Nitric oxide synthase (NOS) enzymes synthesize nitric oxide, a signal for vasodilatation and neurotransmission at low concentrations and a defensive cytotoxin at higher concentrations. The high active site conservation among all three NOS isozymes hinders the design of selective NOS inhibitors to treat inflammation, arthritis, stroke, septic shock and cancer. Our crystal structures and mutagenesis results identified an isozyme-specific induced-fit binding mode linking a cascade of conformational changes to a new specificity pocket. Plasticity of an isozyme-specific triad of distant second- and third-shell residues modulates conformational changes of invariant first-shell residues to determine inhibitor selectivity. To design potent and selective NOS inhibitors, we developed the anchored plasticity approach: anchor an inhibitor core in a conserved binding pocket, then extend rigid bulky substituents toward remote specificity pockets, which become accessible upon conformational changes of flexible residues. This approach exemplifies general principles for the design of selective enzyme inhibitors that overcome strong active site conservation.
Nitric oxide (NO) is a small, diffusible, transient molecule produced from the amino acid L-arginine by three NOS enzymes1,2. The endothelial (eNOS) and neuronal (nNOS) NOS isozymes are constitutively expressed and Ca2+ regulated to provide NO for signaling, including vasodilatation, thermoregulation, neuroprotection and endocrine function. The Ca2+-insensitive inducible NOS isozyme (iNOS) is expressed in response to cytokines or pathogens, and produces NO at a high rate to kill bacteria, viruses and tumor cells. Insufficient NO bioavailability from eNOS and nNOS is associated with hypertension, impotence, atherosclerosis and cardiovascular disease, whereas excess NO from iNOS has been implicated in inflammation, rheumatoid arthritis, inflammatory bowel disease, immune-type diabetes, stroke, cancer, thrombosis and infection susceptibilities3,4. Overproduction of NO by iNOS (and nNOS) has also been linked to neurodegenerative disorders including Parkinson's and Alzheimer's diseases, as well as multiple sclerosis5. Thus, the development of iNOS-specific inhibitors is highly desirable.
The three NOS isozymes share a common modular architecture and conserved active site. The N-terminal catalytic oxygenase module (NOSox) binds cofactors heme and (6R)-5,6,7,8-tetrahydro-L-biopterin (H4B), substrate L-arginine, and a structural zinc ion across the dimer interface6,7,8,9,10,11,12,13,14,15. Upon calmodulin binding16, NOSox accepts electrons from the C-terminal reductase module17,18,19. The nearly complete amino acid conservation and structural similarity among the three NOS isozymes'6,7,8,9,10,11,12,13,14,15 active sites presents a significant challenge for the design of isozyme-specific inhibitors20,21. Moreover, as NO availability is controlled at the synthesis level for signaling or cytotoxicity, NOS isozymes are a paradigmatic system to address the challenges of designing isozyme-specific inhibitors despite conserved binding pockets.
NOS inhibitors selective for iNOS are rare, and they commonly exhibit limited selectivity or substantial toxicity22,23,24,25. In contrast, quinazoline26 inhibitors (1, 2, 3, 4, 5) and aminopyridine27,28 inhibitors (6, 7, 8, 9, 10, 11, 12) have good in vitro potency and selectivity for iNOS. In particular, the spirocyclic quinazoline AR-C102222 (3; Fig. 1) shows excellent selectivity over eNOS (3,000-fold) and exhibits significant protective, anti-inflammatory and antinociceptive activities in rodent models of adjuvant-induced arthritis, pancreatitis29, neuropathy, inflammation and postsurgical pain30. Thus, we have chosen to focus our structural studies on quinazoline and aminopyridine inhibitors.
Here, we combined mutagenesis, biochemistry, crystallography and drug design to elucidate the structural basis for the iNOS selectivity of some quinazoline and aminopyridine inhibitors. We demonstrate that plasticity of an isozyme-specific triad of residues distant from the active site modulates conformational changes of invariant residues nearby the active site to determine the exquisite selectivity of these inhibitors for iNOS. We have designed new potent and selective iNOS inhibitors by applying an 'anchored plasticity approach'. Selective inhibitors are designed with an inhibitor core anchored in a conserved binding pocket, and with rigid bulky substituents that extend to remote specificity pockets accessible upon conformational changes of 'plastic' protein residues. Fundamentally, this anchored plasticity approach is broadly applicable to the discovery of new inhibitors against enzyme families with strong active site conservation.
Inhibitor binding to iNOSox
Quinazoline inhibitors (1 and 2), spirocyclic quinazoline inhibitors (3, 4, 5) and aminopyridine inhibitors (6, 7, 8, 9, 10, 11, 12) are potent (half-maximal inhibitory concentration (IC50) from 10 nM to 1.2 μM) and selective (2.7-fold to 3,000-fold) inhibitors for iNOS over eNOS and nNOS (Fig. 1). These inhibitors all share a cis-amidine–derived core, but have different substituents or tails. To determine the basis for the exquisite iNOS potency of these inhibitors, we solved X-ray structures of mouse iNOSox bound to compounds 1, 2, 3, 4, 5, 6, 7, 8, 9, 10 and 12 and of human iNOSox bound to aminopyridine 9 (Methods and Supplementary Table 1 online).
Quinazolines 1, 2, 3, 4, 5 and aminopyridines 6, 7, 8, 9, 10, 11, 12 belong to different chemotypes but bind similarly in the iNOS active site heme pocket (Fig. 2a–d and Supplementary Fig. 1 online). The NOSox active site is lined by the heme, invariant Glu (Glu371/377, mouse/human iNOS numbering; Fig. 2e) and backwall residues (363–366/369–372; Supplementary Fig. 2 online). In all these inhibitor complexes, the cis-amidine moiety mimics the guanidinium group of substrate L-arginine by making bidentate hydrogen bonds to Glu and stacking with the heme. The interplanar angle between the inhibitor core and the heme is 6° for small compounds 1 and 6 but averages 13° for quinazolines 2, 3, 4, 5, and 20° for aminopyridines 7, 8, 9, 10. Compounds 1–8 make an extra hydrogen bond to Trp366/372 and pack more parallel to the heme plane than compounds 9, 10 and 12 (average interplanar angles of 13° and 26°, respectively), thus possibly explaining their improved potency in iNOS (Fig. 1 and Supplementary Fig. 1). Besides the hydrogen bonds to active site Glu and Trp366/372 and stacking with the heme, compounds 6 and 7 make no further interaction with the protein. However, both compounds are potent iNOS inhibitors, thus highlighting the importance of hydrophobic stacking and hydrogen bonding interactions.
The bulky and rigid tails of compounds 2, 3, 4, 5, 9, 10 and 12 all extend above heme propionate A (Fig. 2b–d) and pack with invariant residues Gln (Gln257/263), Arg (Arg260/266), Pro344/350, Ala345/351 and Arg382/388. Hydrogen bonds tether the extended inhibitor tails to invariant Tyr (Tyr341/347) and either Arg382/388 (compound 2) or a water molecule (compounds 3, 4, 5 and 12). We further explored permissive substitutions of compound 9 (Fig. 1). Changing the 4-methyl group in compound 9 into chlorine (compound 10) slightly decreased the inhibitor potency for all three isozymes. However, substitution with a 4-methoxy group (compound 11) improved potency three-fold for iNOS alone, thus contributing to increased isozyme selectivity (719-fold against eNOS, 339-fold against nNOS). Further elaboration of the piperidine acyl group to a substituted benzamide (compound 12) afforded excellent selectivity against eNOS (1,259-fold) and nNOS (82-fold). Our structural analysis thus suggests that both interactions of the inhibitor core with active site residues and of the inhibitor tail with residues outside the active site heme pocket mediate inhibitor binding.
To determine the roles of residues key to inhibitor binding, we measured the binding affinity and inhibitor potency of moderately selective compound 9 for several human iNOS mutant proteins (Table 1). Mutation of active site Glu into alanine had the most dramatic effect (Kd = 0.4 μM for wild type versus ≫100 μM for E377A mutant), thus revealing the crucial role of this invariant charged side chain in inhibitor binding and substrate binding31. The close match between binding affinity (Kd) in iNOSox and IC50 in full-length iNOS (Table 1) suggests that enzyme inhibition data reflect true binding affinity. Both Gln mutations (Q263A, loss of side chain; and Q263N, smaller side chain with similar functionality) result in slightly decreased iNOS affinity for compound 9 (three-fold and five-fold, respectively), thus corroborating the role of Gln in inhibitor tail binding. The human iNOS Y347F Y373F double mutant displayed a ∼10-fold decrease in inhibitor potency, thus revealing a key role for the Tyr hydrogen bond to the carbonyl of the inhibitor tail. Our combined structural and mutagenesis results thus suggest that heme stacking, as well as hydrogen bonds and hydrophobic interactions within and outside the active site, all contribute substantially to the binding of these inhibitors to iNOS.
A new Gln specificity pocket in iNOSox
Bulky inhibitors promote a cascade of conformational changes up to 20 Å away from the iNOSox active site, resulting in the creation of a new pocket. The superposition and analysis of the available iNOSox structure bound to the NOS reaction intermediate N-hydroxy-L-arginine (NOHA; Protein Data Bank (PDB) entry 1DWW, ref. 11) with our iNOS X-ray structures with small inhibitors (compounds 1, 6 and 7) and large-tailed inhibitors (2, 3, 4, 5, 8, 9, 10 and 12) reveals similar overall protein structures (0.4 Å average r.m.s. deviation for all Cα). In the active site pocket, the cis-amidine moieties of the quinazoline and aminopyridine inhibitors bind like the NOHA and L-arginine guanidinium groups7 (Fig. 2a–d and Supplementary Fig. 3a–c online) via heme stacking and anchoring hydrogen bonds to Glu. Yet outside this pocket invariant first-shell and second-shell residues adopt different side chain conformations in these complexes. To prevent collision with bulky inhibitors, the first-shell Gln side chain rotates around its χ1 and χ2 torsion angles, from a “Gln-closed” position (Fig. 2a) with a hydrogen bond to Tyr, to a “Gln-open” position (Fig. 2b–d) with hydrogen bonds to Arg (r.m.s. deviation values of 3.3 Å and 0.6 Å for Glu and Arg, respectively). Similarly, the Arg side chain rotates closer to second-shell residues Asp274/280 and Asn (Thr277/Asn283). Finally, side chain rotation of Arg382/388 (r.m.s. deviation value of 0.7 Å for Arg382/388) closer to Asp376/382 enhances hydrophobic interactions with the inhibitor tail.
Upon binding of bulky inhibitors, the coupled rotations of first-shell Gln and Arg initiate a cascade of conformational changes, which further propagate to second-shell residues. In the human iNOS Gln-open conformation, the conformational change of first-shell Arg induces rotation of second-shell Asn toward third-shell Phe286 and Val305 (Fig. 2d, Supplementary Fig. 3d and Supplementary Movie 1 online). Thus, the conformational plasticity of human iNOS second-shell Asn allows coordinated movements of first-shell Gln and Arg.
The correlated side chain rotations of Gln, Arg and Arg382/388 to accommodate the rigid bulky tails of compounds 2, 3, 4, 5, 9, 10 and 12, expose a new specificity pocket for enhanced inhibitor binding in iNOS. This “Gln specificity pocket” (Fig. 3a) extends from the active site heme pocket and is lined by residues Gln, Arg, Trp340/346, Tyr, Pro344/350, Ala345/351, Tyr367/373, Asp376/382 and Arg382/388 (Fig. 2b–d). All residues forming the iNOS Gln specificity pocket are strictly conserved among NOS isozymes, with one exception: iNOS Asp376/382, which hydrogen bonds to Arg382/388, is replaced by asparagine in eNOS (Supplementary Fig. 2). Notably, all previously reported NOSox structures present the Gln-closed conformation7,8,11,12,13,14,15 or a disordered Gln conformation10. We thus conclude that the Gln-open conformation and the associated cascade of conformational changes leading to the opening of the new Gln specificity pocket are favored or induced by the binding of quinazoline and aminopyridine inhibitors bearing a rigid and extended tail.
Isozyme differences in inhibitor binding
In iNOSox and eNOSox, the binding modes for moderately selective aminopyridine 9 are dramatically different despite common overall protein structures and active sites (r.m.s. deviation of 0.8 Å between eNOS and iNOS). The inhibitor aminopyridine cores bind similarly in the active site heme pocket of both isozymes (Fig. 3a,b and Supplementary Fig. 3e,f). In eNOS, the bidentate hydrogen bonds to active site Glu anchor the aminopyridine core almost parallel to the heme plane and place the inhibitor bulky tail between the eNOSox heme propionates and close to Val337/346/352 (bovine eNOS/mouse iNOS/human iNOS numbering; Supplementary Fig. 2). This position of the rigid inhibitor tail induces slight side chain rotations of Trp448/457/463 on the propionate A side, and of Tyr476/485/491 on the propionate B side. Thus, both the core and the extended tail of aminopyridine compound 9 interact more closely with the eNOS heme than with the iNOS heme. Notably, in the eNOS–compound 9 complex, the heme is more planar than in iNOS, and a second inhibitor molecule now occupies the pterin site and mimics H4B binding7 (Supplementary Fig. 2 and Supplementary Discussion online). Thus, isozyme-specific differences in both the L-arginine and H4B binding sites of the eNOS complex with aminopyridine 9 could contribute to eNOS inhibition.
In eNOS, binding of compound 9 does not induce the opening of the Gln specificity pocket. Invariant Tyr hydrogen bonds to Gln, thus preventing its hydrophobic interaction with the inhibitor tail (Fig. 3b and Supplementary Fig. 3e). As a consequence, the eNOS complex with compound 9 exhibits the Gln-closed conformation, and the Gln specificity pocket is not observed (Fig. 3a). In contrast, the iNOS complex with compound 9 exhibits the Gln-open conformation, thereby allowing the inhibitor tail to bind in the Gln specificity pocket (Fig. 3b and Supplementary Fig. 3f). Not only first-shell Gln and Arg, but also second-shell Asn, present different conformations in the two complexes.
What prevents the Gln-closed to Gln-open conversion and the opening of the Gln specificity pocket in eNOS? First, we propose that Gln gates the Gln specificity pocket and must adopt the Gln-open side chain conformation to allow inhibitor access. Mutation of Gln into alanine (Q246A, loss of side chain) only marginally enhanced inhibitor potency of compound 9 for eNOS, but did not achieve the potency observed for wild-type iNOS (Table 1). The Q246N mutation had no effect. Thus, removal or rotation of the Gln gate is important, but not sufficient, for potent compound 9 binding to eNOS. Second, we followed, in eNOS, the cascade of conformational changes observed in iNOS upon binding of bulky inhibitors (Fig. 3b). An iNOS-like binding mode for compound 9 in eNOS would induce conformational changes of first-shell Gln and Arg, and second-shell Asn (Fig. 4). However, in eNOS, bulky Leu290 and β-branched rigid Ile271 in the third shell block the side chain rotation of second-shell Asn. Consequently, this Asn conformation prevents the conformational changes of Gln and Arg necessary for the opening of the Gln specificity pocket (Figs. 3 and 4). The triad of second-shell (Asn) and third-shell (leucine and isoleucine) residues are the only nearby residues that are not conserved among NOS isozymes (Fig. 2e and Supplementary Fig. 2). To test our hypothesis for the key role of third-shell isozyme-specific residues, we made the human iNOSox F286I V305L double mutant to mimic the corresponding eNOSox residues. This double mutant exhibited the UV-visible spectral properties of wild-type human iNOSox, thus indicating that these mutations do not perturb enzyme structure or the heme electronic environment. Binding affinity of this mutant iNOSox enzyme for compound 9 dramatically dropped to beneath detection levels, as observed for wild-type eNOS (Table 1), whereas binding of nonselective inhibitor 6 was unaffected (data not shown). Furthermore, the X-ray structure of the human iNOSox double mutant cocrystallized with compound 9 (55-fold molar excess) reveals a Gln-closed conformation and the absence of bound inhibitor (Supplementary Fig. 3g). Our results thus demonstrate the crucial role of third-shell isozyme-specific residues in inhibitor binding and provide a structural basis for the exquisite iNOS specificity of large-tailed quinazoline and aminopyridine inhibitors.
Determinants for inhibitor selectivity
Based on our combined structural and mutagenesis analyses, we propose that differences in the plasticity of second- and third-shell residues between iNOS and eNOS modulate conformational changes of invariant first-shell residues to determine inhibitor selectivity. Together, our mutational and structural results suggest that the Gln specificity pocket accounts for the excellent iNOS selectivity of the bulky aminopyridine and quinazoline inhibitors. In turn, opening of this pocket depends not only on conformational changes of invariant first-shell Gln and Arg, but also on the plasticity of isozyme-specific second-shell Asn (Fig. 4). This hypothesis is supported by several observations. First, the potent small NOS inhibitors (1, 6 and 7), which do not induce the Gln-open conformation, show poor selectivity for iNOS27,28 (Figs. 1 and 2; Supplementary Fig. 1). Second, the bulky, but less rigid, tail of compound 8, which neither hydrogen bonds to Tyr nor induces Arg side chain rotation (Supplementary Fig. 1), binds less deeply in the Gln pocket and exhibits only modest selectivity for iNOS32. Third, eNOS third-shell residues Ile271 and Leu290 block binding of compound 9 in the Gln pocket, as evidenced by our structural and mutagenesis results on the human iNOS double mutant (Table 1 and Supplementary Fig. 3g). Fourth, bulky quinazoline and aminopyridine inhibitors present moderate selectivity against nNOS (7- to 80-fold more selective for iNOS; Fig. 1). We predict that inhibitor binding in nNOS will induce similar side chain rotations for first-shell Gln and Arg, and partial rotation of second-shell Asn toward third-shell Phe506 and Leu525 (Fig. 4). The substitution of small Val305 in human iNOS with bulkier Leu525 in human nNOS will likely restrict side chain rotation of second-shell Asn. We thus conclude that the plasticity of the isozyme-specific triad tunes the inhibitor selectivity by controlling the conformational changes of invariant first-shell Gln and Arg and the formation of the new Gln specificity pocket that can be effectively used for inhibitor binding.
Our results on NOS support an anchored plasticity approach for the design of selective inhibitors. Given a protein of known structure, a set of matching protein sequences (from different species or isoforms), and a binding pocket for a common class of ligand (substrate, cofactor, inhibitor, metabolite or other), we propose the following procedure for selective inhibitor design: (i) identify anchor points for binding in the conserved pocket; (ii) locate variations in sequence and structure outside this pocket; (iii) delineate pathways connecting anchor points to variations (using solvent-accessible channels, for example); (iv) design selective inhibitors that incorporate both a core for anchored binding and extended rigid substituents oriented to exploit protein plasticity along pathways leading to variations (Supplementary Fig. 4 online). The core provides binding affinity via anchoring in nonspecific binding pockets, while the extended substituents determine inhibitor selectivity. Fundamentally, this anchored plasticity approach does not necessarily require serendipitous identification of isoform-specific residue movement. Furthermore, it is readily applicable to key enzyme families, such as kinases, that exhibit overlapping specificities.
Design and synthesis of selective iNOS inhibitors
Our combined results allowed us to propose the anchored plasticity approach for the design of specific inhibitors exploiting conserved binding sites coupled to distant isozyme-specific residues via cascades of conformational changes. This approach differs from other methods to design new NOS inhibitors that only exploit differences in first-shell residues15,33,34. To test the applicability of our results, we rationally designed potent and selective iNOS inhibitors starting from a new and unexploited template with a 5,7-fused heterobicyclic amidine core (compound 13; Supplementary Methods online). This inhibitor is potent but not selective (IC50 = 0.2 μM for iNOS and eNOS, and 0.07 μM for nNOS). Addition of small substituents (compounds 14 and 15) increases the potency but does not significantly affect selectivity (Fig. 1). In contrast, addition of the bulkier and rigid isoquinolinyloxy-methyl tail (compound 16) results in a substantial increase in potency and selectivity for iNOS over eNOS35 (Fig. 1). We determined the X-ray structures of mouse iNOSox bound to compounds 14 and 16, and of human eNOSox bound to compound 15 (Fig. 5 and Supplementary Fig. 4). In all structures, the inhibitor core packs above the heme and makes bidentate hydrogen bonds to invariant active site residue Glu and to Trp366 main-chain carbonyl. As seen for bulky quinazoline compounds (2, 3, 4, 5) and aminopyridine compounds (9, 10, 11, 12), the extended tail of compound 16 packs with first-shell residues Gln, Arg, Pro344, Ala345 and Arg382, and induces the Gln-open conformation (Fig. 5). These results thus demonstrate the applicability of our anchored plasticity approach for the design of new potent and selective NOS inhibitors.
The combined inhibitor screening, structural and mutagenesis results on iNOS and eNOS provide new insights for structure-based design of selective inhibitors. In iNOS, but not in eNOS, binding of inhibitors bearing an extended rigid tail is associated with a cascade of adaptive conformational changes, beginning with movements of invariant first-shell residues and leading to the opening of the new Gln specificity pocket for enhanced potency and selectivity. The conformational changes reveal a specificity pocket that is separate from an otherwise conserved active site heme pocket and not observed in previously determined NOSox X-ray structures. Indeed, others had predicted that NOS inhibitors larger than L-arginine would perturb the hydrogen bonding network with Tyr and Gln and extend into the substrate access channel20,21, in distinct contrast to our results. Here, we show that an isozyme-specific triad of second-shell (Asn) and third-shell residues in NOS tunes the plasticity of invariant first-shell residues (Gln and Arg), and thus determines the exquisite selectivity (125- to 3,000-fold) of the long-tailed aminopyridine, quinazoline and bicyclic thienooxazepine inhibitors for iNOS over eNOS. The most selective spirocyclic quinazoline compound (3; ref. 26) and aminopyridine compound (12; ref. 28 and this study) also show good potency in whole cells (IC50 of 0.9 and 1.9 μM, respectively) and in vivo (ID50 of 3.0 and 1.8 μmol kg−1, respectively) activity assays (Supplementary Fig. 5 online). Further studies will be required to test the in vivo potency of the bicyclic thioenooxazepine compounds. Nevertheless, these highly selective NOS inhibitors are promising tools to investigate specific iNOS-mediated effects both in vivo and in vitro. More specifically, these results on iNOS and eNOS inhibitor structures can be applied to future inhibitor design for the treatment of inflammation, cancer and other diseases, while reducing the risks of disrupting the crucial activity of eNOS in maintaining blood pressure.
The opening of an isozyme-specific pocket in iNOS results from permitted conformational changes of conserved first-shell and second-shell residues upon inhibitor binding. This selective induced-fit movement depends on plasticity differences in conserved residues located far from the substrate binding pocket. Exploiting such changes in flexibility to improve inhibitor potency is likely applicable to other key enzyme systems with overwhelmingly conserved active sites, including human immunodeficiency virus (HIV) reverse transcriptase36, aldose reductase37, cyclooxygenases38,39 and kinases40. In NOS, isozyme-specific second-shell and third-shell residues influence the plasticity of invariant first-shell residues and, thus, determine the isozyme selectivity. In all these enzyme families, the new binding pocket is distinct from the active site and becomes accessible after adaptive conformational changes of conserved residues. Whether binding of the inhibitor induces the new conformation (induced-fit), or the inhibitor simply selects from different protein conformations in equilibrium (conformational selection, as seen in antigen recognition41), the result is an improved protein-inhibitor interaction.
Our results for prototypic NOS isozymes appear generally applicable to understanding and tuning binding affinity and specificity of enzyme inhibitors. Our structures demonstrate that differential residue plasticity can be exploited for conformational changes that create new specificity pockets suitable for the design of isozyme-specific inhibitors. Together, these results have implications for drug discovery, and demonstrate that X-ray crystallography is crucial for revealing subtle, but important, differences in residue plasticity between closely related isozymes (for example, iNOS versus eNOS) or between homologous enzymes from different organisms. The significant roles of second- and third-shell residues in determining the plasticity of conserved first-shell residues will add to the existing challenges of accurately modeling induced fit in proteins. Here, we show that systematic structural results combined with mutagenesis can identify selectivity-determining side chain differences distant from the active site, thus overcoming the barriers that active site conservation poses for isozyme-specific drug design.
Expression and purification of NOSox proteins.
Mouse iNOSox Δ65 (residues 66–498) and human eNOSox (65–492, homologous to mouse iNOSox Δ78) were expressed and purified as described12,42. Bovine eNOSox (53–492; homologous to mouse iNOSox Δ65) was obtained via trypsinolysis of holo-eNOS, which was expressed and purified as published43. Human iNOSox wild-type and mutant constructs (82–508; homologous to mouse iNOSox Δ78) were expressed and purified as described10 with slight modifications. Wild-type and mutant human iNOSox proteins were produced in a pT7 Escherichia coli expression vector based on pET-11a vector (Novagen). BL21(DE3) cells (Stratagene) were grown in the presence of ampicillin at 37 °C until reaching a cell density corresponding to A600 of 0.5–0.8. The culture was then induced with 0.5 mM IPTG, 6.125 mg l−1 ferric citrate and 450 μM ∂-amino levulinic acid, and grown for 3 d at 20 °C before harvesting. Pellets were resuspended in buffer A (10 mM sodium phosphate pH 7.0, 0.1 M NaCl, 1 mM L-arginine, 2 mM DTT, 10 μM H4B), sonicated extensively and subsequently loaded on a heparin column (GE Healthcare). Protein was eluted in a single step by adding 0.3 M NaCl to buffer A. Fractions containing NOSox were concentrated, aliquoted and stored at −80 °C.
Mutations were introduced into the complementary DNAs of human iNOSox and full-length NOS within expression vectors by using the QuickChange site-directed mutagenesis kit (Stratagene). Mutagenic oligonucleotides were designed according to the manufacturer's instructions, and mutations were confirmed by sequencing.
Synthesis of compounds.
Binding affinity, inhibitory potency and in vivo activity.
The inhibitory potency (IC50 in μM) was determined in full-length wild-type and mutant NOS in the presence of cofactors (5 μM FAD, 5 μM FMN, 200 μM BH4, 1 mM CaCl2, 25 μg ml−1 calmodulin) and substrates (3 μM [3H]arginine, 1 mM NADPH) as described26. Inhibitors were pre-incubated with NOS proteins in the presence of cofactors and NADPH for 1 h before L-arginine addition.
Binding affinity (Kd) of human iNOSox for compounds 6 and 9 was measured by imidazole displacement. Briefly, iNOSox was incubated for 2 h with a matrix of concentrations of imidazole and compound 6 or 9. Samples were scanned by UV-visible spectroscopy between 270 and 700 nm. Binding affinity was determined by plotting the absorbance difference A428 – A396 as a function of the imidazole concentration. The decrease in apparent imidazole affinity as the concentration of compound increased was used to determine the binding affinity for compounds 6 and 9.
Compound 12 was further tested for inhibition of NO production in an intact cell assay44 and in a rat model of inflammation via lipopolysaccharide (LPS)-induced nitric oxide production26, as described below. Compound 12 or vehicle was given orally to male conscious rats at time zero. Blood samples were taken after 2, 4 and 6 h. Total plasma nitrite and nitrate concentrations, indicative of NO production, were determined by using the Griess reaction after reduction of nitrate to nitrite by nitrate reductase. Oral administration of compound 12 to rats led to dose-dependent inhibition of elevated plasma NO levels (ID50 = 1.8 μmol kg−1, dose that reduces the NO production by 50%) measured 4 h after LPS administration (Supplementary Fig. 5). All in vivo studies were approved by the AstraZeneca Ethical Review Committee and were conducted under license from the UK Home Office.
Crystallization, data collection and refinement.
Mouse and human iNOSox, and bovine and human eNOSox, were cocrystallized (with 2–5 mM inhibitor) by vapor diffusion as described7,8,10,12. Crystals grew overnight (crystallization pH was 7, 7.2, 6.5 and 6.0 for mouse iNOSox, human iNOSox, bovine eNOSox and human eNOSox, respectively). All crystals were cryocooled after transfer to cryoprotectant solution (mouse iNOSox, 30% glycerol; human iNOSox, 100% MgSO4; bovine eNOSox, 15% glycerol; human eNOSox, 15% 2-methyl-2,4-pentanediol) in a cold nitrogen gas stream. Data were collected at 100 K at the Cornell High Energy Synchrotron Source (CHESS) beamline F1 (compound 1), Stanford Synchrotron Radiation Laboratory (SSRL) beamlines 7-1 (compounds 2, 3, 4 and 8, 9, 10), 9-1 (compounds 5 and 7) and 9-2 (compound 6), MAX-LAB beamline I711 (compounds 14 and 16) and European Synchrotron Research Facility (ESRF) beamlines FIP (compound 15) and ID2 (compound 12). All diffraction datasets were processed using DENZO/SCALEPACK programs45. The crystal structures of human iNOSox10, mouse iNOSox7 and human eNOSox10, with ligands, water and cofactors removed, were used as starting models for molecular replacement with AMoRe46 for human iNOSox, mouse iNOSox, and both bovine and human eNOSox, respectively. During crystallographic refinement of protein structures, the heme geometry, like the amino acid geometry, is restrained by a set of parameters (bond lengths, angles, dihedrals) derived from small-molecule studies and high-resolution protein structures47. These parameters and their associated weights can strongly influence the resulting refined geometries for heme in protein structures48, especially those determined at lower resolution. Hence, direct comparisons of heme distortions from published X-ray structures refined in different ways can be problematic. Parameters from the HICUP database47 were used for refinement of the heme and pterin cofactors (HEC and H4B, respectively). Inhibitors were fit into Sigmaa-weighted49 Fo – Fc electron density maps, which confirmed the expected unmodified chemical structures. Overall structures were obtained by iterative cycles of refinement with CNS50 and manual fitting with O51, Xfit52 or Coot53 (Supplementary Table 1). All superimpositions were performed for residues within a sphere of 10 Å around the inhibitor with the CCP4 program LSQKAB54. The r.m.s. deviations were calculated for all superimposed atoms.
Protein Data Bank: Coordinates and structure factors were deposited with the following accession codes: 3E7I (1), 3E6O (2), 3E7T (3), 3E65 (4), 3E6T (5), 3E67 (6), 3E6N (7), 3E68 (8), 3E7M (mouse iNOSox with 9), 3E7G (human iNOSox wild type with 9), 3EJ8 (human iNOSox F286I L305V mutant with 9), 3E7S (bovine eNOSox with 9), 3E6L (10), 3EAI (12), 3EBD (14), 3EAH (human eNOSox with 15) and 3EBF (16).
Note: Supplementary information and chemical compound information is available on the Nature Chemical Biology website.
Protein Data Bank
We thank K. Panda (Cleveland Clinic) and S. Ghosh (Cleveland Clinic) for preparation of the mouse iNOSox and bovine eNOSox proteins used in this study. Part of this work is based on research conducted at CHESS, SSRL, ESRF and MAX-LAB (Lund University). This work was supported in part by US National Institutes of Health grants (E.D.Getzoff and D.J.S.), and by the Skaggs Institute for Chemical Biology (E.D.Garcin).
The Quicktime movie illustrates the cascade of conformational changes that occur in human iNOSox upon aminopyridine compound 9 binding (Quicktime, 582 kB).
About this article
Nature Structural & Molecular Biology (2011)