Evolutionary and Genomic Microbiology

Bacillus halodurans Strain C125 Encodes and Synthesizes Enzymes from Both Known Pathways To Form dUMP Directly from Cytosine Deoxyribonucleotides

Christian Berg Oehlenschlæger, Monika Nøhr Løvgreen, Eva Reinauer, Emilia Lehtinen, Marie-Louise Lindberg Pind, Pernille Harris, Jan Martinussen, Martin Willemoës
S.-J. Liu, Editor
DOI: 10.1128/AEM.00268-15

ABSTRACT

Analysis of the genome of Bacillus halodurans strain C125 indicated that two pathways leading from a cytosine deoxyribonucleotide to dUMP, used for dTMP synthesis, were encoded by the genome of the bacterium. The genes that were responsible, the comEB gene and the dcdB gene, encoding dCMP deaminase and the bifunctional dCTP deaminase:dUTPase (DCD:DUT), respectively, were both shown to be expressed in B. halodurans, and both genes were subject to repression by the nucleosides thymidine and deoxycytidine. The latter nucleoside presumably exerts its repression after deamination by cytidine deaminase. Both comEB and dcdB were cloned, overexpressed in Escherichia coli, and purified to homogeneity. Both enzymes were active and displayed the expected regulatory properties: activation by dCTP for dCMP deaminase and dTTP inhibition for both enzymes. Structurally, the B. halodurans enzyme resembled the Mycobacterium tuberculosis enzyme the most. An investigation of sequenced genomes from other species of the genus Bacillus revealed that not only the genome of B. halodurans but also the genomes of Bacillus pseudofirmus, Bacillus thuringiensis, Bacillus hemicellulosilyticus, Bacillus marmarensis, Bacillus cereus, and Bacillus megaterium encode both the dCMP deaminase and the DCD:DUT enzymes. In addition, eight dcdB homologs from Bacillus species within the genus for which the whole genome has not yet been sequenced were registered in the NCBI Entrez database.

INTRODUCTION

The biosynthetic route to the nucleotide dUMP, the precursor of dTTP, as shown in Fig. 1, differs between organisms and sets it apart from the rest of nucleotide metabolism (1). This is in agreement with the general belief that uracil was the original DNA base-pairing partner for adenine and that the biosynthesis of dTTP from dUMP arose later in evolution (2). The deoxyribonucleotides dGTP, dATP, and dCTP are all produced in a pathway that goes through the multisubstrate enzyme ribonucleotide reductase. This enzyme is highly regulated in order to maintain the balance of deoxyribonucleotide pools (3–5). dUTP is also formed by ribonucleotide reductase, but because of the toxicity of this nucleotide (6, 7), all organisms express an enzyme with dUTPase activity that hydrolyzes dUTP to dUMP (8).

FIG 1
FIG 1

Aerobic deoxyribonucleotide metabolism, including the regulatory patterns (dotted lines). + and −, activation and inhibition, respectively. The alternative pathways from dCTP/dCMP to dTTP are shown using bold arrows, and both the dcdB- and comEB-dependent pathways are present in B. halodurans. Gene designations are as follows: nrdAB, aerobic ribonucleotide reductase; ndk, nucleoside diphosphate kinase; dcd, monofunctional dCTP deaminase; dut, dUTPase; dcdB, DCD:DUT; comEB, dCMP deaminase; thyA, thymidylate synthase; thyX, NADPH-dependent thymidylate synthase; tdk, thymidine kinase; cdd, cytidine deaminase; tmk, dTMP kinase.

The direct formation of dUMP via ribonucleotide reductase (Fig. 1) is not an efficient process, as the ribonucleotide reductase has a reduced affinity for UDP compared to the affinities of three other canonical ribonucleotides (9–12). Instead, most of the dUMP arises from deamination of cytosine deoxyribonucleotides (13–15). Enteric bacteria synthesize an enzyme with dCTP deaminase activity that converts dCTP into dUTP (EC 3.5.4.13), which accounts for 70 to 80% of the dUMP production (15). The dCTP deaminase is a homotrimeric enzyme structurally related to the trimeric dUTPase and part of the dCTP deaminase/dUTPase superfamily (16–18). The genomes of many Gram-positive bacteria and all eukaryotes encode a dCMP deaminase (EC 3.5.4.12) that shares no resemblance to the enteric dCTP deaminase (14, 19–23). The dCTP deaminase catalyzes dCTP deamination solely by way of amino acid residues present in the active site (16, 18), while the dCMP deaminase, a hexameric enzyme which is related to other cytosine deaminating enzymes, such as cytosine deaminase (24) and cytidine deaminase (25–29), binds a zinc ion that directly participates in the catalytic mechanism (14, 19, 20, 30). One feature shared by the two enzymes, dCTP and dCMP deaminase, is that they are both feedback inhibited by dTTP (17, 19, 31–38). In addition, the dCMP deaminase is allosterically activated by dCTP (19, 33, 36), the substrate of the dCTP deaminase.

Within the last decade, yet another enzyme producing dUMP, an enzyme with dual activity that is able to both deaminate dCTP and hydrolyze dUTP to dUMP without releasing the toxic dUTP intermediate, has been revealed. The first enzyme, dCTP deaminase:dUTPase (DCD:DUT; EC 3.5.4.30), discovered in the archaeon Methanocaldococcus jannaschii (39, 40), constitutes the third member of the dCTP deaminase/trimeric dUTPase superfamily of enzymes. This enzyme is structurally most closely related to the dCTP deaminase from enteric bacteria (41, 42). Later, the genome of Mycobacterium tuberculosis was also shown to encode a DCD:DUT activity (43), extending the presence of the bifunctional enzyme to bacteria.

Analysis of the genome of the Gram-positive bacterium Bacillus halodurans led to the surprising finding that in addition to the comEB gene encoding the dCMP deaminase, as characterized in Bacillus subtilis (14), B. halodurans also possesses a gene encoding a putative dCTP deaminase. In this work, we show that both the comEB genes and the gene from enteric bacteria homologous to dcd encode functional enzymes that are expressed in vivo and that the dcd homolog encodes a bifunctional DCD:DUT. Here we designate the DCD:DUT-encoding gene dcdB, which corresponds to the open reading frame BH0368 in the B. halodurans genome sequence. This is to avoid confusion with the dcd gene encoding the monofunctional dCTP deaminase (44).

MATERIALS AND METHODS

Growth conditions for B. halodurans.B. halodurans strain C125 was grown at 37°C with constant longitudinal shaking with 240 movements per minute in modified LB medium (4 g/liter sodium chloride, 10 g/liter peptone, 5 g/liter yeast extract, pH 8.5, supplied with 100 mM sodium carbonate) or a chemically defined medium originally developed for B. subtilis (45): 0.38% glucose, 0,19% glutamate, 0.19 mg/liter manganese(II) sulfate, 3.4 mg/liter ferric chloride, 9.5 mg/liter thiamine, 38.0 mg/liter tryptophan, 70 g/liter dipotassium phosphate trihydrate, 30 g/liter monopotassium phosphate, 5 g/liter trisodium citrate, 1 g/liter magnesium sulfate heptahydrate, and 10 g/liter sodium sulfate adjusted to pH 8.5. Thymidine or deoxycytidine was added to 100 μg/ml, when indicated. Bacterial growth was monitored at 600 nm.

DNA and RNA methods.For the isolation of chromosomal DNA, an overnight culture of B. halodurans strain C125 grown in modified LB medium was harvested by centrifugation (10,000 × g, 5 min). The pellet was resuspended in 270 μl TNEX (10 mM Tris, pH 8.0, 10 mM NaCl, 10 mM EDTA, 1% Triton X-100). A 30-µl volume of freshly made lysozyme solution (5 mg/ml in water) was added, and the cell suspension was incubated for 2 h at 37°C, at which time 15 μl proteinase K (20 mg/ml in water) was added and incubation at 37°C was continued for 2 h, followed by an additional 2 h at 65°C to inactivate the protease. The following steps were performed at 4°C. The DNA was precipitated by adding 1/20 volume of 5 M sodium chloride and 3 volumes of 96% ethanol. The ethanol was removed by repeated centrifugation (10,000 × g, 5 min), and the DNA was resuspended in 100 to 500 μl TE (10 mM Tris, pH 8.0, 10 mM sodium chloride, 10 mM EDTA) and stored at −20°C.

RNA was extracted from cells grown in minimal medium. Samples corresponding to approximately 2 × 108 cells were immediately chilled to 0°C in prechilled tubes, followed by centrifugation (6,500 rpm, 10 min, 0°C). Pellets were stored at −80°C. Total RNA was extracted using a NucleoSpin RNA II system (Macherey-Nagel) according to the supplier's instructions. Briefly, cells were resuspended in 350 μl RA1 buffer and homogenized with glass beads using a Savant Bio101 FP120 FastPrep instrument. The lysate was applied to the NucleoSpin RNA II column, and residual DNA was removed by DNase. RNA was subsequently eluted with 40 μl of RNase-free H2O. The integrity of the DNA and RNA was verified by agarose gel electrophoresis and subsequent ethidium bromide staining. The concentrations of nucleic acids were determined by measurement of the UV absorption at 260 nm.

cDNA was synthesized from total RNA (185 ng) by random primers using an AffinityScript QPCR cDNA synthesis kit (Stratagene) by incubation at 25°C for 5 min, 42°C for 30 min, and 95°C for 5 min according to the supplier's instructions.

Cloning of the B. halodurans reading frames of comEB and dcdB in pET11a for heterologous expression in Escherichia coli.Genomic DNA from B. halodurans strain C125, prepared as described above, served as a template for PCR, where the primers comEB5′ (5′-GGAATTCCATATGAATCGAATTTCTTGGG-3′) and comEB3′ (5′-CCGGGATCCTTAAGAAGATGTATAAAGACGG-3′) in combination yielded a PCR product harboring the reading frame for B. halodurans comEB. The NdeI and BamHI restriction endonuclease sites (indicated in the primers in italics) were used for cloning into the expression vector pET11a by ligation of the NdeI- and BamHI-digested PCR product and vector. Similarly, a PCR product containing the B. halodurans dcdB reading frame was cloned into the same vector using the same endonuclease restriction sites mentioned above and also shown in italics in the sequences for primers dcd5′ (5′-GGAATTCCATATGATTTTAAGTGGAAAACCC-3′) and dcd3′ (5′-CCGGGATCCTTAAAACGCGTCTTTGTAAATTTCGC-3′). The inserts of both plasmids, pET11a:comEB and pET11a:dcdB, were verified by sequencing and subsequent comparison with the sequences of B. halodurans comEB and dcdB (BH0368) deposited in GenBank.

Enzyme purification. (i) DCD:DUT.E. coli BL21(DE3) cells transformed with pET11a:dcdB, described above, were grown at 37°C in 1 liter of LB medium supplemented with ampicillin (100 mg/ml) to an optical density at 436 nm (OD436) of 1, and expression of dcdB was induced by addition of 1 mM IPTG (isopropyl-β-d-thiogalactopyranoside). Cells were vigorously shaken at 250 rpm overnight and harvested by centrifugation with a GSA rotor in a Sorvall centrifuge (8,000 rpm, 20 min) at 4°C. All subsequent steps were carried out at 4°C, unless otherwise stated. The cells were thawed, suspended in 50 mM potassium phosphate buffer, pH 6.8, and opened while cooled on ice by sonication using a Bandelin Sonopuls sonicator (Germany) at 35% maximum effect for 20 cycles with a 30-s pulse and a 30-s rest. The cell debris was collected by centrifugation at 10,000 rpm for 20 min using an SS34 rotor in a Sorvall centrifuge. Streptomycin sulfate solution (10%, wt/vol) was added dropwise to the supernatant to a final concentration of 1%, and the mixture was stirred for 45 min to precipitate the nucleic acids. The precipitate was collected by centrifugation as described above. Solid ammonium sulfate was added to the supernatant to a final saturation of 18%. The precipitate was collected by centrifugation as described above and dissolved in 13 ml 50 mM potassium phosphate buffer, pH 6.8. The protein solution was dialyzed against 500 ml 50 mM potassium phosphate buffer, pH 6.8, overnight.

The dialyzed solution was loaded onto a DE-52 anion-exchange column (1.4 by 29 cm) which had previously been equilibrated with 50 mM potassium phosphate buffer, pH 6.8. The column was washed with 60 ml 50 mM potassium phosphate buffer, pH 6.8, before elution was performed with 160 ml of a linear NaCl gradient from 0 to 0.4 M in 50 mM potassium phosphate buffer, pH 6.8. Fractions of 10 ml were collected with a GradiFrac system (Amersham Bioscience). Anion-exchange chromatography was carried out at room temperature without any loss of enzyme activity. Fractions containing the B. halodurans DCD:DUT were pooled, and solid ammonium sulfate was added to a saturation of 19%. The precipitate was collected by centrifugation as described above and dissolved in 6.5 ml 20 mM HEPES, pH 6.8. Finally, the protein solution was dialyzed two times against 500 ml 20 mM HEPES, pH 6.8. The protein concentration was estimated by measuring the absorbance at 280 nm using the theoretical extinction coefficient at this wavelength (16,960 M−1 cm−1), calculated from the deduced protein sequence under the assumption that all cysteine residues are reduced. The enzyme was stored in aliquots at −21°C until use.

(ii) dCMP deaminase.E. coli BL21(DE3) cells transformed with pET11a:comEB were grown, induced, and harvested as described above for the dcdB-producing cells. All steps were carried out at 4°C unless otherwise stated. The cells were suspended in 20 ml 1 mM dithiothreitol (DTT), 20 mM Tris-HCl, pH 7.5, and opened by sonication. Streptomycin precipitation was carried out as described above. However, 1% streptomycin also precipitated the B. halodurans dCMP deaminase, so the pellet was resuspended in 20 ml 1 mM DTT, 50 mM potassium phosphate buffer, pH 6.8, and insoluble matter was removed by centrifugation (10,000 rpm, 30 min) in an SS34 rotor. The supernatant was dialyzed against 500 ml 1 mM DTT, 20 mM Tris-HCl, pH 7.5, overnight. Anion-exchange chromatography was carried out as described above, except that 1 mM DTT, 20 mM Tris-HCl, pH 7.5, was used as the buffer. Fractions containing the B. halodurans dCMP deaminase were pooled. Solid ammonium sulfate was added to the pooled fractions to a saturation of 46%. The precipitate was collected by centrifugation in an SS34 rotor as described above, dissolved in 5 ml 1 mM DTT, 20 mM Tris-HCl, pH 7.5, and dialyzed twice against 500 ml 1 mM DTT, 20 mM Tris-HCl, pH 7.5. The protein concentration was estimated as described above using the extinction coefficient 20,400 M−1 cm−1. The enzyme was stored in aliquots at −21°C until use.

Assays of enzyme activity and kinetic data analysis.Enzyme assays were performed in one of two buffers, 20 mM HEPES, pH 6.8, or 20 mM Tris-HCl, pH 7.5, as specified in Results. Both buffers were supplemented with 1 mM DTT and 2 mM MgCl2. Nucleotide concentrations varied, and the specific concentrations are indicated in Results. Assays based on the conversion of [5-3H]dCTP to [5-3H]dUMP were performed as described elsewhere (39). Briefly, samples from different time points were quenched by addition of 0.1 M formic acid and subjected to thin-layer chromatography on polyethyleneimine cellulose plates after adding a mix of nonradioactively labeled dCTP, dUTP, dCMP, and dUMP at 5 nmol each and developed in 0.9 M acetic acid, 0.1 M LiCl. Spots identified by markers corresponding to dCTP and dUMP were cut out and quantified by scintillation counting. Initial rates of dCTP and dCMP deamination, based on the linear portion of the change in the absorbance at 291 nm, were determined in a Zeiss S10 Specord instrument by scanning from 280 nm to 300 nm using an extinction coefficient (Δε291) of 1,335 M−1 cm−1 for the conversion of a cytosine to a uracil nucleotide (46).

Initial rates obtained from experiments with various substrates in the absence or presence of inhibitor were analyzed using equation 1, which describes cooperative substrate saturation. Equation 2 was used for analyzing partial cooperative inhibition, and equation 3 was used for analyzing cooperative inhibition. rate=kcat·[S]nHS0.5nH+[S]nH(1) rate=rateuninh1+([I]I0.5)nH+rateoffset(2) rate=rateuninh1+([I]I0.5)nH(3) where rate is the initial rate, kcat is the maximal rate, S0.5 is the concentration of the substrate (S) for a half-maximal rate, and nH is the Hill coefficient. When activation rather than substrate saturation was determined, the concentration of activator for a half-maximal rate, A0.5, replaced S0.5 in equation 1. Rateuninh. is the rate in the absence of the inhibitor (I), I0.5 is the concentration of I for half-maximal inhibition, and rateoffset is the residual activity of the inhibited enzyme in the presence of an infinitely high inhibitor concentration.

Analysis of dcdB and comEB expression using qPCR.Quantitative PCR (qPCR) was carried out in 25-μl reaction mixtures using a Brilliant II SYBR green QRT-PCR master mix (Stratagene) according to the supplier's instructions. Reactions were performed in triplicate and run on an Mx3005P real-time PCR system (Stratagene) with 300 μM primers, 30 μM reference dye, and 2 μl cDNA. For no-template controls, 2 μl H2O replaced cDNA. To check for DNA contamination, a control reaction with no reverse transcriptase was performed, where RNA replaced cDNA in amounts similar to those originally used for cDNA synthesis. To verify the specificity of the synthesized DNA fragments, melting curves were determined for each reaction by collecting fluorescence data continuously on a 55 to 95°C ramp (0.2°C/s). The rpsJ gene (30S ribosomal protein S10) was used as a reference. The primers used are listed in Table 1.

View this table:
TABLE 1

Primers used for qPCR quantification of gene expression in B. halodurans strain C125

MxPro QPCR software was used to run the qPCR, and the amplification plot method, described in the Microsoft Excel workbook entitled Data Analysis for Real-Time PCR (DART-PCR) (47), was used to determine the results as the amplification efficiency (E value), the quantification cycle value, and the level of expression (R0) for each reaction. This method uses a simple algorithm to calculate the amplification efficiency for every sample individually after each qPCR assay. After normalization of R0 to the R0s for the reference genes, the expression levels relative to the expression level for deoxycytidine-supplemented medium were determined. The mean and standard deviation for three biological replicates were determined. The qPCR nomenclature used here is as described previously (48).

Crystallization of B. halodurans DCD:DUT with dTTP.Initial screening for crystallization conditions was carried out with an Ozma 4K screen from Jena Bioscience by mixing 2 μl protein-TTP solution (2.7 mg/ml B. halodurans DCD:DUT in 19 mM HEPES, pH 6.8, 20 mM MgCl2, 5 mM dTTP) with 1 μl reservoir solution, equilibrated over 500 μl reservoir solution incubated at room temperature. Single crystals were obtained in solution 17 containing 200 mM lithium citrate and 20% polyethylene glycol (PEG) 4000. The crystal used for data collection and structure determination was grown at room temperature in a hanging drop produced by mixing 2 μl protein-TTP solution with 1 μl reservoir solution containing 150 mM lithium citrate and 20% PEG 4000 equilibrated over 1 ml reservoir solution. Under these conditions crystals grew to a size of approximately 150 by 50 by 50 μm within a few days. Crystals were harvested and stored in liquid nitrogen until data collection.

Diffraction data collection and processing.Diffraction data were collected at the MAX-lab I911-2 beam line in Lund, Sweden. The diffraction data were indexed, integrated, scaled, and merged by use of the XDS software package (49). Details on the data collection statistics are provided in Table 2.

View this table:
TABLE 2

Data collection and refinement statistics

Protein structure determination and refinement.The crystal structure of the B. halodurans dCTP deaminase-dUTPase in complex with dTTP was solved by molecular replacement using the MOLREP program (50) in the CCP4 suite (51). A trimer generated from the A chain of the Mycobacterium tuberculosis dCTP deaminase-dUTPase structure in complex with dTTP (PDB accession number 2QXX) was used as the search model (43). A solution in space group P21 with two trimers in the asymmetric unit was obtained. Refinement was carried out in the REFMAC5 program (52) with rigid body refinement, followed by restrained refinements and manual corrections in the WinCoot program (53). Automatic local noncrystallographic symmetry (NCS) restraints were imposed. Residues 1 to 176 were modeled in all chains except F, where no electron density was seen for residues 44 to 48. Electron density for the final residue, residue 177, was visible only in some chains. The density for dTTP and Mg2+ was very clear in all chains. In total, 388 water molecules and 1 PEG molecule were modeled. Refinement statistics are provided in Table 2.

The final model was checked using the pdb_redo web server (54), which improved the structure somewhat, and was validated here using the Structure Analysis and Verification Server (http://services.mbi.ucla.edu/SAVES/). The Ramachandran plot contains 99.4% residues in the most favored regions and 0.6% in the additional allowed regions.

Protein structure accession number.The coordinates and structure factors have been deposited in the Protein Data Bank (PDB) under accession number 4XJC.

RESULTS

Enzyme kinetic analysis of DCD:DUT and dCMP deaminase from B. halodurans.To classify the product of the B. halodurans dcdB gene, identical to the BH0368 open reading frame, as either a monofunctional dCTP deaminase or a bifunctional enzyme, we performed the experiments whose results are shown in Fig. 2. The results demonstrated that the formation of [5-3H]dUMP from [5-3H]dCTP (Fig. 2A) occurred at a rate identical to that for the deamination of dCTP, as measured by determination of the change in the absorbance at 291 nm (Fig. 2B). This observation allows us to conclude that the dcdB gene product of B. halodurans is in fact a bifunctional enzyme, DCD:DUT, similar to that found in archaea and mycobacteria (39–41, 43).

FIG 2
FIG 2

Analysis of the B. halodurans DCD:DUT reaction. Progress curves of[5-3H]dUMP formation from [5-3H]dCTP (A) and deamination of dCTP (B) are shown. Assays were performed as described in Materials and Methods in the presence 0.4 mM [5-3H]dCTP or 0.4 mM dCTP, both at pH 6.8. The enzyme concentrations were 3.4 μM (closed circles) and 1.7 μM (open circles).

Both the B. halodurans dCMP deaminase and DCD:DUT behaved kinetically like previously published enzymes from these two families of nucleotide deaminases (14, 34, 36). Both enzymes showed sigmoid saturation curves for their substrates (Fig. 3A and B), and the dCMP deaminase was activated by dCTP and inhibited by dTTP (Fig. 3C and D), as is common for this enzyme from various organisms (21, 38, 55, 56). Likewise, DCD:DUT was inhibited by dTTP (Fig. 3C), as was previously found for both the monofunctional and the bifunctional deaminase enzymes in this family (17, 43). The results in Fig. 3C and D, showing the kinetics of the dCMP deaminase and DCD:DUT enzymes under close to physiological conditions with respect to pH and nucleotide concentrations (20 μM for dCMP [14, 57]), indicate that both enzymes responded similarly in terms of activity with variations in dTTP and dCTP concentrations. The kinetic analysis verified that both the dCMP deaminase and the DCD:DUT enzymes of B. halodurans are competent, having all the catalytic and regulatory properties needed for producing dUMP at a physiological pH of about 7.5, and that they are regulated in response to changes in the concentration of dCTP and dTTP (Fig. 3C and D).

FIG 3
FIG 3

Substrate saturation and inhibition of B. halodurans dCMP deaminase and DCD:DUT by dTTP. Assays were performed as described in Materials and Methods by determining the change in absorption at 291 nm over time. (A) Saturation of dCMP deaminase with dCMP in the presence of 100 μM dCTP at pH 7.5. Data were fitted to equation 1. The kinetic parameters were as follows: S0.5 = 0.22 ± 0.03 mM, kcat = 1.9 ± 0.1 s−1, and nH = 1.27 ± 0.08. (B) Saturation of DCD:DUT with dCTP at pH 6.8. Data were fitted to equation 1. The kinetic parameters were as follows: S0.5 = 0.047 ± 0.003 mM, kcat = 0.28 ± 0.01 s−1, and nH = 2.3 ± 0.3. (C) Inhibition by dTTP at pH 7.5 of dCMP deaminase in the presence of 100 μM dCMP and 100 μM dCTP (open circles) or 20 μM dCMP and 100 μM dCTP (triangles) and DCD:DUT (closed circles) in the presence of 100 μM dCTP and 100 μM dCMP. Data were fitted to equation 2 (open circles, triangles) or equation 3 (closed circles). The kinetic parameters were as follows: rateuninh = 0.20 ± 0.1 s−1, rateoffset = 0.12 ± 0.01 s−1, I0.5 = 0.012 ± 0.001 mM, and nH = 1.8 ± 0.3 (open circles); rateuninh = 0.09 ± 0.01 s−1, rateoffset = 0.052 ± 0.009 s−1, I0.5 = 0.008 ± 0.002 mM, and nH = 2.0 ± 1.0 (triangles); and rateuninh = 0.208 ± 0.009 s−1, I0.5 = 0.030 ± 0.003 mM, and nH = 1.6 ± 0.2 (closed circles). (D) Activation by dCTP at pH 7.5 of dCMP deaminase in the presence of 100 μM dCMP (open circles) or 20 μM dCMP (triangles) and saturation of DCD:DUT with dCTP in the presence of 100 μM dCMP (closed circles). Data were fitted to equation 1. The kinetic parameters were as follows: A0.5 = 0.07 ± 0.02 mM, kcat = 0.73 ± 0.01 s−1, and nH = 1.1 ± 0.1 (open circles); A0.5 = 0.07 ± 0.04 mM, kcat = 0.18 ± 0.04 s−1, and nH = 1.0 ± 0.2 (triangles); and S0.5 = 0.025 ± 0.003 mM, kcat = 0.20 ± 0.01 s−1, and nH = 1.8 ± 0.3 (closed circles).

Confirming the expression of the B. halodurans dcdB and comEB genes encoding DCD:DUT and dCMP deaminase.In order to investigate whether the pathways involving the dcdB and comEB gene products are indeed active in B. halodurans and whether the expression of the two genes is subject to regulation at the genetic level, the relative amount of the corresponding mRNAs from cells grown in the absence or presence of thymidine or deoxycytidine was quantified by real-time PCR. The presence of deoxycytidine may increase the dCMP pool size and thereby facilitate the flux through the dCMP deaminase reaction to dUMP. If dTMP is obtained directly from thymidine, the formation of dUMP is redundant; it could be speculated that the expression of dcdB and comEB is repressed in the presence of exogenous thymidine. To investigate these possibilities, B. halodurans was grown in minimal medium in the absence and presence of deoxycytidine and thymidine, and RNA was extracted at different times in the exponential growth phase. To quantify gene expression, the isolated mRNA was used as the template in cDNA synthesis, and the cDNA was subsequently analyzed by real-time PCR. Three samples were taken during the exponential growth phase (OD600, between 0.420 and 0.735), and the total RNA was extracted and used as the template in reverse transcription-PCR experiments. We estimated the amounts of mRNA synthesized from dcdB and comEB relative to the amount of mRNA synthesized from rpsJ (30S ribosomal protein S10), as rpsJ was found to be expressed at a constant level during exponential growth (data not shown). The rpsJ gene has previously been used as a reference gene for B. subtilis (58). The fold change in the level of dcdB and comEB repression caused by thymidylate precursors is shown in Fig. 4, and on the basis of the findings, it can be concluded that both genes are indeed transcribed, as the signals were significantly above the background (data not shown). The fact both proteins are active enzymes strongly suggests that both pathways are available for dTTP biosynthesis. Moreover, expression of both genes seems to be repressed to the same degree by both deoxycytidine and thymidine. The large errors in Fig. 4 reflect the low expression levels of both genes (59). Repression of gene expression by thymidine makes physiological sense, since both pathways to dUMP are redundant when thymidine is available (Fig. 1). The gene repression in response to deoxycytidine in the medium may indicate that the dominating metabolism of deoxycytidine taken up by the cell is not phosphorylation to dCMP but deamination to deoxyuridine and subsequent phosphorylation to dUMP (Fig. 1). The genes for both (deoxy)cytidine deaminase (CDD; BH1366) and thymidine/deoxyuridine kinase (TDK; BH3779), necessary for the conversion of deoxycytidine to dUMP, can be identified in the B. halodurans genome. Thus, the presence of either thymidine or deoxycytidine in the medium obviates the need for the dcdB and comEB gene products.

FIG 4
FIG 4

Induction of comEB and dcdB gene expression in the absence of exogenous thymidylate precursors. Cells were grown in minimal medium (MM) in the absence and presence of either thymidine (TdR) or deoxycytidine (CdR). The mRNA encoding dCMP deaminase (comEB; dark gray) and dCTP deaminase (dcdB; light gray) was quantified relative to the level of expression of rpsJ, and the resulting expression levels were compared to the expression levels determined in cells growing in the presence of deoxycytidine. The data for each condition are based on three independently isolated RNA samples from exponentially growing cultures and are presented as the means of the three determinations. The calculated standard deviations are shown as vertical lines.

A brief search of the genus Bacillus in the NCBI Entrez database revealed six species, Bacillus pseudofirmus, Bacillus thuringiensis, Bacillus hemicellulosilyticus, Bacillus marmarensis, Bacillus cereus, and Bacillus megaterium, that, like B. halodurans, encode both a dCMP deaminase (comEB) and a DCD:DUT (dcdB). However, apart from the above-mentioned genus members, Bacillus okhensis, Bacillus simplex, Bacillus selenitireducens, Bacillus methanolicus, Bacillus ginsengihumi, Bacillus aurantiacus, Bacillus aidingensis, and Bacillus flexus all seem to have the dcdB gene, but we could not confirm the existence of a comEB gene or a dCMP deaminase homolog encoded by the genomes of these species.

Crystal structure of B. halodurans DCD:DUT.The crystal structure of B. halodurans DCD:DUT in complex with Mg-dTTP was solved by molecular replacement to a resolution of 2.35 Å. The asymmetric unit contains 6 molecules arranged in two trimers. The homotrimeric form is a common feature of the enzyme family (18). The different subunits align to each other with root mean square deviations (RMSDs) ranging from 0.169 Å (subunits A and B) to 0.309 Å (subunits A and C).

The overall fold of a single subunit is mainly composed of β strands (Fig. 5A). Each subunit contains 14 β strands (β1 to β14), two α helices (α1 and α2), and two small 310 helices (γ1 and γ2). The β strands form four separate antiparallel β sheets: S1, consisting of β1, β7, and β11; S2, consisting of β2, β14, β8, and β19; S3, consisting of β9, β12, β6, β3, and β4; and S4, consisting of β13 and β5. Together these sheets form a distorted β barrel.

FIG 5
FIG 5

(A) Overall fold of the DCD:DUT from B. halodurans. The coloring progresses from the N terminus (blue) to the C terminus (red). (B) Binding of dTTP is clearly seen in a 2Fo − Fc omit map contoured at a level of 0.8σ. (C) Schematic view of the hydrogen bonds to dTTP in DCD:DUT from B. halodurans. (D) A closeup on the nucleotide binding site, where all structures are superposed. Blue, the E. coli dCTP deaminase monomer (PDB accession number 1XS1; dUTP bound); purple, the M. jannaschii DCD:DUT monomer (PDB accession number 2HXD; E145A variant, dUMP NP bound); green, the M. tuberculosis DCD:DUT monomer (PDB accession number 2QXX; dTTP bound); yellow, the S. tokodaii putative DCD:DUT monomer (PDB accession number 2YZJ; dUDP bound); red, the B. halodurans DCD:DUT monomer (PDB accession number 4XJC; dTTP bound). Panels A, B, and D were prepared using the PyMOL molecular graphics system.

The electron density of Mg-dTTP is very clear (Fig. 5B) and reveals the binding site situated in the cleft between two subunits, which is common for the enzyme family, and as such there are three binding sites per trimer. The C terminus from one of the subunits forming the cleft folds down upon the binding site, which almost completely shields the inhibitor from solvent contact. The contacts between the protein and dTTP (Fig. 5C) are highly similar to those seen in the crystal structure of the M. tuberculosis DCD:DUT in complex with Mg-dTTP. Briefly, the bottom of the binding cleft is upholstered with hydrophobic residues on which the nucleoside portion of dTTP rests. The thymine moiety interacts with Val95, Val114, and Ile122, while the deoxyribosyl is aligned against Phe118. In addition to the hydrophobic interactions, the nucleotide portion is also bound to the protein through hydrogen bonds. The O-4 of the thymine moiety forms hydrogen bonds with the backbone carbonyl oxygen and N-D2 of Asn110 and the backbone carbonyl oxygen of Thr123. N-3 donates a hydrogen bond to the carbonyl oxygen of Thr123. O-2 forms hydrogen bonds with the NH backbone group of Thr123, NH2 of Arg103* (the asterisk indicates residues from a neighboring subunit), and N-E2 of Gln164. The 3′-hydroxy group on the deoxyribosyl forms a hydrogen bond with the backbone amide NH group and O-D2 of the catalytic Asp115. The triphosphoryl moiety of dTTP is bound through an extensive hydrogen bonding network involving several water molecules and residues, especially from the C terminus. A magnesium ion is also present in the structure, and it is coordinated octahedrally by three oxygen atoms from the phosphate groups (O-1A, O-2B, and O-3G) and three water molecules (Fig. 5C). Of the three phosphate groups, the α-phosphate engages in most hydrogen bonds. O-1A interacts with three water molecules, two of which are also coordinated to the Mg2+ ion. The third water molecule occupies a position similar to that for the catalytic water molecule observed in structures with dUTPase activity. O-2A is hydrogen bonded to one water molecule, to N-E2 of Gln144*, and to the backbone amide NH group of Ser99*. In the β-phosphate group, O-1B interacts with the side chain of Ser100*, while O-2B, besides interacting with Mg2+ and two of the Mg-coordinated water molecules, interacts with N-E of Arg98*. In the γ-phosphate group, O-2G forms hydrogen bonds with Tyr157 and the backbone amide NH group of Lys160; both of these residues are from the C terminus. O-3G interacts only with Mg2+ and two of the Mg-coordinated waters. O-1G is hydrogen bonded to Lys160, Arg98*, and a single water molecule.

As mentioned above, the C terminus completely covers the binding cleft. While Tyr157 and Lys160 are involved in direct interactions with the inhibitor, other residues of the C terminus are involved in indirect inhibitor binding and make key interactions that are responsible for ordering of the flexible C terminus of the apo-enzyme upon inhibitor binding. Asp175 forms a salt bridge with Arg98*, while Phe177 stacks with Arg141* and interacts with the inhibitor through a water molecule. These contacts are possible only when the inhibitor (or substrate) is bound.

Structural comparison of B. halodurans DCD:DUT to other members of the dCTP deaminase family.The structures of enzymes with dCTP deaminase activity are now available for 7 different organisms, including the two Archaea Methanocaldococcus jannaschii and Sulfolobus tokodaii, the three Gram-negative bacteria Escherichia coli, Anaplasma phagocytophilum, and Burkholderia thailandensis, and the Gram-positive bacteria B. halodurans and M. tuberculosis. A comparison of the B. halodurans DCD:DUT crystal structure to that of other available structures from other organisms was performed together with structure-based pairwise sequence alignments, and the results are shown in Table 3.

View this table:
TABLE 3

Comparison of crystal structures of DCD:DUT from B. halodurans and dCTP deaminases and bifunctional DCD:DUT enzymes from different organisms

The conservation of the overall fold between the different organisms is apparent (Table 3; Fig. 5D), and striking differences are mainly seen in flexible loop regions. The overall RMSD between the C-α atoms in the various structures was calculated using the SUPERPOSE program (60) in the CCP4 program suite (51), and striking differences are mainly seen in flexible loop regions. The most notable difference is seen in the M. jannaschii DCD:DUT structure, which contains an additional α helix and β strand inserted between β strands 5 and 6 compared to the B. halodurans structure. The additional β strand is aligned with β strand 5 and is part of the S4 β sheet. Despite this drastic difference between the two structures, they align very well with a low RMSD of 0.565 Å and a sequence identity of 28.5%. The fit between B. halodurans and S. tokodaii is particularly poor, with an RMSD of 1.594 Å, which probably reflects the low sequence identity of only 19.4%.

The alignment to the E. coli dCTP deaminase structure is intermediate with an RMSD of 0.849 Å, even though the sequence identity at 32.8% is higher than that for the M. jannaschii enzyme. The most notable difference is that the E. coli enzyme contains additional secondary structure elements in the form of a 310 helix and an α helix in the loop between β sheets 3 and 4.

The structures from A. phagocytophilum and B. thailandensis align very well to each other with an RMSD of 0.589 Å, which is not surprising, since the identity between these sequences is 55.9%. They generally contain the same structural elements as the B. halodurans enzyme; the only major difference is an insertion of 9 and 14 extra residues, respectively, between β strands 1 and 2 replacing the 310 helix (γ1) of the B. halodurans enzyme by extended loops. They do, however, align poorly with the B. halodurans structure, with RMSD values of 1.152 and 1.334 Å, respectively, and sequence identities of 17.6 and 18.7%, respectively.

In conclusion, the enzyme that structurally resembles the B. halodurans enzyme the most is that of M. tuberculosis. The RMSD is 0.733 Å, and even though the RMSD is lower for the M. jannaschii structure than the M. tuberculosis structure, the M. tuberculosis structure does not contain any large additional structural elements.

DISCUSSION

For the first time, we show that an organism can possess active enzymes that catalyze both of the two known pathways that form dUMP from cytosine deoxyribonucleotides. B. halodurans can generate dUMP from dUTP directly or from either dCTP or dCMP (Fig. 1). Although the flexibility in encompassing both pathways could somehow be beneficial to the bacterium, it is not obvious why both pathways are present. The regulation of the two genes comEB and dcdB responds similarly to the presence of the nucleosides thymidine and deoxycytidine in the medium (Fig. 4), and in principle, the gene products DCD:DUT and dCMP deaminase respond in the same way (Fig. 3) to dCTP (as a substrate and an activator, respectively) and dTTP (an inhibitor), with the dTTP nucleotide being the ultimate product of the pathway. Construction of dcdB and/or comEB mutant strains of B. halodurans is unlikely to reveal any new information about why both genes are present in this bacterium; the genes seem to be dispensable, and no immediate phenotype is found for comEB in B. subtilis (14) or dcd in E. coli (44), except that dcd strains were recently shown to have a mutator phenotype (61).

This is not the only case of alternative enzyme activities in dTMP synthesis. The thyA and thyX gene-encoded thymidylate synthases are found separately and in combination in both Bacteria and Archaea (62, 63), and both catalyze the same reaction, but the reductive methylation of dUMP at the 5 position is facilitated by oxidation of either N5,N10-methylenetetrahydrofolate to dihydrofolate (the thyA gene product) or NADPH to NADP+ (the thyX gene product).

Alternative pathways for the formation of dUMP that circumvent UDP (or, anaerobically, UTP) reduction by ribonucleotide reductase seem to have been imposed on all organisms. One reasonable explanation for an alternative to the ribonucleotide reductase providing the necessary dUMP via reduction of UDP and subsequent cleavage of dUTP is that this pathway is bound to accumulate even small amounts of dUTP (Fig. 1). dUTP is readily incorporated into DNA by the DNA polymerase in place of dTTP, and extensive incorporation leads to chromosomal decay due to the massive repair of DNA and the creation of double-strand breaks (64, 65). This process in its extreme is also known as thymineless death when caused by the depletion of dTTP, e.g., in mutants where thymidylate synthase is defective (6). With the two cytosine deoxyribonucleotide-dependent pathways, the accumulation of dUTP is avoided by the direct formation of dUMP. That this redirection of dUMP synthesis away from the ribonucleotide reductase/dUTPase pathway (Fig. 1) has evolved with the emergence of dTTP in DNA (2) is indicated by the observation that the affinity of ribonucleotide reductase for UDP is significantly lower than that for the other three ribonucleoside diphosphates (9–12). The most extreme case is the phage T4 anaerobic class III enzyme, which does not show any significant activity in reducing UTP (35).

The structure of the B. halodurans DCD:DUT is very similar to that of other known structures of this enzyme, and the mechanism for inhibition by dTTP seems to closely resemble that of the previously characterized enzymes (17, 43). The dcdB gene is distributed widely among Bacillus species, some of which are found in the Bacillus sensu lato group (B. cereus and B. thuringiensis [66]), but is apparently absent in B. subtilis and other species within the Bacillus sensu stricto group (67). This seems to indicate one of two scenarios for the variation in whether the dcdB gene is present: either a member of the genus acquired the dcdB gene via horizontal gene transfer and formed the branching point within the genus that carries this gene, or a genus member that lost the gene forms the branching point for species from which this gene is absent when separating from ancestors carrying the dcdB gene. Perhaps supporting the first of the two scenarios is the observation from simple database searches that the related streptococci and listeria, both of the class Bacilli, seem to encode only a dCMP deaminase (comEB). Interestingly, a subgroup of staphylococci, which are also of the class Bacilli, whose genomes encode the dcdB gene either have both genes (Staphylococcus pseudintermedius and Staphylococcus hyicus) or do not have the comEB gene (Staphylococcus agnetis and Staphylococcus chromogenes), as revealed by a database search, indicating that a similar event of gene transfer has occurred among staphylococci.

The question of the acquisition or loss of the dcdB gene can be more fully addressed in the future after bioinformatic analysis of more genomes within the Bacillus genus has been performed. The outcome of such an analysis can prove valuable to our understanding of the evolution of the pathways for dTTP biosynthesis, pathways that seem to be more relaxed in terms of adopting alternative new enzyme activities compared to what we usually observe in central metabolism and nucleotide metabolism in particular.

ACKNOWLEDGMENTS

We thank Preben Nielsen (Novozymes) for the Bacillus halodurans C125 strain and MAXLab for beam time.

We thank DANSCATT for funding.

FOOTNOTES

    • Received 26 January 2015.
    • Accepted 3 March 2015.
    • Accepted manuscript posted online 6 March 2015.

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