Metabolic role of a genetically conserved aldehyde dehydrogenase in bacterial assimilation of various primary amines

This preprint has been published as a journal article in Archives of Biochemistry and Biophysics.

DOI: https://doi.org/10.1016/j.abb.2025.110387

This preprint version differs from the final, peer-reviewed publication, which reflects revisions made in response to reviewer feedback. For the final version, please refer to the published article. This manuscript version is made available under the CC-BY-NC-ND license.

Tadashi Nakai^a,b,*, Naoya Miyagi^a, Kota Hisamura^b, Shoya Matsuba^a,b, Kanji Nishimoto^b, Emi Nakai^b, Katsuyuki Tanizawa^c, Toshihide Okajima^c

^a Graduate School of Science and Technology, Hiroshima Institute of Technology, Hiroshima, Hiroshima 731-5193, Japan

^b Faculty of Life Sciences, Hiroshima Institute of Technology, Hiroshima, Hiroshima 731-5193, Japan

^c Institute of Scientific and Industrial Research (SANKEN), Osaka University, Ibaraki, Osaka 567-0047, Japan

*Corresponding author. E-mail address: t.nakai.wj@cc.it-hiroshima.ac.jp (T. Nakai).

Highlights:

• An aldehyde dehydrogenase (qhpH) is conserved near the qhp operon
• Expression of qhpH is induced by primary amines, not by their aldehyde substrates
• Growth of P. denitrificans is unaffected by disruption of qhpH
• Co-expression of qhpH in R. sphaeroides improves growth on n-butylamine
• QhpH oxidizes aldehydes produced by an amine dehydrogenase from the qhp operon

ABSTRACT

Primary amines such as n-butylamine and 2-phenylethylamine serve as good nitrogen, carbon, and energy sources for bacterial growth. In many Gram-negative bacterial species, these amines are first oxidized by a periplasmic enzyme, quinohemoprotein amine dehydrogenase (QHNDH), encoded in an operon termed ‘qhp’, consisting of eight genes (qhpABCDEFGR). A gene predicted to encode an aldehyde dehydrogenase is also highly conserved in the vicinity of the qhp operon. In this study, we found that a 5’-upstream region of the aldehyde dehydrogenase gene in Paracoccus denitrificans has a high promoter activity that responds to n-butylamine supplementation in the culture medium, indicating co-regulation with the qhp genes by the transcriptional regulator QhpR. Hence, we designate this gene as the ninth member of the qhp operon, qhpH. Disruption of qhpH in P. denitrificans neither affected bacterial growth on primary amines, nor impaired QHNDH activity, suggesting the presence of another constitutive aldehyde dehydrogenase(s) compensating for the defect of qhpH. Nevertheless, heterologous expression of qhpH along with the eight qhp genes in an amine non-assimilating bacterium, Rhodobacter sphaeroides, significantly enhanced the growth on n-butylamine, as compared to the slow growth without qhpH. The recombinant QhpH purified from Escherichia coli cells showed high aldehyde dehydrogenase activities toward various aldehydes. These findings demonstrate that the qhpH gene encodes an aldehyde dehydrogenase with broad substrate specificity and is evolutionarily conserved with the qhp operon to play a role in an efficient metabolism of various primary amines in Gram-negative bacteria.

Keywords:

aldehyde dehydrogenase

amine assimilation

quinohemoprotein amine dehydrogenase

Abbreviations: 

QHNDH, quinohemoprotein amine dehydrogenase; n-BA, n-butylamine; LB, Luria broth; ONPG, o-nitrophenyl β-D-galactopyranoside.

Introduction

Bacterial degradation of primary amines initiates with the oxidative deamination of the amino groups by amine dehydrogenases or amine oxidases, producing a 2e⁻-reducing equivalent, ammonia, and the corresponding aldehydes [1]. The aldehyde products are further oxidized by aldehyde dehydrogenases to carboxylic acids, which are then modified with coenzyme A (CoA) [1]. The acyl-CoA compounds produced are subsequently assimilated through the ordinary β-oxidation pathway. One of the amine-oxidizing enzymes, quinohemoprotein amine dehydrogenase (QHNDH), is widely distributed in Gram-negative bacteria and catalyzes the oxidative deamination of various primary amines that serve as energy, carbon, and nitrogen sources [2–4]. QHNDH contains a post-translationally derived quinone cofactor, cysteine tryptophylquinone, within its smallest γ-subunit of the αβγ heterotrimeric protein [5,6].

The structural genes encoding QHNDH constitute an operon termed qhp, which, together with several nearby genes, forms a gene cluster, qhpABCDEFGR [7]. The qhp operon is found in over 1300 bacterial species, mostly Gram-negative bacteria [7]. The qhpA, qhpB, and qhpC genes encode the α-, β-, and γ-subunits of QHNDH, respectively. The qhpD gene encodes an unusual radical S-adenosylmethionine enzyme that catalyzes the sequential formation of three Cys-to-Asp/Glu intra-peptidyl thioether bonds within QhpC [8,9]. The qhpE gene encodes a subtilisin-like serine protease that cleaves the N-terminal 28-residue leader peptide from the crosslinked QhpC [10] before its periplasmic translocation through an efflux ABC transporter encoded by the qhpF gene [7]. The qhpG gene encodes an atypical single-component flavin-dependent monooxygenase that catalyzes the dihydroxylation of an unmodified Trp residue in the protein substrate [11]. The qhpR gene encodes an amine-inducible AraC-like transcriptional regulator [7]. In Paracoccus denitrificans Pd1222, a model soil microorganism belonging to α-proteobacteria, the qhp genes are arranged in an order of GADCBEFR (Fig. 1A) on chromosome 1, and the qhpADCBEF operon and the qhpG gene are transcribed individually under the common control of the qhp promoter [7].

As described above, the biogenesis of QHNDH involves an intricate mechanism, and to date, the eight genes of qhpABCDEFGR are known to be necessary and sufficient for the biogenesis of QHNDH [7,12]. In addition to these eight genes, homologs of an open reading frame Pden_1710, coding for a protein of 467 amino acid residues containing NAD(P)⁺-binding sites (Fig. 1B), are highly conserved in other bacterial species near the qhp genes [7]. Although Pden_1710 has been annotated as a betaine aldehyde dehydrogenase gene (GenBank accession no. ABL69807) in the whole genome analysis of P. denitrificans Pd1222, probably based on sequence similarity, the gene is highly conserved among bacteria possessing the qhp genes and is thus presumed to encode an aldehyde dehydrogenase with substrate specificity suitable for the products of QHNDH, both aliphatic and aromatic aldehydes. However, the metabolic role of Pden_1710 remains to be studied.

In this study, we first analyzed the promoter activity of Pden_1710 and found that its expression is induced by n-butylamine (n-BA) at the same level as the qhpADCBEF genes. Thus, we assigned Pden_1710 to qhpH. Next, through gene disruption, we examined the essentiality of the qhpH gene, particularly in relation to amine assimilation. Furthermore, we heterologously co-expressed qhpH and other qhp genes in an amine non-assimilating bacterium, Rhodobacter sphaeroides, and observed enhanced growth on n-BA, indicating the importance of qhpH in efficient amine utilization. Finally, we overexpressed the qhpH gene in Escherichia coli cells, purified the recombinant QhpH, and studied its substrate specificity to infer its metabolic role.

Fig. 1. Structure of the qhp gene cluster.

(A) The qhp gene cluster on chromosome 1 of P. denitrificans Pd1222 is shown schematically. The gene names initially annotated by the genome project are labeled for each gene. Inverted repeat sequences (I1R/I1F and I3R/I3F) and single-repeat sequence (I2F) are presumed to be the QhpR-binding regions [7]. (B) Amino acid sequence of a putative aldehyde dehydrogenase encoded by Pden_1710. Residues forming the NAD(P)⁺-binding site (highlighted in yellow) and the catalytic residues (marked in red) are shown based on the sequence annotation from GenBank: ABL69807.1. The numbers on the left indicate the residue numbers of the first amino acid in each line. (C) Nucleotide sequences of the intergenic region between Pden_1709 (qhpR) and Pden_1710 (qhpH). The sequences of the I3R/I3F and I2F regions [7] are indicated below the nucleotide sequence (green) with dotted arrows. The predicted −35 and −10 promoter regions of Pden_1710 (qhpH) [7] are highlighted in gray.

Materials and Methods

2.1. Materials, bacterial strains, and culture conditions

E. coli strains DH5α, S17-1, and C41(DE3) were used for plasmid preparation, diparental mating, and protein expression, respectively. E. coli cells were grown aerobically at 37 °C in Luria broth (LB) medium [1% (w/v) polypeptone, 0.5% (w/v) yeast extract, and 0.5% (w/v) NaCl]. P. denitrificans strain Pd1222 and E. coli strain S17-1 were kindly provided by R. J. van Spanning (Vrije Universiteit, The Netherlands). R. sphaeroides NBRC12203 was obtained from the NITE Biological Resource Center (Chiba, Japan). P. denitrificans and R. sphaeroides cells were grown aerobically at 30 °C in LB medium or the minimal mineral medium prepared as described previously [12]. The carbon sources used for growth were 46 mM n-BA, 14 mM n-butyraldehyde, or 20 mM choline chloride. Antibiotics were supplied as needed at the following concentrations: 50 μg mL⁻¹ ampicillin, 50 μg mL⁻¹ kanamycin, 20 μg mL⁻¹ rifampicin, 10 μg mL⁻¹ (for E. coli) or 1 μg mL⁻¹ (for P. denitrificans and R. sphaeroides) tetracycline, and 50 μg mL⁻¹ streptomycin.

2.2. Promoter assay

The promoter activity of the qhpH gene was measured using a β-galactosidase assay with o-nitrophenyl β-D-galactopyranoside (ONPG) as the substrate. To construct a plasmid carrying the promoter region fused to the lacZ gene, the upstream region of the qhpH gene was amplified from the genome of Pd1222 using colony PCR with primers (P1 and P2) containing NdeI and EcoRI restriction sites (Table S1), respectively, based on the nucleotide sequence of Pd1222 chromosome 1 (GenBank accession no. CP000489). The amplified fragment, designated as P_H350, corresponds to the region from −322 to −4 nucleotides upstream of the qhpH gene. Expression plasmids for P. denitrificans were constructed using the broad host range vector pRK415-1 [13], primarily following standard molecular genetic methods. The PCR product was digested with NdeI and EcoRI and inserted into pRK-P_A800-lacZ [7] (replacing P_A800), in which P_A800 includes the promoter of the qhp operon, to yield pRK-P_H350-lacZ. The nucleotide sequence of the inserted promoter region is shown in Fig. 1C. The constructed plasmid was introduced into the wild-type Pd1222 by diparental mating using E. coli S17-1 as donor cells. With the reporter strain, promoter activities were determined by ONPG assays and expressed as Miller units [14], calculated using the formula 1000 × A₄₂₀ / (t × V × OD₆₀₀), where A₄₂₀ represents the absorbance at 420 nm of o-nitrophenol generated from ONPG, t is the reaction time in minutes, V is the reaction volume in milliliters, and OD₆₀₀ is the cell density measured at 600 nm.

2.3. Gene disruption

A plasmid for disruption of the qhpH gene was constructed using the suicide vector pGRPd1 [15] according to the flow chart shown in Fig. 2A. (i) The qhpR-qhpH genes on the genome of P. denitrificans Pd1222 were amplified by PCR using primers P3 and P4 (Table S1), with the genomic DNA as the template. The BamHI/XhoI fragment excised from the PCR product was inserted into pBluescript II SK (+) (Agilent), yielding pBSIISK-qhpR-qhpH. (ii) The kanamycin resistance gene (Km^R) was amplified by PCR using primers P5 and P6 with pUC4K as the template. The HincII fragment excised from the PCR product was inserted into the NcoI and HindIII sites (after blunt-ending) of the resulting plasmid, yielding pBSIISK-qhpR-qhpH::Km^R. (iii) This plasmid was then digested with KpnI and BamHI and inserted into the MCS of pGRPd1-MCS [7], yielding the final plasmid for gene disruption, pGRPd1-qhpH::Km^R.

The qhpH gene of P. denitrificans was disrupted by homologous recombination, essentially as described previously [7,8]. Briefly, donor E. coli S17-1 cells carrying pGRPd1-qhpH::Km^R were conjugated with recipient Pd1222 cells [7,8], and qhpH-disrupted mutants of Pd1222 were selected from Km^R and streptomycin-sensitive colonies. Gene disruption was confirmed by colony PCR using primers P7–P10, which were designed to amplify a DNA fragment covering qhpH interrupted by the Km^R gene (Table S1). Finally, a mutant of Pd1222 carrying the qhpH::Km^R gene in the genome, designated as ΔqhpH, was obtained and used without removing the Km^R gene.

Fig. 2. Flow chart of plasmid construction procedures for gene disruption (A) and heterologous expression (B).

The Roman numerals in parentheses correspond to the procedural steps described in the text.

2.4. Heterologous expression of qhp genes in R. sphaeroides

To co-express qhpH with qhpABCDEFGR, the plasmid pRK-qhpGADCBEFRH was constructed according to the following procedure (Fig. 2B). (i) pRK-P_A800-qhpG [7] was digested with BamHI, blunt-ended using Klenow fragment, and self-ligated to generate pRK-P_A800-qhpG-ΔBamHI. (ii) The Bsu36I/SphI fragment excised from pRK-P_A800-qhpG-ΔBamHI was inserted into pRK-qhpGADCBEFR constructed previously [12], yielding pRK-ΔBamHI-qhpGADCBEFR. (iii) The BamHI/EcoRI fragment excised from pRK-ΔBamHI-qhpGADCBEFR was inserted into pET-15b (Merck Millipore), producing pET-qhpR-qhpH(part). (iv) The latter part of the qhpH gene was amplified by PCR using primers P11 and P12, which contained HindIII and EcoRI sites (Table S1), respectively, with Pd1222 genomic DNA as the template. The HindIII/EcoRI fragment excised from the PCR product was inserted into pET-qhpR-qhpH(part), yielding pET-qhpR-qhpH. (v) The BamHI/EcoRI fragment excised from the resulting plasmid was inserted into pRK-ΔBamHI-qhpGADCBEFR, yielding pRK-qhpGADCBEFRH. The expression plasmid (pRK-qhpGADCBEFRH or pRK-qhpGADCBEFR) was introduced into R. sphaeroides by diparental mating using E. coli S17-1 as donor cells. R. sphaeroides cells harboring the expression plasmid were selected on Sistrom’s minimal medium plates (1.5% agar) [16] containing 1 μg mL⁻¹ tetracycline. The cells were grown aerobically at 30 °C in the minimal mineral medium [12] supplemented with 46 mM n-BA.

2.5. Expression, purification, and assay of recombinant QhpH

To construct an expression plasmid for QhpH, the coding region of qhpH was amplified from Pd1222 genomic DNA using primers P4 and P13, which contain NdeI and BamHI sites, respectively (Table S1). The PCR product was digested and inserted into pET-His₆-TEV-QhpG [11] (replacing QhpG) to yield pET-His₆-TEV-QhpH.

E. coli C41(DE3) cells transformed with pET-His₆-TEV-QhpH were grown at 37 °C with reciprocal shaking at 180 rpm in a modified version of Studier autoinduction medium [17] (2% polypeptone, 0.5% yeast extract, 0.5% NaCl, 0.6% Na₂HPO₄, 0.3% KH₂PO₄, 0.5% glycerol, 0.05% glucose, 0.2% lactose) with ampicillin. When the cell density reached an approximate optical density of 0.1, the temperature was lowered to 25 °C and the cells were further grown for 16 h. The cells were harvested by centrifugation at 6,000g for 10 min and stored at −20 °C until use. Frozen cells (3 g of wet weight from 100 mL culture) were thawed, suspended in buffer A [25 mM Tris-HCl buffer (pH 8) containing 500 mM NaCl and 10% (w/v) glycerol], and then disrupted by sonication. The cell lysate was centrifuged at 20,000g for 60 min at 4 °C. The resulting supernatant was applied to a Ni-affinity column packed with cOmplete His-Tag Purification Resin (Roche), which had been equilibrated with buffer A. After washing the column with 25 mL of buffer A, the bound protein was eluted in a stepwise manner with buffer A containing 25, 50, 75, 100, 125, 150, 175, 200, 225, and 250 mM imidazole. The fractions containing QhpH were pooled. To remove the N-terminal His₆-tag, the protein was digested with His-tagged TEV protease for 3 h at 30 °C. After digestion, imidazole was removed using a PD-10 column (Cytiva). The protease and released His₆-tag were removed by a second Ni-affinity chromatography step. The tag-less QhpH was collected from the flow-through fraction and used as the purified protein. The purified protein was concentrated and stored at −80 °C. The fractions were analyzed by SDS-PAGE using an Any kD denaturing polyacrylamide gel (Bio-Rad).

The activity of QhpH was measured at 25 °C as described previously with minor modifications [18]. The assay mixture contained 50 mM Tris-HCl (pH 7.4), known concentrations of aldehyde substrate, and 1 mM NAD⁺ in a final volume of 1 mL. Reactions were initiated by the addition of substrate. The progress of the reaction was monitored by measuring the increase in absorbance at 340 nm, which occurs upon NADH formation.

2.6. Other methods

The preparation of periplasmic and cytoplasmic fractions of P. denitrificans cells, QHNDH activity and protein assays, and SDS-PAGE were performed as described previously [7,8,10].

Results and Discussion

3.1. Promoter activity of the upstream region of Pden_1710

We previously reported that the upstream region of Pden_1710 (P_H350, Fig. 1C) contains a sequence similar to that of the qhpADCBEF operon promoter [7]. Since expression of the qhp operon is induced by n-BA, we predicted that if Pden_1710 is expressed together with QHNDH, the upstream region would exhibit similar promoter activity in the presence of n-BA. To examine this possibility, we constructed the plasmid pRK-P_H350-lacZ to measure the promoter activity of the upstream region of Pden_1710. We transformed the wild-type P. denitrificans Pd1222 strain with this plasmid and measured promoter activity under different culture conditions in the minimal medium supplemented with either n-BA, n-butyraldehyde, or choline as the sole carbon source. Promoter assays as an indicator of LacZ activity in the reporter strain revealed that the Pden_1710 promoter was significantly activated by n-BA but not by n- butyraldehyde (Table 1). The promoter activity in the presence of n-BA (4.3 × 10⁴ Miller units) was nearly 2000-fold higher than that of n-butyraldehyde (24 Miller units), which was comparable to that of the non-inducible control carbon source, choline. This specificity suggests that the Pden_1710 promoter responds specifically to primary amines rather than to their direct aldehyde products, highlighting its role in regulating the bacterial response to amine metabolism. Furthermore, LacZ activity driven by the Pden_1710 promoter in the presence of n-BA was comparable to that of the qhpADCBEF operon promoter in the P_A800 region (2.9 × 10⁴ Miller units), indicating that the Pden_1710 promoter is as strong as the qhp operon promoter (Table 1). Moreover, the promoter activities for qhpADCBEF and Pden_1710 were also at a higher level than that for qhpG (1.0 × 10⁴ Miller units) [7]. These findings indicate that upon induction with n-BA, the transcription of the qhpADCBEF operon and qhpG/Pden_1710 genes is simultaneously activated by the same transcriptional regulator, QhpR (Fig. 1A). Therefore, the Pden_1710 gene can be regarded as a member of the qhp operon, hence assigned to qhpH following qhpG. The coordinated regulation implies that QhpR plays a central role in their expression in response to n-BA. We conclude that the regulatory arrangement ensures a concerted response to the presence of primary amines in the culture medium, optimizing the bacterial metabolism of these compounds.

Table. 1 Promoter activities.

Inducera	PA800	PH350
n-butylamine	29,400 ± 6,100	43,500 ± 5,500
n-butyraldehyde	5 ± 2	24 ± 4
choline	9 ± 1	13 ± 2

^a Cells of P. denitrificans Pd1222 transformed with pRK-P_A800-lacZ or pRK-P_H350-lacZ were cultured at 30 °C for 48 h in the minimal mineral medium supplemented with 46 mM n-BA, 14 mM n-butyraldehyde, or 20 mM choline. Cells were collected by brief centrifugation, permeabilized by chloroform/SDS treatment. LacZ activity in the permeabilized cells was measured using ONPG as the substrate [14]. P_A800 and P_H350 represent the qhpA and qhpH promoters, containing 800 bp and 350 bp of their respective upstream regions. Values are presented as the mean ± standard deviation from two independent experiments.

3.2. Effect of qhpH disruption on bacterial growth and QHNDH activity

When the wild-type Pd1222 strain was cultured in the minimal mineral medium containing n-BA as the sole carbon and energy source (n-BA minimal medium), it grew efficiently by inducibly producing QHNDH. Conversely, when the Pd1222 strain lost the QHNDH activity by the disruption of a related gene, the strain could not grow with n-BA as the carbon source [7,8,10]. Therefore, the involvement of the disrupted gene in n-BA metabolism can be assessed by examining whether the resulting disrupted strain can grow with n-BA as the carbon source.

To investigate the role of qhpH in n-BA metabolism, we generated a strain in which the qhpH gene of Pd1222 was disrupted by homologous recombination, designated as ΔqhpH. We cultured the ΔqhpH strain in the n-BA minimal medium and measured its growth, comparing it with the wild-type Pd1222 strain. As shown in Fig. 3A, the Pd1222 and ΔqhpH strains exhibited the same level of growth in the n-BA minimal medium. Subsequently, we measured QHNDH activity in the periplasmic fraction of each strain. The ΔqhpH strain showed a similar level of QHNDH activity, 87 ± 9 % (n = 3) of the wild-type strain. These results indicate that QHNDH activity is not affected by the disruption of qhpH. It is known that the qhpDEFGR genes, in addition to the structural genes qhpABC, are essential for the biogenesis of QHNDH because of their involvement in the multiple steps of post-translational modification [7–11]. In contrast, the ΔqhpH strain exhibited the same level of QHNDH activity as Pd1222, indicating that qhpH is not involved in the biogenesis of QHNDH.

Additionally, when the wild-type Pd1222 and ΔqhpH strains were cultured in the minimal medium containing n-butyraldehyde as the sole carbon source, both strains grew equally well, with the growth curves showing that the disruption of qhpH did not affect the growth of P. denitrificans on n-butyraldehyde (Fig. 3B). This suggests the presence of another intrinsic (constitutive) aldehyde dehydrogenase(s) that can compensate for the defect of qhpH and the metabolic redundancy for aldehyde oxidation. In fact, 29 genes including qhpH are annotated as aldehyde dehydrogenases or related enzymes in the Pd1222 genome (Table S2). This redundancy underscores the robust metabolic versatility of P. denitrificans [19], allowing it to adapt to various environmental conditions by utilizing different enzymes for similar metabolic functions.

Fig. 3. Growth of wild-type Pd1222 and qhpH-disrupted (ΔqhpH) strains of P. denitrificans in the minimal medium containing n-BA or n-butyraldehyde.

Each bacterial strain was grown in the minimal medium supplemented with 46 mM n-BA (A) or 14 mM n-butyraldehyde (B). Cell densities, measured as optical density at 600 nm (OD₆₀₀), were plotted against culture time. The growth of wild-type Pd1222 (■), ΔqhpH (△), and ΔqhpC (○), as a negative control [8], is shown. Each bar represents the mean ± standard deviation from two independent experiments.

3.3. Enhanced growth of Rhodobacter sphaeroides co-expressing QhpH and QHNDH

A photosynthetic purple bacterium, Rhodobacter sphaeroides, is a close relative of Paracoccus denitrificans, but can assimilate n-BA only when qhpABCDEFGR is expressed heterologously [12]. In this study, we further examined the effect of the additional expression of qhpH in the recombinant R. sphaeroides, in which qhpABCDEFGR gene cluster was co-expressed. As shown in Fig. 4, R. sphaeroides transformed with pRK-qhpGADCBEFRH exhibited significantly faster growth than those transformed with pRK-qhpGADCBEFR alone. In the absence of the qhp genes, the bacterial strain did not grow at all. These results suggest that the presence or absence of qhpH expression affects the growth rate in the n-BA minimal medium during the heterologous expression of qhp genes in R. sphaeroides. This contrasts with the results observed with P. denitrificans. The observed differences could be due to the following reason: With R. sphaeroides, the maximal OD₆₀₀ reached approximately 0.2 after 24 h of cultivation in the minimal medium containing 0.1% (14 mM) n-butyraldehyde. However, when the same amount of n-butyraldehyde was added in five divided doses at 12-h intervals, the OD₆₀₀ reached approximately 1.2 after 72 h [12]. This suggests that R. sphaeroides possesses aldehyde dehydrogenase(s) that use n-butyraldehyde as a non-preferred substrate. In the genome of R. sphaeroides NBRC 12203, 15 genes are annotated as aldehyde dehydrogenases or related enzymes (not shown). However, their activities and/or expression level are insufficient, leading to an increase in intracellular n-butyraldehyde concentration, which inhibits cell growth due to its toxicity through reacting non-specifically with enzymes, DNA, structural proteins, and other macromolecules [20]. In contrast, P. denitrificans cultured in the minimal medium containing 0.1% n-butyraldehyde grew to a maximal OD₆₀₀ of approximately 0.8 after 24 h without the need to divide the addition of n-butyraldehyde (Fig. 3B). These observations indicate that P. denitrificans has a higher capacity to assimilate n-butyraldehyde compared to R. sphaeroides. This suggests that the activity or expression level of aldehyde dehydrogenases, other than QhpH, that use n-butyraldehyde as a substrate is higher in P. denitrificans. Therefore, the additional qhpH co-expression should affect the growth rate when qhpABCDEFGR are expressed heterologously in R. sphaeroides (Fig. 4).

Fig. 4. Growth of R. sphaeroides (Rsp) harboring QHNDH expression plasmids with or without qhpH.

R. sphaeroides (Rsp) transformed with pRK-qhpGADCBEFRH (■) or pRK-qhpGADCBEFR (△) [12] were grown in the minimal medium supplemented with 46 mM n-BA. As a negative control, Rsp without any plasmid (○) was also grown in the minimal medium. Cell densities, measured as optical density at 600 nm (OD₆₀₀), were plotted against culture time. Each bar represents the mean ± standard deviation from two independent experiments.

3.4. Substrate specificity of recombinant QhpH

To study the substrate specificity of QhpH as an aldehyde dehydrogenase, N-terminally His₆-tagged QhpH was expressed in E. coli cells and purified by Ni-affinity chromatography. Judging from SDS-PAGE analysis (Fig. 5), QhpH after removal of the His₆-tag was purified to essentially homogeneity, and the estimated molecular mass (about 50 kDa) was consistent with that predicted from the translated amino acid sequence (49,767 Da) (Fig. 1B). The final yield of the purified protein was 15 mg from 100-mL culture, corresponding to 1.5 mg of protein per gram of wet cells. Although qhpH was initially annotated as a betaine aldehyde dehydrogenase gene in the whole genome analysis of P. denitrificans Pd1222, the purified QhpH did not exhibit detectable activity with betaine aldehyde as the substrate. To search for real aldehyde substrates, the enzymatic activities of QhpH were measured using various aldehydes as substrates in the presence of NAD⁺ (Table 2). The kinetic constants of QhpH indicate that it functions as an aliphatic and aromatic aldehyde dehydrogenase capable of oxidizing n-butyraldehyde and phenylacetaldehyde as substrates. Among examined substrates, acetaldehyde was the most preferred substrate showing the highest k_cat/K_m value. When n-butyraldehyde was used as a substrate, NADP⁺ did not serve as an electron acceptor. Since QHNDH exhibits broad substrate specificity for various primary amines and converts them to the corresponding aldehydes [3], the finding that QhpH exhibits substrate specificity for these aliphatic and aromatic aldehydes is reasonable for their metabolic cooperation. These results confirm the role of QhpH as an NAD⁺-specific aldehyde dehydrogenase with broad substrate specificity, further supporting its involvement in the oxidation of aldehydes produced by QHNDH in the assimilation of various primary amines.

Since QhpH lacks a signal sequence at the N-terminus (Fig. 1B), it is likely localized in the cytoplasm. In contrast, QHNDH is localized in the periplasm, so the aldehyde produced in the periplasm would need to be translocated to the cytoplasm before being processed by QhpH and other aldehyde dehydrogenases. Similar observations have been reported for amine metabolism in Klebsiella oxytoca [1]. The cytoplasmic localization of QhpH, in contrast to the periplasmic localization of QHNDH, likely reflects the need for a continuous NAD⁺ supply to sustain its catalytic activity.

Table 2. Kinetic parameters for QhpH.

Substratea	Km (µM)	kcat (s−1)	kcat/Km (mM−1 s−1)
n-butyraldehyde	13.9 ± 1.6	1.15 ± 0.04	82.6 ± 9.7
propionaldehyde	70.6 ± 5.6	2.81 ± 0.10	39.8 ± 3.5
acetaldehyde	48.1 ± 1.3	31.1 ± 1.0	647 ± 28
formaldehyde	6950 ± 740	8.72 ± 0.76	1.26 ± 0.17
phenylacetaldehyde	1700 ± 400	3.2 ± 0.7	1.9 ± 0.6
benzaldehyde	94.4 ± 6.3	2.69 ± 0.14	28.5 ± 2.4

^a Apparent kinetic parameters were measured at 25 °C in 50 mM Tris-HCl (pH 7.4) with 1 mM NAD⁺. Data are shown as the mean ± standard error calculated from a linear regression analysis of the plotted data.

Fig. 5. SDS-PAGE analysis of recombinant QhpH purification.

QhpH was expressed in E. coli cells as a fusion protein with an N-terminal His₆-tag. The soluble cell lysate (lane S) was applied to a Ni-affinity column. The flow-through (lane F1) and the eluted fractions (lane E) were collected. The eluted fraction was treated with TEV protease and then passed through a desalting column to remove imidazole and simultaneously cleave the His₆-tag. This was followed by a second Ni-affinity chromatography step; the flow-through fraction (lane F2) was collected and used as the purified protein. The molecular masses of the markers (lane M) are indicated on the left (kDa). The protein bands were stained with colloidal Coomassie blue [21].

Conclusion

Our present study demonstrates that the nine genes of qhpABCDEFGHR form a minimal gene cluster necessary for primary amine assimilation in P. denitrificans. The genetically conserved qhpH gene is co-expressed with the qhp operon and functions as an aldehyde dehydrogenase involved in amine assimilation of a large number of Gram-negative bacteria. Notably, the disruption of qhpH did not impede the growth of P. denitrificans in the minimal medium with n-BA, suggesting the presence of compensatory (constitutive) aldehyde dehydrogenases. This indicates a potential redundancy in metabolic pathways, underscoring the robust metabolic adaptability of this bacterium. Furthermore, the enhanced growth of R. sphaeroides co-expressing qhpH with other qhp genes highlights the importance of qhpH in more efficient amine metabolism than its absence.

CRediT authorship contribution statement

Tadashi Nakai: Writing – original draft, Writing – review & editing, Methodology, Investigation, Validation, Formal analysis, Funding acquisition, Project administration, Supervision, Conceptualization. Naoya Miyagi: Investigation. Kota Hisamura: Investigation. Shoya Matsuba: Investigation. Kanji Nishimoto: Investigation. Emi Nakai: Investigation. Katsuyuki Tanizawa: Writing – review & editing, Supervision, Conceptualization. Toshihide Okajima: Writing – review & editing, Funding acquisition, Supervision, Conceptualization.

Declaration of generative AI and AI-assisted technologies in the writing process

During the preparation of this work, the authors used OpenAI's ChatGPT in order to correct grammatical and spelling errors and enhance the readability and language of the manuscript. After using this tool, the authors reviewed and edited the content as needed and take full responsibility for the content of the published article.

Data Availability statement

The data underlying this article will be shared on reasonable request to the corresponding author.

Funding

This research was supported by the Japan Society for the Promotion of Science KAKENHI under grant numbers JP 16K07691 and JP 22K05419 to T.N., and JP 15K07391 and JP 22K19150 to T.O., and by the Cooperative Research Program of “Network Joint Research Center for Materials and Devices.”

Declaration of competing interest

The authors declare no competing interests.

Acknowledgements

We thank Daiki Momosaki, Riki Otsu, and Hiroaki Fujimoto for their contributions to plasmid construction and recombinant protein purification in the early stages of this work.

Appendix A. Supplementary data

Supplementary data for this article can be found at the end of this file.

References

1. Hacisalihoglu A, Jongejan JA, Duine JA. Distribution of amine oxidases and amine dehydrogenases in bacteria grown on primary amines and characterization of the amine oxidase from Klebsiella oxytoca. Microbiology 1997;143:505-12. https://doi.org/10.1099/00221287-143-2-505
2. Adachi O, Kubota T, Hacisalihoglu A et al. Characterization of quinohemoprotein amine dehydrogenase from Pseudomonas putida. Biosci Biotechnol Biochem 1998;62:469-78. https://doi.org/10.1271/bbb.62.469
3. Takagi K, Torimura M, Kawaguchi K et al. Biochemical and electrochemical characterization of quinohemoprotein amine dehydrogenase from Paracoccus denitrificans. Biochemistry 1999;38:6935-42. https://doi.org/10.1021/bi9828268
4. Takagi K, Yamamoto K, Kano K et al. New pathway of amine oxidation respiratory chain of Paracoccus denitrificans IFO 12442. Eur J Biochem 2001;268:470-6. https://doi.org/10.1046/j.1432-1033.2001.01912.x
5. Datta S, Mori Y, Takagi K et al. Structure of a quinohemoprotein amine dehydrogenase with an uncommon redox cofactor and highly unusual crosslinking. Proc Natl Acad Sci USA 2001;98:14268-73. https://doi.org/10.1073/pnas.241429098
6. Satoh A, Kim JK, Miyahara I et al. Crystal structure of quinohemoprotein amine dehydrogenase from Pseudomonas putida. Identification of a novel quinone cofactor encaged by multiple thioether cross-bridges. J Biol Chem 2002;277:2830-4. https://doi.org/10.1074/jbc.M109090200
7. Nakai T, Deguchi T, Frebort I et al. Identification of genes essential for the biogenesis of quinohemoprotein amine dehydrogenase. Biochemistry 2014;53:895-907. https://doi.org/10.1021/bi401625m
8. Ono K, Okajima T, Tani M et al. Involvement of a putative [Fe-S]-cluster-binding protein in the biogenesis of quinohemoprotein amine dehydrogenase. J Biol Chem 2006;281:13672-84. https://doi.org/10.1074/jbc.M600029200
9. Nakai T, Ito H, Kobayashi K et al. The radical S-adenosyl-L-methionine enzyme QhpD catalyzes sequential formation of intra-protein sulfur-to-methylene carbon thioether bonds. J Biol Chem 2015;290:11144-66. https://doi.org/10.1074/jbc.M115.638320
10. Nakai T, Ono K, Kuroda S et al. An unusual subtilisin-like serine protease is essential for biogenesis of quinohemoprotein amine dehydrogenase. J Biol Chem 2012;287:6530-8. https://doi.org/10.1074/jbc.M111.324756
11. Oozeki T, Nakai T, Kozakai K et al. Functional and structural characterization of a flavoprotein monooxygenase essential for biogenesis of tryptophylquinone cofactor. Nat Commum 2021;12:933. https://doi.org/10.1038/s41467-021-21200-9
12. Nakai T, Tanizawa K, Okajima T. Eight genes are necessary and sufficient for biogenesis of quinohemoprotein amine dehydrogenase. Biosci Biotechnol Biochem 2021;85:2026-9. https://doi.org/10.1093/bbb/zbab117
13. Keen NT, Tamaki S, Kobayashi D et al. Improved broad-host-range plasmids for DNA cloning in gram-negative bacteria. Gene 1988;70:191-7. https://doi.org/10.1016/0378-1119(88)90117-5
14. Miller JH. Experiments in Molecular Genetics, Cold Spring Harbor Laboratory Press, Plainview, NY 1972.
15. van Spanning RJ, Wansell CW, Reijnders WN et al. A method for introduction of unmarked mutations in the genome of Paracoccus denitrificans: Construction of strains with multiple mutations in the genes encoding periplasmic cytochromes c₅₅₀, c_551i, and c_553i. J Bacteriol 1991;173:6962-70. https://doi.org/10.1128/jb.173.21.6962-6970.1991
16. Sistrom WR. The kinetics of the synthesis of photopigments in Rhodopseudomonas sphaeroides. J Gen Microbiol 1962;28:607-16. https://doi.org/10.1099/00221287-28-4-607
17. Studier FW. Protein production by auto-induction in high-density shaking cultures. Prot Exp Pur 2005;41:207-34. https://doi.org/10.1016/j.pep.2005.01.016
18. Nakai T, Kuramitsu S, Kamiya N. Structural bases for the specific interactions between the E2 and E3 components of the Thermus thermophilus 2-oxo acid dehydrogenase complexes. J Biochem 2008;143:747-58. https://doi.org/10.1093/jb/mvn033
19. Bordel S, Martín-González D, Börner T et al. Genome-scale metabolic model of the versatile bacterium Paracoccus denitrificans Pd1222. mSystems 2024;9:e0107723. https://doi.org/10.1128/msystems.01077-23
20. LoPachin RM, Gavin T. Molecular mechanisms of aldehyde toxicity: a chemical perspective. Chem Res Toxicol 2014;27:1081-91. https://doi.org/10.1021/tx5001046
21. Dyballa N, Metzger S. Fast and sensitive colloidal Coomassie G-250 staining for proteins in polyacrylamide gels. J Vis Exp 2009;30:1431. https://doi.org/10.3791/1431

Supplementary Information

Table S1. The oligonucleotides used in this study.

Primer name	Nucleotide sequencea	Restriction site and mutation
P1	5'-ATAGAATTCGGCGTTTCCCGTGGCTGGCGCCGGGT-3'	EcoRI
P2	5'-CGCGCATATGGTCCCTCCCCAAAACGGTCGGGCCA-3'	NdeI
P3	5'-ATATGGATCCTCAGCGGATCACGCTGACCTCTTC-3'	BamHI
P4	5'-ATATGGATCCTCAGCGCGCCTCGTTCACGATGCGG-3'	BamHI
P5	5'-GCGGATAACAATTTCACACAGGAAACAGC-3'
P6	5'-TTAGAAAAACTCATCGAGCATCAAATGAAAC-3'
P7	5'-GGTATTGATAATCCTGATATGAATAAATTGCAGTT-3'
P8	5'-CACCTGATTGCCCGACATTATCGCGAGCCCATTTA-3'
P9	5'-ATGACGGAATATCGATTGCTGATCGACGGCAGGCT-3'
P10	5'-GCAAACAGCCCCGGCGCGACCTTG-3'
P11	5'-GACAAGCTTGAGGCGCAGGCCGAGGAGTTCGCCCGGCTGCTGACGCAGGAA-3'	HindIII, GAGb
P12	5'-CGCGAATTCTCAGCGCGCCTCGTTCACGATGCGG-3'	EcoRI
P13	5'-ATATCATATGACCGAATATCGATTGCTGATCGACGGCAGGC-3'	NdeI, ACCc

^a Restriction sites are underlined. Mismatched nucleotides at the mutated sites are shown in bold.

^b Silent mutation (GAA to GAG) to eliminate an EcoRI site (GAATTC).

^c Silent mutation (ACG to ACC) at the second codon to enhance translation efficiency in E. coli [1].

Reference

[1] Looman AC, Bodlaender J, Comstock LJ et al. Influence of the codon following the AUG initiation codon on the expression of a modified lacZ gene in Escherichia coli. EMBO J 1987;6:2489-92. https://doi.org/10.1002/j.1460-2075.1987.tb02530.x

Table S2. Genes encoding aldehyde dehydrogenases and related enzymes in P. denitrificans Pd1222 genome^a.

Entry	Protein name	Gene name
A1B0L0	Bifunctional protein PutA (L-glutamate gamma-semialdehyde dehydrogenase)	Pden_0943
A1B4L2	Aldehyde dehydrogenase (EC 1.2.1.3) (Acetaldehyde dehydrogenase)	Pden_2366
A1AYF2	Aldehyde dehydrogenase	Pden_0180
A1AYJ3	Succinate semialdehyde dehydrogenase (NAD(P)+) (EC 1.2.1.16)	Pden_0223
A1AYL3	Succinate semialdehyde dehydrogenase (EC 1.2.1.16)	Pden_0243
A1AYQ9	Succinate semialdehyde dehydrogenase (EC 1.2.1.16)	Pden_0289
A1B0T0	Succinate semialdehyde dehydrogenase (NAD(P)+) (EC 1.2.1.16)	Pden_1013
A1B0W8	Aldehyde dehydrogenase (NAD+) (EC 1.2.1.3)	Pden_1051
A1B0W9	Aldehyde dehydrogenase (NAD+) (EC 1.2.1.3)	Pden_1054
A1B1G5	Aldehyde dehydrogenase	Pden_1254
A1B2R3	Betaine aldehyde dehydrogenase (EC 1.2.1.8)	Pden_1710
A1B398	Betaine aldehyde dehydrogenase (EC 1.2.1.8)	Pden_1897
A1B3I9	Aldehyde dehydrogenase	Pden_1991
A1B4A5	Succinate semialdehyde dehydrogenase (EC 1.2.1.16)	Pden_2257
A1B5P5	Aldehyde dehydrogenase	Pden_2755
A1B6Z8	Aldehyde dehydrogenase	Pden_3211
A1B715	Aldehyde dehydrogenase	Pden_3228
A1B745	Aldehyde dehydrogenase	Pden_3260
A1B748	Aldehyde dehydrogenase	Pden_3264
A1B8X0	Aldehyde dehydrogenase (NAD+) (EC 1.2.1.3)	Pden_3898
A1BAD6	Succinate semialdehyde dehydrogenase (EC 1.2.1.16)	Pden_4416
A1BAJ9	Methylmalonate-semialdehyde dehydrogenase (CoA acylating) (EC 1.2.1.27)	Pden_4479
A1BAR7	Aldehyde dehydrogenase	Pden_4548
A1BAX4	Aldehyde dehydrogenase domain-containing protein	Pden_4605
A1BBB9	Aldehyde dehydrogenase	Pden_4752
A1BBN1	Aldehyde dehydrogenase	Pden_4865
A1BBV7	Betaine aldehyde dehydrogenase (EC 1.2.1.8)	Pden_4941
A1BBY5	Succinate semialdehyde dehydrogenase (EC 1.2.1.16)	Pden_4969
A1BBY7	Aldehyde dehydrogenase	Pden_4971

^a The entries, protein names, and gene names were taken from UniProt.