Proc. Nail. Acad. Sci. USA Vol. 88, pp. 4015-4019, May 1991 Biochemistry
Switching substrate preference of thermophilic xylose isomerase from D-xylose to D-glucose by redesigning the substrate binding pocket (glucose isomerase/site-directed mutagenesis/enzyme active site/protein engineering/catalytic efficiency)
MENGHSIAO MENG*t, CHANYONG LEEt§, MICHAEL BAGDASARIAN*t,
AND
J. GREGORY ZEIKUS*t¶II
*Michigan Biotechnology Institute, Lansing, MI 48909; tDepartment of Microbiology, Michigan State University, East Lansing, MI 48824; Department of Biochemistry, Michigan State University, East Lansing, MI 48824; and *Department of Pharmaceutical Chemistry and Biochemistry/Biophysics, University of California, San Francisco, CA 94143
Communicated by T. Kent Kirk, February 15, 1991 (received for review December 12, 1990)
ABSTRACT The substrate specificity of thermophilic xylose isomerase from Clostridium thenmosulfurogenes was examined by using predictions from the known crystal structure of the Arthrobacter enzyme and site-directed mutagenesis of the thermophilexylA gene. The orientation of glucose as a substrate in the active site of the thermophilic enzyme was modeled to position the C-6 end of hexose toward His-101 in the substratebinding pocket. The locations of Met-87, Thr-89, Val-134, and Glu-180, which contact the C-6 -OH group of the substrate in the sorbitol-bound xylose isomerase from Arthrobacter [Collyer, C. A., Henrick, K. & Blow, D. M. (1990) J. Mol. Biol. 212, 211-235], are equivalent to those of Trp-139, Thr-141, Val-186, and Glu-232 in the thermophilic enzyme. Replacement of Trp-139 with Phe reduced the K. and enhanced the kt of the mutant thermophilic enzyme toward glucose, whereas this substitution reversed the effect toward xylose. Replacement of Val-186 with Thr also enhanced the catalytic efficiency of the enzyme toward glucose. Double mutants with replacements Trp-139 -* Phe/Val-186 - Thr and Trp-139 Phe/Val-186 -* Ser had a higher catalytic efficiency (kCt/Kl,) for glucose than the wild-type enzyme of 5- and 2-fold, respectively. They also exhibited 1.5- and 3-fold higher catalytic efficiency for D-glucose than for D-xylose, respectively. These results provide evidence that alteration in substrate specificity of factitious thermophilic xylose isomerases can be achieved by designing reduced steric constraints and enhanced hydrogenbonding capacity for glucose in the substrate-binding pocket of the active site.
Specificity of enzymes toward their substrates is determined in part by molecular residues that provide for binding of the substrate and that maintain substrate steric configuration in the active site. A variety of factors influence enzymesubstrate complementarity and catalytic efficiency including steric fit, charge interactions, hydrogen bonding, and hydrophobic interactions (1). Until recently, the main strategy to reveal and study the molecular basis of these factors was to determine the tertiary structure of the enzyme-substrate complexes by x-ray crystallography. Redesigning proteins by engineering of their genes is now a viable approach that complements structural studies and enables determination of amino acid substitution effects on mutant enzyme function. Thus, substrate specificity has been altered by redesigning the structural frame of an enzyme (1-4), its electrostatic network (5-8), or its hydrophobic interaction with the substrate (9). Catalytic function of an enzyme can also be changed and regulated by modifications of the physical microenvironment of its catalytic site (10, 11). The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Xylose isomerase (D-xylose ketol-isomerase; EC 5.3.1.5) converts D-xylose to D-xylulose during xylose metabolism in various microorganisms (12). This enzyme also catalyzes the conversion of D-glucose to D-fructose in vitro and has been used as an industrial biocatalyst for production of high fructose corn syrup (13). Xylose isomerase displays lower kcat and higher Km values for glucose than those for xylose, and it requires different metal ions for enzyme catalysis on these substrates (i.e., Mn2+ for xylose and Co2+ for glucose) (14-16). The catalytic mechanism for xylose isomerase was originally believed to involve histidine-directed general base catalysis (17). Currently, an alternative mechanism of catalysis has been proposed based on results of x-ray crystallographic studies on Arthrobacter or Streptomyces enzymes (18-21) and biochemical properties exhibited by thermophilic enzymes obtained by site-directed mutagenesis of the xylA gene from Clostridium thermosulfurogenes (22). The enzymatic interconversion of aldose to ketose by xylose isomerases involves binding of the substrate in the ring form, substrate ring opening, isomerization of the linear intermediate, intermediate ring closure, and release of the product. The isomerization step is proposed to proceed by a metal ion-assisted hydride shift mechanism (18-22), and this step, rather than ring opening, is rate determining (22). D-xylose and D-glucose have identical atomic configuration, except for the presence of an additional -CH2OH group at the C-6 position in the glucose molecule. This extra hydroxymethyl group must therefore be responsible for the differences in the catalytic efficiency exhibited by xylose isomerase toward glucose versus xylose. We have cloned and overexpressed a gene encoding the thermophilic xylose isomerase of C. thermosulfurogenes in the mesophilic host, Escherichia coli, which enables very simple purification of preparative amounts of homogenous gene product (16). We have identified the active site histidine residue of the enzyme and the rate-limiting step in the isomerization reaction (22). The crystal structure of the Arthrobacter xylose isomerase has been determined at 2.3 A resolution (18, 21). Several amino acid residues, revealed by this structure, could constitute potential steric hindrance for the binding of a six-carbon substrate to the active site pocket of the enzyme. In this work we have substituted several key residues other than histidine (adjacent to the C-6 -OH group of glucose) in the active site of the thermophilic xylose isomerase. By the analysis of kinetic properties of the re-
§Present address: Department of Biochemical Process Research and Development, Merck Sharp & Dohme Research Laboratories, Rahway, NJ 07065. 'To whom reprint requests should be addressed at: Michigan Biotechnology Institute, 3900 Collins Road, P.O. Box 27609, Lansing, MI 48909.
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bation of the reaction mixture at 650C for 15 min, 0.5 ml of 0.5 M perchloric acid was added, and the products were determined by the cysteine/carbazole/sulfuric acid method (28). Km and VmS, were determined from Lineweaver-Burk and from Eadie-Hofstee plots. kat (i.e., turnover number per active site of the enzyme) was determined from the equation kcat[Elo = Vmax, where [Elo = total enzyme concentration
sulting mutant enzymes, we found indication of extensive similarities between the structure of the active domain of xylose isomerase from Arthrobacter and that from the thermophilic Clostridium. By changing some of the key amino acids in the substrate-binding pocket of the active site, we have changed substrate kinetic specificity constants of the thermophilic xylose isomerase enzyme. Notable, some of the designed, or factitious, enzymes display significantly higher catalytic efficiency toward glucose (the industrial substrate) than xylose (the natural substrate).
(29).
Computer-Aided Molecular Modeling. Atomic coordinates for the structure of xylose isomerase from Arthrobacter strain B3728 at 2.3 A resolution were kindly provided by D. M. Blow (21). The structure of the active site containing the six-carbon substrate analogue sorbitol was visualized on the IRIS-4D25 computer (Silicon Graphics Computer System, Mountain View, CA) with the aid of the INSIGHT II graphic program (Biosym Technologies, San Diego, CA).
MATERIALS AND METHODS Strains, Plasmids, and Chemicals. E. coli strain HB101 [FhsdS20 ara-l recA13 proA12 lacYl galK2 rpsL20 mtl-l xyl-5] (23) was used for expression of the C. thermosulfurogenes xylose isomerase gene present in the plasmid pCMG11-3 (22); E. coli strain TG1 [thi supE hisDS A(lac-proAB)/F' traD36 proA+B+ lacIq lacZAM15] in conjunction with bacteriophage M13mp19 (24) was used for oligonucleotide-directed mutagenesis and nucleotide sequence determination as described (22). All chemicals were of reagent grade. DNA Manipulation. Restriction endonucleases and other enzymes for. DNA manipulation were from Bethesda Research Laboratories or from New England Biolabs. The oligonucleotide-directed mutagenesis kit was from Amersham. The following oligonucleotides (obtained from Genosys, Woodlands, TX) were used for generation of sitedirected mutants: 5'-ACGAAAGTYTTGNNNGGTACTGCGAAT-3', where NNN = TTT for Trp-139 -- Phe and TAT for Trp-139 -* Tyr substitution; 5'-ACGAAAGTTTTGTGGGGTNNNGCGAATCTTTTCTCC-3', where NNN = TCT for Thr-141 -) Ser substitution; 5'-GGCGAAAACTAC NNlTTCTGGGGTGGA-3', where NNN = ACA for Val-186--+ Thr, TCA for Val-186--* Ser, and GCA for Val-186 Ala substitution. Synthesis of mutant genes were performed by the method of Sayers et al. (25), and their nucleotide sequences were confirmed by the dideoxy chain termination method (26). The 1.4-kilobase EcoRI/BamHI fragments containing the mutant genes were excised from the M13mp19 double-stranded replicative form DNA, inserted into the vector pMMB67EH (27), and introduced into E. coli strain HB101. Enzyme Purification and Assays. Wild-type and mutant xylose isomerases, expressed by E. coli HB101, were purified as described previously through the DEAE-Sepharose step, which gave enzymes homogeneous on SDS/PAGE (16). For determination of glucose isomerase activity, the reaction mixtures (1.0 ml) contained 50 mM Mops (pH 7.0), 10 mM MgSO4, 1 mM CoCl2, D-glucose (Km = 0.3-2.0), and enzyme (3-5 ,g). For xylose isomerase activity, reaction mixtures (1.0 ml) contained, in the same buffer, 10 mM MnSO4, D-xylose (Km = 0.3-2.0), and enzyme (3-5 ,ug). After incu-
E.c.
Putative Structure of the Active Site. Assuming that conservation of the primary sequence in the active site between xylose isomerases of different origins (Fig. 1) reflects similarities in tertiary structure of the binding-site domain (Fig. 2), the following residues in the thermophilic xylose isomerase might constitute steric hindrance for effective binding of D-glucose: Trp-139, Thr-141, Val-186, and Glu232. In the Arthrobacter enzyme, the structure ofthe enzyme complex with the six-carbon competitive inhibitor sorbitol indicated that the C-6 hydroxymethyl group of the substrate is oriented toward the bottom of the substrate-binding pocket and is adjacent to the residues Met-87, Thr-89, and Val-134. The distances to these atoms from the C-6 -OH group of sorbitol is of the order of 3.4-3.7 A (21). The residues Thr-89, Val-134, and Glu-180 from the Arthrobacter enzyme, are highly conserved among different sequences from divergent origins and correspond to Thr-141, Val-186, and Glu-232, respectively, in the Clostridium enzyme. Met-87 is conserved aknong the enzymes of the Arthrobacter type, but it is replaced and conserved by Trp-139 in the Clostridium- and Bacillus-type isomerases (see Fig. 1). To prove that the residues discussed above are indeed part of the substratebinding pocket of the thermophilic Clostridium isomerase, we have replaced each of these residues with smaller amino acids. Catalytic properties of the mutant enzymes, resulting from the substitution of Trp-139, Thr-141, and Val-186, are described below. The role of Glu-232 will be the subject of a separate communication. Properties of the Mutant Enzymes. Table 1 presents catalytic constants of the site-directed mutants constructed to understand the relationship between specific amino acids in the active-site pocket and enzyme-substrate interactions. Replacement of Trp-139 with Phe, a smaller residue, produced an enzyme that had a higher catalytic efficiency for
131 ETFVMWGGREG ------ KTLVLWGGREG ------ KTYVAWGGREG VPNATTNLFTHPVF ------ KTYVAWGGREG VPNATTNLFTHPVF ------ KTYVAWGGREG
85 VPMVTTNLFSHPVF VPNVTTNLFTHPVF VPKATTNLFTHPVF
A. r. A.m. S.v. S. o. S. g.
C.t. B. s.
RESULTS
137 VLWGTANLFSNPRF --LLWNTANMFTNPRF --- LLNGTANCFTNPRY ---
------
183 ENYVFWGGREG
------
-----------
179 LZPKPNEP
------
--
------
IZPKPNEP -IZPKPNEP --
-----------
------
IZPKPNQP -IZPKPNEP --
231 IZPKPKEP -ENYVFWGGREG ------ IZPKPKEP -ENYVLWGGREG -----IIPKPQEP -.
------
FIG. 1. Alignment of amino acid sequences of the substrate-binding region from different xylose isomerases. Boldfaced letters indicate residues changed in this work. A.r., Arthrobacter strain B3726; A.m., Ampullariella strain 3876; S.v., Streptomyces violaceoniger; S.o., Streptomyces olivochromogenes; S.g., Streptomyces griseofuscus; C.t., C. thermosulfurogenes; B.s., Bacillus subtilis; E.c., E. coli (see ref.
22).
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FIG. 2. Stereo analysis ofthe active-site region of xylose isomerase from Arthrobacter strain B3728. The carbon backbone of sorbitol (green), analog, was positioned to be perpendicular to the plane of the paper by positioning C-1 in front and C-6 in back, which is the bottom of the active-site pocket. The hydroxyl groups at C-2 and C4 are coordinated with a Mg2+ metal ion (red), which are surrounded by the negatively charged amino acid residues (blue). Van der Waals dot surfaces are shown for the atoms in the side chains of Met-87, Thr-89, Val-134, and Glu-180 (yellow), which are adjacent (within 3.9 A distance) to the C-6-OH atoms of the sorbitol (green). The active-site histidine that has been proposed to form a hydrogen bond to the C-5-OH of the substrate during the reaction is shown in red (see refs. 20-22). as a glucose
glucose than the wild-type enzyme (Fig. 3). This property resulted from a decrease of the Km and an increase in kcat. It should be noted that substitutions at this position increased the Km for xylose and decreased the catalytic efficiency for this substrate under the assay conditions employed. Thus, Trp-139 constitutes a steric hindrance for binding of the substrate with the larger molecule. We speculate that mutant isomerases with either Phe or Tyr at position 139 exhibit lower catalytic efficiencies toward xylose because the binding pocket is more spacious and this increases the freedom of movement of the xylose molecule, decreasing its binding efficiency. The enlargement of the binding pocket also decreases binding energy between the enzyme and the transition state resulting in the decrease of kcat. Mutant enzymes in which Val-186 was replaced by Thr, a polar residue, had a slightly lower Km and a higher kcat for glucose. Placement of a Ser residue, which has a smaller side chain but otherwise is equivalent to Thr, in this position did not improve the catalytic efficiency for glucose. Likewise, replacement with Ala did not significantly change either Km or kcat. Thus it would appear that Val-186 does not strictly hinder glucose binding, but the effect of placing a Thr in this position may be to improve the catalytic efficiency for glucose by providing additional hydrogen bonding, presumably to the C-6 -OH group of the substrate. Replacement of Thr-141 with Ser increased the Km for both xylose and glucose and resulted in Lower catalytic efficiency for glucose.
This is consistent with the view that Thr-141 hydrogen bonds to the substrate but does not strictly hinder the binding of glucose.
DISCUSSION This work is relevant to both the scientific understanding of xylose (glucose) isomerase catalysis and to the applied use of this enzyme. First, the mechanism of catalysis for xylose isomerase has become controversial in view of the x-ray crystallographic data and recent interpretations that question the previously proposed biochemical reaction mechanisms for this enzyme (19, 20). Our analysis of site-specific mutant xylose isomerases, performed in this work and reported previously (22), when taken together with structural data obtained at higher resolution (21), provides additional insight into the xylose isomerase catalysis mechanism. Second, a molecular strategy has been delineated to improve on the industrially important catalytic reaction of the enzyme (i.e., fructose manufacture from glucose)-namely, to design an in vitro function via protein engineering based on understanding of the structure and the catalytic mechanism of the enzyme. Although the overall homology between the isomerase of Arthrobacter and that of Clostridium is only 26% (22), it was possible to predict the probable location of several essential amino acid residues in the Clostridium isomerase on the basis of sequence homology and the tertiary structure of the
Table 1. Catalytic properties of wild-type and mutant enzymes obtained by substitution of amino acids in the active center of C. thermosulfurogenes xylose isomerase Glucose Xylose Mutant enzyme changes wt
Trp-139 --Phe Trp-139 Tyr Val-186 Thr Val-186 Ser Val-186 Ala Trp-139 Phe/Val-186 Trp-139 Phe/Val-186 Thr-141 Ser wt, wild type. *Double mutant.
Thr* Ser*
Ki, mM 110 ± 7.6 65 ± 7.4 91 ± 12 91 ± 7 140 ± 7 100 ± 11 29 ± 3.7 58 ± 3.9 160 ± 19
kat min-1 ± ± ± ± ± ± ± ± 470 ±
640 970 540 880 790 540 950 720
85 40 25 86 15 56 106 20
31
kcat/Km, min-I-mM-1 5.8 15 6.0 9.7 5.7 5.3 32.9 12.4 2.9
Km, mM
kcat, min-1
± 2.2 ± 1.1 ± 13.2 ± 1.7 ± 8.7 ± 0.7 ± 2.4 ± 0.4 68 ± 12
1100 620 360 740 780 1000 780 250
12 46 110 13 49 27 36 63
± ± ± ± ± ± ± ± 1500 ±
106 25 25 40
96 15 66 5
162
kcat/Kmg min-'mM-1 97.2 13.6 3.2 55.4 15.9 36.4 21.6 4.0 21.7
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Proc. Natl. Acad. Sci. USA 88 (1991)
by the properties of the mutant enzymes with Val-186 -* Ser and, particularly, Val-186 -+ Ala substitutions. The results of crystallographic studies on xylose isomerases bound to different substrate analogs were interpreted to point to two alternative orientations of the substrate in the active site of the enzyme (17, 20, 21). Results of this work provide functional evidence supporting the orientation in which the C-5 atom of glucose is located near the His-101 residue at the bottom of the active site pocket and the C-1 and C-2 hydroxyls can form coordination bonds with the metal ions. The results presented here do not give a comprehensive picture of all molecular interactions between the substrate and the enzyme in the binding site of xylose isomerase. They merely provide indications that require further direct enzyme structural measurements on the thermophilic Clostridium xylose isomerase molecule. Biochemical analysis of additional site-directed mutant xylose isomerases is required to understand the function of metal centers in the enzyme catalysis and stability.
i2
J 0)
0
0
0-
amt
LL>
2
.> > > 0
0)
Enzymes FIG. 3. Diagram illustrating amino acid changes of substrate preference from xylose (Xyl) to glucose (Glc) associated with the amino acid substitutions in the substrate-binding pocket of xylose isomerase. The ratios of catalytic efficiency (katt/Km) of enzymes with xylose versus that with glucose, shown in Table 1, are expressed in a logarithmic scale. The negative values shown by factitious enzymes indicate more favored enzyme specificity toward glucose than xylose, which is required of "true" glucose isomerase. Amino acids are indicated by the single-letter code.
Arthrobacter isomerase resolved by x-ray diffraction (18, 20, 21). Thus, in a previous report, the function of His-101 was elucidated and found in agreement with the structural data (22). In the present work good evidence was obtained that Thr-141, Trp-139, and Val-186 are part of the substratebinding site and that they play roles in substrate binding and in delineating the borders of the active-site pocket, based on a functional test (i.e., the change of kinetic properties of the mutant enzymes resulting from substitution of these amino acids). We do not know at present whether the enzymatic isomerization of xylose and glucose proceeds via the formation of a Michaelis-type enzyme-substrate complex. In fact, the results of crystallographic studies of the enzyme bound to D-xylose indicated that a non-Michaelis-type complex is formed (20). However, a lower apparent Km of the mutant enzymes in which Trp-139 was replaced by smaller residues indicates an increased affinity of the enzyme for glucose or for its reaction intermediates. We interpret it, therefore, as a result of the increased volume of the substrate-binding pocket, which now accommodates more readily the extra hydroxymethyl group of the glucose molecule. The observed changes in kcat are more difficult to explain on the basis of the available data. We can conclude that the stabilization of the transition state has changed in the mutant enzymes as a result of structural changes in the binding site. Although it is not surprising that the kcat has changed in these mutants, the molecular basis of these changes must await further structural studies. The Val-186 residue does not seem to hinder substrate binding in the Clostridium xylose isomerase. The increased affinity for glucose upon replacement of Val-186 with Thr seems to result from the ability of this residue to provide an additional hydrogen bond to the substrate. This is indicated
This work resulted from an equal contribution by the first two authors. We thank D. M. Blow for providing structural coordinates of Arthrobacter isomerase before submission to the data bank, C. S. Craik for reading the manuscript, R. J. Fletterick for the use of computer modeling equipment, J. H. McKerrow for the use of his laboratory facilities, and C. Bystroff for helpful discussions. This work was supported by grants from the U.S. Department of Agriculture (89-01053 to Michigan Biotechnology Institute), Center for Microbial Ecology, a National Science Foundation Science and Technology Center at Michigan State University, the Research Excellence Fund from Michigan State University, and by National Science Foundation Grant BCS-8897179 (Dr. C. S. Craik as the principal investigator) for C.L. 1. Craik, C. S., Largman, C., Fletcher, T., Roczniak, S., Barr, P. J., Fletterick, R. & Rutter, W. (1985) Science 228, 291-297. 2. Wilks, H. M., Hart, K. W., Feeney, R., Dunn, C. R., Muirhead, H., Chia, W. N., Barstow, D. A., Atkinson, T., Clarke, A. R. & Holbrook, J. J. (1988) Science 242, 1541-1544. 3. Bone, R., Silen, J. L. & Agard, D. A. (1989) Nature (London) 339, 191-195. 4. Scrutton, N. S., Berry, A. & Perham, R. N. (1990) Nature (London) 343, 3843. 5. Wilkinson, A. J., Fersht, A. R., Blow, D. M., Carter, P. & Winter, G. (1984) Nature (London) 307, 187-188. 6. Wells, J. A., Powers, D. B., Bott, R. R., Graycar, T. P. &
Estell, D. A. (1987) Proc. Natl. Acad. Sci. USA 84, 1219-1223. 7. Dean, A. D. & Koshland, Jr., D. E. (1990) Science 249, 10441046. 8. Evnin, L. B., Vdsquez, J. R. & Craik, C. S. (1990) Proc. Natl.
Acad. Sci. USA 87, 6659-6663. 9. Estell, D. A., Graycar, T. P., Miller, J. V., Powers, D. B., Burnier, J. P., Ng, P. G. & Wells, J. A. (1986) Science 233,
659-663. 10. Hurley, J. H., Dean, A. M., Sohl, J. L., Koshland, Jr., D. E.
& Stroud, R. M. (1990) Science 249, 1012-1016.
11. Higaki, J. N., Haymore, B. L., Chen, S., Fletterick, R. & Craik, C. S. (1990) Biochemistry 29, 8582-8586.
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13. Antrim, R. L., Colliala, W. & Schnyder, B. (1979) in Applied Biochemistry and Bioengineering, ed. Wangard, E. B. (Academic, New York), pp. 97-155. 14. Danno, G. (1970) Agric. Biol. Chem. 34, 1805-1814. 15. Danno, G. (1971) Agric. Biol. Chem. 35, 997-1006. 16. Lee, C. & Zeikus, J. G. (1991) Biochem. J. 273, 565-571. 17. Carrell, H. L., Glusker, J. P., Burger, V., Manfre, F., Tritsch, D. & Biellmann, J.-F. (1989) Proc. Natl. Acad. Sci. USA 86, 4440-4444. 18. Henrick, K., Collyer, C. A. & Blow, D. M. (1989) J. Mol. Biol. 208, 129-157. 19. Farber, G. K., Glasfeld, A., Tiraby, G., Ringe, D. & Petsko, G. A. (1989) Biochemistry 28, 7289-7297. 20. Collyer, C. A. & Blow, D. M. (1990) Proc. NatI. Acad. Sci. USA 87, 1362-1366. 21. Collyer, C. A., Henrick, K. & Blow, D. M. (1990) J. Mol. Biol. 212, 211-235.
Biochemistry: Meng et al. 22. Lee, C., Bagdasarian, M., Meng, M. & Zeikus, J. G. (1990) J. Biol. Chem. 265, 19082-19090. 23. Boyer, H. W. & Roulland-Dussoix, D. (1969) J. Mol. Biol. 41, 459-472. 24. Yanish-Perron, C., Vieira, J. & Messing, J. (1985) Gene 33, 103-119. 25. Sayers, J. R., Schmidt, W. & Eckstein, F. (1988) Nucleic Acids Res. 16, 791-802.
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26. Sanger, F., Nicklen, S. & Coulson, A. R. (1977) Proc. Nati. Acad. Sci. USA 74, 5463-5467. 27. Furste, J. P., Pansegrau, W., Frank, R., Blocker, H., Scholz, P., Bagdasarian, M. & Lanka, E. (1986) Gene 48, 119-131. 28. Dische, Z. & Borenfreund, E. (1951) J. Biol. Chem. 192, 583-587. 29. Fersht, A. R. (1985) Enzyme Structure and Mechanisms (Freeman, San Francisco, CA), 2nd Ed.
Switching substrate preference of thermophilic xylose isomerase from D-xylose to D-glucose by redesigning the substrate binding pocket (glucose isomerase/site-directed mutagenesis/enzyme active site/protein engineering/catalytic efficiency)
MENGHSIAO MENG*t, CHANYONG LEEt§, MICHAEL BAGDASARIAN*t,
AND
J. GREGORY ZEIKUS*t¶II
*Michigan Biotechnology Institute, Lansing, MI 48909; tDepartment of Microbiology, Michigan State University, East Lansing, MI 48824; Department of Biochemistry, Michigan State University, East Lansing, MI 48824; and *Department of Pharmaceutical Chemistry and Biochemistry/Biophysics, University of California, San Francisco, CA 94143
Communicated by T. Kent Kirk, February 15, 1991 (received for review December 12, 1990)
ABSTRACT The substrate specificity of thermophilic xylose isomerase from Clostridium thenmosulfurogenes was examined by using predictions from the known crystal structure of the Arthrobacter enzyme and site-directed mutagenesis of the thermophilexylA gene. The orientation of glucose as a substrate in the active site of the thermophilic enzyme was modeled to position the C-6 end of hexose toward His-101 in the substratebinding pocket. The locations of Met-87, Thr-89, Val-134, and Glu-180, which contact the C-6 -OH group of the substrate in the sorbitol-bound xylose isomerase from Arthrobacter [Collyer, C. A., Henrick, K. & Blow, D. M. (1990) J. Mol. Biol. 212, 211-235], are equivalent to those of Trp-139, Thr-141, Val-186, and Glu-232 in the thermophilic enzyme. Replacement of Trp-139 with Phe reduced the K. and enhanced the kt of the mutant thermophilic enzyme toward glucose, whereas this substitution reversed the effect toward xylose. Replacement of Val-186 with Thr also enhanced the catalytic efficiency of the enzyme toward glucose. Double mutants with replacements Trp-139 -* Phe/Val-186 - Thr and Trp-139 Phe/Val-186 -* Ser had a higher catalytic efficiency (kCt/Kl,) for glucose than the wild-type enzyme of 5- and 2-fold, respectively. They also exhibited 1.5- and 3-fold higher catalytic efficiency for D-glucose than for D-xylose, respectively. These results provide evidence that alteration in substrate specificity of factitious thermophilic xylose isomerases can be achieved by designing reduced steric constraints and enhanced hydrogenbonding capacity for glucose in the substrate-binding pocket of the active site.
Specificity of enzymes toward their substrates is determined in part by molecular residues that provide for binding of the substrate and that maintain substrate steric configuration in the active site. A variety of factors influence enzymesubstrate complementarity and catalytic efficiency including steric fit, charge interactions, hydrogen bonding, and hydrophobic interactions (1). Until recently, the main strategy to reveal and study the molecular basis of these factors was to determine the tertiary structure of the enzyme-substrate complexes by x-ray crystallography. Redesigning proteins by engineering of their genes is now a viable approach that complements structural studies and enables determination of amino acid substitution effects on mutant enzyme function. Thus, substrate specificity has been altered by redesigning the structural frame of an enzyme (1-4), its electrostatic network (5-8), or its hydrophobic interaction with the substrate (9). Catalytic function of an enzyme can also be changed and regulated by modifications of the physical microenvironment of its catalytic site (10, 11). The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Xylose isomerase (D-xylose ketol-isomerase; EC 5.3.1.5) converts D-xylose to D-xylulose during xylose metabolism in various microorganisms (12). This enzyme also catalyzes the conversion of D-glucose to D-fructose in vitro and has been used as an industrial biocatalyst for production of high fructose corn syrup (13). Xylose isomerase displays lower kcat and higher Km values for glucose than those for xylose, and it requires different metal ions for enzyme catalysis on these substrates (i.e., Mn2+ for xylose and Co2+ for glucose) (14-16). The catalytic mechanism for xylose isomerase was originally believed to involve histidine-directed general base catalysis (17). Currently, an alternative mechanism of catalysis has been proposed based on results of x-ray crystallographic studies on Arthrobacter or Streptomyces enzymes (18-21) and biochemical properties exhibited by thermophilic enzymes obtained by site-directed mutagenesis of the xylA gene from Clostridium thermosulfurogenes (22). The enzymatic interconversion of aldose to ketose by xylose isomerases involves binding of the substrate in the ring form, substrate ring opening, isomerization of the linear intermediate, intermediate ring closure, and release of the product. The isomerization step is proposed to proceed by a metal ion-assisted hydride shift mechanism (18-22), and this step, rather than ring opening, is rate determining (22). D-xylose and D-glucose have identical atomic configuration, except for the presence of an additional -CH2OH group at the C-6 position in the glucose molecule. This extra hydroxymethyl group must therefore be responsible for the differences in the catalytic efficiency exhibited by xylose isomerase toward glucose versus xylose. We have cloned and overexpressed a gene encoding the thermophilic xylose isomerase of C. thermosulfurogenes in the mesophilic host, Escherichia coli, which enables very simple purification of preparative amounts of homogenous gene product (16). We have identified the active site histidine residue of the enzyme and the rate-limiting step in the isomerization reaction (22). The crystal structure of the Arthrobacter xylose isomerase has been determined at 2.3 A resolution (18, 21). Several amino acid residues, revealed by this structure, could constitute potential steric hindrance for the binding of a six-carbon substrate to the active site pocket of the enzyme. In this work we have substituted several key residues other than histidine (adjacent to the C-6 -OH group of glucose) in the active site of the thermophilic xylose isomerase. By the analysis of kinetic properties of the re-
§Present address: Department of Biochemical Process Research and Development, Merck Sharp & Dohme Research Laboratories, Rahway, NJ 07065. 'To whom reprint requests should be addressed at: Michigan Biotechnology Institute, 3900 Collins Road, P.O. Box 27609, Lansing, MI 48909.
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bation of the reaction mixture at 650C for 15 min, 0.5 ml of 0.5 M perchloric acid was added, and the products were determined by the cysteine/carbazole/sulfuric acid method (28). Km and VmS, were determined from Lineweaver-Burk and from Eadie-Hofstee plots. kat (i.e., turnover number per active site of the enzyme) was determined from the equation kcat[Elo = Vmax, where [Elo = total enzyme concentration
sulting mutant enzymes, we found indication of extensive similarities between the structure of the active domain of xylose isomerase from Arthrobacter and that from the thermophilic Clostridium. By changing some of the key amino acids in the substrate-binding pocket of the active site, we have changed substrate kinetic specificity constants of the thermophilic xylose isomerase enzyme. Notable, some of the designed, or factitious, enzymes display significantly higher catalytic efficiency toward glucose (the industrial substrate) than xylose (the natural substrate).
(29).
Computer-Aided Molecular Modeling. Atomic coordinates for the structure of xylose isomerase from Arthrobacter strain B3728 at 2.3 A resolution were kindly provided by D. M. Blow (21). The structure of the active site containing the six-carbon substrate analogue sorbitol was visualized on the IRIS-4D25 computer (Silicon Graphics Computer System, Mountain View, CA) with the aid of the INSIGHT II graphic program (Biosym Technologies, San Diego, CA).
MATERIALS AND METHODS Strains, Plasmids, and Chemicals. E. coli strain HB101 [FhsdS20 ara-l recA13 proA12 lacYl galK2 rpsL20 mtl-l xyl-5] (23) was used for expression of the C. thermosulfurogenes xylose isomerase gene present in the plasmid pCMG11-3 (22); E. coli strain TG1 [thi supE hisDS A(lac-proAB)/F' traD36 proA+B+ lacIq lacZAM15] in conjunction with bacteriophage M13mp19 (24) was used for oligonucleotide-directed mutagenesis and nucleotide sequence determination as described (22). All chemicals were of reagent grade. DNA Manipulation. Restriction endonucleases and other enzymes for. DNA manipulation were from Bethesda Research Laboratories or from New England Biolabs. The oligonucleotide-directed mutagenesis kit was from Amersham. The following oligonucleotides (obtained from Genosys, Woodlands, TX) were used for generation of sitedirected mutants: 5'-ACGAAAGTYTTGNNNGGTACTGCGAAT-3', where NNN = TTT for Trp-139 -- Phe and TAT for Trp-139 -* Tyr substitution; 5'-ACGAAAGTTTTGTGGGGTNNNGCGAATCTTTTCTCC-3', where NNN = TCT for Thr-141 -) Ser substitution; 5'-GGCGAAAACTAC NNlTTCTGGGGTGGA-3', where NNN = ACA for Val-186--+ Thr, TCA for Val-186--* Ser, and GCA for Val-186 Ala substitution. Synthesis of mutant genes were performed by the method of Sayers et al. (25), and their nucleotide sequences were confirmed by the dideoxy chain termination method (26). The 1.4-kilobase EcoRI/BamHI fragments containing the mutant genes were excised from the M13mp19 double-stranded replicative form DNA, inserted into the vector pMMB67EH (27), and introduced into E. coli strain HB101. Enzyme Purification and Assays. Wild-type and mutant xylose isomerases, expressed by E. coli HB101, were purified as described previously through the DEAE-Sepharose step, which gave enzymes homogeneous on SDS/PAGE (16). For determination of glucose isomerase activity, the reaction mixtures (1.0 ml) contained 50 mM Mops (pH 7.0), 10 mM MgSO4, 1 mM CoCl2, D-glucose (Km = 0.3-2.0), and enzyme (3-5 ,g). For xylose isomerase activity, reaction mixtures (1.0 ml) contained, in the same buffer, 10 mM MnSO4, D-xylose (Km = 0.3-2.0), and enzyme (3-5 ,ug). After incu-
E.c.
Putative Structure of the Active Site. Assuming that conservation of the primary sequence in the active site between xylose isomerases of different origins (Fig. 1) reflects similarities in tertiary structure of the binding-site domain (Fig. 2), the following residues in the thermophilic xylose isomerase might constitute steric hindrance for effective binding of D-glucose: Trp-139, Thr-141, Val-186, and Glu232. In the Arthrobacter enzyme, the structure ofthe enzyme complex with the six-carbon competitive inhibitor sorbitol indicated that the C-6 hydroxymethyl group of the substrate is oriented toward the bottom of the substrate-binding pocket and is adjacent to the residues Met-87, Thr-89, and Val-134. The distances to these atoms from the C-6 -OH group of sorbitol is of the order of 3.4-3.7 A (21). The residues Thr-89, Val-134, and Glu-180 from the Arthrobacter enzyme, are highly conserved among different sequences from divergent origins and correspond to Thr-141, Val-186, and Glu-232, respectively, in the Clostridium enzyme. Met-87 is conserved aknong the enzymes of the Arthrobacter type, but it is replaced and conserved by Trp-139 in the Clostridium- and Bacillus-type isomerases (see Fig. 1). To prove that the residues discussed above are indeed part of the substratebinding pocket of the thermophilic Clostridium isomerase, we have replaced each of these residues with smaller amino acids. Catalytic properties of the mutant enzymes, resulting from the substitution of Trp-139, Thr-141, and Val-186, are described below. The role of Glu-232 will be the subject of a separate communication. Properties of the Mutant Enzymes. Table 1 presents catalytic constants of the site-directed mutants constructed to understand the relationship between specific amino acids in the active-site pocket and enzyme-substrate interactions. Replacement of Trp-139 with Phe, a smaller residue, produced an enzyme that had a higher catalytic efficiency for
131 ETFVMWGGREG ------ KTLVLWGGREG ------ KTYVAWGGREG VPNATTNLFTHPVF ------ KTYVAWGGREG VPNATTNLFTHPVF ------ KTYVAWGGREG
85 VPMVTTNLFSHPVF VPNVTTNLFTHPVF VPKATTNLFTHPVF
A. r. A.m. S.v. S. o. S. g.
C.t. B. s.
RESULTS
137 VLWGTANLFSNPRF --LLWNTANMFTNPRF --- LLNGTANCFTNPRY ---
------
183 ENYVFWGGREG
------
-----------
179 LZPKPNEP
------
--
------
IZPKPNEP -IZPKPNEP --
-----------
------
IZPKPNQP -IZPKPNEP --
231 IZPKPKEP -ENYVFWGGREG ------ IZPKPKEP -ENYVLWGGREG -----IIPKPQEP -.
------
FIG. 1. Alignment of amino acid sequences of the substrate-binding region from different xylose isomerases. Boldfaced letters indicate residues changed in this work. A.r., Arthrobacter strain B3726; A.m., Ampullariella strain 3876; S.v., Streptomyces violaceoniger; S.o., Streptomyces olivochromogenes; S.g., Streptomyces griseofuscus; C.t., C. thermosulfurogenes; B.s., Bacillus subtilis; E.c., E. coli (see ref.
22).
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FIG. 2. Stereo analysis ofthe active-site region of xylose isomerase from Arthrobacter strain B3728. The carbon backbone of sorbitol (green), analog, was positioned to be perpendicular to the plane of the paper by positioning C-1 in front and C-6 in back, which is the bottom of the active-site pocket. The hydroxyl groups at C-2 and C4 are coordinated with a Mg2+ metal ion (red), which are surrounded by the negatively charged amino acid residues (blue). Van der Waals dot surfaces are shown for the atoms in the side chains of Met-87, Thr-89, Val-134, and Glu-180 (yellow), which are adjacent (within 3.9 A distance) to the C-6-OH atoms of the sorbitol (green). The active-site histidine that has been proposed to form a hydrogen bond to the C-5-OH of the substrate during the reaction is shown in red (see refs. 20-22). as a glucose
glucose than the wild-type enzyme (Fig. 3). This property resulted from a decrease of the Km and an increase in kcat. It should be noted that substitutions at this position increased the Km for xylose and decreased the catalytic efficiency for this substrate under the assay conditions employed. Thus, Trp-139 constitutes a steric hindrance for binding of the substrate with the larger molecule. We speculate that mutant isomerases with either Phe or Tyr at position 139 exhibit lower catalytic efficiencies toward xylose because the binding pocket is more spacious and this increases the freedom of movement of the xylose molecule, decreasing its binding efficiency. The enlargement of the binding pocket also decreases binding energy between the enzyme and the transition state resulting in the decrease of kcat. Mutant enzymes in which Val-186 was replaced by Thr, a polar residue, had a slightly lower Km and a higher kcat for glucose. Placement of a Ser residue, which has a smaller side chain but otherwise is equivalent to Thr, in this position did not improve the catalytic efficiency for glucose. Likewise, replacement with Ala did not significantly change either Km or kcat. Thus it would appear that Val-186 does not strictly hinder glucose binding, but the effect of placing a Thr in this position may be to improve the catalytic efficiency for glucose by providing additional hydrogen bonding, presumably to the C-6 -OH group of the substrate. Replacement of Thr-141 with Ser increased the Km for both xylose and glucose and resulted in Lower catalytic efficiency for glucose.
This is consistent with the view that Thr-141 hydrogen bonds to the substrate but does not strictly hinder the binding of glucose.
DISCUSSION This work is relevant to both the scientific understanding of xylose (glucose) isomerase catalysis and to the applied use of this enzyme. First, the mechanism of catalysis for xylose isomerase has become controversial in view of the x-ray crystallographic data and recent interpretations that question the previously proposed biochemical reaction mechanisms for this enzyme (19, 20). Our analysis of site-specific mutant xylose isomerases, performed in this work and reported previously (22), when taken together with structural data obtained at higher resolution (21), provides additional insight into the xylose isomerase catalysis mechanism. Second, a molecular strategy has been delineated to improve on the industrially important catalytic reaction of the enzyme (i.e., fructose manufacture from glucose)-namely, to design an in vitro function via protein engineering based on understanding of the structure and the catalytic mechanism of the enzyme. Although the overall homology between the isomerase of Arthrobacter and that of Clostridium is only 26% (22), it was possible to predict the probable location of several essential amino acid residues in the Clostridium isomerase on the basis of sequence homology and the tertiary structure of the
Table 1. Catalytic properties of wild-type and mutant enzymes obtained by substitution of amino acids in the active center of C. thermosulfurogenes xylose isomerase Glucose Xylose Mutant enzyme changes wt
Trp-139 --Phe Trp-139 Tyr Val-186 Thr Val-186 Ser Val-186 Ala Trp-139 Phe/Val-186 Trp-139 Phe/Val-186 Thr-141 Ser wt, wild type. *Double mutant.
Thr* Ser*
Ki, mM 110 ± 7.6 65 ± 7.4 91 ± 12 91 ± 7 140 ± 7 100 ± 11 29 ± 3.7 58 ± 3.9 160 ± 19
kat min-1 ± ± ± ± ± ± ± ± 470 ±
640 970 540 880 790 540 950 720
85 40 25 86 15 56 106 20
31
kcat/Km, min-I-mM-1 5.8 15 6.0 9.7 5.7 5.3 32.9 12.4 2.9
Km, mM
kcat, min-1
± 2.2 ± 1.1 ± 13.2 ± 1.7 ± 8.7 ± 0.7 ± 2.4 ± 0.4 68 ± 12
1100 620 360 740 780 1000 780 250
12 46 110 13 49 27 36 63
± ± ± ± ± ± ± ± 1500 ±
106 25 25 40
96 15 66 5
162
kcat/Kmg min-'mM-1 97.2 13.6 3.2 55.4 15.9 36.4 21.6 4.0 21.7
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by the properties of the mutant enzymes with Val-186 -* Ser and, particularly, Val-186 -+ Ala substitutions. The results of crystallographic studies on xylose isomerases bound to different substrate analogs were interpreted to point to two alternative orientations of the substrate in the active site of the enzyme (17, 20, 21). Results of this work provide functional evidence supporting the orientation in which the C-5 atom of glucose is located near the His-101 residue at the bottom of the active site pocket and the C-1 and C-2 hydroxyls can form coordination bonds with the metal ions. The results presented here do not give a comprehensive picture of all molecular interactions between the substrate and the enzyme in the binding site of xylose isomerase. They merely provide indications that require further direct enzyme structural measurements on the thermophilic Clostridium xylose isomerase molecule. Biochemical analysis of additional site-directed mutant xylose isomerases is required to understand the function of metal centers in the enzyme catalysis and stability.
i2
J 0)
0
0
0-
amt
LL>
2
.> > > 0
0)
Enzymes FIG. 3. Diagram illustrating amino acid changes of substrate preference from xylose (Xyl) to glucose (Glc) associated with the amino acid substitutions in the substrate-binding pocket of xylose isomerase. The ratios of catalytic efficiency (katt/Km) of enzymes with xylose versus that with glucose, shown in Table 1, are expressed in a logarithmic scale. The negative values shown by factitious enzymes indicate more favored enzyme specificity toward glucose than xylose, which is required of "true" glucose isomerase. Amino acids are indicated by the single-letter code.
Arthrobacter isomerase resolved by x-ray diffraction (18, 20, 21). Thus, in a previous report, the function of His-101 was elucidated and found in agreement with the structural data (22). In the present work good evidence was obtained that Thr-141, Trp-139, and Val-186 are part of the substratebinding site and that they play roles in substrate binding and in delineating the borders of the active-site pocket, based on a functional test (i.e., the change of kinetic properties of the mutant enzymes resulting from substitution of these amino acids). We do not know at present whether the enzymatic isomerization of xylose and glucose proceeds via the formation of a Michaelis-type enzyme-substrate complex. In fact, the results of crystallographic studies of the enzyme bound to D-xylose indicated that a non-Michaelis-type complex is formed (20). However, a lower apparent Km of the mutant enzymes in which Trp-139 was replaced by smaller residues indicates an increased affinity of the enzyme for glucose or for its reaction intermediates. We interpret it, therefore, as a result of the increased volume of the substrate-binding pocket, which now accommodates more readily the extra hydroxymethyl group of the glucose molecule. The observed changes in kcat are more difficult to explain on the basis of the available data. We can conclude that the stabilization of the transition state has changed in the mutant enzymes as a result of structural changes in the binding site. Although it is not surprising that the kcat has changed in these mutants, the molecular basis of these changes must await further structural studies. The Val-186 residue does not seem to hinder substrate binding in the Clostridium xylose isomerase. The increased affinity for glucose upon replacement of Val-186 with Thr seems to result from the ability of this residue to provide an additional hydrogen bond to the substrate. This is indicated
This work resulted from an equal contribution by the first two authors. We thank D. M. Blow for providing structural coordinates of Arthrobacter isomerase before submission to the data bank, C. S. Craik for reading the manuscript, R. J. Fletterick for the use of computer modeling equipment, J. H. McKerrow for the use of his laboratory facilities, and C. Bystroff for helpful discussions. This work was supported by grants from the U.S. Department of Agriculture (89-01053 to Michigan Biotechnology Institute), Center for Microbial Ecology, a National Science Foundation Science and Technology Center at Michigan State University, the Research Excellence Fund from Michigan State University, and by National Science Foundation Grant BCS-8897179 (Dr. C. S. Craik as the principal investigator) for C.L. 1. Craik, C. S., Largman, C., Fletcher, T., Roczniak, S., Barr, P. J., Fletterick, R. & Rutter, W. (1985) Science 228, 291-297. 2. Wilks, H. M., Hart, K. W., Feeney, R., Dunn, C. R., Muirhead, H., Chia, W. N., Barstow, D. A., Atkinson, T., Clarke, A. R. & Holbrook, J. J. (1988) Science 242, 1541-1544. 3. Bone, R., Silen, J. L. & Agard, D. A. (1989) Nature (London) 339, 191-195. 4. Scrutton, N. S., Berry, A. & Perham, R. N. (1990) Nature (London) 343, 3843. 5. Wilkinson, A. J., Fersht, A. R., Blow, D. M., Carter, P. & Winter, G. (1984) Nature (London) 307, 187-188. 6. Wells, J. A., Powers, D. B., Bott, R. R., Graycar, T. P. &
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