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Journal of Bacteriology, November 2005, p. 7511-7517, Vol. 187, No. 21
0021-9193/05/$08.00+0     doi:10.1128/JB.187.21.7511-7517.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

QscR-Mediated Transcriptional Activation of Serine Cycle Genes in Methylobacterium extorquens AM1

Marina G. Kalyuzhnaya1* and Mary E. Lidstrom1,2

Department of Chemical Engineering,1 Department of Microbiology, University of Washington, Seattle, Washington 98195-17502

Received 18 May 2005/ Accepted 8 August 2005


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
QscR, a LysR-type regulator, is the major regulator of assimilatory C1 metabolism in Methylobacterium extorquens AM1. It has been shown to interact with the promoters of the two operons that encode the majority of the serine cycle enzymes (sga-hpr-mtdA-fch for the qsc1 operon and mtkA-mtkB-ppc-mclA for the qsc2 operon), as well as with the promoter of glyA and its own promoter. To obtain further insights into the mechanisms of this regulation, we mapped transcriptional start sites for the qsc1 and qsc2 operons and for glyA via primer extension analysis. We also identified the specific binding sites for QscR upstream of the qsc1 and qsc2 operons and glyA by DNase I footprinting. The QscR protected areas were located at nucleotides –216 to –165, nucleotides –59 to –26, and nucleotides –72 to –39 within the promoter-regulatory regions upstream of transcriptional starts of, respectively, qsc1, qsc2 and glyA. To examine the nature of the metabolic signal that may influence QscR-mediated regulation of the serine cycle genes, Pqsc1::xylE translational fusions were constructed and expression of XylE monitored in the wild-type strain, as well as in knockout mutants defective in a variety of methylotrophy functions. The data from these experiments pointed toward formyl-H4F being a coinducer of QscR and possibly the major signal in the regulation of the serine cycle in M. extorquens AM1. The ability of formyl-H4F to enhance the binding of QscR to a specific region upstream of one of the serine cycle operons was demonstrated in gel retardation experiments.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Methylobacterium extorquens AM1 is an aerobic facultatively methylotrophic bacterium that uses reduced C1 compounds as sole sources of carbon and energy. The availability of both the genomic sequence (http://www.integratedgenomics.com/genomereleases.html#6), and an array of genetic tools (19, 20, 22) make M. extorquens AM1 an attractive model for biochemical and genetic studies in methylotrophy. Extensive genome-based analysis of methylotrophy in M. extorquens AM1 has revealed that over 100 genes are involved in C1 metabolism, and these belong to a number of specialized metabolic modules (11). One of these modules is the serine cycle for formaldehyde assimilation (QSC), the main assimilatory pathway for methylotrophic growth (1). Six of the serine cycle genes are clustered at one end of a large methylotrophy island, and these form two different operons (2, 4-9, 11, 16). The first operon (qsc1) encodes the genes for serine glyoxylate aminotransferase (sga), hydroxypyruvate reductase (hpr), methylene-H4MPT/methylene-H4F dehydrogenase (mtdA) and methenyl-H4F cyclohydrolase (fch). The second operon (qsc2) encodes the genes for malate thiokinase (mtkA and mtkB), phosphoenolpyruvate carboxylase (ppc) and malyl-CoA lyase (mclA). Genes encoding the remaining reactions of the serine cycle, i.e., glyA (serine hydroxymethyltransferase), mdh (malate dehydrogenase), gckA (glycerate kinase), and eno (enolase) are not parts of methylotrophy gene clusters (7, 8, 11). We have previously shown that expression of the majority of the serine cycle genes is regulated by a LysR-type transcriptional regulator, QscR (16). QscR controls expression of the assimilatory module via coordinate transcription of several operons. Therefore QscR coordinates the expression of the module in respond to physiological shift from multicarbon to methylotrophic growth. This regulator is essential for transcription from the qsc1 operon, and is also required for methanol-dependent induction of transcription from the qsc2 operon and from glyA. Since two of the H4F-linked C1 transfer module genes (mtdA and fch) are a part of the qsc1 operon, QscR is also essential for activation of this module during growth on C1 substrates. The linkage and co-regulation of two enzymes from this C1-transfer pathway with serine cycle enzymes suggested that both modules are functionally connected. This hypothesis has been supported by flux analysis data indicating that the H4F-linked C1 transfer module is involved in conversion of formate to methylene-H4F, the starting substrate for the serine cycle (18).

QscR is expressed at low levels under both methanol or succinate growth conditions, suggesting that the protein might respond to a physiological signal or signals that lead to altered binding specificity and increased expression of serine cycle genes. So far, none of many tested metabolites, intermediates of the serine cycle or energy metabolism, have exerted positive effects on the binding specificity of QscR, although the addition of 50 µM NADP to the QscR binding assay resulted in a tenfold decrease of DNA binding (16).

The overall goal of our ongoing investigation is to understand the genetic and physiological mechanisms involved in the regulation of serine cycle gene expression in M. extorquens. Here, we present the transcript analyses of the serine cycle regulons in M. extorquens AM1. Moreover, we present genetic and physiological in vivo studies pointing to potential effector molecules generated in the H4F pathway. Here, we show that formyl-H4F contributes to QscR-mediated control of most serine cycle genes in M. extorquens AM1.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacterial strains, plasmids and culture conditions. Strains of M. extorquens AM1 used in this study are listed in Table 1. These were grown in a minimal medium described previously (15). Succinate (20 mM) or methanol (100 mM) was used as substrate. Mutants CM280K.1, CM279K.1, and CM275K.1 were grown in succinate-containing medium in the presence of 10 mM methanol. The following antibiotic concentrations were used for M. extorquens (µg ml–1): Tet, 12.5; Kan, 100; rifamycin, 50. For serine cycle enzyme induction in mutant strains unable to grow on C1 compounds, succinate-grown cells were collected by centrifugation at 5,000 rpm for 5 min, washed with sterile medium and exposed to methanol at 30°C with shaking for 20 min, 2 h, or 16 to 18 h.


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  		TABLE 1. Bacterial strains used in this work

 
Escherichia coli strains JM109 (34), BL21(DE3) (Novagen, Madison, WI), S17-1 (30) and TOP 10 (Invitrogen) were routinely cultivated at 37°C in Luria-Bertani medium (29). The following antibiotic concentrations were used: Tet, 12.5 µg ml–1; Kan, 100 µg ml–1; Amp, 100 µg ml–1.

The following cloning vectors were used: pRK2013 (13) as a helper plasmid, pCR2.1 (Invitrogen) for cloning of PCR products, pCM130 (19) for promoter fusion construction, pET21d (Novagen) as expression vector.

Triparental or biparental matings between E. coli and M. extorquens were performed overnight on nutrient agar at 30°C (3, 14). Cells were then washed with sterile medium and plated on selective medium at appropriate dilutions. Rifamycin was used for E. coli counterselection.

Construction of promoter fusions. The followed primers were used for the amplification of putative promoter regions of a varying size upstream of the first gene of the qsc1 operon: qsc1-1f, 5'-AAACTGCAGCTTCGGCGTTTGGGCGGTCT-3'; qsc1-2f, 5'-AAACTGCAGCGGCGGGGGAGGCATAACGG-3'; qsc1-3f, 5'-AAACTGCAGCGACATCGCAGAGGTACGG; and qsc1R, 5'-CCCAAGCTTGGAACGAACAGGTGGTTGCGT-3'. The primers were specifically designed to create a PstI site at the 5' end of the fragment and a HincIII site at the 3' end of fragment. These sites were used to clone the resulting fragments into the pCM130 vector. The resulting Pqsc1::xylE fusions were transferred into M. extorquens AM1 strains (Table 1) via conjugation, and expression of the qsc1operon was determined by measuring catechol 2,3-dioxygenase (XylE).

Enzyme assays. For cell extract preparation, cells were pelleted by centrifugation at 5,000 rpm for 8 min, resuspended in 1 ml of 25 mM KH2PO4 · Na2HPO4 buffer (pH 7.5), and disrupted by passing through a French pressure cell at 1.2 x 108 Pa. Cell extracts were centrifuged at 14,000 rpm for 25 min at 4°C to remove cell debris. The assays were performed routinely at 30°C in 1.5-ml cuvettes (path length, 1 cm) in a total volume of 1 ml. XylE activity was determined spectrophotometrically at 375 nm as described previously (17). One mU of XylE activity corresponds to 1 nmol produced catechol 2-hydroxymuconic semialdehyde min–1 per mg protein ({varepsilon} = 33 mM–1 cm–1). Serine cycle enzyme activities (Sga and Hpr) were assayed as described previously (4, 5). Methylene tetrahydrofolate (H4F) dehydrogenase (MtdA) was assayed as described previously (31). One mU of Sga, Hpr, and MtdA activities corresponds to 1 nmol oxidized NAD(P)H min–1 per mg protein ({varepsilon} = 6.22 mM–1 cm–1). Protein concentration was measured by the Biuret reaction (33) using bovine serum albumin as a standard or spectrophotometrically.

Transcriptional start site mapping. Total RNA from exponentially growing M. extorquens was isolated using TRIzol reagent according to the manufacturer's instructions (Invitrogen) and kept at –80°C as an ethanol precipitate. Transcriptional start sites were mapped by means of primer extension, using the First Strand cDNA synthesis kit for RT-PCR, as recommended by the manufacturer (Roche). Ten micrograms of RNA was used per reaction. Oligonucleotide primers were labeled with [{gamma}-32P]ATP using T4 polynucleotide kinase (25). The following primers were used in the transcription start site mapping for qsc1, glyA, and qsc2, respectively: sga1, 3'GAGCGGCGAAGCGGCCGTCG-3'; sga2, 5'-ACCGTACCTCTGCGATGTC-3'; gly1, 5'-GCAAGATGAGCCGAGAAGAAG-3'; gly2, 5'-GGCGATCTCGGGATCGGTCT-3'; mtk1, 5'-ACCCCGAACGCTCGCGAGCAG-3'; mtk2, 5'-GGGCTGAAAGCCACGG-3'.

DNase I footprint analysis. QscR was purified as described previously (16). The probes for DNase I footprint analysis were prepared by PCR amplification. The specific primer pairs for amplification have been described before (16) and were designed to contain the additional restriction sites PstI (the upstream primer) and HindIII (the downstream primer). The DNA fragments were first labeled at both ends using T4 polynucleotide kinase (Promega). To obtain a single end-labeled DNA, probes were then digested with HindIII or PstI to release one of the labeled ends. The G-A DNA sequence ladders were generated using the chemical cleavage procedure of Maxam and Gilbert as described previously (24).

Gel retardation assays. Gel retardation assays were performed essentially as described previously with the 32P-labeled Pqsc1-1 and Pqsc1-2 promoter fragments. The purified QscR (0.5 mg) was preincubated with varying amounts (5 to 50 µM) of formate, formaldehyde, formyl-H4F, methenyl-H4F or methylene-H4F for 10 min, followed by 20 min of incubation at room temperature with labeled DNA in gel shift binding buffer (Promega): 5 mM Tris-HCl (pH 7.5), 50 mM NaCl, 1 mM MgCl, 0.5 mM dithiothreitol, 0.5 mM EDTA, 4% glycerol, 0.05 µg/ml poly(dI-dC) · poly(dI-dC). Mixtures were then loaded onto a 6% nondenaturing polyacrylamide gel in 0.5x TBE and electophoresed at 270 V. Gels were subsequently dried and exposed to X-ray film (Kodak). The images have been analyzed by using Kodak 1D Image Analysis software (Kodak).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Mapping of transcriptional start sites of the qsc1 and qsc2 operons, and of glyA. Primer extension mapping experiments were carried out using total RNA extracted from cells in the early exponential phase of growth on methanol or succinate. With both RNA samples, the primer extension product for qsc1 was located at position –343 relative to the putative translational start of sga, the first gene in the qsc1 operon (Fig. 1A; also succinate data not shown). The location of the dominant initiation start site for the qsc2 operon depended on the growth conditions. Cells grown on succinate had a dominant start site at position –80 (TS1) with a minor start site at –46 (TS2) with respect to the start codon of mtkA, the first gene of the qsc2 operon. In methanol-grown cells, the dominant transcript was initiated at the –46 position, with a weaker start site at the –80 position (Fig. 1B). Primer extension analysis of glyA transcripts from both RNA samples revealed three transcriptional start points located at positions –84(TS1), –63(TS2), and -38(TS3) relative to the putative translational start site. In methanol-grown cells, TS1 and TS2 dominated, while in succinate-grown cells, TS3 dominated (Fig. 1C).



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  		FIG. 1. Mapping the transcription initiation sites for qsc1 (A), qsc2 (B), and glyA (C). Shown are results of primer extension experiments with total RNA extracted from methanol (m) or succinate (s) grown cells M. extorquens AM1. Arrows indicate the transcription initiation sites.

 
DNA sequence analysis of the promoter-regulatory regions of qsc1, qsc2, and glyA operons identified potential {sigma}70 type –10 and –35 promoter consensus sequences, although as noted before (35), the –10 sequences are divergent and tend to be more G+C rich than the E. coli consensus sequence (Table 2).


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  		TABLE 2. Alignment of the putative –10 and –35 regions of qsc1, qsc2, and glyA with the ones of known {sigma}70-type promoters from M. extorquens AM1 (mxaF, mxaW) and the E. coli {sigma}70 consensus sequence

 
Determination of DNA binding sites for QscR. Previous results clearly indicate that QscR is a transcriptional activator for the qsc1 and qsc2 operons and glyA and a repressor of qscR (16). Using gel shift assays, in the same study it was shown that QscR binds to the promoter-regulatory regions of the serine cycle genes. In order to map the binding sites for QSC genes in M. extorquens AM1 DNase I protection assays were carried out using partially purified preparations of QscR. Three previously characterized promoter regions for qsc2 (upstream mtkA region), glyA, and qscR, fragments of 520 bp, 567 bp, and 273 bp containing sequences upstream of the respective genes, were used (16). Since the previously described qsc1 (16) promoter region (650 bp) was too big to be used in footprint experiments, the region required for transcriptional regulation of the qsc1 operon was narrowed down using XylE transcriptional fusion plasmids containing 441 bp (Pqsc1-1), 294 bp (Pqsc1-2), and 160 bp (Pqsc1-3) of sequence upstream of the qsc1 transcription start. The plasmids were transferred into M. extorquens AM1, and the plasmid-containing strains were then assayed for catechol 2,3-dioxygenase activities under methylotrophic growth conditions. Both the Pqsc1-1 and Pqsc1-2 regions resulted in normal expression of the qsc1 operon, while the Pqsc1-3 fusion resulted in low XylE activity (data not shown). These results suggest that the important region for expression of the qsc1 operon lies between 160 and 294 bp upstream of the transcription start site, and the Pqsc1-2 fragment was used to further refine the QscR binding site.

The QscR-protected region for the qsc1 promoter-regulatory region was located between nucleotides –216 to –165 with respect to the transcription start site of the qsc1 region (Fig. 2A). The DNase I sensitivity site was found within this region, at –184 bp. This protected region covered two putative consensus LysR-type binding sites (T-N11-A) (Fig. 3). QscR protected the sequence spanning nucleotides -72 to –39 upstream of the transcriptional start of glyA from DNase I digestion (Fig. 2C). In the protected area, two consensus LysR-type motifs have been identified (Fig. 3).



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  		FIG. 2. DNase I footprint analysis of the binding of QscR to the qsc1 (A), qsc2 (B), glyA (C), and qscR (D) promoter-operator regions. Standard lanes (G/A) contain a Maxam and Gilbert cleavage ladder of the probe used in the experiment. The amount of QscR (mg) in each reaction mixture is shown on top. Bars indicate regions of protection.

 


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  		FIG. 3. Summary of mapping and DNase I protection results for QscR binding to the qsc1, qsc2 and glyA promoter-operator regions. Bars (R site and A site) indicate putative QscR binding sites. Position of a LysR-motif (T-N11-A) in bold. –35 and –10 regions are underlined and bold. At the +1 transcription initiation site arrows indicate translation starts.

 
As shown in Fig. 2B, QscR protected the qsc2 promoter from DNase I digestion at –26 to –59 bp relative to the transcriptional start site qsc2-1 or –22 to +10 relative to the second qsc2 transcriptional start. The major hypersensitive site was present at position –39 (relative to the first transcriptional start) on the top strand. This protected region covered one of the three putative LysR type (T-N11-A) binding sites of the promoter region (Fig. 3). In the protected area, nucleotides between –26 and –38 were strongly protected, and this site covered the T-N9-A motifs.

A labeled probe spanning nucleotides –148 to +125 (relative to the translation site of QscR) was used to identify the QscR binding site on the qscR promoter regions. The DNase I protected region was located at nucleotides +29 to –18 of qscR relative to translation site of QscR (Fig. 2D) and covered two consensus LysR-type motifs (Fig. 3).

The protected regions of all the promoters tested were represented by AT-rich sequences. However, we were not able to identify a consensus sequence for the four binding sites other than the general LysR-type regulator consensus T-N11-A (Fig. 3).

Identification of a co-inducer molecule for QscR. LysR-type transcriptional activators generally require a coinducer, and the latter is often a metabolite of the pathway regulated. However, none of the previously tested serine cycle intermediates, i.e., serine, hydroxypyruvate, glyoxylate, acetyl-CoA or phosphoenolpyruvate, or energy metabolism intermediates, i.e., NAD, NADH, NADPH, ATP or ADP, were shown to enhance QscR binding activity (16). In order to obtain further insights into the nature of the putative positive regulator, a set of mutant strains with lesions at various steps of C1 metabolic pathways were employed (Table 1; Fig. 4). These were used to test for expression of the qsc1 operon, by measuring XylE activity resulting from the Pqsc1-1::xylE fusion, and by measuring activities of the key regulatory targets, Sga, Hpr, and MtdA. We observed no effect on transcription of the qsc1 promoter in mutants with lesions in genes for the serine cycle enzymes, glyA, sga, hpr, ppc, or mcl (Tables 3 and 4). Mutants with lesions in genes for the H4MPT-linked C1 transfer pathway (fae, dmrA, and mtdB) were also tested. Since those mutants are characterized by high sensitivity to methanol (10, 22, 27, 32), they were incubated with methanol for only 20 min. Increased expression of the qsc1 promoter was observed in these mutants. The levels of XylE in these mutants declined during longer incubation periods, presumably due to formaldehyde toxicity. Mutants with defects in the H4F-linked C1 transfer pathway were also analyzed. In the mtdA (CM275K.1) and fch (CM279K.1) mutants, low levels of XylE were measured, and these agreed with the activities of Sga and Hpr, which were also low in these mutants (Tables 3 and 4). The activity of MtdA was also lowered in the fch mutant. In contrast, XylE levels were increased fivefold compared to the wild-type levels in the ftfL mutant (CM216K.1). The activities of Sga, Hpr, and MtdA were also increased in this construct (Table 3). These data suggest that a metabolic signal regulating QscR must be produced in the H4F-linked C1 transfer pathway, and this intermediate likely accumulates in the ftfL mutant. Since the reaction catalyzed by FtfL is reversible, this intermediate could be either formyl-H4F or formate. Lower levels of XylE compared to wild-type levels were measured in the mutant defective in all three formate dehydrogenases, which is known to accumulate formate (12). This result argued against formate being a coinducer for QscR, making formyl-H4F the major candidate for this role.



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  		FIG. 4. Model for transcriptional regulation of the serine cycle operons in M. extorquens AM1, mediated by QscR and formyl-H4F. (A) Diagram showing the relevant methylotrophic pathways and enzymes; (B) model for QscR control of the known QscR-dependent genes.

 

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  		TABLE 3. Activities of Sga, Hpr, and MtdA in wild-type and mutant backgrounds

 

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  		TABLE 4. Activity of XylE from a Pqsc1-xylE transcription fusion in wild-type and mutant backgrounds

 
In order to verify this hypothesis we directly tested the effect of H4F-linked C1 transfer pathway intermediates on DNA-binding properties of QscR. Formaldehyde, methylene-H4F, methenyl-H4F, formyl-H4F and formate were included in gel retardation assays using the qsc1 promoter region (Pqsc1-1) and purified QscR. No significant effect on the DNA-binding activity of QscR was observed with formaldehyde, methylene-H4F or methenyl-H4F, while formate caused an inhibitory effect on binding (Fig. 5A). However, the addition of 5 µM formyl-H4F resulted in an approximately twofold increase in DNA binding by QscR, further supporting the hypothesis of formyl-H4F being the physiological co-inducer of QscR (Fig. 5A and B).



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  		FIG. 5. (A) Gel retardation of the qsc1 promoter fragment in the presence of purified QscR (0.5 mg) (lanes 2 through 8) and effectors in the following lanes (50 µM): 3, formate; 4, formaldehyde; 5, H4F; 6, H4F+formaldehyde; 7, methenyl-H4F; 8, formyl-H4F. (B) Gel retardation of the qsc1-2 promoter fragment in the presence of 0.5 mg purified QscR (lanes 2 through 6) and increasing amounts formyl-H4F is shown in the following lanes: 3, 5 µM; 4, 10 µM; 5, 20 µM; 6, 40 µM. Lane L contains the 32P-labeled 1-kb DNA ladder (Invitrogen). Lane 1 contains the qsc1 (A) or qsc1-2 (B) promoter fragments without QscR.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Most {alpha}-proteobacterial methylotrophs use the serine cycle as the principal pathway for assimilation of C1 units during methylotrophic growth. Physiological studies have shown that the expression of the serine cycle in facultative methylotrophic bacteria depends on the availability of C1 substrates. The discovery that QscR is a transcriptional regulator of the enzymes of the serine cycle in M. extorquens AM1 (16, 22) suggested that this protein conveys key physiological signals generated during growth on C1 compounds to the transcription apparatus. In order to analyze QscR-mediated control in more detail, we have carried out mapping of promoter and regulatory regions involved in QscR-mediated transcription.

We demonstrated that, like other LysR-type transcriptional regulators, QscR specifically binds to AT-rich regions near promoters of the genes subject to regulation. The QscR-binding site was determined in the region between -216 and –165 bp relative to the transcriptional start site of the qsc1 operon. For the qsc2 region, the DNase I protection region for QscR overlaps with the qsc2-TS2 transcriptional start site, suggesting that binding of QscR to this site might interfere with the expression from qsc2-TS2. In keeping with this hypothesis, preferential transcription was observed from the qsc2-TS1 transcription start site during methylotrophic growth. Thus, under methylotrophic growth conditions, expression of the qsc2 operon is likely to be predominantly regulated by QscR. The analysis of the promoter-regulatory region of glyA indicated the presence of two methanol-inducible transcriptional starts (glyA-1 and glyA-2). The glyA-1 start has a putative –35 region overlapping with the QscR binding site, suggesting that QscR might play a role in the relative use of these two starts.

We have demonstrated previously that QscR in M. extorquens AM1 acts as an autorepressor of its own expression (16). The 46-bp protection zone of QscR within the qscR promoter-operator region was shown to cover a portion of the QscR coding region, suggesting a mechanism by which QscR may interfere with its own expression.

Since the level of expression of qscR did not change during the shift from growth on a multicarbon substrate to methylotrophic growth (16), a coinducer molecule was hypothesized, to account for the increased expression of the serine cycle genes under these conditions. By assessing qsc1 promoter activity in various mutants of M. extorquens AM1, we identified formyl-H4F as the most prominent candidate for the role of the co-inducer, and we directly confirmed by gel retardation experiments that formyl-H4F is able to increase the DNA-binding activity of QscR. Based on these data and based on the recent data from flux analysis in M. extorquens AM1 (18), we present a working model for transcriptional regulation of the serine cycle in M. extorquens AM1 (Fig. 4). When the switch from a multicarbon growth mode to a methylotrophic growth mode first occurs, flux measurements show that the formaldehyde produced starts flowing through the H4MPT-linked C1 transfer pathway (18), resulting in production of ATP from the methanol oxidation and NAD(P)H from the methylene-H4MPT dehydrogenase reaction (Fig. 4A). We hypothesize that this flux indicates the energetic status of the cell, and the readiness for C1 assimilation. NADP pools should then drop, releasing QscR in a noninhibited form. Flux measurements show that under these conditions, very little formaldehyde is converted directly to methylene H4F, with the ratio of the direct conversion route to the H4MPT-dependent route being 8:1 (18). Flux measurements also show that formate is produced in the H4MPT-linked pathway (28), and although most of it is oxidized via formate dehydrogenases, resulting in production of additional NADH (12), a small portion (12%) is converted into formyl-H4F, via the FtfL reaction (Fig. 5A) (18, 21). FtfL is known to be present at high levels during growth on multicarbon substrates (21). Under these conditions, formyl-H4F would build up, activating QscR via direct binding, and the activated QscR (QscR*) would then bind to the regulated promoters, resulting in increased transcription of both the serine cycle genes and the H4F-linked C1 transfer pathway genes (Fig. 5B). When cells are actively growing on methanol, it has been shown that the ratio of methylene-H4F produced by the direct route to that produced by the H4MPT-dependent route is high (15:1), but even so, 6% of the total methanol carbon assimilated comes via the H4MPT-dependent route, demonstrating a continued flux through formyl-H4F under these conditions (18). QscR apparently measures the formyl H4F/formate ratio, which would provide a means of balancing the transcription of serine cycle genes with needs based on energy status. When the C1 substrate is depleted, the level of intracellular formyl-H4F drops and this leads to decreased transcription from both the serine cycle genes and the H4F-linked C1 transfer pathway genes. So far, this model agrees with the flux analysis data (18). Further testing of the model will rely on experiments, currently under way, to measure intracellular pools of intermediates specific to methylotrophic metabolism, as well as pools of redox metabolites.


   ACKNOWLEDGMENTS
 
We thank L. Chistoserdova, C. Marx, N. Korotkova, and T. Strovas for their thoughtful discussion of our work, M. Zhang for assistance in transcriptional start site mapping and DNase I footprint analysis, and C. Caperelli for providing us formyl-H4F.

This work was supported by the NIH (grant GM58933).


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Chemical Engineering, University of Washington, Seattle, WA 98195-1750. Phone: (206) 616-6954. Fax: (206) 616-5721. E-mail: mkalyuzh{at}u.washington.edu. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Anthony, C. 1982. The biochemistry of methylotrophs. Academic Press, Inc., London, United Kingdom.
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Journal of Bacteriology, November 2005, p. 7511-7517, Vol. 187, No. 21
0021-9193/05/$08.00+0     doi:10.1128/JB.187.21.7511-7517.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.



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