Urease Inhibition in the Presence of N-(n-Butyl)thiophosphoric

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Urease inhibition in the presence of N-(n-butyl)thiophosphoric triamide, a suicide substrate: structure and kinetics Luca Mazzei, Michele Cianci, Umberto Contaldo, Francesco Musiani, and Stefano Ciurli Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.7b00750 • Publication Date (Web): 31 Aug 2017 Downloaded from http://pubs.acs.org on September 1, 2017

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Urease inhibition in the presence of N-(n-butyl)thiophosphoric triamide, a suicide substrate: structure and kinetics Luca Mazzei,† Michele Cianci,‡ Umberto Contaldo,† Francesco Musiani,† and Stefano Ciurli*,† †

Laboratory of Bioinorganic Chemistry, Department of Pharmacy and Biotechnology, University

of Bologna, Italy. ‡

Department of Agricultural, Food and Environmental Sciences, Università Politecnica delle

Marche, Ancona, Italy.

ABSTRACT The nickel-dependent enzyme urease is a virulence factor for a large number of pathogenic and antibiotic-resistant bacteria, as well as a negative factor for the efficiency of soil nitrogen fertilization for crop production. The use of urease inhibitors to contrast these effects requires the knowledge, at the molecular level, of their mode of action. The 1.28-Å resolution structure of the enzyme-inhibitor complex obtained upon incubation of Sporosarcina pasteurii urease with N-(nbutyl)thiophosphoric triamide (NBPT), a molecule largely utilized in agriculture, reveals the presence of the mono-amidothiophosphoric acid (MATP) moiety, obtained upon enzymatic hydrolysis of the diamide derivative of NBPT (NBPD) to yield n-butyl amine. MATP is bound to the two Ni(II) ions in the active site of urease using a µ2-bridging O atom and terminally bound

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O and NH2 groups, with the S atom of the thiophosphoric amide pointing away from the metal center. The mobile flap modulating the size of the active site cavity is found in the closed conformation. Docking calculations suggest that the interaction between urease in the open flap conformation and NBPD involves a role for the conserved αArg339 in capturing and orienting the inhibitor prior to flap closure. Calorimetric and spectrophotometric determinations of the kinetic parameters of this inhibition indicate the occurrence of a reversible slow-inhibition mode of action, characterized, for both bacterial and plant ureases, by a very small value of the dissociation constant of the urease-MATP complex. No need to convert NBPT to its oxoderivative NBPTO, as previously proposed, is necessary for urease inhibition.


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INTRODUCTION Urease (urea aminohydrolase, E.C. 3.5.1.5) is a nickel-dependent enzyme found in a large variety of organisms, including plants, algae, fungi, and several prokaryotes.1-5 It is involved in

the global nitrogen cycle, catalyzing the rapid hydrolytic decomposition of urea (Scheme 1):6,7 Scheme 1 Urease catalysis causes an overall pH increase3-5 that has negative consequences both on human health8 and agriculture.9 Urease causes a significant decrease in the efficiency of soil nitrogen fertilization with urea, due to ammonia volatilization as well as root damage caused by the increase in pH. Moreover, urease is the main virulence factor for a large variety of human pathogens, such as Helicobacter, Staphylococcus, Clostridium, Vibrio, Mycobacterium, Yersinia, Escherichia, Proteus, Ureaplasma, Klebsiella, Pseudomonas, Corynebacterium, Providencia, Morganella, and Cryptococcus spp. Among the twelve antibiotic-resistant priority pathogens listed in 2017 by the World Health Organization,10 ten are ureolytic bacteria that take advantage of the urease activity to survive in the host organism. This observation imposes urease to the attention of the scientific community as a target to develop new drugs for the treatment of important bacterial infections. The enzymatic hydrolysis of urea occurs in two steps: the first, strictly requiring urease, consists of the decomposition of urea to give ammonia and carbamate, followed by the spontaneous reaction of carbamate to give a second molecule of ammonia and bicarbonate


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(Scheme 2).3-5 The enzymatic reaction has a half-time of a few microseconds, with a kcat/KM that is 3 × 1015 times higher than the rate of the uncatalyzed reaction, making urease the most

efficient hydrolase known.11 Scheme 2 The available crystal structures of ureases from several bacteria and higher plants show a typical trimeric assembly.3-5 Each assembly is in turn composed by a single chain, [(α)3] in higher plants, as in the cases of jack bean (Canavalia ensiformis) urease (JBU)12 and pigeon pea (Cajanus cajan) urease,13 by two chains [(αβ)3] in the case of Helicobacter pylori,14 and by three chains [(αβγ)3] in the cases of Sporosarcina pasteurii (SPU) and Klebsiella aerogenes.3-5 The minimal trimer eventually forms [(α)3]2 dimers in higher plants or nearly spherical [(αβ)3]4 tetramers in H. pylori.14 Each trimeric assembly hosts three conserved active sites, containing two Ni(II) ions (Fig. 1A).3-5 The urease inhibitors structurally characterized so far5 can be roughly divided in two main families, based on the mode of interaction with the enzyme. The first class contains molecules such as phosphate, diamidophosphate (DAP), thiols, sulfite, fluoride as well as hydroxamic, citric and boric acids, which bind the Ni(II) ions in the active site.3-5 The second class of inhibitors is composed of molecules such as β-mercapto-ethanol (BME),15 1,4-benzoquinone,16 and catechol17 that have been proven to bind to the Sγ atom of the conserved αCys322 [SPU numbering]. This residue is located onto a mobile helix-turn-helix motif (flap) that is essential


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for catalysis by regulating the transit of substrate and products through the active site cavity (Fig. 1B), and for modulating the position of the essential αHis323 with respect to the substrate during catalysis. BME can actually be ascribed to both classes, because it is able to further use its thiol moiety to coordinatively bridge the Ni(II) ions in the active site.15-17

Figure 1. (A) Coordination geometry of the two Ni(II) ions in native SPU active site (PDB code 4CEU54). (B) Reconstruction of the flap closure (transparent ribbons) starting from the open (light and dark blue ribbons, PDB code 4CEU) and ending in the closed (PDB code 3UBP) conformations in SPU, highlighting the side chains of αLys220*, αCys322, and αHis323. Phosphoramides are a class of well-known and very potent urease inhibitors, acting with a slow-binding inhibition mechanism.18 The inhibition process appears to involve a first hydrolytic event performed in situ by urease, with the subsequent trapping of a tetrahedral moiety that


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blocks the enzyme active site by mimicking the transition state that would be formed during the enzyme reaction of urea hydrolysis. It has been demonstrated that, irrespective of the starting compound, the inhibition is always brought about by DAP.19-21 In this context, significant information on the structural basis of urease inhibition by phosphoramides was provided by the X-ray crystal structure of SPU in complex with DAP after the treatment of the enzyme with phenylphosphorodiamidate (PPD).22 N-diaminophosphinothioylbutan-1-amine, commonly known as N-(n-butyl)thiophosphoric triamide (NBPT hereafter), is a particular derivative of such compounds, where the oxygen atom of the phosphoryl scaffold has been replaced by a sulfur atom. NBPT is mostly used in agriculture as a nitrogen stabilizer to reduce ammonia volatilization,23 even though significant negative aspects of this practice have been reported.24-30 Several studies have been conducted on the efficacy of NBPT in inhibiting ureases from different sources and in reducing ammonia volatilization upon urea addition, either in vitro or in soils.31-36 These studies commonly assert that NBPT has little or no effect as a urease inhibitor, while it must be converted to its oxoanalogue, N-(n-butyl)phosphoric triamide (NBPTO), in order for a strong inhibition to occur.31-36 Also, the latter conversion appears to take place predominantly in soils, rather than in solution.35 This common idea has been partially challenged by evidences demonstrating that NBPT itself is able to inhibit plant urease without necessarily being converted to NBPTO.37 In addition to these controversies, none of these studies clarified the identity of the actual species that is responsible for urease inhibition in the presence of NBPT, with some speculation suggesting that the moiety interacting with urease is DAP in the case of NBPTO and the unaltered inhibitor in the case of NBPT.37


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The present study reports the X-ray crystal structure of urease from S. pasteurii, a widespread and highly ureolytic soil bacterium, inhibited in the presence of NBPT. The structure, determined at 1.28 Å resolution, demonstrates that the sulfur derivative of NBPT is capable to interact with the nickel ions in the urease active site, undergoing an in situ hydrolysis that generates a tetrahedral moiety blocking the active site in a fashion similar to that proposed for phosphoramides. A structural analysis, corroborated by docking calculations, also suggests that NBPT must be converted to N-(n-butyl)thiophosphoric diamide (NBPD) prior to the formation of the structurally established complex. An extensive kinetic study of the inhibition of SPU and JBU in the presence of NBPT was then carried out to elucidate the nature of the urease inhibition mechanism and to compare, under identical assay conditions, bacterial and plant ureases. These results prove that a slow-binding inhibition mechanism occurs for both enzymes, with very similar kinetic parameters regardless of the biological source of the enzyme. These results clarify a long-standing controversy and pave the way to improve and develop derivatives of phosphoramides as general urease inhibitors for agricultural and medical applications.

MATERIALS AND METHODS Enzyme and inhibitor sources. SPU was expressed and purified from S. pasteurii following a previously described procedure.38 SPU quantification was carried out by measuring the urease activity using a pH-STAT method39 and considering its specific activity of 2500 units mg-1 and Mr = 250 kDa.40 JBU type C-3, powder (≥600,000 units/g) was purchased from Sigma-Aldrich SRL, Saint Louis, MO (U. S. A.), and quantified following the manufacturer’s information. NBPT and NBPTO were purchased from Apollo Scientific Apollo Scientific Ltd, Bredbury (U. K.) and Santa Cruz Biotechnology, Inc., Heidelberg (Germany), respectively. EDTA (2 mM)


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was always added to the protein solution in order to prevent possible inhibition effects by traces of transition metal ions in solution. Crystallization, data collection and structural determination. A 11 mg mL-1 urease solution in 20 mM HEPES buffer at pH 7.5, containing 50 mM Na2SO3 and 2 mM EDTA, was incubated for 1 h in the presence of increasing concentrations of NBPT (in the range 1-16 mM) dissolved in the same buffer. Subsequently, 2 µL of each SPU-NBPT solution was diluted with 2 µL of a precipitant solution consisting in 1.6-2.0 M ammonium sulfate dissolved in 50 mM sodium citrate buffer at pH 6.3, and containing the same concentration of NBPT. Crystallization trials were performed at 293 K using the hanging-drop method, equilibrating the drop against 1 mL of the precipitant solution using 24-well XRL Plates [Molecular Dimensions, Suffolk (U. K.)]. Rice-shaped protein crystals appeared in most of the crystallization conditions within few weeks and grew to a size of 0.1 x 0.1 x 0.3 mm3. Crystals were scooped up using cryoloops and transferred to a cryoprotectant solution of 20% ethylene glycol dissolved in 50 mM sodium citrate buffer at pH 6.3, also containing 2.4 M ammonium sulfate and the same concentration of NBPT present in the crystallization drop. The crystals were then flash-cooled and stored in liquid nitrogen. Diffraction data were collected at 100 K using synchrotron radiation at the EMBL P13 beamline of the Petra III storage ring, c/o DESY, Hamburg (Germany).41 Reflection images were recorded by performing helical scans along the crystal to achieve higher resolution by minimizing radiation damage. Data processing and reduction was carried out with XDS42 and AIMLESS.43 The crystals were isomorphous with respect to those of native urease and other complexes of the same enzyme. The crystal structure of SPU in complex with DAP (PDB code 3UBP, 2.00 Å resolution)22


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devoid of solvent molecules and ligands, was used as an initial model for the rigid body refinement of the αβγ SPU trimer, carried out using Refmac.44 Model building and water or ligand addition/inspection were conducted using Coot.45,46 The structure was isotropically refined, including the hydrogen atoms in the riding positions, and then anisotropically refined to final R and Rfree of 11.90 and 13.68, respectively. The X-ray diffraction data and final refinement statistics are given in Table 1. Figures were generated using PyMol (The PyMOL Molecular Graphics System, Version 1.8 Schrödinger, LLC.), and CrystalMaker (http://www.crystalmaker.com). The structure was deposited in the Protein Data Bank with the accession code 5OL4. Docking calculations. The DOCK 6.8 program suite47-49 was used, following a previously published procedure.50 The computational strategy involved four steps: i) the active site region was modeled as a set of overlapping spheres of variable radii; ii) an electrostatic potential grid was computed within the volume delimited as above; iii) the possible orientations of a ligand inside the reaction site were calculated together with their relative ligand-receptor interaction energies; and iv) the calculated binding poses were clustered and analyzed. The procedure took advantage of the tools included in the UCSF Chimera software.51 The molecular surface of urease was calculated using a probe atom with a radius of 1.0 Å52 and a dot density of 8 points/Å. A radius of 0.83 Å was used for the Ni(II) ions.53 The cavity surface was calculated considering the active site mobile flap in the closed conformation, taken from the structure reported in the present study, and in the open conformation (as found in PDB code 4CEU54). The reaction site cavity was filled with spheres having a radius variable from 1.0 to 4.0 Å. Partial charges were assigned to standard SPU residues by using the AMBER ff14SB force field.55 Partial charges for non-standard residues (i.e. the carbamylated lysine) and for ligands


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were calculated using the NWChem software56 at the HF/6-31G(d) level, and fitted using the RESP (restrained electrostatic potential) procedure.57 The grid calculation was performed in a box of 8 and 13 Å for SPU with the flap in the closed and in the open conformation, respectively. The grid points spacing was 0.3 Å, the cut-off distance for the interaction energy was 12 Å, and the dielectric factor was set to 4. The docking was performed by comparing the distances between the spheres used to fill the active site cavity and the distances of the atoms of the ligand (the all-atom model was used here). A value of 0.3 Å was used as convergence limit for all the calculations. The ligand molecules were considered flexible by using the fragmentation procedure implemented in DOCK. For each ligand, a total of 200 binding poses were generated. The root means square deviations (RMSD) of the calculated binding poses were calculated with respect to the crystallographic structure, when available, or the binding pose having the lowest interaction energy. The docked ligands were clustered using a cut-off of 0.5 Å.

Table 1. Data collection, processing and refinement statistics Data collection Wavelength (Å)

0.9537

Detector

DECTRIS Pilatus 6M

Crystal-to-Detector distance (mm)

225.3

Oscillation angle (degrees)

0.100

Number of images

1300

Space group

P6322

Unit cell (a, b, c, Å)

131.74, 131.74, 188.93

Resolution range (Å)a

97.67 – 1.28


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Total number of reflectionsa

3479631 (171907)

Unique reflectionsa

243949 (11805)

Multiplicitya

14.3 (14.6)

Completenessa (%)

99.4 (98.1)

Rsyma,b (%)

11.7 (226.0)

Rpima,c (%)

3.3 (62.5)

Mean I half-set correlation CC(1/2)a

0.999 (0.673)

Mean I/σ(I)a

15.9 (1.5)

Refinement statistics Monomers in the asymmetric unit

3

Rfactord (%)

11.90

Rfreed (%)

13.68

Cruickshank’s DPI for coordinate 0.032 errore based on Rfactor (Å) Wilson plot B-factor (Å2)

10.4

Average all atom B-factorf (Å2)

16.645

RMS (bonds)d

0.08

RMS (angles)d

1.40

Total number of atoms

7202

Total number of water molecules

782

Solvent content (%)

55.52

Matthews Coefficient (Å3/Da)

2.76

Ramachandran plotg Most favored regions (%)

89.8

Additionally allowed regions (%)

9.5

Generously allowed regions (%)

0.6

Disallowed regions (%)

0.2


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a

Highestresolution bin in parentheses; R = ∑ ∑ I − 〈I〉/ ∑ ∑ I , where I is the intensity of a reflection, and 〈I〉 is the mean intensity of all symmetry related reflections j; ½ c R ... = ∑1/N − 1 ∑ I − 〈I〉 / ∑ ∑ I , where I is the intensity of a reflection, 〈I〉 is the mean intensity of all symmetry related reflections j, and N is the multiplicity;69 and d Taken from REFMAC;44 Rfree is calculated using 5% of the total reflections that were randomly selected and excluded from refinement; b

e

DPI =

!"#

%$∙ '(!) ∙ *+,-. /⅓ 182

234567

9:;< /2=39367 >

, where Natoms is the number of the atoms

included in the refinement, Nrefl is the number of the reflections included in the refinement, Dmax is the maximum resolution of reflections included in the refinement, compl is the completeness of the observed data, and for isotropic refinement, Nparams ≈ 4Natoms;70 f Taken from BAVERAGE;71 g Taken from PROCHECK.71 Calorimetric studies on the enzymatic urea hydrolysis by JBU. The kinetic parameters for the enzymatic urea hydrolysis and NBPT inhibition were derived by calorimetric experiments carried out using a high-sensitivity VP-ITC (ITC: Isothermal Titration Calorimetry) microcalorimeter (MicroCal LLC, Northampton, MA, U. S. A.). For all experiments, the reference cell was filled with deionized water, and the temperature of the reference and sample cells was set and stabilized at 298 K. Stirring speed was 300 rpm, and thermal power was monitored every 2 s using high instrumental feedback. All urease and urea solutions were buffered using 50 mM HEPES, pH 8.0, containing 2 mM EDTA. The apparent molar enthalpy (∆@ A ) of the reaction of urea hydrolysis by JBU, a parameter necessary to derive BC and DEAF from calorimetric data, was determined through the so-called M1 experiment, as previously described for the case of SPU.38 A 1.25 nM solution of JBU and 12 mM urea were used in the sample cell (V = 1.4093 mL) and in syringe, respectively. A first injection of 10 µL urea was carried out, giving a final substrate concentration of 0.085 mM in the cell. After the thermal signal (µcal s-1) returned to the baseline level, indicating that the consumption of the substrate was complete, a second and a third injection were performed, following the same procedure. Numerical integration of the area under the three single peaks was


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carried out using the native Origin package provided by the calorimeter manufacturer, and the average value thus obtained was used to calculate the apparent molar enthalpy for urea hydrolysis in the experimental conditions used. The kinetic parameters BC and DEAF for the reaction of urea hydrolysis by JBU were determined through the so-called M2 experiment, similarly to what previously reported for SPU.38 A 25 pM solution of JBU and 0.5 M urea were used in the cell and in the syringe, respectively. A succession of 12 injections of 5 µL urea solution (with the exception for the first injection, that was of 2 µL volume) were carried out every 150 s, a time necessary to allow the thermal trace to reach a steady-state level after each injection and, at the same time, maintain pseudo-first-order reaction conditions. The thermal power obtained from the baseline shift was averaged using the last 15 s prior to the subsequent injection to obtain an accurate measurement. The calculated thermal power for each injection was converted to the corresponding reaction rate and corrected considering the enzyme dilution. The data analysis was based on the classic Michaelis-Menten equation, the latter being used to fit the experimental data and to derive the kinetic parameters KM and kcat using non-linear regression analysis implemented in the Origin package. Kinetic studies: progress-curves measured by calorimetry. A reverse M1 experiment was carried out by injecting 15 µL of a 10 nM SPU (or 3 nM JBU) solution from the syringe into the sample cell that contained a 100 mM urea solution in the presence of increasing concentrations of NBPT (0.2 - 1 mM for SPU and 0.5 - 1.5 mM for JBU, respectively). The final enzyme concentration in the sample cell was 100 pM for SPU (30 pM for JBU). The resulting thermal power (TP, µcal s-1; see a representative thermogram reported in Fig. S1 of the Supporting Information) was integrated over a 3600 s time period, starting from the time at which it reached


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its minimum. The resulting total heat (µcal) was converted to the corresponding concentration of urea consumed during the reaction by considering the measured ∆@ A for the reaction of urea hydrolysis by SPU and JBU. Kinetic studies: steady-state data by absorption spectroscopy. The experiments were carried out at room temperature by using a spectrophotometric assay and an Agilent Cary 60 UV-Vis spectrophotometer, following a previously described procedure.16,17 The pH indicator cresol red was used to monitor the overtime increase of pH due to urease activity. In the case of SPU-NBPT experiments, a solution of 50 nM SPU dissolved in 2 mM HEPES buffer at pH 8.0, containing 1 mM Na2SO3, was diluted 50 times using a 2 mM HEPES buffer solution at pH 8.0, containing 30 mg L-1 cresol red and 2 mM EDTA (Buffer CR). In the case of JBU-NBPT experiments, a solution of 65 nM JBU dissolved in 20 mM HEPES buffer, at pH 8.0, was diluted 50 times in the same Buffer CR. Then, solutions containing different concentrations of NBPT, in 2 mM HEPES buffer at pH 8.0, were added to either the SPU or the JBU urease solutions prepared as described above (final concentration of NBPT in the range 0-400 µM). The time zero of the reaction was considered as the time of the mixing of the urease and the inhibitor solutions. Subsequently, at fixed times, aliquots of these enzyme-inhibitor solutions were taken and added to a solution containing urea to a final concentration of 100 mM, and the time-dependent absorbance change at 573 nm was monitored. The enzyme activity was calculated as the slope of the linear portion of the absorbance vs. time curve. The pH variation within the observation time is small (ca. 0.2 pH units) and linear with time, suggesting that this pH change does not significantly affect enzyme catalysis, consistently with previous studies on SPU39 and JBU.58 The values of the residual activity at different time points were then calculated by normalizing each value to the activity measured at time zero. Residual percentage activities were determined


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as a function of the reaction time between enzyme and inhibitor prior to addition of substrate urea. RESULTS AND DISCUSSION This study was initiated from the determination of the high-resolution structure of SPU obtained using synchrotron X-ray diffraction data recorded on crystals grown in solutions containing the enzyme and NBPT. The analysis of the structural data then raised additional questions that prompted us to investigate further the reaction between urease and this inhibitor, using kinetic measurements. Structural analysis of SPU inhibited in the presence of NBPT. The structure of SPU cocrystallized in the presence of NBPT has been obtained here at the highest resolution ever reached for this enzyme (1.28 Å). The structure reveals the well-known quaternary structure of S. pasteurii urease, consisting of a (αβγ)3 trimer of trimers where the α subunit is formed by an (αβ)8-barrel domain and a β-type domain, the β subunit is mainly characterized by β strands, and the γ subunit consists of αβ domains. The refined crystallographic model closely matches that of native urease (PDB code 4CEU),54 with a global RMSD between their backbones equal to 0.17, 0.20 and 0.14 Å for the α, β and γ subunits, respectively. A more detailed per-residue analysis is presented in Fig. S2 of the Supplementary Information. A comparison between the backbones of the native and inhibited enzymes in terms of pairwise RMSD per residue (Fig. S2A) displays a substantial superimposition of both the β and γ subunits. However, a significant difference in the range 310-340 of residues belonging to the α subunit is evident. This region corresponds to a conserved highly flexible helix-loop-helix motif, also known as mobile flap, which controls the access of the substrate into the active site cavity of urease (Fig. 1B).4 Several crystallographic studies on native SPU, as well as other inhibited forms of the enzyme, have described the flap in


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an open conformation.4 A remarkable exception is given by the X-ray crystal structure of SPU

inhibited by DAP, a transition state analogue that is generated in situ by the enzymatic hydrolysis of phenylphosphorodiamidate (PPD).22 This model has the flap in a closed conformation, the latter being proposed to stabilize the intermediate of the catalysis. The evaluation of the pairwise RMSD per residue between the backbones of the DAP-inhibited SPU and the present structure (Fig. S2B) yields small values for the region comprised in the 310-340 residue range of the α subunit, indicating that the mobile flap in the NBPT-inhibited SPU is in the closed conformation. Figure 2. (A) Atomic model of the active site of SPU inhibited in the presence of NBPT. The nickel-coordination environment is shown superimposed to the final 2Fo–Fc electron density map


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contoured at 1.5 σ (cyan), while the unbiased Fo–Fc omit map corresponding to the ligand is shown contoured at 3.0 σ (orange). Carbon, nitrogen, oxygen and nickel atoms are grey, blue, red and green, respectively. (B) The same atomic model and nickel coordination environment are shown superimposed to the anomalous map contoured at 3.0 σ.

The electron density in the vicinity of the enzyme active site is very well defined, because of the high resolution and quality of the diffraction data (Table 1, Fig. 2A). The overall architecture of the active site, in terms of the residues that directly interact with the two Ni(II) ions, is highly conserved with respect to the native enzyme.22,54 The two Ni(II) ions are well ordered [B-factors of 11.2 and 10.2 Å2 for Ni(1) and Ni(2), respectively, with full occupancy] and are separated by 3.7 Å, a distance similar to that found in the native enzyme (3.6 Å). The two Ni(II) ions are bridged by the carboxylate group of the carbamylated αLys220*, which is bound to Ni(1) by Oθ1 (at 2.0 Å) and to Ni(2) by Oθ2 (at 2.0 Å). Ni(1) is further coordinated by αHis249 Nδ (at 2.0 Å) and by αHis275 Nε (at 2.1 Å), whereas Ni(2) is bound to αHis137 Nε (at 2.1 Å), αHis139 Nε (at 2.1 Å) and αAsp363 Oδ1(at 2.1 Å). The unbiased omit electron density map, shown in orange in Fig. 2A (calculated with Fourier coefficients Fo-Fc and phases from the refinement of inhibited SPU structure using the DAP-inhibited SPU structure as an initial model after removal of the DAP moiety and other solvent/ligand molecules), as well as the 2Fo-Fc electron density map (Fig. 3A), reveal the presence of additional electron density around the two Ni(II) ions that does not match the solvent molecules usually present in the active site of native SPU. This electron density has a tetrahedral shape and indicates the presence of a non-protein ligand that completes the coordination environment of the metal ions by binding to the two Ni(II) centers through three atoms, with a fourth atom pointing away from the bimetallic center. This tetrahedral arrangement


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exactly replaces the cluster of four water molecules existing in the active site of enzyme in its resting state. Considering the presence of NBPT in the crystallization solution, attempts to fit this entire molecule into the electron density corresponding to the omit map failed, because the latter lacks the portion corresponding to the NBPT butyl group, indicating the loss of n-butyl amine. This loss could occur following the spontaneous or the enzymatic hydrolysis of NBPT in aqueous solution. Phosphoric amides are known to undergo spontaneous hydrolysis.59,60 This type of reaction, in the case of NBPT, could results in the loss of ammonia or n-butyl amine. 1H NMR spectra on samples of NBPT stored at room temperature for four weeks ruled out the hydrolysis of n-butyl amine. Therefore, the loss of the long chain amine must be induced by SPU. The ability of ureases to hydrolyze amides and esters of phosphoric acid is well-documented in the literature.19 A crystallographic study on the inhibition of SPU by phenylphosphorodiamidate (PDB code 3UBP) demonstrated that PPD undergoes a hydrolytic event in the presence of urease to yield phenol and diamidophosphate, the latter inhibiting the enzyme by acting as a stable analogue of the tetrahedral transition state or intermediate that would form during the hydrolysis of urea.22 NBPT could thus convert to either N-(n-butyl)thiophosphoric diamide (NBPD), or to N-(nbutyl)thiophosphoric acid (NBPA), depending on the number of phosphoramide groups that are lost by spontaneous hydrolysis (Scheme 3).


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Scheme 3. Schematization of the putative hydrolytic event catalyzed by urease on NBPT derivatives. The three NBPT derivatives that can be originated by the uncatalyzed reaction occurring in solution and the products generated by the subsequent enzyme hydrolysis are represented on the left and on the right, respectively. The hydrolysis performed by urease on all these derivatives results in the release of a molecule of n-butyl amine. In turn, the structure of the moiety filling the tetrahedral omit map depends on the identity of the molecule that is actually hydrolyzed in the presence of SPU. Therefore, three independent refinement procedures were carried out, by modeling i) di-amidothiophosphoric acid (DATP), ii) mono-amidothiophosphoric acid (MATP) or iii) thiophosphoric acid (TPA, Scheme 3) into the additional electron density corresponding to the omit map, and the results were analyzed. The large scattering factor of the phosphorus and sulfur atoms, as well as the anomalous signals displayed in Fig. 2B, allowed to unequivocally locate their respective position as the central and distal atoms, pointing away from the bimetallic center. 31P NMR spectra of NBPT revealed that, in the experimental conditions used, a single peak is observed at 64.0 ppm, and this species does not evolve to NBPTO (25.0 ppm) in the course of four weeks, indicating that, in the crystallization conditions used, the P=S bond is not transformed to P=O. Moreover, assuming that, as concluded in the case of the PPD-to-DAP conversion by SPU,22 the release of n-butyl


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amine from NBPT is a consequence of a nucleophilic attack on the P atom by the Ni-bridging hydroxide, the electron density found between the Ni(II) ions must be ascribed to an O atom. These conclusions are schematically reported in Scheme 4. The identity of the remaining ligands L1 and L2 was then sought through the analysis of possible H-bonding networks. As shown in Scheme 4, L1 must have two non-bonding electron pairs available, one deputed to coordinate Ni(1) and the other to receive a hydrogen bond from αHis222 Nε (at 2.7 Å) that is protonated, as inferred by the interaction of αHis222 Nδ with the peptide NH group of αAsp224 (at 2.9 Å). In such a situation, a hypothetical P-NH2 group would not satisfy this requirement, because its only lone pair would be needed to coordinate Ni(1); on the other hand, a P-OH group bound to Ni(1) would satisfy this criterion. A similar analysis reveals the presence of two hydrogen bonding acceptor atoms in the vicinity of L2, namely the backbone carbonyl O atoms of αAla170 and αAla366 (both at 3.0 Å, Scheme 4). This implies that L2 acts as a two-hydrogen bonding donor, suggesting the presence of a P-NH2 group bound to Ni(2). This structural analysis suggests that MATP is the moiety bound in the active site of SPU, with two O atoms located at the bridging


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(LB) and L1 positions, respectively, and a P-NH2 group located at the L2 position, with the S atom located at the distal LD position, pointing towards the active site entrance. Scheme 4. Structural scheme showing the hydrogen-bonding network between the Ni(II)-bound thiophosphoramide derivative and the surrounding residues at the active site of SPU. Distances

are in Angstroms. To further support this conclusion, a statistical analysis of the B-factor values for the atoms bound to the Ni(II) ions in a number of alternative refinements was carried out, and the results are provided in Table 2. The value provided for the L1 atom is significantly smaller in the case of DATP (9.1 Å2) than in the case of MATP (10.8 Å2) or TPA (11.1 Å2), and even smaller than that of the Ni(II) atoms, a result that excludes the presence of a P-NH2 group in the L1 position. The refined B-factor value provided for the L2 atom is significantly higher in the case of TPA (13.1 Å2) than that found for MATP (10.7 Å2), the latter being very close to the B-factor of Ni(2) (10.2 Å2), a result that excludes the presence of a P-OH group at the L2 position. Altogether, these structural data are consistent with the presence of MATP as the inhibitor found in the active site of urease crystallized in the presence of NBPT. This conclusion implies that the reactive species


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interacting with and inhibiting the enzyme is the n-butyl phosphoric diamide (NBPD), generated in solution by spontaneous hydrolysis of NBPT. The final refinement procedure was therefore performed by modeling MATP in the active site of the inhibited SPU crystal structure (Fig. 3). MATP binds to Ni(1) and Ni(2) by its OL1 and NL2 atoms, respectively. The second MATP oxygen atom (OB) symmetrically bridges the two Ni(II) ions, while the sulfur atom points away from the binuclear metallo-center towards the active site cavity opening (Fig. 3A). Refined selected distances and angles are given in Table S1.

Table 2. B-factor values for the independently refined DATP, MATP and TPA at 1.28 Å Ligand

DATP

MATP

TPA

Ni(1)

11.2

11.2

11.2

Ni(2)

10.2

10.2

10.2

P

11.6

11.2

11.2

S

13.2

13.0

13.2

L1

9.1

10.8

11.1

L2

11.2

10.7

13.1

LB

11.7

11.3

11.8


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Figure 3. Atomic model of the active site of MATP-bound SPU. (A) Nickel-coordination environment shown superimposed on the final 2Fo–Fc electron density map contoured at 1.5 σ. The map of the inhibitor is shown in blue. (B) The crystallographic structure of the same environment is represented. Putative hydrogen bonds are shown as thin blue lines. Spheres are drawn using the relative atomic radii values in CrystalMaker. Carbon, nitrogen, oxygen, sulfur, phosphorous and nickel atoms are grey, blue, red, yellow, orange and green, respectively. An extended network of hydrogen bonds stabilizes the MATP molecule within the active site cavity (Fig. 3B). As previously described for the case of PPD-to-DAP conversion in the presence


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of urease,22 the Ni(1)-bound MATP oxygen atom (OL1) receives a hydrogen bond from αHis222 Nε (at 2.7 Å), while the Ni(2)-bound MATP NH2 group (NL2) acts as a donor of two hydrogen bonds to the carbonyl backbone O atoms of αAla170 (at 3.0 Å, not shown) and αAla366 (at 3.0 Å), respectively. Furthermore, the nickel-bridging MATP oxygen atom (OB) is at hydrogenbonding distance from αAsp363 Oδ2 atom (2.6 Å), implying the presence of a proton shared between the latter and the nickel-bridging MATP oxygen. Finally, the sulfur atom of MATP is at hydrogen-bond distance from the αHis323 Nε atom (3.2 Å). The P-S bond length (1.9 Å) in MATP is consistent with the presence of a double bond between the phosphorus and the sulfur atoms, thus suggesting the protonated state of αHis323 Nε. In order to establish the protonation state of the urease-MATP complex, docking calculations were performed using a previously established protocol.50 In particular, the structure of the active site of SPU in the closed flap conformation that yielded the omit map shown in Fig. 2 was used as the receptor, while four possible protonation states of MATP were tested (Fig. S3): neutral (MATPn), monoanionic (MATP1a and MATP1b, differing for the position of the negative charge), and dianionic (MATP2). The protonation state of αHis323 Nε was also probed: a first set of calculations were performed considering αHis323 protonated only on Nδ, as suggested by the vicinity with αAsp224, or on both Nδ and Nε, as suggested by the additional proximity of the MATP sulfur atom. The results of these two sets of calculations are reported in Fig. S4 and Fig. S5, respectively. A summary of the results of the docking calculations is also reported in Table S2. In all cases, with the exception of the docking of MATPn, the inclusion of a hydrogen atom on αHis323 Nε (and thus the introduction of a positive charged residue in the reaction cavity) stabilizes the ligand, as proven by the largest number of binding poses found in the most populated cluster. This result differs from the previously established neutral charge on the same


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residue in the case of DAP, a difference that can be ascribed to the presence of a S atom (in MATP) vs. an -NH2 group (in DAP).50 In all types of protonation states, the best score structures reproduce the MATP position found in the crystal structure. However, the comparison of the number of resulting structure in the most populated cluster of each tested docking suggests the presence of the monoanionic MATP in the inhibited SPU reaction site, with the negative charge located on the O atom bridging the Ni(II) ions. These results are in agreement with the same calculations performed for DAP.50 The conformational change involving the closing or the opening of the flap does not significantly affect the position of the active site residues directly involved in the binding of the Ni(II) ions. However, this event drastically changes the positions of some of the amino acid side chains that, even though not directly involved in Ni(II) binding, face the active-site cavity and are considered important for the catalytic mechanism. In particular, αCys322, αHis323 and αHis324, which belong to the mobile flap region, are approximately shifted by 5 Å with respect to the structure of native SPU. The backbone of αAla366 is also affected by the conformational change of the flap, even though it does not belong to the flap region: while in the native SPU its carbonyl O atom is turned away from the Ni(II) ions and points towards the active site entrance, in the SPU-MATP structure it is turned towards the bimetallic active site, at hydrogen-bonding distance (3.0 Å) from the MATP nitrogen atom bound to Ni(2). These structural results further corroborate the currently most accepted model for the reaction mechanism through which urease catalysis the hydrolysis of urea.3-5 In particular, the closing of the flap and the peculiar tetrahedral shape of MATP, similarly to the case of DAP, induce the formation of a strong hydrogen-bond network that blocks the flap in a closed conformation, thus disabling the protein for further substrate hydrolysis.


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In order to probe the structure of the initial complex between urease and all the inhibitors putatively present in solution, docking calculation of NBPT, NBPD, and NBPA were carried out by using SPU in the open flap conformation. All the water molecules present in the reaction site were removed and only the Ni(II) bridging hydroxide was kept in place. The protonation state of all the ligands (see Fig. S6) was probed. In particular, NBPT was considered neutral, the two enantiomers of NBPD were considered as neutral (NBPDna and NBPDnb for the S and R configuration, respectively) or monoanionic (NBPD1a and NBPD1b, deriving from NBPDna and NBPDnb, respectively), and NBPA was considered neutral (NBPAn), monoanionic (NBPA1a and NBPA1b, differing for the position of the negative charge) and dianionic (NBPA2). The protonation state of αHis323 Nε (neutral or positively charged) was also probed. The results of these calculations are reported in Fig. S7 for NBPT, Fig. S8 and S9 for NBPD, and Fig. S10 and S11 for NBPA. A summary of the results of the docking calculations is also reported in Table S3. Considering the high flexibility of the n-butyl group present in all the ligands, the clustering of the binding poses was performed considering only the P atom and the four atoms bound to it. As in the case of the docking calculations performed on SPU with the closed flap conformation, the inclusion of a hydrogen atom on αHis323 Nε stabilizes the negatively charged ligands, as proven by both the largest number of binding poses found in the most populated cluster and by an improvement of the energy scores. Interestingly, in all but two best binding poses, the ligands tend to position themselves with one phosphorus binding atom pointing toward Ni(1), the S atom making one or two H-bonds to αArg339, and the n-butyl chain pointing toward the outside of the reaction cavity. The comparison of the number of resulting structure in the most populated cluster of each tested docking suggests that the monoanionic NBPD in the S configuration (NBPD1a) is the


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molecule that best suits the SPU reaction cavity in the open conformation (Fig. 4). It can be hypothesized that, upon flap closure, the H-bonds between the S atom of NBPD and αArg339 are substituted with a H-bond with αHis323 Nε, with the whole molecule being pushed towards the Ni(II) ions and the bridging hydroxide. The nucleophilic attack of the latter on the P atom of NBPD eventually causes the release of n-butyl amine and the entrapment of MATP in the reaction site. This result is in total agreement with the crystallographic analysis of the reaction product bound to the active site of the inhibited SPU, and support the idea that NBPD, and not

NBPT, is the species acting as urease inhibitor. Figure 4. Best score binding pose for the docking simulations of NBPD1a in the open flap SPU reaction site considering αHis323 protonated on Nδ and Nε. SPU surface is colored from light blue to violet for regions nearest than 3 Å and farther than 15 Å from the Ni(II) ions, respectively. Atoms are colored accordingly to atom type. Ni(II) coordination bonds are in red and H-bonds are in blue.


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Kinetic studies on the inhibition of NBPT on urease. In order to evaluate the possible mechanisms of urease inhibition in the presence of NBPT, a series of kinetic measurements, based on calorimetric and spectrophotometric techniques, were carried out using urease from microbial (SPU) and plant (JBU) origin. The utilization of calorimetry to monitor the progress of the enzymatic reaction, in the absence and presence of the inhibitor, requires the determination of the apparent molar reaction enthalpy (∆@ A ) for the hydrolysis of urea. This value had already been measured using SPU,38 and was determined here also using JBU adopting an identical experimental set up. In particular, an experiment (M1) was carried out, in which a single addition of the substrate into a concentrated solution of the enzyme caused a decrease of the instrumental thermal power necessary to maintain the reference and the sample cell at a constant temperature, indicating an exothermic reaction. Complete consumption of the substrate occurred in ca. 1000 s (Fig. S12A). Two additional injections of substrate into the reaction cell provided curves with identical profiles, showing negligible inhibition by products in these experimental conditions. The integration of the curves yielded ∆@ A = −9.4 ± 0.3 kcal mol−1, which compares well with the value obtained for SPU (−11.3 ± 0.2 kcal mol−1).38 Another type of experiment (M2) was then performed using a diluted enzyme solution in the sample cell and carrying out multiple injections of a concentrated substrate solution. These experiments revealed an initial increase in thermal power due to the endothermic heat of substrate dilution, followed by a decrease of thermal power required to maintain isothermal conditions for the exothermic reaction (Fig. S12B). The amount of heat generated by the enzymatic reaction is equivalent to the decrease in thermal power after each injection, steadily increasing as the substrate concentration increases. Monitoring the reaction rate as a function of substrate concentration yielded a series of points that were fit to the Michaelis-Menten equation (Fig. S12C), yielding BC = 3.8 ± 0.1 mM and DEAF


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= (3.00 ± 0.18) x 104 s-1 for JBU at pH 8.0. These data are in agreement with previously reported values (2.9-3.6 mM, 5.8 x 103 s-1) obtained for JBU at pH 7.0.18 These parameters were 11.7 ± 0.2 mM and (9.90 ± 0.05) x 103 s-1 in the case of SPU, determined using the same protocol.38 The fast- or slow-binding inhibition mode of SPU and JBU in the presence of NBPT was then established following an approach that involves the use of progress curve experiments. The fast enzyme inhibition mechanism can be described as an equilibrium involving the enzyme (E) and inhibitor (I) to form the enzyme-inhibitor (EI) complex (Scheme 5), with values of the kinetic association and dissociation rates (DG and D/G ) that are of the same order of magnitude as DH and D/H , the rate constants relative to the formation of the enzyme-substrate (ES) complex. In the case of slow-binding reversible inhibition, two possibilities arise: i) mechanism A, by which the formation of the EI complex occurs through a single equilibrium regulated by smaller DG and D/G rate constants as compared to DH and D/H , and ii) mechanism B, which involves the initial formation of an EI complex through a rapid equilibrium regulated by DG and D/G , followed by a slow isomerization step, governed by DI and D/I , to form a different and more stable complex, E*I. In the case of reversible slow-binding inhibitors, the inhibition strength varies with time, and for this reason, they are also referred to as time-dependent inhibitors.61,62


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Scheme 5. The use of progress curve experiments permits the establishment of the type of inhibition among the three possibilities described above. Indeed, in the presence of a fast equilibrium inhibition process, substrate consumption (∆S increases linearly over time. By contrast, in the presence of a slow-binding inhibitor, substrate consumption exhibits a three-phase timedependent behavior: i) in the initial phase of the reaction, the consumption of substrate increases linearly with time; here, the equilibrium between enzyme and inhibitor has not yet been established, so that the slope of the curve yields the initial velocity (KL ) of the enzyme reaction; ii) in the intermediate phase, a significant deviation from linearity as a function of time emerges, at which stage enzyme and inhibitor are interacting but the system has not yet reached the steady-state; iii) in the late phase of the reaction, the increase of substrate consumption with time reverts to linearity, with a slope that provides the value of the steady-state velocity (KM ), reached after the equilibrium of the EI complex has been fully established. The obtained progress curves that describe the consumption of urea over time (∆S), determined at increasing concentrations of NBPT (Fig. 5A,B), revealed a clear deviation from


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linearity in the presence of the inhibitor regardless of the concentration used, indicating that the interaction between either JBU or SPU with NBPT involves a slow-binding mechanism. The curves were analyzed using non-linear fits of the experimental data to Eq. 1,61,62

∆S = KM N +

PQ /P7 R5S7

1 − TU-−D$VM N

Eq. 1

which provided the values of K , K and DWX (the pseudo first-order constant that governs the conversion from the initial phase to the steady-state phase) for each concentration of inhibitor (Table 3).


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Figure 5. Inhibition of ureases in the presence of NBPT determined by progress-curve experiments. (A and B) Progress-curve plots representing the consumption of urea over time at increasing concentrations of NBPT, in the case of SPU (A) and JBU (B), respectively. (C and D) Linear plots of DWX as a function of NBPT concentration in the case of SPU and JBU, respectively. In both panels, the lines represent the result of an exponential or linear fit of the data.


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Table 3. Initial and steady-state rates, and pseudo first-order constant of SPU and JBU, inhibited in the presence of NBPT, determined by calorimetry (pH 8.0, 25 °C). SPU 0.0

[NBPT] (mM) YZ (x 104 mM s-1)

Y[ (x 104 mM s-1) \]^[ (x 104 s-1)

0.2

0.4

0.6

0.8

1.0

8.42 ± 0.02

8.42 ± 0.01

8.25 ± 0.01

9.13 ± 0.01

8.49 ± 0.01

8.10 ± 0.01

-

1.91 ± 0.01

1.07 ± 0.01

0.79 ± 0.01

0.60 ± 0.01

0.51 ± 0.01

-

6.89 ± 0.02

10.3 ± 0.02

14.2 ± 0.03

18.0 ± 0.03

22.8 ± 0.04

\/_ (x 104 s-1)

1.37 ± 0.14

\_ (M-1 s-1)

21.4 ± 2.2

`∗a (µM)

6.4 ± 0.9 JBU 0.0

[NBPT] (mM) YZ (x 104 mM s-1)

Y[ (x 104 mM s-1) \]^[ (x 104 s-1)

0.5

0.75

1.0

1.25

1.5

7.97 ± 0.01

8.96 ± 0.01

9.26 ± 0.01

8.98 ± 0.01

8.63 ± 0.01

8.20 ± 0.01

-

0.572 ± 0.001

0.383 ± 0.001

0.266 ± 0.001

0.153 ± 0.001

0.105 ± 0.001

-

5.75 ± 0.01

7.61 ± 0.01

9.50 ± 0.02

11.8 ± 0.03

13.8 ± 0.04

\/_ (x 104 s-1)

0.27 ± 0.08

\_ (M-1 s-1)

28.7 ± 2.0

`∗a (µM)

0.94 ± 0.27

The linear (Eq. 2) or hyperbolic (Eq. 3) dependence of DWX as a function of inhibitor concentration could then be used to distinguish between mechanism A from mechanism B, respectively:61,62 R d

c D$VM = D/G + Hef/g

Eq. 2

h

R d

i D$VM = D/I + g3== Hef/g Q

h ed


Eq. 3

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Here, [S] is the concentration of substrate used in the experiment (100 mM), BC is the !jj

Michaelis-Menten constant, and BL

, present in the case of mechanism B only, represents the

equilibrium constant for the fast dissociation of the EI complex, and is given by the D/G /DG ratio. In both mechanisms, the value of the inhibition constant, referred to as BL∗ , is given as the equilibrium constant of the slow enzyme-inhibitor complex dissociation, and corresponds to the D/G /DG ratio (for mechanism A) or to the D/I /DI ratio (for mechanism B). The values for the initial velocities K show a large invariance with respect to the concentration of NBPT used, a behavior consistent with mechanism A. Moreover, the plot of the DWX values as a function of the concentration of NBPT (Fig. 5C,D), show that the pseudo-first order time constants linearly vary as a function of the concentration of NBPT, confirming the inhibition mechanism A. The values determined for K , K and DWX obtained from the fit of the data to Eq. 1 at each concentration of NBPT were then used to estimate the enzyme-inhibitor slow dissociation constant D/G (Scheme 5, Table 3) by using Eq. 4:61,62 P

D/G = D$VM P7

Eq. 4

Q

The value of D/G found for JBU [(0.27 ± 0.08) x 10-4 s-1, Table 3] is consistent with that previously reported using different experimental techniques [(0.15 ± 0.05) x 10-4 s-1],37 and confirms that the species that inhibits urease in the presence of NBPT, identified as MATP by Xray diffraction studies, hardly dissociates from the urease active site. Moreover, the same parameter determined here for SPU is ca. one order of magnitude larger than that measured for JBU, indicating that the dissociation event in the case of bacterial urease is more favorable than


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for plant urease. An analogous behavior has been reported for a series of phosphoramides, where the P=S group is replaced by a P=O group.20 The same study showed that the dissociation constant is invariant for all the phosphoramide inhibitors tested, within the same urease source (either SPU or JBU), supporting the idea that the actual species that dissociates from the active site of urease is diamidophosphate (DAP).20 In the present case, this species is MATP for both SPU and JBU. The rate constant DG can in turn be derived by Eq. 2, using the known values of [I], [S], BC and D/G, for SPU and JBU, respectively (Table 3). The value of k3 is the rate of the slower of the processes involving i) urease-NBPD association, ii) flap closure, and iii) inhibitor conversion to n-butyl amine and MATP, which could occur in sequence or in a concerted manner. These values appear to be largely invariant with respect to the source of urease tested, suggesting that the reaction of the inhibitor, shown by crystallography and docking calculations to be NBPD, with the active site of ureases, and the consequent enzymatic formation of MATP bound to the Ni(II) ions in the active site as established by crystallographic evidence, can be generalized among ureases from different organisms, without significant species-specific effect. The measured values of DG are similar but slightly different from the value previously reported for the inhibition of JBU in the presence of NBPT (110 ± 10 M−1 s−1).37 Probably, this apparent discrepancy can be ascribed to the different experimental setup used, and in particular to the different pH (pH 7.0 or 8.0) used for the experiments. Again, important features emerge from the comparison of the rate constants for the association between SPU and JBU with the phosphoramide inhibitors. Among the several compounds that have been tested,20 the DG values are in the same order of magnitude for bacterial and plant urease. This suggests that, for each


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type of phosphoramide, the formation of the EI complex is kinetically equivalent for ureases belonging to different organisms. Finally, the ratio D/G /DG was used to estimate the corresponding values for the urease-inhibitor equilibrium dissociation constants, BL∗ (Table 3). The value of BL∗ for SPU is ca. six times larger than that determined for JBU. Considering that DG is similar for the two ureases, this difference is mainly ascribable to the different values of D/G . In other words, the two enzymes bind the inhibitor (NBPD) with a similar affinity, but SPU is more inclined to release the inhibition product (MATP) than JBU. The results obtained by calorimetry were further validated by estimating the values of DG by following a spectrophotometric assay protocol.16,17 In this type of experiment, the enzyme is preincubated with increasing concentrations of NBPT in the absence of substrate (in this way, [S]/ BC = 0 in Eq. 2 and 3), and subsequently by initiating the enzymatic assays with the addition of urea and then measuring the rate of enzymatic reaction (K# ) after different pre-incubation times. In order to obtain the pseudo first-order rate constants (D) for the inactivation of the enzyme at each [I], the data can be analyzed by using Eq. 5:

K# = K$ TU-−DN

Eq. 5

Here, Kl is the enzyme activity, measured as an initial velocity, at time zero of pre-incubation. The values of the rate constants for the enzyme-inhibitor slow association and slow dissociation (DG and D/G for mechanism A, or DI and D/I for mechanism B, Scheme 5) can be obtained by fitting the D values determined at different [I] and by using simplified forms of Eqs. 2 and 3 (derived by setting [S]/ BC = 0) for mechanism A and B, respectively:


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D = D/G + DG I

Eq. 6

R d

i D = D/I + g3== ed Q

Eq. 7

As in the previous case, the linear (Eq. 6) or hyperbolic (Eq. 7) dependence of D as a function of [I] can be used to distinguish between mechanism A or B. The results of such experiments are shown in Fig. 6.

Figure 6. Inhibition of ureases in the presence of NBPT as determined by spectrophotometric essays. (A and B) SPU and JBU residual activities vs. time plots at different NBPT concentrations, respectively. The insets report D values as a function of NBPT concentration in the case of SPU and JBU, respectively. In both panels, the lines represent the result of an exponential or linear fit of the data.


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The values for the percentage residual urease activity as a function of pre-incubation time were optimally fitted to a single exponential decay (Eq. 5, Fig. 6). The resulting pseudo-first order constants D (s−1) showed a linear dependence on the concentration of NBPT (insets in Fig. 6), corroborating the previous analysis of the progress curve calorimetric experiments and confirming the slow-binding inhibition mechanism A. The value for the second-order kinetic constant DG resulting from the linear fit of the D vs. inhibitor concentration plot (Eq. 6) is 29.9 ± 0.4 M−1 s−1 and 24.2 ± 0.3 M−1 s−1 for the inhibition of NBPT on SPU and JBU, respectively. Due to the nature of the experiment, in which a pre-incubation of the enzyme with the inhibitor is performed prior to the measurement of the urease activity by the addition of urea as a substrate, the values of DG found with this analysis is reasonably closer to the true value of the association rate constant, with respect to the linear fit of D vs. inhibitor concentration in the progress-curve calorimetric experiments. In any case, these values are in agreement with the values of DG determined from the progress-curve calorimetric experiments by calculating it with Eq. 2, which considers the contribution of the substrate to the enzyme-inhibitor association rate. Ureases share some similarities with purple acid phosphatases (PAPs), another class of metallo-hydrolases, and in particular the presence of a dinuclear active site with two Ni(II) ions for ureases and either Zn(II)/Fe(III) or Fe(II)/Fe(III) for PAPs.63,64 In analogy with the present case, fluoride has been reported to act as slow binding inhibitor of PAPs containing a Fe(II)/Fe(III) dinuclear active site,65,66 but also as fast binding inhibitor in the case of PAPs containing a Zn(II)/Fe(III) center66 and urease.67 Considering that the inhibition mode of fluoride does not involve a chemical transformation of the inhibitor as in the present case, the slow or fast binding of fluoride with dinuclear metallo-hydrolases must be associated with structural changes within the active site. Indeed, the binding mode of fluoride to these enzymes has been


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established by crystallography both in the case of urease67 and of purple acid phosphatase68 to involve the substitution of the hydroxide ion bridging the two metal ions in the active site, which acts as the nucleophile in the enzymatic hydrolysis mechanism of both enzymes. Therefore, the fast or slow fluoride binding to this class of enzymes must be associated with the rate of exchange of the bridging hydroxide, which in turn depends on the identity and oxidation state of the metal ions present in the active site. CONCLUSIONS Structural and kinetic investigations on the inhibition of urease in the presence of N-(nbutyl)thiophosphoric triamide (NBPT) support the hypothesis that its mono-deaminated form, NBPD, is the species that interacts with the enzyme using a slow-inhibition time-dependent mechanism. All data support the hypothesis that urease selects NBPD to enter the active site, making specific interactions with conserved residues in the active site channel, and leading to a hydrolytic event that releases n-butyl amine upon closure of the mobile flap that modulates the entrance of substrate in the active site (and the exit of the reaction products). The formation of a tetrahedral MATP moiety that mimics the transition state of the reaction of urea hydrolysis stabilizes the mobile flap in a closed conformation, thus precluding the protein from further substrate hydrolysis. This study thus provides the structural basis for the efficacy of thiophosphoric amides in inhibiting urease, a role so far largely ascribed only to their oxoderivatives. The molecular details provided by this work are useful to pave the way for structurebased rational design of more efficient urease inhibitors for human and animal health as well as for improved soil nitrogen fertilization efficiency.


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ASSOCIATED CONTENT Supporting Information. The Supporting Information is available free of charge. Plots of thermal power of JBU activity; Comparison between the α, β and γ subunits in the native and inhibited SPU; additional structural parameters; and results of the docking calculations (PDF) AUTHOR INFORMATION Corresponding Author *E-mail: [email protected] Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ACKNOWLEDGMENT X-ray diffraction data were collected under the beam time award number MX-401/iNEXT 1733 from the European Molecular Biology Laboratory (EMBL, Petra III, 399. c/o DESY, Hamburg, Germany). We thank Dr. Antonio Gonzalez Vara, from the Dept. of Pharmacy and Biotechnology of the University of Bologna, for his assistance in S. pasteurii cell growth. Luca Mazzei was supported by a Ph.D. fellowship from the University of Bologna. ABBREVIATIONS SPU, Sporosarcina pasteurii urease; JBU, Canavalia ensiformis (jack bean) urease; BME, 2mercapto-ethanol; DAP, diamidophosphate; PPD, phenylphosphorodiamidate; NBPT, N-(nbutyl)thiophosphoric triamide; NBPD, N-(n-butyl)thiophosphoric diamide; NBPA, N-(n-


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butyl)thiophosphoric acid; DATP, di-amidothiophosphoric acid; MATP, monoamidothiophosphoric acid; TPA, thio-phosphoric acid (TP); NBPTO, N-(n-butyl)phosphoric triamide; RMSD, root mean square deviation. REFERENCES (1)

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Keegan, R. M.; Krissinel, E. B.; Leslie, A. G.; McCoy, A.; McNicholas, S. J.; Murshudov, G. N.; Pannu, N. S.; Potterton, E. A.; Powell, H. R.; Read, R. J.; Vagin, A.; Wilson, K. S. (2011) Overview of the CCP4 suite and current developments, Acta Crystallogr. D, 67, 235-242.


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Biochemistry

Table of Contents graphic 47x26mm (300 x 300 DPI)


Biochemistry

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Figure 1. (A) Coordination geometry of the two Ni(II) ions in native SPU active site (PDB code 4CEU54). (B) Reconstruction of the flap closure (transparent ribbons) starting from the open (light and dark blue ribbons, PDB code 4CEU) and ending in the closed (PDB code 3UBP) conformations in SPU, highlighting the side chains of αLys220*, αCys322, and αHis323. 117x161mm (299 x 299 DPI)


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Biochemistry

Figure 2. (A) Atomic model of the active site of SPU inhibited in the presence of NBPT. The nickelcoordination environment is shown superimposed to the final 2Fo–Fc electron density map contoured at 1.5 σ (cyan), while the unbiased Fo–Fc omit map corresponding to the ligand is shown contoured at 3.0 σ (orange). Carbon, nitrogen, oxygen and nickel atoms are grey, blue, red and green, respectively. (B) The same atomic model and nickel coordination environment are shown superimposed to the anomalous map contoured at 3.0 σ. 154x246mm (299 x 299 DPI)


Biochemistry

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Figure 3. Atomic model of the active site of MATP-bound SPU. (A) Nickel-coordination environment shown superimposed on the final 2Fo–Fc electron density map contoured at 1.5 σ. The map of the inhibitor is shown in blue. (B) The crystallographic structure of the same environment is represented. Putative hydrogen bonds are shown as thin blue lines. Spheres are drawn using the relative atomic radii values in CrystalMaker. Carbon, nitrogen, oxygen, sulfur, phosphorous and nickel atoms are grey, blue, red, yellow, orange and green, respectively. 367x907mm (72 x 72 DPI)


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Biochemistry

Figure 4. Best score binding pose for the docking simulations of NBPD1a in the open flap SPU reaction site considering αHis323 protonated on Nδ and Nε. SPU surface is colored from light blue to violet for regions nearest than 3 Å and farther than 15 Å from the Ni(II) ions, respectively. Atoms are colored accordingly to atom type. Ni(II) coordination bonds are in red and H-bonds are in blue. 96x90mm (299 x 299 DPI)


Biochemistry

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Figure 5. Inhibition of ureases in the presence of NBPT determined by progress-curve experiments. (A and B) Progress-curve plots representing the consumption of urea over time at increasing concentrations of NBPT, in the case of SPU (A) and JBU (B), respectively. (C and D) Linear plots of k_obs as a function of NBPT concentration in the case of SPU and JBU, respectively. In both panels, the lines represent the result of an exponential or linear fit of the data. 246x189mm (299 x 299 DPI)


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Biochemistry

Figure 6. Inhibition of ureases in the presence of NBPT as determined by spectrophotometric essays. (A and B) SPU and JBU residual activities vs. time plots at different NBPT concentrations, respectively. The insets report k values as a function of NBPT concentration in the case of SPU and JBU, respectively. In both panels, the lines represent the result of an exponential or linear fit of the data. 159x240mm (300 x 300 DPI)


Urease Inhibition in the Presence of N-(n-Butyl)thiophosphoric

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