacteria: Ecophysiology and Biochemistry. 496–563.
  19. 19. Vignais PM, Billoud B (2007) Occurrence, classification and biological function of hydrogenases: an overview. Chem Rev 107: 4206–4272.
  20. 20. Vignais PM, Billoud B, Meyer J (2001) Classification and phylogeny of hydrogenases. FEMS Microbiol Rev 25: 455–501.
  21. 21. Burgdorf T, Lenz O, Buhrke T, van der Linden E, Jones AK, Albracht SPJ, Friedrich B (2005) [NiFe]-hydrogenases of Ralstonia eutropha H16: Modular enzymes for oxygen-tolerant biological hydrogen oxidation. J Mol Microbiol Biotech 10: 181–196.
  22. 22. Ballantine SP, Boxer DH (1986) Isolation and characterization of a soluble active fragment of hydrogenase isoenzyme 2 from the membranes of anaerobically grown Escherichia coli. Eur J Biochem 156: 277–284.
  23. 23. Lukey MJ, Parkin A, Roessler MM, Murphy BJ, Harmer J, et al. (2010) How Escherichia coli is equipped to oxidize hydrogen under different redox conditions. J Biol. Chem 285: 3928–3938.
  24. 24. Sawers RG (2005) Formate and its role in hydrogen production in Escherichia coli. Biochem Soc Trans 33: 42–46.
  25. 25. Hedderich R, Forzi L (2005) Energy-converting [NiFe] hydrogenases: more than just H2 activation. J Mol. Microbiol. Biotechnol 10: 92–104.
  26. 26. Ng G, Tom CG, Park AS, Zenad L, Ludwig RA (2009) A novel endo-hydrogenase activity recycles hydrogen produced by nitrogen fixation. PLoS One 3: e4695.
  27. 27. Lechner S, Conrad R (1997) Detection in soil of aerobic hydrogen-oxidizing bacteria related to Alcaligenes eutrophus by PCR and hybridization assays targeting the gene of the membrane bound (NiFe) hydrogenase. FEMS Microbiol. Ecol 22: 193–206.
  28. 28. Constant P, Chowdhury SP, Pratscher J, Conrad R (2010) Streptomycetes contributing to atmospheric molecular hydrogen soil uptake are widespread and encode a putative high-affinity [NiFe]-hydrogenase. Environ Microbiol. 12. : 821–829.
  29. 29. Boyd ES, Spear JR, Peters JW (2009) [FeFe] hydrogenase genetic diversity provides insight into molecular adaptation in a saline microbial mat community.Appl. Environ. Microbiol 75: 4620–4623.
  30. 30. Boyd ES, Hamilton TL, Spear JR, Lavin M, Peters JW (2010) [FeFe]-hydrogenase in Yellowstone National Park: evidence for dispersal limitation and phylogenetic niche conservatism. ISME J 2010 Jun 10 10: [Epub ahead of print].
  31. 31. Cummings L, Riley L, Black L, Souvorov A, Resenchuk S, et al. (2002) Genomic BLAST: custom-defined virtual databases for complete and unfinished genomes. FEMS Microbiol Lett 216: 133–138.
  32. 32. Venter JC, Remington K, Heidelberg JF, Halpern AL, Rusch D, et al. (2004) Environmental genome shotgun sequencing of the Sargasso Sea. Science 304: 66–74.
  33. 33. Rusch DB, Halpern AL, Sutton G, Heidelberg KB, Williamson S, et al. (2007) The Sorcerer II Global Ocean Sampling expedition: Northwest Atlantic through eastern tropical Pacific. PLoS Biol 5(3): e77.
  34. 34. Yooseph S, Sutton G, Rusch DB, Halpern AL, Williamson SJ, et al. (2007) The Sorcerer II Global Ocean Sampling expedition: Expanding the universe of protein families. PLoS Biol 5: e16.
  35. 35. Appel J, Schulz R (1998) Hydrogen metabolism in organisms with oxygenic photosynthesis - hydrogenases as important regulatory devices for a proper redox poising? J Photochem Photobiol B: Biol 47: 1–11.
  36. 36. Tamagnini P, Leitao E, Oliveira P, Ferreira D, Pinto F, Harris DJ, Heidorn TT, Lindblad P (2008) Cyanobacterial hydrogenases: diversity, regulation and applications. FEMS Microbial Rev 31: 692–720.
  37. 37. Ludwig M, Schulz-Friedrich R, Appel J (2006) Occurrence of hydrogenases in cyanobacteria and anoxygenic photosynthetic bacteria: implications for the phylogenetic origin of cyanobacterial and algal hydrogenases. J Mol Evol 63: 758–768.
  38. 38. Altschul SF, Madden TL, Schaffer AA, Zhang J, Zhang Z, et al. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402.
  39. 39. Gutekunst K, Hoffmann D, Lommer M, Egert M, Suzuki I, et al. (2006) Metal dependence and intracellular regulation of the bidirectional NiFe-hydrogenase in Synechocystis sp PCC 6803. Int J Hydr. Energ 31: 1452–1459.
  40. 40. Tripp HJ, Bench SR, Turk KA, Foster RA, Desany BA, Niazi F, Affourtit JP, Zehr JP (2010) Metabolic streamlining in an open-ocean nitrogen-fixing cyanobacterium. Nature 464: 90–94.
  41. 41. Fontecilla-Camps JC, Volbeda A, Cavazza C, Nicolet Y (2007) Structure/function relationship of [NiFe]- and [FeFe]-hydrogenases. Chem Rev 107: 4273–4303.
  42. 42. Sorokin YI (2002) The Black Sea: Ecology and Oceanography, Backhuys Publishers, Leiden, The Netherlands.
  43. 43. Lenz O, Friedrich B (1998) A novel multicomponent regulatory system mediates H2 sensing in Alcaligenes eutrophus. Proc Natl Acad Sci USA 95: 12474–12479.
  44. 44. Elsen S, Duché O, Colbeau A (2003) Interaction between the H2 sensor HupUV and the histidine kinase HupT controls HupSL hydrogenase synthesis in Rhodobacter capsulatus. J Bacteriol 185: 7111–7119.
  45. 45. Buhrke T, Lenz O, Porthun A, Friedrich B (2004) The H2-sensing complex of Ralstonia eutropha: interaction between a regulatory [NiFe] hydrogenase and a histidine protein kinase. Mol Microbiol 51: 1677–1689.
  46. 46. Vignais PM, Elsen S, Colbeau A (2005) Transcriptional regulation of the uptake [NiFe]hydrogenase genes in Rhodobacter capsulatus. Biochem Soc Trans 33: 28–32.
  47. 47. Bernhard M, Schwartz E, Rietdorf J, Friedrich B (1996) The Alcaligenes eutrophus membrane-bound hydrogenase gene locus encodes functions involved in maturation and electron transport coupling. J Bacteriol 178: 4522–4529.
  48. 48. Ludwig M, Schubert T, Zebger I, Wisitruangsakul N, Saggu M, et al. (2009) Concerted action of two novel auxiliary proteins in assembly of the active site in a membrane-bound [NiFe]-hydrogenase. J Biol Chem 284: 2159–2168.
  49. 49. Redwood MD, Mikheenko IP, Sargent F, Macaskie LE (2008) Dissecting the roles of Escherichia coli hydrogenases in biohydrogen production. FEMS Microbiol Lett 278: 48–55.
  50. 50. Arahal DR, Lekunberri I, Gonzalez JM, Pascual J, Pujalte MJ, Pedros-Alio C, Pinhassi J (2007) Neptuniibacter caesariensis gen. nov., sp. nov., a novel genome-sequenced gamma proteobacterium. Int J Syst Evol Microbiol 57: 1000–1006.
  51. 51. Buhrke T, Friedrich B (1998) hoxX (hypX) is a functional member of the Alcaligenes eutrophus hyp gene cluster. Arch Microbiol 170: 460–463.
  52. 52. Whitman WB, Coleman DC, Wiebe WJ (1998) ) Prokaryotes: The unseen majority. Proc Natl Acad Sci USA 95: 6578–6583.
  53. 53. Giovannoni SJ, Stingl U (2005) Molecular diversity and ecology of microbial plankton. Nature 437: 343–348.
  54. 54. Pedros-Alio C (2006) Marine microbial diversity: can it be determined? Trends Microbiol 14: 257–263.
  55. 55. DeLong EF (2005) Microbial community genomics in the ocean. Nat Rev Microbiol 3: 459–469.
  56. 56. Yutin N, Suzuki MT, Teeling H, Weber M, Venter JC, Rusch DB, Beja O (2007) Assessing diversity and biogeography of aerobic anoxygenic phototrophic bacteria in surface waters of the Atlantic and Pacific oceans using the global ocean sampling expedition metagenomes. Environ Microbiol 9: 1464–1475.
  57. 57. Poretsky RS, Hewson I, Sun S, Allen AE, Zehr JP, Moran MA (2009) Comparative day/night metatranscriptomic analysis of microbial communities in the North Pacific subtropical gyre. Environ Microbiol 11: 1358–1375.
  58. 58. Appel J, Schulz R (1996) Sequence analysis of an operon of a NAD(P)-reducing nickel hydrogenase from the cyanobacterium Synechocystis sp. PCC 6803 gives additional evidence for direct coupling of the enzyme to NAD(P)H-dehydrogenase (complex I). Biochim Biophys Acta 1298: 141–147.
  59. 59. Schmitz O, Bothe H (1996) The diaphorase subunit HoxU of the bidirectional hydrogenase as electron transferring protein in cyanobacterial respiration? Naturwissenschaften 83: 525–527.
  60. 60. Troshina O, Serebryakova L, Shermetieva M, Lindblad P (2002) Production of H2 by the unicellular cyanobacterium Gloeocapsa alpicola CALU 743 during fermentation. Int J Hydr Energ 27: 1283–1289.
  61. 61. Appel J, Phunpruch S, Steinmüller K, Schulz R (2000) The bidirectional hydrogenase of Synechocystis sp. PCC 6803 works as an electron valve during photosynthesis. Arch Microbiol 173: 333–338.
  62. 62. Cournac L, Mus F, Bernard L, Guedeney G, Vignais PM, Peltier G (2002) Limiting steps of hydrogen production in Chlamydomonas reinhardtii and Synechocystis PCC 6803 as analysed by light-induced gas exchange transients. Int J Hydrog Energy 27: 1229–1237.
  63. 63. Cournac L, Guedeney G, Peltier G, Vignais PM (2004) Sustained photoevolution of molecular hydrogen in a mutant of Synechocystis sp. Strain PCC 6803 deficient in the type I NADPH-dehydrogenase complex, J Bacteriol 186: 1737–1746.
  64. 64. Gutthann F, Egert M, Marques A, Appel J (2007) Inhibition of respiration and nitrate assimilation enhances hydrogen evolution in Synechocystis sp. PCC 6803. Biochem Biophys Acta 1767: 161–169.
  65. 65. Hoehler TM, Bebout BM, DesMarais DJ (2001) The role of microbial mats in the production of reduced gases on the early Earth. Nature 412: 324–327.
  66. 66. Vincent KA, Cracknell JA, Clark JR, Ludwig M, Lenz O, et al. (2006) electricity from low-level H2 in still air – an ultimate test for an oxygen tolerant hydrogenase. Chem Comm 5033–5035.
  67. 67. Cracknell JA, Vincent KA, Ludwig M, Lenz O, Friedrich B, Armstrong FA (2008) Enzymatic oxidation of H2 in atmospheric O2: the electrochemistry of energy generation from trace H2 by aerobic microorganisms. J Am Chem Soc 130: 424–425.
  68. 68. Gonzalez JM, Moran MA (1997) Numerical dominance of a group of marine bacteria in the α-subclass of the class proteobacteria in coastal seawater. Appl Environ Microbiol 63: 4237–4242.
  69. 69. King GM (2003) Uptake of carbon monoxide and hydrogen at environmentally relevant concentrations by mycobacteria. Appl Environ Microbiol 69: 7266–7272.
  70. 70. Morita RY (2000) Is H2 the universal energy source for long-term survival? Microb Ecol 38: 307–320.
  71. 71. King GM, Weber CF (2007) Distribution, diversity and ecology of aerobic CO-oxidizing bacteria. Nat Rev Microbiol 5: 107–117.
  72. 72. Moran MA, Miller WL (2007) Resourceful heterotrophs make the most of light in the coastal ocean. Nat Rev Microbiol 5: 792–800.
  73. 73. Ploug H, Kuhl M, Buchholz-Cleven B, Jorgensen BB (1997) Anoxic aggregates - an ephemeral phenomenon in the pelagic environment? Aquatic Micro Ecol 13: 285–294.
  74. 74. Maróti G, Tong Y, Yooseph S, Baden-Tillson H, Smith HO, et al. (2009) Discovery of [NiFe] hydrogenase genes in metagenomic DNA: cloning and heterologous expression in Thiocapsa roseopersicina. Appl. Environ. Microbiol 75: 5821–5830.
  75. 75. Iars-Martinez E, Martin-Cuadrado AB, D'Auria G, Mira A, Ferriera S, Johnson J, Friedman R, Rodriguez-alera F (2008) Comparative genomics of two ecotypes of the marine planktonic copiotroph Altermononas macleodii suggests alternative lifestyles associated with different kinds of particulate organic matter. ISME J 2: 1194–1212.
  76. 76. Mitsui A, Suda S (1995) Alternative and cyclic appearance of H2 and O2 photoproduction activities under nongrowing conditions in an aerobic nitrogen fixing unicellular cyanobacterium Synechococcus sp. Curr Microbiol 30: 1–6.
  77. 77. Ortegacalvo JJ, Stal LJ (1991) Diazotrophic growth of the unicellular cyanobacterium Gloeothece sp. PCC 6909 in continuous culture. J Gen Microbiol 137: 1789–1797.
  78. 78. Zehr JP, Waterbury JB, Turner PJ, Montoya JP, Omoregie E, et al. (2001) Unicellular cyanobacteria fix N2 in the subtropical North Pacific Ocean. Nature 412: 635–638.
  79. 79. Montoya JP, Holl CM, Zehr JP, Hansen A, Villareal TA, et al. (2004) High rates of N2 fixation by unicellular diazotrophs in the oligotrophic Pacific Ocean. Nature 430: 1027–1031.
  80. 80. Zehr JP, Bench SR, Carter BJ, Hewson I, Niazi F, et al. (2008) Globally distributed uncultivated oceanic N2-fixing cyanobacteria lack oxygenic photosystem II. Science 322: 1110–1112.
  81. 81. Langlois RJ, Hummer D, LaRoche J (2008) Abundances and Distributions of the Dominant nifH Phylotypes in the Northern Atlantic Ocean. Appl Environ Microbiol 74: 1922–1931.
  82. 82. Fong AA, Karl DM, Lukas R, Letelier RM, Zehr JP, et al. (2008) Nitrogen fixation in an anticyclonic eddy in the oligotrophic North Pacific Ocean. ISME J 2: 663–76.
  83. 83. Paulmier A, Ruiz-Pino D (2009) Oxygen minimum zones (OMZs) in the modern ocean. Prog Oceanogr 80: 113–128.
  84. 84. Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22: 4673–4680.
  85. 85. Felsenstein J (2005) PHYLIP (Phylogeny Inference Package), version 3.6, Distributed by the author, Department of Genome, Sciences, University of Washington, Seattle.
  86. 86. Fitch WM, Margoliash E (1967) Construction of phylogenetic trees. Science 155: 279–284.