Differential susceptibility of Plasmodium falciparum versus yeast and mammalian enolases to dissociation into active monomers Ipsita Pal-Bhowmick, Sadagopan Krishnan and Gotam K Jarori Department of Biological Sciences, Tata Institute of Fundamental Research, Colaba, Mumbai, India Keywords enolase; monomers; Plasmodium falciparum; rabbit muscle; yeast Correspondence G K Jarori, Department of Biological Sciences, TIFR, Homi Bhabha Road, Colaba, Mumbai 400 005, India Fax: +91 22 22804610 Tel: +91 22 22782000 E-mail: gkj@tifr.res.in (Received December 2006, revised 16 January 2007, accepted 12 February 2007) doi:10.1111/j.1742-4658.2007.05738.x In the past, several unsuccessful attempts have been made to dissociate homodimeric enolases into their active monomeric forms The main objective of these studies had been to understand whether intersubunit interactions are essential for the catalytic and structural stability of enolases Further motivation to investigate the properties of monomeric enolase has arisen from several recent reports on the involvement of enolase in diverse nonglycolytic (moonlighting) functions, where it may occur in monomeric form Here, we report successful dissociation of dimeric enolases from Plasmodium falciparum, yeast and rabbit muscle into active and isolatable monomers Dimeric enolases could be dissociated into monomers by high concentrations ( 250 mm) of imidazole and ⁄ or hydrogen ions Two forms were separated using Superdex-75 gel filtration chromatography A detailed comparison of the kinetic and structural properties of monomeric and dimeric forms of recombinant P falciparum enolase showed differences in specific activity, salt-induced inhibition and inactivation, thermal stability, etc Furthermore, we found that enolases from the three species differ in their dimer dissociation profiles Specifically, on challenge with imidazole, Mg(II) protected the enolases of yeast and rabbit muscle but not of P falciparum from dissociation The observed differential stability of the P falciparum enolase dimer interface with respect to mammalian enolases could be exploited to selectively dissociate the dimeric parasite enzyme into its catalytically inefficient, thermally unstable monomeric form Thus enolase could be a novel therapeutic target for malaria Enolase (EC 4.2.1.11) is a glycolytic enzyme that catalyzes the interconversion between 2-phospho-d-glycerate (PGA) and phosphoenolpyruvate Enolases from most organisms exist as homodimers of subunit mass 40–50 kDa [1], the exceptions being homo-octameric enolases from thermophilic bacteria [2–4] A variety of physical data indicate that each dimer contains two active sites The active site in each subunit is completely independent [5,6] As the active site is fully contained in each subunit, attempts to dissociate the dimeric form into an active monomeric form have been made using genetic [7], chemical and physical methods, but without much success Although dissociation of the dimer could be achieved, the monomers formed were found to be inactive [8–11] The formation of active monomers was inferred under conditions of high temperature (40–45 °C) and low protein concentration (in the nanomolar range) [12,13] However, at such low concentrations, neither the kinetic nor the structural characterization of the monomer could be carried Abbreviations Mim, imidazole-generated monomer; Mnt, native monomer; MpH, pH-generated monomer; 2-PGA, 2-phospho-D-glycerate; RMen, rabbit muscle enolase; r-Pfen, recombinant Plasmodium falciparum enolase; Yen, yeast enolase 1932 FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS I Pal-Bhowmick et al out Hydrostatic pressure, which is viewed as a gentle and reversible perturbation, has also been employed in attempts to obtain active monomers from dimeric enolase [8,11,14–17] Although the dissociation of enolase dimers into active monomers was inferred in some of the above studies, it has never been unequivocally demonstrated As none of these attempts had resulted in the isolation of active monomers and their characterization, it was not clear whether the monomeric form is intrinsically inactive or the means of dissociation selectively inactivated it Suggestions have also been made that intersubunit interactions may be essential for completion of the catalytic cycle In recent years, enolase has also been shown to participate in a variety of nonglycolytic (moonlighting) biological functions [18] The oligomeric state of enolase recruited for the moonlighting functions is not known Our interest in the enolase from the malarial parasite (Plasmodium falciparum) originates from the fact that the intraerythrocytic stages of the parasite lack active mitochondria and hence rely solely on glycolysis for their energy needs [19,20] The level of glycolytic flux in parasite-infected cells is  50–100-fold greater than that in uninfected red blood cells, and the activity of many glycolytic enzymes is upregulated, enolase being one of them [21–23] P falciparum enolase (Pfen) could be a potential drug target, as there is only one gene for this enzyme, and it shows greater resemblance to plant enolases than to mammalian enolases In order to examine such possibilities and explore whether it has any moonlighting functions, we have recently cloned the P falciparum enolase gene, overexpressed it, and obtained pure protein [24] We have also raised polyclonal and monoclonal antibodies against recombinant r-Pfen for subcellular localization studies [25] Our observations have shown a diverse subcellular localization (enolase is associated with plasma membrane, cytosol, cytoskeletal elements and nucleus; unpublished results) for enolase, indicating that it may be recruited for certain other nonglycolytic functions One of the conventional approaches in rational drug design has been to make active site-specific inhibitors that can differentiate between host and parasite proteins and bring about selective inhibition of the parasite enzyme The major limitation of this strategy is that active sites are evolutionarily highly conserved, and structural differences may be rather subtle or even nonexistent As there are numerous protein–protein interactions that operate in living systems [26–28], and most proteins exist as oligomers [29] with a predominance of homodimers [30], another approach could be to target protein–protein interfaces for perturbing Dissociation of enolase into active monomers protein and cell functions [31] In cases where oligomeric structure is essential for biological activity, selective disruption of such an interface in a protein of parasite origin can have therapeutic effects Rationally designed peptides that can compete for the interaction between monomeric subunits (peptidomimetics) have yielded encouraging results [32–35] Attempts have also been made to find nonpeptide small molecule inhibitors that effectively interfere with protein–protein interactions [36–38] Here, we have examined the possibility of dissociating malarial parasite enolase into monomers using small molecules, and characterized the properties of the monomeric state We report the successful dissociation of r-Pfen, and yeast and rabbit muscle enolases, into isolatable active monomers Thus we demonstrate that the dimeric structure is not essential for catalysis However, comparative kinetic and structural studies on the monomeric and dimeric forms of r-Pfen showed several interesting differences The monomeric form has low specific activity, is more thermolabile, and is more prone to lose activity in the presence of salts as compared to the dimer Our experiments have also identified conditions under which selective dissociation of the parasite enolase may be accomplished Although the concentrations of dissociating ligand used in this study are rather high and unrealistic for therapeutic applications, the possibility remains that such differences may be exploitable for targeting this enzyme for therapeutic purposes Results Imidazole-induced dissociation of r-Pfen into active monomers Like most enolases, r-Pfen is a homodimer of two 50 kDa subunits [24] Purified dimeric r-Pfen was dialyzed against increasing concentrations (0–250 mm) of imidazole and then subjected to gel filtration chromatography on a Superdex-75 column Figure 1A shows gel filtration chromatograms obtained at several different concentrations of imidazole at pH 6.0 A dimer peak was observed in the absence of imidazole However, with increasing concentrations of imidazole, the monomeric fraction increased, and at  250 mm imidazole, > 95% of r-Pfen was dissociated The effect of pH on imidazole-induced dissociation was examined by dialyzing the enzyme against 50 mm sodium phosphate at different pH values (6.0, 7.0 and 8.0) with increasing concentrations of imidazole The percentage of monomer present in each of these samples was determined from gel filtration chromatograms The results presented in Fig 1B show that lower pH FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS 1933 Dissociation of enolase into active monomers A I Pal-Bhowmick et al r-Pfen 100 [Imidazole] 250 mM B pH 6.0 pH 7.0 75 50 pH 8.0 25 80 50 mM dimer 50 mM 40 50 60 Ve (ml) 70 100 150 200 [Imidazole](mM) 250 C 40 0 80 50 55 60 65 70 75 ( - ) 50 Specific activity ( U/ mg ) 150 mM A 280 (mAU) A 280 (mAU) 50 % monomer monomer 75 80 Ve (ml) Fig Imidazole-induced dissociation of r-Pfen (A) Superdex-75 gel filtration chromatograms obtained at different concentrations of imidazole Enzyme (0.5 mg) was dialyzed ( 14 h) against 50 mM sodium phosphate (pH 6.0) containing different amounts (0–250 mM) of imidazole The column was pre-equilibrated with appropriate buffer, and chromatography was performed at room temperature (20 ± °C) (B) Effect of pH on imidazole-induced dissociation of r-Pfen Each data point is an average of two chromatographic runs (C) Gel filtration chromatogram of r-Pfen in 50 mM sodium phosphate containing 250 mM imidazole (pH 6.0) A280 nm and specific activity (r—r) for each fraction are shown 1934 Yen A A 280 (mAU) dimer (a) dimer 10 (b) monomer (c) 40 B 50 60 Ve(ml) 70 80 RMen dimer A 280 (mAU) favored the dissociation of r-Pfen Figure 1C shows a gel filtration chromatogram of a sample dialyzed against 250 mm imidazole in 50 mm sodium phosphate (pH 6.0) The enzyme activity of each fraction was measured, and the specific activity was plotted along with protein concentration (A280) as a function of elution volume The results showed that both forms (monomeric and dimeric) were catalytically active, with the dimer having  3-fold greater specific activity than the monomer Experiments were also performed in which the effects of adding NaCl to the dissociation buffer were examined The results showed that the presence or absence of salt (300 mm NaCl) did not have any effect on monomer–dimer equilibrium Imidazole-generated monomers of r-Pfen (Mim) ( 10 lm), when extensively dialyzed against imidazole-free buffer, could reassociate to form active dimers To the best of our knowledge, this is the first demonstration of obtaining active monomers of enolase that could be separated from dimers The ability of imidazole to dissociate enolases from other species was also examined Imidazole at 250 mm and pH 6.0 failed to dissociate yeast enolase (Yen) However, inclusion of 300 mm NaCl along with 250 mm imidazole at pH 6.0 resulted in almost complete dissociation of Yen (Fig 2A) In the case of rabbit muscle enolase (RMen), 250 mm imidazole did not dissociate the enzyme As commercial preparations of RMen contain Mg(II), we included mm EDTA along with imidazole This resulted in partial dissociation of (a) 35 15 dimer (b) dimer monomer (c) 40 50 60 Ve(ml) 70 80 Fig Imidazole-induced dissociation of Yen and RMen Samples were in 50 mM sodium phosphate (pH 6.0), and 0.2–0.6 mg of protein was used for each chromatographic run on a Superdex-75 column (A) Yen: (a) no imidazole; (b) 250 mM imidazole; and (c) 250 mM imidazole + 300 mM NaCl (B) RMen: (a) no imidazole; (b) 250 mM imidazole; and (c) 250 mM imidazole + mM EDTA FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS I Pal-Bhowmick et al Dissociation of enolase into active monomers RMen (Fig 2B) However, further inclusion of 300 mm NaCl did not lead to complete dissociation of RMen In this respect, RMen differs from Yen As Mg(II) is an essential cofactor for stabilization of the active conformation and catalysis, extensive kinetic and direct metal ion-binding studies have been performed in the past These studies have shown that each enolase subunit has three binding sites for the divalent cation, namely a conformational site (site I), a catalytic site (site II) and an inhibitory site (site III) [39–42] As binding of divalent cation induces large conformational changes in enolase, we examined the effect of Mg(II) on imidazole-induced dissociation of r-Pfen, Yen and RMen in the presence of different compounds The results are summarized in Table Inclusion of EDTA [to chelate residual Mg(II) in the protein sample] and NaCl in the dissociation buffer did not have any effect on the dimeric state However, the presence of Mg(II) affected the imidazole-induced (or imidazole + NaCl-induced) dissociation of RMen and Yen It is interesting to note that in the presence of 1.5 mm MgCl2, imidazole could dissociate r-Pfen but had no effect on Yen and RMen Table Effect of Mg(II) on monomer–dimer equilibrium in enolase Enolase, 10 lM (0.5 mgỈmL)1), was dialyzed against 50 mM sodium phosphate (pH 6.0) containing different compounds as indicated below Concentrations of these compounds when used were: [imidazole] ¼ 250 mM; [EDTA] ¼ mM; [NaCl] ¼ 300 mM; [MgCl2] ¼ 1.5 mM The Superdex-75 column was pre-equilibrated with the same buffer as used for dialysis Dimeric–monomeric states of enolasesa Enolase P falciparum (r-Pfen) Yeast (Yen) Rabbit muscle (RMen) + + + + + Dimer Dimer Dimer Monomer Monomer Dimer Dimer Dimer Dimer Dimer + Imidazole + NaCl Monomer Monomer + MgCl2 + imidazole + NaCl + MgCl2 + imidazole + EDTA + MgCl2 + imidazole + EDTA + NaCl Monomer Dimer Dimer Dimer Dimer Dimer Partial monomer Partial monomer Dimer Monomer Dimer Monomer Monomer a MgCl2 EDTA EDTA + NaCl Imidazole Imidazole + EDTA Partial monomer Partial monomer When the amount of dimer or monomer is ‡ 90%, it is stated as ‘dimer’ or ‘monomer’ When the amount is ‡ 40%, it is stated as ‘partial monomer’ pH-induced dissociation of enolases The effect of pH on the oligomeric state of enolases was examined by performing gel filtration chromatography on the enzyme, dialyzed against 50 mm sodium phosphate of the desired pH Figure 3A presents the chromatographic profiles of r-Pfen in the pH range 4.5–8.0 Figure 3B shows that 50% dissociation of r-Pfen occurs around pH 5.5 Monomeric r-Pfen generated by low pH (MpH) was also found to be enzymatically active (see below) Low pH could also dissociate Yen and RMen The half-dissociation point for Yen was around pH (Fig 3C,D), whereas for RMen it was around pH 5.5 (Fig 3E,F) Activity and reassociation of MpH and Mim Measurements of enzyme activity in monomeric and dimeric fractions showed that Mim had  3-fold less specific activity than dimers (Fig 1C) As the assay solution did not contain imidazole, it is possible that the observed activity of Mim may have arisen from reassociation to dimers Similarly, MpH could also reassociate at the assay solution pH of 7.4 Such reassociation of monomers into dimers during an enzyme assay would result in an increase in the slope of the reaction progress curve (as the dimer is  3-fold more active than monomers) From these considerations, we can state that during an enzyme assay using MpH or Mim: (a) if there is reassociation of monomers into dimer on the timescale of the enzyme assay, the reaction progress curve will exhibit a time-dependent increase in slope; and (b) as the amount of dimer formed increases as the square of the monomer concentration, a plot of monomer concentration versus activity would have an upward curvature if there was a significant amount of dimer formed during the enzyme assay Figure 4A shows the reaction progress curves at three different concentrations of MpH (0.04, 0.17 and 0.43 lm) The observed increase in slope with time suggests reassociation of MpH into dimers under our assay conditions A similar experiment performed using Mim (Fig 4B) did not show any change in slope with time We measured the concentration dependence of specific activity for MpH at time  0.0 as well as at time > (Fig 4C) The first slope reflects the monomer activity, whereas the slope determined after > reflects the dimer activity With the increasing concentration of MpH, initially we observed a very low specific activity ([MpH] > 0.5 lm) However, at higher protein concentrations, the rate of formation of dimer increased rapidly, and only a single slope, FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS 1935 Dissociation of enolase into active monomers I Pal-Bhowmick et al A 280 (mAU) A 280 (mAU) A 280 (mAU) A B D F Fig pH-induced dissociation of enolases from different organisms Each enzyme (0.2–0.6 mg) was dialyzed against 50 mM sodium phosphate (pH 4.5–8.0) containing 150 mM NaCl and subjected to gel filtration chromatography (A) Gel filtration profile of r-Pfen at different pH values (B) pH versus % monomer (data are from two different experiments) for r-Pfen The r-Pfen used here was prepared by eluting the protein from Ni–nitrilotriacetic acid resin at low pH (C) Gel filtration chromatograms for Yen (D) pH versus% monomer for Yen (E) Gel filtration chromatograms for RMen (F) pH versus % monomer for RMen characteristic of dimer-specific activity, could be observed (Fig 4C) As in the low-concentration range of MpH the observed activity is rather small, it is likely that either MpH is inactive and the measured activity is a reflection of the formation of tiny amounts of active dimer, or the MpH has intrinsically very low activity Irrespective of these two possibilities, the observed second slope reflects the activity for the dimer formed during the assay As expected, the specific activity computed using the second slope did not exhibit the monomer concentration-dependent variation (Fig 4C) These results indicate that MpH rapidly reassociates to form dimer However, similar measurements of concentration dependence of activity for Mim gave a linear increase in activity, suggesting that r-Pfen Mim did not associate rapidly (with respect to assay timescale) to form more active dimers (Fig 4D) A replot of these 1936 data as specific activity versus Mim concentration (Fig 4D) showed that the observed specific activity was low ( 1.8–1.9 unitsỈmg)1, a characteristic of Mim) and was concentration invariant on enzyme assay timescales These observations support the view that Mim associates slowly to form dimers, whereas MpH associates rapidly Thus MpH and Mim differ in their ability to reassociate, indicating that they represent two different conformational states of the monomeric form of the enolase A schematic representation of the dissociation and association of r-Pfen is presented in Scheme 1, where Mnt is the native monomer, which is assumed to be the only monomeric form that can dimerize As Mim mostly remained in the monomeric state under our assay conditions, kinetic characterization of the monomeric forms of enolase was performed using Mim only FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS I Pal-Bhowmick et al A Dissociation of enolase into active monomers B pH monomer (MpH ): reaction progress curves 1.354 1.464 (µM) 1.34 Imidazole monomer (M im ): reaction progress curves (µM) 0.04 1.45 1.44 0.13 1.32 A 240 nm 0.17 1.30 A 240 nm 1.43 1.42 1.41 1.28 0.33 1.40 1.39 0.43 1.255 0.6 1.379 0.00 0.2 0.4 0.6 0.8 1.0 1.2 0.00 1.4 1.50 0.2 0.4 0.6 D pH monomer (MpH ): concentration dependence of activity 1.2 1.4 1.50 Imidazole monomer (Mim ): concentration dependence of activity 0.40 ) 2.5 1.8 1.5 1.0 0.5 1.6 0.30 1.4 0.25 1.2 0.20 1.0 0.15 0.8 Activity (units) (o 2.0 o) 0.35 Specific Activity (units·mg–1) ( Specific Activity (units·mg–1) 1.0 Time (minutes) Time (minutes) C 0.8 0.10 0.6 0.05 0.0 0.0 0.1 0.2 0.3 0.4 0.2 0.5 [Monomer] (µM) 0.3 0.4 0.5 0.6 0.7 [Monomer] (µM) Fig Reaction progress curves for the monomeric r-Pfen-catalyzed reaction Conversion of phosphoenolpyruvate to 2-PGA was monitored at 240 nm (A) MpH (B) Mim Note the time-dependent change in progress curve slopes in (A) (C) Variation in specific activity as a function of [MpH], calculated from first (time  0.0 min) (d) and second (time > min) (j) slopes of reaction progress curves Note the change in specific activity computed from the time  0.0 slope, reflecting the rapid formation of dimer at higher concentrations of MpH (D) Variation in activity (and specific activity) as a function of [Mim] Activity showed a linear increase (o), with specific activity remaining constant (d), suggesting that Mim did not associate to form high-activity dimer on the enzyme assay timescale MpH from Yen and RMen also showed nonlinear reaction progress curves, very similar to those observed for r-Pfen (data not shown) In enzyme assays, Mim from Yen gave nonlinear reaction progress curves displaying an increase in slope with time This is likely to be due to Mg(II)-induced rapid dimerization of yeast Mim As Mg(II) does not affect the dimerization of rPfen monomers, such a change of slope was not observed for Mim prepared from r-Pfen (Fig 4B) Observed low activities of monomeric (Mim and MpH) enolases from all three species would imply that quaternary interactions between two subunits stabilize catalytically more active conformations of each subunit Comparison of kinetic properties of monomeric (Mim) and dimeric forms of r-Pfen As Mim did not reassociate rapidly into dimers, this form of the enzyme was used for comparative kinetic studies Figure 5A shows the variation of enzyme activity as a function of phosphoenolpyruvate concentration Data were fitted to the Michaelis–Menten equation using sigmaplot software The nonlinear t of data gave Vmax ẳ 13.5 0.5 Uặmg)1, and Km (phosphoenolpyruvate) ¼ 0.28 ± 0.03 mm for dimers, and Vmax ẳ 3.7 0.3 Uặmg)1, and Km (phosphoenolpyruvate) ¼ 0.38 ± 0.07 mm for monomers, respectively Thus disruption of subunit–subunit interactions resulted in a significant decline in enzyme activity ( 3-fold), but did not have much effect on Km We also compared the thermal stability of monomeric (Mim) and dimeric forms Equal amounts of protein were incubated at three different temperatures (4 °C, 37 °C and 50 °C), and activity was assayed at different time intervals The dimeric form was stable for a prolonged (£ 250 min) duration at 37 °C, and showed  20–25% inactivation at 50 °C In comparison, the monomeric form was  80% inactivated at 37 °C (£ 250 min) and completely FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS 1937 Dissociation of enolase into active monomers I Pal-Bhowmick et al A Imidazole monomer +H+ (slow) MpH Mnt -imidazole (slow) Native monomer -H+ (fast) pH monomer Scheme Schematic representation of dissociation of the dimeric (D) form of enolase to the imidazole-induced and pH-induced monomeric (Mim or MpH) forms The dimer is shown to dissociate into the native form (Mnt), which has an enzyme activity similar to that of the dimer Lowering of the pH or addition of imidazole (or both) stabilize different conformations of monomers Mim and MpH differ in activity and stability from the dimer and Mnt MpH is rapidly converted into Mnt on raising of the pH, whereas Mim is slow to convert to Mnt, a form that is competent to form dimers inactivated at 50 °C in £ 150 (Fig 5B) Thus, subunit–subunit interface interactions confer higher thermal stability to the protein As enolases from different organisms are known to differ in their response to different salts [43], we examined the effect of various salts on the catalytic activity of the monomeric and dimeric forms To assess the effect of a salt, we assayed the enzyme with assay mixtures containing different concentrations of a salt Figure 6A–C shows the effects of NaCl, KCl and KBr on the activity of the dimeric and monomeric (Mim) forms of r-Pfen NaCl inhibited both forms of the enzyme, but the inhibition was stronger for the monomeric form (Fig 6A) In the case of KCl and KBr, the dimeric form was mildly activated ( 10–20%), whereas the monomeric form was strongly inhibited (Fig 6B,C) For assessing the activating ⁄ inactivating effect of the salts (NaCl, KCl and KBr), the enzyme was incubated in buffer containing several different concentrations of salt for 24 h at 20 ± °C, and then assayed in the absence of the salt The results are presented in Fig 6D–F At median concentrations ( 300 mm), NaCl and KCl had an activating effect on the dimeric form of the enzyme, whereas all three salts had a concentration-dependent inactivating effect on the monomeric form Comparison of CD and fluorescence spectra of monomeric and dimeric enolase The effect of the loss of subunit–subunit interface interactions on the secondary and ⁄ or tertiary structure 1938 0.0 0.2 0.4 0.6 0.8 1.0 [PEP] (m M) B 1.4 1.2 Fractional activity Mim + imidazole (slow) Activity (Units mg -1) 10 Native dimer (D) 1.0 0.8 0.6 0.4 0.2 0.0 50 100 150 200 250 300 Time (min) Fig (A) Comparison of monomer and dimer activity with varying concentrations of phosphoenolpyruvate For activity measurements, 50 lL of lM r-Pfen (monomeric or dimeric) was added to 450 lL of assay mixture As monomeric protein was in 250 mM imidazole, the final concentration of imidazole in the assay mixture was 25 mM The presence of 25 mM imidazole in the assay mixture had no effect on dimer activity Data were fitted to the Michaelis– Menten equation The best-fit parameters are Km (phosphoenolpyruvate) ¼ 0.38 ± 0.07 mM and Vmax (specific activity) ¼ 3.7 ± 0.3 Uặmg)1 for monomer (j), and Km (phosphoenolpyruvate) ẳ 0.28 0.03 mM and Vmax ẳ 13.5 0.5 Uặmg)1 for dimer (h) (B) Temperature dependence of stability of dimeric (open symbols) and monomeric (Mim) (filled symbols) forms of r-Pfen Enzyme was incubated at °C (s or d), 37 °C (n or m) and 50 °C (h or j) The activity was assayed at different time intervals of enolase was probed by recording CD and fluorescence spectra of monomeric and dimeric forms of the protein As monomer preparations made by imidazole treatment contain about 250 mm imidazole, which interferes with fluorescence and CD spectra, it was not possible to record meaningful spectra for Mim Instead, we recorded CD and fluorescence spectra for r-Pfen and Yen MpH, and compared them with the spectra from dimers (Fig 7) CD spectra of monomeric and dimeric forms (r-Pfen and Yen) were very similar (Fig 7A,C) Analysis of r-Pfen CD spectra using the FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS I Pal-Bhowmick et al Dissociation of enolase into active monomers 140 140 A 120 140 B 100 100 80 80 80 60 60 60 40 40 40 20 140 20 140 C 120 100 % Activity 120 20 140 D F E 120 120 120 100 100 100 80 80 80 60 60 60 40 40 40 100 200 300 400 500 600 [NaCl] (mM) 100 200 300 400 500 600 [KCl] (mM) 100 200 300 400 500 600 [KBr] (mM) Fig Effect of NaCl, KCl and KBr on the activity of monomeric (Mim) (j) and dimeric (h) r-Pfen Upper panel (a, b, c): Enzyme samples were assayed in the presence of different concentrations of salts Lower panel (d, e, f): Enzyme samples (200 lL of lM r-Pfen) containing different concentrations of salts were incubated for 24 h at 20 °C, and assayed in the absence of the salt For each assay, 50 lL of the enzyme was added to 450 lL of the assay mixture Activity is expressed as percentage of control where no salt was added to the enzyme sample cdnn program for secondary structure analysis [44] gave helix 36.6%, beta 14.1%, turn 19.3% and random 29.9% for the dimeric form, and helix 31.5%, beta 18.3%, turn 18.4% and random 31.8% for the monomeric form Thus, it appears that dissociation of dimers to form monomers lead to a slight decrease in helical content with a concomitant increase in the beta sheet content Fluorescence emission spectra of the monomic, dimeric and denatured states of r-Pfen and Yen are shown in Fig 7B,D For both of these enzymes, dissociation of enolase into monomers led to a decrease in emission intensity In the case of r-Pfen, the emission maximum was blue-shifted upon dissociation (Fig 7B; compare traces a and b), whereas for Yen, a slight red shift was observed (Fig 7D) Such a red shift may arise because the dimer interface of Yen has Trp56 (which becomes solvent-exposed upon dissociation), whereas the analogous position in Pfen is occupied by Tyr59 Discussion Alignment of enolase sequences from several different organisms have shown that it is a highly conserved protein [18] Furthermore, the comparison of known three-dimensional structures showed complete posi- tional conservation of active site residues across species [5,6,41,45–49] The enolase polypeptide chain folds into two domains, with the small domain having a mixture of a-helices and b-sheets, and the large domain having an a ⁄ b-barrel structure (Fig 8D) The binding of substrate brings these two domains together to constitute an active site As most enolases are homodimeric, the intersubunit interface is formed by contacts between the small domain of one subunit and the large domain of the other subunit If the subunit–subunit interface is large, the dissociated monomers are generally unstable, due to exposure of large hydrophobic surfaces to the solvent [50] The intersubunit interface in enolases is rather small ( 11–13% buried surface) and quite hydrophilic, suggesting that it may be possible to dissociate the dimer into active monomers In the past, several attempts have been made to dissociate the dimeric enolase into active monomers [7–9,11,14–17] However, the active monomeric form could never be obtained in isolation from the dimeric form Failure to obtain active monomers of enolase strengthened the belief that enolases are active only in the dimeric state [51] and that the quaternary structure of enolase is necessary for catalytic activity Thus, any attempt to dissociate the dimer is accompanied by changes in the tertiary and secondary structures that are detrimental to enzyme activity [8] However, the FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS 1939 Dissociation of enolase into active monomers I Pal-Bhowmick et al 4000 4000 2000 A r-Pfen Mean Residue Ellipticity -1 (degrees cm dmol ) Mean Residue Ellipticity (degrees cm2 dmol -1) -2000 -4000 -6000 -8000 b -10000 a -12000 C Yen 2000 -2000 -4000 -6000 -8000 b -10000 -12000 -14000 a -14000 200 210 220 230 240 250 200 260 210 Wavelength (nm) 240 250 260 3.5e+8 B r-Pfen D Yen a 5e+8 b 4e+8 3e+8 c 2e+8 a 3.0e+8 Fluorescence Intensity Fluorescence intensity 230 Wavelength (nm) 7e+8 6e+8 220 b 2.5e+8 c 2.0e+8 1.5e+8 1.0e+8 1e+8 5.0e+7 0.0 320 340 360 380 320 340 360 380 Wavelength (nm) Wavelength (nm) Fig Comparison of CD and fluorescence spectra of monomeric (MpH) and dimeric forms of r-Pfen and Yen (A) CD spectra of (a) dimer and (b) monomer of r-Pfen (B) Fluorescence spectra of (a) dimer, (b) monomer and (c) urea-denatured r-Pfen (C) Yen CD spectra: (a) dimer, (b) monomer (D) Fluorescence spectra of (a) dimer, (b) monomer and (c) urea-denatured Yen Fluorescence emission intensities were normalized for protein concentration results presented here unequivocally establish that both H+ and imidazole are able to dissociate enolase dimers into active monomers that could be isolated from the dimeric form (Figs 1–3) We observed that at any given concentration, the ability of imidazole to dissociate the dimer was enhanced by low pH (Fig 1B) In the absence of imidazole, a change in pH (in the range 6–8) did not have any significant effect on the dissociation of the enzyme (Fig 1B), suggesting that the increased dissociation induced by imidazole at low pH is mostly due to pH-dependent ionization of imidazole Imidazole has a pKa of 6.9, and hence at pH 6.0 it will exist predominantly as an imidazolium ion Thus, the data presented in Fig 1B are commensurate with the idea that it is the imidazolium ion that is effective in dissociating the r-Pfen dimer into monomers (Mim) The pH-dependent dissociation of various enolases indicate that it is the protonation of group(s) at the intersubunit interface with a pKa  5–5.5 that is responsible for the dissociation (Fig 3) An examination of the enolase intersubunit interface showed that it is stabilized by two salt bridges, several 1940 hydrogen bonds, and p–cation (Tyr or Trp-Arg) and hydrophobic interactions A relatively low contribution of the salt bridge in stabilizing the dimer interface was evident from the fact that replacement of an interface glutamate residue with a leucine (E414L) in RMen did not result into the dissociation of dimer DGo for dissociation for the mutant enzyme decreased from 49.7 kJỈmol)1 to 42.3 kJỈmol)1 [7] One may ask the following question: which are the subunit interface interactions that imidazolium and ⁄ or hydrogen ions can possibly disrupt? An examination of the intersubunit interface suggests the following two possibilities (a) There is a hydrogen bond between His191 NE2 and the carbonyl oxygen of Gly15 (or Arg14) where His191 NE2 acts as a proton donor (Fig 8A,B) Imidazole, being an analog of histidine, can compete for this hydrogen bond The importance of this interaction in stabilizing the dimer is evident from the observed changes in the intersubunit hydrogen bond pattern on Mg(II) binding In one Mg(II)-bound form of Yen (Protein Data Bank: 2ONE, 1EBH), a hydrogen bond forms between His191 and Arg14 (Fig 8B) with a bond FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS I Pal-Bhowmick et al Dissociation of enolase into active monomers A B C D Fig Intersubunit interface hydrogen bond involving His191 in enolase crystal structures (A) Yen with two Mg(II) ions bound per subunit (Protein Data Bank: 1EBG) showing hydrogen bond between His191 and Gly15 (B) Yen with one Mg(II) ion bound per subunit (Protein Data Bank: 2ONE) has a hydrogen bond between His191 and Arg14 (C) Neuronal enolase in which two subunits are asymmetrically bound to one and two Mg(II) ions exhibits both types of hydrogen bond The subunit bound to one Mg(II) ion has a hydrogen bond between His189 and Arg14, and the other subunit, which is bound to two Mg(II) ions (Protein Data Bank: 1TE6), has a His189–Gly15 hydrogen bond (D) Ribbon diagram of dimeric Yen (Protein Data Bank: 1EBG), in which yellow and blue represent two subunits Interface residues (Trp56 and Arg184) involved in p–cation interaction are shown in a stick representation length (as measured between two non-hydrogen atoms ˚ in the hydrogen bond) of 3.2–3.3 A In two Mg(II)bound forms (Protein Data Bank: 1ONE, 1EBG), the hydrogen bond is between His191 and Gly15 (Fig 8A) ˚ with a bond length of 2.9–3.0 A Thus, the loss of Mg(II) may favor weaker hydrogen bonds and dissociation, whereas addition of Mg(II) may reverse the process, leading to subunit–subunit association It is interesting to note that in the neuronal enolase crystal structure (Protein Data Bank: 1TE6), where one subunit is bound to one Mg(II) ion and the other subunit has two Mg(II) ions, both types of hydrogen bond (His–Gly and His–Arg) are observed (Fig 8C) Our observations that Mg(II) favors dimer formation are in agreement with earlier reports [15], and suggest that interactions with His191 are critical for dimer stability (b) The imidazolium cation can compete with cationic side chains if a p–cation interaction is present at the dimer interface Recent analysis of dimeric interfaces has shown that p–cation interactions can make significant contributions to the binding energy for protein– protein complex formation, and it is suggested that such interactions be included in the list of criteria for characterizing protein interfaces [52] We examined the FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS 1941 Dissociation of enolase into active monomers I Pal-Bhowmick et al enolase interface using PP server, which identified Trp56 in Yen (Tyr56 in RMen, Tyr59 in Pfen) as an interface residue The program capture (Cation-p Trends Using Realistic Electrostatics) [53] was used to identify interactions between the cationic group of lysine or arginine and the aromatic rings of phenylalanine, tyrosine and tryptophan Such a search on Yen led to the identification of a Trp–Arg (Fig 8D) interaction with a calculated electrostatic energy of Ees ẳ ) 16.3 kJặmol)1 and a Van der Waals energy of Evdw ẳ ) 7.5 kJặmol)1 In the modeled structures of RMen and r-Pfen, similar interactions occur with Tyr56 and Tyr59, respectively The energy of interaction computed for Tyr and Arg in the modeled structures of RMen (Ees ẳ ) 4.4 kJặmol)1 and Evdw ẳ ) 4.2 kJặmol)1) and r-Pfen (Ees ẳ ) 8.8 kJặmol)1 and Evdw ẳ ) 3.8 kJặmol)1) were found to be much smaller than that for Yen Thus, this particular interaction appeared to be much stronger in Yen than in r-Pfen and RMen This could be a possible reason for the inability of imidazole to dissociate Yen, unless 300 mm NaCl is included in the buffer (Table 1), whereas RMen and r-Pfen become dissociated in the absence of salt An aromatic ring can have two possible modes of interaction with imidazole [54], namely p–p and NH–p interactions, whereas the imidazolium cation has an additional possibility of a p–cation interaction [55] The observed greater effectiveness of the imidazolium ion in dimer dissociation may arise from its ability to undergo p–cation and stronger NH–p interactions with an aromatic residue (Trp or Tyr) at the interface The enolase interface has two salt bridges (Glu20– Arg414 and Arg8–Glu417 in Yen; Glu20–Arg411 and Arg8–Glu414 in RMen; Glu23–Arg415 and Arg11– Glu425 in r-Pfen) involving two glutamate residues pH-induced dissociation of the enzyme may arise due to protonation of the side chains of these glutamate residues at low pH (pKa  4.5–5.5) Although r-Pfen is highly homologous to mammalian enolases, we have observed an interesting difference between these two proteins; namely, imidazole can dissociate r-Pfen in presence of Mg(II), whereas RMen could not be dissociated Thus, there is a possibility of selective dissociation of r-Pfen into a lowactivity, unstable monomeric form that can lead to significant reduction in enolase activity Such a partial decrease in enolase activity combined with low stability could severely hamper the glucose-metabolizing capacity of the parasite As activity of glycolytic enzymes is known to be essential for the growth of Plasmodium in the intraerythrocytic stages [56], analogs of imidazole (with higher pKa) that may be able to dissociate parasite enolase at physiologic pH and in the presence of Mg(II) might prove to be parasite growth inhibitors 1942 The comparison of various properties between the dimeric and monomeric forms of enolase showed that the monomeric form of enolase is catalytically inefficient (Kcat ⁄ Km is  5-fold less than that of the dimer) and thermally unstable It is highly unlikely that such a catalytically inefficient form of enolase is utilized for high-flux glycolytic function in the parasite However, in recent years, enolase has also been recognized as a multifaceted protein with a variety of nonglycolytic novel biological functions [18,57] It is likely that, when executing some of these moonlighting functions, enolase may interact with other proteins In such cases, the monomeric form of enolase may be more suitable, as it has a freely accessible subunit-interacting surface One such case is the 48 kDa s-crystallin protein in the eye lens of the lamprey [58], which has been shown to be a monomeric form of a-enolase [59] As enolases undergo several post-translational modifications inside cells, it is likely that some of these may stabilize the monomeric form of enolase to enable it to participate in diverse physiologic functions Materials and methods Materials Hexahistidine-tagged r-Pfen was purified as described previously [24] Yen was purchased from Sigma Chemical Co (St Louis, MO, USA) RMen and all other enzymes were purchased from Boehringer-Ingelheim GmbH (Ingelheim, Germany) Phosphoenolpyruvate was purchased from Sigma Tris, imidazole and magnesium chloride were obtained from USB (Amersham-Buchler, Braunschweig, Germany) All other reagents used in this study were of analytical grade and were used as received Gel filtration chromatography Gel filtration chromatography was performed with a Superdex-75 prep grade column (Hiload HR 16 ⁄ 60, using an AKTA FPLC system supplied by Amersham-Pharmacia Biotech (Kwai Chung, Hong Kong) Before every run, the column was pre-equilibrated with two column volumes of desired buffer The typical flow rate was 1.5 mLỈmin)1, and the fraction volume was mL Blue dextran (2000 kDa), b-amylase (200 kDa), alcohol dehydrogenase (150 kDa), BSA (67 kDa), ovalbumin (43 kDa) and chymotrypsinogen (25 kDa) were used as molecular weight calibration markers Enzyme activity assay Enolase activity was measured by a continuous spectrophotometric assay by monitoring the formation of 2-PGA FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS I Pal-Bhowmick et al from phosphoenolpyruvate at 240 nm using a Perkin-Elmer (Waltham, MA, USA) lambda 40 spectrophotometer (T ¼ 20 ± °C) The assay mixture contained 1.5 mm phosphoenolpyruvate and 1.5 mm MgCl2 in 50 mm Tris ⁄ HCl (pH 7.4) A unit of enzyme was defined as the amount of enzyme that converts lmol of phosphoenolpyruvate into 2-PGA in at 20 °C Appropriate correction was applied for differences in absorbance of phosphoenolpyruvate at 240 nm with varying pH and Mg(II) concentration [60] Kinetic data were fitted to the Michaelis–Menten equation using sigmaplot Fluorescence and CD spectra The fluorescence measurements were carried out by using a SPEX Fluorolog FL1T11 spectrofluorometer (Edison, NJ, USA) equipped with a thermostated cell holder All fluorescence intensities were corrected for variable background emissions and lamp fluctuations (signal ⁄ reference: s ⁄ r) Fluorescence emission intensities were normalized for protein concentration (2–10 lm range) The excitation wavelength used was 295 nm, and the emission was measured in the range 310–390 nm The scan speed was nmỈs)1, and two scans were averaged for each spectrum CD studies were carried out using a Jasco J-810 spectropolarimeter (Tokyo, Japan) Far-UV CD studies were carried out using a mm path-length cuvette with 2–10 lm protein concentration at constant temperature of 20 ± °C Far-UV CD spectra were measured over the 200–270 nm range, and each spectrum was taken by averaging 10 scans at 20 nmỈs)1 scan speed The data are presented as mean residue ellipticities and were calculated using the formula: ẵh ẳ hobs =ð10 ClÞ where hobs is the observed ellipticity in mdeg, l is the path length in cm, and C is the concentration of 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Am J Hum Genet 76, 911–924 58 Stapel SO & de Jong WW (1983) Lamprey 48-kDa lens protein represents a novel class of crystallins FEBS Lett 162, 305–309 59 Wistow GJ, Lietman T, Williams LA, Stapel SO, de Jong WW, Horwitz J & Piatigorsky J (1988) Taucrystallin ⁄ alpha-enolase: one gene encodes both an enzyme and a lens structural protein J Cell Biol 107, 2729–2736 60 Wold F & Ballou CE (1957) Studies on the enzyme enolase I Equilibrium studies J Biol Chem 227, 301–312 FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS 1945 ... enolase into monomers using small molecules, and characterized the properties of the monomeric state We report the successful dissociation of r-Pfen, and yeast and rabbit muscle enolases, into isolatable... dissociation of yeast enolase into enzymatically active monomers Biochim Biophys Acta 327, 176–185 13 Keresztes-Nagy S & Orman R (1971) Dissociation of yeast enolase into active monomers Biochemistry... of CD and fluorescence spectra of monomeric (MpH) and dimeric forms of r-Pfen and Yen (A) CD spectra of (a) dimer and (b) monomer of r-Pfen (B) Fluorescence spectra of (a) dimer, (b) monomer and