Abstract
The three-dimensional structure of the enzyme dihydrodipicolinate synthase (KEGG entry Rv2753c, EC 4.2.1.52) from Mycobacterium tuberculosis (Mtb-DHDPS) was determined and refined at 2.28 Å (1 Å=0.1 nm) resolution. The asymmetric unit of the crystal contains two tetramers, each of which we propose to be the functional enzyme unit. This is supported by analytical ultracentrifugation studies, which show the enzyme to be tetrameric in solution. The structure of each subunit consists of an N-terminal (β/α)8-barrel followed by a C-terminal α-helical domain. The active site comprises residues from two adjacent subunits, across an interface, and is located at the C-terminal side of the (β/α)8-barrel domain. A comparison with the other known DHDPS structures shows that the overall architecture of the active site is largely conserved, albeit the proton relay motif comprising Tyr143, Thr54 and Tyr117 appears to be disrupted. The kinetic parameters of the enzyme are reported: KMASA=0.43±0.02 mM, KMpyruvate=0.17±0.01 mM and Vmax=4.42±0.08 μmol·s−1·mg−1. Interestingly, the Vmax of Mtb-DHDPS is 6-fold higher than the corresponding value for Escherichia coli DHDPS, and the enzyme is insensitive to feedback inhibition by (S)-lysine. This can be explained by the three-dimensional structure, which shows that the (S)-lysine-binding site is not conserved in Mtb-DHDPS, when compared with DHDPS enzymes that are known to be inhibited by (S)-lysine. A selection of metabolites from the aspartate family of amino acids do not inhibit this enzyme. A comprehensive understanding of the structure and function of this important enzyme from the (S)-lysine biosynthesis pathway may provide the key for the design of new antibiotics to combat tuberculosis.
- diaminopimelate pathway
- dihydrodipicolinate synthase (DHDPS)
- lysine biosynthesis
- Mycobacterium tuberculosis
- succinylase branch
INTRODUCTION
Mycobacterium tuberculosis (Mtb), the causative agent of tuberculosis, causes more deaths than any other bacterium [1,2]. The increase in tuberculosis cases worldwide, particularly among immunocompromised individuals, combined with the increase in multidrug-resistant strains, highlights the need for new antituberculosis drugs [3].
Mycobacterial cell walls are characterized by an unusually high DAP (diaminopimelic acid) content. DAP, an intermediate of the (S)-lysine biosynthetic pathway, is a constituent of the short peptide bridges that cross-link peptidoglycan polymer chains. Consequently, the absence of DAP results in cell lysis and death, as has been demonstrated in gene-knockout experiments with Mycobacterium smegmatis [4]. The process of cell wall assembly and the biosynthesis of cell wall components have long been accepted as targets for antibiotic design and many existing antibiotics inhibit key steps therein [5]. Therefore an inhibitor that inactivates any of the enzymes that are unique to the (S)-lysine biosynthetic pathway, preventing the synthesis of these crucial metabolites, would be a very effective antibiotic [6]. Additionally, the absence of the (S)-lysine pathway in mammals means that inhibitors of this pathway would not be expected to have mammalian toxicity [6].
The enzyme DHDPS (dihydrodipicolinate synthase) (KEGG entry Rv2753c; EC 4.2.1.52) catalyses the first unique reaction of (S)-lysine biosynthesis [7]: an aldol condensation between (S)-ASA [(S)-aspartate β-semialdehyde] and pyruvate (Scheme 1) [8]. The product of the reaction is the unstable heterocycle HTPA [(4S)-4-hydroxy-2,3,4,5-tetrahydro-(2S)-dipicolinate], which is thought to undergo a non-enzymatic dehydration to (S)-2,3-dihydrodipicolinate, the substrate of the next enzyme in the (S)-lysine biosynthetic pathway, DHDPR (dihydrodipicolinate reductase) [9,10]. The proposed Ping Pong mechanism of DHDPS involves Schiff base formation between pyruvate and an active-site lysine residue, followed by the release of water and then enamine formation [7], which has been suggested to be essentially irreversible [11]. (S)-ASA then binds to the stable substituted enzyme form, and this event is followed by the release of the product and the regeneration of the free enzyme [10]. A catalytic triad, involving residues from two monomers, was shown to have a crucial role in catalysis; the current hypothesis is that these three residues are involved in the transport of protons from the active site to bulk solvent [12].
All DHDPS enzymes that have had both their structure solved and function confirmed are homotetramers. The three-dimensional structures of DHDPS from Escherichia coli [10,13,14], Nicotiana sylvestris [9], Thermotoga maritima [15] and Bacillus anthracis [16], as well as the structures of five point mutants of the E. coli DHDPS [12,17], have been determined.
Characterized DHDPS enzymes display a quaternary structure best described as a dimer of tight dimers, with many interactions between the two monomers to form the tight dimer, and, in the case of the bacterial enzymes, relatively fewer interactions between the tight dimers that form the tetramer [13]. Recently, the structure of a putative DHDPS from Agrobacterium tumefaciens was determined to be a hexamer (PDB code 2HMC), although the function of this enzyme (and its annotation as a DHDPS) has yet to be confirmed. Despite originating from different kingdoms of life, the structure of the monomeric subunits of E. coli and the plant species N. sylvestris DHDPS enzymes are strikingly similar.
Intriguingly, the quaternary structure of the plant and bacterial species is quite different. The arrangement of the tight dimer is the same, but the arrangement of the two tight dimers to form the tetramer is different. The altered conformation can perhaps be explained by the proposed mechanism of allosteric inhibition. The activity of DHDPS is moderately inhibited by (S)-lysine in some bacteria, such as E. coli [10], whereas, in plants, a greater degree of inhibition is observed [9]. The structures of DHDPS from both E. coli and N. sylvestris complexed with (S)-lysine show that one (S)-lysine molecule is bound to each monomer, but, at the same time, is co-ordinated by residues from both monomeric units at the tight-dimer interface [9,10]. Upon (S)-lysine binding, a significant conformational change occurs in the N. sylvestris DHDPS structure [9], which is in striking contrast with the situation in the E. coli DHDPS, where relatively few residues shift upon (S)-lysine binding [13].
The focus of the present study is to understand the structure–function relationship in Mtb-DHDPS, which will underpin the rational design of antimicrobials. In this paper, we present an in-depth structural and kinetic study of the ‘toolkit’ used by Mtb-DHDPS in catalysis.
EXPERIMENTAL
Materials
Unless otherwise stated, all chemicals were obtained from Sigma Chemical Co., GE Biosciences or Invitrogen, and all enzyme manipulations were carried out at 6 °C or on ice. Protein concentration was measured by a modification of the Bradford method, with improved linearity over a broader range of concentrations [18] using BSA as a standard.
Overexpression and purification of Mtb-DHDPS
For the structural studies, the cloning, expression, purification and crystallization of Mtb-DHDPS has been described previously [19]. In brief, the M. tuberculosis dapA gene (Rv2753c) was cloned and expressed in E. coli cells and purified to homogeneity by affinity and size-exclusion chromatography.
For functional studies, the pETM11 plasmid containing the M. tuberculosis dapA gene was transformed into E. coli BL21(DE3) cells containing the pGroESL plasmid, which codes for the GroEL and GroES chaperones [20]. This step increased the amount of Mtb-DHDPS isolated. Cells were grown overnight at 37 °C and induced with IPTG (isopropyl β-D-thiogalactoside) for 3 h. Both the extraction and elution buffers contained 10 mM pyruvate, which has been used previously to stabilize E. coli DHDPS [11]. Following elution from a 5 ml Ni-charged column, fractions containing DHDPS activity (determined using the o-aminobenzaldehyde assay [21]) were incubated overnight with recombinant TEV (tobacco etch virus) protease [1 mM DTT (dithiothreitol) and 5 mM EDTA] to remove the His6 tag, then dialysed or exchanged into a storage buffer [20 mM Tris/HCl, 2 mM 2-mercaptoethanol and 5% (v/v) glycerol (pH 8)]. Any remaining contaminants were removed by gel filtration, and the active peak, which eluted with a molecular mass of approx. 120 kDa, consistent with a homotetramer, was collected. The purified Mtb-DHDPS was analysed by SDS/PAGE (4–12% gel) and blue native PAGE (4–16% gel) (NuPAGE Bis-Tris gel; Invitrogen) [22], and stored in storage buffer containing 250 mM NaCl at 6 or 22 °C for several weeks without detectable degradation.
Crystallization, structure solution and refinement
Crystals of Mtb-DHDPS were grown in the presence of 28% (w/v) PEG [poly(ethylene glycol)] 4000, 170 mM MgCl2 and 100 mM Tris/HCl (pH 8.5). Crystals were flash-cooled in a nitrogen stream at −173 °C, using 20% (v/v) MPD (2-methyl-2,4-pentanediol) in reservoir solution as cryoprotectant. Diffraction data were collected on the XRD (X-ray diffraction) beamline at the ELETTRA synchrotron (Trieste, Italy) using a MAR CCD (charge-coupled device) (165 mm) detector. The crystals belong to the primitive monoclinic space group P21 with the following unit cell parameters: a=94.79 Å (1 Å=0.1 nm), b=87.37 Å, c=139.85 Å, β=107.78° [19]. Based on the self-rotation function, it was concluded that the asymmetric unit of the crystals contained two tetramers exhibiting D2 symmetry. The structure of Mtb-DHDPS was solved by molecular replacement using a single monomer of T. maritima DHDPS (PDB code 1O5K [15]) as a search model. The molecular replacement solution was then subjected to rigid-body, positional and B-factor refinement protocols as implemented in CNS [23]. At this point the free R-factor had fallen to 46%, indicating the correctness of the solution. The correct amino acid sequence was introduced for one monomer using GUISIDE [24] and the co-ordinates for the remaining seven monomers were generated utilizing the non-crystallographic symmetry. The model was improved further using iterative manual model corrections using the program O [25] and refinement in REFMAC5 [26]. Non-crystallographic symmetry restraints were used throughout the refinement. Water molecules were placed using the program ARP/wARP [24]. The refinement statistics are summarized in Table 1. The quality of the model was checked using the program PROCHECK [27]. The Figures illustrating structural details were prepared using the program PyMOL (DeLano Scientific), unless stated otherwise. The refined co-ordinates, as well as the corresponding structure factor amplitudes, were deposited with the PDB under the accession number 1XXX.
R.m.s.d, root mean square deviation.
Sequence alignments were derived from the three-dimensional alignments of the DHDPS structures, which were carried out using the programs ALIGN [29], STAMP [30] and LSQKAB [31]. Buried surface areas were calculated with DSSP [31], the EBI PISA server [32] and the Protein–Protein Interaction server (http://www.biochem.ucl.ac.uk/bsm/PP/server).
Enzyme activity assay
The activity of DHDPS was studied using a coupled assay as described previously [12], with the following components: pyruvate, (S)-ASA, DHDPR, NADPH and Hepes buffer. (S)-ASA was synthesized using the methods of Roberts et al. [33] and was of high quality (>95%) as judged by 1H-NMR. Control assays were performed to ensure the absence of contaminating NADPH-utilizing enzymes and to ensure an excess of DHDPR. The coupled assay follows the consumption of NADPH by the next enzyme in the (S)-lysine biosynthetic pathway, DHDPR [12,34,35]. Coupling enzyme was purified from T. maritima. All components were incubated for 15 min before initiation of the reaction by the addition of DHDPS. The temperature was kept constant at 30 °C by the use of a circulating water bath and the pH was maintained at 8.0 by 100 mM Hepes buffer (from a stock of 200 mM Hepes, pH 8.0, at 22 °C). Selwyn's test [36] was performed to ensure that enzymes and substrates were stable over the course of the assay (results not shown). The initial rate of NADPH consumption was measured in triplicate, and was reproducible (±10%). The data were fitted with the appropriate kinetic models using the program ENZFITTER (Biosoft).
Enzyme stability
The optimum pH for enzyme activity was determined using a series of 20 mM buffers (Mes, Hepes or Bicine) covering a pH range of 6–9 at 30 °C with ionic strength of 0.15 M (adjusted by the addition of NaCl). The optimum ionic strength for the enzymatic reaction was determined using a series of 20 mM Hepes buffers with various NaCl concentrations, corrected for the amount of HCl/NaOH added to bring the pH to 8.25 at 30 °C. For heat stability assays, DHDPS from M. tuberculosis or E. coli was buffer-exchanged into 20 mM Hepes (pH 8.25, at 30 °C with an ionic strength of 0.15 M) before incubation at various temperatures (30–100 °C) using a solid heat block. Aliquots of 10 μl were taken after an incubation of 5 min, at a range of temperatures (30–100 °C), or at 70 °C with a range of incubation times (0–60 min), and stored on ice or added directly to initiate the coupled assay. The heat stability of DHDPS from M. tuberculosis and E. coli was also analysed with a thermal melt in 15 mM Tris/HCl solutions (ionic strength of 0.15 M) containing 10× Sypro Orange (Molecular Probes) with protein unfolding monitored by fluorescence emission at 575 nm [37]. The wavelength of excitation was 490 nm, and the plate was heated from 20 to 90 °C by an iQ5 Real Time PCR Detection System (Bio-Rad Laboratories).
Analytical ultracentrifugation
Sedimentation velocity experiments were performed with purified Mtb-DHDPS using an An-60 Ti four-hole rotor in a Beckman Coulter Model XL-I analytical ultracentrifuge equipped with a UV–visible absorbance optical system. Double-sector cells with quartz windows were loaded with 400 μl of reference (20 mM Tris/HCl and 0.15 M NaCl, pH 8.0) and 380 μl of sample (0.055 mg·ml−1) and centrifuged at 40000 rev./min and 20 °C. Radial absorbance data were collected at 230 nm every 6 min without averaging. Data were fitted to a continuous size-distribution model using the program SEDFIT [38]. The program SEDNTERP [39] was used to determine the partial specific volume (v) of the sample (0.7402 ml·g−1), buffer density (1.005 g·ml−1), buffer viscosity (1.021 cP), and the standardized sedimentation coefficient (S020,w). The predicted standardized sedimentation values for three different geometries of tetramer were calculated using eqn (1) [40], where M is the predicted molar mass of the tetramer, and F is the geometric factor.
(1)
RESULTS AND DISCUSSION
Overall structure
Mtb-DHDPS is a tetramer comprising four identical subunits arranged in D2 symmetry (Figure 1A). Each monomer (300 amino acid residues) comprises an N-terminal (β/α)8-barrel domain (residues 1–233) and a C-terminal domain (residues 234–300) consisting of three α-helices (Figures 1B–1D). The residues responsible for substrate binding and catalysis are located in the (β/α)8-barrel domain. The crystallographic asymmetric unit contains two tetramers of the enzyme. Each tetramer can be described as a dimer of dimers, with the two monomers A and B (and C and D, Figure 1A) tightly bound to each other to form the tight dimer, and weaker interactions between the AB and CD dimeric units.
(A) The two monomers forming the tight dimer AB are coloured red and light red, and the two monomers forming the second tight dimer CD are coloured green and light green respectively. Monomer structure (B) and topology plot (C) of Mtb-DHDPS. (D) Superposition of DHDPS structures on to the A-subunit of Mtb-DHDPS (1XXX, cyan). R.m.s.d. (root mean square deviation) values (in Å) and number of superposed residues (out of 296), are also given in parentheses: E. coli (1YXC, purple, 1.37, 285); T. maritima (1O5K, orange, 1.32, 286); N. sylvestris (structural data from R. Huber, green, 1.53, 276); B. anthracis (1XKY, grey, 1.17, 287); and A. tumefaciens (2HMC, yellow, 1.75, 272).
The final refined Mtb-DHDPS model contains 2364 residues in total, 1587 water molecules, eight DTT molecules covalently bound to Cys248, eight Mg2+ and eight Cl− ions. The model includes all amino acid residues, with the exception of the first four or five amino acids at the N-terminus, which are not visible in the electron density. Of all residues, 90.4% are in the most favoured regions of the Ramachandran plot [41] and 9.2% of residues are in additionally allowed regions. Only Tyr117 lies in the forbidden region of the Ramachandran plot, with the exact values being ϕ=74.9° and ψ=−52.5° for Tyr117 in chain A. The unusual conformation of Tyr117 is well supported by the electron density in each of the monomers. The same observation has been made previously for the corresponding residue in all other DHDPS enzymes of known structure. Tyr117 lies at the tight-dimer interface of two monomers and takes part in formation of the active site of the neighbouring monomer.
A secondary structure matching search against all structures deposited in the PDB using subunit A of Mtb-DHDPS yielded a total of 21 hits above a Q-score threshold of 0.32. After removing redundancies, 12 entries were left (Q-score=0.57–0.78) (see Supplementary Table 1 at http://www.BiochemJ.org/bj/411/bj4110351add.htm). The most similar structure was found to be DHDPS from B. anthracis followed by DHDPS from T. maritima. At the time of solving the structure of Mtb-DHDPS, only the latter was available in the PDB. This in retrospect validates the choice of the search model for molecular replacement, which was based solely on amino acid sequence comparison. An interesting observation is the apparent relationship of DHDPS to N-acetylneuraminate lyases, which has been noted previously [42].
Active site and metal-binding sites
The active site is located at the centre of each monomer, facing the central cavity of the tetramer. It is situated in a pocket at the C-terminal side of the (β/α)8-barrel, as in all known (β/α)8-enzymes. The active site centres about Lys171, the amino acid that forms a Schiff base with the first substrate, pyruvate. In the E. coli enzyme, the equivalent amino acid is Lys161, as previously identified via tryptic digest studies [7]. Three amino acid residues form the conserved catalytic triad: Tyr143, Thr54, and Tyr117, whereby Tyr117 is contributed from the adjacent monomer across the tight-dimer interface. Both the identity, as well as the relative spatial orientation of these functional groups, is conserved among characterized DHDPS enzymes, but not conserved in the putative A. tumefaciens DHDPS enzyme (Swiss-Prot accession number Q8U6Y1), where the corresponding residues are Tyr136, Ser48 and Leu108 (Figure 2). As mentioned above, the exact function of the A. tumefaciens enzyme remains to be established. A detailed examination of the Mtb-DHDPS active site, however, shows the distance between Tyr143-OH and Thr54-OH is somewhat increased (4.7 Å) compared with E. coli DHDPS (2.7 Å), and the geometry is such that hydrogen-bonding is unlikely (Figure 3). In addition, the active site also contains a methionine residue (Met251) and a cysteine residue (Cys248), which, in Mtb-DHDPS, binds a DTT molecule. These residues sit opposite Lys171, lining the active-site cavity (Figure 3), and could be exploited in rational inhibitor design. We have noted recently the potential for a speciesspecific inhibitor designed to target DHDPS [43]. Whether these motifs are utilized in Mtb-DHDPS catalysis is under investigation. The mechanism of the DHDPS-catalysed reaction has been studied in detail for the E. coli enzyme by X-ray crystallography, NMR [10] and site-directed mutagenesis [12]. After Schiff base formation between pyruvate and the active-site lysine residue, (S)-ASA binds to the stable substituted enzyme form. The product heterocycle (HTPA) is then released and undergoes dehydration and the enzyme is regenerated [10].
Multiple sequence alignment of the DHDPS sequences from M. tuberculosis (1XXX), E. coli (1YXC), T. maritima (1O5K), B. anthracis (1XKY), N. sylvestris (Nsyl) and A. tumefaciens (2HMC). The alignment was derived from the three-dimensional alignments of the DHDPS structures, which were carried out using the programs ALIGN [29] and STAMP [30]. The secondary-structure elements (α-helices and β-strands) observed in the structure of Mtb-DHDPS are indicated and labelled. Amino acid residues in black boxes are conserved in four or more of the six DHDPS enzymes listed. Hash marks (#) indicate those residues involved in catalysis, and an asterisk (*) indicates those residues that are involved in (S)-lysine binding in the E. coli, T. maritima and N. sylvestris enzymes.
The active site, defined by the position of Lys171, is found in a cleft within the (α/β)8-barrel (secondary structure is transparent). Tyr117 from the adjacent monomer reaches through the interface (AB from Figure 1) to interact with Thr54.
An Mg2+ ion is co-ordinated by Ala162-O, His164-O and Ile167-O and three water molecules. In the E. coli (PDB codes 1DHP and 1YXC [13,14]) and B. anthracis (PDB entry 1XKY [16]) DHDPS structures, a K+ ion is found in the same position as the Mg2+ ion in the Mtb-DHDPS structure. The function of this metal-binding site is unclear, especially since it is located far from the enzyme active site. We note that, via the coupled assay, the activity of the enzyme is unaffected by Mg2+ ions (<20 mM). It is likely that the Mg2+ ions seen in the structure are an artefact of the crystallization conditions. A Cl− ion, co-ordinated by Lys174-NZ, Ser179-OG and two water molecules, was also observed in Mtb-DHDPS. In contrast with the model of the E. coli enzyme (PDB code 1YXC [13]), where a Cl− ion is found in the active site, co-ordinated by the catalytic Lys161, Thr44 and two water molecules [13], the Cl−-binding site in Mtb-DHDPS is distal to the active site, leaving its functional significance unclear.
Dimer–dimer interface of Mtb-DHDPS
In order to confirm the biologically significant unit of the enzyme, all the potential interfaces were examined using the EBI PISA server [32], and the Protein–Protein Interaction server. Within the tetrameric unit shown in Figure 1(A), each of the two tight-dimer interfaces (AB and CD) bury approx. 1470 Å2 per monomer, which corresponds to roughly 13% of the surface area of the monomer, whereas the interfaces between monomers A and D, and monomers B and C, bury a surface area of approx. 860 Å2 per monomer (see Supplementary Table 2 at http://www.BiochemJ.org/bj/411/bj4110351add.htm). Polar residues represent 40% of the tight interface area, and 30% of the AD and BC interfaces. For the E. coli enzyme, each of the two tight-dimer interfaces (AB and CD) bury approx. 1300 Å2 per monomer (roughly 11% of the surface area of the monomer), whereas the weaker dimer interfaces AC and BD bury a surface area of approx. 500 Å2. The polar residues represent 35% of the tight-dimer interface area, and 44% of the weak AC and BD interfaces. The T. maritima DHDPS structure buries an interface-accessible area similar to that of the Mtb-DHDPS structure (see Supplementary Table 2). Figure 4(A) shows a schematic diagram of the E. coli DHDPS and Mtb-DHDPS dimer–dimer interface. This comparison suggests that the oligomeric structure of Mtb-DHDPS may be somewhat more stable than the structure of E. coli DHDPS.
(A) Analysis of the dimer–dimer interface of tetrameric Mtb-DHDPS (1XXX) and E. coli DHDPS (1YXC). Interfaces were probed using JavaProtein Dossier [51] and manual inspection. Dotted lines indicate hydrogen-bonding, continuous lines indicate hydrophobic interactions, and circles represent water. The thicker lines indicate that the interactions are in both symmetrical interfaces (i.e. interfaces AD and BC of Figure 1) of the tetramer. Amino acids are identified using single-letter codes. (B) Sedimentation velocity absorbance data plotted as a function of radial position from the axis of rotation (cm) for Mtb-DHDPS at a concentration of 0.055 mg·ml−1 in 20 mM Tris/HCl and 150 mM NaCl, at pH 8.0. The raw data are presented as open symbols (○) plotted at time intervals of 6 min overlaid with the non-linear least squares best-fit (solid line) to a continuous size distribution model [c(M)] [38]. The residuals (top) for the c(M) distribution best-fit are plotted as a function of radial position (cm) from the axis of rotation. (C) The c(M) distribution is plotted as a function of molar mass (kDa) for Mtb-DHDPS. The fit was obtained using a resolution of 200 species between Mmin of 2.5 kDa and Mmax of 350 kDa with v=0.7402 ml·g−1, ρ=1.00499 g·ml−1, η=1.0214 cP, and f/f0=1.21902. The r.m.s.d. (root mean square deviation) and Z test for the fit were 0.003895 and 3.49 respectively.
In order to corroborate the apparent oligomeric state seen in the crystal structure, further study of Mtb-DHDPS in solution was performed via analytical ultracentrifugation. Sedimentation velocity experiments, fitted to a continuous size-distribution model [38], confirmed that Mtb-DHDPS was a single species in solution with a molecular mass of 115 kDa, taken from the ordinate maximum of the peak observed in the c(M) distribution (Figure 4C). Based on c(s) distribution analysis (results not shown), the tetrameric species has a sedimentation coefficient (S020,w) of 6.7 S. We note that, at a concentration of 0.055 mg·ml−1, Mtb-DHDPS shows no dimer in solution. This is in contrast with the E. coli enzyme, which shows a significant proportion of dimer at a similar concentration (0.05 mg·ml−1) [44], suggesting that the dimer–dimer interface for Mtb-DHDPS is indeed stronger than in the E. coli enzyme. Using eqn (1), the spatial arrangement of the Mtb-DHDPS tetramer in aqueous solution (S020,w=6.7 S) is intermediate between square planar (calculated S020,w=6.540 S, F=0.926) and tetrahedral (calculated S020,w=6.903 S, F=0.977), which correlates well with the particular D2-symmetric shape shown in the X-ray crystal model (Figure 1). The homotetrameric structure of Mtb-DHDPS in solution has been corroborated further by gel filtration and blue native PAGE (results not shown).
Enzyme kinetics
The coupled enzyme assay was used to characterize the kinetic properties of Mtb-DHDPS. Initially, the stability and pH optimum for activity was assessed. For enzymatic activity of Mtb-DHDPS, the optimum pH was determined to be 8.25 at 30 °C in Hepes buffer (Figure 5A). Additionally, a steady decrease in activity was seen when the ionic strength was increased above 0.1 M (Figure 5B). In a buffer of 20 mM Hepes (pH 8.25) at 30 °C (ionic strength 0.15 M), Mtb-DHDPS showed a greater thermal stability than E. coli DHDPS, with an apparent Tm (melting temperature) of ∼82 °C compared with ∼57 °C for the E. coli enzyme (Figure 5C). Further experiments showed that Mtb-DHDPS maintained its activity for 40 min when incubated at 70 °C, whereas E. coli DHDPS showed degradation within the first few minutes (results not shown). This result was supported by following protein unfolding monitored by fluorescence emission (Figure 5D). The thermal stability of the Mtb-DHDPS is probably in part due to the greater number of intersubunit contacts as compared with the E. coli enzyme (Figure 4A).
The effect of varying (A) pH with different buffers (⊙ Mes, ○ Hepes and ● Bicine), and (B) ionic strength on enzyme activity was measured using the coupled assay. The data in (B) are normalized to the highest measured rate (vcon) of enzyme activity. (C) Enzyme activity determined using the coupled assay after pre-treatment using a 5 min incubation at increasing temperature. The thermal stability of Mtb-DHDPS (▲) was compared with E. coli DHDPS (Δ) using the apparent initial rate (vapp) divided by rate of enzyme activity without pre-treatment (vcon). Results are means±S.D. of triplicate measurements. (D) The melting temperature as determined by a fluorescence melt for Mtb-DHDPS (○) and E. coli DHDPS (●) was comparable with that determined using activity assays.
A full matrix of initial rates was determined with both substrates varied, and these data were fitted with different kinetic models: ternary complex, Ping Pong and Ping Pong with substrate inhibition. The Ping Pong model provided the best fit (Figure 6), and yielded KM constants for pyruvate (0.17±0.01 mM) and (S)-ASA (0.43±0.02 mM). Interestingly, a 6-fold increase in the Vmax (4.42±0.08 μmol·s−1·mg−1) was observed over E. coli DHDPS, which may reflect the increased importance of DAP in the peptidoglycan layer of mycobacteria [45]. That the ordered Ping Pong kinetic mechanism is conserved in Mtb-DHDPS, is consistent with the ordered kinetic mechanism proposed for other DHDPS enzymes. We note that the Ping Pong mechanism is symmetrical, but speculate that, in the case of Mtb-DHDPS, pyruvate is the first substrate to bind, followed by (S)-ASA. Whether the catalytic triad, as proposed for the E. coli enzyme, operates in this enzyme remains to be elucidated; as we have observed above, this motif is conserved, yet its geometry probably precludes function without structural changes occurring, perhaps during catalysis. The structure of Mtb-DHDPS complexed with various inhibitors or substrates will be helpful in this regard.
The initial velocity was measured using the coupled assay over various concentrations of each substrate, (S)-ASA and pyruvate (× 3.0 mM, ○ 1.5 mM, ⊙ 0.30 mM, ♦ 0.15 mM and ◊0.06 mM). Each point was measured at least in triplicate and the data were fitted with the Ping Pong model (R2 of 0.9945) using the software program Enzfitter (A and B). (C, D) Lineweaver–Burk transformations reflect the trend predicted for the Ping Pong model.
Previous studies suggest that the real power of an enzyme lies not only in the mechanistically important residues that form the catalytic unit, but also in a combination of the very ‘local’ structural features of the catalytic unit and more ‘global’ features, such as the dynamics of the structure and the overall microenvironment of the active site [46]. Despite the high degree of overall structure similarity (both in the solid state and in solution) and active-site conservation, Mtb-DHDPS shares little sequence identity with other DHDPS enzymes, and our data reveal important differences in their structural, biophysical and biochemical properties.
Mtb-DHDPS is not feedback-regulated by (S)-lysine
Except at very high concentrations (50 mM and above), (S)-lysine did not have any effect on the activity of Mtb-DHDPS. The IC50 was determined to be 250 mM, which greatly exceeds that which could be reasonably expected in a cell. Thus, in contrast with plant and Gram-negative DHDPS enzymes [8,35,47–49], Mtb-DHDPS can be considered insensitive to inhibition by (S)-lysine. This observation is explained nicely by structural superposition of the two enzymes and examination of the (S)-lysine-binding site observed in E. coli (Figure 7). Most residues identified as important for the (S)-lysine allosteric binding site are not conserved in Mtb-DHDPS (Table 2) [9,10,13]. Mtb-DHDPS is the only DHDPS enzyme with a known structure in which the asparagine residue of the (S)-lysine-binding site is not conserved, but replaced by a tyrosine residue. Additionally, the (S)-lysine-binding cavity in the M. tuberculosis enzyme is shallow compared with the equivalent site of the E. coli or N. sylvestris enzymes, leaving little space for (S)-lysine to bind. Interestingly, other aspartate family amino acids (DAP, threonine and methionine) showed no significant inhibition of Mtb-DHDPS activity (Table 3).
(A) Surface representation of the tight dimer showing the (S)-lysine-binding site of the E. coli DHDPS (1YXD) with the bound (S)-lysine in yellow. (B) Superposition of the corresponding Mtb-DHDPS (1XXX) residues (in red) on to the (S)-lysine-binding residues (grey) of E. coli DHDPS (1YXD). The bound (S)-lysine residues (from 1YXD) are shown in yellow.
The (S)-lysine-binding residues are identified in E. coli DHDPS (second column), followed by an alignment of corresponding residues in the other known structures. N. sylvestris DHDPS co-ordinates were kindly provided by R. Huber.
The activity of Mtb-DHDPS in the presence of aspartate family amino acids. Each substrate concentration was held at 0.3 mM. Relative activity is relative to that in the absence of inhibitor.
The regulation of the (S)-lysine biosynthetic pathway (if it is indeed regulated in M. tuberculosis) is maintained via an alternative strategy. In E. coli, the DAP pathway is likely to be controlled at both the aspartate kinase and DHDPS catalytic step. Moreover, it has been suggested that E. coli DHDPS expression is regulated by the level of DAP [50]; whether such an approach is adopted by M. tuberculosis remains to be elucidated.
Accordingly, the most valid approach for generating inhibitors and novel antibiotics targeting DHDPS from M. tuberculosis should focus on the highly conserved active-site geometry of the enzyme, rather than the vestigial allosteric cleft (Figure 7).
In conclusion, the increasing prevalence of tuberculosis cases, and especially that of antibiotic resistance, necessitates the development of novel antibiotics. Pivotal to such endeavours is an extensive understanding of the proposed antibiotic targets. In the present paper, we describe a structural and biochemical study of an important antibiotic target from M. tuberculosis. We have found that the active site of Mtb-DHDPS is generally similar to that of E. coli DHDPS. However, we note that the proton-relay residues are such that the hydrogen-bonding network may be disrupted. Given that the residues responsible for substrate binding are generally conserved, lead compounds targeted to the E. coli active site are likely to also be effective against the M. tuberculosis enzyme. Additionally, the presence of a cysteine residue and a methionine residue within the active site may be exploited to develop inhibitors tailored to Mtb-DHDPS. The M. tuberculosis enzyme, unlike E. coli DHDPS, is not allosterically affected by (S)-lysine, which may reflect the requirement for DAP in the bacterial cell wall. Continued investigation into the catalytic mechanisms, the consequences of this enzyme's oligomeric state and the DAP/(S)-lysine biosynthetic pathway in M. tuberculosis are necessary to explain the regulatory mechanism of this essential pathway in bacteria.
Acknowledgments
We thank Dr Jeanne Perry (UCLA, Los Angeles, CA, U.S.A.) for providing genomic Mtb-DNA, Dr Arie Geerlof (EMBL Hamburg) for help with the solubility screens, Dr Santosh Panjikar (EMBL Hamburg) for help in solving the Mtb-DHDPS structure by molecular replacement, Dr Robert Huber (Cardiff University, Cardiff, Wales, U.K.) for providing the structural co-ordinates of N. sylvestris and E. coli DHDPS. We thank Dr George Lorimer (E. I. Dupont De Nemours and Co., Wilmington, DE, U.S.A.) for generously providing the pGroESL plasmid, and Jackie Healy for unassuming and resourceful technical support. G. K. was funded by the TB consortium (http://www.doe-mbi.ucla.edu/TB) for postdoctoral exchange grants and the X-Mtb consortium (http://www.xmtb.org) for funding through BMBF (Bundesministerium für Bildung und Forschung)/PTJ (Projektträger Jülich) grant number BIO/0312992A. G. L. E., J. A. G., M. A. P., M. D. W. G. and R. C. J. D. were supported in part by the Royal Society of New Zealand Marsden Fund (contract UOC303), S. R. A. D. acknowledges financial support from FRST (Foundation for Research, Science and Technology), and M. A. P., J. A. G. and R. C. J. D. were supported in part by the DTRA (Defense Threat Reduction Agency) (project W911NF-07-1-0105).
Footnotes
The structural co-ordinates for Mycobacterium tuberculosis dihydrodipicolinate synthase will appear in the Protein Data Bank under accession code 1XXX.
Abbreviations: DAP, diaminopimelate; DHDPR, dihydrodipicolinate reductase; DHDPS, dihydrodipicolinate synthase; DTT, dithiothreitol; HTPA, (4S)-4-hydroxy-2,3,4,5-tetrahydro-(2S)-dipicolinate; Mtb, Mycobacterium tuberculosis; (S)-ASA, (S)-aspartate β-semialdehyde
- © The Authors Journal compilation © 2008 Biochemical Society
References
April 2008
Volume: 411 Issue: 2
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