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We wish to acknowledge Dr. K. A. Malik, DirectorNIBGE, for providing ... We are thankful to Mr. Munir Malik for typing the manuscript, Tanvir Ahmad for his help in ...
Folia Microbiol. 41 (4), 341-346 (1996)

The Stability of Extracellular [3-Glucosidase from Aspergillus niger Is Significantly Enhanced by Non-covalently Attached Polysaccharides M.H. RASHID and K.S. SIDDIQUI* National Institute for Biotechnology & Genetic Engineering~ P.O. Box 577, Faisalabad, Pakistan Received November 29, 1995 Revised version May 29, 1996

ABSTRACT. The removal of noncovalently bound polysaccharide coating from the extracellular enzymes of Aspergillus niger, by the technique of compartmental eleetrophoresis, had a very dramatic effect on the stability of 13-glucosidase. The polysaceharide-~-glucosidase complex was extremely resistant to proteinases and far more stable against urea and temperature as compared with polysaccharide-free ~giucosidase. The I$-glucosidase-polysaccharide complex was 18-, 36-, 40- and 82-fold more stable against chymotrypsin, 3 mol/L urea, total thermal denaturation and irreversible thermal denaturation, respectively, as compared with polysaccharide-free Igglucosidase. The activation energy of polysaccharide-complexed 13-glucosidase (55 kJ/mol) was lower than polysaccharide-free enzyme (61 kJ/moi), indicating a slight activation of the enzyme by the polysaccharide. No significant difference could be detected in the specificity constant (V/Km) for 4-nitrophenyi ~-D-glucopyranoside between polysaccharide-free and polysaccharide-complexed 15-glucosidase. We suggest that the function of these polysaccharides secreted by fungi including A. niger might be to protect the extraceUular enzymes from proteolytic degradation, hence increasing their life span.

[~-Glucosidase (EC 3.2.1.21) is a component of the cellulase system which converts cellobiose to glucose (Gokhale et al. 1984). Our previous results showed that polysaccharides secreted into the medium byA. niger form noncovalent complexes with the enzymes (Rashid and Siddiqui 1994); therefore, these polysaccharides must be removed from the enzymes before 13-glucosidase purification, characterization and antibody production against any enzyme is undertaken (Lindner 1988; Kamphuis et al. 1992; Evans et al. 1991). These extracellular polysaccharides (EPS) were nondestructively removed from all the extracellular enzymes of A. niger, including 13-glucosidase by our technique of compartmental electrophoresis (Siddiqui et al. 1994). In this paper we report a detailed effect of proteinases, urea, temperature and substrate variation on the 13-glucosidase-polysaccharide complex and polysaccharide-free ~-glucosidase.

MATERIALS AND METHODS All chemicals were purchased from Sigma Chemical Company (USA) and were of molecularbiology grade. Cultivation of A. niger and fl-glucosidase-polysacchatide extraction. Aspergillus niger was grown in shake flasks at 30 *C for 14 d using Vogal's medium containing 2 % (W/V) wheat bran and untreated kallar grass (Leptochloa fusca) in a 1 : 1 ratio as carbon sources as reported earlier (Gokhale et al. 1984, 1988). Mycelia were removed by centrifugation and the clear supernatant was concentrated 8-fold in a Amicon concentrator. After concentrating the sample, the volume was made to 2 L with distilled water and the sample was concentrated again. This step was repeated twice to dialyze out the salts. The dialyzed supernatant was then freeze-dried. Removal of polysaccharides. Non-covalently bound polysaccharides were removed from the enzymes including ~-glucosidase by redissolving half of the freeze dried powder in distilled water and subjecting it to compartmental electrophoresis as described before (Siddiqui et al. 1994). After compartmental electrophoresis, the sample from the anodic compartment containing polysaccharide free ~-glucosidase was again dialyzed and freeze-dried. The complete removal of polysaccharides from 13-glucosidase was checked as described (Rashid and Siddiqui 1994; Siddiqui et al. 1994). Stock solution for stability studies. Five mg of freeze-dried material containing 13-glucosidasepolysaccharide complex (0.9 mg protein) and polysaccharide free ~-glucosidase (1.2 mg protein) were

*Corresponding author.

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separately dissolved in 5 mL of 25 mmol/L MOPS-KOH (pH 7) buffer. Total protein was estimated by the dye-binding method (Bradford 1976). Effect of proteinases. Effect of proteinases on I$-glucosidases was determined as described before for xylose isomerase (Siddiqui et al. 1993) with the following modification. Chymotrypsin or subtilisin (0.2 mg/mL) were added to each of the test tubes containing ~-glucosidase-polysaccharide complex or polysaccharide-free 13-glucosidase and incubated at 30 ~ Aliquots (20-100 ~tL) were withdrawn at different time intervals and immediately assayed for ~-glucosidase activity at 30 ~ (Wood and Bhat 1988). Effect of temperature. Melting temperatures of the i$-glucosidase-polysaccharide complex and polysaccharide free ~-glucosidase were determined as described for xylose isomerase (Rangarajan et al. 1992) with the following modification. Aliquots (200 ~tL) were taken in Eppendorf tubes and incubated at different temperatures for 5 min. After 5 min the tubes were cooled in ice for at least 30 min before assaying for 13-glucosidase activity at 30 ~ Irreversible thermal inactivation of the 13-glucosidase-polysaccharide complex and polysaccharide-free 13-glucosidase were determined by incubating both enzyme solutions at 40 ~ and then different time-course aliquots were withdrawn, cooled on ice for 30 min and then assayed for ~-glucosidase activity at 40 ~ Total thermal denaturation of l~-glucosidase-polysaccharide complex and polysaccharide-free ~-glucosidase were measured by incubating both enzyme solutions at 40 ~ and then the different timecourse aliquots were immediately assayed at 40 ~ for 13-glucosidase. Activation energy. Both types of I$-glucosidases were assayed in 50 mmol/L sodium acetateacetic acid buffer (pH 5) at different temperatures ranging from 10 to 99 ~ (Sanyal et al. 1988). The rate of reaction of polysaccharide-complexed l~-glucosidase is compared with polysaccharide-free 13-glucosidase as follows: log ( k E P s / k ) = (Ea -

Ea,EPS)/2.303 R T

(Eq. 1)

where k is the rate without polysaccharide, kEPS rate with polysaccharide, Ea activation energy, R gas constant and T absolute temperature. Effect of urea. Solid urea was added to 10 mmol/L Tris-HCl (pH 7) buffer to a final concentration of 3 mol/L. Five mg of the freeze-dried I$-glucosidase-polysaccharide complex and polysaccharide-free 13-glucosidase were added to the 3 mol/L urea buffer and incubated at 10 ~ Aliquots (20-100 laL) were withdrawn at different time intervals and assayed immediately for 13-glucosidase activity at 30 ~ All graphs except that involving subtilisin and melting temperatures were fitted to first-order plots by using a 'Sigma plot' computer programme. Measurement of specificity constant (V/Km). ~-Glucosidase-polysaccharide complex and polysaccharide-free ~-glucosidase were assayed in 50 mmol/L sodium acetate buffer (pH 5.5) containing variable amounts of 4-nitrophenyl ~-D-glucopyranoside. The data were plotted according to Lineweaver and Burk as described by Price and Stevens (1982) using a 'Lotus 123 release 4' computer programme.

RESULTS A N D DISCUSSION

In our previous paper (Siddiqui et al. 1994) we observed that the removal of noncovalently bound polysaccharides from 13-glucosidase produced extracellularly byA. niger significantly reduced the shelf life of the enzyme at 4 ~ In the past not many workers have studied the effect of noncovalently attached polysaccharides on enzyme stability. We have now studied the stability of the 13-glucosidasepolysaccharide complex and polysaccharide-free l~-glucosidase and found that noncovalent attachment of extraceUular polysaccharides confers on the enzyme a very significant protection against chymotrypsin (Fig. 1A), subtilisin Carlsberg (Fig. 2), 3 mol/L urea (Fig. 1B), irreversible thermal denaturation (Fig. 1C) and total thermal denaturation (Fig. 1D). There was a difference of 21 ~ in the melting temperatures of polysaccharide-complexed and polysaccharide-free ~-glucosidase (Fig. 3). In the case of subtilisin Carlsberg which is a proteinase of broad specificity, 25 and 75 % of initial enzyme activity was lost in the case of the ~-glucosidase-polysaccharide complex and polysaccharide-free l~-glucosidase, respectively, in less than 10 rain. After 10 rain there was no further decrease in enzyme activity. This was the reason that in case of subtilisin treatment the values could not be fitted to first-order plots whereas the effect of chymotrypsin (Fig. 1A), 3 mol/L urea (Fig. 1B), irreversible (Fig. 1C) and total thermal denaturation on polysaccharide-eomplexed and free l~-glucosidase (Fig. 1D) followed first-

STABILITY AND ACTIVITY OF ~-GLUCOSIDASES FROM A. niger 343

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order kinetics. The calculated values of first-order rate constants (k) and the corresponding half-lives are given in Table I. Noncovalent attachment of polysaccharides gave J3-glucosidase 18-, 36-, 82- and 40-fold more protection against chymotrypsin, 3 mol/L urea, irreversible thermal denaturation and total thermal denaturation, respectively. The removal of covalently bound saccharide (deglycosylation) from cellulases ofHumicola insolens strain YH-8 resulted in a significant decrease in their thermal stability (Hayashida and Yoshioka 1980). Whereas the removal of noncovalently bound polysaccharides from invertase of Sphacelia sorghi also resulted in a decrease in its thermal stability (Dickerson and Baker 1979). 5

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Fig. lPa First-order plots of the effect of chymotrypsin on enzyme activity (%) of the ~glucosidasepolysaccharide complex (closed symbols) and polysaccharide-free 13-gluco6idase (open symbols). Both samples were incubated at 30 ~ in 25 mmol/L MOPS-KOH (pH 7) buffer and the aliquots were assayed immediately at 30 ~ for ~glucosidase in 50 mmol/L sodium acetate (pH 5) buffer. B: First--order plots of the effect of 3 mol/L urea. Both samples were incubated at 10 ~ in 10 mmol/L Tris-HC! (pH 7) buffer containing 3 mol/L urea and the aliquots withdrawn at different time intervals were immediately assayed at 30 ~ for I~-gluco~idase activity. C: First-order plots of the effect of irreversible thermal denaturation. Both samples were incubated at 40 ~ in 25 mmol/L MOPS-KOH (pH 7) buffer and the aliquots withdrawn at different time intervals were cooled in ice for 30 rain before assaying for 13-glucosidase activity at 40 ~ Ik. First-order plots of the effect of total thermal denaturation. Both samples were incubated at 40 ~ in 25 mmol/L MOPS-KOH (pH 7) buffer and the aliquots withdrawn at different time intervals were immediately assayed for ~glucosidase at 40 ~

It has been argued that these fungal EPS could act as energy storage material under unfavorable conditions (Seviour et al. 1992; Dickerson et al. 1970). Another function of these fungal EPS includes protection of cells against water stress (Willets 1971). Based on our results we suggest that an additional and very important function of these EPS is to protect the extracellular enzymes from degradation, hence extending their life span. Polysaccharide-free j3-glucosidases have similar half-lives of 1.5 h, both in the case of total and irreversible thermal denaturation (Table I). In the case of polysaccharide-complexed 13-glucosidase, the half-life of irreversibly denatured enzyme is twice that of totally denatured enzyme. This could indicate that with irreversible thermal denaturation the polysaccharide either does not allow the enzyme to unfold at all or the polysaccharide acts as a template which helps the reversibly denatured enzyme to quickly refold when the sample is cooled in ice before assaying. If we analyze the activation energies (Ea) of both enzymes in the temperature range of 62-99 ~ (Fig. 5, after inflection point), we fmd that

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polysaccharide-complexed l~-glucosidase has a lower Ea (69 kJ/mol) than the polysaccharide-free enzyme (79 kJ/mol). This means that polysaccharide-complexed enzyme catalyzes the reaction 37 times more efficiently than polysaecharide-free enzyme, due to its more rapid thermal denaturation in this temperature range (calculated by Eq. 1).

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Fig. 3. Melting temperatures (~ of the ~glucosidase-polysaccharide complex (closed symbols) and polysaccharide free 13-glucosidase (open symbols). Both samples were incubated at different temperatures for 5 min, cooled in ice for 30 min and assayed for 13-glucosidase activity (%) at 30 ~ in 50 mmol/L sodium acetate (pH 5) buffer. The melting temperature is the temperature at which 50 % enzyme activity was lost.

Fig. 2. Effect of subtilisin Carlsberg on the enzyme activity (%) of ~-glucosidase-polysaccharide complex (closed symbols) and polysaccharide free 13-glucosidase (open symbols). Both samples were incubated at 30~ in 25 mmol/L MOPS-KOH (pH 7) buffer and aliquots were withdrawn at different time intervals to be assayed immediately for 13-glucosidase at 30~ in 50 mmol/L sodium acetate (pH 5) buffer.

Table I. Effect of chymotrypsin, urea and temperature treatment on the first-order rate constants (k) and half-lives (tl/z) of polysaccharide complexed and polysaccharide-free ~glucosidase activitya

Polysaccharide-complexed ~glucosidase

Polysaccharide-free ]3-glucosidase

Treatment

Chymotrypsin Urea, 3 mol/L Temperature (irreversible denaturation) Temperature (total denaturation)

10 -4 • k 1/min

tlA h

10 -3 • k 1/min

tlA h

1.0 1.4 0.9

115_5 82_5 128.3

1.78 5.09 7.38

6_50 2.20 1_56

1.9

60.8

7.57

1.51

ak = (-slope) taken from the respective graphs of In % enzyme activity vs. time. ttA = In 2/k.

We found that the specificity constant (V/Km) of the l$-glucosidase-polysaccharide complex (1.3) and polysaccharide free 13-glucosidase (1.6) were not significantly different (Fig. 4), implying that polysaccharide did not hinder the movement of substrate (4-nitrophenyl ]3-D-glucopyranoside) to the active site of the enzyme. The removal of polysaccharides from 13-glucosidase of Coriolus versicolor resulted in a 20-fold activation of the enzyme while its effect on the stability of the enzyme was not studied (Gallagher and Evans 1990). On the other hand, in the case of 13-glucosidase from Trichoderma reesei the noncovalently attached polysaccharides activated the enzyme (Messner et al. 1990). These workers also did not study the stability of the ]3-glucosidase-polysaccharide complex. Analysis of the activation energy profile of both enzymes in the temperature range of 10-62 ~ (Fig. 5, before the inflection point) revealed that polysaccharide-complexed 13-glucosidase has a lower activation energy (55 kJ/mol) than the polysac-

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STABILITY AND ACTIVITY OF ~-GLUCOSIDASES FROM A. niger 345

charide-free enzyme (61 kJ/mol). This indicated a slight activation of the enzyme by the noncovalently attached polysaccharides. The activation energy of I~-g!ucosidase from A. japonicus was found to be 35 kJ/mol (Sanyal et al. 1988). Analysis of the activation energy profde has the advantage that activity and stability of two forms of the same enzyme could be determined simultaneously as discussed above. I

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Fig. 4. Lineweaver-Burk plot for fl-glucosidase-polysaccharidecomplex (closed symbols) and polysaceharide free ~-glueosidase (open symbols). Both samples were assayed at 30 ~ in 50 mmol/L sodium acetate (pH 5) buffer containing variable amounts of 4-nitrophenyl ~D-glucopyranosideas substrate. The intercept on the y-axis corresponds to I/V, the intercept on the x-axis to -1/Kin; V is seen to be 3.35 nmol mL-1 min -1, Km then 2.62 mmol/L for the I$-glucosidase-~polysaccharidecomplex and 7.70 and 4.65, respectively,for polysaccharide-free~glucosidase. Work on xylose isomerase I showed that thermal unfolding begins InV /. from the enzyme terminus or a surface loop. Such flexible loops or floppy regions are the weakest link in the thermal 3 unfolding pathway and could be probed and identified by their susceptibility to 8 2 proteinases as demonstrated in the case of xylose isomerase (Siddiqui et al. 1993). 1 The neutralization of acidic groups of 0 I xylose isomerase by glycinamide and car2.8 3.1 3.6 bodiimide resulted in increased flexibility 103KIT of C-terminal loop because of breakage of salt bridges. When this modified Fig. 5. Arrhenius plot for the determination of activation energy of xylose isomerase was probed by ther- hydrolysis of p-NPG by fl-glucosidase; closed symbols: polysaccharidemolysin the rate of proteolytic nicking complexed 13-glucosidase,open symbols: polysaccharide-free 13-glucoswas found to be four times that of native idase. enzyme (Siddiqui et al. 1993). Moreover, the melting temperatures of modified xylose isomerase was 14"C less than with the native enzyme (Siddiqni 1990). Similarly, Sauer and his co-workers generated mutants of DNA-binding proteins, 6-cro and 6-repressor (Sauer et al. 1986; Parsell and Saner 1989; Pakula and Sauer 1989). They also found that mutants that were more proteinase susceptible were also less thermostable. In our present study a similar correlation between proteinase susceptibility and thermal inactivation emerged. Removal of noncovalently bound polysaccharide covering from 13-glucosidase renders surface loops more floppy, consequently making the enzyme more susceptible to narrow and broad specificity proteinases. This resulted in a concomitant decrease in the thermostability of polysaccharide-free ~-glucosidase. In the past workers have tried to correlate different parameters, such as surface area, buried in the folded state (Stellwagen and Wilgus 1978), hydrophobie index (Merkler r al. 1981) and the number of hydrogen bonds and salt bridges (Walker et al. 1980) with thermal protein unfolding but no clear rules have emerged. Based on all the above results, it is apparent that there exists a negative correlation between proteinase susceptibility and thermostability. We propose, therefore, that proteins could possibly be

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made more thermostable by making them more proteinase-resistant because it is much easier to identify the flexible region, which controls thermal unfolding, by proteolytic nicking (Siddiqui et al. 1993) and subsequently locking the loop to make it less mobile. The work described is part of the PhD research of M. H. Rashid. This work was financed in part by a grant made by the United States Agency for International Development under PSTC proposal 6-163, USAID grant no. 9365542-G00-89-42-00 and PAEC. We wish to acknowledge Dr. K. A. Malik, DirectorNIBGE, for providing research facilities. We would like to thank M. H. Rashid's External Supervisor, Prof. A. IL Shakoori and Dr. M. I. Rajoka, HeadBiofuel Group for their valuable suggestions. We are thankful to Mr. Munir Malik for typing the manuscript, Tanvir Ahmad for his help in computer graphics. The technical assistance of G. A. Waseer is appreciated.

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