As the second essential enzyme of the folate biosynthetic pathway, the potential antimicrobial target, HPPK (6-hydroxymethyl-7,8-dihydropterin pyrophosphokinase), catalyzes the Mg2+-dependant transfer of pyrophosphate from the cofactor (ATP) to the substrate, 6-hydroxymethyl-7,8-dihydropterin. Recently, we showed that 8-mercaptoguanine (8-MG) bound at the substrate site (KD ∼13 µM), inhibited the S. aureus enzyme (SaHPPK) (IC50 ∼ 41 µM), and determined the structure of the SaHPPK/8-MG complex. Here we present the synthesis of a series of guanine derivatives, together with their HPPK binding affinities, as determined by SPR and ITC analysis. The binding mode of the most potent was investigated using 2D NMR spectroscopy and X-ray crystallography. The results indicate, firstly, that the SH group of 8-MG makes a significant contribution to the free energy of binding. Secondly, direct N9 substitution, or tautomerization arising from N7 substitution in some cases, leads to a dramatic reduction in affinity due to loss of a critical N9-H···Val46 hydrogen bond, combined with the limited space available around the N9 position. The water-filled pocket under the N7 position is significantly more tolerant of substitution, with a hydroxyl ethyl 8-MG derivative attached to N7 (compound 21a) exhibiting an affinity for the apo enzyme comparable to the parent compound (KD ∼ 12 µM). In contrast to 8-MG, however, 21a displays competitive binding with the ATP cofactor, as judged by NMR and SPR analysis. The 1.85 Å X-ray structure of the SaHPPK/21a complex confirms that extension from the N7 position towards the Mg2+-binding site, which affords the only tractable route out from the pterin-binding pocket. Promising strategies for the creation of more potent binders might therefore include the introduction of groups capable of interacting with the Mg2+ centres or Mg2+ -binding residues, as well as the development of bitopic inhibitors featuring 8-MG linked to a moiety targeting the ATP cofactor binding site.
Citation: Chhabra S, Barlow N, Dolezal O, Hattarki MK, Newman J, Peat TS, et al. (2013) Exploring the Chemical Space around 8-Mercaptoguanine as a Route to New Inhibitors of the Folate Biosynthesis Enzyme HPPK. PLoS ONE 8(4): e59535. https://doi.org/10.1371/journal.pone.0059535
Editor: Anil Kumar Tyagi, University of Delhi, India
Received: December 30, 2012; Accepted: February 15, 2013; Published: April 2, 2013
Copyright: © 2013 Chhabra et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: The authors have no support or funding to report.
Competing interests: The authors have declared that no competing interests exist.
Antibiotic resistance is rapidly emerging as one the most significant health challenges of this century . In Europe alone, 25,000 deaths were reported in 2007 as a result of antimicrobial resistance, with an estimated cost of €1.3 billion per year . Compounding this problem is the fact that antibiotic drug discovery is on the decline –a reflection of the considerable challenges associated with identifying both viable new targets as well as drugs to target them, but also a general lack of interest from large pharmaceutical companies. Most alarmingly, Methicillin-resistant S. aureus (MRSA) has evolved globally into a range of strains with varying phenotypes . It has become resistant to both oxacillin and erythromycin, and resistance to levofloxacin is reported to be on the rise . Community-acquired MRSA (caMRSA) is a relatively recent threat among patients without conventional risk factors. The epidemic USA300 strain of caMRSA is exceptionally virulent due to high levels of alpha toxin and the phenol-soluble modulins ; remarkably, it accounts for over half of all illnesses caused by the entire range of S. aureus species.
Logical targets for antimicrobials are essential enzymes that are unique to microorganisms, of which those of the folate biosynthesis pathway are prime examples. Folate is essential for the growth of all living cells, with the reduced form, tetrahydrofolate, used in the biosynthesis of thymidine, glycine and methionine. However, only bacteria and lower eukaryotes synthesize folate de novo; mammals and higher eukaryotes obtain it from their diet by active transport. The folate pathway enzymes, dihydropteroate synthase (DHPS) and dihydrofolatereductase (DHFR) are the targets for the sulfa drugs and Trimethoprim, respectively, which are used to treat diseases such as malaria, pneumocystis pneumonia (PCP), and, more recently, caMRSA infections.
It is well established that point mutations in pathogenic DHPS and DHFR genes contribute to widespread resistance to the aforementioned drugs. Recently, structure-based investigations have identified new inhibitors of DHPS that bind to the pterin site, remote from the sulpha drug site , as well as a new lead candidate for inhibiting the quadruple mutant DHFR enzyme conferring resistance in Plasmoidium falciparum . These studies exemplify the application of modern drug discovery approaches to old targets as a means of generating potential new antibiotics.
An alternative approach to combating resistant isolates is the development of inhibitors for as-yet-to-be-drugged enzymes within the folate pathway. Hydroxymethyl-pterin pyrophosphokinase (HPPK) is one such enzyme, responsible for catalysing the transfer of a pyrophosphate group from the ATP to the pterin substrate, 6-hydroxymethyl-7,8-dihydropterin (HMDP) (Fig. 1A). HPPK structures from many microbial sources have been solved (E. coli, H. influenza, S. pneumoniae, S. cerevisiae, Y. pestis, F. tularensis and S. aureus) –. All have a thioredoxin-like fold containing the binding sites for both the substrate and the ATP cofactor. X-ray structural studies have revealed that major conformational changes, particularly in loop L3, occur throughout the catalytic cycle . Structural and kinetic studies – have also established that ATP binds (KD = 2.6–4.5 µM) prior to the substrate, which binds with sub-micromolar affinity. The pterin stacks between two highly conserved aromatic residues (Tyr or Phe) and both the substrate and cofactor are fixed in position by a multitude of hydrogen bonds; in total, they interact with 26 separate residues.
A) HPPK catalysis. B) Known inhibitors of HPPK.
While much is known about the structure of HPPK, very few small molecule inhibitors have been developed (Fig. 1B). The gem-dimethyl- and 7-phenethyl-substituted pterin analogues, 3 and 4, were reported to be HPPK inhibitors over three decades ago by Woods . They have since been crystallized bound to the E. coli enzyme , , and 3 was utilized in a number of structural studies aimed at understanding the catalytic trajectory of HPPK , . Recent inhibitor design has included the production of bitopic ligands featuring pterin coupled to adenosine via mono- through to tetra-phosphate linkers (5), with the longest linker providing the best affinity (KD = 0.47 µM) and inhibition (IC50 = 0.44 µM) . Bitopic ligands featuring a more drug-like piperidine bridge (6) , or gem-dimethyl pterin in combination with a piperidine or amide-sulphone linker (7  and 8 ), have also been reported, however no gain in potency has been achieved (8 did, however, display a novel binding mode in which the base was flipped).
Very recently, we showed that the simple guanine derivative, 8-mercaptoguanine (8-MG), is able to inhibit HPPK from S. aureus (KD ∼11 µM, IC50 = 41 µM) through interaction with the HMDP pocket . Binding was found to be non-competitive with either the cofactor (ATP) or its non-hydrolyzable analogue, AMPCPP, as judged by both surface plasmon resonance (SPR) and isothermal titration calorimetry (ITC) analysis. A 1.65 Å resolution X-ray crystal structure revealed a high degree of stereo-electronic complementarity between 8-MG and the HMDP-binding pocket, together with an extensive network of hydrogen bonds, accounting for the unusually high binding affinity of the small 8-MG molecule (183 Da) (Fig. 2A, B). Most intriguingly, NMR analysis on the 8-MG/AMPCPP ternary complex provided compelling evidence that the SH group of 8-MG interacts with the L3 loop of SaHPPK, locking it onto a “closed” conformation above the active site .
A–C) Structure of SaHPPK (PDB:1QBC) in complex with 8-MG. A–B Intermolecular interactions between 8-MG and SaHPPK. C) Surface representation of SaHPPK showing the bound 8-MG (blue) overlayed with the closed loop L3 (green) and the bound AMPCPP as observed in the EcHPPK/HMDP/AMPCPP (PDB:1Q0N) complex. D) Ribbon representation of the loop structure of several EcHPPK structures overlayed with SaHPPK (yellow) in complex with 8-MG (red) to illustrate the range of conformations in loops L2 and L3. The interaction of the Trp89 (brown) and the phenethyl inhibitor (cyan) is highlighted (PDB:1DY3) and the position of the HMDP (pink) and AMPCPP (pink) from EcHPPK/HMDP/AMPCPP (PDB:1Q0N). Images were produced using the UCSF Chimera package (www.cgl.ucsf.edu/chimera).
Herein, we report the results of a study interrogating the chemical space available within the active site of SaHPPK and the chemical developability of 8-MG as a HPPK inhibitor. As part of this study, a series of N7- and N9-substituted 8-MG analogues have been synthesized, and their interaction with SaHPPK examined using a combination of SPR, ITC, NMR spectroscopy and X-ray crystallography, in order to determine which of these positions are amenable to the lead optimization extension strategy. Additionally, a small number of C8-sustituted analogues have been studied to allow further assessment of the relative importance of the SH group of 8-MG to the overall binding characteristics of this compound.
Results and Discussion
Structure-based Hypotheses and Design of 8-MG Analogues
As shown in Figure 2A and B, the pyrimidine heterocycle (ring A) of 8-MG is “perfectly tailored” for the pterin-binding pocket of HPPK, as evidenced by full saturation of all hydrogen donors and acceptors, and the sandwiching of the ring between the two phenylalanine residues, Phe54 and Phe123. This ring was therefore considered of limited value as a site for further chemical modification aimed at improving binding affinity and potency. Instead, our efforts focused on exploring the effects of substitution at the N7, C8 and N9 positions of ring B.
Predicting the likely outcome of substituent changes/additions to 8-MG is complicated by the fact that the L2 and L3 catalytic loops in HPPK can adopt a diverse range of conformations, leading to drastic changes in the microenvironment of ring B (Fig. 2D) (loops L2 and L3 are also inherently dynamic in the apo and cofactor-bound states on the micro to millisecond timescale , ). The 8-MG/SaHPPK X-ray structure (PDB: 3QBC) itself displays an extended L3 loop conformation , and is therefore limited in guiding modelling and structure-based design of 8-MG analogues from the N7, C8 and N9 positions. In the first instance, we therefore chose to explore the effect of replacing the mercapto group of 8-MG with a variety of other substituents (compounds 10a–10f, Table 1) in order to probe tolerance to substitution at this position. In part, this was performed in order to test our hypothesis (based on earlier 15N chemical shift and NMR relaxation measurements ), that Gly90 or Trp89 at the tip of the L3 loop form a favorable contact to the mercapto group at the C8 position, which serves to fix L3 into a “closed” conformation resembling that observed in the ternary complex of E. coli HPPK, HMDP and AMPCPP (PDB: 1Q0N) (Fig. 2C) .
Our substituent choices for the N9 position were inspired by the structure of the ternary complex of E. coli HPPK with the phenethyl HMDP analogue (2-amino-6-methoxy-7-methyl-7-phenethyl-7,8-dihydropterin) and AMPCPP (PDB:1DY3) . Within this structure, the phenyl ring of the substrate analogue makes two hydrophobic intermolecular interactions; one edge-on to Trp89 in loop L3 and the other to the side-chain of Leu45 (Val46 in S. aureus) in loop L2. From an overlay of 1DY3 and 3QBC (Fig. 2D), it was conjectured that the appendage of a hydrophobic group to the N9 position of 8-MG could afford similarly favorable interactions with side-chains present in loops L2 and L3. Four hydrophobic substituents of increasing size (CH3, C2H5, CH2C6H5, CH2CH2C6H5) were thus chosen for investigation (compounds 15a–15d, Table 1). In order to deliver a stronger binder, it was recognized that any favorable interaction(s) afforded by these groups would have to more than compensate for the loss of the hydrogen-bond between the N9 H group and the Val 46 carbonyl in the SaHPPK/8-MG complex (Fig. 2A, B).
Analysis of the SaHPPK/8-MG crystal structure revealed a water-filled pocket proximal to the N7 position (Fig. 2B, C). Given the hydrophilic nature of this region, it was postulated that attachment of a suitable polar substituent might enhance binding through provision of additional interactions with the polar side-chains and/or bound waters present, coupled with entropically-favorable water displacement. A small series of 8-MG analogues featuring alcohol, amine and guanidinium pendants attached to N7 were therefore included within our test set (compounds 21a–21e).
Synthesis of 8-MG Analogues
8-(Methylamino)guanosine, 9, synthesized as described in the literature , was hydrolyzed using 1 M HCl to afford the first of the test compounds, 8-(methyamino)guanine (10a). All other derivatives with C8 substitution (10b–10f) were commercially sourced.
The synthetic routes to N9-substituted guanines are well-established, in part because of the use of the N9-substituted drugs, acyclovir and ganciclovir, in the treatment of herpes virus infections . An expedient synthesis of the N9-methyl guanine from 2-amino-6-chloropurine, exploiting the N9-directing effect of the chloro-substituent, has been previously reported and involved alkylation with methyl iodide followed by hydrolysis to install the oxo group . We found this method could also be employed to provide ethyl, benzyl and 2-phenethyl substituents at the N9-position (Fig. 3). Transformation of 13a to the 8-mercapto derivative 15a had been previously been demonstrated by bromination at C8 to provide 14a , followed by treatment with thiourea . We found this similar transformation could be applied to our other derivatives providing the brominated analogues 14b–d and the SH-containing target compounds, 15b–d.
The N7-substituted 8-MG analogues were prepared via alkylation of 8-bromo-N2-acetylguanine (18), formed in two steps from guanosine (16) according to a literature method ,  (Fig. 4). Benzylation of 18 at N9 with benzyl bromide has been reported previously under conditions that required no base . We found that alkylation with other reagents proceeded well when the pH of the reaction mixture was adjusted to 3. These reactions generally yielded a ca. 1∶1 mixture of N7- and N9-substituted isomers, from which the desired N7-alkylated intermediates were isolated following either silica gel or preparative-scale reverse-phase HPLC. Installation of the 8-mercapto group was then achieved by reaction with sodium thiosulfate in the presence of a catalytic quantity of aluminium-trichloride , and the target 8-MG analogues isolated following removal of any protecting groups under the appropriate conditions (Fig. 4). Compound 21e, featuring an ethyl guanidinium group, was prepared from the amino analogue, 21c, through reaction with pyrazole carboxamidine (Fig. 5).
All final compounds were purified by preparative HPLC to >95% purity.
SPR and ITC Analysis of Binding
Initially, the binding of each of the test compounds to SaHPPK was quantitatively assessed using SPR (Fig. S1, S2). Compared with the parent compound (8-MG), SPR data for the synthesized analogues did not appear to be compromised by their solubility in aqueous buffer at the maximum concentration used (126 µM). Moreover, all sensorgrams (Fig. S1 and S2) were of high quality and consistent with near perfect 1∶1 stoichiometric binding of analogues. Table 1 lists the estimated equilibrium dissociation constants (KD). It is clear from the data that replacement of the 8-mercapto group is highly detrimental to binding; compounds 10a and 10b, featuring -NHCH3 and -SCH3 groups at the C8 position, did not bind SaHPPK at all (although binding of compound 10a was detected (KD = 108 µM) in the presence of saturating amounts of ATP), whilst all other C8-substiuted analogues exhibited 15–20-fold lower KD values than 8-MG. This supports the hypothesis that the 8-mercapto group of 8-MG aids binding through the formation of one or more favorable interactions with SaHPPK. The precise nature of this/these interaction(s) remain unclear, however the considerably inferior binding affinity of the C8-OH analogue (10e) suggest that it is unlikely to be a simple hydrogen bond to a loop L3 residue, as we speculated might be the case earlier .
The 8-MG derivatives with simple hydrophobic substituents at the N9 position (15a–d) exhibited 10-20-fold lower affinities for SaHPPK, indicating that any potential positive contribution to binding afforded by these groups is not sufficient to make up for the loss of the intramolecular N9-H·Val46 carbonyl hydrogen bond. Extension of the 8-MG scaffold via the N9 position, therefore, does not appear to be a promising strategy for lead optimization.
Of the N7-substituted 8-MG analogues, compound 21a, with an ethyl alcohol substituent, displayed comparable affinity to 8-MG (KD ∼12 µM), whilst the analogues with amine and guanidinum pendants (21c–21e) displayed slightly weaker binding to SaHPPK; the carboxylate pendant-bearing analogue, 21b, did not bind. This indicates that addition of substituents at the N7 position are tolerated, and that extension from this ring position is likely the most promising avenue for future development of more potent 8-MG analogues. It should be noted, however, that in contrast to 8-MG, the binding of compounds 21a, 21c and 21d was found to be 10-15-fold weaker under saturating ATP conditions, suggesting extensions from ring B into the space towards the Mg2+ centres and the ATP binding site leads to competitive binding with the ATP cofactor. Any future lead optimization studies will need to bear this in mind.
To corroborate the ligand binding affinities determined by SPR, and to determine the enthalpic and entropic contributions to the free energy of binding, ITC experiments were performed for selected compounds (21a and 21c–21e) (Table 2, Fig. S3). KD values in the 17–35 µM range were obtained, in excellent agreement with the values obtained by SPR. Interestingly, compound 21a, which yielded a similar KD value to 8-MG (16.7 µM vs. 12.8 µM), showed a lower binding enthalpy than 8-MG, but its binding to SaHPPK was associated with a much lower entropic penalty. Given that 21a has more rotatable bonds than 8-MG, this may suggest that binding of the latter may lead to a greater degree of immobilization of the catalytic loops within SaHPPK. To investigate the factors contributing to the free energy of binding, we solved the structure of SaHPPK in complex with 21a.
X-ray Structure of SaHPPK in Complex with Compound 21a
Attempts were made to co-crystallize each of the strongest binding compounds (21a and 21c–21e) with SaHPPK, however diffraction quality crystals could only be obtained for compound 21a. These provided excellent quality electron density data, and a high-resolution X-ray structure (1.85 Å) of the SaHPPK/21a binary complex (Fig. 6A) determined via molecular replacement (crystal data and details of the data collection and refinement are provided in Table 3). A head-to-tail protein dimer was found in the asymmetric unit, similar to that observed for the earlier SaHPPK/8-MG structure (PDB: 3QBC) , with the ligand bound to the pterin sites of both protein monomers. The ethyl alcohol pendant projects into the space leading towards the Mg2+ binding site, making two hydrogen bond contacts with a pair of bound water molecules (Fig. 6B). Presumably, these interactions in part compensate for the loss of the hydrogen bond between the N9-H of 8-MG and the backbone carbonyl of Val46, which occurs as a consequence of the tautomerization accompanying alkylation at the N7 position. A water molecule found in the cavity under N7 in the SaHPPK/8-MG structure has been displaced in the SaHPPK/21a structure, and there is a tightly bound water between the hydroxyethyl and Asp97 which orients Asp97 in a similar position to that found of Asp97 of the EcHPPK/AMPCPP/HMDP structure (where Mg2+ sits). Superposition of the SaHPPK/21a structure with that of EcHPPK/AMPCPP/HMDP (PDB: 1QON)  indicates that if Mg2+ ions and ATP were simultaneously bound, the oxygen of the hydroxyethyl pendant of 21a is displaced by ∼1 Å and would lie only 1.5–1.6 Å from one of the metal ions, which is considerably less than the Mg-O bond length observed in the 1QON structure (2.1 Å) and sterically unfavorable (Fig. 6C). This is the likely reason for the cofactor and metal competition observed for 21a.
(A) 2Fo-Fc electron density map of the pterin binding site, contoured at 2.0 sigma showing density for 21a. B) Detail of intermolecular interactions from 21a to SaHPPK and two bound waters. C) Superposition of the EcHPPK/HMDP/AMPCPP (PDB:1QON) structure. Selected loops and sidechains of EcHPPK are shown (blue) along with the bound HMDP (plum) and AMPCPP (pink), two bound magnesium ions (dark cyan) and the oxygen atoms of coordinating waters (grey). SaHPPK/21a is shown colored as in B) with selected sidechains shown (green). Images were produced using the UCSF Chimera package (www.cgl.ucsf.edu/chimera) and PyMOL (Delano Scientific).
Heteronuclear NMR Analysis of Compound 21a Binding to SaHPPK
Titration of 21a into 15N-labelled apo (data not shown) and magnesium-loaded SaHPPK enzyme led to broadening and disappearance of several, common peaks in the 2D 15N HSQC NMR spectrum (Fig. 7A, B), which is characteristic of the intermediate exchange timescale, and indicates that binding of 21a is not magnesium-dependent. This is similar to what was observed for the binding of 8-MG to SaHPPK (Fig. 7A) and is consistent with the fact that density characteristic of magnesium was not observed in the X-ray crystal data of the SaHPPK/21a complex.
A–B) Binding of 8-MG and 21a to magnesium bound SaHPPK are very similar. C–D) Binding of 8-MG and 21a to the AMPCPP bound SaHPPK are very different. Figures A and C are adapted from data in Figure 6A in . The concentration of 15N-labelled SaHPPK was ∼100 µM in all cases. The concentration of magnesium, AMPCPP, 8-MG and 21a was 10 mM, 1 mM, 0.6 mM and 0.6 mM respectively. The assignment of selected substrate site peaks are shown to highlight the effects of binding of the two compounds on the NMR spectra. The sidechain Hε1–Nε1 peak of Trp89 is labelled as W89sc.
The observed intermediate exchange regime for the binding of 8-MG and 21a is possibly dictated in part by the slow µs-ms timescale motion of loop L3 , . While the spectra (Fig. 7A, B) appear to be very similar, however, closer inspection reveals that the sidechain Hε1–Nε1 peak of Trp89 (in loop L3) is only perturbed in the 8-MG bound spectrum (Fig. 7A). This is mechanistically interesting and may indicate that this region of loop L3, adjacent to the substrate-binding loop L2, is involved in binding of 8-MG but not 21a. Following on from this, the observed larger entropic penalty to the free energy of binding of 8-MG as compared to 21a (Table 2) may derive in part from this increase in loop L3 rigidity in the presence of 8-MG, whilst the more favorable enthalpic contribution likely reflects the formation of the N9-H Val46 intermolecular hydrogen bond (as observed in the X-ray structure). Reduced loop L3 involvement in 21a binding, on the other hand, is likely a result of the loss of the N9-H Val46 intermolecular hydrogen bond (due to tautomerization from substitution at N7), which would reduce any dampening of the adjacent loop L2 dynamics. Ligand-induced loop L2 and L3 dampening can be detected and investigated directly by NMR, but in order to do this the NMR timescale needs to be shifted out from the intermediate regime. This was previously accomplished by binding 8-MG to the AMPCPP bound SaHPPK enzyme . The results of our heteronuclear NMR spin relaxation studies reported therein revealed dampening of loop L2 and L3 motion on the fast timescale compared to the apo or the AMPCPP (or ATP)-bound SaHPPK enzyme. Unfortunately, it was not possible to investigate the enzyme dynamics in the same way for the binding of 21a as it was found to be competitive with AMPCPP. From a comparison of the X-ray structures of the 21a/SaHPPK binary complex (this work) with our previous 8-MG/SaHPPK binary structure , the expulsion of a bound water underneath N7 is likely to be thermodynamically favorable for the binding of 21a, and in line with the observed reduced entropic penalty. This may be the reason for the reduced entropic penalty associated with binding of the 21a–c series as a whole; see Table 2.
Repeating the titration in the presence of a saturating amount of the cofactor analogue, AMPCPP (KD = 3 µM), led to broadening of the same pterin site signals and the peak corresponding to the sidechain of Trp89 displayed very little change, in accord with the lack of involvement of this residue in 21a binding, as described previously. In accordance with the SPR data, however, cross peaks in slow exchange, characteristic of formation of a ternary complex (observed in our earlier study of the interaction of 8-MG with SaHPPK in the presence of AMPCPP (Fig. 7C)) were absent (Fig. 7D), indicating that binding of 21a to SaHPPK is competitive with AMPCPP.
8-MG represents a promising scaffold for the potential development of a new antibiotic drug targeting the folate pathway enzyme, HPPK. This study has shown that the 8-mercapto group plays a pivotal role in binding, and ought to be maintained in any future lead optimization studies. Extension from the N9 position within ring B leads to a dramatic loss of affinity and is therefore not a viable site for chemical modification. Substitution at the N7 position, however, is tolerable, as exemplified by N7-hydroxyethyl-8-MG (21a), which was found to bind SaHPPK with comparable affinity to the parent compound. An important caveat is that extension into the space surrounding the N7 atom leads to competitive binding with the ATP cofactor. To provide a meaningful enhancement in potency, future studies will therefore need to focus on the development of N7 pendants that interact strongly with the residues surrounding this pocket. This could include the introduction of groups to bind to the absolutely conserved metal-binding residues, Asp95 and Asp97, within the apo form of the enzyme. An alternate route to an increase in potency could involve changing the nature of ring B of the 8-MG core such that the N9-H Val46 H bond is maintained whilst still allowing extension from the N7 position towards the highly conserved metal-binding residues. We are currently investigating this approach.
Compared to the reported bitopic inhibitors for HPPK , , , both 8-MG and 21a are less potent, yet they have better ligand efficiencies (KD ∼10 µM over 12 and 15 heavy atoms, respectively, compared to KD ∼ 3 µM over 40+ heavy atoms). 8-MG could potentially be linked to adenosine to provide a bitopic ligand with considerably enhanced affinity, though problems associated with linking two subsite binders as a route to higher affinity have been well documented , . Ultimately, incremental step-wise chemical evolution of the 8-MG scaffold in a more conventional manner may prove the most efficient route to developing an inhibitor with superior pharmacodynamic and pharmacokinetic properties.
Finally, it is worth noting that it has recently been shown that 8-MG can also bind to the pterin pocket in DHPS, the adjacent, downstream enzyme to HPPK . The chemical strategies described herein may therefore prove beneficial for the design of more potent DHPS inhibitors based on the 8-MG scaffold, and perhaps even for the development of agents capable of inhibiting multiple enzymes within the folate biosynthesis pathway.
Chemistry - General Methods
Melting points were determined on a Mettler Toledo MP50 melting point system and are uncorrected. The abbreviation dec. indicates that the compound decomposed at the specified temperature. 1H and 13C NMR spectra were recorded on a Bruker Ultrashield 400 Plus at 400 MHz and 101 MHz, respectively. Analytical HPLC was performed on a Waters Alliance 2690 fitted with a Waters 5996 PDA detector and a Phenomenex Luna C8 column (5 µm, 100 Å, 150 × 4.60 mm). Analyses were conducted using a gradient of 0 to 64% acetonitrile in water over 10 min with 0.1% trifluoroacetic acid (TFA) throughout. Preparatory HPLC was performed on a Waters Prep LC 4000 system fitted with a Waters 486 Tunable Absorbance Detector and either a Phenomenex Luna C18 (10 µm, 100 Å, 250 × 30 mm) column or a Phenomenex Luna C8 (10 µm, 100 Å, 50 × 21.2 mm) column. Low resolution mass spectrometry was performed on an Agilent 6120 single quadrapole LCMS system using electrospray ionization. High resolution mass spectrometry was performed on a Waters Premier XE time-of-flight mass spectrometer using electrospray ionization.
Chemistry - Synthesis
A solution of 8-(methylamino)guanosine 9 (50 mg, 0.20 mmol) in 1 M HCl (10 mL) was refluxed for 2 h, then cooled to rt (room temperature). The precipitate was collected by filtration and resuspended in water (5 mL). This mixture was made basic by drop wise addition of 1 M NaOH whereupon the precipitate dissolved. Reverse phase chromatography (C18, 1% TFA in water) provided the title compound as a white solid (30 mg, quantitative). Mp 252–257°C (dec.), 1H NMR (400 MHz, D2O) δ 2.66 (s, 3H).13C NMR (101 MHz, D2O) δ 164.2, 163.6, 162.5, 157.5, 116.2, 30.0. LRMS (ESI): m/z: 181.1 ([M+H]+100%). HRMS (ESI): observed m/z: 181.0837; calculated m/z: 181.0832 [M+H]+.
A solution of 2-amino-6-chloropurine (1.00 g, 5.90 mmol) in DMF (10 mL) was treated with ethyl iodide (472 µL, 5.89 mmol) and K2CO3 (815 mg, 5.89 mmol). After stirring for 15 h at rt the solution was evaporated to dryness under reduced pressure and 2-amino-N9-ethyl-6-chloropurine isolated by silica gel chromatography (CHCl3/MeOH, 95∶5). This material was refluxed in 1 M HCl (20 mL) for 2 h then cooled to rt. The resulting precipitate was collected by filtration, affording the title compound as a white powder (527 mg, 50%). 1H NMR (400 MHz, DMSO-d6) δ 12.03 (s, 1H), 9.34 (s, 1H), 7.58 (s, 2H), 4.15 (q, J = 7.3 Hz, 2H), 1.43 (t, J = 7.3 Hz, 3H). 13C NMR (101 MHz, DMSO-d6) δ 155.8, 153.1, 149.4, 136.7, 107.2, 34.0, 14.2. LRMS (ESI): m/z: 180.1 [M+H]+ (100%).
A solution of 2-amino-6-chloropurine (1.00 g, 5.90 mmol) in DMF (10 mL) was treated with benzyl bromide (700 µL, 5.89 mmol) and K2CO3 (815 mg, 5.89 mmol), and stirred for 15 h at rt. The intermediate 2-amino-N9-benzyl-6-chloropurine was isolated and hydrolyzed as described for the preparation of 13b, to provide the title compound as a white powder (1.30 g, 90%). 1H NMR (400 MHz, DMSO-d6) δ 11.79 (s, 1H), 9.18 (s, 1H), 7.41–7.29 (m, 7H), 5.35 (s, 2H). 13C NMR (101 MHz, DMSO-d6) δ 155.8, 153.8, 150.0, 137.3, 135.3, 129.0, 128.4, 127.8, 108.8, 47.5. LRMS (ESI): m/z: 242.2 [M+H]+ (100%).
A solution of 2-amino-6-chloropurine (1.00 g, 5.90 mmol) in DMF (10 mL) was treated with 2-phenethyl bromide (798 µL, 5.89 mmol) and K2CO3 (815 mg, 5.89 mmol), and stirred for 15 h at rt. The intermediate 2-amino-N9-(2-phenethyl)-6-chloropurine was isolated and hydrolyzed as described for the preparation of 13b to provide the title compound as a white powder (1.35 g, 90%). 1H NMR (400 MHz, DMSO-d6) δ 11.78 (s, 1H), 8.89 (s, 1H), 7.34–7.15 (m, 7H), 4.34 (t, J = 7.3 Hz, 2H), 3.15 (t, J = 7.3 Hz, 2H).13C NMR (101 MHz, DMSO-d6) δ 155.5, 153.6, 149.7, 137.1, 136.9, 128.6, 128.6, 126.8, 108.4, 45.5, 34.1.LRMS (ESI): m/z: 256.2 [M+H]+ (100%).
A solution of compound 13b (212 mg, 0.820 mmol) in glacial acetic acid (15 mL) was treated with N-bromosuccinimide (211 mg, 1.19 mmol). After stirring for 15 h at rt, the solution was poured into a mixture of ice (50 g) and water (100 mL). The precipitate was filtered, washed with water and methanol, then dried to provide the title compound as a yellow powder (122 mg, 58%). 1H NMR (400 MHz, DMSO-d6) δ 10.66 (s, 1H), 6.58 (s, 2H), 3.96 (q, J = 7.2 Hz, 2H), 1.25 (t, J = 7.2 Hz, 3H). 13C NMR (101 MHz, DMSO-d6) δ 155.5, 153.8, 152.0, 120.2, 116.8, 31.4, 14.5. LRMS (ESI): m/z: 258.1 [M+H]+ (100%).
A solution of compound 13c (455 mg, 1.89 mmol) in glacial acetic acid (30 mL) was treated with N-bromosuccinimide (453 mg, 2.55 mmol). After stirring for 15 h at rt, the product was isolated using the procedure described for 14b, providing the title compound as a yellow powder (384 mg, 65%). 1H NMR (400 MHz, DMSO-d6) δ 10.72 (s, 1H), 7.38–7.15 (m, 5H), 6.60 (s, 2H), 5.16 (s, 2H). 13C NMR (101 MHz, DMSO-d6) δ 155.6, 154.0, 152.2, 135.8, 128.6, 127.7, 126.6, 120.8, 115.0, 45.4. LRMS (ESI): m/z: 320.1 [M+H]+ (100%).
A solution of compound 13d (1.21 g, 4.57 mmol) in glacial acetic acid (90 mL) was treated with N-bromosuccinimide (1.10 g, 6.20 mmol). After stirring for 15 h at rt, the product was isolated using the procedure described for 14b, providing the title compound as a yellow powder (914 mg, 60%). 1H NMR (400 MHz, DMSO-d6) δ 10.68 (s, 1H), 7.49–6.89 (m, 5H), 6.59 (s, 2H), 4.15 (t, J = 7.8 Hz, 2H), 3.00 (t, J = 7.8 Hz, 2H). 13C NMR (101 MHz, DMSO-d6) δ 155.5, 153.8, 151.9, 137.3, 128.6, 128.4, 126.6, 120.7, 115.0, 43.9, 34.3. LRMS (ESI): m/z: 334.2 [M+H]+ (100%).
A solution of compound 14b (102 mg, 0.40 mmol) and thiourea (60 mg, 0.80 mmol) in EtOH (5 mL) was refluxed for 15 h. The solvent was removed in vacuo and the residue purified using reverse phase chromatography (C18, isocratic: 0.1% TFA in water) to afford the title compound as an off-white solid (51 mg, 60%). Mp 240–244°C (dec.), 1H NMR (400 MHz, DMSO-d6) δ 12.71 (s, 1H), 10.87 (s, 1H), 6.66 (s, 2H), 4.01 (q, J = 7.1 Hz, 2H), 1.23 (t, J = 7.1 Hz, 3H). 13C NMR (101 MHz, DMSO-d6) δ 155.8, 153.1, 149.4, 136.7, 107.2, 34.0, 14.2. LRMS (ESI): m/z: 212.1 [M+H]+ (100%), HRMS (ESI): observed m/z: 212.0603 [M+H]+; calculated m/z: 212.0601 [M+H]+, RP-HPLC: tR 5.26 min, >98%.
A solution of compound 14c (110 mg, 0.34 mmol) and thiourea (131 mg, 1.70 mmol) in EtOH (5 mL) was refluxed for 15 h. The solvent was removed in vacuo and the residue purified using reverse phase chromatography (C18, isocratic: 0.1% TFA in water) to afford the title compound as an off-white solid (65 mg, 70%). Mp>300°C (dec.), 1H NMR (400 MHz, DMSO-d6) δ 12.87 (s, 1H), 10.94 (s, 1H), 7.41–7.15 (m, 5H), 6.64 (s, 2H), 5.21 (s, 2H). 13C NMR (101 MHz, DMSO-d6) δ 164.7, 154.1, 150.8, 150.0, 136.4, 128.3, 127.3, 127.2, 103.7, 44.9. LRMS (ESI): m/z: 274.1 [M+H]+ (100%), HRMS (ESI): observed m/z: 274.0770 [M+H]+; calculated m/z: 274.757 [M+H]+, RP-HPLC: tR = 7.31 min, >95%.
A solution of compound 14d (500 mg, 1.50 mmol) and thiourea (228 mg, 3.00 mmol) in EtOH (5 mL) was refluxed for 15 h. The solvent was removed in vacuo and the residue purified using reverse phase chromatography (C18, isocratic: 0.1% TFA in water) to afford the title compound as an off-white solid (260 mg, 60%). Mp 291–295°C (dec.), 1H NMR (400 MHz, DMSO-d6) δ 12.76 (s, 1H), 10.90 (s, 1H), 7.35–7.21 (m, 5H), 6.65 (s, 2H), 4.18 (t, J = 7.8 Hz, 2H), 2.99 (t, J = 7.8 Hz, 2H). 13C NMR (101 MHz, DMSO-d6) δ 164.1, 154.0, 150.8, 149.9, 138.0, 128.5, 128.4, 126.4, 103.6, 43.0, 33.1. LRMS (ESI): m/z: 288.2 ([M+H]+100%), HRMS (ESI): observed m/z: 288.09 [M+H]+; calculated m/z: 288.0914 [M+H]+, RP-HPLC: tR = 7.9 min, >98%.
To a suspension of N2-acetyl-8-bromoguanine (18) (100 mg, 0.37 mmol) in DMF (1 mL) was added 2-bromoethanol (50 µL, 0.70 mmol) and DIPEA (32 µL, 0.20 mmol). The reaction was heated at 100°C for 24 h with periodic addition of DIPEA in order to maintain the pH between 3 and 4. The solution was diluted with water (5 mL) and purified and subjected to reverse phase chromatography (C18, 0–4% ACN with 0.1% TFA in water) to isolate the title compound as a white solid (30 mg, 26%). 1H NMR (400 MHz, DMSO-d6) δ 4.31 (t, J = 5.6 Hz, 2H), 3.72 (t, J = 5.6 Hz, 2H), 2.16 (s, 3H).13C NMR (101 MHz, CDCl3) δ 21a3.5, 156.5, 151.5, 147.4, 131.3, 113.7, 59.8, 49.4, 23.7. LRMS (ESI): m/z: 315 [M+H]+, (100%), 321 [M+H]+ (100%).
To a suspension of N2-acetyl-8-bromoguanine (18) (1.00 g, 3.70 mmol) in dry DMF (5 mL) under N2 was added DIPEA (1.30 mL, 7.40 mmol) and methyl bromoacetate (386 µL, 4.10 mmol). The solution was stirred for 20 h at rt, the solvent removed in vacuo, and the residue was coevaporated with methanol (3×). The residue was chromatographed on silica gel (MeOH/DCM, 1∶19) to provide the title compound as a white solid (255 mg, 20%). 1H NMR (400 MHz, DMSO-d6) δ 7.95 (s, 1H), 5.02 (s, 2H), 3.70 (s, 3H), 2.16 (s, 3H). 13C NMR (101 MHz, DMSO-d6) δ 213.5, 168.1, 154.7, 148.9, 147.9, 140.1, 119.6, 52.5, 44.1, 23.7. LRMS (ESI): m/z: 345.9 [M+H]+ (100%), 347 [M+H]+ (100%).
A solution of N2-acetyl-8-bromoguanine (18) (500 mg, 1.85 mmol), DIPEA (960 µL, 5.60 mmol) and N-(2-bromoethyl)phthalimide (560 mg, 1.85 mmol) in DMF (5 mL) were heated at 100°C under N2 for 15 h. The solvent was removed in vacuo, and the residue was coevaporated with methanol (3×). The crude mixture was purified using silica gel chromatography (petroleum spirits/ethylacetate/methanol, 1∶1∶4) to provide the title compound as a white solid (205 mg, 25%). 1H NMR (400 MHz, DMSO-d6) δ 8.00–7.59 (m, 4H), 4.50 (t, J = 7.8 Hz, 2H), 4.04 (t, J = 7.8 Hz, 2H), 2.15 (s, 3H). 13C NMR(101 MHz, DMSO-d6) δ 213.5, 167.3, 156.4, 151.4, 147.3, 134.5, 131.3, 130.5, 123.1, 113.8, 45.7, 37.3, 23.7. LRMS (ESI): m/z: 445 [M+H]+ (100%), 446 ([M+H]+ (100%).
A mixture of N2-acetyl-8-bromoguanine (18) (810 mg, 2.98 mmol), N-(3-bromopropyl)phthalimide (1.10 g, 4.03 mmol), DIPEA (1.60 mL, 9.00 mmol) in DMF (10 mL) was refluxed at 100°C overnight. The solvent was evaporated in vacuo, diluted with water (50 mL) and extracted with chloroform (3 × 50 mL). The pooled organic phases were dried over MgSO4, then evaporated in vacuo. The residue was purified by silica gel chromatography (MeOH/CHCl3, 1∶19) providing the title compound as a white solid (276 mg, 20%). 1H NMR (400 MHz, DMSO-d6) δ 7.93–7.70 (m, 4H), 4.34 (t, J = 7.0 Hz, 2H), 3.63 (t, J = 7.0 Hz, 2H), 2.27–2.04 (m, 5H). 13C NMR (101 MHz, DMSO-d6) δ 213.4, 167.8, 156.3, 151.4, 147.4, 134.3, 131.6, 130.1, 122.9, 113.5, 44.7, 34.6, 28.6, 23.6. LRMS (ESI): m/z: 459 [M+H]+ (100%), 460 [M+H]+ (100%).
To a solution of N2-acetyl-8-bromo-7-(2-hydroxyethyl)guanine (19a) (10 mg, 0.03 mmol) in water (4 mL) and acetonitrile (2 mL) was added sodium thiosulfate (10 mg, 0.10 mmol) and aluminium chloride (0.02 mmol). The solution was refluxed for 24 h, then 1M HCl added and the solution stirred for a further 2 h. The solutionwas subjected to reverse phase chromatography (C18, isocratic 0.1% TFA in water) to isolate the title compound as a white powder (5 mg, 69%). Mp>300°C, 1H NMR (400 MHz, DMSO-d6) δ 10.91 (s, 2H), 6.54 (s, 5H), 4.79 (t, J = 5.7 Hz, 3H), 4.25 (t, J = 6.7 Hz, 6H), 3.64 (dd, J = 6.7, 5.7 Hz, 6H).13C NMR (101 MHz, DMSO-d6) δ 164.1, 154.1, 151.4, 149.5, 105.3, 58.5, 46.4. LRMS (ESI): m/z: 228.1 [M+H]+ (100%), HRMS (ESI): observed m/z: 226.039 [M-H]-; calculated m/z: 226.0404 [M-H]-, RP-HPLC: tR4.14 min, >98%.
2-(8-Mercaptoguanin-7-yl)acetic acid- (21b).
To a solution of methyl-8-bromo-(N2-acetylguanin-7-yl)acetate (19b) (95 mg, 0.28 mmol) in water (4 mL) and acetonitrile (2 mL) was added sodium thiosulfate (200 mg, 1.10 mmol,) and aluminium chloride (0.02 mmol). The solution was refluxed for 2 days, filtered and resuspended in water/methanol/dioxane (2∶1∶4), and the pH of the solution was brought to 13 by adding 1 M NaOH. The solution was stirred at 50°C for 2 h, then subjected to reverse phase chromatography (C18, isocratic 0.1% TFA in water) to isolate the title compound as a white solid (15 mg, 20%). Mp 247–253°C (dec), 1H NMR (400 MHz, DMSO-d6) δ 10.99 (s, 1H), 6.62 (s, 1H), 4.87 (s, 1H).13C NMR (101 MHz, DMSO-d6) δ 168.6, 165.0, 154.2, 151.4, 149.2, 105.0, 45.4. LRMS (ESI): m/z: 242 [M+H]+ (100%), HRMS (ESI): observed m/z: 242.0341 [M+H]+; calculated m/z: 242.0342 [M+H]+, RP-HPLC: tR = 4.23 min, >95%.
To a solution of of N-2-(8-bromo-N2-acetylguanin-7-yl)ethylphthalimide (19c) (160 mg, 0.36 mmol) in water (12 mL) and acetonitrile (8 mL) was added with sodium thiosulfate (447 mg, 1.80 mmol) and aluminium chloride (0.02 mmol). The reaction was refluxed for 2 days, solvent removed in vacuo and the residue resuspended in methanol (1 mL) and hydrazine hydrate (12 µL, 0.36 mmol). The solution was stirred for 15 h, the subjected to reverse phase chromatography (C18, isocratic: 0.1% TFA in water) to provide the title compound as a white solid (10 mg, 32%). Mp 280–287°C (dec), 1H NMR (400 MHz, DMSO-d6) δ 8.22 (s, 1H), 7.85 (s, 2H), 6.78 (s, 2H), 4.41 (t, J = 6.1 Hz, 1H), 3.16 (t, J = 6.1 Hz, 1H).13C NMR (101 MHz, DMSO-d6) δ 164.6, 154.5, 151.7, 150.1, 105.1, 42.5, 38.5. LRMS (ESI): m/z: 227.1 [M+H]+ (100%), HRMS (ESI): observed m/z: 225.0564 [M-H]-; calculated m/z: 225.0564 [M-H]-,RP-HPLC: tR = 2.9 min, >98% (gradient).
A suspension of N-3-(N2-acetylguanin-7-yl)propylphthalimide (19d) (150 mg, 0.33 mmol) in water (12 mL) and acetonitrile (8 mL) was added sodium thiosulphate (400 mg, 1.60 mmol) and aluminium chloride (0.02 mmol). The reaction was refluxed for 2 days. After cooling the mixture was concentrated to dryness under reduced pressure. The reaction was refluxed for 2 days, solvent removed in vacuo and the residue resuspended in methanol (1 mL) and hydrazine hydrate (12 µL, 0.36 mmol). The reaction was stirred for 15 h at rt and the the mixture subjected to reverse phase chromatography (C18, isocratic: 0.1% TFA in water) to provide the title compound as a white solid (12 mg, 44%). Mp 239–243°C (dec.), 1H NMR (400 MHz, D2O) δ 4.41 (t, J = 6.6 Hz, 2H), 3.08 (t, J = 6.6 Hz, 2H), 2.22 (t, J = 6.6 Hz, 2H). 13C NMR (101 MHz, D2O) δ 163.0, 154.4, 153.0, 150.1, 106.0, 42.0, 36.3, 26.5. LRMS (ESI): m/z: 241.1 [M+H]+ (100%), HRMS (ESI): observed m/z: 241.0942 [M+H]+; calculated m/z: 241.0827 [M+H]+, RP-HPLC: tR = 3.76 min, >98%.
A mixture of 7-(2-aminoethyl) 8-mercaptoguanine (21c) (10 mg, 0.04 mmol) and pyrazolecarboxamidine (7 mg, 0.10 mmol) in DMF was stirred at 50°C for 2 days. The resulting mixture was concentrated to dryness under reduced pressure. The crude product was purified using reverse phase chromatography (C18, isocratic: 0.1% TFA in water) to provide the title compound as a white solid (5.00 mg, 45%). Mp 256–262°C (dec), 1H NMR (400 MHz, DMSO-d6) δ 11.18 (s, 1H), 7.64 (t, J = 6.2 Hz, 1H), 6.75 (s, 2H), 4.29 (t, J = 6.4 Hz, 2H), 3.47 (t, J = 6.4 Hz, 2H). 13C NMR (101 MHz, DMSO-d6) δ 164.4, 156.9, 154.3, 151.5, 149.9, 104.8, 45.7, 42.8. LRMS (ESI): m/z: 269.1 [M+H]+ (100%), HRMS (ESI): observed m/z: 269.0936; calculated m/z: 269.0928 [M+H]+, RP-HPLC: tR4.23 min, >98%.
Surface Plasmon Resonance (SPR)
All SPR binding experiments were performed as described previously . The only difference was the use of a sulfhydryl reactive maleimide-activated biotin derivative (Thermo Scientific, 1-biotinamido-4-(4′-[maleimidoethylcyclohexane]-carboxamido)butane. The maleimide-activated biotin was attached to the exposed surface cysteine residue of SaHPPK according to manufacturer’s instructions. The resulting site-specific biotinylated protein was immobilized onto the sensor chip surface using the Biotin capture kit (GE Healthcare). All analogues were serially diluted (either 2- or 3-fold from 126 µM down to 1.5 µM) in SPR binding buffer (50 mM HEPES, 150 mMNaCl, 1 mM TCEP, 0.05% (v/v) Tween-20, 10 mM MgCl2, 5% (v/v) DMSO, pH 8.0) and injected for 30 sec contact time at 60 µL/min and then allowed to dissociate for 60 sec. Binding sensorgrams were processed using the Scrubber (version 2c, BioLogic Software, Campbell, Australia). To determine the binding affinity (equilibrium dissociation constant; KD), responses at equilibrium for each compound were fit to a 1∶1 steady state affinity model available within Scrubber.
Isothermal Titration Calorimetry (ITC)
Experiments were performed using an iTC200 instrument (MicroCal) at 298 K, with the ligands titrated into solutions of SaHPPK using 18×2.2 µL injections. Data were fitted using Origin software to yield the thermodynamic parameters, ΔH, KD and N (the binding stoichiometry), assuming a cell volume of 0.2 mL. These were then used to calculate the Gibb’s free energy of binding, ΔG (-RT.lnKa), and entropy of binding, ΔS (using ΔG = ΔH - TΔS). A stock solution of SaHPPK was dialyzed overnight into 50 mM HEPES, 1 mM TCEP, 10 mM MgCl2, pH 8.0 buffer with the addition of 5% DMSO (v/v) prior to running the experiment. For titrations with compounds 21a–e, SaHPPK was typically at 30 µM and the ligand stocks were at 1–1.5 mM dissolved in the above buffer then diluted into more of the same buffer. There was no apparent issue with limited solubility of 21a–c compromising either the stock solutions or the injected concentrations.
X-ray Crystallization and Structure Determination
Crystallization experiments were performed as described previously . Briefly, co-crystallization was set-up in the JCSG+ Suite commercial crystal screens (Qiagen) at 281 K using sitting-drop vapor-diffusion method with droplets consisting of 150 nL protein solution and 150 nL reservoir solution and a reservoir volume of 50 µL. Crystals of the SaHPPK in complex with 7-(2-hydroxyethyl)guanine (21a) were observed in conditions containing 240 mM sodium malonate and 20% polyethylene glycol 3350. Data were collected at the MX-2 beamline of the Australian Synchrotron (see Table 3 for statistics) using a one degree oscillation angle, 360 frames were obtained for a complete data set. These data were indexed using XDS  and scaled using SCALA .
The SaHPPK structure (3QBC) was used to solve the initial phases of the binary complex by molecular replacement using Phaser . Refinement was performed using REFMAC5  and the Fourier maps (2FO-FC and FO-FC) were visualized in Coot . After several rounds of manual rebuilding, 21a and water molecules were added and the model further refined to a resolution of 1.85 Å (Rfree (%) = 26.4, Rwork (%) = 20.9).
The coordinates of SaHPPK in complex with 21a have been deposited at the Protein Data Bank with accession number 4ad6.
15N-labelled protein samples for NMR spectroscopy were prepared as described . 2D soFast 15N HMQC  NMR experiments were recorded on a Varian Inova 600 MHz NMR spectrometer equipped with a cryoprobe and Z axis gradient on samples of ∼100 µM 15N-labelled SaHPPK dissolved in 50 mM HEPES buffer (pH 8.0, 90% H2O 10% D2O, 1% sorbitol) by titrating in aliquots from a 25 mM stock of 21a dissolved in DMSO-D6.
SPR raw data ( top ) and steady-state response curves ( bottom ) for the binding of C 8- (10a–f), N 9-(15a–d) and N 7-(21a–e) substituted analogues to Sa HPPK.
SPR raw data ( top ) and steady-state response curves ( bottom ) for the binding of compounds 21a, 21c, 21d and 21e to Sa HPPK.
All SaHPPK crystals were grown at the C3 Crystallisation Centre at CSIRO, Parkville, Australia and X-ray data were obtained at the Australian Synchrotron, Victoria, Australia. All NMR data were acquired at the Monash Institute of Pharmaceutical Sciences. We would like to thank Brett Collins for his helpful suggestions regarding ITC measurements and critical reading of this manuscript.
Conceived and designed the experiments: SC NB OD TSP BG JDS. Performed the experiments: SC NB OD MKH JN TSP BG JDS. Analyzed the data: SC NB OD MKH JN TSP BG JDS. Contributed reagents/materials/analysis tools: OD JN TSP BG JDS. Wrote the paper: SC NB TSP BG JDS.
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