Elucidation of the trigonelline degradation pathway reveals previously undescribed enzymes and metabolites

Abstract

Trigonelline (TG; N-methylnicotinate) is a ubiquitous osmolyte. Although it is known that it can be degraded, the enzymes and metabolites have not been described so far. In this work, we challenged the laboratory model soil-borne, gram-negative bacterium Acinetobacter baylyi ADP1 (ADP1) for its ability to grow on TG and we identified a cluster of catabolic, transporter, and regulatory genes. We dissected the pathway to the level of enzymes and metabolites, and proceeded to in vitro reconstruction of the complete pathway by six purified proteins. The four enzymatic steps that lead from TG to methylamine and succinate are described, and the structures of previously undescribed metabolites are provided. Unlike many aromatic compounds that undergo hydroxylation prior to ring cleavage, the first step of TG catabolism proceeds through direct cleavage of the C5-C6 bound, catalyzed by a flavin-dependent, two-component oxygenase, which yields (Z)-2-((N-methylformamido)methylene)-5-hydroxy-butyrolactone (MFMB). MFMB is then oxidized into (E)-2-((N-methylformamido) methylene) succinate (MFMS), which is split up by a hydrolase into carbon dioxide, methylamine, formic acid, and succinate semialdehyde (SSA). SSA eventually fuels up the TCA by means of an SSA dehydrogenase, assisted by a Conserved Hypothetical Protein. The cluster is conserved across marine, soil, and plant-associated bacteria. This emphasizes the role of TG as a ubiquitous nutrient for which an efficient microbial catabolic toolbox is available.

Keywords: LC/MS-Orbitrap; N-heterocycle degradation; bacterial metabolism; functional genomics; trigonelline.

Fig. 1.

Phenotyping of the genome-scale ADP1 mutant collection on TG and succinate as the carbon sources. The mineral medium was supplemented with 10 mM TG or succinate as the carbon source and 18 mM ammonium as the nitrogen source. The medium was inoculated from an overnight culture pregrown on ammonium and succinate. Cell growth from duplicate independent cultures was measured by OD₆₀₀ after 24 h. The squared area represents mutants growing poorly on TG, while presenting significant growth on succinate plus ammonium. Annotation of genes is provided in Table 1.

Fig. 2.

TG degradation gene cluster in ADP1 and S. meliloti 1021. Physical colocalization of genetic loci was observed through the MicroScope platform (65). Transporter genes are all colored gray, irrespective of the transporter family. The same applies to predicted transcriptional regulators (yellow). Uncolored gene symbols are used for genes apparently unrelated to TG degradation. ACIAD, Acinetobacter baylyi ADP1; Sma, Sinorhizobium meliloti 1021.

Fig. 3.

In vitro reconstruction of the TG degradation pathway. The initial substrate concentrations were 500 μM TG, 1.5 mM NADH, and 25 μM FMN in a final volume of 100 μL. Two micrograms of each protein was added. (A) Sample of the reaction mixture in the absence of enzyme. (B) Sample analyzed 30 min after addition of TgnA and TgnB. (C) Sample analyzed 30 min after addition of TgnAB and TgnC. (D) Sample analyzed 30 min after addition of TgnAB, TgnC, and TgnD. (E) Sample analyzed 30 min after addition of TgnAB, TgnC, TgnD, and TgnE. (F) Sample analyzed 30 min after addition of TgnAB, TgnC, TgnD, TgnE, and TgnF. Samples were analyzed by LC/MS in the negative ionization mode. The extracted ion chromatograms (EICs) correspond to the quasimolecular ions ([M-H]⁻) of TG [m/z = 136.040 atomic mass units (amu)], catabolite A (m/z = 170.045 amu), catabolite B (m/z = 186.040 amu), SSA (m/z = 101.024 amu), and succinate (m/z = 117.019 amu) at 5 ppm accuracy.

Fig. 4.

Structural elucidation of MFMB and MFMS. ¹H-¹H ROESY experiments (600 MHz) at 298 K for MFMB [conformers (+/−)-A and (+/−)-A′ in equilibrium] (A) and MFMS [conformers (+/−)-B and (+/−)-B′ in equilibrium] (B) in D₂O. Most significant chemical exchange correlations and NOE correlations are pointed to on both ROESY spectra and the respective catabolite structures (E and F) with black and red arrows. Chemical exchange correlations demonstrate conformational equilibria (e.g., correlation between H_a and H′_a indicates that H_a becomes H′_a during the NMR experiment time scale). The intensity of NOE correlations is related to the spatial distance between protons belonging to the same molecule, and it allows the determination of the C = C bond configuration in MFMB and MFMS catabolites, as well as the 3D conformation of all catabolite isomers. For clarity, several signal intensity levels were combined together: (i) medium level for the general ROESY frames, (ii) high level in green dashed boxes, and (iii) low level in pink dashed box. ROESY original spectra for each intensity level are provided in Dataset S1 A–E. (C) N-methyl region from ¹H NMR spectra (600 MHz) of MFMB at different temperatures in CD₃CN illustrates the dynamic equilibrium between rotamers. Rotation around the N-C_a bond interconverts the position of the formyl group, and hence modifies the chemical environment of the N-methyl protons H_g and H′_g (E, in green). At 294 K, when this interconversion is too slow compared with the ¹H NMR acquisition time scale, the ¹H spectrum of the N-methyl groups consists of two sharp peaks arising from H_g and H′_g. As the temperature is increased and the rate of interconversion becomes faster, the peaks broaden and tend to merge. The same phenomenon occurs for formyl (H_a and H′_a) and vinyl (H_b and H′_b) protons as well (Fig. S5A). (D) Slow proton/deuterium exchanges within MFMB rotamers are evidenced by the disappearance of the H_e + H′_e and H_g + H′_g signals of MFMB in CDCN₃ (i) upon addition of D₂O to the CDCN₃ sample (ii) and subsequent redissolution in CDCN₃ of the lyophilizate (iii). These exchanges indicate that H_e + H′_e and H_g + H′_g are enolizable protons in the α-position of the aldehyde group of open isomers (even at a neutral pH), which are in equilibrium with closed structures. (E and F) Structures of MFMB and MFMS conformers, respectively, as inferred from their NMR data.

Fig. 5.

Steady-state kinetics of NADH oxidation by TgnA in the presence of TG. (A and B) Rates of NADH oxidation versus NADH concentration in the presence of saturating concentrations of TG and FMN. (C and D) Rates of NADH oxidation versus TG concentration in the presence of saturating concentrations of NADH and FMN. (E and F) Rates of NADH oxidation versus FAD concentration in the presence of saturating concentrations of TG and NADH. (G and H) Rates of NADH oxidation versus FMN concentration in the presence of saturating concentrations of TG and NADH. v, the initial rate of the reaction, is expressed in moles of NADH oxidized per second and per mole of enzyme. Curves were drawn using Sigma-Plot software. The Michaelis–Menten equation v = (V_max S)/(K_m + S) applies for plot A, and the Hill equation v = (V_max Sⁿ)/(S₅₀ⁿ + Sⁿ) applies for plots C, E, and G. S₅₀ is the substrate concentration showing half-maximal velocity, n is the Hill coefficient, and V_max is the maximal velocity. (B, D, F, and H) Eadie–Hofstee representation (v versus v/S) of the kinetics.

Fig. 6.

TG degradation pathway in ADP1. TgnA and TgnB, two-component TG oxygenase; TgnC, MFMB dehydrogenase; TgnD, MFMS hydrolase; TgnE, SSA dehydrogenase; TgnF, SSA dehydrogenase-stimulating protein. Data from NMR analysis for MFMB and MFMS are indicated in gray boxes. A and A′ represent the two closed-form conformers of MFMB detected by NMR. B and B′ represent the two conformers of MFMS. TgnF is indicated in brackets since it is not known whether it is an enzyme.

Fig. 7.

Illustration of the taxonomic diversity of homologous predicted TG-degradation gene clusters. Homologous gene clusters in bacterial genomes were retrieved using MicroScope. Shown are gene clusters that contain homologous genes to ADP1 (A) for at least four of the six catalytic proteins of the TG-degradation pathway in γ-, α- and β-Proteobacteria (B–D, respectively) and in Actinobacteria (E). Taxonomic orders are indicated in brackets. Candidate genes for transporters and transcriptional regulators were frequently found to be conserved within these clusters. Genes indicated in white are not predicted to be related to TG degradation.