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J. Biol. Chem., Vol. 280, Issue 34, 30273-30281, August 26, 2005

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NAD⁺-dependent DNA Ligase (Rv3014c) from Mycobacterium tuberculosis

CRYSTAL STRUCTURE OF THE ADENYLATION DOMAIN AND IDENTIFICATION OF NOVEL INHIBITORS^*

Sandeep Kumar Srivastava, Rama Pati Tripathi¶, and Ravishankar Ramachandran||

From the Divisions Molecular and Structural Biology and ^¶Medicinal and Process Chemistry, Central Drug Research Institute, Chattar Manzil, Mahatma Gandhi Marg, Lucknow-226001, India

Received for publication, April 7, 2005 , and in revised form, May 10, 2005.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
DNA ligases utilize either ATP or NAD⁺ as cofactors to catalyze the formation of phosphodiester bonds in nicked DNA. Those utilizing NAD⁺ are attractive drug targets because of the unique cofactor requirement for ligase activity. We report here the crystal structure of the adenylation domain of the Mycobacterium tuberculosis NAD⁺-dependent ligase with bound AMP. The adenosine nucleoside moiety of AMP adopts a syn-conformation. The structure also captures a new spatial disposition between the two subdomains of the adenylation domain. Based on the crystal structure and an in-house compound library, we have identified a novel class of inhibitors for the enzyme using in silico docking calculations. The glycosyl ureide-based inhibitors were able to distinguish between NAD⁺- and ATP-dependent ligases as evidenced by in vitro assays using T4 ligase and human DNA ligase I. Moreover, assays involving an Escherichia coli strain harboring a temperature-sensitive ligase mutant and a ligase-deficient Salmonella typhimurium strain suggested that the bactericidal activity of the inhibitors is due to inhibition of the essential ligase enzyme. The results can be used as the basis for rational design of novel antibacterial agents.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
DNA ligases are vital enzymes in replication and repair and catalyze the formation of a phosphodiester linkage between adjacent termini in double-stranded DNA through similar mechanisms (1). These enzymes can be divided into two classes, viz. NAD⁺- and ATP-dependent ligases, based on the cofactor specificities (2). NAD⁺-dependent DNA ligases, commonly called LigA, are found in bacteria and entomopox-viruses (3, 20), whereas ATP-dependent ligases are ubiquitous (3). Although there is little sequence homology between the eubacterial and eukaryotic enzymes, they exhibit some structural homology in specific domains (4, 5). The mechanistic steps involved in enzymatic action are also broadly conserved. Briefly, in the first step, the mode of action involves an attack on the -phosphorus of ATP or NAD⁺ by the enzyme, releasing pyrophosphate or NMN and forming a ligase-adenylate intermediate. In the second step, the bound AMP is transferred to the 5'-end of DNA to form a DNA-adenylate intermediate. AMP is released in the third step, where the protein catalyzes the joining of the 3'-nicked DNA to the DNA-adenylate intermediate. These steps involve large conformational changes and also encircling and partial unwinding of the nicked DNA substrate (6-8).

Some bacteria code for both NAD⁺- and ATP-dependent DNA ligases (3, 9). Mycobacterium tuberculosis codes for at least three different types of ATP-dependent ligases and a NAD⁺-dependent ligase (10, 11). Gene knockout and other studies have shown LigA to be indispensable in several bacteria, including Escherichia coli, Staphylococcus aureus, Bacillus subtilis, and M. tuberculosis (10, 12-15).

No LigA structure from mycobacterial sources is available to date. However, the crystal structure of the full-length protein is available for the Thermus filiformis enzyme (TfiLigA),¹ whereas structures of the adenylation domain are available for the Bacillus stearothermophilus and Enterococcus faecalis (EfaLigA) enzymes (7, 8, 16). The structures have shown that the enzyme has a modular architecture consisting of distinct domains. The adenylation domain contains the cofactor-binding site and can be divided further into two subdomains. Subdomain 1a contains the NMN-binding pocket, whereas subdomain 1b contains the AMP-binding site. The EfaLigA structures show that the NAD⁺-binding site is generated by a specific spatial disposition of the two subdomains where subdomain 1a is in close proximity to the AMP-binding site in subdomain 1b (7). Structure-based mutagenesis experiments have also led to the identification of residues important for NAD⁺ recognition and support systematic active-site remodeling in different reaction steps (17).

With the problem of multiple drug resistance spreading across the world, it is important to find inhibitors from different chemical classes with new modes of action. In this context, specific inhibitors for NAD⁺-dependent ligases are being identified, as no drug is known to act against this enzyme so far. Other groups have very recently identified compounds belonging to arylamino and pyridochromanone classes as specific inhibitors of NAD⁺-dependent DNA ligases (18, 19). New inhibitors can also be potentially used as broad bactericidal agents, as NAD⁺-specific enzymes have not been identified in eukaryotic genomes and are exclusively found in eubacteria (3) and some viruses (20).

In this work, we report the crystal structure of the adenylation domain of the M. tuberculosis NAD⁺-dependent DNA ligase (MtuLigA) bound to AMP. The structure captures a new spatial disposition of the two subdomains in the protein. The AMP conformation in the crystal structure is different from that observed in the TfiLigA structure, but is similar to the AMP part of NAD⁺ in its co-crystal structure with Efa-LigA. Based on the crystal structure and in silico docking studies, we have identified glycosyl ureides as a new class of DNA ligase inhibitors. In vitro assays and bactericidal activities assayed using specific E. coli and Salmonella typhimurium strains demonstrated that the compounds are able to distinguish between NAD⁺- and ATP-dependent ligases. Although the M. tuberculosis enzyme was inhibited in the low micromolar range, human DNA ligase I was inhibited only at much higher concentrations.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Cloning, Expression, and Purification—The adenylation domain of MtuLigA (Rv3014c) consists of residues 1-328. The DNA sequence encoding this domain was PCR-amplified from genomic DNA of M. tuberculosis H37Rv using forward primer 5'-GGAATTCCATGGGCTCCCCAGACGCCG-3' and reverse primer 5'-ATCGGATCCCTCGGGCGGGTACTTGTAGG-3' containing NcoI and BamHI restriction sites (underlined), respectively. The amplified PCR product was digested and ligated into pQE60 (Qiagen Inc.) digested at same site. Incorporation of NcoI into the forward primer leads to replacement of the first two amino acids in the sequence: valine and serine to methionine and glycine, respectively. The integrity of the insert was verified by sequencing. The construct was transformed into E. coli BL21(DE3) cells (Novagen) and grown in LB medium containing 0.1 mg/ml ampicillin to A₆₀₀ 0.5. Protein expression was induced by addition of 0.8 mM isopropyl -D-thiogalactopyranoside at 28 °C for 8 h. Cells were harvested by centrifugation; resuspended in 50 mM Tris-HCl (pH 8.0), 200 mM NaCl, and 10 mM imidazole (buffer A); and lysed by sonication. The crude lysate was centrifuged at 27,000 x g for 30 min. The supernatant was applied to a nickel-iminodiacetic acid column (Amersham Biosciences) equilibrated with buffer A, and protein was eluted using a 10-500 mM imidazole gradient. Purified fractions were pooled, precipitated using ammonium sulfate (45% saturation), redissolved in a minimum volume of buffer B (50 mM Tris-HCl (pH 8.0), 50 mM NaCl, 1 mM EDTA, and 2 mM dithiothreitol), and loaded onto a Superdex S-200 gel filtration column (Amersham Biosciences) equilibrated with buffer B. Purified protein was pooled and concentrated to 15 mg/ml using a Centricon concentrator (10-kDa cutoff; Amicon, Inc.). Protein concentrations were determined with Bradford reagent (21) using bovine serum albumin as a standard.

Crystallization and Data Collection—Crystals of the MtuLigA adenylation domain were grown by vapor diffusion using the hanging drop method. A drop containing 2 µl each of 12 mg/ml protein solution in buffer B containing 4 mM NAD⁺ and reservoir solution containing 0.1 M NaCl, 0.1 M Na-HEPES (pH 7.6), and 1.5 M (NH₄)₂SO₄ kept for 1 week at 24 °C yielded crystals of typical dimensions (0.7 x 0.5 x 0.2 mm). These were mounted on capillaries, and x-ray data were collected at room temperature on a MAR imaging plate mounted on a Rigaku rotating anode generator. The crystals diffracted weakly to 3.15 Å, and the data were overall complete to 99.4% with an average redundancy of 8.7. Data integration, reduction, and scaling were performed using the DENZO/SCALEPACK suite of programs (22). The data collection statistics are summarized in Table I.

View larger version (36K): [in this window] [in a new window]   FIG. 1. Split stereo rendering of the C atom trace showing the overall fold of the adenylation domain of MtuLigA. Some residues are labeled for clarity, and the two subdomains are also indicated. The bound AMP cofactor is shown in stick representation.

In Vitro Activity—In vitro assays for ligase activity were performed using a 40-bp double-stranded DNA substrate carrying a single-strand nick between bases 22 and 23 (31). This substrate was created in Tris/EDTA buffer by annealing a 22-mer (5'-CCT GGA CAT AGA CTC GTA CCT T-3') and a 18-mer (5'-AGC TGG ATC ACT GGA CAT-3') to a complementary 40-mer (5'-ATG TCC AGT GAT CCA GCT AAG GTA CGA GTC TAT GTC CAG G-3'). The 18-mer was radiolabeled at the 5'-end by incubating 10 µg of the oligonucleotide with 100 µCi of [-³²P]ATP (3000 Ci/mmol; Board of Radiation and Isotope Technology) and 30 units of T4 polynucleotide kinase for 1 h, followed by 10 min at 70 °C (32). The unincorporated label was removed using a Sephadex G-25 column. The labeled 40-bp nicked DNA substrate was used to assay the in vitro inhibitory activity of different compounds against MtuLigA, bacteriophage T4 ligase, and human DNA ligase I.

The full-length MtuLigA protein was cloned into the NdeI/NcoI-digested pET41a vector (Novagen). After expression in E. coli BL21(DE3) cells, the C-terminally His-tagged protein was purified according to standard procedures. The assays were done with 2 ng of the purified protein. Reaction mixtures (15 µl) containing 50 mM Tris-HCl (pH 8.0), 5 mM dithiothreitol, 10 mM MgCl₂, 10% Me₂SO, 2 µM NAD⁺, 2 pmol of ³²P-labeled nicked duplex DNA substrate, and different concentration of compounds were incubated for 1 h at 25 °C.

Reactions were quenched with formamide and EDTA. The reaction products were resolved electrophoretically on 15% polyacrylamide gel containing 8 M urea in 90 mM Tris borate and 2.5 mM EDTA. Autoradiograms of the gels were developed, and the extent of ligation was measured by scanning the gel using ImageMaster 1D Elite software (Amersham Biosciences). All the compounds were dissolved in 100% Me₂SO. The compound solutions comprised 0.1 volume of the ligation reaction mixture; thus, 10% Me₂SO was included in all the control reactions. The activity assay was performed in the same way for T4 ligase in a volume of 15 µl containing 0.05 unit of enzyme (Amersham Biosciences), 2 pmol of labeled template, and 66 µM ATP in 66 mM Tris-HCl (pH 7.6), 6.6 mM MgCl₂,10mM dithiothreitol, and 10% Me₂SO.

View larger version (33K): [in this window] [in a new window]   FIG. 2. An initial 2Fo - Fc electron density map calculated after molecular replacement at 3.15 Å contoured at 1.2 around the AMP-binding site. The cofactor was not included in the calculations. The refined coordinates are superposed onto the map. AMP (pink) and interacting residues (violet) are labeled for clarity. Polar interactions within 3.5 Å are also indicated by dotted lines. The figure was made using Turbo-Frodo.

The IC₅₀ values were determined by plotting the relative ligation activity versus inhibitor concentration and fitting to equation V_i/V₀ = IC₅₀/(IC₅₀ + [I]) using GraphPad Prism®. V₀ and V_i represent the rates of ligation in the absence and presence of inhibitor, respectively, and [I] refers to the inhibitor concentration.

Antibacterial Activity and Inhibition of Ligase in Vivo—The recombinant plasmid pRBL (34) containing the gene for T4 DNA ligase in pTrc99A was transformed into the E. coli GR501 ligA^ts mutant (35). To have the same genetic background, the M. tuberculosis ligA gene was amplified from genomic DNA using primers containing sites for NcoI and HindIII, cloned into NcoI- and HindIII-digested pTrc99A (36), and transformed into E. coli GR501. In growth experiments, the strains expressing MtuLigA or T4 DNA ligase were compared with a control GR501 strain carrying empty pTrc99A without any gene insertions at 37 °C. As reported previously (19) and reproduced by us, the E. coli GR501 ligA^ts strain grows well at 30 °C, whereas it is strongly delayed at 37 °C. Complementation with either MtuLigA or T4 ligase restores the growth of the mutant strain.

Minimum inhibitory concentrations (MICs) of the inhibitors were determined for MtuLigA and T4 DNA ligase in the E. coli GR501 ligA^ts mutant and in S. typhimurium LT2 (37) and its DNA ligase-null mutant derivative, which had been rescued with a plasmid (pBR313/598/8/1b) encoding the T4 DNA ligase gene (38), to check the specificity of compounds for NAD⁺-dependent ligases from other sources as well. Antimicrobial activity was monitored in microtiter plates using a microdilution assay technique in a volume of 200 µl. Approximately 10⁵ colony-forming units/ml in the case of the E. coli ligA^ts mutant and 10⁶ colony-forming units/ml in the case of S. typhimurium LT2 and its LigA^- mutant strain, rescued with T4 DNA ligase, were incubated with different compound concentrations under ambient conditions for 20 h, and MICs were determined on the basis of the presence of any visible growth. The E. coli mutant strain was grown in LB medium, whereas the S. typhimurium strains were grown in nutrient broth. The media contained 20 µg/ml polymyxin B nonapeptide to facilitate passage of the inhibitors across the outer membrane.

Growth Inhibition Studies—To investigate the specificity and sensitivity of the compounds to NAD⁺-dependent ligase, exponentially growing cultures of S. typhimurium LT2 and its DNA ligase-null mutant derivative in nutrient broth were treated at A₆₀₀ = 0.4 with increasing compound concentrations. The effect on the growth and viability of both the strains was compared by monitoring A₆₀₀ and the number of colony-forming units for 4-5 h after addition of the compound. Serially diluted culture aliquots of both strains in phosphate-buffered saline were plated on nutrient agar, and visible colonies were counted after incubating the plates for 15 h at 37 °C.

DNA-Inhibitor Interaction—In this assay, the DNA intercalating properties of the inhibitors were measured by the ability to compete with ethidium bromide for DNA binding. Detection of ethidium bromide displacement from DNA, if any, is based on the strong loss of fluorescence that should occur upon its detachment from DNA (39). The assay mixture contained, in a volume of 100 µl, 5 µg of calf thymus DNA, 5 µM ethidium bromide, 25 mM Tris-HCl (pH 8.0), 50 mM NaCl, and 1 mM EDTA. Upon addition of the inhibitor at increasing concentrations, ethidium bromide fluorescence was immediately detected at an excitation wavelength of 485 nm and an emission wavelength of 612 nm.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The adenylation domain of LigA contains all the residues necessary for AMP/NAD⁺ binding. This domain (consisting of residues 1-328 in MtuLigA) was cloned, expressed, and purified as described. The adenylation domain itself consists of two subdomains. Subdomain 1a is known to be flexible and adopts different spatial dispositions relative to subdomain 1b (7, 8). We therefore carried out independent molecular replacement calculations for the subdomains. The cofactor was not used in the calculations (Fig. 1). Clear connectivity was observed in the initial electron density maps between the two subdomains. We added NAD⁺ under the crystallization conditions, but well defined density only for noncovalently bound AMP was observed in the initial electron density map itself (Fig. 2). The data collection and refinement statistics are summarized in Table I.

Subdomain 1a consists of residues 1-76 and contains residues involved in NMN recognition. This subdomain consists mainly of two helical stretches (Figs. 1 and Fig. 3A). Subdomain 1b contains bound AMP in the crystal structure and consists of residues 77-328. The two domains adopt a novel relative spatial disposition in the structure (Fig. 3). Subdomain 1a in the B. stearothermophilus LigA structure is at one end of the conformation spectrum, whereas in TfiLigA, it is at the other end (Fig. 3B). In EfaLigA, this domain comes in close proximity to the AMP-binding site and generates the complete NAD⁺-binding site. This subdomain in MtuLigA adopts a new spatial disposition between the above two extremes.

Adenylation Site and AMP Conformation—The adenylation domain contains five of six conserved sequence motifs in NAD⁺ ligases (40). These mainly line the AMP/NAD⁺-binding pocket. The active-site lysine (Lys¹²³ in MtuLigA), which covalently binds AMP to form the ligase-adenylate intermediate in the first step of the reaction, is part of the conserved motif I, whereas a Glu residue (Glu¹⁸⁴ in MtuLigA), which apparently discriminates between AMP conformations, is part of motif III (17). Intriguingly, in the present structure, AMP is not observed to form the phosphoamide adduct with the motif I lysine. In fact, the side chain of this residue appears to be more mobile as evidenced by weaker electron density. The only other AMP-bound LigA structure is that of TfiLigA, whereas an NAD⁺-bound structure is available for EfaLigA. Although the model used for molecular replacement calculations was derived from TfiLigA, the AMP molecule adopts a conformation similar to that of the AMP part in the NAD⁺-bound EfaLigA structure (Protein Data Bank code 1TAE [PDB] ). In the TfiLigA structure, the adenylation domain is in an "open" state (Fig. 3B), and the adenosine nucleoside moiety of the covalently bound AMP adopts an anti-conformation. In the NAD⁺-bound EfaLigA structure, the adenylation domain is observed to be in a "closed" state, and the adenosine nucleoside moiety adopts a syn-conformation (41).

View larger version (51K): [in this window] [in a new window]   FIG. 3. A, schematic of the MtuLigA adenylation domain crystal structure. Individual subdomains 1a and 1b are shown in dark blue and cyan, respectively. The bound cofactor is also indicated. The figure were made using MolScript (44). B, superposition of the adenylation domains from B. stearothermophilus LigA (B. st; Protein Data Bank code 1B04 [PDB] ), TfiLigA (T. f; code 1V9P), and EfaLigA (E. f; code 1TAE) onto the MtuLigA (M. tb) structure. Subdomain 1b is shown in cyan, whereas subdomains 1a from B. stearothermophilus LigA (pink), TfiLigA (light blue), EfaLigA (violet), and MtuLigA (dark blue) are color-coded and indicated separately for clarity. The bound NAD+ cofactor in the EfaLigA structure is shown in ball-and-stick representation.

NAD⁺-binding Site in MtuLigA—To structurally identify corresponding residues involved in NAD⁺ recognition on the basis of the EfaLigA structure and also for use in ligand docking calculations, we superposed the individual subdomains of MtuLigA onto the corresponding ones in the NAD⁺-bound Efa-LigA structure (Protein Data Bank code 1TAE [PDB] ). As expected, the NAD⁺ molecule fits well into the generated model (Fig. 4B). The interactions with the AMP part are conserved because of their similar conformations as described above. The nicotinamide moiety has stacking interactions with Tyr²⁹ and Tyr⁴² in the EfaLigA structure. The corresponding interactions are provided by Tyr³¹ and Phe⁴⁴, respectively. Mutating the former tyrosine (which is also highly conserved) in EcoLigA is known to abolish activity (43), whereas mutating the latter tyrosine (Phe⁴⁴ in MtuLigA) is known to reduce activity.

View larger version (26K): [in this window] [in a new window]   FIG. 4. A, conformation differences in the bound AMP in the structures with TfiLigA and MtuLigA. Some surrounding residues of Mtu-LigA (black lines) are indicated and labeled for clarity. Interaction of AMP (red sticks)in MtuLigA with the motif III Glu residue is indicated. The AMP moiety (green sticks) in TfiLigA is covalently linked in its co-crystal structure (interaction not indicated). The figure is shown in split stereo. B, stereo representation of some interacting residues (black lines) in the NAD+-binding site of MtuLigA generated by superposing and adjusting the conformation of residues in subdomain 1a of MtuLigA to the same orientation of the subdomain in the NAD+-bound structure of EfaLigA (Protein Data Bank code 1TAE [PDB] ). The AMP molecule in the MtuLigA structure is represented in red, and the bound NAD+ molecule in the EfaLigA structure is shown in pink. The binding mode predicted by the docking calculations with an inhibitor (compound 2 in this study) is shown in blue.

In silico docking analysis suggested an overlap of the binding sites of NAD⁺ and glycosyl ureides. We therefore evaluated by standard kinetics whether the compounds act competitively with NAD⁺ in the overall nick-sealing reaction in vitro. In the absence of the inhibitor, we determined a K_m of 1.56 µM for NAD⁺ in the presence of 10% Me₂SO in the assay mixture, which agrees well with previously reported data (10). In our inhibition studies, when the amount of NAD⁺ was increased up to 50 µM in the presence of increasing concentrations of compound 2 (0-20 µM) and a saturating DNA concentration (0.85 µM), the kinetics clearly indicated competitive inhibition of NAD⁺ by the compound (Fig. 5A), as also visualized in a double-reciprocal plot (Fig. 5B). Linear regression using the apparent K_m values leads to a K_i of 4.9 µM (Fig. 5C). These results strongly suggest that the binding sites of glycosyl ureides and NAD⁺ overlap with each other.

In Vivo Ligase Inhibition and Antibacterial Activities—To check the selectivity and specificity of the glycosyl ureides for NAD⁺ ligase in vivo, we chose pTrc99A-based systems in E. coli GR501 involving MtuLigA and T4 DNA ligase (34). This strain is known to harbor a temperature-sensitive lig251 mutation in LigA (35). It grows well at 30 °C, but growth is strongly delayed at 37 °C. This deficiency can be overcome, however, by complementing it with NAD⁺- or ATP-dependent ligase (11, 19, 35), which restores the growth of E. coli GR501 at elevated temperatures. This strain has therefore been useful in demonstrating the LigA specificity of inhibitors in vivo (19). We also reproduced earlier results (11, 35) that MtuLigA and T4 ligase complement the growth of the mutant strain at elevated temperatures for use in the in vivo assays. NAD⁺ ligase has also been reported to be essential for survival in the prominent human pathogen S. typhimurium LT2 strain (37). As yet another system for testing the specificity of the inhibitors in vivo and to determine whether the compounds act also against other NAD⁺-dependent ligases, we used the S. typhimurium LT2 strain and its DNA ligase-null derivative (TT15151) (38), which had been rescued with T4 ligase.

The much higher sensitivity of the E. coli GR501 strain harboring only the pTrc99A plasmid (Table IV) to the compounds compared with the corresponding ligase-rescued strains is attributed to the low residual ligase activity in the mutant strain compared with the growth-rescued strains, which possess a high copy number of the overexpressed ligase used to rescue them. This was also observed in the case of pyridochromanones (19). Consistent with the in vitro results, the MICs of the compounds (Table IV) were less for the strain rescued by Mtu-LigA and higher for the strain rescued by T4 ligase. The trend that the tested compounds were more selective for NAD⁺ ligase compared with the ATP-dependent ligase in E. coli GR501 was also seen in the case of the S. typhimurium system. The compounds exhibited more sensitivity to the Salmonella wild-type strain harboring the NAD⁺-dependent ligase compared with its ligase-deficient variant, rescued by the ATP-dependent ligase.

View larger version (15K): [in this window] [in a new window]   FIG. 5. Competitive inhibition of MtuLigA with respect to NAD+ by glycosyl ureides. A and B, activity of MtuLigA measured in the presence of increasing concentrations of compound 2 (0-20 µM) and NAD+ (0-50 µM). The double-reciprocal plot in B clearly indicates competitive binding between NAD+ and glycosyl ureides. C, linear regression plot of the inhibitor concentration versus Km(app). The Ki value is marked with an arrow.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Although we added NAD⁺ under the crystallization conditions, we observed density only for noncovalently bound AMP. The conformation of the cofactor in the current crystal structure mimics that of the AMP moiety in the NAD⁺-bound Efa-LigA structure. The only other corresponding cofactor-bound LigA structure is that of AMP-bound TfiLigA, where AMP is reported to be covalently linked to the motif I lysine in a different conformation (8).

It was suggested previously that a syn- to anti-conformational switch around the adenosine nucleoside of AMP is linked to the progression of the ligase reaction and that the active site is "serially remodeled" in the interactions with NAD⁺ and AMP (17, 41). Based on mutational analysis of the motif III Glu residue in EcoLigA (Glu¹⁸⁴ in MtuLigA), it was observed that contact with this Glu residue is essential in the third step of the reaction, but not in the second step. The syn-conformation exhibited by the moiety in the EfaLigA·NAD⁺ complex in the first step is replaced by the anti-conformation around the moiety in the AMP·TfiLigA complex in the second step of the reaction. In this step, contact with the motif III Glu residue is lost, and subsequently, the adenosine nucleoside moiety in AMP must undergo a conformational change again to syn to interact with this Glu residue in the third step (41). We would therefore expect to find AMP in at least two conformations when subdomain 1a is in the open state, viz. one in which it is covalently attached and the other in which it interacts with the motif III Glu residue. The present structure appears to have captured a snapshot of the syn-switched conformation of AMP after the covalent bond with the motif I lysine is broken in LigA. More structures of wild-type and mutant enzymes, especially with bound DNA substrate, will be very interesting in this context.

With multiple drug resistance spreading across the world, the identification of new classes of inhibitors with novel mechanisms is essential to keep pace with the adaptability of bacterial populations. NAD⁺ ligases are now proven novel targets, and additionally, no drug is as yet known to target them. We were therefore interested in identifying novel inhibitors for the enzyme.

View larger version (66K): [in this window] [in a new window]   FIG. 6. Bactericidal activity of compound 2. A, effect on growth as reflected in changes in A600 of S. typhimurium LT2 upon its exposure to compound 2 at 4-20 µg/ml, representing 0.5-2.5 times the MIC. The arrow indicates the point at which the compound was added. B, comparative viability between S. typhimurium LT2 (upper row) and its corresponding DNA ligase-null derivative (TT15151 (LigA-/T4 Lig+); lower row), rescued with a plasmid containing the gene for T4 ligase, as shown by surviving colony-forming units 4 h after addition of compound 2 to the growth medium. The cells were plated at dilution ratios of 10-4 on nutrient agar, and the indicated MICs correspond to that of S. typhimurium LT2 against the inhibitor. The control (C) and the amount of added inhibitor as multiples of MIC are also indicated.

Pyridochromanones, a different class of compounds, were identified recently as potent inhibitors of LigA (19). These compounds have nanomolar affinities for NAD⁺ ligases, and the most potent compounds could distinguish ATP ligases with >3 orders of magnitude. In contrast to the arylamino compounds, the binding site of pyridochromanones is thought to at least partially overlap with the NAD⁺-binding site. It was reported recently that MtuLigA is also sensitive to pyridochromanones, as expected, as indeed are NAD⁺ ligases from different sources (10, 19).

We used in silico docking against our in-house data base, with the cofactor-binding site of the ligase as the target, for the calculations. It is therefore conceivable that the compounds picked up are those that compete with the NAD⁺ moiety. In support of this hypothesis, the analysis of the docked complexes showed that the identified glycosyl ureides partially mimicked NAD⁺ interactions (Fig. 4B). For example, the O-benzyl moiety of compound 2 possesses a spatial disposition similar to that of the adenosine nucleoside, whereas the tetrahydrofuran moiety of the compound exhibits a spatial disposition similar to that of the ribose sugar of AMP. On the other hand, the docked pyridochromanone-3 contacts residues from subdomain 1a involved in NMN recognition (data not shown), consistent with the report of its binding site overlapping the NAD⁺-binding site. The in silico calculations agree well with the in vitro kinetic experiments; our studies demonstrate that glycosyl ureides acted competitively with NAD⁺ in the ligase reaction. This clearly suggests that the inhibitor and cofactor possess overlapping binding sites.

The glycosyl ureide-based inhibitors identified in this work were able to distinguish between NAD⁺- and ATP-dependent ligases quite well in vitro. They inhibited MtuLigA in the single digit micromolar range, whereas the major human DNA ligase I was inhibited in the 100-200 µM range (Table III). As control inhibitors, we used the antibiotics doxorubicin and chloroquine, which were reported previously to inhibit EcoLigA with IC₅₀ values of 1.3 and 53 µM, respectively (18). Although doxorubicin could not distinguish between NAD⁺ and ATP ligases, chloroquine exhibited exquisite specificity. Doxorubicin and chloroquine inhibited MtuLigA with IC₅₀ values of 5 and 46 µM, respectively (Table III). Analogous to the previous report, our results with MtuLigA also confirm that doxorubicin cannot distinguish between the ligase classes, whereas chloroquine is more specific for MtuLigA.

It is possible that compounds that inhibit the protein in vitro may affect bacterial growth through completely unrelated mechanisms such as nonspecific interference with a variety of essential cellular functions and disturbance of membrane integrity, etc. To exclude such other effects, we carried out assays against specific ligase-deficient E. coli and S. typhimurium strains (Table IV). Incidentally, the latter is also a major human pathogen. Our assays suggested that the observed antibacterial activities of glycosyl ureides are based on inhibition of the ligase enzyme. The ligase-deficient E. coli and S. typhimurium strains harboring MtuLigA, S. typhimurium LigA, temperature-sensitive EcoLigA, and T4 ligase, respectively, were used. The effectiveness of the compounds against them shows that these compounds can inhibit a variety of ligases, as observed in the case of pyridochromanones, most likely due to the conserved nature of the binding site in this class of enzyme.

In summary, we report the x-ray structure of the MtuLigA adenylation domain with bound AMP. We subsequently used structure-based computational approaches to identify glycosyl ureides as novel inhibitors of LigA. Our assay results demonstrate that these compounds represent a new class of inhibitors that can distinguish between NAD⁺- and ATP-dependent ligases and that can potentially be used to develop novel antibacterial therapies.

	FOOTNOTES

 
The atomic coordinates and structure factors (code 1ZAU) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).

^* The work was supported in part by Department of Biotechnology Grant SSP0139 and Council for Scientific and Industrial Research Network Grant CMM0017. This is Communication 6757 from the Central Drug Research Institute. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Recipient of junior and senior research fellowships from the Council for Scientific and Industrial Research.

^|| To whom correspondence should be addressed: Molecular and Structural Biology Div., Central Drug Research Institute, P. O. Box 173, Chattar Manzil, Mahatma Gandhi Marg, Lucknow-226001, India. Tel.:91-522-261-2411 (ext. 4442); Fax: 91-522-262-3405; E-mail: ravi_anitha{at}yahoo.com.

¹ The abbreviations used are: TfiLigA, T. filiformis LigA; EfaLigA, E. faecalis LigA; MtuLigA, M. tuberculosis LigA; MICs, minimum inhibitory concentrations; EcoLigA, E. coli LigA.

	ACKNOWLEDGMENTS

 
We thank Prof. T. P. Singh (All India Institute of Medical Sciences, New Delhi, India) for collection of a data set. The S. typhimurium wild-type LT2 strain and its DNA ligase-null derivative containing the T4 Lig⁺ plasmid pBR313/598/8/1b (TT15151) were kindly provided by Dr. J. R. Roth (University of Utah, Salt Lake City, UT). pTrc99A, the E. coli GR501 ligA^ts mutant, and the pTrc99A-T4 (pRBL) construct were kind gifts of Dr. R. P. Bowater (University of East Anglia, Norwich, UK). The human full-length DNA ligase I construct was kindly gifted by Dr. D. E. Barnes (Clare Hall Laboratories, Herfordshire, UK).

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