Link between the Membrane-Bound Pyridine Nucleotide Transhydrogenase and Glutathione-Dependent Processes in Rhodobacter sphaeroides

Abstract

The presence of a glutathione-dependent pathway for formaldehyde oxidation in the facultative phototroph Rhodobacter sphaeroides has allowed the identification of gene products that contribute to formaldehyde metabolism. Mutants lacking the glutathione-dependent formaldehyde dehydrogenase (GSH-FDH) are sensitive to metabolic sources of formaldehyde, like methanol. This growth phenotype is correlated with a defect in formaldehyde oxidation. Additional methanol-sensitive mutants were isolated that contained Tn5 insertions in pntA, which encodes the α subunit of the membrane-bound pyridine nucleotide transhydrogenase. Mutants lacking transhydrogenase activity have phenotypic and physiological characteristics that are different from those that lack GSH-FDH activity. For example, cells lacking transhydrogenase activity can utilize methanol as a sole carbon source in the absence of oxygen and do not display a formaldehyde oxidation defect, as determined by whole-cell ¹³C-nuclear magnetic resonance. Since transhydrogenase can be a major source of NADPH, loss of this enzyme could result in a requirement for another source for this compound. Evidence supporting this hypothesis includes increased specific activities of other NADPH-producing enzymes and the finding that glucose utilization by the Entner-Doudoroff pathway restores aerobic methanol resistance to cells lacking transhydrogenase activity. Mutants lacking transhydrogenase activity also have higher levels of glutathione disulfide under aerobic conditions, so it is consistent that this strain has increased sensitivity to oxidative stress agents like diamide, which are known to alter the oxidation reduction state of the glutathione pool. A model will be presented to explain the role of transhydrogenase under aerobic conditions when cells need glutathione both for GSH-FDH activity and to repair oxidatively damaged proteins.

Formaldehyde (HCHO) is a toxic compound that organisms encounter frequently in nature. It can be produced by many industrial means or by biological oxidation of methanol, methyl halides, and methylated compounds (16, 19, 25, 30, 50). Cells often have a mechanism to remove formaldehyde due to its ability to damage nucleic acids, proteins, and other cellular components (21, 32, 47). Previous work has identified a glutathione-dependent pathway for formaldehyde oxidation to carbon dioxide in the facultative phototroph Rhodobacter sphaeroides (described below). In this work, a molecular explanation is provided for how different enzyme activities contribute to this glutathione (GSH)-dependent pathway (Fig. 1).

FIG. 1.

Contribution of different enzyme activities to GSH-dependent pathway for formaldehyde oxidation in R. sphaeroides.

The existence of this GSH-dependent pathway for formaldehyde oxidation was suggested by several observations (5–7). Upon addition of [¹³C]formaldehyde to cell suspensions, the production of S-hydroxymethyl-GSH (GS-CH₂OH), formate (HCO₂⁻), and carbon dioxide (CO₂) is observed via nuclear magnetic resonance (NMR) (5). Additionally, growth in the presence of carbon sources like methanol, which are known to be metabolic sources of formaldehyde, results in the induction of a GSH-dependent formaldehyde dehydrogenase and a formate dehydrogenase (GSH-FDH and FDH, respectively) (5, 6). Also, mutations that inactivate the GSH-FDH gene (adhl) prevent cells from utilizing methanol as the sole photosynthetic carbon source, presumably because this strain is unable to generate carbon dioxide for subsequent assimilation by the Calvin-Benson-Bassham cycle (5). Cells lacking GSH-FDH are also unable to grow when methanol is added to a succinate-based minimal medium under aerobic respiratory conditions. One possibility to explain this methanol sensitivity is that formaldehyde accumulates to toxic levels in cells that lack GSH-FDH activity as methanol is oxidized. In this work, a defect in formaldehyde oxidation is correlated with the aerobic methanol sensitivity of cells that lack GSH-FDH activity.

We also demonstrate that inactivation of pntA, which encodes a subunit of the enzyme pyridine nucleotide transhydrogenase, results in aerobic methanol sensitivity. In bacteria, pyridine nucleotide transhydrogenase is typically a membrane-bound enzyme that catalyzes the reversible transfer of hydride equivalents between NAD⁺ and NADP⁺ with concomitant proton translocation across the cytoplasmic membrane (23). Unlike cells that lack GSH-FDH activity, pntA mutations do not lead to a formaldehyde oxidation defect. R. sphaeroides mutants lacking transhydrogenase activity appear to only have an aerobic growth defect, and this phenotype is suppressed when these cells are provided with another metabolic source of NADPH. Loss of transhydrogenase activity under aerobic conditions also leads to an increase in the levels of GSH disulfide (GSSG) and an increase in sensitivity to the oxidative stress agent diamide, which is known to form disulfide bonds in cytoplasmic proteins. From these data, a model is presented to explain why loss of transhydrogenase slows growth of cells when GSH is required for both formaldehyde oxidation by the GSH-dependent pathway and the repair of cytoplasmic proteins that become damaged in the presence of oxygen.

MATERIALS AND METHODS

Strains, plasmids, and growth conditions.

Bacteria and plasmids used in this study are listed in Table 1. R. sphaeroides was grown in Sistrom’s minimal medium (43) at 32°C containing either succinate (35 mM) or glucose (20 mM). For testing of aerobic methanol sensitivity, the medium was supplemented with 100 mM methanol. Photosynthetic growth with 100 mM methanol as a sole carbon source used Sistrom’s medium lacking succinate with 80 mM dimethyl sulfoxide added as an external electron acceptor. Escherichia coli strains were grown at 37°C in Luria-Bertani medium. When necessary, the medium was supplemented with spectinomycin (25 μg/ml), trimethoprim (30 μg/ml), or tetracycline (1 μg/ml) for R. sphaeroides, and ampicillin (100 μg/ml), tetracycline (10 μg/ml), or trimethoprim (300 μg/ml) for E. coli. Mobilization of plasmids from E. coli into R. sphaeroides was performed as described (17).

TABLE 1.

Bacterial strains and plasmids used in this study

Strain or plasmid	Relevant phenotype or genotype	Source or reference
Strains
R. sphaeroides
Ga	crtI	Laboratory strain
Adhl1	Ga derivative, Δadhl::ΩSpr	7
MS21	Ga derivative, Tn5 insertion at codon 474 of pntA (pntA1)	This study
MS65	Ga derivative, Tn5 insertion at codon 128 of pntA (pntA2)	This study
PntA1	Ga derivative, Tpr insertion at codon 285 of pntA (pntA3)	This study
E. coli
DH5α	supE44 ΔlacU169 (φ80lacZΔM15) hsdR178 recA1 endA1 gyrA96 thi-1 relA1 C600::RP-4 2-(Tc::Mu) (Kn::Tn7) thi pro hsdR hsdM+ recA	8
S17-1		42
Plasmids
pUC19	Apr	34
pBR322	Apr	9
pRK415	Tcr Mob+lacZα IncP	26
pSUP202	Apr Tcr Cmr Mob+; pBR322 derivative	42
pLA2917	Knr Tcr RK2 derivative; cos	1
pUI8662	Tcr; pLA2917 derivative, cosmid containing pntAB	This study
pS12	10-kb BamHI fragment from pUI8662 containing pntAB cloned into pUC19	This study
pPnt3	3.5-kb PCR-generated fragment from pS12 cloned into BamHI-HincII sites of pUC19	This study
pPnt4	3.5-kb BamHI-HindIII fragment from pPnt3 cloned into pRK415	This study
pNIC1	6.1-kb BamHI fragment containing R. rubrum pntAA pntAB pntB cloned into pBR322	49
pRTH1	6.1-kb BamHI fragment from pNIC1 cloned into pUC19	This study
pRTH2	6.1-kb BamHI-HindIII fragment from pRTH1 cloned into pRK415	This study
pΔPnt3	0.7-kb Tp cassette cloned into HincII site in pntA at codon 285	This study
pΔPnt4	2.0-kb PstI fragment from pΔPnt3 cloned into pSUP202	This study

Isolation of methanol-sensitive mutants.

R. sphaeroides Ga was mutagenized with a derivative of Tn5 that encodes trimethoprim resistance (Tp^r) (11, 33). The mutant bank was screened for inability to grow on a succinate-based minimal medium supplemented with 100 mM methanol by replica plating. To map the Tn5 insertion in the methanol-sensitive mutants, MS21 and MS65, chromosomal DNA was analyzed by pulsed-field electrophoresis (44). To clone the region flanking the Tn5 insertion, genomic DNA was digested with BamHI and cloned into pUC19, selecting for Tp^r resistance. Primers specific to the IS50 region were used to sequence the DNA flanking the Tn5 insertion. To isolate a wild-type copy of this region, an existing cosmid library of wild-type DNA was mobilized into the methanol-sensitive mutants, screening for cosmids that resulted in methanol-resistant exconjugants. After a cosmid (pUI8662) was found to contain an ≈8-kb BamHI-complementing restriction fragment, this was used to obtain the wild-type DNA sequence of this region.

Constructing a pntAB plasmid.

Primers specific to sequences upstream (5′-NNGGATCCCGTGCCCGCCTTCCTGTG-3′) or downstream (5′-AGGAAGAGGCCGCCGTGTCA-3′) of pntAB were used to amplify a region that contains ∼300 and ∼230 bp of upstream and downstream DNA, respectively. The upstream primer contained a BamHI restriction site (underlined) to facilitate cloning of an ∼3.5-kb PCR product into BamHI-HincII-digested pUC19. After DNA sequencing to ensure the absence of mutations, the plasmid (pPnt3) was digested with BamHI-HindIII, and the resulting ∼3.5-kb restriction fragment was cloned into BamHI-HindIII-digested pRK415 to create pPnt4.

Creating an insertion in pntA.

A gene encoding Tp^r was cloned into a HincII site at codon 285 within pntA to create the pntA3 allele. The resulting plasmid (pΔPnt3) was digested with PstI, producing an ∼2.0-kb restriction fragment that was cloned into the PstI site of the mobilizable suicide plasmid pSup202 (42). After transforming the resulting plasmid (pΔPnt4) into S17-1, it was mobilized into R. sphaeroides. Strains in which pntA3 had replaced the wild-type gene by an even number of crossover events were identified by screening Tp^r colonies for sensitivity to tetracycline (encoded by the suicide plasmid). The presence of the pntA3 allele in PntA1 was confirmed by PCR with primers that flank pntA.

Growth rate determinations.

Cultures were started from single colonies and subcultured into fresh media without antibiotics. Cell growth was monitored by measuring the optical density at 600 nm in a Shimadzu Biospec 1601 spectrophotometer. The data reported are the average doubling times calculated from at least two separate cultures.

Whole-cell ¹³C-NMR spectroscopy.

Experiments were performed as previously reported (5) with the following modifications. Cultures (500 ml) were grown aerobically by bubbling with 69% N₂, 30% O₂, and 1% CO₂. Once the culture reached a density of 5 × 10⁸ to 1.0 × 10⁹ cells/ml, cells were harvested (5,000 × g for 15 min at 4°C), washed once in 0.15 M potassium phosphate buffer (pH 7.5), and suspended in the same buffer at ∼10¹¹ cells/ml.

For photosynthetically grown cells, 2.0 ml of this cell suspension and 1.0 ml of D₂O were added to an NMR tube and bubbled with N₂ gas. At time zero, [¹³C]formaldehyde was added to a final concentration of 10 mM, and dimethyl sulfoxide was added to a final concentration of 80 mM to act as an electron acceptor. The NMR tubes were sealed and placed in front of a light source, and spectra were acquired at the indicated time points.

For aerobically grown cells, suspensions were bubbled with air. At time zero, [¹³C]formaldehyde was added to a final concentration of 10 mM, and 2.0-ml samples were removed and mixed with D₂O (1.0 ml) in NMR tubes at the indicated times.

Decoupled ¹³C-NMR spectra were acquired at 100.6 MHz by using a sweep width of 25,252.525 Hz with a deuterium lock on a Bruker Instruments DMX-400 Avance console, 9.4T wide-bore magnet, NMR spectrophotometer at the National Magnetic Resonance Facility at University of Wisconsin—Madison (NMRFAM). The pulse duration was 15 μs, with a relaxation time of 3.5 s. The spectra were Fourier transformed with 5 Hz of line broadening; data sizes were 16,384 before and after transformation. Peak assignments were aided by an internal standard of tetramethylsilane (TMS) present in the NMR tube in a sealed capillary. To assign peaks, control spectra were obtained with commercially available standards or synthetic material (5).

Diamide sensitivity assays.

Approximately 10⁷ late-exponential-phase cells (culture density of ∼1.5 × 10⁹ cells/ml) that were grown aerobically in the absence of antibiotics were mixed into 5 ml of molten Sistrom’s soft agar (0.7%) and poured onto an agar plate containing the same medium. After the soft agar solidified, a 1-cm-diameter filter disk containing 10 μl of 1 M diamide was placed on the plate. After incubation for 72 to 96 h at 32°C, the resulting zones of growth inhibition (clearing) were measured.

Cell extract preparation, enzyme assays, and protein determination.

Crude cell extracts were prepared from exponential-phase cultures as in reference 5, with the following modifications. Harvested cells were washed in 100 mM potassium phosphate buffer (pH 7.5) and suspended in 3 ml of either 150 mM potassium phosphate buffer (pH 8.5) for GSH-FDH assays, 50 mM potassium phosphate buffer (pH 7.0) for pyridine nucleotide transhydrogenase and isocitrate dehydrogenase assays, or 50 mM Tris-Cl (pH 8.9) for glucose-6-phosphate dehydrogenase assays. Protein concentrations were determined by A_260–280 or by the Bradford method (31). All enzyme and protein assays were performed on a Shimadzu Biospec 1601 spectrophotometer.

GSH-FDH activity was measured as previously described (5), except that potassium phosphate buffer (150 mM [pH 8.5]) was used. To measure pyridine nucleotide transhydrogenase activity, the final reaction mixture contained 50 mM potassium phosphate (pH 7.0), 100 mM NaCl, 0.1 mM NADPH, 0.1 mM acetylpyridine-NAD⁺, and 10 mM β-mercaptoethanol. The assay was started by addition of ∼100 μg of crude extract protein, and a change in A₃₇₅ was used to follow reduction of acteylpyridine-NAD⁺ (29). Isocitrate dehydrogenase activity was measured by monitoring the isocitrate-dependent reduction of NADP⁺ to NADPH. The final reaction mixture contained 50 mM potassium phosphate (pH 7.0), 4 mM isocitrate, 5 mM MgCl₂, and 0.1 mM NADP⁺. The reaction was started by addition of 100 to 200 μg of crude extract protein (12). Glucose-6-phosphate dehydrogenase activity was measured by monitoring the glucose-6-phosphate-dependent reduction of NADP⁺ to NADPH. The final reaction mixture contained 50 mM Tris-Cl (pH 8.9), 10 mM glucose-6-phosphate, 10 mM MgCl₂, 0.2 mM ATP, and 0.2 mM NADP⁺. The assay was started by addition of 100 to 200 μg of crude extract protein (24). GSSG reductase activity was measured by monitoring the GSSG-dependent oxidation of NADPH to NADP⁺. The final reaction mixture contained 100 mM potassium phosphate (pH 7.8), 1 mM EDTA, 1 mM GSSG, and 0.1 mM NADPH. The assay was started by addition of 100 to 200 μg of crude extract protein (28).

GSH and GSSG assays.

Cultures (500 ml) were grown aerobically by bubbling with 69% N₂, 30% O₂, and 1% CO₂. Once the culture reached a density of 5 × 10⁸ to 1.0 × 10⁹ cells/ml, 100 to 200 ml of cells was harvested (5,000 × g for 10 min, 4°C), the cell pellets were washed with buffer (100 mM potassium phosphate, 4.4 mM EDTA [pH 7.5]), and suspended in 1 ml of ice-cold 5% 5-sulfosalicylic acid. The acid extracts were centrifuged at 15,000 × g for 5 min at 4°C to remove cell debris. The supernatant was removed, passed through a 0.2-μm-pore-size filter, and assayed for GSH and GSSG on the same day as sample preparation.

Total GSH and GSSG were assayed by the method of Griffith (18). For assaying total GSH, 10 μl of the acid-soluble extract was added to a mixture of buffer (100 mM potassium phosphate, 4.4 mM EDTA [pH 7.5]), 100 μM NADPH, and 600 μM dithionitrobenzoic acid that was prewarmed to 32°C. The assay was started by the addition of 10 μl of GSSG reductase at a concentration of 300 U/ml. The change in A₄₁₂ over a 2-min period was used to calculate the total GSH after comparing this to a standard curve prepared with known quantities of GSH. GSSG was measured by the same procedure, except that 100 μl of the acid extract was mixed with 2 μl of 2-vinylpyridine and triethanolamine (∼4 to 6 μl to neutralize the acid) for 1 h at room temperature to remove GSH before the assay was initiated. In this case, a second standard curve using samples containing GSSG, 2-vinylpyridine, and triethanolamine was prepared to determine the concentration of GSSG in acid extracts. The amount of GSH in samples was estimated by subtracting GSSG levels from the total GSH value. The amount of GSH or GSSG present per gram (dry weight) of cells was determined by calculating the cellular dry weight (13) by using dry weight estimates for aerobically grown R. sphaeroides reported previously (46).

RESULTS

Loss of GSH-FDH leads to a defect in formaldehyde oxidation.

Mutations that inactivate the GSH-FDH structural gene (adhl) both block the ability of R. sphaeroides to use methanol as a sole photosynthetic carbon source and cause sensitivity to 100 mM methanol under aerobic respiratory conditions (5). Since methanol is oxidized to formaldehyde, the lack of GSH-FDH could cause a defect in oxidation of the formaldehyde that is generated from this carbon source. To test if a lack of GSH-FDH results in a defect in formaldehyde metabolism, the fate of ¹³C-labeled formaldehyde was monitored in suspensions of wild-type cells and those lacking GSH-FDH activity (Adhl1) that were grown under aerobic conditions. In wild-type cells (Fig. 2, left panels), the majority of the [¹³C]formaldehyde (81.9 ppm) was depleted after 90 min with the accumulation of formate (171.0 ppm) and smaller amounts of 5, 10 methylenetetrahydrofolate (66.5 ppm). The time-dependent oxidation of formaldehyde observed at this and other time points in wild-type cells is generally similar to that previously observed when analyzing cells that were grown aerobically in the presence of both succinate and methanol (5).

FIG. 2.

Metabolic fate of formaldehyde in wild-type cells (Ga, [left panels]) and those lacking GSH-FDH activity (Adhl1 [right panels]). Metabolism of [¹³C]formaldehyde was monitored after its addition to suspensions of cells that were grown aerobically utilizing succinate as the sole carbon source. Spectra were collected at the time points indicated after the addition of formaldehyde. All peaks were normalized by using TMS (0 ppm) as an internal standard and assigned with appropriate standards. HCHO, formaldehyde; HCO₂⁻, formate; CH₂-THF, 5, 10 methylenetetrahydrofolate; TMS, tetramethylsilane.

In contrast, 90 min after addition of ¹³C-labeled formaldehyde to a suspension of Adhl1 (Fig. 2, right panels), the majority of the [¹³C]formaldehyde remains, with accumulation of only small amounts of formate and 5, 10 methylenetetrahydrofolate. There is no further metabolism of [¹³C]formaldehyde in cells that lack GSH-FDH even ∼300 min after this compound was added (data not shown). Thus, it appears that aerobic methanol sensitivity of cells that lack GSH-FDH activity is correlated with a defect in formaldehyde oxidation.

The small amount of formate that is produced in Adhl1 is likely due to a GSH-FDH-independent process, since this strain does not contain detectable levels of a GSH-dependent formaldehyde dehydrogenase activity (5, 7). Native polyacrylamide gels stained for formaldehyde dehydrogenase activity show that other proteins are capable of oxidizing formaldehyde in both wild-type cells and Adhl1 (data not shown). However, the methanol sensitivity of Adhl1 indicates that any GSH-independent formaldehyde dehydrogenase activities are insufficient to dissimilate formaldehyde at a rate that alleviates its toxicity.

Screening for methanol-sensitive mutants.

Having shown a mutation that interferes with formaldehyde metabolism by the GSH-dependent pathway can lead to aerobic methanol sensitivity, a Tn5 mutant library was generated and screened for other mutations that produced aerobic methanol sensitivity. This screen identified three independent mutants, two of which (MS21 and MS65) are analyzed in this study (Table 2).

TABLE 2.

Growth phenotypes

Strain	Aerobic								Photosynthetic (anaerobic)
	Succinate		Succinate + MeOH		Glucose		Glucose + MeOH		Succinate		MeOH
	Growtha	Doubling timeb (min)	Growtha	Doubling timeb (min)	Growtha	Doubling timeb (min)	Growtha	Doubling timeb (min)	Growtha	Doubling timeb (min)	Growtha	Doubling timeb (min)
Ga	+	172 ± 3	+	185 ± 7	+	195 ± 4	+	207 ± 21	+	163 ± 10	+	645 ± 10
Adhl1	+	189 ± 1	−		+	210 ± 8	−		+	161 ± 1	−
MS65	+	209 ± 13	−	472 ± 42	+	201 ± 4	+	207 ± 13	+	231 ± 8	+	680 ± 42
MS65 pPnt4	+	198 ± 34c	+	207 ± 38c	+	NDd	+	ND	+	ND	+	ND
PntA1	+	217 ± 16	−	432 ± 85	+	222 ± 25	+	216 ± 8	+	227 ± 2	+	703 ± 26
PntA1 pPnt4	+	222 ± 17c	+	210 ± 17c	+	ND	+	ND	+	ND	+	ND

^aGrowth phenotypes on plates after 5 days of incubation at 30°C. In the case of photosynthetic conditions, growth was assayed either on plates (succinate) or in liquid cultures (methanol).

^bDoubling times (in minutes) during the logarithmic growth phase. The data reported are averages of at least two independent growth experiments.

^cThe medium contained 1 μg of tetracycline per ml.

^dND, not determined.

Pulsed-field electrophoresis mapped the Tn5 insertions in MS21 and MS65 to the same region (coordinate ∼2970) of R. sphaeroides chromosome (data not shown), which is ∼200 kb away from the GSH-FDH structural gene (39). Assays for GSH-FDH activity showed that both MS21 and MS65 contained wild-type levels of this enzyme (Table 3). Thus, the mutations in MS21 and MS65 cause methanol sensitivity without altering GSH-FDH expression.

TABLE 3.

Enzyme specific activities in wild-type cells (Ga) or those lacking GSH-FDH (Adhl1) or transhydrogenase (MS65) activity

Strain	Plasmid	Relevant phenotypea	Sp actb
			GSH-FDH (μmol of NAD+ reduced/min/mg of protein)	Transhydrogenase (μmol of acetylpyridine-NAD+ reduced/min/mg of protein)
Ga	None	MeOHr	12.0 ± 1.2	50.8 ± 5.5
Adhl1	None	MeOHs	<0.01	NDc
MS21	None	MeOHs	17.0 ± 2.1	<0.1
MS65	None	MeOHs	15.4 ± 0.9	<0.1
MS65	pPnt4	MeOHr	13.2 ± 3.1	186.5 ± 20.4
MS65	pRTH2	MeOHr	11.7 ± 2.2	109.2 ± 24.1
PntA1	None	MeOHs	14.2 ± 3.1	<0.1
PntA1	pPnt4	MeOHr	13.0 ± 4.2	110.3 ± 35.4

^aMeOH^r, methanol resistance; MeOH^s, methanol sensitivity.

^bData are averages of at least three independent experiments.

^cND, not determined.

Loss of pyridine nucleotide transhydrogenase activity can cause aerobic methanol sensitivity in R. sphaeroides.

When the DNA sequence of the region flanking the Tn5 element in MS21 and MS65 was determined, both insertions were found to lie within a gene predicted to encode a protein with over 70% amino acid sequence similarity to PntA from E. coli, the α subunit of pyridine nucleotide transhydrogenase (GenBank accession no. AY026033) (14). Immediately downstream of pntA is a gene (pntB) that encodes a homologue of the β subunit of this enzyme (Fig. 3).

FIG. 3.

Organization of the R. sphaeroides pntAB locus. The predicted direction of transcription of each proposed gene is indicated by the arrows. The points of Tn5 insertion in MS21 and MS65 are indicated above pntA. Below pntA is shown the position of the trimethoprim cartridge in PntA1. Distances between each gene are shown below the genetic map. An open reading frame predicted to lie ∼580 bp upstream of pntA, transcribed in the opposite direction, displayed homology to a hypothetical protein predicted to be a kyurenine hydrolase from Pseudomonas aeruginosa (AAG05468.1). An open reading frame ∼280 bp downstream of pntB displayed similarity to a hypothetical protein of unknown function from Agrobacterium tumefaciens (AF065244).

To determine if these mutations inactivated genes that encoded the R. sphaeroides homologue of this enzyme, transhydrogenase specific activities were measured in wild-type cells and the two methanol-sensitive mutants. Crude extracts from wild-type cells contained specific activities comparable to those in other bacteria (29), but extracts from MS21, MS65, or a strain that contains a defined insertion in pntA (PntA1) at codon 285 did not contain any detectable activity in this assay (Table 3).

To test if loss of pntA was causal to aerobic methanol sensitivity, a plasmid (pPnt4) was introduced into MS21, MS65, and PntA1 that contained pntAB with 300 and 230 bp upstream and downstream of pntAB, respectively. As expected, this pntAB plasmid restored both transhydrogenase activity (Table 3) and the ability of these strains to grow on solid medium that contains 100 mM methanol (Table 2). Additionally, a plasmid encoding pyridine nucleotide transhydrogenase from the related α-proteobacterium Rhodospirillum rubrum (pRTH2) (49) restored both enzyme activity (Table 3) and aerobic resistance to 100 mM methanol in a succinate-based minimal medium (data not shown). Transhydrogenase specific activities are reproducibly higher (two- to threefold) in cells containing either of these pnt plasmids, presumably because they are present in approximately six copies per cell (26).

To determine how the loss of transhydrogenase might be causing aerobic methanol sensitivity, growth of these mutants was monitored in liquid cultures under two conditions in which GSH-FDH activity is required (5). In the presence of O₂, the addition of 100 mM methanol to a succinate-based minimal medium has very little effect on growth of wild-type cells, while cells lacking GSH-FDH activity (Adhl1) do not grow under these conditions (Table 2). In contrast, the generation time of cells lacking transhydrogenase activity (MS65 and PntA1) is slowed ∼2.5-fold when methanol is added to a succinate-based minimal medium. When these strains were tested for the ability to use methanol as the sole photosynthetic carbon source, only Adhl1 was unable to grow (Table 2). The fact that cells lacking transhydrogenase display different growth phenotypes when either cometabolizing methanol with succinate under aerobic conditions or utilizing methanol as a sole carbon source under photosynthetic conditions is consistent with the hypothesis that these mutants do not have the same defect in formaldehyde oxidation as those that lack GSH-FDH activity.

Cells lacking transhydrogenase activity are capable of formaldehyde oxidation.

The ability of cells lacking transhydrogenase activity (MS65 and PntA1) to use methanol as a sole photosynthetic carbon source (described above) suggests that they are capable of oxidizing formaldehyde. To test this, whole-cell NMR was used to compare the metabolic fate of formaldehyde in wild-type cells and a representative pntA mutant (MS65). The fate of [¹³C]formaldehyde was monitored in photosynthetic cultures, since these cells must acquire all of their carbon skeletons by the oxidation of this one-carbon compound under these conditions (5). When ¹³C-labeled formaldehyde was added to a suspension of photosynthetic cells, production of formate is observed within 60 min after formaldehyde addition in both wild-type cells and MS65. After 240 min, formaldehyde was not detectable in cell suspensions of either wild-type cells or MS65 (Fig. 4). Thus, cells that lack transhydrogenase activity do not have an observable defect in formaldehyde oxidation under photosynthetic conditions.

FIG. 4.

Metabolic fate of formaldehyde in wild-type cells (Ga [left panels]) and those lacking transhydrogenase activity (MS65 [right panels]). Metabolism of [¹³C]formaldehyde was monitored after its addition to suspensions of cells that were grown photosynthetically with methanol as the sole carbon source. Spectra were collected at the time points indicated after the addition of formaldehyde. All peaks were normalized with TMS (0 ppm) as an internal standard and assigned by using appropriate control compounds. HCHO, formaldehyde; HCOO⁻, formate; HCO₃⁻, bicarbonate; CO₂, carbon dioxide; DMSO, dimethyl sylfoxide.

When aerobically grown cells were analyzed, there was also no discernable defect in the ability of MS65 to metabolize ¹³C-labeled formaldehyde when compared to wild-type cells (data not shown). In considering the metabolic fate of formaldehyde in aerobically grown cells, it should be noted that the cell suspensions used for whole-cell NMR (10¹¹ cells/ml) are probably O₂ limited, even though they were aerated during this analysis. As a consequence, it was difficult to assess if cells lacking transhydrogenase activity have a significant defect in formaldehyde oxidation when exposed to O₂ tensions that they are likely to encounter on plates or in aerated liquid cultures.

Cells lacking transhydrogenase activity appear to require an additional means to synthesize NADPH.

Since the loss of transhydrogenase activity did not block formaldehyde oxidation, additional experiments were performed to determine a cause for the methanol sensitivity of mutants that lack this enzyme. Pyridine nucleotide transhydrogenase is hypothesized to be a source of NADPH for bacteria under aerobic growth conditions (22, 40). If the lack of transhydrogenase activity resulted in the loss of a major source of NADPH, then these cells might be expected to increase levels of alternative NADPH-producing enzymes to compensate for a limitation for NADPH. To investigate this possibility, the specific activities of two other enzymes that can generate NADPH in R. sphaeroides, isocitrate dehydrogenase and glucose-6-phosphate dehydrogenase (12, 15, 45), were measured in wild-type cells and those that lack transhydrogenase activity (MS65). When grown aerobically, the specific activities of each of these enzymes were approximately two- to threefold higher in MS65 cells than those in wild-type cells (Table 4). In contrast, the specific activities of isocitrate dehydrogenase and glucose-6-phosphate dehydrogenase in cells that lack GSH-FDH activity (Adhl1) were comparable to those in wild-type cells (Table 4). This suggests that the observed increases of these NADPH-producing enzymes are not common to all strains that exhibit aerobic methanol sensitivity and that the specific activity of these enzymes is increased in MS65 in order to compensate for a defect in NADPH production.

TABLE 4.

Specific activities of NADPH-producing enzymes in wild-type cells (Ga) and those lacking GSH-FDH (Adhl1) or transhydrogenase (MS65) activity

Strain	Sp acta (μmol of NADP+ reduced/min/mg of protein)
	Isocitrate dehydrogenase	Glucose-6-phosphate dehydrogenase
Ga	69.6 ± 3.3	19.2 ± 4.7
Adhl1	62.0 ± 5.6	20.1 ± 3.5
MS65	151.4 ± 12.6	54.1 ± 9.5

^aData are averages of at least three independent experiments.

If strains lacking transhydrogenase activity were methanol sensitive due to the loss of a major source of NADPH synthesis, then an alternative source of this reduced pyridine nucleotide might rescue this phenotype. In R. sphaeroides, glucose is metabolized exclusively by the Entner-Doudoroff pathway (15, 45). The R. sphaeroides glucose-6-phosphate dehydrogenase is NADPH linked; therefore, 1 mol of NADPH is produced for each mol of glucose that is oxidized by this enzyme. When glucose was substituted for succinate as the sole carbon source, both wild-type cells and those lacking transhydrogenase activity (MS65 and PntA1) grew at comparable rates in the presence of methanol (Table 2). However, the use of glucose as the sole aerobic carbon source did not allow cells that lack GSH-FDH activity to grow in the presence of methanol, suggesting that glucose utilization does not result in a general suppression of aerobic methanol sensitivity (Table 2). The aerobic methanol sensitivity of cells lacking transhydrogenase activity can be rescued by providing an additional metabolic source of NADPH.

Cells lacking transhydrogenase activity are more sensitive to diamide.

In the presence of oxygen, the response to oxidative stress utilizes NADPH for maintenance of the oxidation reduction state of the GSH pool and cytoplasmic proteins. If aerobic methanol sensitivity of cells lacking transhydrogenase was caused by the loss of a major source of NADPH, R. sphaeroides cells that lack transhydrogenase activity might exhibit increased sensitivity to diamide, which is known to oxidize free sulfhydryl groups and alter the oxidation-reduction state of the GSH pool (2, 27).

When the diamide sensitivity of cells grown aerobically utilizing succinate was tested in a filter disk assay, the zone of growth inhibition was larger for cells that lack transhydrogenase activity (MS65) than for either wild-type cells or cells that lack GSH-FDH activity (Adhl1) (Fig. 5). We also tested if aerobic growth in the presence of glucose rescued the diamide sensitivity of cells that lack transhydrogenase activity. The zone of inhibition measured with MS65 when grown aerobically in a glucose-based minimal medium approached that found in wild-type cells and Adhl1 (Fig. 5). This is consistent with the hypothesis that the NADPH produced from glucose oxidation by the Entner-Doudoroff pathway can also allow cells lacking transhydrogenase activity to maintain the oxidation reduction state of both GSH and cytoplasmic proteins when diamide is present under aerobic growth conditions.

FIG. 5.

Growth inhibition by diamide. Shown are the zones of inhibition obtained with cells that were grown aerobically in either a succinate- or glucose-based minimal medium. Results are averages of at least three independent experiments; error bars show the standard deviations. From left to right, the bars show the values for wild-type cells (Ga), cells lacking GSH-FDH activity (Adhl1), and cells lacking transhydrogenase activity (MS65).

Loss of transhydrogenase activity can alter the oxidation reduction state of the GSH pool.

NADPH is used to reduce the GSSG that is produced during repair of oxidative damage (10, 36). Thus even though the addition of GSH under aerobic conditions does not restore methanol resistance to cells that lack transhydrogenase activity (data not shown), it is possible the aerobic methanol sensitivity of these mutants might reflect an impairment in GSSG reduction to GSH. To test if the loss of transhydrogenase activity resulted in a detectable alteration in the oxidation reduction state of the GSH pool, we measured the levels of GSH and GSSG. The total GSH pool (GSH plus GSSG) was comparable in wild-type cells, those lacking GSH-FDH activity (Adhl1), or those lacking transhydrogenase activity (MS65) in a succinate-based minimal medium (Table 5). However, there was approximately two- to threefold more GSSG in MS65 than in either a wild-type strain or Adhl1 when grown aerobically in a succinate-based minimal medium (Table 4). This has the result of changing the half-cell reduction potential of the GSSG-2GSH couple from −223 mV in wild-type cells to −213 mV in MS65 (Table 5). In contrast, both the size of the GSH pool, the levels of GSSG, and reduction potential of the GSSG-2GSH half-cell were comparable when either wild-type, Adhl1, or MS65 cells were grown aerobically in a glucose-based minimal medium (Table 5), indicating that glucose utilization provides a means for cells lacking transhydrogenase activity to maintain the oxidation reduction state of the GSH pool.

TABLE 5.

Oxidation reduction state of the GSH pool of wild-type cells (Ga) and those lacking GSH-FDH (Adhl1) or transhydrogenase (MS65) activity

Strain	Succinate utilizationa				Glucose utilizationa
	GSH (μmol of GSH/g [dry wt])	GSSG (μmol of GSSG/g [dry wt])	GSH/GSSG ratio	Ehcb(mV)	GSH (μmol of GSH/g [dry wt])	GSSG (μmol of GSSG/g [dry wt])	GSH/GSSG ratio	Ehcb(mV)
Ga	3.44 ± 0.35	0.011 ± 0.003	321 ± 47	−223	3.95 ± 0.3	0.009 ± 0.001	441 ± 39	−229
Adhl1	4.50 ± 0.18	0.012 ± 0.001	372 ± 13	−229	4.48 ± 0.4	0.010 ± 0.001	433 ± 42	−230
MS65	4.22 ± 0.39	0.035 ± 0.002	120 ± 11	−213	4.14 ± 0.48	0.011 ± 0.002	386 ± 34	−228

^aData are the averages of GSH and GSSG levels from three separate cultures.

^bE_hc = −240 − (59.1/2) log ([GSH]²/[GSSG]) mV at 25°C (pH 7.0) (41).

To test if this alteration in the oxidation reduction state of the GSH pool might be caused by a difference in expression of the NADPH-linked, GSSG reductase, we measured the specific activity of this enzyme in wild-type cells and those that lacked transhydrogenase activity (MS65). The specific activity of GSSG reductase was similar in a wild-type strain (81.1 μmol of NADPH oxidized/min/mg of protein) and a transhydrogenase mutant (73.4 μmol of NADPH oxidized/min/mg of protein) when assayed in crude cell extracts. Therefore, we think it is unlikely that the loss of transhydrogenase activity alters the GSH oxidation reduction state by affecting levels of the enzyme GSSG reductase. Rather, we favor a model in which the loss of transhydrogenase alters the capacity for GSSG reduction by enzymes like GSSG reductase that use NADPH as a source of reducing power.

When these data are considered together, it appears that the loss of transhydrogenase activity affects the oxidation reduction state of the GSH pool under conditions in which this enzyme is a major source of NADPH (i.e., during aerobic growth in a succinate-based minimal medium). This alteration could result in sensitivity to methanol in the presence of O₂, since these cells require GSH for both formaldehyde oxidation by the GSH-FDH-dependent pathway (5) and to reduce intramolecular disulfide bonds in cytoplasmic proteins (10). It also appears that growth conditions that provide an independent source of NADPH (i.e., aerobic growth in the presence of glucose) or those in which there is a lower demand for reducing oxidatively damaged proteins (photosynthetic growth under anaerobic conditions in the light) provide sufficient NADPH so that formaldehyde metabolism by the GSH-FDH-dependent pathway has no significant effect on growth (see Discussion).

DISCUSSION

These experiments are part of an effort to identify gene products that contribute to the GSH-dependent oxidation of formaldehyde in R. sphaeroides Homologues of R. sphaeroides GSH-FDH are present in a wide variety of bacteria, microbes, plants, and animals (48), so it is likely that proteins that contribute to the GSH-dependent oxidation of formaldehyde in R. sphaeroides will play a similar role in a variety of cells. In this bacterium, the aerobic methanol sensitivity of a mutant lacking GSH-FDH activity is correlated with a defect in formaldehyde oxidation. However, analysis of methanol-sensitive mutants lacking transhydrogenase activity showed that this phenotype was not always associated with a block in formaldehyde oxidation. Rather, it appears that the loss of pyridine nucleotide transhydrogenase can lead to metabolic and growth defects when there is a high demand for GSH in multiple metabolic pathways. The major findings and new questions posed by our observations are presented below.

Transhydrogenase is proposed to provide prokaryotic and eukaryotic cells with a means to transfer reducing equivalents between NADH and NADPH (22, 23, 40). The properties of an R. sphaeroides strain that lacks transhydrogenase activity suggest that this enzyme might have a broader role in this species than in other bacteria where mutants in this enzyme have been analyzed.

The behavior of R. sphaeroides cells that lack transhydrogenase in a succinate-based minimal medium is in some ways reminiscent of the slow growth that is observed when an E. coli strain lacking both transhydrogenase and glucose-6-phosphate dehydrogenase uses glucose as the sole aerobic carbon source (20). It was suggested that E. coli cells lacking these two enzymes grew more slowly in the presence of glucose because loss of these two NADPH-producing enzymes leads to a limitation for NADPH, which is needed for both cellular biosynthesis and to combat oxidative stress under aerobic conditions (20). In contrast, E. coli cells lacking either transhydrogenase alone or this enzyme and glucose-6-phosphate dehydrogenase display wild-type growth rates when utilizing a respiratory carbon source like succinate in the presence of oxygen. This is not the case in R. sphaeroides, since loss of transhydrogenase is sufficient to reduce the growth rate by ∼20% with succinate as an aerobic carbon source. It is possible that a diminished ability of R. sphaeroides to generate NADPH by transhydrogenase-independent routes accounts for differences in the observed growth phenotypes. Another less likely possibility is that there is a greater need for the NADPH that is produced by transhydrogenase activity in R. sphaeroides than in E. coli.

Under anaerobic conditions in the light (i.e., when growing photosynthetically), there is also a reproducible increase in the doubling time (∼40%) of R. sphaeroides strains lacking transhydrogenase activity when succinate is used as the sole carbon source. However, growth of R. sphaeroides cells lacking transhydrogenase activity is similar to that of its wild-type counterpart when methanol is used as the sole photosynthetic carbon source. It is not known why the loss of transhydrogenase activity would produce a detectable growth phenotype when succinate, and not methanol, is used as the sole photosynthetic carbon source. One possibility is that the additional reducing power produced during assimilation of formaldehyde under photosynthetic conditions (5) could provide a way to generate NADPH independent of transhydrogenase. There are no other reports that asses the physiological role of transhydrogenase when cells utilize nonfermentable carbon sources under anaerobic growth conditions, so our remaining remarks will focus on providing a working model to rationalize the metabolic consequences of losing this enzyme activity under aerobic conditions in R. sphaeroides.

The ∼2.5-fold slower growth rate of a transhydrogenase mutant relative to wild-type cells in media containing 100 mM methanol suggests that the addition of this one-carbon compound exacerbates the defect caused by the loss of this enzyme. Our data lead us to propose that the loss of transhydrogenase removes a major source of NADPH that is required for the function of the GSH-FDH pathway and other GSH-dependent processes in the presence of oxygen.

Several observations support the notion that the transhydrogenase mutant is lacking a major source of NADPH when it is grown in a succinate-based minimal medium under aerobic conditions. The specific activities of other enzymes that produce NADPH are increased in a transhydrogenase mutant compared to those in the wild type. In addition, the use of glucose as the sole aerobic carbon source restored wild-type growth rates to transhydrogenase mutants in the presence of methanol. Since glucose is metabolized by the Entner-Doudoroff pathway in R. sphaeroides, it provides a metabolic source of NADPH to compensate for the loss of transhydrogenase activity in the presence of oxygen.

This model can also explain why formaldehyde oxidation by a GSH-dependent pathway has a negative effect on aerobic growth of cells lacking transhydrogenase. NADPH, the product of transhydrogenase activity, is also a source of reducing power for enzymes such as GSSG reductase and thioredoxin, which maintain the oxidation reduction state of GSH in the cytoplasm (3, 4, 10, 36, 37). Under aerobic conditions, GSH is used to reduce intramolecular disulfide bonds in cytoplasmic proteins (10). If the loss of transhydrogenase removes a major source of NADPH, then one would expect this strain to have increased sensitivity to oxidative stress agents that cause disulfide bond formation within cytoplasmic proteins (27, 38). As predicted, cells lacking transhydrogenase are more sensitive to diamide, a sulfhydryl-oxidizing agent that is known to stimulate disulfide bond formation in cytoplasmic proteins of other bacteria. Even in the absence of diamide, cells lacking transhydrogenase activity have a larger pool of GSSG than wild-type cells when grown aerobically in a succinate-based medium. Consistent with this model, aerobic growth in the presence of glucose repairs the sensitivity of cells lacking transhydrogenase to diamide, and it restores the oxidation-reduction state of the GSH pool to that found in wild-type cells, presumably due to the production of NADPH by the Entner-Doudoroff pathway.

Based on these results, one might expect to have obtained methanol-sensitive mutants that have Tn5 insertions that disrupt genes encoding GSSG reductase or those involved in GSH biosynthesis. To date, we have not isolated any mutants that either map to these loci or are repaired by the addition of GSH to growth media. In E. coli, loss of GSSG reductase alone does not significantly alter the oxidation reduction state of the GSH pool, and it has been shown that other disulfide-reducing activities, such as thioredoxin, also play a role in maintaining the oxidation reduction state of the GSH pool (4, 10). If thioredoxin can also play such a role in R. sphaeroides, then loss of GSSG reductase would not significantly affect the oxidation reduction state of the GSH pool and would not lead to methanol sensitivity. In R. sphaeroides, it appears that thioredoxin is essential, suggesting that GSH and presumably pathways that maintain the oxidation reduction state of such a low-molecular-weight cytoplasmic thiol might be necessary for aerobic growth (35). If this is true, it might also explain why we did not isolate strains carrying mutations within genes that are involved in GSH biosynthesis during this study.

In this model, the metabolic defect of cells lacking transhydrogenase is exacerbated by the need to oxidize formaldehyde by a GSH-dependent pathway when oxygen is present. When cells cometabolize methanol with succinate in the presence of oxygen, GSH is needed in large amounts for several pathways. Under these conditions, one can imagine that a significant portion of the GSH pool is needed to support a high flux of carbon skeletons through the GSH-FDH pathway, especially since whole-cell NMR shows that it takes the equivalent of approximately one doubling time for this suspension to oxidize ∼10 mM formaldehyde. When a significant portion of the GSH pool is needed for formaldehyde metabolism, it is easy to envision why cells lacking transhydrogenase will have even greater difficulty combating the damage that results from an aerobic lifestyle.

In contrast, when cells use methanol as the sole photosynthetic carbon source, there appears to be no or little effect of the loss of transhydrogenase, presumably because oxidative damage to proteins is minimal in the absence of oxygen. Since large amounts of GSH should not be needed to reduce cytoplasmic disulfide bonds under these conditions, a high rate of flux through the GSH-dependent pathway for formaldehyde oxidation does not slow growth of cells lacking transhydrogenase. Therefore, reduced demands for GSH in the absence of oxygen allow cells that lack transhydrogenase activity to grow with wild-type doubling times, even when they need a need a high rate of flux through the GSH-dependent pathway for formaldehyde oxidation of R. sphaeroides.

When considering our data, it is also conceivable that the increase in GSSG levels might inhibit activity of GSH-FDH, thus limiting the oxidation of formaldehyde in a transhydrogenase mutant when oxygen is present at high levels. However, it has been shown that concentrations of GSSG as high as 7 mM (which is well above both the apparent K_m of this enzyme for GSH and the intracellular levels of GSSG that we have measured) do not inhibit GSH-FDH activity in vitro, even after preincubation of the enzyme with this compound for 90 min (48). Therefore, it does not seem likely that GSSG acts as an effective substrate analog of GSH or as a negative allosteric regulator of GSH-FDH activity in cells that contain or lack transhydrogenase activity.

In summary, we have shown that loss of pyridine nucleotide transhydrogenase activity results in a conditional growth defect of R. sphaeroides. The aerobic growth defect of this strain appears to reflect the fact that the product of transhydrogenase activity, NADPH, is required to maintain the oxidation reduction state of the cytoplasmic GSH pool. This growth defect is exacerbated under conditions in which there are multiple demands for GSH in the cell (i.e., formaldehyde oxidation and the repair of oxidatively damaged proteins). This is unlike the situation in E. coli, where the loss of transhydrogenase alone does not result in a discernible growth phenotype under the laboratory conditions that have been tested (20). In the future, it will be interesting to see if other mutations that could interfere with the oxidation reduction state of the GSH pool produce similar phenotypes and to determine if the loss of transhydrogenase has a similar effect on the numerous other prokaryotic and eukaryotic cells that contain a GSH-dependent pathway for formaldehyde oxidation.