Skip to main content

Carboxypeptidase G and pterin deaminase metabolic pathways degrade folic acid in Variovorax sp. F1



Folic acid (FA) is a synthetic vitamin (B9) and the oxidized form of a metabolic cofactor that is essential for life. Although the biosynthetic mechanisms of FA are established, its environmental degradation mechanism has not been fully elucidated. The present study aimed to identify bacteria in soil that degrade FA and the mechanisms involved.


We isolated the soil bacterium Variovorax sp. F1 from sampled weed rhizospheres in a grassland and investigated its FA degradation mechanism. Cultured Variovorax sp. F1 rapidly degraded FA to pteroic acid (PA), indicating that FA hydrolysis to PA and glutamate. We cloned the carboxypeptidase G (CPG) gene and found widely distributed paralogs within the Variovorax genus. Recombinant CPG preferred FA and deaminofolic acid as substrates, indicating its involvement in FA degradation by Variovorax. Prolonged culture of Variovorax sp. F1 resulted in decreased rates of deaminofolic acid (DFA) and deaminopteroic acid (DPA) accumulation. This indicated that the deamination reaction also comprised a route of FA degradation. We also identified an F1 gene that was orthologous to the pterin deaminase gene (Arad3529) of Agrobacterium radiobacter. The encoded protein deaminated FA and PA to DFA and DPA, which was consistent with the deamination activity of FA and PA in bacterial cell-free extracts.


We discovered that the two enzymes required for FA degradation pathways in isolates of Variovorax sp. F1 comprise CPG and pterin deaminase, and that DFA and PA are intermediates in the generation of DPA.

Peer Review reports


Vitamins are essential coenzymes that regulate cellular metabolism. Humans and other animals cannot synthesize most essential vitamins so they must be ingested from foods, to avoid deficiencies that can cause various symptoms and pathological states [1, 2]. Thus, the physiological functions of vitamins have been established [3]. Autotrophic plants and microorganisms produce vitamins via known synthetic mechanisms. Chemically synthesized vitamins support an increasing demand for food supplements [4,5,6]. The amounts of vitamins in living organisms are homeostatic and maintained, but the molecular mechanisms of vitamin decomposition by ecological systems are not fully understood. Examples are recent findings of bacterial genes encoding proteins that degrade thiamine (vitamin B1) [7], riboflavin (vitamin B2) [8], nicotinamide (vitamin B3) [9], pyridoxine (vitamin B6) [10], and L-ascorbate (vitamin C) [11, 12].

Folic acid (FA, vitamin B9) is a synthetic N-pteroyl-p-aminobenzoylglutamate and precursor of tetrahydrofolic acid (THFA). Its related compounds are involved in cellular one-carbon metabolism [13, 14], and it is essential for human functions. A deficiency of THFA and its related compounds results in anemia, stunted growth, and neural tube defects in newborns. Folic acid is therefore a therapeutic agent and nutrient supplement that is particularly important for pregnant women [15, 16]. Folic acid is chemically synthesized on an industrial scale, whereas its microbial fermentation is under development [17]. The structure of FA comprises pterin, p-aminobenzoic acid (PABA), and glutamate moieties [18]. The de novo biosynthetic pathway of THFA starts from guanosine triphosphate (GTP), and mediates dihydropterin, which is conjugated with p-aminobenzoic acid and glutamate to generate the reduced forms of FA, dihydrofolic acid and tetrahydrofolic acids. Multiple conjugation reactions of glutamate generate polyglutamate derivatives. The extent of polyglutamylation varies among species and it regulates cofactor affinity for enzymes and subcellular compartmentation in plants [19].

Despite extensive investigation into biosynthesis of THFA and its related compounds, the mechanisms of their degradation in animals and the environment are not fully understood. Tetrahydrofolic acids and their products derived from plants and microorganisms like other natural compounds, are considered to be decomposed in the material cycles of soil. Tetrahydrofolic acids are quite easily oxidized to FA in fertile aerobic soils. Therefore, elucidating the taxonomic distribution and physical location of bacteria that degrade FA is important for understanding microbial community involvement in the FA material cycle. Furthermore, understanding the bacterial enzymes and genes responsible for FA degradation is important to determine the molecular mechanisms of THFA mineralization by soil ecosystems.

A Pseudomonad bacterium isolated in 1967 hydrolyzed FA to pteroic acid (PA) and glutamate [20], then a protein in the carboxypeptidase G family (CPG; EC catalyzing this reaction was identified [20,21,22,23] and their encoding genes were cloned from several Pseudomonas species [24]. Pterin deaminase (PDA; EC deaminates FA to deaminofolic acid (DFA) in cell-free extracts of Alcaligenes metalcaligenes [25], A. faecalis [26], Flavobacterium polyglutamicum [27], Pseudomonas sp. Fo8 [28], and Bacillus megaterium [29]. Genetic evidence of bacterial PDA for FA had remained obscure before homology modeling and identification of the Arad3529 protein from Agrobacterium radiobacter K84 that deaminates FA [30]. Some PDA deaminates PA to deaminopteroic acid (DPA) [25, 26, 28], but whether PDA participates in PA deamination in A. radiobacter K84 cells has remained obscure. N-(4-aminobenzoyl)-L-glutamic acid (ABG) is an oxidation product of THFA [31], and Escherichia coli BN101 produces its ABG hydrolase, AbgAB, that seems inert against FA [32]. Besides these findings, bacteria that utilize both a carboxypeptidase and a deaminase to degrade FA are limited to the tentatively classified Pseudomonas sp. Fo8 that was discovered in 1974 [28]. However, the culture properties of this bacterium, the genetic basis of the mechanism of FA degradation and its classification have remained unknown.

Here, we screened a series of soil samples for FA-degrading bacteria. We identified the novel FA-degrading bacterium Variovorax sp. F1, which consumed FA and accumulated PA in vitro, indicating that it utilizes glutamate liberated from FA. The accumulation of both DFA and DPA in cultures indicated that Variovorax sp. F1 deaminates FA and PA. We cloned the Variovorax sp. F1 genes for CPG and PDA, and recombinant CPG (rCPG) that hydrolyzed FA and deaminofolic acid (DFA). Recombinant PDA (rPDA) deaminated FA and PA to DFA and DPA, respectively, indicating two pathways of FA degradation in the soil bacterium Variovorax sp. F1.


Isolation of soil bacteria that degrade FA

Six environmental soil samples were cultured in M9 Minimal Medium containing FA as the sole carbon source (M9-FA medium) to enrich bacteria that can degrade FA. Orange FA disappeared and yellow insoluble pigments appeared on plates containing M9-FA agar inoculated with eight cultured samples (Fig. 1A). Cultures in liquid M9-FA medium also consumed FA to form yellow insoluble pigments, which were solubilized in 0.1 M NaOH, and confirmed as PA by high performance liquid chromatography (HPLC) equipped with an alkaline-tolerant column and a photodiode array detector. All isolates consumed most of the FA and some accumulated less PA than the amount of consumed FA (Fig. 1B). We further analyzed F1 among the eight strains. The nucleotide sequence of the 16S rRNA gene of the F1 strain was 99.9% identical to that of Variovorax paradoxus NBRC 15149T and V. boronicumulans NBRC 103145T. Phylogenetic analyses indicated that the F1 strain is a β-proteobacterium related to the genus Variovorax (Fig. 1C) and was identified as Variovorax sp. F1. Partial sequencing of the 16 S rRNA gene revealed that the other seven isolates were related to the Variovorax and Xenophilus genera and belonged to the Comamonadaceae family (Table S1).

Fig. 1
figure 1

Isolation of FA degrading bacteria. A Growth of eight isolates on M9-FA agar plates at 28 °C for 48 h. B Degradation of FA to PA by isolates cultured in M9-FA medium at 28 °C for 48 h. Filled bars, FA; unfilled bars, PA. Error bars, standard errors of means (n = 3, p < 0.05). *P < 0.05 (control FA vs. accumulated PA). C Nucleotide sequences of selected strains were aligned, and phylogenetic trees were constructed by neighbor-joining method [33] using MEGA X software [34]. Numbers along branches indicate 1000 bootstrap replicates D Time-dependent decomposition of FA by typical culture of Variovorax sp. F1 in M9-FA medium at 28 °C for 48 h. , FA; ■, PA; ▲, DFA; , DPA. Bars, cell growth estimated as total cellular proteins. Error bars, standard errors of means (n = 3). DFA, deaminofolic acid; DPA, deaminopteroic acid; FA, folic acid; PA, pteroic acid

We investigated time-dependent changes in FA and its degradation products in cultured F1 (Fig. 1D). This strain consumed > 90% of the initial 10 mM FA within 36 h when cultured in liquid M9-FA medium. The accumulation of ~ 8 mM PA indicated that the strain almost stoichiometrically converted FA to PA. The growth of Variovorax sp. F1 was measured as total protein in cultures using the Bradford method because insoluble FA and PA interfere with conventional measurements of cell mass based on optical density. The results indicated concomitant cell growth and FA conversion to PA, which implied that Variovorax sp. F1 degrades FA to PA and glutamate and utilizes it as a carbon source for growth. This notion is supported by fact that the rates of F1 growth were similar in medium containing glutamate as carbon and nitrogen sources and FA as a carbon source (M9-FA medium). Analysis of prolonged cultures using HPLC produced additional compounds that eluted at the same retention times as DFA and DPA prepared and identified herein (see Materials and methods). Incubation for 36 h resulted in the relatively low accumulation of DFA and DPA (~ 70 and < 5 μM, respectively), whereas culture for 264 h increased these amounts to 180 μM and 1.4 mM, respectively. These results suggested that the F1 strain respectively hydrolyzed FA and DFA to PA and DPA, then deaminated FA and PA to DFA and DPA (Fig. 2).

Fig. 2
figure 2

Proposed FA degradation pathway in Variovorax sp. F1. Schematic model of FA degradation by Variovorax sp. F1. ABG, N-(4-aminobenzoyl)-L-glutamic acid; CPG, carboxypeptidase G; DFA, deaminofolic acid; DPA, deaminopteroic acid; FA, folic acid; PA, pteroic acid; PABA, p-aminobenzoic acid; PDA, pterin deaminase

Variovorax sp. F1 CPG for FA utilization

We considered that Variovorax sp. F1 would produce a counterpart of CPG2 in Pseudomonas sp. RS-16 [23] and other bacteria [22,23,24,25,26,27,28,29,30,31,32, 35, 36] to hydrolyze the C-terminal glutamate portion of FA. We therefore cloned the CPG2 paralog of the F1 strain. Database searches predicted CPG proteins from the Variovorax spp. SCN67–85, KB5, OV084, YR634, and root434, with amino acid sequence identities of 94.0–95.2%. We amplified the Variovorax sp. F1 CPG gene by PCR using custom-designed primer sets for conserved nucleotide sequences and total DNA of Variovorax sp. F1. The gene comprised an open-reading frame encoding 415 amino acids that was 93.5% identical to CPG2 of the most intensively studied Pseudomonas sp. RS-16 [37], and 94.7 and 97.6% identical respectively, to the CPGs predicted from V. paradoxus strain EPS (Varpa5372) and V. boronicumulans (CKY39_29385). The results of phylogenetic analyses showed that the CPG of the F1 strain was the most closely related to the predicted CPG of V. boronicumulans and V. paradoxus (Fig. 3). The phylogenetic tree contained more than one CPG from individual Variovorax strains in different clades. The amino acid sequence of the CPG of the F1 strain was 44.1–45.9% identical to those of CPGs with uncharacterized FA hydrolytic capacity located in different clades.

Fig. 3
figure 3

Phylogenetic relationship of CPG from Variovorax sp. F1. Amino acid sequences of predicted CPG from selected strains were aligned and phylogenetic trees were constructed by neighbor-joining [33] using MEGA X software [34]. Numbers along branches indicate 1000 bootstrap repeats. Gene IDs are shown after strain names in parenthesis

We generated a recombinant CPG (rCPG) of Variovorax sp. F1 with a 6 × histidine tag on the amino terminus using an E. coli expression system. The purified rCPG resolved on SDS-PAGE as a single band at 45 kDa (Fig. 4A). The reaction of rCPG and FA generated PA (Fig. 4B), indicating that rCPG hydrolyzes FA to PA and glutamate like the Pseudomonas CPG2. The initial velocity of the reaction against 5 mM FA was 52 ± 3 μmol min− 1 mg− 1 (Fig. 4B), and comparable to that of the known CPG (40–725 μmol min− 1 mg− 1) [21,22,23]. The reaction was inhibited by 1 mM EDTA, which agrees with the Zn2+-dependent reaction of Pseudomonas CPG2 [23].

Fig. 4
figure 4

Enzymatic properties of rCPG derived from Variovorax sp. F1. A SDS-PAGE gel. Lanes: M, molecular weight marker; rCPG, Recombinant CPG (1.8 μg) from Variovorax sp. F1 strain. BD HPLC analyses of reactions containing 5 μg mL− 1 rCPG and various substrates at 30 °C for 15 min. Substrates (5 mM) each are B FA; C DFA; D ABG. Traces: 1, substrates; 2, products; 3, reaction. E Specific activities of rCPG against FA, DFA, and ABG. Error bars indicate standard errors of means (n = 3). *P < 0.05. ABG, N-(4-aminobenzoyl)-L-glutamic acid; DFA, deaminofolic acid; DPA, deaminopteroic acid; FA, folic acid; PA, pteroic acid; rCPG, recombinant carboxypeptidase G

Substrate specificity of rCPG

The substrate specificity of rCPG against ABG derivatives was investigated considering that rCPG hydrolyzes FA at the amide bond within the ABG residue. The substrates FA and ABG are commercially available, and we prepared DFA in-house. Briefly, we deaminated commercial FA in a preparative scale using recombinant PDA from the F1 strain to obtain DFA and purified it to > 99%. The reaction of rCPG and DFA generated another compound on HPLC analysis (Fig. 4C). This compound eluted at the same retention time as DPA, which we prepared and identified from a parent mass ion peak and a mass ion fragment at m/z = 312.2 ([M–H]) and 176.1, respectively, using LC-MS/MS (Fig. S1A; Materials and methods). These results indicated that rCPG hydrolases DFA to DPA.

Reactions of rCPG with FA, DFA and ABG (0.1 mM each) produced stoichiometric amounts of DPA and PABA (Fig. 4C, D) at rates of 1.0 ± 0.1 and 1.2 ± 0.1 μmol min− 1 mg− 1, respectively (Fig. 4E). These results indicated that FA, DFA and ABG are substrates of rCPG. The initial velocity of the rCPG reaction linearly increased as substrate concentrations increased to 30, 50, and 20 mM FA, DFA, and ABG, respectively (Fig. S2). Since the initial velocities of the enzyme did not reach saturation according to substrate concentrations, the Michaelis (Km) and kinetic (kcat) constants for these reactions could not be determined.

Identification of PDA from Variovorax sp. F1

The accumulation of DFA and DPA in culture medium of the F1 strain (Fig. 1D) indicated their production via FA and PA deamination. We searched for a Variovorax F1 protein orthologous to A. radiobacter K84 PDA (Arad3529) which was the only PDA identified to date as an enzyme that deaminates FA to DFA. Database searches identified a predicted deaminase (WP 068679287) in many Variovorax spp. that has 36.0% amino acid sequence identity with Arad3529. We aimed to construct a set of primers with a conserved nucleotide sequence among V. paradoxus orthologs to clone the corresponding gene in F1 strain. However, we found a conserved sequence for the primer corresponding to the 5′-, but not the 3′-end of the gene. We therefore designed a 3′-end primer with reference to conserved endoribonuclease genes located downstream of the predicted PDA genes in the V. paradoxus genome. The Variovorax sp. F1 PDA gene was amplified by PCR using the primers for the conserved nucleotide sequences and total DNA of Variovorax sp. F1. The DNA fragment contained a gene encoding 399 amino acid residues that were 76.7 and 37.0% identical, respectively, to the protein predicted from V. paradoxus S110 (Vapar5141) and Arad3529. These proteins were hydrolases that act on non-peptide carbon-nitrogen bonds (EC 3.5), which are diverse among bacteria. The Variovorax paradoxus S110 genome encodes 89 such hydrolases. Our phylogenetic analyses classified many of these enzymes based on the molecular structures of the substrates that they hydrolyze (Fig. 5). Variovorax sp. F1 PDA and Arad3529 were located in a branch of putative hydrolases that act on cyclic amidines (EC 3.5.2).

Fig. 5
figure 5

Phylogenetic relationships of deaminase from Variovorax sp. F1 PDA. Amino acid sequences of putative hydrolases that catalyze non-peptide carbon-nitrogen bond cleavage (EC 3.5) from V. paradoxus S110 were aligned, and phylogenetic trees were constructed by neighbor-joining [33] using MEGA X software [34]. Numbers along branches indicate 500 bootstrap repeats. Gene IDs are color-coded according to their enzyme families as: hydrolases acting on linear amides (red), cyclic amides (blue), linear amidines (green), cyclic amidines (yellow), and nitriles (gray)

We produced recombinant PDA (rPDA) from Variovorax sp. F1 using an E. coli expression system. Purified rPDA resolved as a single 45 kDa band on SDS-PAGE (Fig. 6A). The reaction between rPDA and FA generated a novel compound that was separated by HPLC (Fig. 6B, left). The LC-MS findings showed a parent mass ion peak at m/z = 441.2 ([M–H]), which corresponded to the molecular mass (Mr 442) of DFA (Fig. S1B). The mass ion fragments at m/z = 312.2 and 176.0 generated by this compound were consistent with the structure of deaminated pterin (Fig. S1B). The reaction of rPDA and PA generated another compound (Fig. 7C), which eluted at the same retention time as DPA (Fig. S1A). These results showed that this novel enzyme deaminated FA and PA and was Variovorax sp. F1 PDA.

Fig. 6
figure 6

Enzymatic properties of rPDA from Variovorax sp. F1. A SDS-PAGE gel. Lanes: M, molecular weight marker; rPDA, Recombinant PDA (2 μg) from Variovorax sp. F1 strain. B HPLC analyses of PDA reactions containing 0.01 μg mL− 1 rPDA and 1 mM substrate (left, FA; right, PA) at 30 °C for 2 min. Traces: 1, substrates; 2, products; 3, reaction. C Initial velocity of rPDA reaction determined to calculate Km and kcat values of rPDA for FA (left) and PA (right) as substrates. Data were fitted to Michaelis-Menten equation

Fig. 7
figure 7

Reactions of cell-free extract of Variovorax sp. F1 with various substrates. Folic acid (A) degradation of DFA (B), PA (C), and ABG (D) in cell-free extracts prepared from F1 strain cultured in M9-FA medium. Reactions proceeded in 50 mM Tris-HCl (pH 7.5) containing 0.2 mM ZnSO4, cell-free extract (5 μg mL− 1 protein) and substrates at 30 °C for 4 h. No DPA was degraded. , FA; ■, PA; ▲, DFA; , DPA; , ABG; , PABA. Error bars, standard errors of means (n = 3)

Steady-state kinetics of rPDA

The initial velocity of the PDA for deaminating FA and PA (1 mM each) was respectively 11.7 ± 1.2 and 0.31 ± 0.05 μmol min− 1 mg− 1. The reaction kinetics fit the Michaelis-Menten equation, with Km and kcat values of 0.28 ± 0.06 mM and 10.1 ± 0.4 s− 1 for FA, and 1.5 ± 0.6 mM and 0.62 ± 0.06 s− 1 for PA (Fig. 6C). The Km values were comparable to the concentrations of FA and PA in the cultures. These results indicated that Variovorax sp. F1 PDA uses both FA and PA as substrates but preferentially deaminates FA. This finding supports the phylogenetically close relationship of F1 PDA to the predicted cytosine and creatine amidases (Vapar3881, Vapar4756, Vapar2654), the latter of which shares a guanidine moiety with pterins that are deaminated by PDAs.

Cell-free activity reveals potential FA-degradation mechanism

The discovery of CPG and PDA genes in the F1 strain and accumulation of DFA and DPA in the culture suggested that the CPG-dependent cleavage of glutamate residues and deamination of the pterin moiety constitute an FA-degradation pathway. Therefore, we validated these activities in cell-free extracts of the cultured F1 strain that degraded FA. Reactions between cell-free extracts and FA resulted in decreased FA with a specific activity of 49 ± 2 nmol min− 1 mg− 1 (Fig. 7A). The reaction products were identified as essentially equal amounts of PA and DFA, indicating that the cells had CPG and PDA activities. Reaction with the cell-free extract resulted in the stoichiometric conversion of DFA to DPA (Fig. 7B) The reaction rate was faster than that for FA hydrolysis to PA (59 ± 6 vs. 20 ± 1 nmol min− 1 mg− 1; Fig. 7A), which agreed with our findings that bacterial rCPG hydrolyzed FA and DFA (Fig. 4D). We also found PA deamination activity in cell-free extracts with a specific activity of 37 ± 2 nmol min− 1 mg− 1 (Fig. 7C). This reaction produced DPA, which was the final product of the long-term culture (Fig. 1C). The cell-free extracts decomposed minimal amounts of DPA under these reaction conditions. Reactions between the cell-free extract and ABG, which is a product of THFA oxidation [31], generated PABA at a rate of 39 ± 1 nmol min− 1 mg − 1 (Fig. 7D). These results indicated that the F1 strain degrades FA to DPA, and that PA and DFA are intermediates in the mechanism of bacterial FA degradation (Fig. 2).


Ecological systems maintain the homeostasis of almost all types of biological molecules including those that are bulk-produced to generate physiologically active compounds. Among them, FA is a synthetic vitamin and popular food supplement that natural microorganisms can decompose. This study found that Variovorax sp. F1 in soil degrades FA via two distinct CPG and PDA pathways that mediate PA and DFA to produce DPA and supply this bacterium with glutamate as carbon/energy sources for growth. These findings are consistent with bacterial CPG hydrolysis, FA deamination, and FA degradation to DPA that have not been explored for decades [28]. Conventional HPLC using acidic water and solvents is inappropriate for analyzing extremely soluble FA and related compounds. An alkaline-tolerant HPLC system enabled analysis of alkali-soluble compounds and their high-throughput quantitation. Consequently, we could identify the genus, enzymes, and culture properties associated with bacterial FA degradation.

Bacterial FA degradation was long considered to involve CPG and deaminase, but the reconstitution of reactions by two enzymes originating from a single Variovorax species is novel. The CPG of Variovorax sp. F1 was phylogenetically similar to that of Pseudomonas sp. RS-16 (CPG2) and explains why FA breakdown generated PA as a major product at a relatively rapid rate (~ 48 h; Fig. 1D). We found that DFA and DPA were also degradation products of FA (Fig. 2), indicating that the bacterium deaminated FA before CPG decomposed DFA to DPA. Time-dependent changes in the accumulation of less DFA and DPA (deaminase products) than PA (CPG product) in the culture broth revealed a slower deamination rate than that catalyzed by CPG (Fig. 1D). We reconstituted the bacterial PDA deamination reactions of FA and PA to DFA and DPA, respectively (Figs. 6 and 7). As far as we can ascertain, this is the first PDA for which the gene was cloned from FA-degrading bacteria, and the second example of a PDA with amino acid sequences that are similar to those of A. radiobacter PDA (Arad3529). The F1 PDA was located in the same branch as other predicted Variovorax proteins and Arad3529 (Fig. 5). Thus, our findings will enable the future discovery of other novel PDAs (EC from proteins that are predicted deaminases for cyclic amidines (EC 3.5.4).

Cultured Variovorax sp. F1 consumed more FA than the degraded products (PA, DFA, and DPA) identified herein (Fig. 1C), indicating that this bacterium further catabolized these compounds. Lumazine-6-carboxylic acid and/or ABG are also candidate FA degradation intermediates that might be similarly generated in some bacteria [28]. The oxidative cleavage of DFA, DPA, and FA followed by deamination might generate lumazine-6-carboxylic acid. The rapid catalytic activity of rCPG against ABG (Fig. 5D) indicated the bacterial degradation of ABG to PABA and glutamate. Neither ABG nor lumazine-6-carboxylic acid were detectable in the cultured F1 strain, hence the mechanism of their production in this strain remains elusive.

The degradative mechanism of FA in ubiquitous Variovorax soil bacteria is likely to participate in the natural homeostasis of FA, THFA, and related compounds. Our extensive screen of sampled weed rhizospheres resulted mostly in Variovorax species (Table S1) and proteins with high similarity to CPG encoded by the Variovorax genome (Fig. 3). This implied that FA degrading activity is extant in Variovorax genera. Variovorax belongs to the recently identified Comamonadaceae family of bacteria [38] that thrive in fertile soils rich in organic matter mostly derived from plants [39, 40]. Thus, these bacteria should participate in degrading plant-derived materials in nature. Variovorax includes plant growth-promoting rhizobacteria that mutually interact with plants by producing enzymes that degrade plant hormone intermediates [41, 42]. The present study showed that Variovorax bacteria together with plants in the rhizosphere decomposed the plant-derived, physiologically active compound THFA that is oxidized to FA. The considerable accumulation of PA and DPA in cultured Variovorax raises the question of whether and how emerging groups of bacteria decompose these degradants in various environments.


The mechanisms through which soil bacteria degrade synthetic folic acid (FA) have remained unexplored. The present study isolated the novel soil-bacterium Variovorax sp. F1, which produced carboxypeptidase G that liberated glutamate residues of FA and deaminofolic acids, and a pterin deaminase that deaminated FA and PA. We consider that both enzymes comprise the bacterial mechanism of FA degradation.

Materials and methods

Strains, culture, and media

Soil microorganisms that degrade FA were enriched by culture in Minimal M9 Medium comprising 10 mM KH2PO4, 10 mM KCl, 20 mM NH4Cl, 10 mM MgSO4, and 0.1% trace elements [43] (pH 7.2) (M9-FA medium) with 10 mM FA added as a carbon source. We replaced FA with PA, DPA and DFA (10 mM each in 0.1 M NaOH) in some cultures. Samples (0.1 g) from a weed rhizosphere (Table S1) were aerobically cultivated at 28 °C in 3 mL of M9-FA medium in 20-mL test tubes for 24 h with agitation at 120 rpm. Thereafter, cultures (30 μL) in 3 mL of fresh M9-FA were passaged at least four times under the same conditions, then broth from the enriched cultures and soil samples was spread over M9-FA agar plates. Isolates were cultured in Luria-Bertani (LB) medium overnight, then 1% of each was inoculated into 500-mL flasks containing 100 mL of M9-FA medium at 28 °C with agitation at 120 rpm. Bacterial growth was measured as total protein in culture pellets using the Protein Assay Dye Reagent (Bio-Rad Laboratories, Hercules, CA, USA) as described by the manufacturer. Briefly, total proteins were stained using the Bradford reagent, and concentrations were determined as Coomassie Blue dye absorption at 595 nm and compared with a standard curve of bovine serum albumin.

Determination of FA, PA, DFA and DPA

Samples were dissolved in 0.1 M NaOH containing 1 M NaCl (pH 12.5), and centrifuged at 20,400×g and 4 °C for 5 min to remove insoluble materials before separation by anion exchange HPLC under the following conditions: column, TSKgel SAX column (6.0 mm × 15.0 cm) (Tosoh Bioscience, Tokyo, Japan); linear gradient of 100 to a 60:40 ratio of aqueous 0.1 M NaOH containing 1 M NaCl (pH 12.5) to 50% acetonitrile in 0.1 M NaOH; column temperature, 30 °C; flow rate, 1.0 mL min− 1. Folic acid, PA, DFA and DPA were detected in eluates as absorption at 254 nm using a 1260 Infinity system equipped with a photodiode array detector (Agilent Technologies, Santa Clara, CA, USA).

Preparation of DFA and DPA

Folic acid (0.88 g) in 0.1-L 50 mM Tris-HCI (pH 7.5) was incubated with 5 μg mL− 1 Variovorax sp. F1 rPDA at 30 °C for 12 h. The rPDA was denatured with 1 M NaOH, then the pH of the reaction was reduced to < 4.0 with 1 M HCl to precipitate DFA with > 99% purity. Deaminopteroic acid was produced by incubating the DFA (~ 0.5 g) with 5 μg mL− 1 rCPG in 0.1-L 50 mM Tris-HCI (pH 7.5) containing 0.2 mM ZnSO4 at 30 °C for 6 h, then purified as described above to > 99%. The DFA and DPA were confirmed by liquid chromatography-mass spectrometry (LCMS). An LCMS 8030 spectrometer (Shimadzu Co., Kyoto, Japan) was equipped with a Purospher® STAR RP-18 endcapped column (particle size 5 μm, Merck KGaA, Darmstadt, Germany) and the flow rate of a 40-min linear gradient from 0 to 40% acetonitrile in 0.05% formic acid was 0.8 mL min− 1. Mass ions were detected in the negative mode under the following conditions: probe voltage, 3.5 kV; detection range, m/z = 10–500 for DFA (precursor m/z 441) and 10–400 (precursor m/z 312) for DPA; column temperature, 40 °C; desolvation line temperature, 250 °C; heat block temperature, 400 °C; nebulizer gas, 3 L min− 1; drying gas, 15 L min− 1.

Protein sequence alignments and phylogenetic analysis

Amino acid sequences obtained from GenBank databases were aligned using CLUSTAL W [44]. A phylogenetic tree was constructed using MEGA X [34] and the neighbor-joining method [33] with 1000 bootstrap resampling replicates. Amino acid sequences with > 44% similarity to CPG in the F1 strain were selected from the Kyoto Encyclopedia of Genes and Genomes (KEGG) [45], and representative sequences were selected from redundant sequence pools derived from strains without species names. Amino acid sequences related to the F1 PDA were selected from Variovorax paradoxus S110 proteins in the KEGG database.

Preparation of recombinant CPG (rCPG) from Variovorax sp. F1

Nucleotide sequences of putative CPG2 genes from various Variovorax bacteria were compared with conserved sequences among the genes using the Basic Local Alignment Search Tool (BLAST), and the (5’→3′) primers: ACCATCATCACCACAGCCAGGATCCGATGCGTCCGAGCATCCAT and TTAAGCATTATGCGGCCGCAAGCTTTCATTTGCCAGCAC. A DNA fragment encoding the CPG of the Variovorax sp. F1 CPG gene was amplified by PCR in a mixture containing these primers, bacterial total DNA, and Ex Taq® DNA Polymerase (Takara, Kyoto, Japan) at 94 °C for 5 min followed by 30 cycles of 98 °C for 10 s, 55 °C for 30 s, 72 °C for 1 min with an additional 7 min at 72 °C for the final cycle. The amplified DNA was fused to pRSFDuet-1 (Merck KGaA) and digested with BamHI and HindIII using NEBuilder HiFi DNA Assembly Master Mix (New England Biolabs, Inc., Ipswich, MA, USA). Escherichia coli BL21 (DE3) (Merck KGaA), harboring the fused plasmid was incubated in LB medium for 12 h, then portions (1 mL) were cultured in 100 mL of fresh LB medium at 37 °C until the OD600 reached 0.5–0.6. Isopropyl β-d-1-thiogalactopyranoside (IPTG; final concentration, 0.1 mM) was added to induce rCPG production, then cultures were shaken for 18 h at 80 rpm and 28 °C.

Cells were harvested by centrifugation at 6,500×g for 10 min at 4 °C, washed twice with 5 mL of 20 mM sodium phosphate (pH 7.4) and 20 mM imidazole, then sonicated on ice for 200 s at 30% output on a 35% duty cycle using a Branson Sonifier® 250 (Branson Ultrasonics Corp., Brookfield, CT, USA). After centrifugation for 10 min at 6,500×g, the supernatant was applied to a 1-mL HisTrap™ FF crude column (GE Healthcare, Chicago, IL, USA), washed with 20 mM sodium phosphate (pH 7.4) containing 0.2 M NaCl and 20 mM imidazole, then rCPG was eluted with 20 mM sodium phosphate (pH 7.4) containing 0.2 M NaCl and 200 mM imidazole. The eluates were concentrated to 1 mL and the solvents were replaced with 20 mM Tris-HCl (pH 7.4) using an Amicon® Ultra-4 Centrifugal Filter Unit Ultracel-30 (Merck KGaA). Proteins were resolved by SDS-PAGE as described by Laemmli [46].

Isolation of Variovorax sp. F1 PDA gene and preparation of rPDA

Orthologs to the predicted deaminase conserved among multi-Variovorax species (WP 068679287) were compared with the predicted genes of V. paradoxus strains and their nucleotide sequences. Conserved sequences were extracted to design the (5’→3′) primers ATGAAGCTCGAGGCCGTCCGC and TACTCGTACCCGTTCGGTTAC respectively corresponding to the 5′ ends of the V. paradoxus genes and the downstream endoribonucleotidase gene. We amplified a DNA fragment encoding the Variovorax sp. F1 PDA gene by PCR using the same primers, total DNA and other conditions used to amplify the CPG genes. The PDA genes were amplified using the rPDA primers ATTTCATATGAAGCTCGAGGCCGTCCGC and CTAACTCGAGTCATGCAATGT TCTCCTGTGA, then amplicons were digested with NdeI and XhoI, cloned into pET28a (Novagen, Madison, WI, USA), and introduced into E. coli BL21 (DE3). Transformants were cultured in LB medium at 30 °C until the OD600 reached 0.5–0.6, after which IPTG (final concentration, 0.2 mM) was added to induce gene expression overnight at 25 °C. The rPDA was purified as described for rCPG.

Enzyme assays of rCPG and rPDA

Enzyme reactions of CPG proceeded in 50 mM Tris-HCl (pH 7.5) containing 0.2 mM ZnSO4 and appropriate amounts of rCPG at 30 °C, then the outcomes were analyzed by HPLC as described above. Substrates in the reaction buffer were incubated at 30 °C for 5 min, followed by reactions with a final concentration of 5 μg mL− 1 rCPG for 5 min. Enzyme reactions of rPDA proceeded in 50 mM Tris-HCl (pH 7.5) with appropriate amounts of rPDA at 30 °C, and were analyzed by HPLC under the same conditions. The enzyme concentration was determined by the Bradford method using Protein Assay Dye Reagent (Bio-Rad Laboratories, Hercules, CA, USA) as described by the manufacturer.

Preparation and analysis of cell-free extract of Variovorax sp. F1

Variovorax sp. F1 was cultured in M9-FA medium at 28 °C for 24 h, harvested by centrifugation at 5100×g for 10 min, and washed with ice-cold 20 mM Tris-HCI (pH 7.2). Cells were resuspended in 5 mL of 50 mM Tris-HCI (pH 7.2) and disrupted as described above. The supernatants were filtered through a 0.45-μm CA syringe filter (Merck KGaA) to obtain cell-free extracts. The preparation (typically, 0.1 mg mL− 1 cell-free extract) in 50 mM Tris-HCl (pH 7.5) containing 0.2 mM ZnSO4 was reacted with purified FA, PA, DFA and AGB that were subsequently quantified by HPLC as described above.

Availability of data and materials

The datasets generated during the current study are available in the GenBank repository (, accession numbers MZ914412, LC718122, and MZ934696).


16S rRNA:

16S ribosomal RNA


N-(4-Aminobenzoyl)-L-glutamic acid


Carboxypeptidase G


Deaminofolic acid


Deoxyribonucleic acid


Deaminopteroic acid


Folic acid


High performance liquid chromatography


Isopropyl β-D-1-thiogalactopyranoside


Kyoto Encyclopedia of Genes and Genomes


Liquid chromatography-mass spectrometry


Pteroic acid


p-aminobenzoic acid


Pterin deaminase


Recombinant CPG


Recombinant pterin deaminase


Sodium dodecyl sulfate-polyacrylamide gel electrophoresis


Tetrahydrofolic acid


  1. McDowell LR. Vitamins in animal and human nutrition. 2nd ed. Ames: Iowa State University Press; 2000.

    Book  Google Scholar 

  2. Shulpekova Y, Nechaev V, Kardasheva S, Sedova A, Kurbatova A, Bueverova E, et al. The concept of folic acid in health and disease. Molecules. 2021;26:3731.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Shenkin A. Basics in clinical nutrition: physiological function and deficiency states of vitamins. Eur J Clin Nutr Metab. 2008;3:e275–80.

    Article  Google Scholar 

  4. Pappenberger G, Hohmann HP. Industrial production of L-ascorbic acid (vitamin C) and D-isoascorbic acid. Adv Biochem Eng Biotechnol. 2014;143:143–88.

    Article  CAS  PubMed  Google Scholar 

  5. Parker GL, Smith LK, Baxendale IR. Development of the industrial synthesis of vitamin a. Tetrahedron. 2016;72:1645–52.

    Article  CAS  Google Scholar 

  6. Woodward RB. The total synthesis of vitamin B12. Pure Appl Chem. 1973;33:145–78.

    Article  CAS  PubMed  Google Scholar 

  7. Abe M, Ito SI, Kimoto M, Hayashi R, Nishimune T. Molecular studies on thiaminase I. Biochim Biophys Acta. 1987;909:213–21.

    Article  CAS  PubMed  Google Scholar 

  8. Kanazawa H, Shigemoto R, Kawasaki Y, Oinuma KI, Nakamura A, Masuo S, et al. Two-component flavin-dependent riboflavin monooxygenase degrades riboflavin in Devosia riboflavina. Appl Environ Microbiol. 2018;200:e00022–18.

    Article  Google Scholar 

  9. Hu C, Zhao S, Li K, Yu H. Microbial degradation of nicotinamide by a strain Alcaligenes sp. P156 Sci Rep. 2019;9:3647.

    Article  CAS  PubMed  Google Scholar 

  10. Yuan B, Yoshikane Y, Yokochi N, Ohnishi K, Yagi T. The nitrogen-fixing symbiotic bacterium Mesorhizobium loti has and expresses the gene encoding pyridoxine 4-oxidase involved in the degradation of vitamin B6. FEMS Microbiol Lett. 2006;234:225–30.

    Article  CAS  Google Scholar 

  11. Yew WS, Gerlt JA. Utilization of L-ascorbate by Escherichia coli K-12: assignments of functions to products of the yjf-sga and yia-sgb operons. J Bacteriol. 2002;184:302–6.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Campos E, de la Riva L, Garces F, Gimenez R, Aguilar J, Baldoma L, et al. The yiaKLX1X2PQRS and ulaABCDEFG gene systems are required for the aerobic utilization of L-ascorbate in Klebsiella pneumoniae strain 13882 with L-ascorbate-6-phosphate as the inducer. J Bacteriol. 2008;190:6615–24.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Brown GM. Biogenesis and metabolism of folic acid. In: Greenberg DM, editor. Metabolic pathways, vol. 4. 3rd ed. NY: Academic Press; 1970. p. 383–410.

    Chapter  Google Scholar 

  14. Fox JT, Stover PJ. Folate-mediated one-carbon metabolism. Vitam Horm. 2008;79:1–44.

    Article  CAS  PubMed  Google Scholar 

  15. Lucock M. Folic acid: nutritional biochemistry, molecular biology, and role in disease processes. Mol Genet Metab. 2000;71:121–38.

    Article  CAS  PubMed  Google Scholar 

  16. MRC Vitamin Study Research Group. Prevention of neural tube defects: results of the medical research council vitamin study. Lancet. 1991;338:131–7.

    Article  Google Scholar 

  17. Serrano-Amatriain C, Ledesma-Amaro R, López-Nicolás R, Ros G, Jiménez A, Revuelta JL. Folic acid production by engineered Ashbya gossypii. Metab Eng. 2016;38:473–82.

    Article  CAS  PubMed  Google Scholar 

  18. Shane B. Folate and vitamin B12, metabolism: overview and interaction with riboflavin, vitamin B6, and polymorphisms. Food Nutr Bull. 2008;29(suppl 1):S5–16.

    Article  PubMed  Google Scholar 

  19. Rossi M, Amaretti A, Raimondi S. Folate production by probiotic bacteria. Nutrients. 2011;3:118–34.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Goldman P, Levy CC. Carboxypeptidase G: purification and properties. Proc Natl Acad Sci U S A. 1967;58:1299–306.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. McCullough JL, Chabner BA, Bertino JR. Purification and properties of carboxypeptidase G1. J Biol Chem. 1971;246:7207–13.

    Article  CAS  PubMed  Google Scholar 

  22. Albrecht AM, Boldizsar E, Hutchison DJ. Carboxypeptidase displaying differential velocity in hydrolysis of methotrexate, 5-methyltetrahydrofolic acid, and leucovorin. J Bacteriol. 1978;134:506–13.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Sherwood RF, Melton RG, Alwan SM, Hughes P. Purification and properties of carboxypeptidase G2 from Pseudomonas sp. strain RS-16. Use of a novel triazine dye affinity method. Eur J Biochem. 1985;148:447–53.

    Article  CAS  PubMed  Google Scholar 

  24. Minton NP, Atkinson T, Sherwood RF. Molecular cloning of the Pseudomonas carboxypeptidase G2 gene and its expression in Escherichia coli and Pseudomonas putida. J Bacteriol. 1983;156:1222–7.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Levenberg B, Hayaishi O. A bacterial pterin deaminase. J Biol Chem. 1959;234:955–61.

    Article  CAS  PubMed  Google Scholar 

  26. McNutt WS. The enzymic deamination and amide cleavage of folic acid. Arch Biochem Biophys. 1963;101:1–6.

    Article  CAS  Google Scholar 

  27. Pratt AG, Crawford EJ, Friedkin M. The hydrolysis of mono-, di-, and triglutamate derivatives of folic acid with bacterial enzymes. J Biol Chem. 1968;243:6367–72.

    Article  CAS  PubMed  Google Scholar 

  28. Rappold H, Bacher A. Bacterial degradation of folic acid. Methods Enzymol. 1980;66:652–6.

    Article  PubMed  Google Scholar 

  29. Takikawa S, Kitayama-Yokokawa C, Tsusue M. Pterin deaminase from bacillus megaterium: purification and properties. J Biochem. 1979;85:785–90.

    Article  CAS  PubMed  Google Scholar 

  30. Fan H, Hitchcock DS, Seidel RD 2nd, Hillerich B, Lin H, Almo SC, et al. Assignment of pterin deaminase activity to an enzyme of unknown function guided by homology modeling and docking. J Am Chem Soc. 2013;135:795–803.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Reed LS, Archer MC. Oxidation of tetrahydrofolic acid by air. J Agric Food Chem. 1980;28:801–5.

    Article  CAS  Google Scholar 

  32. Hussein MJ, Green JM, Nichols BP. Characterization of mutations that allow p-aminobenzoyl-glutamate utilization by Escherichia coli. J Bacteriol. 1998;180:6260–8.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406–25.

    Article  CAS  PubMed  Google Scholar 

  34. Kumar S, Stecher G, Li M, Knyaz C, Tamura K. MEGA X: molecular evolutionary genetics analysis across computing platforms. Mol Biol Evol. 2018;35:1547–9.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. Sadeghian I, Hemmati S. Characterization of a stable form of carboxypeptidase G2 (glucarpidase), a potential biobetter variant, from Acinetobacter sp. 263903-1. Mol Biotechnol. 2021;63:1155–68.

    Article  CAS  PubMed  Google Scholar 

  36. Rashidi FB, AlQhatani AD, Bashraheel SS, Shaabani S, Groves MR, Dömling A, et al. Isolation and molecular characterization of novel glucarpidases: enzymes to improve the antibody directed enzyme pro-drug therapy for cancer treatment. PLoS One. 2018;13:e0196254.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Minton NP, Atkinson T, Bruton CJ, Sherwood RF. The complete nucleotide sequence of the Pseudomonas gene coding for carboxypeptidase G2. Gene. 1984;31:31–8.

    Article  CAS  PubMed  Google Scholar 

  38. Willems A, De Ley J, Gillis M, Kersters K. Comamonadaceae, a new family encompassing the acidovorans rRNA complex, including Variovorax paradoxus gen. Nov., comb. nov., for Alcaligenes paradoxus (Davis 1969). Int J Syst Bacteriol. 1991;41:445–50.

    Article  Google Scholar 

  39. Nguyen TM, Kim J. Description of Variovorax humicola sp. nov., isolated from a forest topsoil. Int J Syst Evol Microbiol. 2016;66:2520–7.

    Article  CAS  PubMed  Google Scholar 

  40. Im WT, Liu QM, Lee KJ, Kim SY, Lee ST, Yi TH. Variovorax ginsengisoli sp. nov., a denitrifying bacterium isolated from soil of a ginseng field. Int J Syst Evol Microbiol. 2010;60:1565–9.

    Article  CAS  PubMed  Google Scholar 

  41. Finkel OM, Salas-González I, Castrillo G, Conway JM, Law TF, Teixeira PJPL, et al. A single bacterial genus maintains root development in a complex microbiome. Nature. 2020;587:103–8.

    Article  CAS  PubMed  Google Scholar 

  42. Sun SL, Yang WL, Fang WW, Zhao YX, Guo L, Dai YJ. The plant growth-promoting rhizobacterium Variovorax boronicumulans CGMCC 4969 regulates the level of indole-3-acetic acid synthesized from indole-3-acetonitrile. Appl Environ Microbiol. 2018;84:e00298–18.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Hutner SH, Provasoli L, Schatz A, Haskins CP. Some approaches to the study of the role of metals in the metabolism of microorganisms. Proc Am Philos Soc. 1950;94:152–70

    CAS  Google Scholar 

  44. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 1994;22:4673–80.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  45. Kanehisa M, Goto S. KEGG: Kyoto encyclopedia of genes and genomes. Nucleic Acids Res. 2000;28:27–30.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  46. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature. 1970;227:680–5.

    Article  CAS  PubMed  Google Scholar 

Download references


We thank Norma Foster (English Express) for critical reading of the manuscript.


This work was supported by JSPS KAKENHI Grant Number JP19H05687 to NT.

Author information

Authors and Affiliations



YY isolated the bacteria. YY, YD, and NM cloned the genes and prepared and analyzed recombinant proteins. YY, YD, NT, SM and NT conducted the investigation and drafted the manuscript. The author(s) read and approved the final manuscript.

Corresponding author

Correspondence to Naoki Takaya.

Ethics declarations

Ethics approval and consent to participate

This study did not require human or animal participation and no data were collected.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1: Fig. S1.

Determination of DFA and DPA. Fig. S2. Dependence of rCPG activity on substrate concentration.

Additional file 2: Supplementary Table S1.

Bacteria isolated from grassland weed rhizosphere.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

You, Y., Doi, Y., Maeda, N. et al. Carboxypeptidase G and pterin deaminase metabolic pathways degrade folic acid in Variovorax sp. F1. BMC Microbiol 22, 225 (2022).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI:


  • Vitamin B9
  • Pterin deaminase
  • Pteroic acid
  • Deaminofolic acid
  • Variovorax