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IMP dehydrogenase (IMPDH) catalyzes two very different chemical transformations, a dehydrogenase reaction and a hydrolysis reaction. The enzyme toggles between the open conformation required for the dehydrogenase reaction and the closed conformation of the hydrolase reaction by moving a mobile flap into the NAD site. Despite these multiple functional constraints, the residues of the flap and NAD site are highly diverged, and the equilibrium between open and closed conformations (Kc) varies widely. In order to understand how differences in the dynamic properties of the flap influence the catalytic cycle, we have delineated the kinetic mechanism of IMPDH from the pathogenic protozoan parasite Cryptosporidium parvum (CpIMPDH), which was obtained from a bacterial source through horizontal gene transfer, and its host counterpart, human IMPDH type 2 (hIMPDH2). Interestingly, the intrinsic binding energy of NAD+ differentially distributes across the dinucleotide binding sites of these two enzymes as well as in the previously characterized IMPDH from Tritrichomonas foetus (TfIMPDH). Both the dehydrogenase and hydrolase reactions display significant differences in the host and parasite enzymes, in keeping with the phylogenetic and structural divergence of their active sites. Despite large differences in Kc, the catalytic power of both the dehydrogenase and hydrolase conformations are similar in CpIMPDH and TfIMPDH. This observation suggests that the closure of the flap simply sets the stage for catalysis rather than plays a more active role in the chemical transformation. This work provides the essential mechanistic framework for drug discovery.
Inosine-5′-monophosphate dehydrogenase (IMPDH)1 catalyzes the penultimate step in guanine nucleotide biosynthesis, the NAD+-dependent conversion of inosine-5′-monophosphate (IMP) to xanthosine-5′-monophosphate (XMP). This reaction controls the size of the guanine nucleotide pool, which in turn controls proliferation (1). IMPDH inhibitors are used as immunosuppressive, anticancer, and antiviral agents (2,3). Recently, IMPDH has emerged as an attractive target for drugs against Cryptosporidium parvum, the intracellular eukaryotic parasite that is a major cause of diarrhea and malnutrition in the developing world and a serious pathogen of AIDS and other immunosuppressed patients in the industrialized world (4–6). The purine salvage pathway of C. parvum relies solely on adenosine; guanine nucleotides are produced by the consecutive actions of adenosine kinase, AMP deaminase, IMPDH, and GMP synthase (7–9). CpIMPDH appears to have been obtained from epsilon proteobacteria via horizontal gene transfer and thus is highly diverged from the human counterparts hIMPDH1 and hIMPDH2 (7,10,11). This situation contrasts with other Apicomplexan parasites such as Plasmodium and Toxoplasma, which have more complex purine salvage pathways and contain eukaryotic IMPDHs similar to the human enzymes (12). Presumably, the bacterial CpIMPDH confers an evolutionary advantage uniquely suited to the metabolic demands of Cryptosporidium.
The IMPDH reaction requires two different chemical transformations, a redox step where Cys319 attacks IMP producing NADH and the covalent intermediate E-XMP* and a hydrolysis step that produces XMP (Figure (Figure1;1; Tritrichomonas foetus IMPDH numbering) (13). When NADH departs, a mobile flap docks in the vacant dinucleotide site, carrying the catalytic base, Arg418, into the active site. Thus the flap toggles the enzyme between dehydrogenase (flap open, dinucleotide site exposed) and hydrolase conformations (flap closed, dinucleotide site occluded). Compounds such as mycophenolic acid inhibit IMPDH by competing with the flap for the vacant NAD+ site, thus trapping E-XMP* (Figure (Figure1).1). Similarly, NAD+ inhibits by forming a nonproductive E-XMP*·NAD+ complex.
Despite such multiple functional constraints, the residues comprising the flap and NAD+ site vary widely among enzymes from different sources, as does the equilibrium between open and closed conformations (14–16). We have proposed that this divergence is a response to naturally occurring IMPDH inhibitors such as mycophenolic acid, which traps E-XMP*, and mizoribine monophosphate, which stabilizes the closed conformation. Indeed, a mutation that causes resistance to mycophenolic acid renders the enzyme hypersensitive to mizoribine, demonstrating how chemical warfare at the microbial level might drive evolution (17). IMPDHs that are resistant to mycophenolic acid have high values of kcat, which indicates that the equilibrium between open and closed conformations controls both inhibition and the catalytic cycle (13). For efficient catalysis, the affinity of the flap and NAD+ must be balanced so that the productive E·IMP·NAD+ complex can form but the nonproductive E-XMP*·NAD+ complex does not accumulate. Together, these observations pose an evolutionary conundrum: for catalytic success, a mutation must simultaneously increase (or simultaneously decrease) the affinity of both the flap and NAD+.
CpIMPDH provides a convenient system to test these ideas: while the active site of the hydrolase conformation of CpIMPDH is very similar to that of the well-characterized TfIMPDH (Figure (Figure2),2), the equilibrium between open and closed conformations varies dramatically (4 for CpIMPDH, 140 for TfIMPDH, and ≤0.2 in hIMPDH2; (16,18)).
Here we delineate the kinetic mechanism of CpIMPDH and hIMPDH2 in order to investigate how differences in the dynamic properties of the flap influence the catalytic cycle and to identify functional differences that might explain why C. parvum discarded its eukaryotic IMPDH in favor of the bacterial version. These experiments demonstrate that, despite large differences in equilibrium between open and closed conformations, the catalytic power of the hydrolase conformation is similar among CpIMPDH and TfIMPDH. The kinetic mechanism derived in this work has already proven invaluable in inhibitor discovery (19).
IMP, XMP, GMP, 3-acetylpyridine adenine dinucleotide (APAD+), and NADH were purchased from Sigma. NAD+ was purchased from Roche. DTT was purchased from Research Organics, Inc. Tris, glycerol, EDTA, trichloroacetic acid, and KCl were purchased from Fisher Scientific. HA nitrocellulose filters were purchased from Millipore. [8-14C]IMP was obtained from Moravek Biochemicals, Inc. D2O and DCl were purchased from Cambridge Isotope Laboratories, Inc. EICARMP was a gift from A. Matsuda (Hokkaido University, Japan).
Free acid forms were used for all substrates, products, and ligands with the following exceptions. The sodium form of NADH was converted into a TrisH+ form by cation exchange using Dowex 50WX4 resin (Sigma). [8-14C]IMP was used as the ditriethylammonium salt.
CpIMPDH was expressed and purified as previously described (16). Purity was >95% as assessed by SDS−PAGE. The concentration of CpIMPDH was determined by Bio-Rad assay using IgG as a standard and by active site titration with the irreversible inactivator EICARMP (20) (data not shown). The Bio-Rad assay overestimates enzyme concentration by a factor of 2.6. This is comparable to the correction factors measured for the IMPDHs from Escherichia coli(20) and T. foetus(18).
Standard IMPDH assays were performed in assay buffer (50 mM Tris-HCl, pH 8.0, 100 mM KCl, and 1 mM DTT) at 25 °C. The production of NADH and reduced 3-acetylpyridine adenine dinucleotide (APADH) was monitored spectrophotometrically at 340 nm (ϵ = 6.22 mM−1 cm−1) and 363 nm (ϵ = 9.1 mM−1 cm−1), respectively. Steady-state parameters for the NAD+ reaction were determined by measuring initial velocities while varying both substrates. Initial velocities were fit by the Michaelis−Menten equation (eq 1) or an equation including uncompetitive inhibition (eq 2) using SigmaPlot (Systat Software, Inc.)
where v is the initial velocity, Vm is the maximal velocity, KM is the Michaelis constant for substrate (S), and Kii is the inhibition constant for S. Steady-state parameters with respect to NAD+ were determined by deriving the apparent Vm values from plots of initial velocity versus IMP concentration (eq 1). These were replotted versus NAD+ concentration, and the steady-state parameters were calculated from eq 2. Similarly, apparent Vm values from initial velocities versus NAD+ concentration (eq 2) were replotted versus IMP concentration, and the Michaelis constant for IMP was derived from eq 1. The apparent steady-state parameters for APAD+ were derived from initial velocity versus dinucleotide concentration at saturating IMP (500 μM) using eq 1.
Ligand was successively added to 0.1−0.15 μM CpIMPDH in assay buffer at 25 °C. The equilibrium protein fluorescence was measured with a Hitachi F-2000 fluorescence spectrophotometer. In order to minimize inner filter effects, excitation wavelengths of 290 nm for IMP and NADH and 295 nm for NAD+, XMP, and GMP were used. Emission was monitored at 344 nm for IMP, NAD+, XMP, and GMP and at 342 nm for NADH. Inner filter corrections were calculated by the formula
where Fc is the corrected fluorescence, Fo is the observed fluorescence, and Aex and Aem are the absorbencies at the excitation and emission wavelengths, respectively. Fc/Fo was less than 2 for all ligand concentrations used. Since purines are known fluorescent quenchers (21), nonspecific quenching was measured by titrating ligand against a solution of l-Trp and was described by the Stern−Volmer relation
where F0 is the unquenched fluorescence, F is the fluorescence at a given quencher (Q) concentration, and KQ is the quenching constant. The experimentally determined values of KQ with l-Trp were used to calculate correction factors (F0/F) that were applied to the binding data. The values of KQ−1 for IMP, XMP, GMP, NAD+, and NADH are 5.5, 0.53, 1.1, 4.3, and 0.24 mM, respectively. The corrected fluorescence due to binding (Fbind) was calculated using eq 5.
The program DynaFit (22) was used to describe Fbind versus ligand concentration with a simple one-step binding mechanism.
Assay buffer was prepared in either H2O or D2O. The pH meter readings for the D2O solutions were adjusted by adding 0.4. Initial velocities were measured for varying dinucleotide concentrations in assay buffer with 500 and 400 μM IMP for CpIMPDH and hIMPDH2, respectively. Steady-state parameters for the CpIMPDH reaction were derived from eq 2 and eq 1 for NAD+ and APAD+, respectively, and eq 2 was used for hIMPDH2.
The kinetics of XMP binding were measured with an Applied Photophysics SX.17MV stopped-flow spectrophotometer by monitoring the changes in protein fluorescence (excitation = 295 nm, 320 nm cutoff filter). Equal volumes of enzyme and XMP were combined at time 0 in assay buffer at 25 °C giving a final concentration of 0.4 μM enzyme and 30 or 120 μM XMP.
Pre-steady-state experiments were performed monitoring either NADH absorbance (340 nm) or fluorescence (340 nm excitation, 420 nm cutoff filter). Enzyme and saturating IMP were preincubated and combined with an equal volume of NAD+ at time 0. For CpIMPDH, NAD+ was varied against a final concentration of 500 μM IMP and 2.7 and 1 μM enzyme for the absorbance and fluorescence experiments, respectively. Fluorescence experiments with hIMPDH2 used final concentrations of 20−200 μM NAD+ combined with 4.0 μM enzyme and 2 mM IMP. Absorbance experiments also used 2 mM IMP but with 1, 1.75, or 3.5 μM enzyme with 400 μM NAD+ or 3.5 μM enzyme with 1 mM NAD+.
St is the signal at time t, A1 and A2 are the amplitudes of the first and second phases respectively, kobs1 and kobs2 are the observed first-order rate constants for the first and second phases respectively, v is the steady-state rate, and b is a background term. The hyperbolic dependence of kobs on NAD+ concentration was described by
where kmax is the maximal value of kobs and Kapp is the concentration of [NAD+] at 1/2 kmax.
Accumulation of the covalent E-XMP* complex was measured from reactions with 100 μM [8-14C]IMP, 500 μM NAD+, and 1 μM CpIMPDH in assay buffer at 25 °C. Reactions were quenched during the steady state prior to 10% completion by precipitation with cold TCA to a final concentration of 10%. Enzyme was captured on 0.45 μm HA nitrocellulose filters (prewet with 10% TCA) and washed with cold 10% TCA. Radioactivity was measured by scintillation counting. Control reactions lacked either enzyme or NAD+.
We determined the steady-state parameters for CpIMPDH using active site titrated enzyme and varying both substrate concentrations (Table (Table1).1). These values are in good agreement with our previous report (16). Importantly, the values of kcat for NAD+ and APAD+ are comparable, suggesting that these two reactions share a common rate-limiting step, most likely the hydrolysis of E-XMP*. Solvent isotope effects on kcat of 2.6 and 3.3 were observed for the NAD+ and APAD+ reactions, respectively (Table (Table1),1), which further indicates that the hydrolysis of E-XMP* is rate-limiting.
We measured the binding of substrates and products to CpIMPDH by monitoring changes in intrinsic protein fluorescence. CpIMPDH contains two Trp residues (Trp90 and Trp368). The corresponding residues in TfIMPDH (Phe99 and Gln470, respectively) are located ~25 Å from the active site. Nevertheless, all ligands produce decreases in CpIMPDH fluorescence. Corrections for inner filter effects and nonspecific quenching were made as described in Materials and Methods. In all cases, the data are well described by a simple binding mechanism. The values of Kd for IMP, XMP, and GMP are similar (~5 μM, Table Table2,2, Figure S1; the notation “S” denotes figures, etc., in the Supporting Information). In contrast, NADH binds with greater affinity than NAD+ by a factor of 6 (Table (Table2).2). The values of Kd for both IMP and NAD+ are significantly lower than their respective KM values. This observation suggests that both substrates are “sticky”, i.e., react to form products faster than they are released from the enzyme.
We also investigated the kinetics of XMP association and dissociation. Despite the fact that XMP quenches the intrinsic fluorescence of CpIMPDH, we were unable to observe a signal in the stopped-flow spectroscopy experiments. We conclude that the binding of XMP is too fast to measure (≥200 s−1).
A pre-steady-state burst of NADH production is observed when the CpIMPDH reaction is monitored by absorbance (Figure (Figure3A).3A). The progress curve is well described by a single exponential phase followed by a linear steady state (Figure S2A). The value of kobs for the burst phase displays a hyperbolic dependence on NAD+ concentration (Figure S2B, Table Table3).3). Since absorbance monitors both free and enzyme-bound NADH (23), the maximum value of kobs (kburst = 41 ± 2 s−1) is a composite rate constant including both hydride transfer and NADH release. Therefore, 41 s−1 serves as a lower limit for the rate of hydride transfer.
Fortuitously, purines are strong quenchers so that the fluorescence of NADH is quenched in the E-XMP*·NADH complex and is only observed when NADH is released from the enzyme (23). A burst of NADH production is also observed when the CpIMPDH reaction is monitored by fluorescence, and again the progress curve is well described by a single exponential burst phase followed by a linear steady state (Figures (Figures3B3B and S3A). The value of kobs is hyperbolically dependent on NAD+ concentration with a maximum value of 16 s−1 (Figure S3B, Table Table33).
The above experiments indicate that hydride transfer is fast and hydrolysis is rate-limiting in the CpIMPDH reaction, which predicts that E-XMP* will accumulate in the steady state. Therefore, we measured the incorporation of radioactivity into the protein during the reaction with [8-14C]IMP and NAD+ (500 μM). This NAD+ concentration was chosen to avoid formation of the nonproductive E-XMP*·NAD+ complex. No radioactivity is incorporated into protein if NAD+ is omitted from the reaction. Under these conditions, 25 ± 3% of the enzyme is trapped as E-XMP*, which is comparable to the burst amplitude of NADH produced at similar NAD+ concentrations (30 and 21% at 600 and 300 μM NAD+, respectively).
Values for the individual rate constants in the kinetic mechanism of CpIMPDH were derived by fitting the absorbance and fluorescence stopped-flow progress curves using the program DynaFit (22) as described in the Supporting Information. The individual rate constants are shown in Scheme 1. Signal response factors for the fluorescence data were calculated for each concentration of NAD+ to account for inner filter effects by normalizing the linear steady-state regions with the absorbance data. The rate constants for the association of NAD+ to E·IMP (Scheme S1, k5, k6), hydride transfer (k7, k8), and NADH release (k9) are well constrained. As noted above, the release of XMP is fast (≥200 s−1); the individual rate constants (kclose, kopen) describing the movement of the flap were chosen to be arbitrarily fast such that the value of Kc = 4 (16). The individual rate constants for NAD+ binding to E-XMP* (k11, k12) are not well determined, but the ratio k12/k11 is consistently 200−400 μM. Taking into account the competition with the flap, this value is in reasonable agreement with experiment (400 μM × 5 = 2 mM; Kii = 4.9 mM, Table Table1).1). The results of the global fits are summarized in Figure Figure3,3, Table S1, and Scheme 1. Using these values, the calculated value of kcat is 1.8 s−1 (eq 11, Supporting Information), in agreement with the measured value of 2.6 s−1. The calculated value of KMNAD+ is 40 μM (eq 12, Supporting Information), similar to the measured value of 140 μM.
Although hydrolysis is rate-limiting in the reaction of hIMPDH2 (24,25), the pre-steady-state reaction had not been characterized in detail. When the reaction was monitored by absorbance, the progress curves display a burst of NADH production and are well described by a single exponential with a linear steady state (Figures (Figures4A4A and S4A). The amplitude of the burst phase is 0.9 ± 0.1 [NADH]/[E]. The value of kobs = 14 ± 1 s−1 at saturating substrate concentrations (Table S2), which is 35 times greater than the value of kcat (0.4 s−1).
When the reaction was monitored by fluorescence, the progress curve exhibits three phases, a lag followed by a burst and a linear steady state, which are well described by eq 7 (Figures (Figures44 and S4A). Replots of the values of kobs1 and kobs2 versus [NAD+] are both hyperbolic, with maximum values of 23 ± 1 s−1 and 12 ± 1 s−1 for the lag and burst phases, respectively (Table (Table44).
These observations suggest that the hydrolysis of E-XMP* is also rate-limiting in the reaction of hIMPDH2, which predicts that a solvent deuterium isotope effect will be observed on kcat; we do observe a solvent deuterium isotope effect of 1.8 ± 0.1, in contrast to a previous report from another laboratory (25). This discrepancy may result from the different reaction conditions (100 mM Tris-HCl, pH 7.7, and 10 mM KCl at 37 °C whereas our experiments use 50 mM Tris-HCl, pH 8.0, 100 mM KCl, and 1 mM DTT at 25 °C).
Global Fits of the hIMPDH2 Reaction. Values for the individual rate constants in the kinetic mechanism of hIMPDH2 were derived as described above and in the Supporting Information. The individual rate constants derived from the global and local fit analyses are shown in Scheme 1 and Table S1. The rate constants for the association of NAD+ to E·IMP (Scheme S2, k5, k6), hydride transfer (k7, k8), and NADH release (k9) are well constrained. As with CpIMPDH, the individual rate constants for NAD+ binding to E-XMP* (k11, k12) are not well determined; the values of k11 and k12 were set arbitrarily fast such that k12/k11 = Kii = 590 μM (24). Importantly, the magnitude of the solvent isotope effect indicates that hydrolysis is not solely rate-limiting in the hIMPDH2 reaction. Assuming the intrinsic isotope effect is 3, as observed in the reaction of CpIMPDH (as well as in the reaction of serine proteases (26)), then kHOH = 1 s−1 and k = 0.7 s−1 for the second rate-limiting step (27). Both hydride transfer and the release of NADH are much faster than 0.7 s−1. Unfortunately, hIMPDH2 does not contain any Trp residues, so the kinetics of ligand binding cannot be directly measured; nevertheless, given the low affinity of XMP (Kd = 140 μM (28)), the release of XMP is unlikely to be rate-limiting. Therefore, we propose that the second rate-limiting step is a conformational change. Since the open flap conformation is favored in hIMPDH2 (Kc ≤ 0.2) (14), we have provisionally assigned 0.7 s−1 to kclose.
We have now characterized the kinetic mechanisms of TfIMPDH, CpIMPDH, and hIMPDH2; Scheme 1 displays a “kinetic alignment” of these enzymes. The interchange between open and closed conformations of the flap is the most divergent step in these catalytic cycles (Kc in Scheme 1). Since the flap competes with NAD+ for the dinucleotide site, the affinity of NAD+ must be balanced against that of the flap; otherwise nonproductive complexes such as E-XMP*·NAD+ will accumulate. When the fraction of enzyme in the open conformation is taken into account, the “intrinsic binding affinity” of compounds that bind in the NAD site can be considerably greater than the observed binding affinity (Table (Table5).5). TfIMPDH dramatically illustrates this principle: although the observed affinity of NAD+ is the lowest of the three enzymes, the intrinsic affinity is the highest. Similar results are observed for the intrinsic affinity of β-methylene-tiazofurin adenine dinucleotide (β-Me-TAD, Figure Figure5).5). Curiously, the intrinsic affinities of NAD analogues for CpIMPDH and hIMPDH2 are very similar despite the divergent structures of these dinucleotide sites. Interestingly, the intrinsic affinity of β-Me-TAD differentially distributes across the dinucleotide sites of these three enzymes. The intrinsic affinity of tiazofurin is similar in TfIMPDH and CpIMPDH, as expected given that the structures of the nicotinamide subsites are essentially identical. In contrast, the nicotinamide subsite of hIMPDH2 contains several substitutions, and the intrinsic affinity of tiazofurin is significantly lower. The opposite situation is observed at the adenosine subsite: the intrinsic affinity of ADP is much higher for TfIMPDH than CpIMPDH. This difference is likely to arise from interactions with the adenine ring of ADP, which is sandwiched between Arg241 and Trp269 in TfIMPDH. These residues are Asn144 and Asn171 in CpIMPDH. In hIMPDH2, the adenine is sandwiched between His253 and Phe282, and ADP displays an intrinsic affinity similar to CpIMPDH.
Remarkably, despite large differences in the dynamics of the flap, the values of kHOH are identical for CpIMPDH and TfIMPDH. This observation suggests that the dynamics of the flap set the stage for the hydrolysis reaction but do not play an active role in the chemical transformation. The rate constants for hydride transfer are also similar for CpIMPDH and TfIMPDH. The residues in the immediate vicinity of the hypoxanthine and nicotinamide groups are virtually identical for these two enzymes, so this congruence is also not surprising. In fact, with the exception of the equilibrium between open and closed conformations and the binding of NAD+, the kinetic mechanisms of CpIMPDH and TfIMPDH are essentially identical. In contrast, both hydride transfer and hydrolysis are slower in hIMPDH2 than in either of the parasite enzymes. The equilibrium between the E·IMP·NAD+ and E-XMP*·NADH complexes also has changed, which indicates that the transition state for this reaction has also changed. Several substitutions can be found at the active site that may account for this behavior. The most intriguing of these is the substitution of Glu431 with Gln; we have proposed that Glu431 is part of a proton relay that activates water and might also function to activate Cys319 (L. Hedstrom and W. Yang, unpublished). The substitution of Glu431 with Gln does decrease the value of kcat as expected (29).
We are aware of four other enzyme systems where the detailed kinetic mechanisms have been determined for orthologues: triose-phosphate isomerase, dihydrofolate reductase, purine nucleoside phosphorylase, and adenosine deaminase. Product release is rate-limiting in the cases of dihydrofolate reductase and triose-phosphate isomerase, and the values of kcat are similar in orthologues (30–33). In contrast, the values of kcat vary by as much as 2 orders of magnitude, and the chemical transformations are rate-limiting in purine nucleoside phosphorylase and adenosine deaminase (34,35). Here the structures of the transition state change, as we envision the transition structure must change in hIMPDH2 versus CpIMPDH and TfIMPDH. Interestingly, substitutions quite remote from the active site contribute to these changes (36).
Why does C. parvum utilize a bacterial IMPDH? Interestingly, many eukaryotic pathogens, including Trypanomastids, Giardia lamblia, Entamoeba histolytica, and Trichomonas vaginalis, have remodeled and simplified their metabolic pathways while adopting bacterial genes, so this phenomenon is widespread (37–40). One simple hypothesis is that the higher values of kcat associated with bacterial IMPDHs promote greater flux through the guanine nucleotide biosynthetic pathways. Our experiments suggest that this is not the case: though the rate constants for individual steps in the reactions of CpIMPDH and hIMPDH2 are often significantly different, the flux through both enzymes will be comparable in vivo, assuming that the physiological concentrations of NAD+ and IMP in the parasite are similar to those of most organisms ([IMP] = 20−60 μM (41); [NAD+] = 0.3−2 mM (42)). Therefore, it seems unlikely that C. parvum chose the bacterial enzyme to increase the production of guanine nucleotides. Instead, a moonlighting function may have provided the impetus for the switch. Virtually all IMPDHs contain a subdomain. The subdomain is not required for enzymatic activity (43–45), and its function is currently under debate. CpIMPDH lacks the subdomain and, therefore, may be missing the moonlighting functions of the host enzyme. Intriguingly, the subdomain has recently been shown to regulate the distribution of purine nucleotides between the adenine and guanine nucleotide pools in E. coli(46), and it seems reasonable to expect this regulation would be different in C. parvum given its unusual purine salvage pathway. We have found that the subdomain mediates the association of IMPDH with single-stranded nucleic acids (47) and may well have an as yet unappreciated role in RNA metabolism that would also be different in C. parvum.
The kinetic mechanism derived herein has already proved invaluable in designing a high throughput screen to identify selective inhibitors of CpIMPDH (19). We used Scheme 1 to define assay conditions such that the dinucleotide site was empty in the predominant enzyme forms, thus selecting for inhibitors that bind to the most diverged site on the enzyme. These results demonstrate how a detailed understanding of enzyme kinetics can be important in drug discovery.
Equilibrium fluorescence binding data as well as local and global fit analyses of the absorbance and fluorescence stopped-flow data for CpIMPDH and hIMPDH2. This material is available free of charge via the Internet at http://pubs.acs.org.