Acceleration of Enzymatic Reaction of Trypsin through the Formation

ACS Catalysis 2015 5 (8), 4503-4513. Abstract | Full ... Langmuir 2014 30 (13), 3826-3831 ... Journal of Molecular Catalysis B: Enzymatic 2015 115, 13...
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Biomacromolecules 2005, 6, 627-631

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Communications Acceleration of Enzymatic Reaction of Trypsin through the Formation of Water-Soluble Complexes with Poly(ethylene glycol)-block-Poly(r,β-aspartic acid) Akifumi Kawamura, Yuriko Yoshioka, Atsushi Harada,* and Kenji Kono Department of Applied Materials Science, Graduate School of Engineering, Osaka Prefecture University, 1-1 Gakuen-cho, Sakai, Osaka 599-8531, Japan Received December 19, 2004; Revised Manuscript Received January 29, 2005

The amidase activity of bovine pancreas trypsin in water-soluble complexes with poly(ethylene glycol)block-poly(R,β-aspartic acid) (PEG-PAA) was evaluated by a colorimetric assay using L-lysine p-nitroanilide as a substrate. The enzymatic reaction of trypsin was accelerated through the complexation with PEG-PAA. By determining the kinetic parameters of the enzymatic reaction of trypsin, it was confirmed that the catalytic rate constant of the complexed trypsin was 15 times higher than that of the native trypsin. From the evaluation of pH dependence of initial reaction rate, it was indicated that this acceleration was induced by a stabilization of the imidazolium ion of the His residue in the catalytic site, the Asp-His-Ser triad, of trypsin due to the Asp units of PEG-PAA. The hydrogen bonded Asp-His pairs are critical constituents in several key enzymatic reactions including serine protease and apurinic endonucleases, and it was expected that the acceleration of the catalytic reaction might occur for other enzymes by the formation of water-soluble complexes with PEG-PAA. Introduction The complexes of natural compounds including DNA and enzymes with synthetic polymers through intermolecular interactions are interesting research topics from both fundamental and applied standpoints. Recently, the focus on the complexes of natural compounds with block ionomers in aqueous medium has been increasing especially because of their potential utilities as functional materials including carriers for biologically active compounds in a drug delivery system.1 The mixtures of block ionomers with oppositely charged synthetic and natural polymers at optimized mixing ratios formed polyion complex micelles, which have a waterinsoluble polyion complex core surrounded by a polymer brush corona layer. The complexes of enzymes with block ionomers was first reported using egg white lysozyme as the enzyme and poly(ethylene glycol)-block-poly(R,β-aspartic acid) (PEG-PAA).2 Their mixtures spontaneously formed polyion complex micelles, in which the polyion complex core of the lysozyme and PAA blocks was surrounded by a PEG brush layer. The nanoscopic core of this system worked as a unique reaction field for the lysozyme, and the enzymatic reaction of the * To whom correspondence should be addressed. Tel & Fax: +81-72254-9328. E-mail: [email protected].

lysozyme was remarkably enhanced through the change in the binding specificity between the lysozyme and substrate.3 Also, when bovine pancreas trypsin was incorporated into the core by the complexation with PEG-PAA, the enzymatic reaction of trypsin was also enhanced in the core of the PIC micelles, although its mechanism was still unclear.4 We tried to evaluate the mechanism of the enhanced enzymatic reaction of trypsin in the micelles, and it was found that the enhancement of the enzymatic reaction of trypsin was induced not according to the effect of the incorporation into the core of the micelles but according to the effect of the complexation with the block ionomers, PEGPAA. Also, it was confirmed that this acceleration of the amidase activity of trypsin was not due to the apparent change in the Michaelis constant, at which a concentration effect was often reported on the enzyme-polymer conjugates, but due to a change in the catalytic rate constant. Such a phenomenon might be observed due to the formation of water-soluble complexes of trypsin and block ionomers, which showed a higher dispersibility compared with conventional polyion complexes formed from trypsin and anionic polyelectrolytes. It is expected by forming water-soluble complexes of other kinds of enzymes with block ionomers that interesting phenomena, which have not been discovered until now, will be observed.

10.1021/bm049198w CCC: $30.25 © 2005 American Chemical Society Published on Web 02/15/2005

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Table 1. Physicochemical Properties of Used Polymers in This Study

EA15 EA24 EA37 EA68 PAA

Mna

Mw/Mnb

13700 14800 16300 19800 6600

1.06 1.12 1.12 1.05 1.25

Mn of PEGb

Mw/Mn of PEGb

12000 12000 12000 12000

1.02 1.02 1.02 1.02

a These values were calculated from the peak area ratio between PEG and PAA block in 1H NMR spectra. b These values were determined from GPC based on a standard curve of PEG.

Experimental Section Materials. Four types of poly(ethylene glycol)-blockpoly(R,β-aspartic acid)s (PEG-PAA; PEG Mn was fixed to 12 000) having different polymerization degrees of the PAA block (15, 24, 37, and 68; code names were EA15, EA24, EA37, and EA68, respectively), poly(R,β-aspartic acid) homopolymer (PAA; polymerization degree was 57) were synthesized as already described2a,5 (the physicochemical properties of used polymers were summarized in Table 1). Bovine pancreas trypsin and L-lysine p-nitroanilide were purchased from Sigma (St. Louis, MO), and used without further purification. Preparation of Trypsin/PEG-PAA Complexes. A total of 8 mg/mL of trypsin and varying concentration of PEGPAA in sodium phosphate buffer (10 mM, pH 7.4) were separately prepared as stock solutions, and then stored under a cooled condition (4 °C) to prevent the autolysis of the trypsin. Both solutions with same volume were mixed so as to be the fixed trypsin concentration (4 mg/mL) and varying PEG-PAA concentration, in which the mixing ratio of trypsin and PEG-PAA was defined as the molar ratio of the number of Asp residues of PEG-PAA versus the total number of Arg and Lys residues of trypsin, and the mixtures were stored at 4 °C. After 30 min of mixing at 4 °C, the mixtures were used in further evaluations. Characterization of Trypsin/PEG-PAA Complexes. Dynamic and static light scattering measurements were carried out using a DLS-700 spectrometer (Otsuka Electronics Co., Ltd., Japan) equipped with an Ar ion laser (λ ) 488 nm) at 25 °C. The detection angle for the dynamic light scattering was fixed at 90°. The static light scattering measurements were performed at five detection angles (30°, 60°, 90°, 120°, and 150°), and the scattering intensity at a zero detection angle was then extrapolated. Evaluation of Amidase Activity. The amidase activity of trypsin was evaluated using L-lysine p-nitroanilide as a substrate at 25 οC.6 A total of 100 µL of the complex solutions including 4 mg/mL of trypsin was added to 900 µL of the substrate solution including 5.6 mM of L-lysine p-nitroanilide. The reaction rates of the native trypsin and the complexed trypsin with PEG-PAA were determined by monitoring the change in absorbance at 410 nm, at which the wavelength for the extinction coefficient of p-nitroanilide is 8800 cm-1 M-1, after mixing of the complex solution and substrate solution. The initial reaction rate was determined from the slope of the change in absorbance at 410 nm from 100 to 200 s after mixing sample solution and substrate

Figure 1. Change in the relative enzymatic activity of trypsin with the mixing ratio for the trypsin/PEG-PAA complexes. (b, EA15; 2, EA24; 9, EA37; [, EA68; the concentrations of trypsin and substrate were fixed to 16.8 µM and 5 mM, respectively.)

solution. The relative activity was defined as the ratio of initial reaction rate of the samples against that of native trypsin. Also, the trypsin/PAA complex solution was turbid, and it was not able to treat the absorbance at 410 nm quantitatively. Results and Discussion The complexes of trypsin and PEG-PAA were prepared at varying mixing ratios. All of the prepared mixing solutions of trypsin and PEG-PAA remained transparent solutions, although the mixture of trypsin and the PAA homopolymer became turbid and underwent precipitation. The amidase activity of trypsin in the water-soluble complexes with PEGPAA was evaluated by a colorimetric assay using L-lysine p-nitroanilide as a substrate. Figure 1 shows the change in the relative activity of trypsin complexed with PEG-PAA having various PAA blocks with the mixing ratio, where the relative activity was the ratio of the initial reaction rate of the complexed trypsin versus that of the native trypsin. The complexation with all of the PEG-PAAs induced an acceleration of the enzymatic reaction of trypsin, although the acceleration degree depended on the mixing ratio and the PAA block length of the PEG-PAA. Also, such acceleration of the enzymatic reaction of trypsin was not observed from the mixture of trypsin and PEG homopolymer. Obviously, the acceleration degree increased with an increase in the PAA block length. The relative activities of the complexed trypsin with EA15, EA24, EA37, and EA68 were 2, 12, 14, and 24 at the mixing ratios showing the highest relative activity, respectively. Also, the mixing ratios showing the highest relative activity depended on the PAA block length, which were 15, 10, 10, and 5 for EA15, EA24, EA37, and EA68, respectively. This suggests that PEG-PAA, having the longer PAA block length, effectively accelerated the enzymatic reaction of trypsin through complexation. From the dependence of the highest relative activity and its mixing ratio on the PAA block length, it was expected that the state of the trypsin/PEG-PAA complexes might change with the varying PAA block length in PEG-PAA. To characterize the trypsin/PEG-PAA complexes, dynamic and static light scattering measurements were carried out for the complexes prepared at the varying mixing ratios. Figure 2 shows the change in the relative scattering intensity, which was normalized by the scattering intensity of the native trypsin solution as the control, with the mixing ratio for the

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Table 2. Enzymatic Activity of the Native Trypsin and the Complexed Trypsin with PEG-PAA (EA68) mixing ratio 0 2 5 7.5 10

kcat [min-1]a

Km [M]a 10-3

1.20 ( 0.01 × 1.49 ( 0.04 × 10-3 3.58 ( 0.08 × 10-3 1.41 ( 0.04 × 10-3 9.43 ( 0.07 × 10-4

(1.0)b (1.2)b (3.0)b (1.2)b (0.8)b

0.560 ( 0.003 3.31 ( 0.09 8.47 ( 0.19 3.78 ( 0.11 2.96 ( 0.02

kcat/Km [M-1 min-1] (1.0)b (5.9)b (15.1)b (6.8)b (5.3)b

467 2220 2370 2680 3130

(1.0)b (4.8)b (5.1)b (5.7)b (6.7)b

a These values were determined from the Lineweaver-Burk plots and the experimental error was calculated from the correlation coefficients of the Lineweaver-Burk plots. b These values were relative values versus the values at zero of the mixing ratio (native trypsin).

Figure 3. Lineweaver-Burk plots of the native trypsin (2) and the complexed trypsin with PEG-PAA (b). (The complexes were prepared at 5 of the mixing ratio by using EA68.) Figure 2. Change in the relative scattering intensity of the trypsin/ PEG-PAA complexes with the mixing ratio. (b, EA15; 2, EA24; 9, EA37; [, EA68)

trypsin/PEG-PAA complexes. All of the trypsin/PEG-PAA complexes exhibited the maximum relative scattering intensity. Incidentally, when the mixture of trypsin and PEG-PAA did not form the complexes, i.e., no interaction between trypsin and PEG-PAA, the relative scattering intensity did not increase with an increase in the mixing ratio and remained almost unity. The trends in the change of the relative scattering intensity with the mixing ratio were similar to the trends in the change of the relative activity shown in Figure 1, although the mixing ratios showing the highest relative scattering intensity were shifted to a smaller mixing ratio compared with those showing the highest relative enzymatic activity. The mixing ratio showing the highest relative scattering intensity was the optimal mixing ratio for the preparation of the PIC micelles, and the PIC micelles being multimolecular assemblies were detected at the mixing ratios showing the highest relative scattering intensity by dynamic light scattering measurements. The average diameters of the PIC micelles prepared from trypsin and PEGPAA (EA24, EA37, and EA68) were 110, 90, and 70 nm,4 respectively. However, there was no detection of the PIC micelles having an average diameter of several tens of nanometers for the trypsin/EA15 complexes due to the low scattering intensity even at the mixing ratio showing the maximum relative scattering intensity. It should be noted here that the mixing ratio, which shows the maximal relative activity, is higher than the mixing ratio showing the maximum relative scattering intensity. For example, in the case of the trypsin/EA68 complexes, the mixing ratios showing the maximum relative activity and relative scattering intensity were 5 and 0.75, respectively. This means that the optimal mixing ratios were different between the enzymatic function and the micellization. In other words, for the acceleration of enzymatic reaction of trypsin found in this

study, it was not important to take the micellar structure, i.e., core-shell structure, but it is important to form the water-soluble complexes. Rather than the effect of a reaction field, it is postulated that it depended on the effect on a rather smaller size scale, e.g., the specific binding site and catalytic site. To understand the reason for the acceleration, the Michaelis constant (Km), the catalytic rate constant (kcat), and kcat/Km were determined from the Lineweaver-Burk plot for the complexes prepared from trypsin and EA68 at various mixing ratios. Figure 3 shows the Lineweaver-Burk plots of the native trypsin and the complexed trypsin with EA68 at 5 of the mixing ratio as a typical example of the complexed trypsin. Both the Lineweaver-Burk plots for the native and complexed trypsin exhibited good linearities (the correlation coefficients were greater than 0.95), indicating that the enzymatic reaction of trypsin can be kinetically analyzed based on the Michaelis-Menten equation. The determined Km, kcat, and kcat/Km values are summarized in Table 2. At 5 of the mixing ratio, at which the enzymatic reaction of trypsin was the most accelerated in Figure 1, both the Km and kcat values were maximized. It was considered that the factor with the reverse influence on the Km and kcat values makes the Km and kcat values have maximum values by a change in the mixing ratio. The change in the Km value, which mainly reflects the change in the binding between a substrate and an enzyme, could be explained by the steric hindrance in the interaction between a substrate and trypsin in the complexes from the result of the relative scattering intensity shown in Figure 2. The scattering intensity is sensitive to change in the molecular weight of a solute, and the change in the relative scattering intensity shown in Figure 2 reflects the change in the association number of trypsin and PEGPAA in the complex. For a higher association number, the interaction between trypsin and substrate might be negatively influenced, although it was difficult to clearly show any correlation between the association number and steric

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hindrance. By increasing the mixing ratio to 10, the Km value of the complexed trypsin was close to that of the native trypsin, suggesting that the association number of trypsin might be close to unity. Similar changes in the scattering intensity and the association number were observed for the complexes of plasmid DNA (pDNA) and PEG-block-poly(L-lysine) (PEG-PLL).7 In this case, the formation of pDNA/ PEG-PLL complexes, which entrapped a single molecule of pDNA, at the mixing ratio including excess PEG-PLL versus stoichiometric mixing ratio was indicated. That is, the pDNA/ PEG-PLL complexes with excess PEG-PLL were formed in a noncooperative manner. Trypsin/PEG-PAA complexes at excess PEG-PAA might also be formed in a noncooperative manner and decrease the association number of trypsin with an increase in the mixing ratio. It is difficult to explain the change in the kcat value only from the results of the light scattering measurements shown in Figure 2. To explain the change in the kcat value, it needs to take the catalytic mechanism of trypsin into consideration. It is well-known that the triad of residues, Asp, His, and Ser, plays an important role in a catalytic reaction of serine proteases such as trypsin, chymotrypsin and elastase.8 In the catalytic site of trypsin, the nucleophilicity of the hydroxyl group of Ser195 increases through a proton-transfer process in the Asp-His dyad, in which the carboxylate group of Asp102 stabilizes the imidazolium ion of His57.9 The importance of the Asp residue in the catalytic triad was confirmed by replacing Asp102 of trypsin with the Asn residue, and the kcat value of the mutant trypsin was ∼104 times lower than that of the native trypsin at a neutral pH condition.10 Also, the enzymatic activity of trypsin strongly depended on the pH, because His and Asp residues are weak cationic and anionic amino acid residues and have an equilibrium between the protonated and nonprotonated states. For the trypsin/PEG-PAA complexes, the Asp units of the complexed PEG-PAA might locate near the catalytic site of trypsin, i.e., the Asp-His-Ser triad, and a part of the Asp units might be able to support the stabilization of the imidazolium ion of the His residue. The kcat value increased with an increase in the mixing ratio in the range of less than 5, since the probability of Asp units, which participate in stabilization, increased. However, the complexed PEG-PAA might also have a negative effect on the catalytic reaction of trypsin, and might induce a decrease in the local pH of the complexes. A decrease in the pH from neutral pH shifts the equilibrium of the His and Asp residues to the protonated sides, and the enzymatic activity of trypsin then decreases. Consequently, the kcat value decreased at a greater than 5 of the mixing ratio. Also, although the kcat/Km value, which expresses the enzymatic activity of the entire processes, continued gradually increasing with an increase in the mixing ratio, this increase was a result of the balance among the effect of the stabilization of the imidazolium ion of the His residue and a decrease in the local pH of the Asp units of the complexed PEG-PAA and the effect of steric hindrance of the catalytic sites. Further, to clarify the interaction of PEG-PAA with the catalytic site of trypsin, pH effect on initial reaction rate was compared between native and complexed trypsin. Figure 4

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Figure 4. Plots of initial reaction rate of the native trypsin (2) and the complexed trypsin with PEG-PAA (b) against pH. (The complexes were prepared at 5 of the mixing ratio by using EA68.)

shows the influence of pH to initial reaction rate of native and complexed trypsin. As mentioned above, the enzymatic activity of trypsin strongly depended on the pH. Obviously, the amidase activity of both native and complexed trypsin depended on pH, and initial reaction rate decrease with a decrease in pH. However, the pH profile of initial reaction rate of the complexed trypsin shift to lower pH region, indicating that PEG-PAA interacted with the catalytic site of tyrpsin, the Asp-His-Ser triad. There were two possibilities of the shift of the pH profile, essential and apparent change. The essential change was the stabilization of the imidazolium ion of His in the catalytic site through the interaction of PEGPAA. The apparent change was the local environmental change around the catalytic site, i.e., the local pH change. If the local pH change might be induced through the complexation with PEG-PAA, the initial reaction rate might decrease with a decrease in pH. Consequently, the shift of the pH profile shown in Figure 4 was due to essential change and indicated the interaction of PEG-PAA with the catalytic site. When the acceleration of the enzymatic reaction of trypsin was induced by such a site specific effect, it was thought that the acceleration was also observed in the trypsin/PAA complexes, but in trypsin/PAA complexes, it was difficult to monitor the enzymatic reaction of trypsin due to the turbidity of the trypsin/ PAA complexes as mentioned above. By the formation of the water-soluble complexes from block ionomer and enzyme with a high dispersibility, it became possible to determine the enzyme reaction constants, Km and kcat, and to consider the acceleration mechanism. It is expected that the use of block copolymers, i.e., block copolymerization, serves as the one technique of the analysis of a structure as well as a function about a complex with biomacromolecules, such as enzymes and DNA. Conclusions The amidase activity of trypsin in the water-soluble complexes formed with PEG-PAA was evaluated using L-lysine p-nitroanilide as a substrate, and it was confirmed that the enzymatic reaction of trypsin was accelerated through the complexation with PEG-PAA. The degree of the acceleration was dependent on the mixing ratio of trypsin and PEG-PAA as well as the composition of PEG-PAA. To determine the mechanism of the acceleration, the kinetic parameters of the enzymatic reaction of trypsin were determined from the Lineweaver-Burk plot. Interestingly,

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the enzymatic reaction of trypsin in the complexes was enhanced by not the apparent change in the Km values, i.e., the concentration effect, but by the increase in the kcat value. That is, the enzymatic function of trypsin was essentially enhanced through the complexation with PEG-PAA. Also, the increase in the kcat value might be induced by the stabilization of the imidazolium ion of the His residue in the catalytic Asp-His-Ser triad. The Asp-His-Ser triad is a common catalytic site of serine protease including not only trypsin but also chymotrypsin and elastase, and the Asp-His dyad plays an important role in other kinds of enzymes such as apurinic endonuclease 1, which is a human DNA repair enzyme that cleaves adjacent to the abasic sites in DNA.12 The results obtained in this study suggest that PEG-PAA might have the potential ability to accelerate the enzymatic function of not only trypisn but also various kinds of enzymes through the stabilization of the Asp-His dyad in the catalytic sites by the formation of water-soluble complexes. Acknowledgment. This study was supported by the Industrial Technology Research Grant Program from New Energy and Industrial Technology Development Organization (NEDO) of Japan.

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References and Notes (1) (a) Kabanov, A. V.; Kabanov, V. A. AdV. Drug DeliVery ReV. 1998, 30, 49. (b) Kataoka, K.; Harada, A.; Nagasaki, Y. AdV. Drug DeliVery ReV. 2001, 47, 113. (c) Co¨lfen, H. Macromol. Rapid Commun. 2001, 22, 219. (2) (a) Harada, A.; Kataoka, K. Macromolecules 1998, 31, 288. (b) Harada, A.; Kataoka, K. Langmuir 1999, 15, 4208. (3) Harada, A.; Kataoka, K. J. Am. Chem. Soc. 2003, 125, 15301. (4) Jaturanpinyo, M.; Harada, A.; Yuan, X.; Kataoka, K. Bioconjugate Chem. 2004, 15, 344. (5) Nishiyama, N.; Kataoka, K. J. Controlled Release 2001, 74, 83. (6) Erlanger, B. F.; Kokowsky, N.; Cohen, W. Arch. Biochem. Biophysics 1961, 96, 271. (7) Itaka, K.; Yamauchi, K.; Harada, A.; Nakamura, K.; Kawaguchi, H.; Kataoka, K. Biomaterials 2003, 24, 4495. (8) (a) Blow, D., H.; Birktoft, J. J.; Hartley, B. S. Nature 1969, 221, 337. (b) Hunkapiller, M. W.; Smallcombe, S. H.; Whitaker, D. R.; Richards, J. H. J. Biol. Chem. 1973, 248, 8306. (c) Robillard, G.; Shulman, R. G. J. Mol. Biol. 1974, 86, 519. (d) Scheiner, S.; Lipscomb, W. N. Proc. Nat. Acad. Sci. U.S.A. 1976, 73, 432. (e) Markley, J. L.; Ibanez, I. B. Biochemistry 1978, 17, 4627. (9) (a) Kossiakoff, A. A.; Spencer, S. A. Biochemistry 1981, 20, 6462. (b) Wellner, N.; Zundel, G. J. Mol. Struct. 1994, 317, 249. (10) Craik, C. S.; Roczniak, S.; Largman, C.; Rutter, W. J. Science 1987, 237, 909. (11) Lowry, D. F.; Hoyt, D. W.; Khazi, F. A.; Bagu, J.; Lindsey, A. G.; Wilson D. M., III. J. Mol. Biol. 2003, 329, 311.

BM049198W