Enzymatic Conversion of Biomass for Fuels Production - American

Chapter 4

Role of Cellulose-Binding Domain of Cellobiohydrolase I in Cellulose Hydrolysis T. R .

Downloaded by NORTH CAROLINA STATE UNIV on October 26, 2012 | http://pubs.acs.org Publication Date: October 7, 1994 | doi: 10.1021/bk-1994-0566.ch004

1

1

Donner ,

2

B . R . Evans , K . A .

2

Affholter ,

a n d J. W o o d w a r d

2,3

D e p a r t m e n t o f C h e m i s t r y , University of T u l s a , T u l s a , OK 74104 C h e m i c a l Technology D i v i s i o n , O a k Ridge N a t i o n a l Laboratory, O a k Ridge, TN 37831-6194

2

The kinetics of the cellulose-binding domain's ability to adsorb onto microcrystalline cellulose (Avicel) and its effects on the surface structure of the cellulose fibers were studied. The catalytic domain of Trichoderma reesei cellobiohydrolase I was rendered inactive by modification with a water-soluble carbodiimide. After modification, the cellobiohydrolase I possessed no ability to hydrolyze the model substrate p-nitrophenyl-β-D-cellobioside (PNPC). However, the modified cellobiohydrolase I was still capable of adsorbing onto microcrystalline cellulose. Scanning electron microscopy showed that whereas native C B H I smoothed the surface of cotton fibers, catalytically inactivated C B H I was without effect.

The cellulase enzyme complex secreted by the fungus Trichoderma reesei catalyzes the hydrolysis of insoluble crystalline substrates to glucose (1). These complexes can be separated by various chromatographic procedures into three major types of catalytic activity: cellobiohydrolase, endoglucanase, and 6-glucosidase. Approximately 80% of this cellulase complex is composed of two cellobiohydrolases, termed C B H I and II, of which C B H I comprises 60% (2). Small-angle X-ray scattering studies have shown that both cellobiohydrolase I and II are shaped like a tadpole (3,4) consisting of a cellulose-binding domain, or "tail" region, and a catalytic domain, or "head" region. The exact mechanism by which these domains hydrolyze insoluble cellulose fibers, including their effect on the microscopic structure of the insoluble cellulose, is unknown. However, it has been shown recently that the cellulose-binding domain of a bacterial endoglucanase plays a direct role in the disruption of the Ramie cellulose fiber structure during hydrolysis (5). This apparent disruption of Ramie cellulose fibers by the cellulose-binding domain supports the conclusion that the cellulose-binding domain's purpose in the degradation of insoluble cellulose may be two-fold. Not only does the cellulose-binding domain secure the enzyme to the •^Corresponding author 0097-6156/94/0566-0075S08.00/0 © 1994 American Chemical Society

In Enzymatic Conversion of Biomass for Fuels Production; Himmel, M., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1994.

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surface of cellulose fibers, but its apparent ability to disrupt the surface fibers may render crystalline celluloses more susceptible to hydrolysis. The "tail" region of cellobiohydrolase I possibly possesses a "wedge-like structure" (6), which aids it in physically prying apart the surface fibers of crystalline cellulose. It was postulated over 40 years ago by E l w y n Reese (7) that the Q factor of cellulases renders crystalline celluloses susceptible to hydrolysis. These recent data support the conclusion that the cellulose-binding domain could be the Q factor of cellulases. This study was performed to determine the physiological function of the cellulose-binding domain of cellobiohydrolase I from T. reesei. Materials and Methods Purification and Assay of Cellobiohydrolase I. The crude T. reesei cellulase preparation was obtained as a gift from Novo Nordisk Bioindustrials. C B H I was purified from this preparation by chromatofocusing as described previously (8). Fractions containing C B H I were further purified using a P D - 1 0 column with Sephadex equilibrated with 50 m M ammonium carbonate p H 5.0. The C B H I was then lyophilized. It was assayed as described previously using £-nitrophenyl-P-Dcellobioside ( P N P C ) as the substrate (9). Modification of Cellobiohydrolase I. Approximately 5.0 mg of C B H I was dissolved in 3.0 m L of nanopure water, p H 5.0. One-ethyl-3-(3dimethylaminopropyl)-carbodiimide ( E D C ) was added to the solution to a final concentration of 0.1 M . The solution was then incubated at room temperature (25°C) for 48 h, and the p H was maintained at 5.0 by titrating with 0.5 M HC1. A t intervals of time, the catalytic activity of the modified enzyme was tested using the model substrate P N P C . Quenching of the reaction occurred during the assay, which was conducted in 50 m M sodium acetate buffer, p H 5.0. A 25-pL of the reaction mixture containing approximately 40 pg protein was used in the assay mixture (0.5 m L total volume) for the determination of residual C B H I activity. Scanning Electron Microscopy (SEM). In separate test tubes, two batches of cotton fibers (1.0 mg) were incubated at 4 0 ° C in either 0.25 m L modified C B H I (2.38 mg/mL) or 0.06 m L native C B H I (10.56 mg/mL) for 72 h. Fifty m M sodium acetate, p H 5.0, was added to each reaction to a total reaction volume of 1.0 m L . Flat pieces of both native and modified enzyme-treated cotton fibers were mounted on separate aluminum stubs and coated at a 15°-angle with Pt-C. The samples were examined with an E T A C autoscan scanning electron microscope operated at 20 k V . Samples were observed at magnifications of 600 and 1200. Analytical Techniques. The protein concentration was determined using the Coomassie Blue reagent (Bio-Rad Laboratory) according to the method of Bradford using almond P-glucosidase as the standard. Catalytic activity of C B H I after incubation with E D C was determined at 5 5 ° C and p H 5.0 by measuring the hydrolysis of 20 m M P N P C at p H 5.0. The fluorescence spectra of both the native and modified


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Cellulose-Binding Domain of Cellobiohydrolase I

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enzyme were recorded using a Perkin-Elmer L S - 5 spectrofluorimeter. The samples were excited at 280 nm, and the fluorescence emitted was measured between 310 nm and 480 nm. Both the modified and native enzyme were tested for their ability to adsorb onto microcrystalline cellulose (Avicel PH-105, 10 mg/mL). For details, see reference 10. The absorbance at 280 nm was recorded for each sample before the addition of A v i c e l . After the incubation of C B H I with A v i c e l , the reactions were spun down in a centrifuge, and the supernatant was filtered through a 0.2-pm Acrodisc. Binding of the enzyme to A v i c e l was determined by measuring the absorbance of the supernatant at 280 nm and extrapolating the bound protein from that of the remaining free protein. Results and Discussion Inactivation of C B H The purified native C B H I was reacted with E D C for approximately 48 h. Throughout the reaction, the p H was adjusted with 0.5 M HC1 to keep the optimal p H for the reaction at 5.0. Complete inactivation of the enzyme was observed after 48 h, although >95% of activity was lost after 3 h (Figure 1). It should be noted that the modified enzyme possessed no catalytic activity at both p H 5.0 and 7.5. After modification, C B H I also lacked the ability to degrade barley P-glucan as determined by viscosity measurement (10), as well as P N P C . The reason for the inactivation of C B H I by E D C is apparently due to the modification of an essential carboxyl group within the catalytic site (77). The number of carboxyl groups in C B H I modified by reaction with E D C was not determined. Characterization of C B H I ^ h i * , and C B H I . Isoelectric focusing of the modified C B H I displayed a dramatic increase in its isoelectric point when compared to accepted values for the p i of the native enzyme (data not shown). This apparent alkalinity may be attributed to the conversion of the carboxyl groups of the native enzyme to the N-acylurea derivatives (72). Under mild, rather than denaturing, conditions only the most reactive carboxyl group sites w i l l be modified. The measured isoelectric point of the modified C B H I was found to be between p H 8.5 and 9.3, while that of the native C B H I remained consistent, with accepted values between p H 3.6 and 4.0. natjve

The fluorescence spectra of both the modified and native C B H I, excited at 280 nm, gave peaks of maximum emission at 340 nm. However, the peak at 340 n m of the modified C B H I was dramatically lower than that of native C B H I (Figure 2). This change in fluorescence may be due to a conformational change of the enzyme during the modification with E D C . S D S - P A G E analysis was used to determine the molecular weights of both modified and native C B H I. It was found that each enzyme had a molecular weight of approximately 67,000 g/mol, which was in basic agreement with the accepted value for T. reesei C B H I. Adsorption Isotherms. The binding of native and modified C B H I to A v i c e l as a function of protein concentration is shown in Figure 3. It is obvious that although the modified enzyme does bind to A v i c e l , it does so to a considerably lesser extent than native C B H I. The reason for this is unknown but may be related to the difference




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O R N L DWG 9 3 A - 2 0 1

0

320

340

360

380

400

420

440

460

480

W A V E L E N G T H O F EMISSION (nm)

F i g u r e 2.

Fluorescence spectra of native and chemically modified C B H I. Protein concentration 100 pg/mL.


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100

200

300

400

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PROTEIN ADDED (/^g

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The adsorption of native and modified C B H I to A v i c e l as a function of protein concentration.

in conformation that exists between the native and modified enzymes. Another explanation could be the possible reaction between E D C and tyrosine residues i n the cellulose-binding domain of C B H I (72). Tyrosines are important for binding of C B H I to cellulose (13). The concentration of adsorption sites on A v i c e l for C B H I was reported to be 70 pg/mg A v i c e l (14). The data shown here for native C B H I indicate that saturation of A v i c e l had not occurred. S E M Observations. Scanning electron micrographs of cotton linters that were untreated and treated with native or modified C B H I are shown i n Figures 4 and 5. The interpretation of these observations is that untreated cotton linters possess macrofibrils protruding from the surface of the fiber which are removed by the catalytic action of native C B H I. This results in the smoothing appearance of the fiber surface and is similar to the observations made by D i n et al. (5) when the Ramie cellulose fibers were treated with the catalytic domain of an endoglucanase from Cellulomonas firm. N o such action takes place resulting from the incubation of modified C B H I on the cotton linters. There is also no evidence for any disruption of the fiber surface by either the native or modified enzyme. Conclusions The chemical modification of T. reesei C B H I by E D C results i n a loss in its catalytic activity. The enzyme thus modified had no obvious effect on the appearance of cotton linters. The conclusion that the cellulose-binding domain of C B H I has no physical effect on the surface of cotton cellulosic fibers is tempered by the finding that modification also resulted in a change in the conformation of the enzyme and also in the reduced binding of the enzyme to cellulose.



DONNER ET AL.

Cellulose-Binding

Domain of Cellobiohydrolase

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Acknowledgments The authors thank Donna Reichle for editorial assistance and Debbie Weaver for secretarial assistance. This work was supported by the Chemical Sciences Division, Office of Basic Energy Sciences, U . S . Department of Energy, under contract D E A C 0 5 - 8 4 O R 2 1 4 0 0 with Martin Marietta Energy Systems, Inc., and the National Renewable Energy Laboratory.


Literature Cited

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

Wood, T.M. Biochem. Soc. Trans. 1992, 20, 46-53. Goyal, A.; Ghosh, B.; Eveleigh, D. Bioresource Technol. 1991, 36, 37-50. Schmuck, M.; Pilz, I.; Hayn, M . ; Esterbauer, H . Biotechnol. Lett. 1986, 8, 397402. Abuja, P.M.; Pilz, I.; Claeyssens, M.; Tomme, P. Biochem. Biophys. Res. Comm. 1988, 156, 180-185. Din, N . ; Gilkes, N.R.; Tekant, B.; Miller, R.C. Jr.; Warren, A.J.; Kilburn, D.G. BioTechnology 1991, 9, 1096-1099. Kraulis, P.J.; Clore, G.M.; Nilges, M . ; Jones, T.A.; Peterson, G.; Knowles, J.; Gronenborn, A . M . Biochemistry 1989, 28, 7241-7257. Reese, E.T.; Siu, R.G.H.; Levinson, H.S. J. Bacteriol. 1950, 59, 485-497. Offord, D.A.; Lee, N.E.; Woodward, J. Appl. Biochem. Biotechnol. 1991, 28/29, 377-386. Woodward, J.; Lee, N.E.; Carmichael, J.S.; McNair, S.L.; Wichert, J . M . Biochim. Biophys. Acta 1990, 1037, 81-85. Woodward, J.; Affholter, K.A.; Noles, K.K.; Troy, N.T.; Gaslightwala, S.F. Enzyme Microb. Technol. 1992, 14, 625-630. Tomme, P.; Claeyssens, M . FEBS Lett. 1989, 243, 239-243. Means, G.E.; Feeney, R.E. In Chemical Modification of Proteins; Holden-Day: San Francisco, C A , 1971. Teeri, T.T.; Reinikainen, T.; Ruohonen, L.; Jones, T.A.; Knowles, J.K.C. J. Biotechnol. 1992, 24, 169-176. Tomme, P.; Heriban, V.; Claeyssens, M . Biotechnol. Lett. 1990, 12, 525-530.

R E C E I V E D M a r c h 1, 1994


Enzymatic Conversion of Biomass for Fuels Production - American

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