Expanded structural and stereospecificity in peptide synthesis with chemically modified mutants of subtilisin

Article abstract of DOI:10.1016/S0957-4166(99)00255-4

Employing the strategy of combined site directed mutagenesis and chemical modification, we previously generated chemically modified mutant enzymes (CMMs) of subtilisin Bacillus lentus (SBL). We now report the use of these SBL-CMMs for peptide coupling rea

Full text of DOI:10.1016/S0957-4166(99)00255-4

Pergamon
Tetrahedron: Asymmetry 10 (1999) 2563–2572
Expanded structural and stereospecificity in peptide synthesis
with chemically modified mutants of subtilisin
Kanjai Khumtaveeporn, Grace DeSantis and J. Bryan Jones ∗
Department of Chemistry, University of Toronto, 80 St George Street, Toronto, Ontario M5S 3H6, Canada
Received 7 May 1999; accepted 10 June 1999
Abstract
Employing the strategy of combined site directed mutagenesis and chemical modification, we previously
generated chemically modified mutant enzymes (CMMs) of subtilisin Bacillus lentus (SBL). We now report the
use of these SBL-CMMs for peptide coupling reactions. The SBL-CMMs exhibit dramatically altered substrate
specificity, including the acceptance of D-amino acid acyl donors, generating dipeptides containing D-Phe, D-Ala
and D-Glu in up to 66% yield, which was not possible using wild-type SBL (WT-SBL)0. In addition, SBL-CMMs
accommodate α-branched amino acids such as L-Ala-NH2 as acyl acceptors in their S1 pockets, which WT-SBL
will not. © 1999 Elsevier Science Ltd. All rights reserved.
1. Introduction
Enzymatic, protease-catalyzed peptide coupling offers an attractive alternative to chemical solution or
solid phase peptide synthesis.1 Since enzymes are operational under mild reaction conditions and offer
high regio- and stereoselectivity, the need for extensive protection strategies is minimized.2 The serine
proteases, which are a well-characterized group of hydrolytic enzymes, have demonstrated particular
potential in peptide ligation.1
Serine protease-catalyzed peptide coupling requires the formation of a covalent acyl enzyme inter-
mediate between the acyl donor substrate (RCOOR0) and the hydroxyl group of the catalytic serine
residue, forming an acyl enzyme intermediate (RCOO-Enz) as outlined in Scheme 1. The acyl enzyme
intermediate is then aminolyzed by the amino terminus of the acyl acceptor (R00NH2), thereby effecting
the formation of a peptide bond. However, in order to assure the success of this strategy the competing
hydrolysis of the acyl enzyme intermediate, and also hydrolysis of the resulting peptide product (i.e.
the enzyme’s amidase activity), must be minimized. Furthermore, it is desirable to expand the substrate
specificity of proteases in order to enhance their applicability to unnatural substrate analogues.
∗
Corresponding author. E-mail: jbjones@alchemy.chem.utoronto.ca
0957-4166/99/$ - see front matter © 1999 Elsevier Science Ltd. All rights reserved.
PII: S0957-4166(99)00255-4

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K. Khumtaveeporn et al. / Tetrahedron: Asymmetry 10 (1999) 2563–2572
Scheme 1.
Chemical modification,3 genetic engineering,4 the addition of water-miscible organic solvents,5 en-
zyme immobilization, the use of high pH reaction conditions6 and site directed mutagenesis7 have already
been explored with the goal of creating better peptide ligation biocatalysts. However, further progress is
0
necessary since serine proteases have rather restricted P1 and P1 side-chain specificities, especially in
their preference for the L-amino acid stereoisomers.1 The incorporation of D-amino acids into peptides
is of interest due to their prevalence in numerous biologically active compounds.8 The current work is
directed towards removing the above specificity limitations, with the aim of expanding the applicability
of serine proteases for peptide coupling reactions.
Recently, we have successfully exploited the combination of site directed mutagenesis and chemical
modification to modify the representative serine protease subtilisin Bacillus lentus (SBL).9 This strategy
involves the introduction of a cysteine residue at non-catalytic active site locations by site-directed
mutagenesis. Since wild-type SBL (WT-SBL) does not contain any cysteine residues, the introduced
cysteine provides a unique thiol for modification. The thiol group of the introduced cysteine is modified
by reaction with a variety of methanethiosulfonate reagents to produce chemically modified mutant
enzymes (CMMs). For this study, the S166C mutant, which is located as the bottom of the S1 pocket
0
and modulates P1 specificity, and the M222C mutant, which is located at the entrance to the S1 pocket
and is adjacent to the catalytic triad residues, were subjected to chemical modification as outlined in
Scheme 2.10,11
Scheme 2.
Several subtilisin CMMs (a–d) with significantly lowered amidase activity and which retain or surpass
WT esterase activity levels were identified.12,13 We now report the application of these selected CMMs
as catalysts in peptide coupling reactions which demonstrate both increased yields and broadened
specificity, in particular with respect to the incorporation of the unnatural D-amino acid series.

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2565
2. Results and discussion
The CMMs previously identified as having high esterase and lowered amidase activity12,13 were
selected as suitable candidate catalysts for peptide ligation. All of the CMMs (Scheme 2) chosen for
the current peptide ligation study have esterase-to-amidase selectivity ratios up to 48-fold higher than
WT.12,13 Each of the CMMs were prepared and fully characterized as described previously.12,13
The peptide coupling reactions between acyl donors 1–4 and acyl acceptors 5 and 6 (Scheme 3) were
conducted in 50% aqueous DMF. The acyl donors 1–4 were chosen as representative examples of large
and small hydrophobic, negatively charged and positively charged P1 side-chain-containing acyl donors,
respectively, and permit the broad evaluation of the S1 pocket specificity. The small amino acid amides
0
5 and 6 were chosen as the acyl acceptors since the S1 pocket of subtilisin has narrow specificity and
accepts α-branched amino acids only poorly.14 The coupling reactions are shown in Scheme 3 and the
results are summarized in Table 1.
Scheme 3.
The higher yield of Z-L-Phe-Gly-NH2 (7) compared to Z-L-Ala-Gly-NH2 (8), Z-L-Glu-Gly-NH2 (9)
and Z-L-Lys-Gly-NH2 (10) with WT-SBL as the ligation catalyst (Table 1) attests to the preference of
the S1 pocket of SBL for large hydrophobic residues.15 Furthermore, the yield of dipeptide 7 is excellent
with each of the four S166-CMMs as a ligation catalyst, which is reflective of the high resistance of this
+
pocket to attempts to change its specificity. Interestingly, the M222C-SCH2CH2NH3 CMM, which has
Table 1
WT-SBL and SBL-CMM-catalyzed coupling of L-amino acids (1–4) and glycinamide (5)a

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K. Khumtaveeporn et al. / Tetrahedron: Asymmetry 10 (1999) 2563–2572
Table 2
WT-SBL and SBL-CMM-catalyzed coupling of L-amino acids (1–4) and L-alaninamide (6)a
the highest esterase-to-amidase selectivity ratio of the CMMs evaluated here, gave only a 33% yield of
dipeptide 7 after 5 h, and was accompanied by the recovery of a 41% yield of the starting material 1.
The yields reported in Table 1 illustrate that Z-L-Ala-OBn (2) and Z-L-Glu-OMe (3) are poorer acyl
donors than Z-L-Phe-OBn (1). However, the yields of dipeptides Z-L-Ala-Gly-NH2 (8) and Z-L-Glu-
Gly-NH2 (9) could generally be improved by increasing the reaction time from 1 h to 5 h. On the other
hand, the yield of dipeptide Z-L-Glu-Gly-NH2 (9) was not increased by longer reaction time when WT
or S166C-SCH2C6F5 was used as the ligation catalyst. Nevertheless, Z-L-Glu-Gly-NH2 was isolated
+
in quantitative yield after 5 h when S166C-S-CH2CH2NH3 , with its S1 pocket possessing a positive
charge complementary to the negative one of Glu-P1 residue, was employed as the ligation catalyst,
versus an only 62% yield with WT. The enhanced yield due to favorable S1–P1 electrostatic interaction
+
is in line with our previous observation of rate enhancement of the S166C-SCH2CH2NH3 CMM with
the complementarily negatively charged P1-Glu containing substrate Suc-AAPE-p-NA.16 In all cases,
M222C-SCH2CH2NH3+ proved to be a poor ligation catalyst.
Of the five CMMs evaluated, three CMMs gave an improved yield of the Z-L-Lys-Gly-NH2 (10)
dipeptide compared to WT. The almost quantitative yield of dipeptide 10 which results from the use of the
complementary negatively charged S1 pocket S166C-SBnCOOH ligation catalyst is attributed to a favor-
able electrostatic substrate–enzyme interaction, again consistent with previous kinetic experiments.16 It
is noteworthy that dipeptides 7–10 were isolated in good yield when S166C-SBn was used as the ligation
catalyst. This is attributed to the high esterase-to-amidase ratio of this enzyme.
With the success of improved yields attributable to complementary P1–S1 electrostatic interaction in
hand, the ability of the selected CMMs to accept α-branched amino acid acyl acc0eptors was investigated.
The smallest α-branched amino acid, L-Ala-NH2 (6) was used to probe the S1 subsite tolerance. The
0
constricted nature of the S1 site was reflected by the longer reaction time of 24 h required when L-
Ala-NH2 was used as the acyl acceptor instead of Gly-NH2 (5) (Table 2). In all cases, the reaction of
the Z-L-Phe-OBn (1) acyl donor was slower with 6 than 5. After 24 h only a 57% yield of dipeptide Z-
+
L-Phe-L-Ala-NH2 (11) was isolated with the WT ligation catalyst. The S166C-SCH2CH2NH3 ligation
catalyst gave improved dipeptide yields. In fact the excellent performance of this catalyst is exemplified
by the fact that it is the only one which yields any Z-L-Ala-L-Ala-NH2 (12) and 0Z-L-Glu-L-Ala-NH2
(13) dipeptides. These results contrast the previously reported preference of the S1 pocket of subtilisin
Bacillus lentus for Ala over Gly.15 In all cases yields obtained using Gly-NH2 (5) as the acyl acceptor are
higher, and the reaction faster (Table 1), compared to using L-Ala-NH2 (6) (Table 2) as the acyl acceptor.
As the serine proteases, and WT-SBL in particular, have a normal preference for L-amino acid acyl

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2567
donors, the ability of the CMM approach to extend the applicability of SBL catalyzed peptide ligation
to include D-amino acid ester (15–18) acyl donors (Scheme 4) was explored (Table 3). Excitingly,
compared with the exclusive L-stereospecificity of the WT enzyme, the stereospecificity of all of the
S166C-CMMs was broadened, resulting in good yields of D-amino acid containing dipeptides (20–23).
While each of these enzymes still shows a preference for L-amino acids, yields of up to 66% of the D-
amino acid containing dipeptide Z-D-Phe-Gly-NH2 (20) were achieved using S166C-SBn as the ligation
catalyst. This expansion of stereospecificity demonstrates a truly dramatic CMM-induced change in
SBL’s acceptance of D-amino acids.
Scheme 4.
The use of any of the CMM peptide ligation catalysts effected rather similar yields of the Z-D-Phe-
Gly-NH2 (20) and Z-D-Ala-Gly-NH2 (21) dipeptides, suggesting that the mode of expansion of the
stereospecificity is similar for all the CMMs. Since both large neutral, negatively and positively charged
side chains were introduced into S1 by chemical modification, and no discernible specificity pattern was
observed, the mode of action of the observed stereospecificity relaxation is attributed to an altered binding
mode of the D-amino acid acyl donors, such that carbobenzoxy (Z)-group becomes able to bind in the S1
pocket.
When Z-D-Glu-OMe (17) was used as the acyl donor, 3 to 10% yields of dipeptide Z-D-Glu-Gly-
NH2 (22) were obtained with all CMM catalyzed reactions. While these yields are low, they represent
a significant improvement over the zero yield with WT. Furthermore, replacing the Z-group of the Z-
D-Phe-Gly-NH2 (15) acyl donor with the acetyl group, as in Ac-D-Phe-Gly-NH2 (19), resulted in lower
yields (Table 3). This effect may be due to the differences in desolvation energies of the acetyl and
Z-groups or to poorer binding of the acetyl group compared to the Z-group in the S1 pocket.
To determine if any new beneficial interaction between the Z-D-Phe-OBn acyl donor and the S166C-
Table 3
WT-SBL and SBL-CMM-catalyzed coupling of D-amino acids (15–19) and glycinamide (5)a

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K. Khumtaveeporn et al. / Tetrahedron: Asymmetry 10 (1999) 2563–2572
SBn CMM, which is not present in the WT, could be responsible for the enhanced reaction, molecular
modeling analysis was employed. The energy of the structure of the ES complex of each of the Z-L-Phe-
OBn and Z-D-Phe-OBn substrates bound to each of WT and S166C-SBn was minimized. The L-series of
substrates were positioned in the enzyme active sites of WT and S166C-SBn such that the Z-group was
oriented toward the S2 pocket, the benzyl side chain of phenylalan0ine was oriented into the S1 pocket
and the -OBn leaving group of the ester was oriented toward the S1 pocket. In contrast, the D-series of
substrates were positioned in the enzyme active sites of WT and S166C-SBn such that the benzyl side
chain of phenylalanine was oriented toward the S2 pocket, the Z-group was oriented into the S1 pocket
0
and the -OBn leaving group of the ester was oriented toward the S1 pocket. In the energy minimized
structures, for three of the four structures the orientation of the P1 and P2 and P10 oriented residues were
0
quite similar. For the fourth ES complex between Z-D-Phe-OBn and S166C-SBn, the -OBn0P1 moiety
0
0
was better oriented into the S1 pocket than for the others, for which P1 points toward S2 . Since the
S166C-SBn CMM shows the greatest expansion of stereospecificity with the Z-D-Phe-OBn acyl donor
it is interesting that this ES pair adopts a different preferred binding mode to the others. However, the
beneficial influence of the altered binding is not clear from the modeling.
3. Conclusion
The synthetic utility of chemically modified mutant enzymes for peptide synthesis has been demons-
trated. Using the CMM approach we were able to incorporate a variety of amino acids as acyl donors into
dipeptide products. These included charged P1 amino acids which typically are not accepted into the S1
0
pocket of WT-SBL. The specificity of the S1 pocket of subtilisin was broadened to accept α-branched
amino acids as efficient acyl acceptors. Moreover, significant broadening of S1 pocket stereospecificity
was also observed. Most dramatic is that all of the chemically modified mutant enzymes could accept
D-amino acids as acyl donors while the WT enzyme accepted none. These results illustrate that the
combined site-directed mutagenesis and chemical modification approach represents a powerful tool for
broadening the use of enzymes in preparative organic synthesis applications, and further studies are in
progress.
4. Experimental
4.1. General methods
WT-subtilisin Bacillus lentus and mutant enzymes, S166C and M222C were purified9 and prepared as
previously reported.9c,13 Protected amino acids were purchased from Sigma or Bachem and were used as
received. All solvents were reagent grade and distilled prior to use. Thin layer chromatography analysis
and purification were performed on pre-coated Merck silica gel (60 F-254) plates (250 µm) visualized
1
with UV light or iodine. H and 13C NMR spectra were recorded on a Varian Gemini 200 (200 MHz
1
1
for H and 50.3 MHz for 13C) or Unity 400 (400 MHz for H and 100 MHz for 13C) spectrometer and
chemical shifts are given in ppm (δ) using CDCl3 or DMSO-d6 as an internal standard. High resolution
mass spectra (HRMS) were recorded using Micromass ZAB-SE (FAB+). Optical rotations were measured
with a Perkin–Elmer 243B polarimeter.

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2569
4.2. General procedure for peptide ligation
To a solution of amino acid acyl donor (0.1 mmol) in DMF (0.4 mL) and water (0.4 mL), glycinamide
hydrochloride (0.3 mmol) or alaninamide hydrochloride (0.2 mmol) and Et3N (0.083–0.125 mL, 0.3–0.4
mmol) were added, followed by addition of a solution of 1 mg of active enzyme (0.0037 mmol, 0.037
equiv.), as determined by titration with phenylmethanesulfonyl fluoride (PMSF),17 in buffer solution (10
mmol MES, 1 mmol CaCl2, pH 5.8). The resulting total volume of reaction was 1.0–1.2 mL, with an
apparent pH of 9.7. The reaction was left stirring at room temperature for the period of time indicated
in Tables 1–3. In the case of D-amino acids being used as acyl donors, after 24 h, 1 mg more of active
enzymes as well as an equal amount of DMF were added. After the reaction was finished, the mixture
was then concentrated in vacuo and subjected to purification using preparative TLC (5–10% MeOH in
CH2Cl2). Yields are given in Tables 1–3. The absence of background ligation was established by omitting
enzyme from reaction mixture. Properties of the products are as follows.
4.2.1. Z-L-Phe-Gly-NH2 (7)14a,18
1H NMR (CDCl3) δ 3.10 (m, 2H, CH2Ph), 3.85 (2×d, J=2, 5 Hz, 2H, NHCH2CO), 4.40 (m, 1H,
NHCHCO), 5.05 (s, 2H, OCH2Ph), 5.50, 5.70, 6.25, 6.90 (4×br s, 4H, NH), 7.20–7.40 (m, 10H, 2×Ph);
13C NMR (CDCl3) δ 38.2, 42.7, 56.6, 67.3, 127.2, 128.1, 128.3, 128.6, 128.8, 129.2, 135.8, 136.0, 156.3,
171.2, 171.6. HRMS (FAB+) MH+ calcd for C19H22N3O4: 356.1610; found: 356.1613. [α]D30=−4.3 (c
0.81, MeOH).
4.2.2. Z-L-Ala-Gly-NH2 (8)19
1H NMR (CDCl3) δ 1.40 (d, J=7 Hz, 3H, CH3), 3.85 (dd, J=1.7, 5 Hz, 2H, NCH2CO), 4.20 (m, 1H,
NHCHCO), 5.10 (dd, J=1.6, 2 Hz, 2H, OCH2Ph), 5.80, 6.60, 7.20 (3×br s, 4H, NH), 7.30–7.40 (m,
5H, Ph); 13C NMR (CDCl3) δ 17.7, 42.2, 50.6, 66.2, 127.6, 128.0, 136.0, 155.9, 171.3, 172.8. HRMS
(FAB+) MH+ calcd for C13H18N3O4: 280.1297; found: 280.1310. [α]D26=−8.44 (c 0.97, MeOH); lit.
[α]D23=−8.5 (c 2.0, MeOH).
4.2.3. Z-L-Glu-Gly-NH2 (9)20
1H NMR (DMSO-d6) δ 1.75–1.95 (m, 2H, CH2CH2COOH), 2.30 (m, 2H, CH2CH2COOH), 3.65 (dd,
J=0.5, 1.7 Hz, 2H, NHCH2CO), 4.10 (m, 1H, NHCHCO), 5.00 (s, 2H, OCH2Ph), 7.20–7.40 (m, 5H,
Ph), 12.40 (br s, 1H, COOH); 13C NMR (DMSO-d6) δ 27.2, 30.3, 41.9, 54.2, 65.6, 128.4, 128.5, 128.7,
128.8, 136.9, 156.2, 170.8, 171.7, 174.0. HRMS (FAB+) MH+ calcd for C15H20N3O6: 338.1352; found:
338.1332. [α]D28=−10.2 (c 1.16, MeOH); lit. [α]D25=−10.2 (c 1.0, MeOH).
4.2.4. Z-L-Lys-Gly-NH2 (10)
1H NMR (DMSO-d6) δ 1.50 (m, 2H, CH2(CH2)3NH2), 1.60–1.85 (m, 4H, CH2CH2CH2CH2NH2),
2.00–2.18 (m, 2H, (CH2)3CH2NH2) 3.30 (m, 2H, NHCH2CO), 4.40 (m, 1H, NHCHCO), 5.10 (s, 2H,
OCH2Ph), 6.05, 6.20 (2×br s, 2H, NH), 7.20–7.40 (m, 5H, Ph); 13C NMR (DMSO-d6) δ 28.1, 29.0,
32.2, 42.3, 45.9, 53.8, 66.7, 128.1, 128.6, 136.7, 155.6, 172.2, 175.3. HRMS (FAB+) MH+ calcd for
C16H24N4O4: 337.1876; found: 337.1842. [α]D28=+6.97 (c 0.55, MeOH).
4.2.5. Z-L-Phe-L-Ala-NH2 (11)18,21
1H NMR (DMSO-d6) δ 1.20 (d, J=7 Hz, 3H, CH3), 2.90 (m, 2H, CH2Ph), 4.30 (m, 2H, 2×NHCHCO),
4.90 (s, 2H, OCH2Ph), 5.75, 6.10, 6.45 (3×br s, 4H, NH), 6.90–7.40 (m, 10H, 2×Ph); 13C NMR (DMSO-
d6) δ 18.5, 37.4, 48.1, 56.2, 66.2, 126.3, 127.4, 127.7, 128.1, 128.3, 129.2, 137.1, 138.2, 155.9, 171.2,

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K. Khumtaveeporn et al. / Tetrahedron: Asymmetry 10 (1999) 2563–2572
174.1. HRMS (FAB+) MH+ calcd for C20H24N3O4: 370.1766; found: 370.1753. [α]D29=−8.86 (c 0.57,
MeOH).
4.2.6. Z-L-Ala-L-Ala-NH2 (12)22
1H NMR (DMSO-d6) δ 1.20 (2×d, J=7 Hz, 6H, 2×CH3), 4.10, 4.20 (m, 2H, 2×NHCHCO), 5.00 (s,
2H, OCH2Ph), 7.20–7.40 (m, 5H, Ph); 13C NMR (DMSO-d6) δ 18.1, 18.4, 47.9, 50.1, 65.4, 127.7, 128.4,
137.0, 155.8, 172.0, 174.1. HRMS (FAB+) MH+ calcd for C14H20N3O4: 294.1454; found: 294.1457.
[α]D25=−20.4 (c 0.77, MeOH).
4.2.7. Z-L-Glu-L-Ala-NH2 (13)
1H NMR (DMSO-d6) δ 1.20 (d, J=7 Hz, 3H, CH3), 1.82–2.00 (m, 2H, CH2CH2COOH), 2.30 (m,
2H, CH2CH2COOH), 4.00, 4.20 (m, 2H, 2×NHCHCO), 5.00 (s, 2H, OCH2Ph), 6.20–7.40 (m, 5H, Ph);
13C NMR (DMSO-d6) δ 18.4, 26.2, 30.2, 47.9, 53.1, 65.4, 126.5, 127.7, 127.8, 128.1, 128.4, 137.0,
156.2, 173.6, 173.8, 174.1. HRMS (FAB+) MH+ calcd for C16H22N3O6: 352.1509; found: 352.1478.
[α]D25=−16.7 (c 0.76, MeOH).
4.2.8. Z-D-Phe-Gly-NH2 (20)
1H and 13C NMR data are identical to (7). HRMS (FAB+) MH+ calcd for C19H22N3O4: 356.1610;
found: 356.1608; [α]D30=+4.12 (c 1.17, MeOH).
4.2.9. Z-D-Ala-Gly-NH2 (21)23
1H and 13C NMR data are identical to (8). HRMS (FAB+) MH+ calcd for C13H18N3O4: 280.1297;
found: 280.1298; [α]D27=+10.5 (c 0.72, MeOH); lit. [α]D=+10.5.
4.2.10. Z-D-Glu-Gly-NH2 (22)
1H and 13C NMR data are identical to (9). HRMS (FAB+) MH+ calcd for C15H20N3O6: 338.1352;
found: 338.1348; [α]D28=+10.77 (c 1, MeOH).
4.2.11. Ac-D-Phe-Gly-NH2 (24)24
1H NMR (DMSO-d6) δ 1.90 (s, 3H, CH3), 3.05 (m, 2H, NHCH2CO), 3.65 (2×d, J=7, 15 Hz,
2H, CHCH2Ph), 4.40 (q, J=7 Hz, 1H, NHCHCO), 7.15–7.25 (m, 5H, Ph); 13C NMR (DMSO-d6) δ
22.4, 29.1, 45.9, 54.7, 126.1, 127.9, 128.9, 137.3, 155.2, 171.2, 171.5. HRMS (FAB+) MH+ calcd for
C13H17N3O3: 264.1348; found: 264.1321. [α]D30=−4.38 (c 0.80, MeOH).
4.3. Molecular modeling
The X-ray structure of subtilisin Bacillus lentus25 was used as the starting point for calculations on
the wild type and chemically modified mutant enzymes. The enzyme setup was performed with Insight
II.26 To create initial coordinates for the minimization, hydrogens were added at the pH used for kinetic
measurements. This protonated all Lys and Arg residues and the N-terminus and deprotonated all Glu and
Asp residues and the C-terminus. In addition, the active site His64 was protonated. The model system
with the Ala-Ala-Pro-Phe (from crystal structure)25 product inhibitor bound in the S1–S4 pocket was
solvated with a 5 Å layer of water molecules giving a total number of water molecules of 1143 in this
system. The overall charge of the enzyme–inhibitor complex resulting from this setup was +4 for the WT
enzyme. Energy simulations were performed with the Discover program,27 on a Silicon Graphics Iris
Indigo computer, using the consistent valence force field function (CVFF). A non-bonded cutoff distance

K. Khumtaveeporn et al. / Tetrahedron: Asymmetry 10 (1999) 2563–2572
2571
of 18 Å with a switching distance of 2 Å was employed. The non-bonded pair list was updated every
20 cycles and a dielectric constant of 1 was used in all calculations. The energy of the structure of the
WT enzyme was minimized in stages, with initially only the water molecules being allowed to move,
then the water molecules and the amino acid side chains, and then the entire enzyme. The mutated and
chemically modified enzymes were generated using the Builder module of Insight. Then the amino acid
side chains were minimized, and then all of the atoms were minimized. The AAPF product inhibitor was
subsequently replaced with the appropriate acyl donor substrate Z-L/D-Phe-OBn and then the energy of
this structure was minimized as above.
Acknowledgements
We are grateful to the National Sciences and Engineering Research Council of Canada and Genencor
International Inc. for generous funding, and Genencor for providing WT-, S166C- and M222C-SBL
enzymes. We also thank Dr. Erika Plettner for preliminary experiments and Dr. Benjamin G. Davis for
helpful discussion.
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