| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
í
ovsk
1,2
1 Institute of Organic Chemistry and Biochemistry, Academy of Science of the Czech Republic, 166 10 Praha 6, Czech Republic
2 Department of Biochemistry, School of Science, Charles University, 128 43 Praha 2, Czech Republic
3 Biological Testing Branch, Science Applications International Corp. (SAIC), National Cancer Institute-Frederick Cancer Research Development Center (NCI-FCRDC), Frederick, Maryland 21784, USA
Reprint requests to: Jan Konvalinka, Institute of Organic Chemistry and Biochemistry, Flemingovo nám. 2, 166 10 Praha 6, Czech Republic; e-mail: jan.konvalinka{at}uochb.cas.cz; fax: 420-220-183-203.
(RECEIVED May 7, 2003; FINAL REVISION July 8, 2003; ACCEPTED July 8, 2003)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.03171903.
 |
Abstract |
|---|
 TOP Abstract IntroductionResultsDiscussionMaterials and methodsReferences |
|---|
Keywords: Retroviral protease; dimerization; HIV protease; fireman’s grip; kinetic assay; fluorescence; pressure
Abbreviations: HIV-1-PR(T), wild-type HIV-1-PR • HIV-1-PR(S), HIV-1-PR harboring Thr26Ser mutation • MAV-PR(S), wild-type MAV-PR • MAV-PR(T), MAV-PR harboring Ser38Thr mutation • MPMV-PR(T), wild-type MPMV-PR, 12-kD form • DABCYL, 4-[[4'-(dimethylamino)phenyl]azo]-benzoic acid • EDANS, 5-[(2'aminoethyl)-amino]naphtalenesulfonic acid • MAV, myeloblastosis-associated virus • MPMV, Mason-Pfizer monkey virus • PAL, peptide amide linker, 5-(4-Fmoc-aminomethyl-3,5-dimethoxyphenoxy)valeric acid • PheSta, phenylstatin • PR, protease • RSV, Rous sarcoma virus • TBTU, O-(benzotriazol-1-yl)-N,N,N',N'-tetramethyluronium tetrafluoroborate • Fmoc, 9-fluorenylmethoxycarbonyl • TFA, trifluoroacetic acid • PMSF, phenylmethylsulfonyl fluoride
|
Introduction |
|---|
|
TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
|---|
Because the PRs are active in the form of noncovalently bound homodimers, chemical compounds preventing dimerization became possible antiviral agents. Moreover, the dimerization seems to be the key event in the activation of the polyprotein processing in retroviruses (Kräusslich 1991). Hence, many analyses of dimerization equilibrium and rate constants have been performed, especially for HIV-1 PR. The experimental approaches can be divided into two groups: substrate-independent methods, which involve techniques such as ultracentrifugation (Holzman et al. 1991; Grant et al. 1992; Towler et al. 1998; Xie et al. 1999; St
í
ovsk et al. 2000), and substrate (inhibitor)-dependent methods, based on kinetic measurements, most often fluorometric ones (Cheng et al. 1990; Zhang et al. 1991; Jordan et al. 1992; Kuzmi
1993; Darke et al. 1994; Pargellis et al. 1994; Uhlíková et al. 1996).
The method of sedimentation equilibrium is attractive due to its independence of substrate. The result is therefore less influenced by stabilization of the dimeric form by substrate binding. On the other hand, the experiments require relatively high concentrations of pure protein. The Kd values for HIV-1 and HIV-2 proteases determined by this method are summarized in Table 1
. The majority of these values are in the micromolar range. These results, however, seem to be inconsistent with the fact that most activity assays routinely use nanomolar concentrations of HIV-PRs for activity determinations.
|
As shown in Table 1
, results of both Kd and rate constants determined kinetically with fluorogenic substrate or inhibitor have been reported by several groups for both HIV-1 and HIV-2 proteases (Cheng et al. 1990; Zhang et al. 1991; Jordan et al. 1992; Kuzmi 1993; Darke et al. 1994; Pargellis et al. 1994; Uhlíková et al. 1996. For summary, see Darke 1994). The reported Kd values from kinetic studies are typically of the order of 10-7 to 10-9 M, that is, on average, three orders of magnitude lower than the values derived from sedimentation analysis. The largest difference between two HIV-1 Kd values is five orders of magnitude. This major disagreement may be partially explained by the incorrect assumption of very fast association and dissocia tion processes in the publication by Jordan et al. (1992), in which the lowest values were determined. Another reason for the observed differences may be the inconsistency of experimental conditions in different experiments, especially pH varying from 4.5 to 7.0. However, there still remain discrepancies in the reported Kd values that are difficult to explain.
There are two major structural features believed to be responsible for the dimer stability of PRs: the N and C termini intertwined in an antiparallel ß-sheet, and a hydrogen bond scaffold supporting the active site called the fireman’s grip. It is formed by the Asp-Thr(Ser)-Gly triplet in which the oxygen of side chain of the active site Thr (Thr26 in the case of HIV-1 PR) forms a hydrogen bond with the main-chain amide of the active site Thr (Thr26') of the other molecule in the dimer. Similar interactions take place between the side chain of this Thr (Thr26) and the main-chain carbonyl of the preceding Leu (Leu24' in the case of HIV-1 PR) of the other molecule in the dimer. Both hydrogen bonds are dependent on the side-chain hydroxyls of the Thr residues. An X-ray crystallographic model of the three-dimensional structure of HIV-1 PR and a detailed view of the active site region depicting the described hydrogen bond network are shown in Figure 1. This complex structure maintains the productive conformation of the active site (Pearl and Blundell 1984; Pearl and Taylor 1987).
|
íovsk et al. 2000) provides evidence that the fireman’s grip contributes to dimerization, which could thus be responsible for these differences in activity. Stíovsk et al. (2000) showed, by using sedimentation analysis and activity assays, that mutation of Thr26 of wild-type HIV-PR to Ala and, in lesser extent, also to Ser and Cys decreases the dimer stability considerably. This finding makes it important to address whether this is a general phenomenon, that is, whether the presence of either Thr or Ser in the active site triplet Asp-Thr(Ser)-Gly indeed regulates the dimer stability of PRs. Therefore, we undertook a comparative analysis of the equilibrium and rate constants of dimerization of three prototype PRs, from HIV, MAV, and MPMV. We examined the hypothesis that the presence of either Thr or Ser in the active site triplet of PRs plays a regulatory role in the PR dimerization and therefore in its activation. We also asked whether Ser-Thr changes affect the Kd of the dimer by affecting the association or the dissociation rate constants. Answering this question may give clues to the folding pathway that leads to the active PR dimer and thus triggers polyprotein processing in retroviruses. |
Results |
|---|
|
TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
|---|
. Both substrates are suitable for wild-type MAV-PR (MAV-PR[S]). For the other enzymes, however, only peptide FS1 can be used because the cleavage of FS2 is too slow or does not occur at all.
|
 | (1) |
The equilibrium concentration of dimer in solution is given by the equation
 | (2) |
where Kd is the equilibrium constant, [D] the concentration of dimer, and E the overall (analytical) concentration of the enzyme considered as monomer. We introduce the constants 
and ß (Kuzmi 1993):
 | (3a) |
 | (3b) |
Equation 2 now has the form
 | (4) |
Thus, the equilibrium constant has to be calculated from the ratio of the initial and equilibrium dimer concentration with the aid of equation 4. Let us denote Qeq as the ratio [Deq]/[D0], where [Deq] is the equilibrium dimer concentration and [D0] is the initial dimer concentration, that is, the equilibrium concentration of the stock solution multiplied by the dilution factor. Qeq fulfills the following equation:
 | (5) |
After some rearrangements, the quadratic equation for Kd is obtained, which has two roots, one of them physically meaningful:
 | (6) |
 | (7a) |
 | (7b) |
 | (7c) |
Here E is the cuvette concentration, and E0 is the concentration of the stock solution.
Equation 6 was used for the practical determination of the dimerization equilibrium constant Kd. The parameter Qeq was calculated from the initial (tpreincubation = 0) and limit (tpreincubation 
) values of reaction rates. Final enzyme concentrations after the dilution were chosen to cover the significant range properly, that is, to reach different values of the degree of dimer dissociation from low to high. Substrate concentration was kept as low as possible, generally below the value of Km, to prevent possible substrate stabilization of the dimer.
A typical dependence of the initial rate on the preincubation time is shown in Figure 2, together with the inferred equilibrium rate. The value of Kd was calculated for every series of measurements, and the final result was then obtained as an arithmetic average of these values. These data, together with number of determinations and the initial (stock) and final concentrations, are presented in Table 3. A schematic plot of these values for all the studied enzymes is given in Figure 3. The wild-type forms of HIV-PR and MPMV-PR prefer the dimer form more than the wild type of MAV-PR, with the difference being approximately an order of magnitude. The two enzymes that can be compared to their fireman’s grip mutants indicate that the dimers of T-variants of PRs are about an order of magnitude more stable than those of the S-variants. Wild-type MPMV-PR (MPMV-PR[T]) is not compared with its mutant, but the value of Kd for this enzyme is close to that of the other T-variants.
|
|
|
 | (8) |
Using [M] = E - 2[D] ([M] is the monomer concentration), this equation can be solved yielding the time dependence of dimer concentration, [D]
 | (9) |
The symbol 
is defined as
 | (10) |
and 0 and ß0 are defined analogously to and ß in equation 3, a and b, where E is substituted by E0.
To determine the kinetic constants k1 and k2, we make use of the approximate, but generally accepted, relation among these constants and Kd:
 | (11) |
By rearranging equation 9 with the aid of equation 11, we obtain the equation
 | (12) |
where Q, the ratio of current and initial dimer concentrations, is a generalization of the experimental quantity Qeq from equation 5 and is determined as a ratio of the reaction rate at a certain time and at the initial time. The experimental quantity W depends linearly on time through the proportionality constant k2. Thus, this constant can be determined by linear regression. By using equation 11, we obtain the other rate constant k1.
The determination of the rate constant k2 makes use of equation 12. In contrast to Kd, the values of initial reaction rate for all preincubation times are necessary. For every experimental point, the quantity W is calculated and linear regression of its dependence on time is performed. The constant k2 is obtained as a slope of this dependence. The calculated values of the rate constant k2 are shown in Table 3. In addition, values of the rate constant of dissociation k1 are presented there. The last quantity presented in this table is the halftime of dimer dissociation provided that the dissociation is an independent process and no association takes place. Thus, this quantity is calculated in the following way:
 | (13) |
It is, in fact, only a different expression of k2 with a more transparent meaning.
The final averaged values of kinetic parameters are presented in Table 3. It can be seen that MPMV-PR(T) is the most quickly dissociating enzyme, whereas the other two wild-type enzymes dissociate more slowly, MAV-PR(S) being slightly faster than HIV-PR(T). Comparison of wild-type enzymes and fireman’s grip mutants of HIV-PR and MAV-PR shows that the dissociation rate constants are less affected by the mutations than the are Kd values. Thus, although HIV-PR(S) dissociates approximately twice as quickly as HIV-PR(T), there is no difference between the rate of dissociation of MAV-PR(S) and MAV-PR harboring Ser38Thr mutation (MAV-PR[T]). In contrast, the association rate constant k1, calculated from the values of Kd and k2 (see Table 3), is strongly affected by the mutations. In summary, despite the higher experimental error of k1 determination, the results show that the ThrSer mutation in fireman’s grip leads to the decrease of the rate of association rather than to the increase of the rate of dissociation.
|
Discussion |
|---|
|
TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
|---|
). Moreover, the concentration range in which neither of the two forms prevails is rather limited, approximately one order of magnitude on each side. For example, when the enzyme concentration E = 10Kd, 80% of the protein is present as dimer, whereas for E = 0.1Kd, the dimer forms only 15% of the total enzyme. For the determination of low values of Kd (10-7 to 10-9), the experimental enzyme concentration has to be kept close to these values, which consequently complicates the detection of the response, and thus, it allows only fluorescent detection in kinetic studies. Methods such as NMR or light scattering, which require high protein concentrations, are almost excluded from these studies unless the protease has a dimerization defect leading to a much higher Kd (Ishima et al. 2001; Schatz et al. 2001; Louis et al. 2003). The results presented in this article were obtained under optimal conditions of pH and ionic strength for each wild-type enzyme, and the substrate concentrations were kept below the Kd of the dimer. Two different substrates for MAV PR gave similar results. Other artifacts that might interfere with the interpretation also are unlikely. Autoprocessing, precipitation, or denaturation of the enzyme or its adsorption on the walls of preincubation tubes are excluded by those experiments performed with higher enzyme concentrations, because they run to a well-pronounced equilibrium.
In the experimental results presented, there are two principal sources of the experimental error, the error of enzyme titration and the error of determination of the ratio of equilibrium and initial rates. The error of Kd was estimated using the equation
 | (14) |
where 
Kd means the error of the equilibrium constant, E0 is the error of the initial concentration obtained from the nonlinear regression procedure, and Q is the error estimate of the equilibrium and initial rates ratio. The error of enzyme titration depends on the availability of a tight-binding inhibitor of the corresponding enzyme. In the case of MAV and MPMV enzymes, for which only less tightly binding inhibitors are available, the error is higher. Furthermore, the fluorometric detection of enzyme activity, especially the signal-to-noise ratio, is supported either by high enzyme activity or lower dimer stability, because the experiment then can be carried out with more enzyme. Accordingly, the lowest experimental error was obtained for the enzymes with high activity and low Kdvalue (such as HIV-1-PR[T]) and the highest for MAV-PR(T) and MPMV-PR(T).
As regards the rate constant determination, we decided to determine experimentally the dissociation constant k2 and then calculate k1 by using the value of of Kd. The reason is that dimer dissociation is the prevalent process immediately after the dilution of the enzyme solution, and k2 is thus less affected by the experimental error. Moreover, dimer dissociation obeys first-order kinetics, which are concentration independent. This fact precludes the influence of enzyme-titration error on k2, when only short-time measurements are taken into account. The values of k1, on the other hand, are influenced by all the errors of the both previous determinations, which makes them less reliable for enzymes in which both the measurements are troublesome, as, for example, MPMV-PR(T).
Because of the complexity of the mathematical procedure used to determine k2 and its weak dependence on the enzyme concentration, the experimental error of this constant was estimated as a standard deviation of the mean of a set of independent experimental results. The errors of the dependent quantities, 
1/2 and k1, were evaluated by using the following expressions:
 | (15a) |
 | (15b) |
The determined values of Kd (see Fig. 3) differ considerably among the studied enzymes. Physically meaningful dependencies can, however, be observed. First, the Kd of wild-type HIV-1-PR (HIV-1-PR[T]) agrees well with the other kinetically determined values of this constant in similar conditions, especially as published by Pargellis et al. (1994) and Uhlíková et al. (1996). However, the majority of the sedimentation experiments provide values that are higher by three to four orders of magnitude. We hypothesize that this disagreement is caused by the pressure dependence of the equilibrium constant, as found, for example, in the case of LexA represor (Mohana-Borges et al. 2000). High pressure in the cuvette forces proteins to decrease their volume, leading to unfolding or dissociation of multimeric structures (Silva et al. 2002). The equilibrium constant Kd depends on pressure via the standard molar Gibbs reaction energy rG0, Kd = exp(-rG0/RT), where R is the molar gas constant (R = 8.314 J mole-1K-1) and T is thermodynamic temperature. The partial derivative of rG0 with respect to the pressure is 
rG0/p = rV0, where rV0 is the molar reaction change of volume. Thus, if an approximation is made that this quantity is independent of pressure, the pressure dependence of rG0 is as follows: rG0(p) = rG0(p0) + rV0 (p-p0). Now it is possible to compare results of kinetic and sedimentation studies, and estimate the value of rV0 that might cause the observed difference. If an example of the kinetic result of the present study (Kd = 15 nM) and the sedimentation result of Stíovsk et al. (2000; Kd = 77 µM) are taken, the former gives rG0 = 45kJ/mole; the latter, rG0 = 24 kJ/mole. The pressure during ultracentrifugation is in the order of 5 MPa. Thus, the corresponding rV0 is ~-4.3 x 10-3 m3/mole, which is -7.1 x 10-27 m3 per a reaction of one dimer molecule. According to the calculation method presented by Zamyatnin (1972), the volume of the dimer is about 2.7 x 10-26 m3. Similar volume estimates can be deduced also from the Protein Data Bank crystal structures. Hence, the calculated volume change after dissociation of a single PR dimer decreases by one fourth. Although this estimate of the volume change is higher than the values measured for other proteins (Foguel et al. 1998; Mohana-Borges et al. 2000; Suarez et al. 2001), the volume change might account for the experimentally observed high Kd values, when sedimentation analysis is used for Kd determination.
The dimer of HIV-1-PR(T) is by more than an order of magnitude more stable than that of MAV-PR(S), but is comparable with MPMV-PR(T) dimer. This observation means that in the concentration range between ~10-6 and 10-9 M, the HIV-1 and MPMV PRs are considerably more active as a direct consequence of the process of dimerization. Dimerization could play an important role in regulation of activity. The concentration of the protease in the viral particle, estimated to be ~0.1 mM (Konvalinka et al. 1995), at first glance seems too high for such regulation. However, dimerization might be important for activity regulation in the cell before the formation of an immature viral particle, although the quantification of this process is difficult. It was reported that N-terminal extension of the protease may change both its activity and dimerization properties (Zybarth and Carter 1995; Schatz et al. 2001). The requirement for dimerization prevents the cleavage of cellular proteins and the premature processing of viral polyproteins by viral PR. It was shown (Burstein et al. 1991; Kräusslich 1991) that introduction of a tethered PR dimer into a retroviral provirus lead to the loss of infectivity and particle formation, clearly due to the premature activation of the polyprotein processing.
Our experiments show that the dimers of the T-version of MAV and HIV PRs are more stable than are their S-analogs, with the difference in Kd being about an order of magnitude for both enzymes. The structural reason for this phenomenon is far from clear, but the observation itself is in correspondence with the previous virological expectations. Thus, the expression strategy of a retrovirus (PR in frame with its substrate or not) correlates well with the amino acid after the active-site aspartate (serine or threonine), as well as with the Kd value. Hence, the change in this amino acid might be a direct cause of the different dimer stability, which in turn helps to regulate the replication of the virus. Our results thus strongly support the hypothesis that ThrSer mutation in fireman’s grip destabilizes the dimer of the protease and vice versa, and this mechanism is used in the nature for regulation of proteolytic activity of PRs.
The determined values of the kinetic constant k2 (and also 
) summarized in Table 3 show that the increased dimer stability of the T-version of PRs is caused by a higher rate of the dimer association rather than slower dissociation. Thus, the respective amino acid does not probably stabilize the dimer considerably but plays an important role in the mechanism of dimer association. It is conceivable that association and dissociation are not strictly reciprocal but proceed by different pathways. Provided that the dimer is kept together mainly by the interactions in the N- and C-terminal dimerization domain and the flap region (see Fig. 1A), it is not surprising that the influence of the fireman’s grip mutation to the dissociation process is small. However, these two domains are probably insufficient to induce the dimer formation on their own. We hypothesize that the fireman’s grip provides an aid to dimerization, mediating the initial contact of the two monomer molecules and adjusting them to the proper conformation and/or orientation. Several possible mechanisms for this phenomenon can be proposed. One of them can be inferred from a detail of this structure (Fig. 1B). This domain is capable of binding the two chains quite firmly by a network of hydrogen bridges and forms some predimeric intermediate. Recent structural analyses (Ishima et al. 2001; Louis et al. 2003) have demonstrated the existence of a folded monomer conformationally very similar to the subunits of a dimer. Thus, the conformation change of the monomer subunit is likely to be small and the initial intermediate may play a role in orientation of the particles to the proper positions. The fireman’s grip domain is situated in an optimal position for this purpose (as opposed to, e.g., dimerization domain at the PR termini), because it is close to the mass center of the dimer. Hence, once the subunits are connected in this region and even oriented properly, the dimerization process might be facilitated.
The remarkable difference between threonine and serine in the fireman’s grip motif also can be anticipated from Figure 1B. Methyl groups of threonines may come into a tight contact during the dimer formation. An interaction of these groups may restrict the rotation around the joint connecting the two monomers and, therefore, stabilize the nascent dimer. The conformation of the threonine side chains need not change in a major way during this process, because even in the final dimer structure, they are still close to one of the favorable rotamers, as can be deduced from the rotamer library (http://www.fccc.edu/research/labs/dunbrack/bbdep.html; see also Dunbrack Jr. and Karplus 1993; Dunbrack Jr. and Cohen 1997). If threonine is substituted by serine, the methyl groups are missing. The nascent intermediate is lacking their interaction, and its geometry may therefore be less favorable for establishing the contacts with the other domains important for dimerization.
|
Materials and methods |
|---|
|
TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
|---|
í Hejnar (Institute of Molecular Genetics, Academy of Sciences of the Czech Republic, Prague, Czech Republic), and the DNA coding for the Ser38Thr mutant of MAV-PR was kindly provided by Moshe Kotler (Department of Molecular Genetics, Hebrew University, Hadassah Medical School, Jerusalem, Israel). The DNA coding regions of MAV-PR(S) and MAV-PR(T) were PCR-amplified and cloned into the expression vector pET 22b (Novagen) by using 5'-AGAAGCTTCTCGAGCTATAAATTTGTCAAGCG-3' as a downstream primer and 5'-TAGGTACCATATGGGATCCCCGGGACATTAT-3' as the upstream primers for MAV-PR, cloned with 65 codon extension at the 5'-end, and 5'-TAGGTACCATAT GTTAGCGATGACAATGGAG-3' for the mutated MAV-PR(T) (without extension). Restriction endonucleases XhoI and NdeI were used, respectively. Coding areas of the constructs were checked by DNA sequencing. The expression and purification protocol follows the procedure of Pichová et al. (1992) with minor modifications. Bacterial cells BL21(DE3) were transformed by the expression plasmids and grown to the optical density of 1.0. The expression was induced by 1 mM IPTG (isopropyl-ß-D-galactopyranoside); after 3 h of expression cells were harvested, washed by PBS buffer (10 mM PO43-, 100 mM NaCl at pH 7.2), and resuspended in buffer A (50 mM Tris-HCl, 50 mM NaCl, 5 mM EDTA at pH 8.0) with the addition of phenylmethylsulfonyl fluoride (PMSF, 10 µg/mL). Cells were disrupted by two cycles of freezing/thawing (-70°C), 30-min incubation with chicken egg lysozyme (0.5 mg/mL), and another 30-min incubation with 1% sodium deoxycholate (0.05 mL per 1 mL of the cell suspension) followed by three 1-min sonication cycles. Suspension was centrifuged for 10 min at 1000g at 4°C; pellet was resuspended in 3 mL buffer B (50 mM Tris-HCl, 50 mM PO43-, 30 mM NaCl, 2 mM EDTA, 0.1% [v/v] mercaptoethanol at pH 7.5) and then dissolved in buffer B containing 9 M urea. The solution was dialyzed against buffer C (20 mM Tris-HCl, 10 mM PO43-, 20mM NaCl, 1 mM EDTA, 0.05% [v/v] mercaptoethanol, 5% [v/v] glycerol at pH 7.0) and centrifuged (20,000g, 4°C, 30 min). The refolding was repeated three times. Supernatants were collected and purified by batchwise anion-exchange chromatography on DEAE-Sephadex in buffer C, and the unbound material was further dialyzed into the buffer D (10 mM sodium acetate, 1 mM EDTA, 0.05% 2-mercaptoethanol at pH 5.0), loaded on the column of SP-Sepharose and eluted by NaCl gradient (0 to 0.4M) in buffer D.
HIV-1-PR(T) and HIV-1-PR harboring Thr26Ser mutation (HIV-1-PR[S]) were prepared as described in Stíovsk et al. (2000). MPMV-PR (the 12-kD form; Zábransk et al. 1998) was a kind gift of Iva Pichová (Institute of Organic Chemistry and Biochemistry, Academy of Sciences of the Czech Republic, Prague, Czech Republic).
Titration of enzyme by tight-binding inhibitor
One milliliter of assay buffer AB (0.1 M sodium acetate, x M NaCl, 1 mM EDTA, 10%[v/v] glycerol at pH 5.0; x = 2.0 for both wild-type and mutated MAV-PR[S] and [T] and 0.3 for the other enzymes) was mixed with a constant amount (typically 10 to 20 µL of 3 mM stock solution) of the peptide substrate AlaThrHisGlnValTyr (p-nitro-Phe) ValArgLysAla (5 mg/mL) and a variable amount (0 to 100 µL) of a tight binding competitive inhibitor (Majer et al. 1993). Enzyme-specific peptide inhibitors were used for this purpose: QF34, 1 µM (Konvalinka et al. 1997) for HIV-1-PR(T) and HIV-1-PR (S); ProProCysVal (PheSta) AlaMetThrMet, 13 µM for MAV-PR(S) and MAV-PR (T); and ProTyrVal (PheSta) AlaMetThr, 36 µM for MPMV-PR(T). The reaction was started by the addition of 10 µL of the enzyme and was monitored spectrophotometrically. Typically, the reaction was performed for several different values of inhibitor concentration, and the initial velocity was determined for each of them. The enzyme concentration was determined by nonlinear regression of the dependence of the initial velocity on the inhibitor concentration.
Fluorogenic substrates for kinetic studies
Two peptide substrates were designed by using an internally quenching pair of fluorescent groups EDANS (donor) and DABCYL (acceptor) as reported (Matayoshi et al. 1990). The sequences of the two substrates are as follows: FS1, (Glu[EDANS]ThrHisGln ValTyr
PheValArgLysAlaLys[DABCYL]-NH2); FS2, (Glu[EDANS] ThrProGlnValTyrPheValArgLysAlaLys [DABCYL]-NH2; the arrow denotes the cleavage site). The fluorogenic substrates were synthesized on the solid phase by using standard O-(benzotriazol-1-yl)-N,N,N',N'-tetramethyluronium tetrafluoroborate (TBTU) mediated 9-fluorenylmethoxycarbonyl (Fmoc) chemistry on polystyrene support with peptide amide linker (PAL), 5-(4-Fmoc-aminomethyl-3,5-dimethoxyphenoxy)valeric acid. Both the fluorophore (EDANS) and the quencher (DABCYL) were introduced onto the corresponding amino acids (Glu and Lys) prior to their attachment; hence, FmocGlu(EDANS)-OH and FmocLys (DABCYL)-OH were used for the coupling. The peptides were cleaved off the resin and simultaneously deprotected by 5% thioanisole, 3% ethandithiole, 2% anisole, and 90% trifluoroacetic acid (TFA) and were finally purified by high-performance liquid chromography (Vydac C18 column) using water/acetonitrile/0.1% TFA as mobile phase in gradient elution (Gulnik et al. 1997).
Kinetic characterization of fluorogenic substrates
A series of reactions was performed in which increasing amounts of the substrate (typically 3 to 15 µL of 2 mM stock solution) was mixed with 3 mL of assay buffer AB (see above), and the reaction was started by addition of 3 µL of the enzyme. The fluorescence increase was monitored for 300 sec, and the initial reaction rate was determined. The constants Km and kcat were calculated by fitting the data to the Michaelis-Menten equation by nonlinear regression.
Determination of dimerization parameters
A stock concentration of the enzyme was dialyzed into the assay buffer AB (see above) and titrated by tight-binding inhibitor. To determine the dimerization parameters Kd and k2, several series of individual reaction runs were performed, each of them for different enzyme concentration. Every experiment consisted of several reactions differing in the preincubation time. The reaction for the zero preincubation time: 3 mL of the assay buffer was mixed with known amount (1 or 2 µL) of fluorogenic substrate (~2 mM), and the reaction was started by the addition of the corresponding amount of enzyme. The course of the reaction was monitored fluorometrically for 300 sec. The reaction mixtures for the other reactions were prepared mixing 3 mL of the assay buffer with the corresponding amount of the enzyme in plastic tubes. These samples were preincubated in 37°C, each one for a different time (2 to 75 min). After preincubation, the content of the tube was transferred into the cuvette, and substrate was added in the same amount as in the first reaction; the reaction course was monitored. Initial reaction rate was determined graphically as the negative value of the slope of this dependence for treaction = 0. After completing the whole series, the limit reaction rate for tpreincubation was estimated. The dimerization parameters were derived from the set of initial reaction velocities for all these series by a mathematical procedure described in Results.
|
Acknowledgments |
|---|
Zábransk for providing MPMV protease. We are grateful to Jií Hejnar and Moshe Kotler for providing the DNA coding for the wild-type and mutant MAV protease. We also thank Pavel Majer for his kind assistance in the synthesis of fluorogenic substrates and Martin Lepík for his help in preparation of Figure 1. This work was performed under the research project Z4 055 905 and supported by grants from the Ministry of Health Care of the Czech Republic (NI/6339-3) and from the Fifth Framework of the European Commission (QLRT-2000-02360). The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.
|
References |
|---|
|
TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
|---|
Burstein, H., Bizub, D., and Skalka, A.M. 1991. Assembly and processing of avian retroviral Gag polyproteins containing linked protease dimers. J. Virol. 65: 6165–6172.
Cheng, Y.S., Yin, F.H., Foundling, S., Blomstrom, D., and Kettner, C.A. 1990. Stability and activity of human immunodeficiency virus protease: Comparison of the natural dimer with a homologous, single-chain tethered dimer. Proc. Natl. Acad. Sci. 87: 9660–9664.
Darke, P.L. 1994. Stability of dimeric retroviral proteases. Methods Enzymol. 241: 104–127.[Medline]
Darke, P.L., Jordan, S.P., Hall, D.L., Zugay, J.A., Shafer, J.A., and Kuo, L.C. 1994. Dissociation and association of the HIV-1 protease dimer subunits: Equilibria and rates. Biochemistry 33: 98–105.[CrossRef][Medline]
Dunbrack Jr., R.L. and Cohen, F.E. 1997. Bayesian statistical analysis of protein side-chain rotamer preferences. Protein Sci. 6: 1661–1681.[Abstract]
Dunbrack Jr., R.L. and Karplus, M. 1993. Backbone-dependent rotamer library for proteins: Application to side-chain prediction. J. Mol. Biol. 230: 543–574.[CrossRef][Medline]
Dunn, B.M. 1998. Human immunodeficiency virus 1 retropepsin. In Handbook of proteolytic enzymes (eds. A.J. Barret et al.), pp. 919–928. Academic Press, London.
Foguel, D., Silva, J.L., and de Prat-Gay, G. 1998. Characterization of a partially folded monomer of the DNA-binding domain of human papilloma virus E2 protein obtained at high pressure. J. Biol. Chem. 273: 9050–9057.
Grant, S.K., Deckman, I.C., Culp, J.S., Minnich, M.D., Brooks, I.S., Hensley, P., Debouck C., and Meek, T.D. 1992. Use of protein unfolding studies to determine the conformational and dimeric stabilities of HIV-1 and SIV proteases. Biochemistry 31: 9491–9501.[CrossRef][Medline]
Gulnik, S.V., Suvorov, L.I., Majer, P., Collins, J., Kane, B.P., Johnson, D.G., and Erickson, J.W. 1997. Design of sensitive fluorogenic substrates for human cathepsin D. FEBS Lett. 413: 379–384.[CrossRef][Medline]
Holzman, T.F., Kohlbrenner, W.E., Weigl, D., Rittenhouse, J., Kempf, D., and Erickson, J. 1991. Inhibitor stabilization of human immunodeficiency virus type-2 proteinase dimer formation. J. Biol. Chem. 266: 19217–19220.
Ishima, R., Ghirlando, R., Tozser, J., Gronenborn, A.M., Torchia, D.A., and Louis, J.M. 2001. Folded monomer of HIV-1 protease. J. Biol. Chem. 276: 49110–49116.
Jordan, S.P., Zugay, J., Darke, P.L., and Kuo, L.C. 1992. Activity and dimerization of human immunodeficiency virus protease as a function of solvent composition and enzyme concentration. J. Biol. Chem. 267: 20028–20032.
Konvalinka, J., Strop, P., Velek, J., Cerna, V., Kostka, V., Phylip, L.H., Richards, A.D., Dunn, B.M., and Kay, J. 1990. Sub-site preferences of the aspartic proteinase from the human immunodeficiency virus, HIV-1. FEBS Lett. 268: 35–38.[CrossRef][Medline]
Konvalinka, J., Bláha, I., 
krabana, R., Sedláek, J., Pichová, I., Kaprálek, F., Kostka, V., and trop, P. 1991. Subsite specificity of the proteinase from myeloblastosis associated virus. FEBS Lett. 282: 73–76.[CrossRef][Medline]
Konvalinka, J., Hoejí, M., Andreánsk, M., Novek, P., Pichová, I., Bláha, I., Fábry, M., Sedláek, J., Foundling, S., and trop, P. 1992. An engineered retroviral proteinase from myeloblastosis-associated virus acquires pH dependence and substrate specificity of the HIV-1 proteinase. EMBO J. 11: 1141–1144.[Medline]
Konvalinka, J., Litterst, M.A., Welker, R., Kottler, H., Rippmann, F., Heuser, A.M., and Krausslich, H.G. 1995. An active-site mutation in the human immunodeficiency virus type 1 proteinase (PR) causes reduced PR activity and loss of PR-mediated cytotoxicity without apparent effect on virus maturation and infectivity. J. Virol. 69: 7180–7186.[Abstract]
Konvalinka, J., Litera, J., Weber, J., Vondráek, J., Hradilek, M., Souek, M., Pichová, I., Majer, P., trop, P., Sedláek, J., et al. 1997. Configurations of diastereomeric hydroxyethylene isosteres strongly affect biological activities of a series of specific inhibitors of human-immunodeficiency-virus proteinase. Eur. J. Biochem. 250: 559–566.[Medline]
Kräusslich, H.G. 1991. Human immunodeficiency virus proteinase dimer as component of the viral polyprotein prevents particle assembly and viral infectivity. Proc. Natl. Acad. Sci. 88: 3213–3217.
Kuzmi, P. 1993. Kinetic assay for HIV proteinase subunit dissociation. Biochem. Biophys. Res. Commun. 191: 998–1003.[CrossRef][Medline]
Louis, J.M., Ishima, R., Nesheiwat, I., Pannell, L.K., Lynch, S.M., Torchia, D.A., and Gronenborn, A.M. 2003. Revisiting monomeric HIV-1 protease: Characterization and redesign for improved properties. J. Biol. Chem. 278: 6085–6092.
Majer, P., Urban, J., Gregorova, E., Konvalinka, J., Novek, P., Stehlikova, J., Andreansky, M., Sedlacek, J., and Strop, P. 1993. Specificity mapping of HIV-1 protease by reduced bond inhibitors. Arch. Biochem. Biophys. 304: 1–8.[CrossRef][Medline]
Matayoshi, E.D., Wang, G.T., Krafft, G.A., and Erickson, J. 1990. Novel fluorogenic substrates for assaying retroviral proteases by resonance energy transfer. Science 247: 954–958.
Mohana-Borges, R., Pacheco, A.B., Sousa, F.J., Foguel, D., Almeida, D.F., and Silva, J.L. 2000. LexA repressor forms stable dimers in solution: The role of specific DNA in tightening protein–protein interactions. J. Biol. Chem. 275: 4708–4712.
Pargellis, C.A., Morelock, M.M., Graham, E.T., Kinkade, P., Pav, S., Lubbe, K., Lamarre, D., and Anderson, P.C. 1994. Determination of kinetic rate constants for the binding of inhibitors to HIV-1 protease and for the association and dissociation of active homodimer. Biochemistry 33: 12527–12534.[CrossRef][Medline]
Pearl, L. and Blundell, T. 1984. The active site of aspartic proteinases. FEBS Lett. 174: 96–101.[CrossRef][Medline]
Pearl, L.H. and Taylor, W.R. 1987. A structural model for the retroviral proteases. Nature 329: 351–354.[CrossRef][Medline]
Pichová, I., trop, P., Sedláek, J., Kaprálek, F., Bene, V., Trávníek, M., Pavlíková, L., Souek, M., Kostka, V., and Foundling, S. 1992. Isolation, biochemical characterization and crystallization of the p15gag proteinase of myeloblastosis associated virus expressed in E. coli. Int. J. Biochem. 24: 235–242.[CrossRef][Medline]
Ridky, T.W., Bizub-Bender, D., Cameron, C.E., Weber, I.T., Wlodawer, A., Copeland, T., Skalka, A.M., and Leis, J. 1996. Programming the Rous sarcoma virus protease to cleave new substrate sequences. J. Biol. Chem. 271: 10538–10544.
Rose, J.R., Babe, L.M., and Craik, C.S. 1995. Defining the level of human immunodeficiency virus type 1 (HIV-1) protease activity required for HIV-1 particle maturation and infectivity. J. Virol. 69: 2751–2758.[Abstract]
Schatz, G.W., Reinking, J., Zippin, J., Nicholson, L.K., and Vogt, V.M. 2001. Importance of the N terminus of Rous sarcoma virus protease for structure and enzymatic function. J. Virol. 75: 4761–4770.
Silva, J.L., Oliveira, A.C., Gomes, A.M., Lima, L.M., Mohana-Borges, R., Pacheco, A.B., and Foguel, D. 2002. Pressure induces folding intermediates that are crucial for protein-DNA recognition and virus assembly. Biochim. Biophys. Acta 1595: 250–265.[CrossRef][Medline]
Stíovsk, K., Tessmer, U., Langner, J., Konvalinka, J., and Kräusslich, H.G. 2000. Systematic mutational analysis of the active-site threonine of HIV-1 proteinase: Re-thinking the "fireman’s grip" hypothesis. Protein Sci. 9: 1631–1641.[Abstract]
Suarez, M.C., Lehrer, S.S., and Silva, J.L. 2001. Local heterogeneity in the pressure denaturation of the coiled-coil tropomyosin because of subdomain folding units. Biochemistry 40: 1300–1307.[CrossRef][Medline]
Towler, E.M., Gulnik, S.V., Bhat, T.N., Xie, D., Gustschina, E., Sumpter, T.R., Robertson, N., Jones, C., Sauter, M., Mueller-Lantzsch, N., et al. 1998. Functional characterization of the protease of human endogenous retrovirus, K10: Can it complement HIV-1 protease? Biochemistry 37: 17137–17144.[CrossRef][Medline]
Uhlíková, T., Konvalinka, J., Pichová, I., Souek, M., Kraüsslich, H.G., and Vondráek, J. 1996. A modular approach to HIV-1 proteinase inhibitor design. Biochem. Biophys. Res. Commun. 222: 38–43.[CrossRef][Medline]
Wlodawer, A., Miller, M., Jaskolski, M., Sathyanarayana, B.K., Baldwin, E., Weber, I.T., Selk, L.M., Clawson, L., Schneider, J., and Kent, S.B. 1989. Conserved folding in retroviral proteases: Crystal structure of a synthetic HIV-1 protease. Science 245: 616–621.
Xie, D., Gulnik, S., Gustchina, E., Yu, B., Shao, W., Qoronfleh, W., Nathan, A., and Erickson, J.W. 1999. Drug resistance mutations can effect dimer stability of HIV-1 protease at neutral pH. Protein Sci. 8: 1702–1707.[Abstract]
Zábransk, A. 1999. "Analysis of the protease from Mason-Pfizer monkey retrovirus." PhD thesis, Institute of Organic Chemistry and Biochemistry, Prague, Czech Republic.
Zábransk, A., Andreánsky, M., Hruková-Heidingsfeldová, O., Havlíek, V., Hunter, E., Ruml, T., and Pichová, I. 1998. Three active forms of aspartic proteinase from Mason-Pfizer monkey virus. Virology 245: 250–256.[CrossRef][Medline]
Zamyatnin, A.A. 1972. Protein volume in solution. Prog. Biophys. Mol. Biol. 24: 107–123.[CrossRef][Medline]
Zhang, Z.Y., Poorman, R.A., Maggiora, L.L., Heinrikson, R.L., and Kezdy, F.J. 1991. Dissociative inhibition of dimeric enzymes: Kinetic characterization of the inhibition of HIV-1 protease by its COOH-terminal tetrapeptide. J. Biol. Chem. 266: 15591–15594.
Zybarth, G. and Carter, C. 1995. Domains upstream of the protease (PR) in human immunodeficiency virus type 1 Gag-Pol influence PR autoprocessing. J. Virol. 69: 3878–3884.[Abstract]
CiteULike 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  M. B. Clark, M. Janicke, U. Gottesbuhren, T. Kleffmann, M. Legge, E. S. Poole, and W. P. Tate Mammalian Gene PEG10 Expresses Two Reading Frames by High Efficiency 1 Frameshifting in Embryonic-associated Tissues J. Biol. Chem., December 28, 2007; 282(52): 37359 - 37369. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Sperka, P. Boross, H. Eizert, J. Tozser, and P. Bagossi Effect of mutations on the dimer stability and the pH optimum of the human foamy virus protease Protein Eng. Des. Sel., August 1, 2006; 19(8): 369 - 375. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||
