The role of the flap residue, threonine 77, in the activation and catalytic activity of pepsin A

M. Okoniewska, T. Tanaka and R.Y. Yada1

University of Guelph, Ontario Agricultural College, Department of Food Science, Guelph, Ontario, N1G 2W1, Canada


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Flexible loops, often referred to as flaps, have been shown to play a role in catalytic mechanisms of different enzymes. Flaps at the active site regions have been observed in the crystal structures of aspartic proteinases and their residues implicated in the catalytic processes. This research investigated the role of the flap residue, threonine 77, in the activation of pepsinogen and the catalytic mechanism of pepsin. Three mutants, T77S, T77V and T77G, were constructed. Differences in amino acid polarity and hydrogen bonding potential were shown to have an influence on the activation and catalytic processes. T77S activated at the same rate and had similar catalytic parameters as the wild-type pepsin. The activation rates of T77V and T77G were slower and their catalytic efficiencies lower than the wild-type. The results demonstrated that the threonine 77 polar side chain played a role in a proteolysis. The contribution of the side chain to zymogen activation was associated with the proteolytic cleavage of the prosegment. It was postulated that the hydroxyl group at position 77 provided an essential hydrogen bond that contributed to proper substrate alignment and, indirectly, to a catalytically favorable geometry of the transition state.

Keywords: aspartic proteinases/flap loop/pepsin/pepsinogen/zymogen activation


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Many enzymes have loops, floppy tails, mobile lids or moving, hinged domains which appear to contribute specifically to enzyme catalysis. Flexible loops were identified to contribute to enzyme specificities and were found to be essential for efficient catalytic processes (Kempner, 1993Go). The role of particular loop residues and their specific contribution to protein function, with the exception of glycine and proline which are known to contribute to structural flexibility, has not been clearly defined.

Porcine pepsin A (EC 3.4.23.1) belongs to a family of enzymes known as aspartic proteinases. Aspartic proteinases represent an excellent opportunity to investigate structure and function relationships of proteins owing to the availability of a large number of structural and kinetic data, different degrees of amino acid sequence homology and different specificities among the group members.

Pepsin is secreted as a zymogen called pepsinogen. The zymogen has an additional 44 amino acid peptide, referred to as the propeptide or prosegment, covalently bound to the N-terminus of pepsin. The propeptide is held in place in the pepsinogen binding cleft mostly by electrostatic interactions between its basic residues and acidic residues of the pepsin part of the molecule. The enzyme is activated when negatively charged residues become neutralized in an acidic environment which subsequently leads to proteolytic cleavage of the prosegment (Sielecki et al., 1991Go; Hartsuck et al., 1992Go). The activation process involves a conformational change of pepsinogen, proteolytic removal of the prosegment and dissociation of the cleaved prosegment or its fragments (Glick et al., 1986Go, 1991Go). During the activation process, a large conformational change occurs in the molecule which results from the removal of the prosegment and from the displacement of the 1–13 pepsin strand from the top to the bottom of the molecule. In the active enzyme, I–G–D–E–P–L strand takes the place of L–V–K–V–P–L prosegment strand in the interdomain ß-sheet which is located just behind the active site region (Sielecki et al., 1990Go, 1991Go). The sequence differences of the strands can result in different catalytic environments in pepsinogen and pepsin.

The tertiary structure of aspartic proteinases consists of two homologous lobes composed predominantly of ß-sheets. The extended substrate binding cleft and catalytic aspartic acid residues are located between the two lobes. Two antiparallel ß-strands form a flexible loop, L71–G82, which is located at the entrance to the active site and is commonly known as the flap. The flap projects over the cleft forming a channel into which substrate binds (Cooper et al., 1990Go; Sielecki et al., 1990Go). Although the active site is conformationally formed in the zymogen, the subsites adopt their final shape during the activation. The side chain of threonine 77 is closer to the active site in pepsin than in pepsinogen and it cannot assume its pepsin position until the activation peptide has been removed. In the active enzyme, the flap assumes the lowest position upon substrate binding. The greatest movement has been observed at the turn of the flap, G76–G78, which changes its position in relation to the rest of the loop (Figure 1Go). C{alpha} atoms of G76–T77–G78 change positions by 4.8 Å on average (Sielecki et al., 1991Go; Andreeva et al., 1995Go). HIV-1 protease has been shown to have two very flexible active site flaps that move by 7 Å upon substrate binding to enclose the reaction environment (Miller et al., 1989Go; Navia et al., 1989Go).



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Fig. 1. Position of the flap in pepsinogen (blue) (Sielecki et al., 1991Go), pepsin (yellow) (Sielecki et al., 1990Go) and pepsin bound to inhibitor (gray) (Chen et al., 1992Go). The threonine 77 side chain changes position from its high location in pepsinogen (cyan) to a lower position in free pepsin (orange) and the lowest position in pepsin bound to an inhibitor (red). The molecules were aligned by superimposition of catalytic aspartates depicted in pink. Inhibitor is shown as CPK model in white. The hydrogen bond of the 77 hydroxyl group to an inhibitor is represented as a green dashed line.

 
There are enzyme–substrate interactions involving residues of the flap which provide proper substrate orientation (James and Sielecki, 1985Go; Sielecki et al., 1990Go; Lin et al., 1993Go). A number of hydrogen bonds between an aspartic proteinase molecule and its substrate are formed with the flap and most of them are located at the tip of the flap, residues 76–78. In porcine pepsin, threonine 77 provides the hydrogen bonds for S1 and S2 subsites (Sielecki et al., 1990Go).

Three flap residues, tyrosine 75, glycine 76 and threonine 77, contribute directly to a particular subsite specificity by hydrogen binding to substrate residues. The presence of aspartate 77 in microbial enzymes has been used to explain their trypsin-like specificity, i.e., ability to hydrolyze bonds with lysine in the P1 position. Electrostatic interactions between a substrate lysine amine group and enzyme aspartic carboxylate have been implicated in this specificity (James and Sielecki, 1985Go). Shintani et al. (1997) engineered a pepsin loop sequence to match fungal enzymes and confirmed that the presence of aspartate 77 results in trypsin-like activity of fungal enzymes; however, this sequence was not exclusively responsible for efficient catalysis as compared with aspergillopepsin I.

In the present study, the role of flap residue threonine 77 in pepsin activation and catalytic mechanism were investigated. Three mutations of threonine 77 (serine, valine and glycine) were constructed to reveal contributions of a hydroxyl group in position 77 to enzyme activation and the catalytic process. Threonine 77 was substituted with serine, an amino acid carrying a hydroxyl group, and valine, a residue of almost the same volume as serine but without a hydroxyl group and, therefore, hydrogen bonding potential. The T77S mutation at the tip of the pepsin loop sequence would mimic that seen in gastricsin and cathepsin D (Faust et al., 1985Go; Hayano et al., 1988Go), while T77G attempted to introduce loop flexibility similar to that found in HIV-1 protease (Miller et al., 1989Go) (Table IGo). The information obtained from these studies would contribute to a better understanding of activation and catalytic mechanisms, and also to enzyme–substrate interactions in the family of aspartic proteinases.


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Table 1. Flap sequences of selected aspartic proteinases
 

    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Materials

Restriction enzymes with corresponding buffers were purchased from New England Biolabs (Mississauga, ON, Canada) and from Boehringer Mannheim Canada (Laval, QC, Canada). All the chromatography resins were purchased from Pharmacia Biotech (Uppsala, Sweden). Protein assay reagents, SDS–PAGE reagents and equipment were purchased from Bio-Rad Laboratories (Hercules, CA). Substrate I, Lys–Pro–Ala–Glu–Phe–Phe(NO2)–Ala–Leu, was synthesized in the Institute for Molecular Biology and Biotechnology, McMaster University (Hamilton, ON, Canada). Substrate II, Leu–Ser–Phe(NO2)–Nle–Ala–Leu, was purchased from Sigma Chemical (St Louis, MO). Additional materials were obtained either from Fisher Scientific (Mississauga, ON, Canada) or Sigma Chemical.

Plasmid construction and cloning

Mutations of pepsinogen were performed using the Kunkel method (Kunkel, 1985Go). Site-directed mutagenesis was performed using bacterial strains of Escherichia coli CJ236 (dut ung), JM109 (wild-type) and bacteriophage M13mp19 RF (replicative form) DNA. The primers that introduced desired mutations and new restriction enzyme sites for T77S were CTATGGCT* CCGGA*AGCATGA (BspE1), for T77V CCATCACA*TATGGCG*T*CGGTAGC (Nde1) and for T77G CCATCACA*TATGGCG*G*CGGTAGC (Nde1). Base replace-ments are indicated by asterisks and restriction enzyme sites are underlined. The mutations were confirmed on agarose gel after digestion with appropriate restriction enzymes and DNA sequencing by the automated M13-dideoxy method. The mutated pepsinogen genes were expressed as a fusion protein with thioredoxin (fusion protein, Trx-PG) in E.coli GI724 using pTFP1000 plasmid (Tanaka and Yada, 1996Go) by introduction of a short fragment containing the mutation in place of the wild-type counterpart. Fusion protein production was verified using Western blotting.

Protein expression and purification

Transformed cells were pre-cultured in RM medium [0.8% (w/v) casamino acids, 1% (v/v) glycerol, 1*M9 Salt, 0.1 mM MgCl2] at 30°C for 18 h and grown in LB medium (1% Bacto tryptone, 0.5% Bacto yeast extract, 1% NaCl, 0.150 µg/ml ampicillin, pH 7.5) at 30°C for 10 h. Cells were collected by centrifugation at 15 000 g for 10 min, resuspended in 20 mM Tris–HCl (pH 8.0) with 2 M urea and disrupted by sonication in a Model 300 Sonic Dismembrator (Fisher Scientific, Mississauga, ON, Canada). Cell lysate was collected by centrifugation at 20 000 g for 20 min. The purification procedure was performed using three ion-exchange columns. Nucleic acids were precipitated from the cell lysate with protamine sulfate (10% of protein concentration) and removed by centrifugation (20 000 g, 20 min). The supernatant was applied at a 1 ml/min flow rate on a CM-Sepharose column (22x2.5 cm i.d.) connected directly to a DEAE-Sephacel column (30x2.5 cm i.d.). The two columns were disconnected after washing with 0.10 M NaCl in 20 mM Tris–HCl, 2 M urea (pH 8.0) and the second column was subsequently washed with a stepwise gradient of NaCl (0.15, 0.20, 0.35 M) in 20 mM Tris–HCl (pH 8.0) and 2 M urea at 1 ml/min. The purest fractions of the fusion protein, as detected by SDS–PAGE, were eluted with 0.35 M NaCl, pooled and diluted three to four times to decrease the ionic strength. This preparation was applied on a DEAE-Sepharose column (25x2.5 cm i.d.) equilibrated with 50 mM phosphate buffer (pH 6.5) and washed with a stepwise gradient of NaCl (0.10, 0.20, 0.25, 0.30 M NaCl), at 1 ml/min. The Trx-PG fractions eluted with 0.30 M NaCl were subjected to further concentration on an Amicon Model 8200 concentrator using a 10 kDa cut-off refined cellulose membrane (Amicon, Beverly, MA) to a final volume of ~10 ml. The sample was then dialyzed against 20 mM Tris–HCl (pH 8.0), filtered through 0.22 µm nitrocellulose filter and stored at 4°C.

Protein concentration

Protein concentration was determined using two methods: (1) measuring the absorbance at 280 nm using a molar absorptivity of 27 500 l/mol.cm (Lin et al., 1989Go) or (2) a Bio-Rad Dc protein assay based on the Lowry method. Calibration curves were generated using bovine serum albumin.

SDS–PAGE and Western blot analysis

SDS–PAGE was performed using Bio-Rad Mini-Protean cell according to Laemmli (1970) on a 12% acrylamide gel at 30–40 mA. Proteins were stained with Coomassie Blue R-250. Western blots were run on a Bio-Rad Mini-Trans blot cell at 100 V. Proteins were transferred to a 0.2 µm PVDF membrane.

Activation of Trx-PG and purification of pepsin

Fusion protein was incubated with 0.09 M HCl for 1 h at room temperature. Activated pepsin samples were then purified from thioredoxin-activation fragments as described by Tanaka and Yada (1996) by gel filtration chromatography on Sephadex G50. Two buffers were used: 20 mM sodium acetate (pH 5.3) for kinetic assays and 10 mM sodium acetate (pH 5.3) for circular dichroism (CD) spectroscopic analysis. Active samples were identified by the milk clotting assay (McPhie, 1976Go) and only the first active fraction that eluted in the void volume was used for further pepsin analysis.

Kinetic analysis

Kinetic measurements were performed on a DU 640 spectrophotometer (Beckman Instruments, Fullerton, CA) with a temperature control unit. Two synthetic substrates were used as defined in the Materials section. For substrate I, a change in absorbance was measured at 300 nm, 37°C, in 100 mM sodium citrate buffer (pH 2.1) according to Dunn et al. (1986) and Lin et al. (1992). The substrate range was 4–150 µM. For substrate II, a change in absorbance was measured at 310 nm, 37°C, in 100 mM sodium citrate buffer (pH 3.95). The second substrate range was 10–375 µM. A minimum of six concentrations for each substrate range was used to determine initial rates. The initial slopes of progress curves were measured to give {Delta}A/min. The non-linear least-squares method was used to fit data and to calculate the kinetic constants Km and kcat (Sakoda and Hiromi, 1976Go). The results were calculated based on a minimum of two measurements for each enzyme.

Trx-PG activation kinetics and the pH dependence of activation were determined as described by Tanaka and Yada (1996, 1997). The starting fusion protein concentrations were 0.2 mg/ml for Trx-PG activation kinetics and 0.3 mg/ml for pH dependence of activation studies. Activation rate constants were calculated based on the hydrolysis of synthetic substrate I, using the Guggenheim method (Guggenheim, 1926Go). The pH dependence of activation was determined using the milk clotting assay (McPhie, 1976Go). The activity of samples was tested over 10 pH values, starting at pH 1.1 and then from pH 1.5 to 5.5 in 0.5 pH unit increments. The results were calculated based on three measurements for each enzyme.

Structural analysis

Far-UV CD spectra were generated using 800 µl of 0.1 mg/ml purified enzyme solutions with a Jasco 600 spectropolarimeter (Japan Spectroscopic, Tokyo, Japan). Pepsin samples in a 0.1 cm pathlength cuvette were scanned six times from 190 to 250 nm at room temperature with continuous nitrogen flushing. The buffer blank and enzyme solutions were degassed prior to the analyses. Secondary structures were determined using the Jasco Protein Secondary Structure Estimation Program based on the method of Chang et al. (1978).

Energy minimization and molecular dynamics were performed using Discover 2.9.5 (Biosym Technologies, San Diego, CA) on an IBM Risc System/6000 computer. The coordinates of porcine pepsin (entry name 4PEP by Sielecki et al., 1990Go) from the Protein Data Bank (Brookhaven, MA) were used as initial data for the calculations. The residues of the wild-type were replaced with corresponding mutant residues and energy minimization was performed in a consistent valence force field by the steepest descents method for 10 000 iterations with a maximum derivative of 0.001 kcal/mol. Molecular dynamics simulations were performed on the energy minimized molecule models for 10 ps (10 000 iterations) and coordinates were collected every 50 fs at a temperature of 300 K. The distances between appropriate {alpha}-carbons for all the combinations of recorded coordinates were calculated every 50 fs to give average and maximum displacements. The flexibility at each {alpha}-carbon was judged based on the comparison of maximum {alpha}-carbon displacements of each mutant and the wild-type.


    Results and discussion
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
All the mutants and the wild-type had similar elution profiles (results not shown). Proteins were purified to homogeneity as indicated by a fusion protein single band on SDS–PAGE and Western blotting of active pepsin samples (Figure 2Go).



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Fig. 2. (A) SDS–PAGE gel of purified pepsinogen fused with thioredoxin (Trx-PG): 1, the wild- type; 2, T77S; 3, T77V; 4, T77G. (B) Western blot of Trx-Pg and purified active pepsins: 1, the wild-type fusion protein (not active); 2, T77S; 3, T77V; 4, T77G; 5, commercial pepsin.

 
Kinetic parameters were measured using two substrates and are presented in Table IIGo. Generally, the same trend in kinetic constants was observed for both substrates, demonstrating that the changes were not substrate specific but could be attributed to differences in the environments of the active site. Km and kcat values were the same, within experimental error, for the wild-type and T77S, indicating that the mutation had no effect on substrate binding and overall efficiency of the catalysis. Since T77S did not show any major change in catalytic parameters as compared with the wild-type, it is proposed that any of the two amino acids, serine or threonine, could contribute efficiently to the catalysis. Hence a selection for either one does not give any evolutionary advantage to the enzyme. Consequently, the presence of serine in position 77 in gastricsin and cathepsin D (Table IGo) (Faust et al., 1985Go; Hayano et al., 1988Go) is thought not to contribute to catalytic differences in the aspartic proteinase family in any significant way.


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Table 2. Kinetic parameters for wild-type and recombinant pepsin
 
Increases in Km values were observed for T77V and T77G with a simultaneous decrease in kcat values, demonstrating that the environment of the catalytic site has changed in these two mutants. Since changes in kcat values were more pronounced than those in Km, it could be suggested that the hydroxyl group of threonine 77 contributed to stabilization of a transition state. The reaction catalyzed by pepsin was thought to take place with the formation of an oxyanion tetrahedral intermediate that was proposed to be stabilized by hydrogen bonds of residues 76 and possibly 77 (Pearl, 1987Go). Hence it is logical to speculate that the absence of a hydroxyl group in T77V and T77G mutants created a different distribution of hydrogen bonds that changed the enzyme–substrate interaction pattern and resulted in decreased catalytic rates. Furthermore, the T77S mutation which retained the hydroxyl group and, therefore, hydrogen bond potential had catalytic parameters similar to those of the wild-type. This is consistent with the argument of James and Sielecki (1985) that Pro is not favored in the pepsin P1 position since Pro is unable to act as a hydrogen bond donor owing to the cyclic attachment of its side chain to the {alpha}-imino nitrogen atom. Our results support previous observations (James and Sielecki, 1985Go; Pearl, 1987Go) that the residue at position 77 participates in hydrogen bonding to a substrate. However, it cannot be concluded that the hydroxyl group at position 77 directly stabilizes the transition state. Additionally, the crystal structures of pepsin–inhibitor complexes (Chen et al., 1992Go; Bailey et al., 1993Go; Fujinaga et al., 1995Go) demonstrate that the threonine 77 side chain is in too distant a location to interact directly with the tetrahedral intermediate. Moreover, the crystallographic studies suggest that threonine 77 interacts with a substrate backbone in S1 and S2 subsites. Based on the evidence presented above, we concluded that the interaction of threonine 77 helps to maintain proper substrate geometry during catalysis. In this indirect way, the hydrogen bond at position 77 may contribute to the most catalytically favorable position of the transition state. The supportive evidence for our argument comes from observations of Fruton (1976) and James and Sielecki (1985), who demonstrated that hydrogen bonding between the enzyme and a substrate was critical to the proper positioning of the scissile bond relative to the catalytic aspartyl residues. This proper alignment resulted from secondary interactions in the pepsin binding cleft (other than with a scissile peptide bond) and increased the rate of catalysis.

Research by Shintani et al. (1997) indicated that the introduction of a second hydrogen bond potential in position 77 (e.g. T77D pepsin mutation) resulted in an enzyme that had a lower catalytic efficiency than the wild-type pepsin and aspergillopepsin. The authors suggested that the presence of aspartate 77 created a different pattern of hydrogen bond network in pepsin's catalytic site which confirmed our observations. The apolar mutations, T77V and T77G, eliminated hydroxyl oxygen while T77D introduced an additional oxygen atom. In T77D, carboxylic oxygens had a greater electronegative charge than the hydroxyl group alone and, therefore, created a geometrically different hydrogen bond network that could lead to a distorted position of a substrate with respect to catalytic aspartates. This is in agreement with observations made previously (Fruton, 1976Go; James and Sielecki, 1985Go) that a delicate network of hydrogen bonds is responsible for substrate alignment necessary for the most productive catalytic process.

The polar group at position 77 in porcine pepsin may constitute a major force contributing to the closed loop conformation by hydrogen bonding with a substrate backbone. This interaction would fix the tip of the loop in one position and create an enclosed catalysis environment. In the absence of hydrogen bonding potential, as in the case of T77V and T77G, the tip of the loop lacks the ability to interact with a substrate in the same fashion as in the wild-type or T77S. Therefore, the catalytic environment of T77V and T77G is less compact than in the wild-type, which subsequently results in a lower catalytic efficiency of the two mutants. The hypothesis is supported by observations of Andreeva et al. (1995) where hydrogen bonds created by threonine 77 seem to have a different pattern in free and bound enzyme. It is proposed that the hydroxyl group at position 77 serves as a key force that locks the pepsin loop in closed conformation once the substrate is placed in the active site. The `locking' action is mediated by a hydroxyl group that creates a hydrogen bond(s) with a substrate in P1 and P2 positions.

The pH requirements of activation were examined by Western blotting and milk clotting activity (Figure 3Go) after a constant length of acidification. All the enzymes demonstrated maximum activation at the most acidic pH and had similar pH activation profiles. The enzymes could be activated in the pH range 1.1–4.5. No milk clotting activity or pepsin band on the Western blot were observed at either pH 5.0 or 5.5. The results showed that the mutant enzymes did not change with respect to their pH range of activity as compared with the wild-type. Thus, the solvent did not gain additional accessibility to the active site, which indicates that the position of the loop in the fusion protein was not affected by mutations. The side chain of threonine 77 in pepsinogen is in a different position to that in pepsin. The maximum change of the side chain position is induced by residues G2–D3–E4–P5–L6–E7–N8 that are inserted between a strand of the flap and the E107–S110 strand in the zymogen (Sielecki et al., 1991Go; Andreeva et al., 1995Go). Amino acids in the strand responsible for displacement, G6–N8, are susceptible to pH changes. If any structural changes occurred in the zymogen they should be reflected in the pH dependence of the activation process.



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Fig. 3. pH dependence of fusion protein activation determined by milk clotting assay. Each point represents the average of three measurements. Error bar = standard error.

 
The activation process of three mutants and the wild-type was studied at pH 1.1, 2.0 and 3.0 to investigate if the amino acid side chain in position 77 influenced the rate of zymogen activation. Tanaka and Yada (1997) investigated the activation of pepsinogen fused with thioredoxin in comparison with recombinant pepsinogen and showed that the fusion protein exhibited dominant self-activation (unimolecular reaction), in contrast to recombinant pepsinogen, which exhibited both unimolecular and bimolecular reactions. First-order rate constants were calculated for each enzyme and are presented in Table IIIGo. Consistent with previous results (Al-Janabi et al., 1972Go; Tanaka and Yada, 1997Go), the reactions were the fastest at pH 1.1, intermediate at pH 2.0 and the slowest at pH 3.0 for all the enzymes. The activation process was similar, within experimental error, for the wild-type and T77S under all pH conditions. The activation rates of T77V and T77G were slower than those for the wild-type under all three pH conditions. The two apolar mutants, T77V and T77G, had comparable activation rates that were much smaller than those for the wild-type and T77S, indicating that the polar group in position 77 participates in the activation process. As discussed above, the kinetic parameters of T77V and T77G were shown to be different from those of T77S and the wild-type pepsin. The apolar mutants displayed lower substrate specificity and catalyzed reactions at a slower rate whereas T77S was similar to the wild-type. The combined results of kinetic and activation experiments indicated that T77V and T77G mutations contributed to less efficient enzyme catalysis and slower activation processes than the wild-type at pH <=3.0. The slower activation was explained as the direct effect of the slower catalytic rate. Hence the importance of the 77 polar group in the activation process is associated with the catalytic process and the proteolytic cleavage of the prosegment.


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Table 3. First-order rate constants for activation of thioredoxin–pepsinogen fusion proteins calculated based on a hydrolysis of the synthetic substrate 1
 
The first events in the activation process involve conformational change, proteolytic cleavage and dissociation of the propeptide (Glick et al., 1986Go, 1991Go). The two mutants, T77V and T77G, have been shown to have lower catalytic efficiency which resulted in slower activation rates of both mutants as compared with the wild-type. Hence the combined results of activation studies and the kinetic measurements indicated that in very acidic conditions the proteolytic cleavage of the prosegment represents a limiting step in the conversion of fusion protein to pepsin. The findings are consistent with observations of Al-Janabi et al. (1972) and Glick et al. (1986, 1989) that the activation rates are pH dependent.

CD results are presented in Figure 4Go. All the mutants had spectra overlapping with the wild-type and their calculated secondary structure contents were virtually the same as for the wild-type (data not shown), indicating that no structural change occurred upon the mutations.



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Fig. 4. Far-UV CD spectra of the wild-type and mutant pepsins. Spectra were generated using 0.1 mg/ml purified enzyme solutions in 10 mM sodium acetate buffer (pH 5.3). Pepsin samples in a 0.1 cm pathlength cuvette were scanned six times from 190 to 250 nm at room temperature with continuous nitrogen flushing.

 
Energy minimization and molecular dynamics calculations were performed. The results indicated no major change in molecular flexibility among the wild-type and the mutants, although the carbon atom at position 294 showed less movement in T77G while the two other mutants displayed higher flexibility than the wild-type (Figure 5Go). Position 294 is located at the tip of the loop that forms an entrance to the active site cleft in the C-terminal lobe and displays large conformational variability in pepsinogen and pepsin (Sielecki et al., 1990Go; Cooper and Newman, 1991Go). Since this loop and the flap are topologically similar to the active site flap loops in HIV-1 protease, this difference is intriguing.



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Fig. 5. The difference in maximum movement of {alpha}-carbons during molecular dynamics calculations. Each line represents the difference in maximum {alpha}-carbon displacement between an individual mutant and the wild-type (Å). T77S, red line; T77V, blue line; T77G, black line.

 
Based on the above results of pH requirements of activation, CD and molecular dynamics, it was concluded that no major conformational change occurred in any of the fusion protein and pepsin mutants.

Conclusions

A polar character of the residue in position 77 is important in the proteolytic cleavage of the propeptide in the same way as in the enzyme catalytic mechanism. The hydroxyl group provides an essential hydrogen bonding potential and contributes in this way to a delicate hydrogen bond network that aligns the substrate and is indirectly responsible for a proper geometry of the transition state. The kinetic parameters are very sensitive to any changes in hydrogen bond patterns in the vicinity of the active site. A disrupted hydrogen bond network leads to less favorable substrate positioning with respect to the catalytic aspartates and, therefore, less efficient catalysis. The polar group at position 77 appears to lock the flap loop when a substrate is bound and in this way encloses the catalytic environment. The presence of serine in position 77 instead of threonine in gastricsin and cathepsin D is not thought to influence catalytic differences in the aspartic proteinase family. The lower hydrogen bond potential in the HIV-1 protease flaps may explain the need for two very flexible flaps covering the active site compared with one in eukaryotic enzymes.


    Acknowledgments
 
The authors thank Brian Bryksa for his help with various laboratory analyses. The financial support of the Natural Sciences and Engineering Research Council of Canada is gratefully acknowledged.


    Notes
 
1 To whom correspondence should be addressed. E-mail: ryada{at}uoguelph.ca Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
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Received June 10, 1998; revised September 21, 1998; accepted October 2, 1998.





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