Respiratory membrane endo-hydrogenase activity in the microaerophile Azorhizobium caulinodans is bidirectional.

BACKGROUND: The microaerophilic bacterium Azorhizobium caulinodans, when fixing N(2) both in pure cultures held at 20 µM dissolved O(2) tension and as endosymbiont of Sesbania rostrata legume nodules, employs a novel, respiratory-membrane endo-hydrogenase to oxidize and recycle endogenous H(2) produced by soluble Mo-dinitrogenase activity at the expense of O(2). METHODS AND
FINDINGS: From a bioinformatic analysis, this endo-hydrogenase is a core (6 subunit) version of (14 subunit) NADH:ubiquinone oxidoreductase (respiratory complex I). In pure A. caulinodans liquid cultures, when O(2) levels are lowered to <1 µM dissolved O(2) tension (true microaerobic physiology), in vivo endo-hydrogenase activity reverses and continuously evolves H(2) at high rates. In essence, H(+) ions then supplement scarce O(2) as respiratory-membrane electron acceptor. Paradoxically, from thermodynamic considerations, such hydrogenic respiratory-membrane electron transfer need largely uncouple oxidative phosphorylation, required for growth of non-phototrophic aerobic bacteria, A. caulinodans included.
CONCLUSIONS: A. caulinodans in vivo endo-hydrogenase catalytic activity is bidirectional. To our knowledge, this study is the first demonstration of hydrogenic respiratory-membrane electron transfer among aerobic (non-fermentative) bacteria. When compared with O(2) tolerant hydrogenases in other organisms, A. caulinodans in vivo endo-hydrogenase mediated H(2) production rates (50,000 pmol 10(9)·cells(-1) min(-1)) are at least one-thousandfold higher. Conceivably, A. caulinodans respiratory-membrane hydrogenesis might initiate H(2) crossfeeding among spatially organized bacterial populations whose individual cells adopt distinct metabolic states in response to variant O(2) availability. Such organized, physiologically heterogeneous cell populations might benefit from augmented energy transduction and growth rates of the populations, considered as a whole.

Introduction

Given the relatively low (−414 mV) potential in aqueous solution for the standard hydrogen bio-electrochemical half-cell, hydrogen gas (H2) is a strong electron (e–) donor, whereas in the back-reaction, combining H+ ions are weak e– acceptors. Hydrogenases, which catalyze this reaction, are widely distributed among bacteria [1]. Diverse aerobic bacteria employ O2-tolerant (group-1) hydrogenases as e– donor, oxidizing substrate H2 at the expense of substrate O2 as preferred e– acceptor, driving oxidative phosphorylation. These group-1 hydrogenases, both soluble and membrane-associated, typically include globular, heterodimeric catalytic proteins. In the stably membrane-associated group-1 hydrogenases, each catalytic heterodimer is first exported and then stably complexes with a membrane-integral diheme b-type cytochrome [2]. Resulting heterotrimeric complexes are typically denoted ‘uptake hydrogenases’ as they operate in vivo as unidirectional catalysts of H2 oxidation. Product H+ ions, released on the exterior (exo) face of cell membranes, directly contribute to trans-membrane proton-motive force absolving these activities of any, obvious chemiosmotic (ion-pumping) workload. These stably membrane-associated, group-1 heterotrimeric complexes may be termed exo-hydrogenases.

The reversible or H2 evolving (group-4) hydrogenases, also membrane-associated, are encoded by completely divergent gene-sets. The group-4 hydrogenases are typically employed by anaerobic bacteria to produce H2 as fermentative end-product and in so doing, facilitate overall cellular oxidation-reduction balance [1]. In anaerobes, the hydrogenesis (H2 production from H+ ions) reaction involves direct coupling of group-4 hydrogenases as e– acceptor with various e– donors such as formate and carbon monoxide dehydrogenases as membrane-integral complexes [3], [4]. In contrast to group-1 membrane-associated uptake hydrogenases, the catalytic heterodimers of group-4 hydrogenases are oriented to the cytosolic (endo) face of cell membranes [5], [6] and so may be termed endo-hydrogenases.

However, endo-hydrogenases are not exclusive to fermentative anaerobes. We recently reported on a novel endo-hydrogenase in the aerobic microaerophile Azorhizobium caulinodans which requires oxidative phosphorylation for growth. Indeed, A. caulinodans employs both membrane-associated exo- and endo-hydrogenases when respiring with H2 as e– donor. In chemolithotropic cultures with exogenous H2 as sole energy source, A. caulinodans primarily relies on exo-hydrogenase activity [7]. Archetype A. caulinodans strain ORS571 was originally isolated as N2-fixing endosymbiont of stem- and root- nodules in Sesbania rostrata, an annual legume indigenous to the Atlantic coastal Sahel [8]. A. caulinodans ORS571 may be cultured diazotrophically (N2 as sole N-source) and organotrophically (oxidizable organic acids as C- and energy source) under a reduced (2%) atmosphere [9]. Its sole N2 fixing activity, Mo-dinitrogenase, also produces stoichiometric H2 in an ATP-dependent process [10], [11]. In such diazotrophic liquid cultures, respiratory-membrane uptake hydrogenase activity allows input of endogenous H2 as fuel for oxidative phosphorylation, recovering invested ATP. In contrast to use of exogenous H2, in endogenous H2 uptake, endo-hydrogenase activity predominates [7]. In Sesbania rostrata (legume) nodules actively fixing N2, A. caulinodans endosymbionts employ both exo- and endo-hydrogenases to recycle endogenous H2 produced by Mo-dinitrogenase activity [12]. In these cases, both exo- and endo-hydrogenases function as uptake hydrogenases.

However, as we demonstrate here, A. caulinodans endo-hydrogenase in vivo activity is bidirectional, reversing in response to physiological O2 availability. Given sufficient O2, endo-hydrogenase operates in H2 uptake mode. Under strict O2 limitation, endo-hydrogenase reverses and operates in hydrogenesis mode at extraordinarily high in vivo rates. Hitherto, endo-hydrogenase mediated hydrogenesis has been masked as it occurs in N2 fixing pure cultures in which Mo-dinitrogenase activity itself also produces H2 [10], [11]. Because exo-hydrogenase invariably operates in H2 uptake mode, exo- and endo-hydrogenases then function at cross-purposes, yielding a novel and seemingly paradoxical physiology.

Results

Hyq endo-hydrogenase is a Core Homolog of L-type Respiratory Complex I

The Azorhizobium caulinodans hyq operon (Entrez Gene identifier: AZC4360–AZC4355) encodes an endo-hydrogenase including six discrete structural proteins as well as a transcriptional activator [7]. From SUPERFAMILY analysis, a hidden Markov model library of protein structures [13], the six catalytic A. caulinodans Hyq proteins all have close homologs among the Nuo proteins of NADH:quinone oxidoreductase, commonly referred to as ‘L-type’ respiratory complex I (Table 1). Bacterial respiratory complex I typically includes 14 subunits equally divided into membrane-integral (LO) and cytosol-interfacing, membrane-peripheral (L1) subcomplexes [14], [15]. In pairwise primary amino acid sequence alignments, four Hyq proteins (HyqBCEF) and four LO subcomplex NuoHJLM proteins are ∼60% conserved (Table 1; Figs. S1, S2, S3, S4, S5, S6). For the L1 subcomplexes, two (HyqGI) proteins are homologs of three (NuoCDB) proteins. From SUPERFAMILY analysis, HyqG corresponds to a fused NuoC::D (SSF56762). The HyqG (504 residues) N-terminal domain (residues 1–156) is homologous to NuoC, and its C-terminal domain (residues 157–504) is homologous to NuoD. Because its (SSF56762) superfamily also includes the group-1 hydrogenase catalytic (large) subunit, HyqG together with HyqI presumably catalyze hydrogenase activity. HyqI, a small FeS protein, is a NuoB (SSF56770) homolog; all HyqI orthologs show conserved cys-55, cys-58 (Cys-X-X-Cys), cys-112 and cys-152 residues likely coordinating a N2-type, high-potential 4Fe4S center, which in respiratory complex I serves as immediate e– donor to membrane quinone [16], [17]. The binding site for complex I membrane quinone, its e– acceptor, is a cavity formed between a four-helix bundle of NuoD, the H1 helix of NuoB, and transmembrane helix 1 of NuoH [14], [15], [18], all of which elements are conserved in Hyq endo-hydrogenases (Table 1; Figs. S1, S2, S3, S4, S5, S6). By inference, the Hyq endo-hydrogenase of microaerophiles constitutes a core L-type H2:ubiquinone oxidoreductase (Fig. 1).

Table 1

A. caulinodans Nuo (NADH:quinone oxidoreductase) and Hyq (endo-hydrogenase) structural homologs.

A. caulinodans complex I	EntrezGene identifier	T. thermophilus complex I	A. caulinodans hydrogenase	EntrezGene identifier	Identity††%	Conserved††%
L1 subcomplex (membrane-peripheral)
NuoB	AZC_1668	Nqo6	HyqI	AZC_4355	31	66
NuoC	AZC_1669	Nqo5	HyqG(N-term.)†	AZC_4356	23	57
NuoD	AZC_1670	Nqo4	HyqG(C-term.)‡		26	62
NuoE	AZC_1671	Nqo2
NuoF	AZC_1672	Nqo1
NuoG	AZC_1674	Nqo3
NuoI	AZC_1676	Nqo9
LO subcomplex (membrane-integral)
NuoA	AZC_1667	Nqo7
NuoH	AZC_1675	Nqo8	HyqC	AZC_4359	23	58
NuoJ	AZC_1677	Nqo10	HyqE	AZC_4358	18	54
NuoK	AZC_1678	Nqo11
NuoL	AZC_1679	Nqo12	HyqB	AZC_4360	24	55
NuoM	AZC_1680	Nqo13	HyqF	AZC_4357	22	61
NuoN	AZC_1667	Nqo14

5′-end of hyqG encodes residues 1–156;

3′-end of hyqG encodes residues 157–504;

CLUSTAL 2.1 pairwise alignments.

Figure 1

Structure-function rendering of L-type Hyq endo-hydrogenase by analogy and homology to respiratory complex I.

Inferred membrane ubiquinone (Q) or ubiquinol (QH2) binding at the interface of HyqC, HyqG and Hyq I requires partial (14Å) extraction from the respiratory membrane hydrophobic phase; yellow rods represent linked transmembrane and transverse α-helices [14]. Any HyqG catalytic site remains speculative; in vivo activity is in principle fully reversible (see Discussion).

In Growth-optimized A. caulinodans Diazotrophic Liquid Cultures held at 20 µM DOT, endo-hydrogenase Activity Serves in vivo as Respiratory Membrane e– donor for Uptake of Endogenous H2

To recapitulate, A. caulinodans operates distinct, respiratory-membrane exo- and endo-hydrogenases; unlinked ΔhyqRI7 (endo-hydrogenase) and ΔhupSL2 (exo-hydrogenase) complete deletion alleles of relevant structural genes were previously isolated. In growth-optimized liquid diazotrophic cultures open to the environment, exo-hydrogenase mutants grow normally, whereas endo-hydrogenase mutants grow slowly [7]. To more accurately measure relative contributions of both exo- and endo-hydrogenase activities to in vivo recycling of H2 produced by Mo-dinitrogenase activity, H2 evolution rates of diazotrophic liquid batch cultures under continuous sparge have now been measured. A. caulinodans strains were batch cultured at 29°C in defined liquid media lacking utilizable-N; N2 as sole N-source was provided by continuous sparge with (2% O2, 5% CO2, bal. N2) gas mixture optimized for A. caulinodans N2-dependent growth (Materials). Dissolved O2 tension (DOT) in these sparged cultures held steady in the range of 18–20 µM O2 as measured potentiometrically with a Clark-type polarographic electrode (Thermo-Orion 97–08). Culture exit gas streams were periodically sampled and analyzed for evolved H2 by gas chromatography (Materials). In these diazotrophic cultures, both A. caulinodans ΔhyqRI (endo-hydrogenase) mutant 66132 and ΔhyqRI, ΔhupSL (exo-, endo-hydrogenase) double-mutant 66204 showed tenfold elevated H2 evolution rates relative to both hyq +, hup + parent 61305R and ΔhupSL exo-hydrogenase mutant 66081 (Table 2B).

Table 2

H2 evolution by A. caulinodans diazotrophic cultures.

A. caulinodans	Genotype	H2 evolved†	relative H2 evolved‡
(A) N2 and NO3– as N-sources (20 µM DOT)
66204	ΔhyqRI ΔhupSL	460	46.±5.0
66216R	nifK ΔhupSL	10	1.0±0.2
(B) N2 as sole N-source (20 µM DOT; growth optimized)
61305R	nif+ hyq+ hup+	12	1.0±0.2
66081	ΔhupSL	16	1.3±0.3
66132	ΔhyqRI	175	15.±1.6
66204	ΔhyqRI ΔhupSL	540	45.±5.0
(C) N2 as sole N-source (<1 µM DOT; microaerobic)
61305R	nif+ hyq+ hup+	7,100	1.0±0.2
66081	ΔhupSL	61,000	9.0±1.0
66132	ΔhyqRI	2,600	0.4±0.04
66204	ΔhyqRI ΔhupSL	14,000	2.2±0.4
(D) N2 and NO3– as N-source (<1 µM DOT; microaerobic)
60107R	nifA	1,100	1.0±0.2
66216R	nifK ΔhupSL	56,600	51.±3.0

pmol 109·cells−1 min−1 (typical, single experiment);

multiple experiments.

Sparge rates for all liquid cultures were standardized to allow culture atmosphere exhaust rates of 0.5 min–1. In principle, for these growth-optimized diazotrophic cultures, relative abilities of exo- and endo-hydrogenases to recycle endogenous H2 might vary with sparge rates. Increased sparge rates proportionally decreased exit gas H2 levels of all cultures; relative H2 evolution rates among cultures were not affected. Culture sparges were slowed to the minimum rate still maintaining stable 20 µM DOT. Nonetheless, ΔhyqRI single mutant 66132 still evolved tenfold more H2 than did ΔhupSL exo-hydrogenase mutant 66081. Because ΔhyqRI mutants invariably evolved more H2 than did ΔhupSL mutants at 20 µM culture DOT, endo-hydrogenase activity is disproportionately responsible for recycling endogenous H2 produced by Mo-dinitrogenase activity in growth-optimized liquid cultures.

Similarly, when defined media were supplemented with 5 mM L-glutamine, measurable H2 evolution by all strains was negligible. In A. caulinodans cultures, L-glutamine sufficiency yields complete repression of the N2 fixation regulon, including nifD genes encoding Mo-dinitrogenase [19] as well as hyq + genes encoding endo-hydrogenase [7]. A. caulinodans ORS571 wild-type also grows aerobically with either nitrate or nitrite as sole utilizable N-source; both nitrate (AZC0679) and nitrite (AZC0680–AZC0682) reductases are soluble and assimilatory; neither nitrate nor nitric oxide serves as respiratory e– acceptor nor are these activities suggested by analysis of the complete genome sequence [20]. Similarly, A. caulinodans test strains were aerobically cultured in defined medium supplemented with 5 mM nitrate as utilizable N-source. Upon reaching a cell density of ∼1×108 ml–1, exponentially growing liquid cultures were shifted to 2% O2 sparge and H2 levels of exit gases were monitored as before. In this protocol, H2 was evolved by nifK ΔhupSL double- mutant 66216R at baseline levels (Table 2A). By comparison, exo-, endo-hydrogenase double-mutant 66204 evolved H2 at levels corresponding to those of optimized diazotrophic cultures (Table 2A,2B). Accordingly, physiological H2 evolution at 20 µM DOT was thus entirely owed to and benchmarked optimal Mo-dinitrogenase activity.

In Microaerobic (<1 µM DOT) A. caulinodans Cultures, in vivo endo-hydrogenase Activity Reverses, Driving Hydrogenic Respiratory Membrane e– transfer at Extraordinarily High Rates

For A. caulinodans chemostat cultures sparged with 0.2% or more O2, elevated H2 production is not observed [21]. This critical O2 level corresponds to ≥0.9 µM DOT, allowing A. caulinodans 57100 to be continuously cultured with succinate as C-source and N2 as N-source with O2 rate-limiting for growth [22]. In similar continuous cultures at <1 µM DOT, A. caulinodans 57100 dinitrogenase activity levels decrease twofold [23] when compared to optimum (10–20 µM DOT) diazotrophic culture conditions [21], [22], [23]. Accordingly, similar diazotrophic liquid batch cultures were initially sparged with 2% O2, 5% CO2, bal. N2 for 24 hr allowing cell densities to reach ∼1×108 ml–1, at which point sparge gas O2 levels were decreased to 0.11%. In response, culture DOT levels declined precipitously, breaching 1 µM DOT, true microaerobic physiology, defined as DOT insufficient to sustain conventional cytochrome aa oxidase activity [2]. A. caulinodans microaerobic cultures employ two ultra-high O2 affinity terminal oxidases, cytcbb 3 and cytbd, to maintain active oxidative phosphorylation [25]. When microaerobic cultures were supplemented with 5 mM L-glutamine and sampled periodically for viable cell counts by plating (Materials), all strains maintained exponential growth for 72+ hr; for all strains, microaerobic cell doubling-times were 8.1±1.5 hr at 29°C. Indeed, when strains were inoculated at low cell densities (∼1×106 ml–1) and cultured microaerobically (0.11% O2 sparge) in minimal defined medium supplemented with 2.5 mM L-glutamine, all strains and cultures grew completely, and measurable H2 evolution in sparged culture exit gases was insignificant. Whereas, when microaerobic cultures were supplied with sparged (95%) N2 gas as sole N-source, no diazotrophy (cell doubling-times >20 hr) for any strain was measured. Neither was microerobic growth observed when cultures were supplemented with 5 mM nitrate or nitrite. When 5 mM ammonium was supplied, microaerobic growth of test strains was variable. Strain 60107R nifA and 66132 Δhyq cultures both yielded cell-doubling times of 9.5±0.5 hr; parental strain 61305R cultures yielded cell-doubling times of 14±0.8 hr; for all other strains tested, cell-doubling times exceeded 20 hr. In all cases, microaerobic growth with ammonium as N-source inversely correlated with H2 evolution rates (Discussion).

Methylene blue (3,7-bis[dimethylamino]-phenothiazin-5-ium) serves as alternative e– acceptor for respiratory complex I and, when reduced, as e– donor to cytc-dependent cytochrome oxidases, bypassing cytochrome bc (respiratory complex III) activity and uncoupling oxidative phosphorylation [26]. Accordingly, methylene blue was deployed in microaerobic culture samples as in vivo respiratory e– transfer probe. At experimentally sampled time points, culture samples were withdrawn into a gas tight syringe containing anoxic (colorless) methylene blue solution (2 µM final); all culture samples initially turned visibly blue. However, when enclosed syringes were then held at 29°C, within 60 min all culture samples turned completely colorless (anoxic). When thus sampled, all microaerobic A. caulinodans cultures supplied excess succinate as C- and energy source retained respiratory-membrane e– transfer activity for the duration of experiments (days).

Exit gas streams of sparged microaerobic cultures were sampled and H2 evolution was again measured by gas chromatography. Relative to optimized diazotrophic cultures (20 µM DOT), H2 evolution of microaerobic (<1 µM DOT), diazotrophic cultures dramatically increased. In parental 61305R cultures, microaerobic H2 evolution rates increased more than fiftyfold. For ΔhupSL (exo-hydrogenase) mutant 66081, H2 evolution rates increased almost four-thousandfold, which output persisted for 72+ hr. Yet, in endo-hydrogenase mutant 66132 cultures, H2 evolution rates increased only fifteen-fold (Table 2C). H2 evolution rates of all microaerobic batch cultures were sustained 72+ hr given sufficient oxidizable organic-C (succinate) as energy substrate. In conclusion, the extraordinarily high H2 evolution rates of strain-specific microaerobic cultures required both endo-hydrogenase present and exo-hydrogenase absent.

Recall that Mo-dinitrogenase operates in concert with endo-hydrogenase activity given optimum (20 µM DOT) diazotrophic physiology. To what extent does Mo-dinitrogenase activity contribute to H2 output by microaerobic (<1 µM DOT) cultures? To test this hypothesis, similar microaerobic shift experiments were conducted with cultures grown with and maintained on 5 mM nitrate as N-source, which allows full transcriptional derepression of the N2 fixation regulon [19]. Upon microaerobic shift, nitrate-grown nifK ΔhupSL double-mutant 66216R likewise showed exceedingly high H2 output (Table 2D). Note that for all strains, microaerobic cultures with nitrate fail, implying nitrate is not then a competing e– acceptor. Moreover, nitrate itself has no inducible effect on H2 evolution; nifA null mutant 60107R, which entirely lacks ability to derepress the N2 fixation regulon [19], shows negligible H2 output in microaerobic nitrate-supplemented culture (Table 2D). Thus, Mo-dinitrogenase activity itself contributes little (<10%) of the exceedingly high H2 output by ΔhupSL mutant microaerobic cultures. Moreover, because ΔhyqRI ΔhupSL double-mutant 66204 retains 20% microaerobic H2 evolution rates when compared to ΔhupSL single-mutant 66081 (Table 2C), an additional, uncharacterized H2 source is then operative. As it is absent in nifA mutant 60107R (Table 2D), this additional microaerobic H2 source also seems associated with the N2 fixation regulation. In summary, endo-hydrogenase activity is itself responsible for ∼80% of the H2 evolved, i.e., net hydrogenic respiratory membrane e– transfer rates, by microaerobic cultures in which the N2 fixation regulon is derepressed.

Discussion

In summary, A. caulinodans endo-hydrogenase is a bidirectional catalyst whose in vivo activity reverses in response to physiological O2 availability. In optimized diazotrophic (20 µM DOT) cultures, endo-hydrogenase operates in H2 uptake mode, consuming endogenous H2 as respiratory e– donor. In microaerobic (<1 µM DOT) cultures, membrane-integral endo-hydrogenase switches to hydrogenic respiratory-membrane e– transfer mode, employing H+ ions as terminal e– acceptor. Accordingly, endo-hydrogenase shares a microaerobic terminal oxidase role with cytochrome cbb 3 and cytochrome bd [25], whereas respiratory complexes I (NADH:quinone oxidoreductase) and II (succinate dehydrogenase) serve as net e– donors. Measured in vivo respiratory-membrane endo-hydrogenase H2 production rates (45,000 pmol 109·cells−1 min−1) are orders of magnitude higher than previously observed for O2 tolerant hydrogenases in non-fermentative microorganisms. In A. caulinodans exo-hydrogenase mutants lacking microaerobic H2 uptake activity, respiratory membrane endo-hydrogenase mediated H2 production in liquid batch cultures persists at high rates for 72+ hr. Sum totals of evolved H2 (2e– reduction) at 72 hr are 230±30 µmol per 109 cells, representing net oxidation of some 25% of total (340 µmol) succinate supplied these cultures as sole organotrophic energy source and quantitatively converted to poly-β-hydroxbutyrate as organic end-product (5e– oxidation per succinate; [24], [27]).

Similarly, hydrogenic Mo-dinitrogenase activity, at most 10% of hydrogenic endo-hydrogenase activity, consumes almost tenfold more NADH on a mole:mole basis (4 NADH for reductant; 5+ NADH as substrate for oxidative phosphorylation to make the required 16 ATP) [10], [11]. Then, Mo-dinitrogenase activity itself consumes similar amounts of succinate. Unsurprisingly, for all strains tested, diazotrophic (N2 as sole N-source) microaerobic liquid batch cultures open to the environment fail (cell doubling-times >20 hr). Whereas, all strains may be successfully cultured microaerobically with L-glutamine provided as N-source absent all hydrogenesis. When provided ammonium and N2 as N-sources, some strains grow microaerobically, albeit slowly as significant N2 fixation persists; any growth inversely correlates with hydrogenesis by diazotrophic microaerobic cultures (Table 2C). When atmospheric N2 is entirely replaced by argon, ammonium-supplemented microaerobic cultures indeed grow [27], hence Mo-dinitrogenase activity is explicitly responsible for failed microaerobic growth. Earlier, we reported A. caulinodans microaerobic diazotrophic liquid suspension cultures showed increased spectrophotometric absorbance at 600 nm [25]. In more recent experiments, growth in microaerobic diazotrophic liquid suspension cultures was measured by removing samples and aerobically plating for viable cell counts on rich media (Materials) as, in these cultures, viable cell counts do not correlate with increased spectrophotometric absorbance at 600 nm. Likewise, colony growth tests on solid media reflect multiple cell physiology states, and colony growth is facilitated by more efficient H2 recycling at increased cell densities [12].

Among bacteria with known genome sequences, eight genera (A. caulinodans, Azospirillum brasilense, Beijerinckia indica, Bradyrhizobium japonicum; Rhizobium leguminosarum bv. viciae, Rhodopseudomonas palustris, Rhodocista centenaria, Xanthobacter autotrophicus), all microaerophiles capable of N2 fixation, possess orthologous hyq+ operons encoding endo-hydrogenase [12]. Of the six, inferred endo-hydrogenase subunits, five have close homologs in L-type respiratory complex I (Table 1). The sixth (HyqG) protein is homologous to a fused NuoC/D protein. The presumed HyqG H2 catalytic site shared among conserved group-4 hydrogenases is yet undetermined. The group-4 HyqG superfamily (SSF56762) also includes the group-1 exo-hydrogenase catalytic subunit, which possesses a heteronuclear Ni,Fe catalytic center coordinated by four, completely conserved Cys residues, of which two bridge the catalytic Ni and Fe = C = O binuclear center [28]. To the contrary, inferred HyqG proteins from eight microaerophilic genera all lack both N-terminal and C-terminal Cys-X-X-Cys motifs. Rather, three Cys residues (A. caulinodans Cys-258, Cys-491, and Cys-497) are completely conserved by the HyqG family (Fig. S7). The NuoD (Nqo4) proteins of bacterial respiratory complex I, also members of this same Superfamily, neither possess the Cys-X-X-Cys pairs nor do they exhibit a Ni,Fe binuclear center. Therefore, that HyqG actually carries a binuclear Ni,Fe catalytic site seems uncertain, if not unlikely. The binding site for respiratory complex I membrane ubiquinone, its ultimate e– acceptor, is a cavity formed between a four-helix bundle of NuoD (Nqo4), the H1 helix of NuoB (Nqo6), and transmembrane helix 1 of NuoH (Nqo8) [14], [18], all of which motifs are conserved in the Hyq endo-hydrogenase (HyqG, HyqI, and HyqI, respectively).

By inference, the Hyq endo-hydrogenase is a membrane-integral H2:quinone oxidoreductase. Given the strong reducing potential of the biochemical standard hydrogen electrode (E o′ = –0.414V) relative to that of ubiquinone (E o′ = +0.070V) respiratory-membrane hydrogenesis is a highly endergonic process under standard conditions. In Rps. palustris, membrane physiology has been modeled under a variety of dynamic steady-state conditions including microaerobic respiration, whose membrane ubiquinone pools are necessarily highly (>90%) reduced [29]. If membrane ubiquinone pools were poised some 90% reduced, a ubiquinon/ubiquinol half-cell potential (E′) of +0.040V (at 25°C) would obtain. By inference, the balanced reaction for endo-hydrogenase mediated H2 uptake, including trans-membrane H+ pumping, may be written:where N connotes endo and P connotes exo membrane faces. In purified, reconstituted vesicles, respiratory complex I activity, including H+ translocation, is fully reversible [30]. If the same holds true for endo-hydrogenase in vivo, its hydrogenesis mode activity may be written:

This activity would tap steady-state membrane proton-motive force (Δp), modeled in Rps. palustris microaerobic respiratory membranes as Δp = 0.195V [29]. Were 2H+() counter-transported during steady-state hydrogenesis, E′ values at 25°C would be effectively lowered from +0.040V to −0.350V, at which steady-state potential the operative hydrogen half-cell H2 partial pressure (pH2) would approach 0.7 kPa at 25°C. Indeed, when exit gases of sparged A. caulinodans hydrogenic cultures maintained at 30°C were analyzed, pH2 levels reproducibly approached 0.7 kPa as sustained hydrogenesis rates. In all likelihood, high-level endo-hydrogenase dependent H2 production requires both highly reduced membrane ubiquinone pools and high Δp values. In A. caulinodans microaerobic liquid batch cultures, elevated H2 production requires supplementation with excess, primary C-source (succinate, L-malate, or L-lactate), which presumably drive reduction of respiratory membrane ubiquinone pools at relatively high rates. Moreover, if indeed consumptive of membrane Δp, respiratory membrane hydrogenesis only operates when sufficient O2 is also available as respiratory e− acceptor to regenerate high Δp (respiratory complex I, III, and IV activities). Regardless, any respiratory-membrane hydrogenesis would necessarily largely uncouple oxidative phosphorylation.

For A. caulinodans microaerobic respiration, both available (limiting) O2 and H+ ions simultaneously serve as e– acceptors. A. caulinodans then employs multiple cytc- and ubiquinol-oxidases [25], resulting in varied respiratory membrane proton-translocation yields. So, no fixed stoichiometries of H2 relative to H2O production may be deduced. Moreover, parceling out relative in vivo contributions as respiratory membrane e– acceptors is problematic, given restricted choice. When one or more e– acceptor activities are absent due to mutation, compensatory flux to soluble e– acceptors, such as N2 (Mo-dinitrogenase), NO3 − (nitrate reductase), and CO2 (both rubisco and CO dehydrogenase) then obtains. As one example, H2 evolution rates for exo- endo-hydrogenase double-mutant 66204 increased twenty-five-fold when shifted from growth optimal (20 µM DOT) to microaerobic (<1 µM DOT) conditions, presumably owed to restricted choice of available e– acceptors (i.e., relative absence of O2). Moreover, restricted choice also extends to e– donors. Because it successfully reoxidizes >80% of H2 then produced by combined (>90%) endo-hydrogenase and (<10%) Mo-dinitrogenase activities, exo-hydrogenase operates as a relatively more competitive microaerobic respiratory membrane e– donor. Conceivably, these changes in cellular microaerobic respiratory membrane physiology might simply reflect more-reduced ubiquinone pools. Alternatively, respiratory membranes might build exo- and/or endo-hydrogenases into macromolecular complexes which would preclude simple diffusion control of respiratory e− transfer by membrane ubiquinone pools.

In A. caulinodans, the two, respiratory membrane hydrogenases possess distinct physiological roles; group-4 endo-hydrogenase activity is bidirectional and strictly correlates with diazotrophy and endogenous H2 uptake, whereas group-1 exo-hydrogenase activity is unidirectional and also allows chemoautotrophy with exogenous H2 as energy source [7]. Among capable anaerobes, fermentative membrane hydrogenesis is well described [1], [2], [5]. Whereas, among obligate aerobes, respiratory membrane hydrogenesis as a sustained physiological process seems counterproductive, as it significantly uncouples oxidative phosphorylation. Indeed, in diazotrophic microaerobic A. caulinodans cultures, exo-hydrogenase and endo-hydrogenase are both highly active and thus seemingly operative at cross-purposes.

However, liquid culture experiments, which strive to allow all bacterial cells a similar physiological milieu, are contrived. In reality, bacterial cell populations experience a dimensional world. We suggest, as one possibility, concomitant H2 evolution and H2 uptake might prove useful if partitioned among aerobic and microaerobic bacterial cells in dimensional populations. Varying O2 microenvironments within organotrophic bacterial colonies or biofilms might de facto segregate metabolic physiology, allowing internal O2-restricted cells to evolve H2 and external O2-sufficient cells to take up and use that H2, driving oxidative phosphorylation. Superficial H2 oxidizing, O2 rich cells might then redirect environmental organic-C sources away from catabolism (oxidative phosphorylation substrate) towards anabolism (C-assimilation), augmenting growth rates and proliferation of dimensionally organized and specialized cell populations considered as a whole.

Materials and Methods

Bacterial Strains and Media

Azorhizobium caulinodans ORS571 wild-type (strain 57100; ATCC No. 43989), was originally isolated from Sesbania rostrata stem-nodules [8]. Strain 61305R [32], a 57100 derivative carrying an IS50R insertion in the (catabolic) nicotinate dehydrogenase structural gene served as ‘virtual’ wild-type for reported experiments; 61305R uses supplied (3 µM) nicotinate only as anabolic substrate for synthesis of pyridine nucleotides, for which 57100 is auxotrophic [33]. Precise, in-frame deletion mutagenesis of A. caulinodans target genes was conducted out by “crossover PCR” as previously described [34]. Strain 66216R was constructed as for strain 66081 with strain 60057R as parent (Table 3). Defined media for all cultures was basal NIF medium (7.5 mM potassium phosphate pH 6.3, 1 mM MgSO4, 0.5 mM CaCL2, 2 µM ferric citrate, 3 µM nicotinate, 1 µM sodium molybdate, 1 µM pantothenate, 0.1 µM D-biotin, and Hutner’s “44′ trace elements [35]) supplemented with 20 mM potassium succinate as sole C- and energy source, and 2.5 mM ammonium bicarbonate as N-source. Strains (whose lineage does not include 61305R) and which actively catabolize nicotinate were supplemented with 0.1 mM nicotinate in aerobic cultures; in microaerobic cultures, A. caulinodans wild-type 57100 does not measurably catabolize nicotinate.

Table 3

Azorhizobium caulinodans strains.

Strain	Genotype	Ref.
57100	ORS571 wild-type	[8]
60035	57100 nifD35::Vi	[31]
60035R	60035 nifD35::IS50R
60057R	60057 nifK57::IS50R
60107R	57100 nifA107R
61305R	57100 Nic–, 6-OH-Nic+	[32]
66081	61305R hupΔSL2	[7]
66132	61305R hyqΔRI7	[7]
66204	61305R hupΔSL2 hyqΔRI7	[7]
66216R	60057R hupΔSL2

Physiological Growth Measurements and Evolved H2 Analyses

Starter cultures of A. caulinodans strain 61305R and its derivatives were aerobically cultured in minimal defined NIF liquid medium [8] supplemented with: 0.3 mM ammonium as sole, limiting N-source and 3 µM nicotinate at 37°C until growth arrest (cell densities ∼1×108 cells ml−1). For kinetic measurements of diazotrophy, arrested starter cultures were each diluted one-hundredfold in 20 ml NIF medium; serum vials (30 ml capacity) were sealed with silicone rubber septa, sparged continuously (10 ml min−1) with defined gas mixtures (e.g. 2% O2, 5% CO2, bal. N2), and incubated at 29°C. At least three times per cell-doubling period, culture samples were removed, serially diluted, plated on rich GYPC medium [9], and incubated aerobically 48 hr at 37°C; colonies were counted in triplicate. In vivo H2 uptake activities were inferred, coupled to Mo-dinitrogenase activity as H2 donor, by comparing rates of H2 evolution from A. caulinodans hup + hyq + (wild-type), ΔhupSL (exo-hydrogenase) mutant, ΔhyqRI (endo-hydrogenase) mutant, and ΔhupSL ΔhyqRI double-mutant cultures (Table 3). Collectively, both hydrogenases account for all in vivo H2 uptake activity [12]. (Amperometric cell-free assay of isolated endo-hydrogenase H2 uptake activity with exogenous H2 as substrate is not at hand.) To measure evolved H2, sparged culture exit gas streams were sampled and analyzed by gas chromatography (RPC1; Peak Laboratories LLC) fitted with an HgO (reducing compound) photometer as detector [36] and a fixed volume (25 µl) sampling loop. Molar H2 evolution rates were inferred from measured dilution rates of culture atmospheric volumes. Total cellular protein was measured by the bicinchoninic acid procedure (Sigma-Aldrich Co.); for A. caulinodans and related microaerophiles employing oxidative metabolic gearing, mean total cell proteins levels (135±15 fg) are largely independent of cell physiology [24].

HyqB and NuoL (CLUSTAL W2) alignment.

(EPS)

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HyqC and NuoH (CLUSTAL W2) alignment.

(EPS)

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HyqE and NuoJ (CLUSTAL W2) alignment.

(EPS)

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HyqF and NuoM (CLUSTAL W2) alignment.

(EPS)

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HyqG and NuoC/D (CLUSTAL W2) alignment.

(EPS)

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HyqI and NuoB (CLUSTAL W2) alignment.

(EPS)

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HyqG orthologs (CLUSTAL W2) multiple alignment. From the top, species are: Rhizobium leguminosarum (Rhleg), Azorhizobium caulinodans (Azoca), Xanthobacter autotrophicus (Xanpy2), Beijerinckia indica (Beind), Bradyrhizobium japonicum (Braja), Rhodospirillum centenum, (Rhoce), Rhodopseudomonas palustris (BisA53), Azospirillum brasilense (Azobr), Rhodopseudomonas palustris (HaA2), Rhodopseudomonas palustris (BisB18), Rhodopseudomonas palustris (BisB5).

(EPS)

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