Choline degradation in Paracoccus denitrificans: identification of sources of formaldehyde

Trusha Parekh; Marcus Tsai; Stephen Spiro

doi:10.1128/jb.00081-24

. 2024 Mar 19;206(4):e00081-24. doi: 10.1128/jb.00081-24

Choline degradation in Paracoccus denitrificans: identification of sources of formaldehyde

Trusha Parekh ¹, Marcus Tsai ¹, Stephen Spiro ^1,^✉

Editor: George O'Toole²

PMCID: PMC11025334 PMID: 38501746

ABSTRACT

Paracoccus denitrificans is a facultative methylotroph that can grow on methanol and methylamine as sole sources of carbon and energy. Both are oxidized to formaldehyde and then to formate, so growth on C1 substrates induces the expression of genes encoding enzymes required for the oxidation of formaldehyde and formate. This induction involves a histidine kinase response regulator pair (FlhSR) that is likely triggered by formaldehyde. Catabolism of some complex organic substrates (e.g., choline and L-proline betaine) also generates formaldehyde. Thus, flhS and flhR mutants that fail to induce expression of the formaldehyde catabolic enzymes cannot grow on methanol, methylamine, and choline. Choline is oxidized to glycine via glycine betaine, dimethylglycine, and sarcosine. By exploring flhSR growth phenotypes and the activities of a promoter and enzyme known to be upregulated by formaldehyde, we identify the oxidative demethylations of glycine betaine, dimethylglycine, and sarcosine as sources of formaldehyde. Growth on glycine betaine, dimethylglycine, and sarcosine is accompanied by the production of up to three, two, and one equivalents of formaldehyde, respectively. Genetic evidence implicates two orthologous monooxygenases in the oxidation of glycine betaine. Interestingly, one of these appears to be a bifunctional enzyme that also oxidizes L-proline betaine (stachydrine). We present preliminary evidence to suggest that growth on L-proline betaine induces expression of a formaldehyde dehydrogenase distinct from the enzyme induced during growth on other formaldehyde-generating substrates.

IMPORTANCE

The bacterial degradation of one-carbon compounds (methanol and methylamine) and some complex multi-carbon compounds (e.g., choline) generates formaldehyde. Formaldehyde is toxic and must be removed, which can be done by oxidation to formate and then to carbon dioxide. These oxidations provide a source of energy; in some species, the CO₂ thus generated can be assimilated into biomass. Using the Gram-negative bacterium Paracoccus denitrificans as the experimental model, we infer that oxidation of choline to glycine generates up to three equivalents of formaldehyde, and we identify the three steps in the catabolic pathway that are responsible. Our work sheds further light on metabolic pathways that are likely important in a variety of environmental contexts.

KEYWORDS: choline, glycine betaine, formaldehyde, dimethylglycine, two-component regulatory system, sarcosine

INTRODUCTION

Paracoccus denitrificans is a metabolically versatile Alphaproteobacterium that is frequently detected in terrestrial and aquatic environments, in waste treatment reactors, and in association with animals and plants (1 –5). The organism can grow on C1 compounds (methanol and methylamine) as sole sources of carbon and energy and has been used as a model for studies of the genetics and biochemistry of methylotrophy (6, 7). Both methanol and methylamine are oxidized to formaldehyde by pyrroloquinoline quinone-dependent periplasmic enzymes. The formaldehyde is internalized, probably by a specific transport system (8), and is oxidized to formate by a cytoplasmic glutathione (GSH)-dependent formaldehyde dehydrogenase (GD-FALDH) (7, 9). P. denitrificans also has a gene (Pden_1186, hpbM) encoding a predicted GSH-independent formaldehyde dehydrogenase (10, 11). Formate is oxidized to carbon dioxide, which is assimilated by the Calvin cycle (11, 12), although other species in the Paracoccus genus may use the serine cycle (13).

Transcription of the genes required for the oxidation of methanol, methylamine, and formaldehyde is regulated by two two-component systems comprising histidine kinases (FlhS and MoxY) and their cognate response regulators (FlhR and MoxX, respectively). MoxXY (also called MxaXY) is required for transcription of the genes encoding methanol dehydrogenase and associated proteins, and moxXY mutants have a growth defect on methanol but not methylamine (14). FlhSR-deficient mutants are unable to grow on both methanol and methylamine (15), and the FlhSR proteins are believed to activate transcription of the operons encoding the methanol, methylamine, and formaldehyde dehydrogenases, and MoxXY (7). The signal that triggers the FlhSR signal transduction pathway has not been identified biochemically, but it is suggested that both FlhS and MoxX respond to formaldehyde (7). The DeepTMHMM algorithm (16) predicts FlhS to be located in the cytoplasm, which is, therefore, the likely site of formaldehyde sensing.

FlhSR-deficient mutants are also unable to grow on choline as a source of carbon and energy (15). The explanation for this phenotype is that the oxidation of choline to glycine is accompanied by the production of formaldehyde, and the flhSR mutants are poisoned because they are unable to upregulate expression of the formaldehyde dehydrogenase. Thus, ectopic expression of the GD-FALDH rescues the choline growth defect of flhS and flhR mutants (15). Choline catabolism in P. denitrificans has been variously suggested to be accompanied by the production of “several” (17), two (18) or three (19) equivalents of formaldehyde.

Under aerobic conditions, choline is degraded in the cytoplasm through betaine aldehyde, glycine betaine, dimethylglycine. and sarcosine to glycine (Fig. 1; 20). Each of the three steps from glycine betaine to glycine involves the removal of one methyl group and is a potential source of formaldehyde. Glycine betaine can be oxidatively demethylated by a glycine betaine monooxygenase that generates dimethylglycine and formaldehyde (21, 22). In other species, a glycine betaine:homocysteine methyl transferase donates the methyl group to homocysteine and so makes methionine rather than formaldehyde (23, 24). Dimethylglycine can be oxidatively demethylated by a dimethylglycine dehydrogenase (or oxidase). These are tetrahydrofolate (THF)-linked enzymes that generate methylene-THF (rather than formaldehyde) as the reaction product (25). There is also a dimethylglycine demethylase that has not been characterized biochemically but is a potential source of formaldehyde (20, 21). Sarcosine is oxidized either by a monomeric or a tetrameric sarcosine oxidase. The former generates formaldehyde as the reaction product, while the tetrameric sarcosine oxidase is THF dependent and so makes methylene-THF (26). The THF-dependent dimethylglycine and sarcosine oxidases can make formaldehyde as a reaction product in vitro in the absence of THF (25, 26); whether these enzymes are a source of formaldehyde in vivo is not known. Depending upon the combination of enzymes used and whether or not the THF-dependent enzymes are sources of formaldehyde in vivo, the conversion of choline to glycine may be accompanied by the formation of between zero and three equivalents of formaldehyde (20, 24). P. denitrificans also has a characterized pathway for the degradation of proline betaine, which is predicted to generate two equivalents of formaldehyde through the action of proline betaine and methyl proline demethylases (10).

Fig 1 — Pathways for catabolism of choline, stachydrine (L-proline betaine), and C1 substrates in *Paracoccus denitrificans*. Metabolic pathways are proposed based on previous literature (see citations in the text), the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (27), and results presented in this paper. Enzymes that are known or likely sources of formaldehyde are highlighted in orange. Solid arrows denote experimentally supported reactions; dashed lines are predictions based on the genome annotation. Based on the data presented in this paper, we suggest that formaldehyde generated from glycine betaine and dimethylglycine is oxidized predominantly by GD-FALDH, while the GSH-independent enzyme oxidizes formaldehyde generated from L-proline betaine and methyl-L-proline. The genes encoding the enzymes shown in this diagram are depicted in Fig. S1.

While studying the regulation of formaldehyde metabolism in P. denitrificans, we became interested to identify the source(s) of the formaldehyde generated during choline catabolism. Reconstructing the pathway from the genome sequence is challenging for several reasons: (1) For some steps, there are multiple candidate enzymes encoded in the genome, (2) some potentially relevant genes are duplicated, and (3) definitively assigning biochemical functions to gene products is not always possible. Therefore, we have adopted an empirical approach (informed by the genome sequence) to determine the likely pathway for choline degradation in P. denitrificans and the source(s) of formaldehyde. To do so, we have taken advantage of the fact that flhSR mutants have a defect for growth on substrates that generate formaldehyde. We present evidence to indicate that the major sources of formaldehyde are two orthologous monooxygenases that oxidatively demethylate glycine betaine to form dimethylglycine and the subsequent oxidative demethylations of both dimethylglycine and sarcosine.

RESULTS

Formaldehyde is a by-product of the oxidation of glycine betaine and dimethylglycine

It has been reported previously that flhS and flhR mutants are unable to grow on solid media containing choline as the sole source of carbon and energy (15). We have reconstructed unmarked deletions in flhS and flhR and confirmed that these mutants are unable to grow on solid minimal medium containing choline (Fig. S4). In liquid media, the flhS and flhR mutants grow normally to an optical density of around 0.3 before arresting (Fig. 2). This pattern is consistent with the accumulation of a toxic intermediate (likely to be formaldehyde).

Fig 2 — Growth phenotypes of Δ*flhS* and Δ*flhR* mutants. Wild-type *P. denitrificans* and Δ*flhS* and Δ*flhR* deletion mutants were grown in minimal medium containing either (A) choline, (B) glycine betaine, (C) dimethylglycine, or (D) sarcosine as sole sources of carbon and energy (substrate concentrations were adjusted such that carbon was equimolar). Growth data are the mean of three biological replicates, and the error bars represent standard deviations.

We established that P. denitrificans grows well on intermediates of the choline degradation pathway: glycine betaine, dimethylglycine, and sarcosine (Fig. 2; Table 1). In liquid media, the flhS and flhR mutants show growth defects on glycine betaine similar to the choline phenotypes (Fig. 2), whereas both mutants grow normally on dimethylglycine and sarcosine. The patterns are similar on solid media, except that the flhS and flhR mutants exhibit a partial defect for growth with dimethylglycine as the substrate (Figs. S5 to S7). We could detect significant concentrations of formaldehyde in the supernatants of flhS and flhR mutants grown on choline, glycine betaine, and dimethylglycine (Table 1). Thus far, we conclude that formaldehyde is generated as a product of the oxidation of both glycine betaine and dimethylglycine, but that the amount of formaldehyde accumulated in dimethylglycine-grown cells is insufficient to impose a phenotype on the flhS and flhR mutants.

TABLE 1.

Growth phenotypes on solid media and formaldehyde accumulation in liquid culture supernatants

	Growth phenotype^a						Formaldehyde accumulation^b
	Wild type	ΔflhS	ΔflhR	ΔgbcAB	ΔhpbC2B2	ΔgbcAB ΔhpbC2B2	Wild type	ΔflhS	ΔflhR
Succinate	+++	+++	+++	+++	+++	+++	−	−	−
Choline	+++	−	+	+++	+	−	−	++	++
Glycine betaine	+++	−	+	+++	−	−	−	++	++
Dimethylglycine	+++	++	++	+++	+++	+++	−	+	+
Sarcosine	+++	+++	++	+++	+++	+++	−	−	−
Stachydrine	+++	+++	+++	+++	+++	+++	−	−	−

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^{^a}

Streak plates were incubated for approximately 72 h at 30°C. Images of representative plates and of complementation tests are in Fig. S4 to S8.

^{^b}

−, not detectable, below the detection limit of the assay (0.6 µM/OD unit); +, detectable in the range 10–13 μM/OD unit; ++, detectable in the range 13–28 μM/OD unit.

All growth phenotypes on solid (Figs. S4 to S7) and liquid (not shown) media associated with the flhS and flhR deletions could be complemented by expression of the corresponding genes in trans from recombinant plasmids.

Formaldehyde generated from choline degradation activates the FlhSR signaling pathway

It has been proposed that formaldehyde generated during choline degradation activates transcription of genes required for formaldehyde oxidation (15). To confirm that choline-derived formaldehyde activates FlhSR and to determine the source(s) of formaldehyde, we took advantage of the fact that the promoter of the structural gene (mxaF) encoding the methanol dehydrogenase is activated by formaldehyde, by a mechanism that requires FlhSR (7, 15). We constructed an mxaF-lacZ promoter–reporter fusion and confirmed the expected low activity in cells grown on succinate and >10-fold higher activity in cells grown on choline (Table 2). The promoter fusion also had >10-fold higher activities in cells grown on glycine betaine and dimethylglycine, confirming that degradation of these substrates generates formaldehyde (Table 2). In sarcosine-grown cells, mxaF promoter activity was intermediate between that measured in succinate- and dimethylglycine-grown cells, suggesting that sarcosine oxidation is also a source of formaldehyde. In all cases, induction of the mxaF promoter was eliminated in flhS and flhR mutants, and introduction of the corresponding genes on plasmids complemented or partially complemented the mutant phenotypes. These patterns of mxaF promoter activity correlate well with the formaldehyde accumulation data and mutant growth phenotypes and further suggest that degradation of glycine betaine, dimethylglycine, and sarcosine produces sufficient formaldehyde to switch on an FlhSR-activated promoter.

TABLE 2.

β-Galactosidase activities expressed from the mxaF-lacZ reporter fusion

	β-Galactosidase^a
	Wild type	ΔflhS	ΔflhR	ΔgbcAB	ΔhpbC2B2	ΔgbcAB ΔhpbC2B2
Succinate	41 (2)	39 (3)	40 (1)	ND^b	ND	ND
Choline	574 (84)	20 (17)	30 (36)	438 (41)	134 (16)	65 (9)
Glycine betaine	646 (61)	11 (9)	2 (1)	258 (49)	118 (4)	83 (5)
Dimethylglycine	413 (115)	22 (4)	44 (33)	319 (102)	77 (20)	73 (7)
Sarcosine	200 (4)	30 (25)	51 (23)	81 (18)	104 (4)	107 (2)
Stachydrine	434 (101)	16 (5)	28 (18)	ND	ND	ND

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^{^a}

β-Galactosidase activity was measured in cells grown on the indicated carbon sources. Units of activity are as defined by Miller (28). β-Galactosidase activities are the means of triplicate assays from three independently grown cultures with the standard deviation in parentheses.

^{^b}

ND, not done.

We also measured the activity of the GSH-dependent formaldehyde dehydrogenase, the expression of which is believed to be activated by FlhSR (15). Similar to the pattern of mxaF-lacZ activity, GD-FALDH activity was low in extracts of cells grown on succinate, high in cells grown on choline, glycine betaine, and dimethylglycine, and intermediate in cells grown on sarcosine (Table 3). Note that if each oxidative demethylation generates one equivalent of formaldehyde, then growth on glycine betaine, dimethylglycine, and sarcosine would be accompanied by production of three, two, and one equivalents of formaldehyde, respectively. Accumulation of less formaldehyde is consistent with the absence of flhSR mutant phenotypes on dimethylglycine and sarcosine and the evidence for reduced FlhSR activity in cells grown on these substrates, compared to glycine betaine and choline. Induction of GD-FALDH activity was lost in flhS and flhR mutants (Table 3), and mutant phenotypes could be complemented by expression of the corresponding genes from a plasmid (data not shown).

TABLE 3.

GSH-dependent formaldehyde dehydrogenase activities

	GSH-dependent formaldehyde dehydrogenase (μ mol NADH/min/mg protein)^a
	Wild type	ΔflhS	ΔflhR	ΔgbcAB	ΔhpbC2B2	ΔgbcAB ΔhpbC2B2
Succinate	75 (8)	6 (2)	27 (13)	ND^b	ND	ND
Choline	361 (107)	53 (7)	39 (7)	95 (35)	264 (60)	25 (12)
Glycine betaine	539 (73)	9 (8)	21 (12)	82 (36)	207 (99)	26 (13)
Dimethylglycine	313 (62)	37 (18)	29 (3)	244 (106)	145 (49)	53 (31)
Sarcosine	135 (39)	16 (10)	14 (6)	140 (90)	32 (18)	20 (7)
Stachydrine	63 (23)	34 (5)	11 (7)	ND^b	ND	ND

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^{^a}

GSH-dependent formaldehyde dehydrogenase activities are the means of assays from three independently grown cultures with the standard deviation in parentheses.

^{^b}

ND, not done.

Our measurements of mxaF promoter activity and GD-FALDH enzyme activity both implicate the oxidation of glycine betaine, dimethylglycine, and sarcosine as the sources of formaldehyde. The predicted dimethylglycine dehydrogenase encoded by Pden_1940 is likely to be required for dimethylglycine catabolism, and there are at least two candidate sarcosine oxidases encoded in the genome (Fig. S1). We focused on identifying the enzyme(s) required for growth on glycine betaine.

Overlapping functions of enzymes processing glycine betaine

In Pseudomonas aeruginosa and Pseudomonas syringae, a two-gene operon required for growth on glycine betaine encodes a predicted redox enzyme, GbcAB (21, 29). In Chromohalobacter salexigens, this enzyme has been characterized as a formaldehyde-generating glycine betaine monooxygenase (named BmoAB) comprising a Rieske non-heme iron oxygenase (BmoA) and a reduced nicotinamide adenine dinucleotide (NADH)-dependent flavin reductase, BmoB (22). In P. denitrificans, the predicted proteins most similar to GbcAB are encoded by Pden_4896 and 4897. We constructed a deletion mutant lacking these two genes and found only a modest defect for growth on choline and glycine betaine (Fig. 3).

Fig 3 — Growth phenotypes of Δ*gbcAB*, Δ*hpbC2B2,* and Δ*gbcAB* Δ*hpbC2B2* mutants. Wild-type *P. denitrificans* and Δ*gbcAB*, Δ*hpbC2B2,* and Δ*gbcAB* Δ*hpbC2B2* mutants were grown in minimal medium containing (A) choline, (B) glycine betaine, (C) dimethylglycine, and (D) sarcosine as sole sources of carbon and energy (substrate concentrations were adjusted such that carbon was equimolar). Growth data are the mean of three biological replicates, and the error bars represent standard deviations.

A previous study of stachydrine catabolism in P. denitrificans characterized two additional members of the GbcAB family (10). HpbC1B1 (encoded by Pden_1188–89) and HpbC2B2 (Pden_2831–32) are both annotated as stachydrine (L-proline betaine) demethylases (or monooxygenases). Deletion of both pairs of genes was required to eliminate growth on stachydrine, the implication being that both enzymes are able to oxidize stachydrine to methyl-L-proline (10). Since HpbC2B2 are more similar to GbcAB than HpbC1B1, we deleted the hpbC2B2 (Pden_2831–32) genes and found that the mutant exhibits markedly biphasic growth when cultured on choline and glycine betaine (Fig. 3). When subcultured into fresh medium, biphasic growth of the hpbC2B2 mutant repeated, from which we infer that the extended lag phase and biphasic pattern are not due to the accumulation of mutations. The double mutant lacking both enzymes has a null phenotype for growth on choline and glycine betaine (Fig. 3; Fig. S4 and S5), implying that both GbcAB and HpbC2B2 can oxidize glycine betaine. This further suggests that HpbC2B2 has a relaxed substrate specificity being able to act on both glycine betaine (this work) and stachydrine (10).

Catabolism of stachydrine is predicted to generate two equivalents of formaldehyde (10). We found that flhS and flhR mutants have no growth defect on stachydrine (Fig. S2 and S8), as is the case for growth on dimethylglycine (Fig. 2), which we suggest also generates two equivalents of formaldehyde. From both these growth substrates, sufficient formaldehyde is made to activate the mxaF promoter (Table 2), but insufficient to impose a phenotype on the flhSR mutants (Fig. 2; Fig. S2).

The predicted GSH-independent formaldehyde dehydrogenase is encoded by a gene (Pden_1186, hpbM) immediately downstream of the genes encoding the stachydrine demethylase HpbC1B1 and a proline betaine epimerase (10). We therefore speculated that expression of HpbM might be specifically induced by growth on stachydrine. We measured only background levels of GD-FALDH in extracts of cells grown on stachydrine but increased activities of a GSH-independent formaldehyde dehydrogenase compared to cells grown on all other substrates (Table 4). Our results are consistent with transcriptomics data that show upregulation of hbpM in cells grown on stachydrine versus cells grown on methanol (10). However, the same transcriptomics data also suggest upregulation of genes encoding the GD-FALDH by growth on stachydrine (10), which we do not see reflected in the enzyme activity.

TABLE 4.

GSH-independent formaldehyde dehydrogenase activities

	GSH-independent formaldehyde dehydrogenase (μ mol NADH/min/mg protein)^a
	Wild type	ΔflhS	ΔflhR
Succinate	4 (0.1)	2 (0.6)	3 (0.9)
Choline	5 (2)	1 (0.6)	2(0.2)
Glycine betaine	6 (2)	5 (1)	4 (0.4)
Dimethylglycine	20 (3)	7 (4)	9 (1)
Sarcosine	13 (5)	6 (4)	14 (7)
Stachydrine	74 (21)	8 (3)	28 (4)

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^{^a}

GSH-independent formaldehyde dehydrogenase activities are the means of assays from three independently grown cultures with the standard deviation in parentheses.

DISCUSSION

Considering the literature, genome annotations and metabolic predictions in the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (27), and our data, we propose a choline metabolic pathway shown in Fig. 1, comprising enzymes that are the products of the genes and operons in Fig. S1. Choline degradation is initiated with choline oxidation to betaine aldehyde by a choline dehydrogenase (BetA) and then to glycine betaine by a betaine aldehyde dehydrogenase (BetB). In P. denitrificans, these enzymes are likely encoded by Pden_1896 and Pden_1897, respectively. Upstream of these genes is Pden_1898, which is annotated to encode BetI, a repressor that likely mediates choline regulation of the betAB genes (30).

Our results indicate that the conversion of glycine betaine to dimethylglycine can be catalyzed primarily by HpbC2B2 (encoded by Pden_2832–2831) and, in its absence, by the GbcAB (Pden_4897–4896). This step is a major source of formaldehyde in the choline degradation pathway, and FlhSR signaling is required for expression of the enzyme(s) responsible for further catabolism of formaldehyde.

Conversion of dimethylglycine to sarcosine is likely catalyzed by a THF-dependent dimethylglycine dehydrogenase (encoded by Pden_1940). This enzyme can generate formaldehyde in vitro (25) and, our evidence suggests, also in vivo. In either case, our evidence indicates that this reaction does not liberate sufficient formaldehyde to inhibit growth of the flhSR mutants.

The P. denitrificans genome does not encode a predicted monomeric sarcosine oxidase but has two sets of genes annotated to encode the subunits of a tetrameric enzyme (Pden_0512, 0514–0516, and Pden_4909–4906), with the latter likely being co-expressed with a 5,10-methylene THF dehydrogenase (Pden_4905) from a shared promoter. Although the tetrameric sarcosine oxidase is THF-linked, it can make formaldehyde as a reaction product in vitro (26). Our evidence suggests that this enzyme is also a source of formaldehyde in vivo.

In this work, we have successfully used mutant phenotypes and promoter and enzyme activities as probes for the endogenous generation of formaldehyde. We infer that up to three equivalents of formaldehyde may be produced during choline degradation in P. denitrificans. Direct quantitation of this stoichiometry would require measurement of formaldehyde production. This is challenging in part because of the pathways for the onward catabolism of formaldehyde. Ultimately, isotope-based metabolic flux analysis and metabolic modeling (11) could be used to provide a complete description of the metabolic fate of the methyl groups abstracted from quaternary amines such as choline and stachydrine.

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions

Bacterial strains and plasmids used in this study are detailed in Table S1. Paracoccus denitrificans Pd1222 (31) was used as the wild-type strain. Escherichia coli S17-1 (32) was used as the host for the introduction of plasmids into P. denitrificans by conjugation (33). L broth (10 g of tryptone.liter⁻¹, 5 g of yeast extract.liter⁻¹, 5 g of NaCl.liter⁻¹) was used for the routine propagation of E. coli and P. denitrificans at 37°C or 30°C, respectively.

For phenotypic assays, cultures were grown in a defined minimal medium (34) containing sodium succinate (20 mM), methanol (50 mM), choline chloride (32 mM), glycine betaine (32 mM), dimethylglycine (40 mM), sarcosine (54.3 mM), stachydrine hydrochloride (12 mM), and L-proline (32 mM). Aerobic cultures were grown in 200 µL of minimal medium in 96-well plates shaken at 420 rpm. Growth was assayed every 2–4 h by measurement of light scatter at 650 nm (OD650). Growth was also assessed on minimal medium agar streak plates.

Genetic manipulations

Plasmid pSTBlue-1 (Novagen) was used as the vector for routine DNA manipulations. Triparental matings were performed by mixing log-phase P. denitrificans, E. coli S17-1 transformed with recombinant plasmid derivatives, and E. coli DH5α (pRK2013) in a 2:1:1 ratio and incubating on L agar for 24 h at 30°C. P. denitrificans exconjugants were selected by rifampicin and indicated antibiotic(s). For conjugation and subsequent selection, salt-free L broth and a low-salt (4 g of sodium chloride.liter⁻¹) L agar were used.

Unmarked in-frame deletions were made by allelic replacement using pK18mobsacB (35). Briefly, ~500-bp regions flanking the gene(s) to be deleted were amplified separately and then ligated together by PCR using overlapping regions added to the primers. The ligated fragment was cloned into pK18mobsacB then mobilized into P. denitrificans as described above. Selection for kanamycin resistance yielded strains in which the plasmid was integrated into the chromosome by a single homologous recombination event. Recombinants with double crossovers were then isolated on the basis of sucrose resistance and kanamycin sensitivity; deletion mutations were confirmed by PCR.

For complementation tests, genes were cloned into the broad host-range expression vector pIND4, which allows induction of expression by isopropyl β-D-1-thiogalactopyranoside (IPTG) (36). A promoter-lacZ reporter fusion was constructed by cloning 300 bp upstream of the start codon of the methanol dehydrogenase structural gene mxaF into pMP220 (37). After triparental mating into P. denitrificans, selection for recombinant pIND4 and pMP220 plasmids was done using 25 µg/mL of kanamycin or 4 µg/mL of tetracycline, respectively. PCR confirmation was done with isolated plasmids.

Formaldehyde measurement

Formaldehyde accumulation in supernatants collected from stationary phase cultures was measured using a modified NASH assay (38). Ninety-six-well plates were centrifuged at 5,000 g for 5 min at 4°C after the completion of growth measurements. Triplicate 125-µL aliquots of each supernatant were mixed with 125 µL of NASH reagent (0.3% acetic acid and 0.2% acetylacetone added freshly to a 3% ammonium acetate solution) in a clear round bottom 96-well plate and incubated at 37°C for 50 min for color development. Formaldehyde standards ranged from 0.01 to 1 mM. The plate was equilibrated to room temperature, and the absorbance at 412 nm was recorded. Concentrations are reported after normalization to the final culture densities.

Enzyme assays

β-galactosidase expressed from the mxaF promoter-lacZ fusion was assayed as described by Miller (28). Cultures (three independent replicates) were grown in 96-well plates, 0.05-1 mL samples in 0.9-mL reaction mix were made permeable with chloroform and SDS and assayed according to the standard protocol (28); the absorbance at 420 nm was measured in 96-well plates.

Glutathione-dependent formaldehyde dehydrogenase and GSH-independent formaldehyde dehydrogenase were assayed in extracts prepared from 5- to 50-mL cultures. Mid- to late-log phase cultures were harvested at 6,000 g at 4°C for 20 min, and cells were washed twice with sodium phosphate buffer (pH 7.4). The wet cell pellet was resuspended in approximately three volumes of phosphate buffer along with 2 mM dithiothreitol (DTT) and 2 mM phenylmethylsulfonyl fluoride (PMSF). The cell suspension was sonicated with a cycle of 20-s on/20-s off for 2–5 min in an ice-water bucket. The sonicated suspension was clarified by centrifugation at 16,000 × g at 4°C for 20 min.

Enzyme activity was measured at 30°C using a Cary UV spectrophotometer. The 3mL reaction mixture contained 50–200 µL of cell lysate in 100 mM sodium phosphate buffer (pH 7.4) and 1 mM NAD⁺. For the GSH-independent formaldehyde dehydrogenase, assays were started by the addition of 10 mM formaldehyde. For the GD-FALDH, formaldehyde addition was followed by 20 mM GSH. Specific activity was calculated as previously described using 6.22 as the millimolar extinction coefficient of NADH at 340 nm (39). Enzyme activities were assayed in triplicate in extracts of three independently grown cultures. Protein concentrations in cell lysates were measured with the Pierce BCA assay kit.

ACKNOWLEDGMENTS

We are grateful to Sneha Narvekar for her contributions to strain constructions.

This work was supported in part by the C. L. and Amelia A. Lundell endowment.

Footnotes

This article was submitted via the Active Contributor Track (ACT). Stephen Spiro, the ACT-eligible author, secured reviews from Gary Sawers, Martin Luther University of Halle-Wittenberg Institute of Biology/Microbiology, and David J. Richardson, University of East Anglia.

Contributor Information

Stephen Spiro, Email: stephen.spiro@utdallas.edu.

George O'Toole, Geisel School of Medicine at Dartmouth, Hanover, New Hampshire, USA.

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/jb.00081-24.

Supplemental material. jb.00081-24-s0001.pdf.

Table S1 and Figures S1 to S8.

jb.00081-24-s0001.pdf^{(1.1MB, pdf)}

DOI: 10.1128/jb.00081-24.SuF1

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5. Castelino M, Eyre S, Moat J, Fox G, Martin P, Ho P, Upton M, Barton A. 2017. Optimisation of methods for bacterial skin microbiome investigation: primer selection and comparison of the 454 versus MiSeq platform. BMC Microbiol 17:23. doi: 10.1186/s12866-017-0927-4 [DOI] [PMC free article] [PubMed] [Google Scholar]
6. Baker SC, Ferguson SJ, Ludwig B, Page MD, Richter OM, van Spanning RJ. 1998. Molecular genetics of the genus Paracoccus: metabolically versatile bacteria with bioenergetic flexibility. Microbiol Mol Biol Rev 62:1046–1078. doi: 10.1128/MMBR.62.4.1046-1078.1998 [DOI] [PMC free article] [PubMed] [Google Scholar]
7. Van Spanning RJM, de Vries S, Harms N. 2000. Coping with formaldehyde during C1 metabolism of Paracoccus denitrificans. J Mol Catal B Enzym 8:37–50. doi: 10.1016/S1381-1177(99)00065-X [DOI] [Google Scholar]
8. Köstler M, Kleiner D. 1989. Assimilation of methylamine by Paracoccus denitrificans involves formaldehyde transport by a specific carrier. FEMS Microbiol Lett 53:1–4. doi: 10.1016/0378-1097(89)90356-x [DOI] [PubMed] [Google Scholar]
9. Ophem PW, Duine JA. 1994. NAD- and co-substrate (GSH or factor)-dependent formaldehyde dehydrogenases from methylotrophic microorganisms act as a class III alcohol dehydrogenase. FEMS Microbiol Lett 116:87–93. doi: 10.1111/j.1574-6968.1994.tb06680.x [DOI] [Google Scholar]
10. Kumar R, Zhao S, Vetting MW, Wood BM, Sakai A, Cho K, Solbiati J, Almo SC, Sweedler JV, Jacobson MP, Gerlt JA, Cronan JE. 2014. Prediction and biochemical demonstration of a catabolic pathway for the osmoprotectant proline betaine. mBio 5:e00933-13. doi: 10.1128/mBio.00933-13 [DOI] [PMC free article] [PubMed] [Google Scholar]
11. Bordel S, Martín-González D, Börner T, Muñoz R, Santos-Beneit F. 2024. Genome-scale metabolic model of the versatile bacterium Paracoccus denitrificans Pd1222. mSystems 9:e0107723. doi: 10.1128/msystems.01077-23 [DOI] [PMC free article] [PubMed] [Google Scholar]
12. Cox RB, Quayle JR. 1975. The autotrophic growth of Micrococcus denitrificans on methanol. Biochem J 150:569–571. doi: 10.1042/bj1500569 [DOI] [PMC free article] [PubMed] [Google Scholar]
13. Dziewit L, Czarnecki J, Prochwicz E, Wibberg D, Schlüter A, Pühler A, Bartosik D. 2015. Genome-guided insight into the methylotrophy of Paracoccus aminophilus JCM 7686. Front Microbiol 6:852. doi: 10.3389/fmicb.2015.00852 [DOI] [PMC free article] [PubMed] [Google Scholar]
14. Harms N, Reijnders WN, Anazawa H, van der Palen CJ, van Spanning RJ, Oltmann LF, Stouthamer AH. 1993. Identification of a two-component regulatory system controlling methanol dehydrogenase synthesis in Paracoccus denitrificans. Mol Microbiol 8:457–470. doi: 10.1111/j.1365-2958.1993.tb01590.x [DOI] [PubMed] [Google Scholar]
15. Harms N, Reijnders WN, Koning S, van Spanning RJ. 2001. Two-component system that regulates methanol and formaldehyde oxidation in Paracoccus denitrificans. J Bacteriol 183:664–670. doi: 10.1128/JB.183.2.664-670.2001 [DOI] [PMC free article] [PubMed] [Google Scholar]
16. Hallgren J, Tsirigos KD, Pedersen MD, Almagro Armenteros JJ, Marcatili P, Nielsen H, Krogh A, Winther O. 2022. DeepTMHMM predicts alpha and beta transmembrane proteins using deep neural networks. Bioinformatics. doi: 10.1101/2022.04.08.487609 [DOI]
17. Ras J, Van Ophem PW, Reijnders WN, Van Spanning RJ, Duine JA, Stouthamer AH, Harms N. 1995. Isolation, sequencing, and mutagenesis of the gene encoding NAD- and glutathione-dependent formaldehyde dehydrogenase (GD-FALDH) from Paracoccus denitrificans, in which GD-FALDH is essential for methylotrophic growth. J Bacteriol 177:247–251. doi: 10.1128/jb.177.1.247-251.1995 [DOI] [PMC free article] [PubMed] [Google Scholar]
18. de Vries GE, Harms N, Maurer K, Papendrecht A, Stouthamer AH. 1988. Physiological regulation of Paracoccus denitrificans methanol dehydrogenase synthesis and activity. J Bacteriol 170:3731–3737. doi: 10.1128/jb.170.8.3731-3737.1988 [DOI] [PMC free article] [PubMed] [Google Scholar]
19. Harms N, van Spanning RJ. 1991. C1 metabolism in Paracoccus denitrificans: genetics of Paracoccus denitrificans. J Bioenerg Biomembr 23:187–210. doi: 10.1007/BF00762217 [DOI] [PubMed] [Google Scholar]
20. Wargo MJ. 2013. Homeostasis and catabolism of choline and glycine betaine: lessons from Pseudomonas aeruginosa. Appl Environ Microbiol 79:2112–2120. doi: 10.1128/AEM.03565-12 [DOI] [PMC free article] [PubMed] [Google Scholar]
21. Wargo MJ, Szwergold BS, Hogan DA. 2008. Identification of two gene clusters and a transcriptional regulator required for Pseudomonas aeruginosa glycine betaine catabolism. J Bacteriol 190:2690–2699. doi: 10.1128/JB.01393-07 [DOI] [PMC free article] [PubMed] [Google Scholar]
22. Shao Y-H, Guo L-Z, Zhang Y-Q, Yu H, Zhao B-S, Pang H-Q, Lu W-D. 2018. Glycine betaine monooxygenase, an unusual Rieske-type oxygenase system, catalyzes the oxidative N-demethylation of glycine betaine in Chromohalobacter salexigens DSM 3043. Appl Environ Microbiol 84:e00377-18. doi: 10.1128/AEM.00377-18 [DOI] [PMC free article] [PubMed] [Google Scholar]
23. Barra L, Fontenelle C, Ermel G, Trautwetter A, Walker GC, Blanco C. 2006. Interrelations between glycine betaine catabolism and methionine biosynthesis in Sinorhizobium meliloti strain 102F34. J Bacteriol 188:7195–7204. doi: 10.1128/JB.00208-06 [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Lidbury I, Kimberley G, Scanlan DJ, Murrell JC, Chen Y. 2015. Comparative genomics and mutagenesis analyses of choline metabolism in the marine Roseobacter clade. Environ Microbiol 17:5048–5062. doi: 10.1111/1462-2920.12943 [DOI] [PMC free article] [PubMed] [Google Scholar]
25. Leys D, Basran J, Scrutton NS. 2003. Channelling and formation of ‘active’ formaldehyde in dimethylglycine oxidase. EMBO J 22:4038–4048. doi: 10.1093/emboj/cdg395 [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Wagner MA, Schuman Jorns M. 1997. Folate utilization by monomeric versus heterotetrameric sarcosine oxidases. Arch Biochem Biophys 342:176–181. doi: 10.1006/abbi.1997.0106 [DOI] [PubMed] [Google Scholar]
27. Kanehisa M, Sato Y, Kawashima M, Furumichi M, Tanabe M. 2016. KEGG as a reference resource for gene and protein annotation. Nucleic Acids Res 44:D457–D462. doi: 10.1093/nar/gkv1070 [DOI] [PMC free article] [PubMed] [Google Scholar]
28. Miller JH. 1992. A short course in bacterial genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. [Google Scholar]
29. Li S, Yu X, Beattie GA. 2013. Glycine betaine catabolism contributes to Pseudomonas syringae tolerance to hyperosmotic stress by relieving betaine-mediated suppression of compatible solute synthesis. J Bacteriol 195:2415–2423. doi: 10.1128/JB.00094-13 [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Rkenes TP, Lamark T, Strøm AR. 1996. DNA-binding properties of the BetI repressor protein of Escherichia coli: the inducer choline stimulates BetI-DNA complex formation. J Bacteriol 178:1663–1670. doi: 10.1128/jb.178.6.1663-1670.1996 [DOI] [PMC free article] [PubMed] [Google Scholar]
31. de Vries GE, Harms N, Hoogendijk J, Stouthamer AH. 1989. Isolation and characterization of Paracoccus denitrificans mutants with increased conjugation frequencies and pleiotropic loss of a (nGATCn) DNA-modifying property. Arch. Microbiol 152:52–57. doi: 10.1007/BF00447011 [DOI] [Google Scholar]
32. Simon R, Priefer U, Pühler A. 1983. A broad host range mobilisation system for in vivo genetic engineering: transposon mutagenesis in Gram negative bacteria. Nat Biotechnol 1:784–791. doi: 10.1038/nbt1183-784 [DOI] [Google Scholar]
33. Shearer N, Hinsley AP, Van Spanning RJ, Spiro S. 1999. Anaerobic growth of Paracoccus denitrificans requires cobalamin: characterization of cobK and cobJ genes. J Bacteriol 181:6907–6913. doi: 10.1128/JB.181.22.6907-6913.1999 [DOI] [PMC free article] [PubMed] [Google Scholar]
34. Hahnke SM, Moosmann P, Erb TJ, Strous M. 2014. An improved medium for the anaerobic growth of Paracoccus denitrificans Pd1222. Front Microbiol 5:18. doi: 10.3389/fmicb.2014.00018 [DOI] [PMC free article] [PubMed] [Google Scholar]
35. Sullivan MJ, Gates AJ, Appia-Ayme C, Rowley G, Richardson DJ. 2013. Copper control of bacterial nitrous oxide emission and its impact on vitamin B12-dependent metabolism. Proc Natl Acad Sci U S A 110:19926–19931. doi: 10.1073/pnas.1314529110 [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Ind AC, Porter SL, Brown MT, Byles ED, de Beyer JA, Godfrey SA, Armitage JP. 2009. Inducible-expression plasmid for Rhodobacter sphaeroides and Paracoccus denitrificans. Appl Environ Microbiol 75:6613–6615. doi: 10.1128/AEM.01587-09 [DOI] [PMC free article] [PubMed] [Google Scholar]
37. Spaink HP, Okker RJH, Wijffelman CA, Pees E, Lugtenberg BJJ. 1987. Promoters in the nodulation region of the Rhizobium leguminosarum Sym plasmid pRJ1JI. Plant Mol Biol 9:27–39. doi: 10.1007/BF00017984 [DOI] [PubMed] [Google Scholar]
38. Kleeberg U, Klinger W. 1982. Sensitive formaldehyde determination with NASH's reagent and a ‘tryptophan reaction’. J Pharmacol Methods 8:19–31. doi: 10.1016/0160-5402(82)90004-3 [DOI] [PubMed] [Google Scholar]
39. Ogushi S, Ando M, Tsuru D. 1984. Substrate specificity of formaldehyde dehydrogenase from Pseudomonas putida. Agric Biol Chem 48:597–601. doi: 10.1271/bbb1961.48.597 [DOI] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental material. jb.00081-24-s0001.pdf.

Table S1 and Figures S1 to S8.

jb.00081-24-s0001.pdf^{(1.1MB, pdf)}

DOI: 10.1128/jb.00081-24.SuF1

[B1] 1. Du H, Jiao N, Hu Y, Zeng Y. 2006. Diversity and distribution of pigmented heterotrophic bacteria in marine environments. FEMS Microbiol Ecol 57:92–105. doi: 10.1111/j.1574-6941.2006.00090.x [DOI] [PubMed] [Google Scholar]

[B2] 2. Russell JA, León-Zayas R, Wrighton K, Biddle JF. 2016. Deep subsurface life from North Pond: enrichment, isolation, characterization and genomes of heterotrophic bacteria. Front Microbiol 7:678. doi: 10.3389/fmicb.2016.00678 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B3] 3. Wang Z, Zhang X-X, Lu X, Liu B, Li Y, Long C, Li A. 2014. Abundance and diversity of bacterial nitrifiers and denitrifiers and their functional genes in tannery wastewater treatment plants revealed by high-throughput sequencing. PLoS One 9:e113603. doi: 10.1371/journal.pone.0113603 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B4] 4. Dias ACF, Dini-Andreote F, Hannula SE, Andreote FD, Pereira E Silva M de C, Salles JF, de Boer W, van Veen J, van Elsas JD. 2013. Different selective effects on rhizosphere bacteria exerted by genetically modified versus conventional potato lines. PLoS One 8:e67948. doi: 10.1371/journal.pone.0067948 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B5] 5. Castelino M, Eyre S, Moat J, Fox G, Martin P, Ho P, Upton M, Barton A. 2017. Optimisation of methods for bacterial skin microbiome investigation: primer selection and comparison of the 454 versus MiSeq platform. BMC Microbiol 17:23. doi: 10.1186/s12866-017-0927-4 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B6] 6. Baker SC, Ferguson SJ, Ludwig B, Page MD, Richter OM, van Spanning RJ. 1998. Molecular genetics of the genus Paracoccus: metabolically versatile bacteria with bioenergetic flexibility. Microbiol Mol Biol Rev 62:1046–1078. doi: 10.1128/MMBR.62.4.1046-1078.1998 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B7] 7. Van Spanning RJM, de Vries S, Harms N. 2000. Coping with formaldehyde during C1 metabolism of Paracoccus denitrificans. J Mol Catal B Enzym 8:37–50. doi: 10.1016/S1381-1177(99)00065-X [DOI] [Google Scholar]

[B8] 8. Köstler M, Kleiner D. 1989. Assimilation of methylamine by Paracoccus denitrificans involves formaldehyde transport by a specific carrier. FEMS Microbiol Lett 53:1–4. doi: 10.1016/0378-1097(89)90356-x [DOI] [PubMed] [Google Scholar]

[B9] 9. Ophem PW, Duine JA. 1994. NAD- and co-substrate (GSH or factor)-dependent formaldehyde dehydrogenases from methylotrophic microorganisms act as a class III alcohol dehydrogenase. FEMS Microbiol Lett 116:87–93. doi: 10.1111/j.1574-6968.1994.tb06680.x [DOI] [Google Scholar]

[B10] 10. Kumar R, Zhao S, Vetting MW, Wood BM, Sakai A, Cho K, Solbiati J, Almo SC, Sweedler JV, Jacobson MP, Gerlt JA, Cronan JE. 2014. Prediction and biochemical demonstration of a catabolic pathway for the osmoprotectant proline betaine. mBio 5:e00933-13. doi: 10.1128/mBio.00933-13 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B11] 11. Bordel S, Martín-González D, Börner T, Muñoz R, Santos-Beneit F. 2024. Genome-scale metabolic model of the versatile bacterium Paracoccus denitrificans Pd1222. mSystems 9:e0107723. doi: 10.1128/msystems.01077-23 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B12] 12. Cox RB, Quayle JR. 1975. The autotrophic growth of Micrococcus denitrificans on methanol. Biochem J 150:569–571. doi: 10.1042/bj1500569 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B13] 13. Dziewit L, Czarnecki J, Prochwicz E, Wibberg D, Schlüter A, Pühler A, Bartosik D. 2015. Genome-guided insight into the methylotrophy of Paracoccus aminophilus JCM 7686. Front Microbiol 6:852. doi: 10.3389/fmicb.2015.00852 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B14] 14. Harms N, Reijnders WN, Anazawa H, van der Palen CJ, van Spanning RJ, Oltmann LF, Stouthamer AH. 1993. Identification of a two-component regulatory system controlling methanol dehydrogenase synthesis in Paracoccus denitrificans. Mol Microbiol 8:457–470. doi: 10.1111/j.1365-2958.1993.tb01590.x [DOI] [PubMed] [Google Scholar]

[B15] 15. Harms N, Reijnders WN, Koning S, van Spanning RJ. 2001. Two-component system that regulates methanol and formaldehyde oxidation in Paracoccus denitrificans. J Bacteriol 183:664–670. doi: 10.1128/JB.183.2.664-670.2001 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B16] 16. Hallgren J, Tsirigos KD, Pedersen MD, Almagro Armenteros JJ, Marcatili P, Nielsen H, Krogh A, Winther O. 2022. DeepTMHMM predicts alpha and beta transmembrane proteins using deep neural networks. Bioinformatics. doi: 10.1101/2022.04.08.487609 [DOI]

[B17] 17. Ras J, Van Ophem PW, Reijnders WN, Van Spanning RJ, Duine JA, Stouthamer AH, Harms N. 1995. Isolation, sequencing, and mutagenesis of the gene encoding NAD- and glutathione-dependent formaldehyde dehydrogenase (GD-FALDH) from Paracoccus denitrificans, in which GD-FALDH is essential for methylotrophic growth. J Bacteriol 177:247–251. doi: 10.1128/jb.177.1.247-251.1995 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B18] 18. de Vries GE, Harms N, Maurer K, Papendrecht A, Stouthamer AH. 1988. Physiological regulation of Paracoccus denitrificans methanol dehydrogenase synthesis and activity. J Bacteriol 170:3731–3737. doi: 10.1128/jb.170.8.3731-3737.1988 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B19] 19. Harms N, van Spanning RJ. 1991. C1 metabolism in Paracoccus denitrificans: genetics of Paracoccus denitrificans. J Bioenerg Biomembr 23:187–210. doi: 10.1007/BF00762217 [DOI] [PubMed] [Google Scholar]

[B20] 20. Wargo MJ. 2013. Homeostasis and catabolism of choline and glycine betaine: lessons from Pseudomonas aeruginosa. Appl Environ Microbiol 79:2112–2120. doi: 10.1128/AEM.03565-12 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21. Wargo MJ, Szwergold BS, Hogan DA. 2008. Identification of two gene clusters and a transcriptional regulator required for Pseudomonas aeruginosa glycine betaine catabolism. J Bacteriol 190:2690–2699. doi: 10.1128/JB.01393-07 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B22] 22. Shao Y-H, Guo L-Z, Zhang Y-Q, Yu H, Zhao B-S, Pang H-Q, Lu W-D. 2018. Glycine betaine monooxygenase, an unusual Rieske-type oxygenase system, catalyzes the oxidative N-demethylation of glycine betaine in Chromohalobacter salexigens DSM 3043. Appl Environ Microbiol 84:e00377-18. doi: 10.1128/AEM.00377-18 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B23] 23. Barra L, Fontenelle C, Ermel G, Trautwetter A, Walker GC, Blanco C. 2006. Interrelations between glycine betaine catabolism and methionine biosynthesis in Sinorhizobium meliloti strain 102F34. J Bacteriol 188:7195–7204. doi: 10.1128/JB.00208-06 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B24] 24. Lidbury I, Kimberley G, Scanlan DJ, Murrell JC, Chen Y. 2015. Comparative genomics and mutagenesis analyses of choline metabolism in the marine Roseobacter clade. Environ Microbiol 17:5048–5062. doi: 10.1111/1462-2920.12943 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B25] 25. Leys D, Basran J, Scrutton NS. 2003. Channelling and formation of ‘active’ formaldehyde in dimethylglycine oxidase. EMBO J 22:4038–4048. doi: 10.1093/emboj/cdg395 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B26] 26. Wagner MA, Schuman Jorns M. 1997. Folate utilization by monomeric versus heterotetrameric sarcosine oxidases. Arch Biochem Biophys 342:176–181. doi: 10.1006/abbi.1997.0106 [DOI] [PubMed] [Google Scholar]

[B27] 27. Kanehisa M, Sato Y, Kawashima M, Furumichi M, Tanabe M. 2016. KEGG as a reference resource for gene and protein annotation. Nucleic Acids Res 44:D457–D462. doi: 10.1093/nar/gkv1070 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B28] 28. Miller JH. 1992. A short course in bacterial genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. [Google Scholar]

[B29] 29. Li S, Yu X, Beattie GA. 2013. Glycine betaine catabolism contributes to Pseudomonas syringae tolerance to hyperosmotic stress by relieving betaine-mediated suppression of compatible solute synthesis. J Bacteriol 195:2415–2423. doi: 10.1128/JB.00094-13 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B30] 30. Rkenes TP, Lamark T, Strøm AR. 1996. DNA-binding properties of the BetI repressor protein of Escherichia coli: the inducer choline stimulates BetI-DNA complex formation. J Bacteriol 178:1663–1670. doi: 10.1128/jb.178.6.1663-1670.1996 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B31] 31. de Vries GE, Harms N, Hoogendijk J, Stouthamer AH. 1989. Isolation and characterization of Paracoccus denitrificans mutants with increased conjugation frequencies and pleiotropic loss of a (nGATCn) DNA-modifying property. Arch. Microbiol 152:52–57. doi: 10.1007/BF00447011 [DOI] [Google Scholar]

[B32] 32. Simon R, Priefer U, Pühler A. 1983. A broad host range mobilisation system for in vivo genetic engineering: transposon mutagenesis in Gram negative bacteria. Nat Biotechnol 1:784–791. doi: 10.1038/nbt1183-784 [DOI] [Google Scholar]

[B33] 33. Shearer N, Hinsley AP, Van Spanning RJ, Spiro S. 1999. Anaerobic growth of Paracoccus denitrificans requires cobalamin: characterization of cobK and cobJ genes. J Bacteriol 181:6907–6913. doi: 10.1128/JB.181.22.6907-6913.1999 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B34] 34. Hahnke SM, Moosmann P, Erb TJ, Strous M. 2014. An improved medium for the anaerobic growth of Paracoccus denitrificans Pd1222. Front Microbiol 5:18. doi: 10.3389/fmicb.2014.00018 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B35] 35. Sullivan MJ, Gates AJ, Appia-Ayme C, Rowley G, Richardson DJ. 2013. Copper control of bacterial nitrous oxide emission and its impact on vitamin B12-dependent metabolism. Proc Natl Acad Sci U S A 110:19926–19931. doi: 10.1073/pnas.1314529110 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B36] 36. Ind AC, Porter SL, Brown MT, Byles ED, de Beyer JA, Godfrey SA, Armitage JP. 2009. Inducible-expression plasmid for Rhodobacter sphaeroides and Paracoccus denitrificans. Appl Environ Microbiol 75:6613–6615. doi: 10.1128/AEM.01587-09 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B37] 37. Spaink HP, Okker RJH, Wijffelman CA, Pees E, Lugtenberg BJJ. 1987. Promoters in the nodulation region of the Rhizobium leguminosarum Sym plasmid pRJ1JI. Plant Mol Biol 9:27–39. doi: 10.1007/BF00017984 [DOI] [PubMed] [Google Scholar]

[B38] 38. Kleeberg U, Klinger W. 1982. Sensitive formaldehyde determination with NASH's reagent and a ‘tryptophan reaction’. J Pharmacol Methods 8:19–31. doi: 10.1016/0160-5402(82)90004-3 [DOI] [PubMed] [Google Scholar]

[B39] 39. Ogushi S, Ando M, Tsuru D. 1984. Substrate specificity of formaldehyde dehydrogenase from Pseudomonas putida. Agric Biol Chem 48:597–601. doi: 10.1271/bbb1961.48.597 [DOI] [Google Scholar]

PERMALINK

Choline degradation in Paracoccus denitrificans: identification of sources of formaldehyde

Trusha Parekh

Marcus Tsai

Stephen Spiro

Roles