Genetics of the glutamate-mediated methylamine utilization pathway in the facultative methylotrophic beta-proteobacterium Methyloversatilis universalis FAM5


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The ability of some microbial species to oxidize monomethylamine via glutamate-mediated pathways was proposed in the 1960s; however, genetic determinants of the pathways have never been described. In the present study we describe a gene cluster essential for operation of the N-methylglutamate pathway in the methylotrophic beta-proteobacterium Methyloversatilis universalis FAM5. Four major polypeptides from protein fractions displaying high activities of N-methylglutamate synthetase, N-methylglutamate dehydrogenase and γ-glutamylmethylamide synthetase were selected for mass spectrometry-based identification. The activities of enzymes were associated with the presence of peptides identified as ferredoxin-dependent glutamate synthase (GltB2), large subunit of putative heterotetrameric sarcosine oxidase (SoxA) and glutamine synthetase type III (GSIII) respectively. A gene cluster (8.3 kb) harbouring gltB2, soxA and gsIII-like genes was amplified from M. universalis FAM5, sequenced and assembled. Two partial and six complete open reading frames arranged in the order soxBDAG-gsIII-gltB132 were identified and subjected to mutational analysis, functional and metabolic profiling. We demonstrated that gltB-like and sox-like genes play a key role in methylamine utilization and encode N-methylglutamate synthetase and N-methylglutamate dehydrogenase respectively. Metabolic, enzymatic and mutational analyses showed that the gsIII-like gene encodes γ-glutamylmethylamide synthetase; however, this enzyme is not essential for oxidation of methylamine.


Methylated amines, the reduced organic forms of nitrogen ([CH3]nNHx), are formed in nature as by-products of decomposition of proteins (Lee, 1988; Neff et al., 2002). These compounds are essential intermediates in the metabolism of caffeine/theanine/theobromine producing plants (Konishi et al., 1972; Yamamoto et al., 2007), and serve as central components in the formation/breakdown of compatible solutes, such as glycine betaine, proline betaine and trimethylamine oxide in marine animals, halophilic algae and bacteria (Oren, 1990; Brad et al., 2002). In the human body methylamine plays a significant part in central nervous system disturbances and has a role in general toxicity caused by oxidative stress (Asatoor and Kerr, 1961; Yu and Dyck, 1998; Mitchell and Zhang, 2001). It is the simplest aliphatic amine found in human urine, with the average daily output of methylamine being 11.00 ± 8.17 mg (Mitchell and Zhang, 2001). Methylated amines are good solvents and common precursors for organic synthesis, thus they are pervasively used in the chemical, pharmaceutical, rubber, plastic, dye-stuff, textile, cosmetics and metal industries (Stellman, 1998). On average, in pristine ecosystems methylated amines are present at low concentrations, likely due to rapid microbial degradation (Oremland et al., 1982; King et al., 1983). This paradigm shifts when a significant source (natural or anthropogenic) of methylated compounds is present (Neff et al., 2002). Major global sources of methylated amines emitted into the environment are oceans (0.6 Tg of N) (Lee, 1988), animal husbandry operations (0.15 Tg of N) (Schade and Crutzen, 1995) and biomass-burning (0.06 Tg of N) per year (Neff et al., 2002). Methylated amines could also accumulate in sediments, groundwaters, and organic-rich soils as a result of hydrolysis of N-methyl carbamate pesticides, which are used worldwide (Kiene and Capone, 1986). However, relatively little is known about the environmental fate of these compounds. Thus, the understanding of global cycling of methylated amines has major environmental implications (Neff et al., 2002).

The microbial oxidation of methylated amines is an important component of environmental cycling of C and N as well as an essential step in preventing formation of methane, the second most important greenhouse gas following carbon dioxide. The ability to use methylamine as a sole source of carbon, nitrogen and energy is widely distributed in microorganisms. The first efforts towards an understanding of the biochemical nature of methylamine oxidation in bacteria were undertaken in the 1960s, demonstrating the presence of two major routes for methylamine oxidation (Fig. 1): (i) direct oxidation to formaldehyde and ammonia carried out by a periplasmic quinoprotein methylamine dehydrogenase (MADH) in proteobacteria (Eady and Large, 1968; Chistoserdov et al., 1994; Van Der Palen et al., 1995; Gak et al., 1997) and by a methylamine oxidase (MAO) in Gram-positive bacteria (Zhang et al., 1993; Langley et al., 2006); and (ii) conversion of methylamine to N-methylglutamate (NMG) (Shaw et al., 1966; Hersh et al., 1971; Pollock and Hersh, 1971; Bamforth and Large, 1977a), N-methylalanine (Lin and Wagner, 1975) or γ-glutamylmethylamide (Kung and Wagner, 1969; Kimura et al., 1992), followed by oxidation of the respective methylated amino acids to formaldehyde and regeneration of the corresponding amino acid. To this day, only genetics of the enzymes for direct methylamine oxidation, MADH (Chistoserdov et al., 1994; Van Der Palen et al., 1995; Gak et al., 1997) and MAO (Zhang et al., 1993) have been studied.

Figure 1.

A. Methylamine utilization pathways in proteobacteria. In grey – methylamine utilization pathways/reactions that were not detected in Methyloversatilis universalis FAM5. 1, Methylamine dehydrogenase (, amine dehydrogenase); 2, Methylamine oxidase (EC, primary-amine oxidase); 3, N-methylglutamate synthase (EC, Methylamine-glutamate N-methyltransferase (NMGS); 4, N-methylglutamate dehydrogenase (EC, N-methyl-l-glutamate:oxidoreductase (NMGDH); 5, γ-glutamylmethylamide synthetase (EC, l-glutamate:methylamine ligase (ADP-forming) (GMAS); 6, γ-glutamylmethylamide-dissimilating enzyme system. *Based on in vitro assays, end-products of oxidative demethylation of N-methylglutamate were shown to be formaldehyde and glutamate (Hersh et al., 1971; 1972; Bamforth and Large, 1977a). We demonstrate here that in M. universalis FAM5 in the presence of H4folate, N-methylglutamate is converted to glutamate and 5,10-methylene-H4folate.
B. Organization of gene clusters for glutamate-mediated pathway in proteobacteria. Homologous genes have the same colour. Designations: luxR and trc, putative transcriptional regulators purU, formyltetrahydrofolate deformylase; folD, methylenetetrahydrofolate dehydrogenase/cyclohydrolase; soxBCAG, putative sarcosine oxidase operon; glIII, putative glutamine synthatase III; gltB132, putative glutamate synthase.

The NMG pathway for methylamine utilization was first proposed in 1966 to explain the growth of Pseudomonas spp. (now Aminobacter sp.) on methylamine while displaying no MADH activity (Shaw et al., 1966). Two enzymes, named NMG synthetase (EC Methylamine-glutamate N-methyltransferase, NMGS) and NMG dehydrogenase [EC N-methyl-L-glutamate:(acceptor) oxidoreductase, NMGDH], were found to be involved, by first converting methylamine into NMG (Pollock and Hersh, 1971; 1973) and then cleaving it to formaldehyde and glutamate. Enzymatic systems involved in NMG synthesis/oxidation were eventually purified and characterized (Hersh et al., 1971; 1972; Pollock and Hersh, 1971; 1973; Bamforth and Large, 1977a,b; Boulton et al., 1980). Two types of NMG oxidizing enzymes were described: (i) particle-bound NMGDH, that was active with 2,6-dichlorophenol-indophenol (DCPIP) as electron acceptor (Hersh et al., 1971; Bamforth and Large, 1977a); and (ii) soluble NAD-activated enzyme (Loginova et al., 1974). A soluble flavohaemoprotein active with 2,6-dichlorophenol-indophenol was also reported (Boulton et al., 1980). Several mutants of Pseudomonas aminovorans deficient in the ability to grow on methylamine were generated (Bamforth and O'Connor, 1979). These mutants showed multiple lesions in C1-oxidation and assimilatory pathways, suggesting that genetic systems for NMG pathway and serine cycle are interconnected via regulatory machinery, or structurally, as an operon on a plasmid.

Activities of the NMG pathway enzymes have been detected in a variety of methylotrophic proteobacteria and were shown to be induced during growth on methylated amines (Kung and Wagner, 1970; Hersh et al., 1971; Doronina et al., 2004; Miller et al., 2005; Trotsenko et al., 2007). While some methylotrophs display relatively high activities of the NMG pathway enzymes even in the presence of functional MADH (Doronina et al., 2004), for organisms lacking MADH the NMG pathway has been proposed as a major route for methylamine oxidation (Shaw et al., 1966; Kung and Wagner, 1970; Hersh et al., 1971). However, genes encoding the NMG pathway enzymes have never been identified. Here we present the first description of genetic determinants for key enzymes of the NMG pathway and mutant evidence for their roles in bacterial utilization of methylated amines.


N-methylglutamate and γ-glutamylmethylamide are major intermediates of methylamine metabolism in M. universalis FAM5

The previous data suggested that M. universalis FAM5 lacks the activity of MADH, and may possess an alternative pathway for methylamine utilization (Kalyuzhnaya et al., 2006). In order to identify the mechanism by which M. universalis grows on methylamine, we performed LC-MS/MS analysis of methanol and methylamine grown cells. Comparative targeted metabolic profiling revealed marked differences in the amino acid composition (Table 1). Cells grown on methylamine as a source of carbon/energy accumulated significant amounts of NMG (2.67 ± 0.85 µmol g−1 of dry weight) and γ-glutamylmethylamide (4.23 ± 1.38 µmol g−1 of dry weight). These compounds were associated with growth on methylamine and were not detected in cells grown on methanol. Methylamine grown cells also displayed elevated concentrations of glutamate (1.17 ± µmol g−1 of dry weight), while intracellular concentrations of glutamine and alanine were minimal (Table 1). These data suggest that M. universalis FAM5 employs glutamate-mediated pathways for methylamine utilization.

Table 1.  Intracellular concentrations of intermediates of amino-acid mediated pathways in M. universalis FAM5.
MetaboliteAmount (µmol/mg of dry weight)
  1. Note: Cells were grown on methanol (met); methylamine (ma); or grown on methanol and induced with methylamine for 12–16 h (met→ma).

  2. ND, metabolite was not detected.

Glutamate1.17 ± 0.300.18 ± 0.040.37 ± 0.08
N-methylglutamate2.67 ± 0.85ND0.35 ± 0.07
Glutamine0.14 ± 0.030.10 ± 0.010.30 ± 0.05
γ-glutamylmethylamide4.23 ± 1.38ND14.99 ± 4.04
Alanine0.35 ± 0.050.63 ± 0.100.37 ± 0.08

N-methylglutamate pathway enzymes and γ-glutamylmethylamide synthetase are induced during growth on methylated amines

In order to identify enzymes involved in methylamine utilization in M. universalis FAM5, we carried out initial experiments with cell extracts of M. universalis FAM5 grown on methylamine and methanol. We tested previously described methods for detection of N-methylated amino acid oxidases/dehydrogenases (NMGDH) activity (Hersh et al., 1971; Bamforth and Large, 1977a; Wagner et al., 1997), and optimized the assay using protein extract (or the protein fraction with the highest enzyme activities, 20–30% ammonium sulphate saturation, see below) from M. universalis FAM5 cells grown with methylamine as a sole carbon and energy source. We were not able to detect any activity with DCPIP as electron acceptor, as described by Bamforth and Large (1977a) for NMGDH from Pseudomonas aminovorans. The assay used for detection of the NMG oxidizing enzyme in M. universalis FAM5 was based on measurements of the rate of formaldehyde production from NMG, using the standard Nash reagent. The assay is similar to the enzymatic assays widely used for detection of N-methyl amino acid oxidizing enzymes (Wagner et al., 1997). The optimum pH for the enzymatic reaction is 7.6. The rate of formaldehyde production was 35% higher when sodium phosphate buffer was used as opposed to the Tris-HCl buffer (data not shown). The addition of NAD to the reaction mixture increased the formaldehyde production rate from 175 to 288 nmol min−1 mg−1 (∼64% increase). However, NAD was not reduced during this reaction. Similar results were obtained with protein fractions partially purified by ammonium sulphate precipitation and gel filtration. Addition of potassium cyanide to the reaction mixture had no effect on the reaction kinetics. Thus, these data indicate that NAD may be essential for the stability of the NMG converting enzyme. This resembles the properties of heterotetrameric sarcosine oxidases that also require NAD+ for stability/folding of the large subunit (Eschenbrenner et al., 2001). The optimized assay was used for enzymatic profiling of M. universalis FAM5 grown on methanol and methylamine. Activity of the NMGDH was almost 10-fold higher in methylamine grown cultures versus methanol grown cultures (Table 2). Activity of γ-glutamylmethylamide synthetase (EC; also known as L-glutamate:methylamine ligase (ADP-forming), GMAS), an alternative system described for methylamine utilization, was measured by monitoring γ-glutamylhydroxamate formation (O'Neal and Joy, 1974; Yamamoto et al., 2007). A significant increase in the rate of γ-glutamylhydroxamate formation was observed in methylamine-grown cells when compared with methanol growth (Table 2). These data indicated a potential role of a glutamine synthetase (GS)/GMAS-like enzyme in methylamine utilization. The activity was strongly dependent on the presence of Mg2+ ions. No activity was detected when Mg2+ ions were substituted with Mn2+. Activities of γ-glutamylmethylamide dissimilating enzyme (Kimura et al., 1995) or alanine dehydrogenase (Lin and Wagner, 1975) were not detected.

Table 2.  Activities in nmol min−1 mg protein−1 of the key enzymes of the N-methylglutamate pathway in wild-type and mutant strains of Methyloversatilis universalis FAM5.
  1. Enzymatic measurements were performed on protein extract from cell grown on methanol (met); methylamine (ma); or grown on methanol and induced with methylamine for 12–16 h (met→ma). Activities measured in methanol grown cells are given in parentheses.

  2. In bold: enzyme activity was measured in protein fraction from methylamine grown cells recovered at 0–20% ( *), 20–30% ( **), or 30–40% ( ***) ammonium sulfate saturation.

  3. –, not tested. NMGS, N-methylglutamate synthase; NMGDH, N-methylglutamate dehydrogenase; GMAS, γ-glutamylmethylamide synthetase; SOX, sarcosine oxidase; H4F, tetahydrofolate.

NMGS22.3 ± 3.2111.3 ± 5.270.5 ± 4.655.7 ± 4.611.8 ± 2.311.3 ± 3.4
NMGDH19.3 ± 2.7288.0 ± 5.4131.34 ± 27.2< 3< 377.4 ± 12.283.6 ± 3.174.1 ± 24.0
24.2 ± 5.3*
340.3 ± 67.8**
87.4 ± 17.2***
NMGDH+H4F 26.2 ± 14.9      
GMAS21.7 ± 2.9205.2 ± 12.977.3 ± 12.770.1 ± 16.355.9 ± 2.220.4 ± 6.875.0 ± 14.168.4 ± 19.8
< 12*
27.2 ± 3.2**
190.8 ± 15.3***
SOX 9.3 ± 4.6

In a variety of proteobacteria the initial catabolism of methylated amines (such as trimethylamine and dimethylamine) usually results in the production of methylamine as one of the end-products of demethylation reactions (Eady et al., 1971; Meiberg and Harder, 1978). During exponential growth of M. universalis FAM5 on trimethylamine, the following enzymes were induced: trimethylamine and dimethylamine mono-oxygenases, NMGDH and GMAS (Table S2). No activities of primary amine or N-methylalanine dehydrogenases were detected. Thus, the metabolism of methylamine, a product of the dimethylamine conversion, is likely to proceed via the NMG pathway.

Identification of proteins induced during growth on methylamine

In order to identify proteins in M. universalis FAM5 that carry out the activities described above, cell extracts of a methylamine-grown culture were fractioned by ammonium sulphate treatment (0–20%, 20–30%, 30–40% saturation), and each fraction was compared with similarly fractionated extracts from methanol-grown cells by SDS-PAGE. Activities of NMGDH and GS/GMAS in corresponding protein fractions were assayed as described above. The highest rate of NMGDH was observed in the protein fraction recovered at 20–30% saturation (Table 2). Three major proteins, approximately 95, 46 and 20 kDa (band 1, 2 and 3 respectively) were overproduced in methylamine-grown cells compared with methanol (Fig. 2). The highest activity of γ-glutamylhydroxamate formation was detected in the protein fraction recovered at 30–40% ammonium sulphate saturation. One major differentially produced polypeptide (about 45 kDa, labelled as band 2a) was observed in this protein fraction. Polypeptide bands were cut out from gels and subjected to mass spectrometric analysis. The resultant MS/MS peptide spectra were matched against two data sets: a non-redundant data set that includes data from sequenced microbial genomes (NR), and a data set consisting of polypeptide sequences derived from a Lake Washington sediment metagenome (LWS). Our previous analysis of the metagenomic database revealed a relatively high coverage for a number of methylotrophs, including a strain of M. universalis very closely related to FAM5 (> 99% at the 16S rRNA gene level) (Kalyuzhnaya et al., 2008). Therefore, it was reasonable to expect that at least partial sequences for the proteins of interest are present in the metagenomic data set. Indeed, the search against the LWS metagenomic data provided a higher number of unique peptides with higher spectral counts for all three tested protein bands than the NR database (Table 3). A search of the spectrum of the 95 kDa polypeptide (band 1) against the LWS data set resulted in two major hits: a glycine cleavage system T/aminomethyltransferase (SoxA/GcsT) and a hypothetical protein of unknown function (Table 3). An aminomethyltransferase-like peptide was also identified as one of the most probable matches for band 1, when the NR data set was used for the search. Band 2 contained the beta subunit of ATP synthase and putative ferredoxin-dependent glutamate synthase (GltB2) (data not shown). The spectral count for band 2a was divided between GltB2 (5 unique peptides and 37 spectral count) and glutamine synthetase (GSIII) with 2 unique peptides and 49 spectral count (Table 3). The 20 kDa polypeptide (band 3) was matched to a homologue of formaldehyde activating enzyme (Fae 2) in both analyses.

Figure 2.

SDS-PAGE (4–25%) analysis of crude extract and protein fractions after ammonium sulphate precipitation (20%, 30% and 40%) of cell extract obtained from Methyloversatilis universalis FAM5 grown on methanol (m) or methylamine (ma). Arrows indicate bands representing proteins of significantly different abundance.

Table 3.  Detected peptides statistics and identification of proteins from M. universalis FAM5 differentially expressed during growth on methylamine.
GelNCBI non-redundant data setLMS metagenomic data seta
bandDescription (protein ID, predicted function, strain)Unique peptidesbSpectral countscDescription (protein ID, predicted function)Unique peptidesbSpectral countsc
  • a. 

    M. Hackett and L. Chistoserdova, unpublished data sets.

  • b. 

    Only peptides with spectral counts higher then 10 are listed.

  • c. Spectral counting is a frequency measurement based on the number of redundant peptide sequences observed for a given protein. In our usage of the term only fragmentation mass spectra that map uniquely to a given protein are counted. Redundancy here refers to counting multiple observations of the same peptide in different HPLC fractions and at different elution times. It has been shown that spectral counting correlates, with certain caveats, with relative protein abundance (Liu et al., 2004).

11. YP160847.1, Alpha-ketoglutarate decarboxylase, Azoarcus sp. EbN1s1141. 2006973257, Glycine cleavage system, T protein/aminomethyltransferase468
2. YP326317.1, Hypothetical protein, Natronomonas pharaonis DSM 21601142. 2006864371, Hypothetical protein111
3. YP405620.1, Aspartate kinase III, Shigella dysenteriae Sd197112
4. YP544565.1, T protein/ aminomethyltransferase, Methylobacillus flagellatus KT111
2a1. ZP03267511.1, Ferredoxin dependent glutamate synthase, Burkholderia sp H1603341. 2006973258, Glutamine synthetase249
2. ATP synthase F1, Alpha subunit, Haemophilus somnus 23364132. 2006873269, Glutamate synthase537
31. NP868826.1, Homologue of formaldehyde-activating enzyme, fae2 Rhodopirellula baltica SH 12411. 2006964245, Homologue of formaldehyde-activating enzyme, fae2463
2. 2006813921, Homologue of formaldehyde-activating enzyme, fae2673

Identification and sequencing of soxBDAG-gsIII-gltB132 from M. universalis FAM5

The genomic characterization of the newly identified proteins revealed that genes encoding three out of four detected peptides (SoxA/GcsT, GltB3 and GSIII) are parts of a single conserved soxBDAG-gsIII-gltB13gltS cluster on a plasmid in Burkholderia phymatum STM815, a nitrogen-fixing symbiotic bacterium (Elliott et al., 2007), and on the chromosome of Methylobacillus flagellatus KT, a well-characterized C1-oxidizer (Chistoserdova et al., 2007) (Fig. 1B). The sequence information from B. phymatum STM815 and M. flagellatus KT was used to design degenerate primers sets for amplification of the homologous genes from M. universalis FAM5. Five genes and the corresponding intergenic regions were amplified, sequenced and assembled in an 8.2 kb gene cluster. Two partial and six complete open reading frames were identified. blast searches revealed high homology (71–89% amino acid sequence identity) between putative proteins from M. universalis FAM5 and B. phymatum SMT815 with the exception of a small subunit of a putative sarcosine oxidase (SoxG), sharing only 42% of amino acid sequence identity (Table 5). Corresponding genes were arranged in the following order soxBDAG-gsIII-gltB132, the same organization as the gene cluster in B. phymatum SMT815 (Fig. 1B). Polypeptides encoded by putative soxA and gsIII genes from M. universalis FAM5 displayed a high degree of sequence identity (98–99%) to polypeptides that were positively matched in the MS/MS analysis (Table 3), thus confirming the correct identification of the genes that encode proteins induced during growth on methylamine.

Table 5.  Gene description and assigned physiological function.
Best NCBI hitPredicted functionIdentitiesGeneAssigned gene functionPredicted protein MW, (kDa)Previously described enzymes MW, (kDa)
  • a. 

    No prediction can be made as only partial orf sequence obtained.

N-methylglutamate pathway: step I
 gltB1Glutamine amidotransferase class-II, Burkholderia phymatum STM81579%mgsAN-methyl glutamate synthase, subunit A33.4730–35 kDa (Pollock and Hersh, 1971)
 gltB3Glutamate synthase alpha subunit, Leptothrix cholodnii SP-675%mgsBN-methyl glutamate synthase, subunit B23.91
N-methyl glutamate synthase, subunit B
 gltFFerredoxin-dependent glutamate synthase, Burkholderia phymatum STM81576%mgsCN-methyl glutamate synthase, large subunit Ca
N-methylglutamate pathway: step II
 soxBFAD dependent oxidoreductase, Burkholderia phymatum STM81589%mgdAN-methyl glutamate dehydrogenase/oxidoreductase, subunit Aa108–130 kDa Homotetrameric enzyme (Bamforth and Large, 1977b; Boulton et al., 1980)
 soxDSarcosine oxidase delta subunit heterotetrameric, Burkholderia phymatum STM81575%mgdBN-methyl glutamate dehydrogenase/oxidoreductase, subunit B9.93
 soxA2006973257, Glycine cleavage system T protein/aminomethyl transferase, LWS metagenome98%mgdCN-methyl glutamate dehydrogenase/oxidoreductase, large subunit C100.22
GcvT/SoxA, Glycine cleavage system T-protein/aminomethyltransferase, Burkholderia phymatum STM8165–77%   
 soxGHypothetical protein Bphy_5917, Burkholderia phymatum STM81542%mgdDN-methyl glutamate dehydrogenase/oxidoreductase, subunit D23.53
Synthesis of gamma-glutamylmethylamide
 gltIII2006973258, Glutamine synthetase, type III, LWS metagenome99%gmsGamma-glutamylmethylamide synthetase48.8255 kDa Homo-octomer (Kimura et al., 1992)
Glutamine synthetase, type III, Burkholderia phymatum STM81571%   
Gamma-glutamylmethylamide Methylovorus mays69%   51 kDa (Yamamoto et al., 2007)

The soxBDAG subcluster is essential for N-methylglutamate oxidation

The first four open reading frames are predicted to encode flavocoenzymes that are usually annotated as heterotetrameric sarcosine oxidase (TSOX). However, the specific physiological function of the majority of the TSOX-like proteins in proteobacteria is mainly unknown. The SoxB and SoxA-like proteins from M. universalis FAM5 are distantly related to characterized small and large catalytic subunits of TSOX from Corynebacterium sp. P-1 (42% and 28% amino acid sequence similarity respectively) (Chlumsky et al., 1995). Phylogenetic analysis of the SoxB-like proteins and other related proteins in the FAD-dependent oxidoreductase family involved in the oxidation of methylated amino acids, such as monomeric sarcosine oxidase (MSOX), small catalytic subunit of TSOX, N-methyltryptophan oxidase (MTOX) is shown in Fig. S1. The putative SoxB from M. universalis FAM5 and B. phymatum STM815 are separated from known enzymes and cluster together with similar proteins from other obligate methylotrophic bacteria such as Methylophaga and M. flagellatus KT. Enzymatic tests with cell extracts of M. universalis FAM5 grown on methylamine showed a very low activity of sarcosine oxidation (SOX, Table 2), indicating that TSOX-like proteins detected here might play a different biochemical role. Thus, it was reasonable to suggest that SoxBDAG from M. universalis FAM5 may be involved in oxidation of NMG.

To better understand the role of the newly identified genes in C1-utilization, mutant strains of M. universalis FAM5 defective in soxB and soxA were generated as described in Experimental procedures. The resulting strains were named NMGP1 (soxB:kan) and NMGP2 (ΔsoxA:kan) (Table S1). Growth rates of the mutants in mineral medium supplemented with methanol were similar to the wild-type, but both NMGP1 and NMGP2 strains failed to grow on mono-, di- or trimethylamines.

As these mutant strains were not able to grow on methylamine, metabolic profiling and NMGDH activity measurements were performed in cultures grown on methanol and induced with methylamine for 12–16 h. Control experiments with the wild-type showed that this length of time was sufficient for induction of NMGDH activity to a level of fivefold compared with methanol cultures and accumulation of intermediates of methylamine metabolism. NMGP1 and NMGP2 mutants did not exhibit any detectable activity of the formaldehyde production, indicating that the genes are essential for NMG oxidation (Table 2). Furthermore, intracellular concentrations of NMG in the mutant strains considerably exceeded the wild-type background (1.6-fold for NMGP1 and > 44-fold for NMGP2, Table 4). Overall, the data indicated that TSOX-like proteins are essential for oxidation of NMG. All four genes of the cluster were assigned to NMGDH function and named mgdABCD (Table 5).

Table 4.  Comparison of intracellular intermediates of amino-acid mediated pathways in M. universalis FAM5 mutant strains.
  1. Note: mutant strains and wild-type were grown on methanol and induced with methylamine for 12 h. The data represent ratios of peak area of corresponding metabolite in mutant strain versus wild-type. TR, trace; ND, not detected.

Glutamate0.064 ± 0.0160.25 ± 0.0740.37 ± 0.0850.25 ± 0.092
N-methylglutamate1.681 ± 0.38644.67 ± 14.5750.996 ± 0.2760.21 ± 0.054
Glutamine0.100 ± 0.0232.330 ± 0.530.457 ± 0.1020.966 ± 0.197
γ GlutamylmethylamideTR0.043 ± 0.016ND0.196 ± 0.078

The synthesis of 5,10-methylenetetrahydrofolate (CH2-H4folate) as a final product of methyl group oxidation is a most distinguishable characteristic of heteroterameric sarcosine oxidases (Chlumsky et al., 1995; Wagner and Jorns, 1997). Since an aminomethyltransferase folate-binding domain was also detected near the C-terminus of the large subunit of NMGDH, we tested if this enzyme could use H4folate as a substrate and produce CH2-H4folate as an end-product of NMG oxidation. The ability of NMGDH to utilize H4folate was investigated by measuring the rate of formaldehyde production in the presence or absence of H4folate, as carried out previously for TSOX and MSOX enzymes (Wagner and Jorns, 1997; 2000). We demonstrated that addition of 10 µM H4folate to the reaction caused a significant decrease in the rate of formaldehyde production (Table 2). Moreover, the amount of H4folate required for inhibition of formaldehyde production by NMGDH was fivefold lower than the concentration of H4folate used in a similar assay with TSOX system (Wagner and Jorns, 1997). The formation of methylenetetrahydrofolate(MTHF) was also directly monitored in a coupled assay with methylenetetrahydrofolate dehydrogenase (Fig. S2). In the presence of NMG the rate of MTHF formation was 10-fold higher, compared with the control reaction for the spontaneous formation from formaldehyde and H4Folate. These results suggest that NMGDH, like TSOX, may be a complex bifunctional enzyme that catalyses oxidation of NMG and the transfer of the oxidized methyl group to H4folate.

The gsIII-like gene from M. universalis FAM5 encodes γ-glutamylmethylamide synthetase

The next gene in the cluster encodes a GSIII-like protein that lacks the ammonia-binding domain and has high similarity to a recently identified GMAS from the methylotrophic bacterium Methylovorus mays No9 (Yamamoto et al., 2007; 2008). The activity of the GMAS-like enzyme was elevated in M. universalis FAM5 during growth on methylamine. We showed that γ-glutamylmethylamide is one of the major intermediates of methylamine metabolism in this culture (Table 1). In order to elucidate the role of the GSIII-like enzyme in synthesis of γ-glutamylmethylamide, a new mutant strain (NMGP3) lacking the gsIII gene was generated. NMGP3 failed to grow on 30 mM methylamine (Fig. S2). Cell extracts of methanol-grown/methylamine-induced cells of mutant strain displayed a twofold increase in NMGS and NMGDH activities while the activity of glutamine synthetase stayed at the methanol growth level (Table 2). γ-Glutamylmethylamide was not detected in the intracellular pool of free amino acids from NMGP3 mutant strain (Table 4). However, the mutant strain was able to accumulate NMG at the wild-type level. These data strongly suggest that gsIII-like gene from M. universalis FAM5 encodes GMAS.

Further phenotypic characterization of the NMGP3 strain revealed that it is able to utilize di- and trimethylamines. Weak growth of the mutant was observed at low concentration of substrate (5 mM). Prolonged incubation of the Δgms strain on solid methylamine (30 mM) containing medium resulted in selection of Δgms mutants that regained the ability to grow at high concentrations of methylamine. The growth rates of the two revertant (Δgms+) strains in liquid culture were fourfold lower than the growth rate of the wild-type. These partial revertants were found to have increased glutamine synthetase activity (data not shown). The data suggest that the ability of Δgms strains to grow on methylamine could be recovered by a spontaneous suppressor mutation or upregulation of alternative glutamine synthetase.

Putative gltB132 subcluster is essential for formation of N-methylglutamate

The last three genes of the cluster are predicted to encode a class II glutamine amidotransferase (gltB1), a glutamate synthase alpha subunit (gltB3) and a ferredoxin-dependent glutamate synthase beta subunit (gltB2) (Table 5). The genes belong to a family of archaeal gltBs identified in a large number of archaeal and bacterial genomes; however, biochemical and genetic evidence for their function(s) are still missing. Like archaeal gltBs, putative polypeptides encoded by gltB132, (33, 23 and 48 kDa, respectively) are much shorter than the typical bacterial glutamate synthase, composed of two subunits (150 and 50 kDa) (Vanoni and Curti, 1999). In order to define the specific role of this part of the gene cluster in operation of the NMG pathway, two new deletion mutants were constructed: ΔgltB1:kan (named NMGP4) and ΔgltB3:kan (NMGP5). As only partial genetic information for gltB3 is available, we could not obtain a mutant in this gene. NMGP4 and NMGP5 strains failed to use mono-, di- and trimethylamines for growth and lost the ability to produce NMG from methylamine and glutamate in vitro (Table 2). Intracellular concentrations of NMG in mutant strains were reduced three- to fivefold compared with wild-type (Table 4). It must be mentioned that intracellular concentrations of glutamate and γ-glutamylmethylamide were also reduced, while glutamine stayed at its wild-type level. No induction of NMG synthase activity above methanol growth background was observed in both gltB-like mutants.

Thus, phenotypic, metabolic and enzymatic data indicate that gltBs-like genes are essential for formation of NMG in M. universalis FAM5, and most likely are involved in the formation of this intermediate of glutamate-mediated methylamine utilization pathway. These genes were annotated as mgsABC (Table 5).


The NMG and γ-glutamylmethylamine (GMS) pathways for methylamine utilization were first proposed in the 1960s to explain methylamine growth of several microbial species, which did not demonstrate any detectable activities of MADH or oxidase (Shaw et al., 1966; Kung and Wagner, 1970). Enzymes of both pathways were purified and characterized (Pollock and Hersh, 1971; 1973; Bamforth and Large, 1977a,b; Boulton et al., 1980); however, genes responsible for these specific metabolic functions were not identified. The results presented here provide the first genetic overview of glutamate-mediated oxidation of methylated amines in the methylotrophic beta-proteobacterium Methyloversatilis universalis FAM5. An 8-gene cluster essential for methylamine growth of the culture was identified, and specific metabolic functions were assigned to the genes (Table 5). The first four genes of the cluster, mgdABCD, exhibited modest sequence homology to a soxBDAG operon and were shown to encode NMGDH. Predicted subunit composition (44, 100, 12 and 10 kDa), solubility and inability of the enzyme to use DCPIP as an electron acceptor show that NMGDH from M. universalis FAM5 is different from previously characterized enzyme from Pseudomonas aminovorans, which was predicted to be a homotetrameric (subunit molecular weight ∼130 kDa) integral membrane protein (Bamforth and Large, 1977b). However, similar to the previously described enzyme, it is predicted to be a flavohaemoprotein. So far, we could not find any suitable explanation for our inability to observe a DCPIP-linked oxidation of NMG in protein extract or partially purified enzyme from M. universalis FAM5. At the same time, NMGDH from M. universalis FAM5 resembles the soluble NAD-activated enzyme from methylotrophic alpha-proteobacteria Hyphomicrobium vulgare (Loginova et al., 1974). This soluble enzymatic system seems to be quite widespread among different phyla of bacteria (Doronina et al., 2004; Miller et al., 2005; Trotsenko et al., 2007, N.V. Doronina and Y.A. Trotsenko, pers. comm.); however, it still awaited detailed enzymatic characterization in terms of kinetic parameters of NMG oxidation. Our preliminary studies indicated that the enzyme from M. universalis FAM5 is also similar, in most respects, to heteroterameric sarcosine oxidase: it is predicted to be a complex, multisubunit enzyme of the of amine oxidizing flavoenzymes family, requires NAD, has a putative binding site for tetrahydrofolates, and likely produces methylenetetrahydrofolate as the end-product of methyl group oxidation. The latter feature of this enzymatic system in question is quite captivating. Formaldehyde was assumed to be the end-product of NMG oxidation (Hersh et al., 1971; Bamforth and Large, 1977a; Boulton et al., 1980). However, previous in vivo studies have demonstrated that there is no significant production of formaldehyde from the oxidation of NMG, and there were speculations that this step in methylamine utilization is tightly coupled to formaldehyde metabolism (Loginova et al., 1974; Jones and Bellion, 1991). The lack of product inhibition by glutamate or formaldehyde on the partially purified ‘solubilized’ enzyme has also raised a possibility that the end-product of the reaction is N-methylene-glutamate rather than glutamate or formaldehyde (Hersh et al., 1972). Furthermore, the low rate of formaldehyde oxidation in Pseudomonas sp. MS led to speculation that oxidation of methylamine in this bacterium with well-characterized particulate NMGDH occurs as H4folate derivatives (Kung and Wagner, 1970). We showed that mutation of aminomethyltransferase, a subunit similar to the methylene-H4folate forming subunit of TSOX, results in failure of M. universalis FAM5 to grow on methylamines or produce active NMGDH. These data are first genetic evidence that support the direct formation of methylenetetrahydrofolate as an end-product of NMG oxidation. Hence, the NMG pathway operating in M. universalis FAM5 may be the second example of methylotrophic pathway, after chloromethane degradation pathway (Studer et al., 2002), which involves H4folate derivatives instead of formaldehyde, as key intermediates of C1-oxidation.

N-methylglutamate synthetase from Pseudomonas MS, a well-characterized enzymatic system, was described as an FMN-containing enzyme composed of approximately 12 subunits (molecular weight 30–35 kDa) (Pollock and Hersh, 1971; 1973). NMG synthetase from M. universalis is predicted to be a multisubunit enzyme with molecular weight of subunits approximately 24, 33 and 48 kDa. The large subunit has a FMN-binding domain, thus like in the previously described enzymatic systems, it seems to be a complex flavoprotein. Mutation analyses demonstrated that both mgsA (encoding 24 kDa subunit) and mgsB (gene for 33 kDa subunit) are essential for growth of M. universalis FAM5 on methylated amines. These mutations resulted in significant reduction of the intracellular pool of NMG; however, they did not abolish the formation of NMG completely. The most obvious explanation of the NMG accumulation would be that mutant strains are still able to form a partially active enzyme. Whether all three genes are in fact encoding functional subunits of NMGS remains to be established by purification of the enzyme in order to determine the type and number of subunits and cofactors and to characterize individual subunits in terms of their catalytic activity and specific function. The data presented here provide the genetic basis for the following thorough study of this complex enzyme.

The published data on the function of bacterial GMAS are controversial. It was believed that γ-glutamylmethylamide serves as a temporary storage sink for methylamine (Konishi et al., 1972). It was also believed that γ-glutamylmethylamide is an intermediate of NMG synthesis, and that it serves as the true substrate for the NMG synthase (Large and Bamforth, 1988). Later, it was suggested that GMAS may be involved in attenuation of the ammonia flow, a side product of methylamine oxidation (Jones and Bellion, 1991). Without a doubt such a mechanism would seem to be essential during growth on methylamine due to an unsuitably high ratio of N:C compared with that needed for growth. On the other hand, the enzyme involved in dissimilation of γ-glutamylmethylamide to ketoglutarate, formaldehyde and ammonia has been purified and characterized, thus also suggesting a role of the enzyme in methylamine oxidation (Kimura et al., 1995). The γ-glutamylmethylamide dissimilation step was not detectable in M. universalis FAM5, indicating that, in this strain, GMA most likely is not an intermediate of C1- oxidation. We defined the function of GMAS using a mutagenesis approach. The mutational evidence strongly suggests that γ-glutamylmethylamide is not an intermediate of the NMG pathway, since the mutation of the gmas gene did not influence the formation of NMG. Our data support the hypothesis that GMAS plays a role in balancing carbon/nitrogen flow since the Δgmas strain was still able to grow at low concentrations of methylated amines.

Comparative genomic studies indicated that genes similar to the NMG pathway gene are present in several species of bacteria from different microbial phyla: Burkholderia phymatus SMT815, Methylobacillus flagellatus KT, Pseudomonas mendocina, Thiomicrospira crunogena XCL-2 and Rubrobacter xylanophilus DSM. Despite the high phylogenetic diversity of these microbes, the mgdABCD-gms-mgsABC clusters share high sequence identity (Fig. 1B). Methylotrophic capabilities have never been reported for Rubrobacter xylanophilus DSM, while B. phymatum ST815 has been shown to use methylated amines (L. Chistoserdova, unpubl. data). The ability of P. mendocina strains to oxidize methylated compounds is known and has an established biotechnological potential (Pandey et al., 2006). NMG pathway genes in these microbes are clustered together with two genes for key enzymes of folate-linked C1-metabolism, purU and folD (Fig. 1B). Other genes involved in the single carbon metabolism, such as serine cycle, were identified downstream of folD in B. phymatum STM815. In this bacterium all genetic components of methylamine oxidation (mgdABCD-gmas-mgsABC, purU, folD) and assimilation (serine cycle genes) are located on a plasmid, thus implicating that this function could be subject to lateral gene transfer. The presence of similar proteobacterial-like genes in Rubrobacter xylanophilus DSM could be an example of such genetic exchanges. From all of the described microbes possessing this highly conserved gene cluster, only M. flagellatus KT has MADH, a canonical enzyme for methylamine oxidation. The genomes of B. phymatum STM815 and Pseudomonas mendocina lack any known system for methylamine oxidation, and it is most likely that the NMG pathway plays a key role in methylamine utilization in these bacteria.

Experimental procedures

Bacterial strains, plasmids and culture conditions

Methyloversatilis universalis strains used in this study are listed in Table S1. These were grown in a minimal medium described previously (Kalyuzhnaya et al., 2006). Succinate (20 mM), methylamine (30 or 5 mM), and methanol (25 mM) were used as growth substrates. For NMG-pathway enzymes induction in mutant strains unable to grow on methylamine, methanol-grown cells were collected by centrifugation at 5000 r.p.m. for 5 min, washed with sterile medium and exposed to methylamine at 30°C with shaking for 12 h. The following antibiotic concentrations were used for M. universalis (µg ml−1): tetracycline (Tet), 1.0; kanamycine (Kan), 100; chloramphenicol (Cmp), 10–15.

Escherichia coli strains were routinely cultivated at 37°C in Luria–Bertani medium (BD Difco). The following antibiotic concentrations were used: Tet, 12.5 µg ml−1; Kan, 100 µg ml−1; and ampicillin (Amp), 100 µg ml−1. The following cloning vectors were used: pCR2.1 (Invitrogen) for cloning of polymerase chain reaction (PCR) products, and pCM184 for generation of mutant strains (Marx and Lidstrom, 2002). Biparental matings between E. coli and M. universalis were performed on defined solid minimal medium supplemented with 10% nutrient broth (BD Difco) and methanol (25 mM) at 30°C for 48 h. Cells were then washed with sterile medium and plated on selective medium: defined mineral medium supplemented with methanol (25 mM), Kan (100 µg ml−1) and Cmp (10 µg ml−1). Chloramphenicol was used for E. coli counter-selection.

LC-MS/MS analysis of intracellular metabolites of M. universalis FMA5 grown on methanol and methylamine

Samples (10 ml at OD = 0.25–0.3) of cell culture exponentially grown with methanol or methylamine were rapidly transferred into 30 ml of cold (−40°C) quenching solution: 60% methanol (v/v) in 70 mM HEPES buffer (pH 6.8). The quenched biomass was collected by centrifugation for 10 min at 10 000 r.p.m. at −20°C in Dupont Sorvall RC5B refrigerated centrifuge (Waltham, MA, USA). Cell pellets were used for intracellular metabolites extraction using previously described method (1) with a few modifications: 1 ml of boiling HEPES buffered ethanol solution (75% v/v ethanol/water, pH 5.2) was added to cell pellets and incubated at 100°C for 5 min. The extracted cell suspensions were incubated on ice for 5 min and the cell debris was removed by centrifugation at 4500 g for 5 min. The cell-free metabolite extract was centrifuged at 20 817 g for 8 min. The supernatant was transferred into a 2 ml glass vial and lyophilized in a vacuum centrifuge (CentriVap Concentrator System, Labconco, MO, USA). Dried sample were dissolved in 100 µl purified water for LC-MS/MS analysis. LC-MS/MS experiments were carried out on a Waters LC-MS system (Milford, MA, USA) consisting of a 1525 µ binary HPLC pump with a 2777C autosampler as described previously (Yang et al., 2009). LC solvents for the pentafluorophenylpropyl-bonded silica column (Luna PFPP, 150 mm × 2 mm, 3 µm, Phenomenex, Torrance, CA, USA) were the following: mobile phase A consisted of 0.1% formic acid in water, while mobile phase B was acetonitrile. The following gradient was used 100% A for 8 min, 100–0% A for 2 min, 0% A for 5 min, 0–100% A for 2 min, 100% A for 8 min. Flow rate was 0.15 ml min−1. The multiple reaction monitoring pairs of standard metabolites were glutamate (ESI+, 148.1→84.0), NMG (ESI+, 162.1→98.0), glutamine (ESI+, 147.1→130.1) and alanine (ESI+, 89.9→43.9). γ-Glutamylmethylamide was identified by selected ion monitoring at m/z 161.1 and further confirmed by daughters scan. Dry weight of methanol and methylamine grown cultures was determined as described (Stepan-Sarkissian and Grey, 1990).

Enzyme assays

All spectrophotometric measurements were performed using a BioSpec-1601 spectrophotometer (Shimadzu, Japan). Crude cell protein extracts were prepared as described above. GMAS (EC; also known as L-glutamate:methylamine ligase) activity was assayed by measuring the formation of γ-glutamylhydroxamate at pH 7.5 as described (Yamamoto et al., 2007). Reaction mixtures (total volume 0.5 ml) were incubated at 30°C for 20 min. The reaction was terminated by addition of 1 ml of the stop solution (g l−1): FeCl3 × 6H2O, 54; trichloroacetic acid, 19.6; 12 N HCl, 20 ml (O'Neal and Joy, 1974). Reaction tubes were centrifuged to remove any precipitation. Accumulation of γ-glutamylhydroxamate was recorded at 540 nm. The γ-glutamylmethylamide dissimilating enzyme was assayed as described (Kimura et al., 1995). The optimized assay for NMGDH included (0.5 ml): 50 mM sodium phosphate buffer (pH 7.6), 5 mM NMG, 0.5 mM of NAD and protein extract/fraction (0.2–0.5 µg). The reaction was initiated by addition of NMG, and after incubation at room temperature for 15–20 min, terminated by addition of 0.5 ml Nash reagent (Nash, 1953). Accumulation of formaldehyde was recorded spectrophotometrically at 412 nm. Activity of sarcosine oxidation was tested in similar assay, in which NMG was replaced by sarcosine (10 mM). When required, a small aliquot (100–500 nmol) of anoxic stock solution of tetrahydrofolate (H4folate; Sigma Life Sciences) was added to the aerobic reaction mixture just prior to assay initiation. Protein extract/fractions were omitted from control reactions, which were performed in both the presence and the absence of H4folate. Activity of NMG formation was tested in a modified NMGDH assay. The reaction mixture included: 50 mM sodium phosphate buffer (pH7.6), 10 mM of sodium glutamate, 10 mM methylamine, 5 mM MgCl ions, 0.5 mM of NAD and protein extract/fraction. The reaction was initiated by addition of methylamine, and terminated after incubation at room temperature for 30 min by the addition of 0.5 ml Nash reagent (Nash, 1953). Activities are reported in nmol min−1 mg protein−1. Protein concentrations were determined spectrophotometrically (Stoscheck, 1990).

Identification of proteins involved in methylamine utilization

Methanol- and methylamine-grown cells of M. universalis FAM5 (OD600 = 0.5–0.7) were harvested by centrifugation at 4500 g at 4°C for 25 min. Cells were re-suspended in 50 mM potassium phosphate buffer (pH 7.5) and passed two times through a French pressure cell at 1.2 × 108 Pa. Centrifugation was performed at 15 000 g for 30 min at 4°C to remove cell debris. Crude extracts were treated with (NH4)2SO4 and 0–20%, 20–30%, 30–40% and 40–70% saturation fractions were collected. Each protein fraction was subjected to enzyme activity measurements and denaturing polyacrylamide (4–24%) gel electrophoresis (SDS-PAGE) as described (Laemmli, 1970). Proteins were visualized by staining with Coomassie brilliant blue R250 (Amersham Biosciences). Major polypeptide bands with molecular masses of approximately 20, 46 and 95 kDa were excised from the gel and subjected to LC/LC-MS/MS peptide mass fingerprinting.

Mass spectrometry

Protein bands 2 and 3 were analysed at the Biological Mass Spectrometry and Proteomics Facility in the Department of Biological Sciences, University of Warwick ( Protein bands of interest were processed on the MassPrep robotic protein handling system. The tryptic digests were analysed by means of nanoLC-ESI-MS/MS and the lockmass-corrected data used to interrogate suitable databases using a 50 p.p.m. error tolerance. Protein sequence of gms genes from Methylovorus mays No9 was appended to the UniProtKB database release 13.3 (May08) and the search engine ProteinLynx Global Server (v2.3) was used to suggest protein identities.

Protein bands 1, 2a and 3 were in-gel digested as described previously (Shevchenko et al., 1996) with minor modifications. Briefly, the gel spots were distained with 200 mM NH4HCO3 in 50% acetonitrile, washed 4 times with water, shrunk with 100% acetonitrile and dried in a Speed-Vac (Jouan Model 1022, St-Herblain, France). Trypsin solution was prepared in 50 mM NH4HCO3 and 5 mM CaCl2 in 10% acetonitrile to 20 ng of trypsin per microlitre. An adequate volume of trypsin solution was added to cover the gel spot, incubated on ice for 1 h then incubated at 37°C for 13 h. After digestion, the gel spots were sonicated 15 min with 20 mM NH4HCO3 once, followed by 5% formic acid in 50% acetonitrile three times. The combined extracts were concentrated by Speed-Vac, diluted to about 10% acetonitrile and concentrated again to about 15 µl, followed by C18 Zip-Tip (Millipore, Billerica, MA, USA) purification according to the manufacturer's protocol. The peptide solutions, after Zip-Tip cleaning, were concentrated by Speed-Vac, diluted to about 10% acetonitrile and concentrated again to about 20 µl. The resultant sample (3 µl) was loaded pneumatically into a 75 µm i.d. × 360 µm o.d. capillary HPLC column packed with 10.2 cm of 5 µm AQUA C18 (Phenomenex, Torrance, CA, USA). LC/MS analysis was performed using an LTQ mass spectrometer (Thermo Electron Corp. San Jose, CA, USA), Magic 2002 HPLC (Michrom BioResouces, Auburn, CA, USA) and an in-house built micro-ESI interface. Solvent A was 0.5% acetic acid in water, B, 0.5% acetic acid in acetonitrile. The gradient started at 5% B, held 5 min, increased to 10% B in 1 min and held for 4 min, to 40% B in 45 min, to 80% B in 1 min and held for 9 min, then down to 5% B in 5 min and held for 10 min. The flow rate was about 0.3 µl min−1. The MS1 scan range was 400–2000 m/z units, each MS1 scan was followed by 10 MS2 scans for the 10 most intense ions in the MS1 scan. Default parameters under the Xcalibur 1.4 data acquisition software (Thermo) were used, with the exception of an isolation width of 3.0 m/z units and normalized collision energy of 40%. Raw mass spectral data were searched by SEQUEST (Eng et al., 1994). The ORF database included normal and reversed sequences from microbial genomes from NCBI (Concise Microbial Protein Database, state of curation as it existed on 20 January 2009, 3.5 Gigabytes total size for forward and reversed sequences) or Lake Washington composite metagenome (LWM, 96 Mbytes) (DOE Joint Genome Institute, and the human subset of the nrdb (NCBI). The SEQUEST search results were filtered by DTASelect Ver. 1.9 (Tabb et al., 2002), then input into a FileMaker script developed in-house for further processing. The parameters were selected to yield a false-positive rate (Peng et al., 2003) of about 4% based on hits to the reversed LWM sequences (fully tryptic peptides only, Sequest Xcorr thresholds of 1.9, 2.2 and 3.3 for singly, doubly and triply charged peptides respectively). A minimum of two unique peptides was required to identify each protein, following the definition of uniqueness used by DTASelect in which each charge state of a given precursor peptide is considered separately (Tabb et al., 2002). Proteins shown in Table 4 with a single unique peptide, using a more restrictive definition of uniqueness (Xia et al., 2006), were identified with multiple charge states associated with the same amino acid sequence. Only peptides with many repeated observations (spectral counts) typically seen for the primary components of gel spots were retained in the final data set.

DNA manipulations

DNA was isolated using the QIAamp DNA Mini kit (Qiagen). Plasmid DNA was purified using the GeneJet Plasmid Miniprep Kit (Fermentas). E. coli transformation, restriction enzyme digestion and ligation reactions were carried out as described by Maniatis et al. (1982). PCR amplifications were performed using Taq polymerase (Qiagen) in accordance with the manufacturer's instructions. Degenerate primers for amplification of putative soxA, soxB, gsIII, gltB1, gltB3 and gltB2 genes were designed based on conservative regions of corresponding genes from Burkholderia phymatum STM815 plasmid (NCBI accession number CP001045), Methylobacillus flagellatus KT (CP000284), Pseudomonas mendocina YMP (CP000680) and Methylovorus mays No9 (BAF99006). DNA sequence alignment for gene clusters was performed in VectorNTI AlignX package (Invitrogen). After amplification of gene fragments, primer sets specific to the gene from M. universalis FAM5 were designed and used to amplify intergenic regions (Table S3). The gene cluster was assembled using Vector NTI Contig Express software (Invitrogen). Primers used for amplification of the NMG pathway genes and for sequencing of the cluster are listed in Table S1.

Phylogenetic analysis

Amino acid sequences were aligned using the ClustalX program ( For phylogenetic analyses, the Phylip package ( was used. Maximum likelihood, distance and parsimony methods were employed, and 1000 bootstrap analyses were performed.

Construction of donor plasmids and generation of mutant strains

Primers used for PCR amplification of upstream or downstream regions (approximately 450–500 bp) of each targeted gene are listed in Table S1. Amplified fragments were cloned into pCR2.1, sequenced and then subcloned into pCM184 using appropriate restriction sites (Table S1). After verification of the nucleotide sequence, the plasmids were transformed into E. coli S17–1, and the resulting donor strains were mated with wild-type M. universalis FAM5 via biparental mating. The KanR recombinants were selected on methanol plates and checked for resistance to Tet. Tet-sensitive (TetS) colonies were chosen as possible double-cross-over recombinants. The identity of the double-cross-over mutants was further verified by diagnostic PCR with primers specific to the insertion sites. Mutant phenotypes were assessed on solid media containing methanol or methylamine as carbon source. The ability of mutant strains to grow on methylamine was also tested in liquid medium. The growth was monitored using a growth analyser Bioscreen C (GrowthCurves USA).


Authors would like to acknowledge the contributions of the Biological Mass Spectrometry and Proteomics Facility in the Department of Biological Sciences, University of Warwick. Authors are very grateful to Mary Lidstrom, Yurii Trotsenko and Stéphane Vuilleumier for many insightful suggestions on the manuscript. We thank Fred Taub for his assistance with the Concise Microbial Database and FileMakerTM applications. This work was supported by Royalty Research Fund (award# 65-1818) and by the National Science Foundation (MCB-0842686). Mass spectrometry analyses were also supported in part by NSF Grant MCB-06044269 and NIGMS Grant GM-58933.