Original articleJack bean (Canavalia ensiformis) urease. Probing acid–base groups of the active site by pH variation
Introduction
Urease (urea amidohydrolase, EC 3.5.1.5) catalyzes the hydrolysis of urea to produce ammonia and CO2. Present in many plants, bacteria, fungi and algae and in soil as a soil enzyme, the enzyme thus plays an important role in the overall metabolism of nitrogen in nature. Primarily, urease allows plants and microorganisms to utilize urea, internally derived or external, to generate ammonia as a nitrogen source for growth [7], [19], [29]. Of great moment in enzymology, urease obtained from jack bean (Canavalia ensiformis) was the first enzyme ever crystallized (1926) [36] and the first nickel-containing enzyme identified (1975) [11].
The reaction catalyzed by urease is deceptively simple (Scheme 1). It consists of the fast hydrolysis of urea to ammonia and carbamate, followed by a spontaneous decomposition of the carbamate to ammonia and carbonic acid [7], [19].
Otherwise, owing to its resonance stabilization urea is highly stable in aqueous solutions and resists decomposition. The half-time of the uncatalyzed decomposition of urea is of the order of 3.6 years, and follows a different mechanism that yields the elimination products (Scheme 1). These peculiar features render urease the most proficient enzyme identified to date [14]. For its enzymatic hydrolysis urease utilizes an active site containing a binuclear nickel center bridged by a carbamylated lysine and a hydroxide ion (Scheme 1) as was shown by the crystallographic structures resolved for three different bacterial ureases, from Klebsiella aerogenes [20], Bacillus pasteurii [4] and Helicobacter pylori [18]. The nearly superimposable active sites in these ureases imply that this structure of an active site is common to all ureases, whether of bacterial, plant or fungal origin.
A long history and extensive research notwithstanding, urease has a catalytic mechanism that is still a matter of debate [7], [19]. Final elucidation of the mechanism is of importance for strategies to combat undesirable effects brought about by the enzyme, to which belong reaction-generated ammonia and an increase in pH. These were shown to have profound medical and agricultural implications [29]. Several ureolytic bacteria have been recognized as pathogenic factors in human/animal infections of urinary and gastrointestinal tracts. In the former they are involved in the urinary stone formation, catheter encrustation and pyelonephritis, and in the latter in chronic active gastritis, peptic ulcers, both induced by H. pylori, and in hepatic coma. In agriculture urease is essential for converting urea fertilizers to utilizable ammonia. Too rapid a hydrolysis, however, results both in plant damage by ammonia toxicity and in the alkalization of soil, and finally in the loss of nitrogen by ammonia volatilization, thereby creating severe environmental and economic problems.
One pragmatic approach to the elucidation of enzyme mechanisms is the analysis of pH-variation studies of enzyme steady-state kinetic parameters. Such an analysis provides information on the ionization states of the components of the enzyme reaction, i.e. of the free enzyme, enzyme–substrate complex, and substrate, and thus helps to resolve the involvement of their acid–base functional groups in the catalytic mechanism [9], [10].
The active site cavity of ureases features several ionizable amino acid residues that are conserved principally in all known ureases and that are thought to participate in the catalytic reaction. Accordingly, these groups along with the Ni-bound water molecules (Scheme 1), should be considered responsible for the observed pH profiles of urease kinetic parameters. Remarkably, despite numerous reports on the pH profiles of urease steady-state kinetic parameters, there is no consensus among the investigators on the number of acid–base groups required for the catalysis, their pKas values and functions exercised in the catalytic mechanism. An array of shapes of the profiles has been obtained and interpreted by an array of ways to provide disparate results.
For most ureases the Michaelis constants KM, falling in value in the range 1–4 mM [5], [12], [13], [17], [23], [26], [30], [32], [33], [35], [37], [39], have been found to be only slightly dependent on pH [5], [12], [17], [23], [35], [39]. Unlike KM, the maximum reaction rate vmax is known to be strongly pH dependent. Most frequently, bell-shaped vmax-pH profiles in the pH range ca. 4.5–10.5 with the optimum pH around 7–8 have been reported for plant, bacterial and fungal ureases [1], [5], [13], [16], [17], [23], [25], [26], [27], [28], [31], [32], [33], [35], [37], [39] and analyzed in terms of two macroscopic pKa values, one on the acidic and the other on the basic side of the curve. Less frequently, three pKas have been obtained by combining vmax-pH curves noted in different buffers with the data derived from KM-pH curves [1], [27]. In few instances, for urease from H. pylori [6], [15], [38], from soybean leaf [21] and from jack bean [6], [22], irregular vmax-pH profiles exhibiting an additional optimum or a hump on the acidic side have been reported, and in some profiles they have been seemingly overlooked [23], [28], [38], [39]. This irregularity implies that the commonly accepted bell-shaped vmax-pH profiles might contain some more complex features, concealed under the data. In two studies, one on jack bean [12] and the other on K. aerogenes urease [30] this irregularity has been interpreted as involving three and four ionizing groups, respectively, in the urease reaction mechanism.
In addition to the common pH-dependence studies of KM and vmax, there are other methods of identifying enzyme ionizing groups [8], [9], among them the method based on the pH-dependence of the inhibition constant Ki of enzyme competitive inhibitors. The method is valuable in that it is capable of differentiating between the enzyme groups involved in binding from those involved in the catalytic reaction [8], [9]. This is because a competitive inhibitor can mimic substrate binding to enzyme but is unable to go through any catalytic steps, and accordingly, the pH-dependence of Ki can be used to determine binding pKas for substrate. Unlike vmax-pH and KM-pH, the method has not been exploited in studies of urease.
Aiming at defining the pKa values and the functions of ionizable groups of the urease active site in the catalysis that could help elucidate its mechanism, in this study we investigated the pH effects on jack bean urease by performing pH-dependent measurements of both the steady-state kinetic parameters and the inhibition constant of a urease competitive inhibitor, boric acid, using organic (MES, HEPES and CHES) buffers. We compare the data with those obtained previously in phosphate buffers (vmax-pH in 22 mM buffer and -pH) [23] and extended here for higher phosphate concentrations and for boric acid inhibition. The buffers proposed are distinct in that while organic buffers are noninhibitory [22], phosphate buffer is a simple competitive inhibitor of jack bean urease at pH < 7.5 [23]. By combining the results we demonstrate that three ionizable groups of the urease active site are involved in the catalytic mechanism. A group with a pKa = 5.3 is required for substrate binding and catalysis, a group with a pKa = 9.1 has a role in the catalysis, while the third group featuring a pKa = 6.6 participates in the binding and/or modulates the structure of the active site.
Section snippets
pH profiles of vmax and KM
The pH profiles of vmax and vmax/KM obtained in the buffers examined are compiled in log–log plots in Fig. 1. Both the maximum reaction rate vmax (Fig. 1a) and its ratio to the Michaelis constant vmax/KM (Fig. 1b) decreased at low and high pH with limiting slopes close to +1 and –1, respectively, thereby revealing that two functional groups are involved in the enzymatic process.
The data fitted to Eq. (3) gave the following pairs of values: pK2 = 5.06 ± 0.06 and pK1 = 9.26 ± 0.06 from the vmax profile,
Materials
Jack bean urease (Sigma type III, activity 22 units mg−1 solid), MES, HEPES, CHES buffers (SigmaUltra) and urea (Sigma Molecular Biology) were used for all experiments. NaH2PO4·H2O and Na2HPO4·12H2O, Na2EDTA and boric acid were from POCh, Poland. MES, HEPES and CHES buffer stock solutions (200 mM) were prepared by dissolving the required amounts in water and had their pH values adjusted with a NaOH solution. The pH ranges were: 5.00–7.03 for MES, 6.47–8.55 for HEPES and 8.36–9.55 for CHES, as
Acknowledgements
This work was supported by the KBN grant no. PB 7/T09A/048/20. S.C. acknowledges funds from the Italian MIUR (PRIN 2003).
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