©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Enhancement of the Endo--1,4-glucanase Activity of an Exocellobiohydrolase by Deletion of a Surface Loop (*)

(Received for publication, October 27, 1994)

Andreas Meinke (§) Howard G. Damude Peter Tomme Emily Kwan Douglas G. Kilburn Robert C. Miller Jr. R. Antony J. Warren Neil R. Gilkes (¶)

From the Department of Microbiology and Immunology, and Protein Engineering Network of Centres of Excellence, University of British Columbia, Vancouver, British Columbia, V6T 1Z3, Canada

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

In the commonly accepted mechanism for enzymatic hydrolysis of cellulose, endo-beta-1,4-glucanases randomly cleave glucosidic bonds within glucan polymers, providing sites for attack by exo-cellobiohydrolases (EC 3.2.1.91). It has been proposed that hydrolysis by Trichoderma reesei cellobiohydrolase II is restricted to the ends of cellulose polymers because two surface loops cover its active site to form a tunnel. In a closely related endoglucanase, E2 from Thermomonospora fusca, access to the substrate appears to be relatively unhindered because the carboxyl-proximal loop is shortened, and the amino-proximal loop is displaced. The hypothesis was examined by deletion of a region in Cellulomonas fimi cellobiohydrolase A corresponding to part of the carboxyl-proximal loop of T. reesei cellobiohydrolase II. The mutation enhanced the endoglucanase activity of the enzyme on soluble O-(carboxymethyl)cellulose and altered its activities on 2`,4`-dinitrophenyl-beta-D-cellobioside, insoluble cellulose, and cellotetraose.


INTRODUCTION

The fungal and bacterial beta-1,4-glucanases involved in cellulose degradation are usually comprised of two or more structural and functional domains; a catalytic domain joined to a cellulose-binding domain is a common arrangement(1) . The enzymes range in size from about 20 to 120 kDa, but nearly all can be classified into a few distinct families based on sequence identity of their catalytic domains alone(1, 2) . Family 6, originally called family B, contains eight known members; these include both exoglucanases (cellobiohydrolases) and endoglucanases(3) .

The catalytic domain of cellobiohydrolase II (CBH II), (^1)a family 6 exo-beta-1,4-glucanase from Trichoderma reesei, contains a central alpha/beta barrel, similar to that of triose-phosphate isomerase(4) . Two extensive surface loops extend from the carboxyl end of the barrel and enclose the active site to form a tunnel-shaped structure (Fig. 1). Amino acid sequence alignment suggests that the two surface loops covering the CBH II active site are shortened in the family 6 endoglucanases. These observations led to the hypothesis that the basic difference between exo- and endoglucanases is the accessibility of their active sites to internal beta-1,4-glucosidic bonds in polymeric substrates. The tunnel-shaped active site of an exoglucanase restricts hydrolysis to beta-1,4-glucosidic bonds at the ends of cellulose molecules(4) .


Figure 1: alpha-Carbon skeletons of the T. reesei CBH II and T. fuscaE2 catalytic domains. The views are chosen to illustrate differences in the accessibilities of the two active sites. C and N, respectively, indicate the carboxyl- and amino-proximal loops that cover the active site of CBH II.



Structural analysis of the catalytic domain of Thermomonospora fuscaE2, a family 6 endo-beta-1,4-glucanase, subsequently provided support for this proposal(5) . Although the general topology of the E2 alpha/beta barrel is very similar to that of CBH II, the E2 active site is an open cleft structure (Fig. 1). The carboxyl-proximal loop covering the CBH II active site (Val-Ala, Fig. 2) is indeed shortened in E2, whereas the amino-proximal loop (Pro-Asp in CBH II), although present, is bent back so that it no longer covers the active site.


Figure 2: Partial amino acid sequence of CbhADeltaC, a deletion mutant of CbhA, and its alignment with corresponding regions of the catalytic domains of CbhA and other known family 6 beta-1,4-glucanases. The region in CbhA that was deleted is underlined. CbhA (GenBank L25809) and CenA (GenBank M15823) are from C. fimi, CBH II (GenBank M16190) from T. reesei, CEL3 (GenBank L24519) from Agaricus bisporus, CelA (Swiss-Prot P26414) from Microbispora bispora, Cel1 (Swiss-Prot P33682) from Streptomyces halstedii; CasA (GenBank L03218) from Streptomyces sp. KSM-9, and E2 (GenBank M73321) from Thermomonospora fusca. CbhA, CBH II, and CEL3 are exocellobiohydrolases; CenA, Cel1, CasA, and E2 are endoglucanases. Each sequence is numbered from the first residue of the mature enzyme; hyphens indicate gaps introduced to improve the alignment. Asterisks mark a pair of cysteine residues known to form one of two disulfide bonds in the catalytic domains of CenA, CBH II, and E2; the arrow indicates the putative general base catalyst(4) .



The hypothesis described above implies that deletion of the loops covering its active site would allow an exoglucanase to hydrolyze internal beta-1,4-glucosidic bonds in cellulose molecules. We tested this prediction by constructing a mutant of cellobiohydrolase A (CbhA), a family 6 exo-beta-1,4-glucanase from Cellulomonas fimi, in which residues corresponding to part of the CBH II carboxyl-proximal active site loop are deleted.


EXPERIMENTAL PROCEDURES

Construction of a Plasmid Encoding CbhADeltaC

The sequence of plasmid pALM1 (3) was changed by site-directed mutagenesis to introduce NcoI and NheI sites (underlined) at the start of the cbhA gene, using the oligonucleotide 5`-CACCCGGAGGACCCCATGGCTAGCCTCGGCAAGCGAGCA-3`. This allowed insertion of an NcoI-EcoRI fragment containing cbhA into the NcoI- and EcoRI-digested plasmid pDAM2-1; this vector was chosen because it was known to give high level expression of another C. fimi beta-1,4-glucanase gene, cenD(6) . The resulting plasmid (pALM3) served as template for further mutagenesis. Mutation of cbhA to cbhADeltaC used the oligonucleotide 5`-CCCCCGGGTGAGTCCGACGGCGCATGCGACCCGACGTTCGTCTCGCCC-3`. The SphI site (underlined) was introduced as a silent mutation to facilitate screening. The resulting plasmid, pALM3.2, encoded CbhADeltaC, a mutant in which CbhA residues corresponding to part of the carboxyl-proximal active site loop of CBH II are deleted (Fig. 2).

Preparation and Assay of Enzymes

CbhA(3) , CenA(7) , and Abg (8) were purified from Escherichia coli containing appropriate recombinant plasmids by affinity chromatography on microcrystalline cellulose and anion-exchange chromatography, as described previously. CbhADeltaC was purified from E. coli DH5alpha F` (pALM3.2), as described for CbhA(3) . Purifications were monitored by SDS-polyacrylamide gel electrophoresis(9) .

Purified CbhA and CbhADeltaC were examined for the presence of free thiol groups by titration with 5,5`-dithiobis(2-nitrobenzoic acid)(10) . Fifty µl of buffer (100 mM potassium phosphate, pH 7.3, 1 mM EDTA, 6 M guanidine hydrochloride) containing 3 mM 5,5`-dithiobis(2-nitrobenzoic acid) were added to 1 ml of buffer containing 10 nmol of enzyme, and the mixture was incubated at 25 °C. Formation of nitrothiobenzoate was monitored from its absorbance at 412 nm over a period of 1 h. The lower limit of detection was 0.1 nmol of free thiol.

Simultaneous determination of specific fluidity ( = ) and reducing sugar during Cm-cellulose hydrolysis has been described(11) . Reaction mixtures contained 2 µM CbhA, 2 µM CbhADeltaC, 3.6 nM CenA or 300 nM Abg, and 3.2% (w/v) Cm-cellulose (Sigma; low viscosity grade; nominal degree of substitution, 0.7; nominal degree of polymerization, 400) in 50 mM sodium citrate, pH 7; mixtures were incubated at 37 °C.

Michaelis-Menten parameters for the hydrolysis of 2`,4`-dinitrophenyl-beta-D-cellobioside were determined by spectrophotometric measurement of the rate of dinitrophenolate release in 50 mM potassium phosphate, pH 7, at 37 °C (12) ; substrate concentrations were from 0.2 to 10 times K(m).

Hydrolysis of insoluble cellulose was assayed by incubation of 1.5 mg of acid-swollen CF1 cellulose (13) and 1.5 nmol of enzyme in 1.5 ml of 5 mM potassium phosphate, pH 7, at 37 °C. Aliquots were removed at 2, 6, and 24 h and centrifuged to sediment residual cellulose. The resulting supernatants were incubated for 5 min at 100 °C to prevent further hydrolysis of soluble sugar prior to analysis. Total soluble sugar in supernatants was determined by the phenol sulfuric acid assay(14) ; component sugars were analyzed by high performance liquid chromatography(6) .

Cellotetraose hydrolysis was examined by incubation of 50 pmol of enzyme and 40 nmol of cellotetraose in 75 µl of 5 mM potassium phosphate, pH 7, at 37 °C for 30 min. Reactions were stopped by incubation at 100 °C for 5 min and the products analyzed by high performance liquid chromatography.

Structural Representations

Coordinates for CBH II were from the Protein Data Bank (acquisition number 3CBH); coordinates for E2 were generously provided by Drs. Spezio, Wilson, and Karplus.


RESULTS AND DISCUSSION

General Characterization of CbhADeltaC

The region Ser-Met of CbhA (Fig. 2) was deleted to give the mutant protein CbhADeltaC. The deleted residues correspond to 11 of the 20-amino acid residue carboxyl-proximal loop covering the active site in CBH II ( Fig. 1and Fig. 2). Purified CbhADeltaC co-migrated with CbhA on an SDS-polyacrylamide gel with an apparent molecular mass of about 85 kDa, in agreement with its predicted molecular mass(3) . No contaminating polypeptides were visible on a heavily loaded gel (data not shown).

A disulfide bond occurs between Cys and Cys in C. fimi CenA, a family 6 endo-beta-1,4-glucanase (Fig. 2); a second bond is formed between Cys and Cys(7) . Bonds occur between corresponding residues in CBH II (4) and E2(15) ; presumably, they are common to all family 6 catalytic domains because the 4 cysteine residues are strictly conserved(3) . The 11-residue deletion of CbhADeltaC is immediately adjacent to Cys (Fig. 2); possible interference with disulfide bond formation was examined using 5,5`-dithiobis(2-nitrobenzoic acid). No free thiol groups were detected in either CbhADeltaC or CbhA, indicating that all disulfide bonds are correctly formed in the mutant and wild-type enzymes.

Relative Exo- and Endohydrolytic Activities of CbhA and CbhADeltaC

The preference of a beta-1,4-glucanase for terminal or internal glucosidic bonds can be evaluted from a plot of specific fluidity () versus reducing sugar production during hydrolysis of soluble Cm-cellulose. The hydrolysis of any beta-1,4-glucosidic bond generates one new reducing sugar group, but the random action of an endoglucanase results in a greater increase in specific fluidity than attack by an exoglucanase(11) . The slope of the plot for CbhA was 17.1 bulletmlbulletmmol (Fig. 3). The slope for CbhA is considered representative of exoglucanase activity because a similar low slope (13.3 bulletmlbulletmmol) was obtained with Abg (Fig. 3); Abg is an Agrobacterium sp. beta-1,4-glucosidase that cleaves glucosyl residues from the nonreducing end of cellobiose and longer beta-1,4-glucans by an exclusive exohydrolytic mode of attack(16) . The data for CbhA are consistent with a mechanism in which the enzyme preferentially removes unsubstituted cellobiosyl residues from the ends of Cm-cellulose polymers. We have previously shown that >95% of the soluble sugar released from microcrystalline cellulose by CbhA is cellobiose(3) . In contrast to CbhA and Abg, CenA gave a slope of 98.0 bulletmlbulletmmol; a similar high slope (88.1 bulletmlbulletmmol) was reported for CenD, another C. fimi endo-beta-1,4-glucanase(17) .


Figure 3: Relationship between specific fluidity () and total reducing sugar during hydrolysis of Cm-cellulose by CbhA, CbhADeltaC, CenA, or Abg. The insets show the rate of reducing sugar production for each enzyme.



The plot for CbhADeltaC had a slope of 44.5 bulletmlbulletmmol (Fig. 3). Therefore, deletion of amino acid residues 373-387 from CbhA enhances its endoglucanase activity. These data support the hypothesis that the accessibility of the active site is a major determinant of exo- or endohydrolytic activity in family 6 beta-1,4-glucanases, a conclusion hitherto based on structural interpretations alone. Access to the CbhADeltaC active site may be still partially restricted, possibly by the amino-proximal loop, because the slope of the CbhADeltaC plot was lower than that for CenA. However, bond cleavage preference may be influenced by additional factors. For example, crystallographic data suggest differences in the binding of small ligands to the four subsites in the active sites of CBH II and E2(5) .

Effects of Loop Deletion on Other Enzymatic Properties

Both enzymes showed very slow turnover on Cm-cellulose (Fig. 3, insets), but the rate of hydrolysis by CbhADeltaC (0.6 mol of productbulletminbulletmol of enzyme, calculated from reducing sugar production after 30 min) was lower than that for CbhA (1.7 molbulletminbulletmol). Similarly, the k for CbhADeltaC on 2`,4`-dinitrophenyl-beta-D-cellobioside, a small soluble substrate analog, was reduced by about 5-fold, although the K(m) was unchanged (Table 1). The deletion in CbhADeltaC occurs only 2 amino acid residues from Asp, the putative general base catalyst, and results in the loss of several charged residues (Fig. 2). Therefore, the reduced catalytic efficiency of CbhADeltaC is most easily explained by perturbation of the active site environment.



The rate of hydrolysis of insoluble cellulose by CbhADeltaC (56 mg of productbullethbulletµmol of enzyme, calculated from total sugar solubilized after 2 h) was also lower than that of CbhA (105 mgbullethbulletµmol) (Fig. 4), but analysis of the soluble products revealed a further difference in the activities of the two enzymes. The CbhADeltaC reaction mixture contained a significant quantity of cellotetraose (Fig. 5A); in contrast, cellotetraose was not seen in the CbhA reaction mixture, neither after 6 h of hydrolysis (Fig. 5A) nor at earlier times (data not shown). It appears that cellotetraose accumulates in the reaction mixture because CbhADeltaC is unable to degrade this substrate efficiently (Fig. 5B). Under the same conditions, cellotetraose was completely degraded to cellobiose by CbhA. Mechanistic interpretation of these data is not yet possible, but they clearly show that deletion of the loop has multiple effects on the activity of CbhA.


Figure 4: Release of total soluble sugar from acid-swollen cellulose by CbhA (), CbhADeltaC (bullet), or CenA ().




Figure 5: High performance liquid chromatographic analysis of products from cellulose (A) or cellotetraose (B). Soluble cello-oligosaccharides released from acid-swollen cellulose were analyzed after hydrolysis by CbhA, CbhADeltaC, or CenA for 6 h (see Fig. 4); products from cellotetraose were analyzed after hydrolysis by CbhA or CbhADeltaC for 30 min. Glucose (1) and cellobiose (2) were each resolved as single peaks; cellotriose (3) and cellotetraose (4) were partially resolved into their alpha- and beta-anomers.




FOOTNOTES

*
This work was supported by a grant from the Natural Sciences and Engineering Research Council and by the Protein Engineering Network of Centres of Excellence, Canada. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Present address: Dept. of Microbiology and Genetics, Dr. Bohrgasse 9, University of Vienna, 1030 Vienna, Austria.

To whom correspondence should be addressed: Dept. of Microbiology and Immunology, 6174 University Blvd., University of British Columbia, Vancouver, British Columbia V6T 1Z3, Canada.

(^1)
The abbreviations used are: CBH II, T. reesei cellobiohydrolase II; Abg, Agrobacterium sp. beta-1,4-glucosidase; CbhA, C. fimi cellobiohydrolase A; CenA, C. fimi endoglucanase A; E2, T. fusca endoglucanase 2.


ACKNOWLEDGEMENTS

We thank Dr. Michael Spezio for advice and assistance.


REFERENCES

  1. Gilkes, N. R., Henrissat, B., Kilburn, D. G., Miller, R. C., Jr., and Warren, R. A. J. (1991) Microbiol. Rev. 55, 303-315
  2. Henrissat, B., and Bairoch, A. (1993) Biochem. J. 293, 781-788 [Medline] [Order article via Infotrieve]
  3. Meinke, A., Gilkes, N. R., Kwan, E., Kilburn, D. G., Warren, R. A. J., and Miller, R. C., Jr. (1994) Mol. Microbiol. 12, 413-422 [Medline] [Order article via Infotrieve]
  4. Rouvinen, J., Bergfors, T., Teeri, T., Knowles, J. K. C., and Jones, T. A. (1990) Science 249, 380-386 [Medline] [Order article via Infotrieve]
  5. Spezio, M., Wilson, D. B., and Karplus, P. A. (1993) Biochemistry 32, 9906-9916 [Medline] [Order article via Infotrieve]
  6. Meinke, A., Gilkes, N. R., Kilburn, D. G., Miller, R. C., Jr., and Warren, R. A. J. (1993) J. Bacteriol. 175, 1910-1918 [Abstract]
  7. Gilkes, N. R., Claeyssens, M., Aebersold, R., Henrissat, B., Meinke, A., Morrison, H. D., Kilburn, D. G., Warren, R. A. J., and Miller, R. C., Jr. (1991) Eur. J. Biochem. 202, 367-377 [Abstract]
  8. Wakarchuk, W. W., Kilburn, D. G., Miller, R. C., Jr., and Warren, R. A. J. (1986) Mol. & Gen. Genet. 205, 146-152
  9. Gilkes, N. R., Warren, R. A. J., Miller, R. C., Jr., and Kilburn, D. G. (1988) J. Biol. Chem. 263, 10401-10407 [Abstract/Free Full Text]
  10. Riddles, P. W., Blakely, R. L., and Zerner, B. (1983) Methods Enzymol. 91, 49-60 [Medline] [Order article via Infotrieve]
  11. Gilkes, N. R., Langsford, M. L., Kilburn, D. G., Miller, R. C., Jr., and Warren, R. A. J. (1984) J. Biol. Chem. 259, 10455-10459 [Abstract/Free Full Text]
  12. Kempton, J. B., and Withers, S. G. (1992) Biochemistry 31, 9961-9969 [Medline] [Order article via Infotrieve]
  13. Coutinho, J. B., Gilkes, N. R., Warren, R. A. J., Kilburn, D. G., and Miller, R. C., Jr. (1992) Mol. Microbiol. 6, 1243-1252 [Medline] [Order article via Infotrieve]
  14. Chaplin, M. F. (1986) in Carbohydrate Analysis: A Practical Approach (Chaplin, M. F., and Kennedy, J. F., eds) pp. 1-36, IRL Press, Oxford
  15. McGinnis, K., and Wilson, D. B. (1993) Biochemistry 32, 8151-8156 [Medline] [Order article via Infotrieve]
  16. Day, A. G., and Withers, S. W. (1986) Biochem. Cell Biol. 64, 914-922 [Medline] [Order article via Infotrieve]
  17. Meinke, A., Gilkes, N. R., Kilburn, D. G., Warren, R. A. J., and Miller, R. C., Jr. (1993) in Genetics, Biochemistry, and Ecology of Lignocellulose Degradation (Shimada, K., Ohmiya, K., Kobayashi, Y., Hoshino, S., Sakka, K., and Karita, S., eds) pp. 286-297, Uni Publishers, Tokyo

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.