Two forms of the glucoamylase of Aspergillus niger

ARCHIVES

OF

BIOCHEMISTRY

Two

Forms

AND

of the

Glucoamylase

D. R. LINEBACK, Department

of Biochemistry Received

134, 539653

BIOPHYSICS

of Aspergillus

I. J. RUSSELL,

and Nutrition, June

(1969)

University

19, 1969; accepted

AND

of

C. RASMUSSEN

Nebraska,

August

nige:

Lincoln,

Nebraska

68609

11, 1969

Two forms of the glucoamylase (or-1,4-glucan glucohydrolase, E.C.3.2.1.3) of Asgillus niger have been purified by chromatography on DEAE-cellulose ion-exchange columns. The purified enzymes possessed a high degree of purity as indicated by paper electrophoresis, sedimentation velocity, and disc-gel electrophoresis. The two enzymes had the same pH optima when starch was used as the substrate, temperature stability at elevated temperatures, action patterns on malto-oligosaccharides, antigenicity, and NHX-terminal amino acid. They differed in electrophoretic mobility, isoelectric point, and, to a small degree, in stability at room temperature for prolonged periods. Glucoamylase I was not dissociable into subunits in the presence of urea, acid, or flmercaptoethanol under the conditions studied. Both forms of the enzyme were present in the crude glucoamylase preparation and did not arise as artifacts of the purification procedure.

The glucoamylase ((Y-1,4-glucan glucohydrolase, E.C.3.2.1.3) of Aspergillus niger consists of two forms (1, 2) which have been isolated and purified by chromatography on DEAE-cellulose ion-exchange columns (1). One of the enzymes (glucoamylase I)” has been partially characterized (1, 4, 5). However, the second enzyme (glucoamylase II)3 has not been studied in detail. Both enzymes have been reported to be glycoproteins containing n-mannose, D-glucose and Dgalactose (6). The carbohydrate moieties appear to be 0-glycosidically linked through n-mannose to hydroxyl groups of serine and threonine in the polypeptide chain (7). 1 This work is published, with the approval of the Director, as paper No. 2616 Journal Series, Nebraska Agricultural Experiment Station. A portion of this work has been presented before the Division of Biological Chemistry at the 150th National Meeting of the American Chemical Society, Atlantic City, New Jersey, September 1965. 2 Present address: Department of Grain Science and Industry, Kansas State University, Manhattan, Kansas 66502. 3 The nomenclature used for the two forms of glucoamylase is that recommended by Webb (3).

Preliminary to a more detailed characterization of the glycoprotein structure of the two forms of the glucoamylase from A. niger, it was of interest to establish a more refined purification procedure and to determine extensively the physical properties of these enzymes. The purification of both glucoamylases has been re-investigated and a number of properties of the purified enzymes have been determined. MATERIALS

AND

METHODS

Materials. A generous supply of Takamine Diazyme concentrate was provided by the Miles Chemical Co., Elkhart, Indiana, from whom the glucose oxidase reagent strips (Clinisticks) were also obtained. DEAE-cellulose (reagent grade, 0.9 meq/g) was purchased from Brown Co., Berlin, New Hampshire. Light green SF yellowish dye was obtained from Fischer Scientific Co., Fairlawn, New Jersey, while Dyes K-150 and K-160 were purchased from Kensington Scientific Corp., Berkeley, California. The n-glucose oxidase reagent (Glucostat) was a product of the Worthington Biochemical Corp., Freehold, New Jersey. Coomassie Brilliant Blue R 250 dye was purchased from Colab Laboratories, Inc., Chicago Heights, Illinois. The p-nitrophenyl glycosides were ob539

540

LINEBACK,

RUSSELL,

tained from Pierce Chemical Co., Rockford, Illinois. Chromatographic procedures. DEAE-cellulose, which had been wetted by standing overnight in water, was resuspended and allowed to stand for several minutes. The supernatant fluid and fine particles were removed through a glass tube with suction. This procedure was repeated approximately six times. The resin was then filtered and washedsuccessively in a Buchner funnel with 0.5 M HCl, distilled water, 0.5 M NaOH, and water. The resin was filtered after each washing and fine particles removed as described above if the filtration rate decreased significantly. After the final washing, fine particles were again removed by several treatments. The DEAE-cellulose was filtered, and the resulting pad suspended in 0.025 M citratephosphate buffer of the desired pH. The mixture was equilibrated by stirring and then adjusted to the desired st,arting pH with 1 M citric acid. Columns (3.5 X 45 cm) were prepared by pouring the stirred solution into the columns and allowing the resin to settle to the desired height. The columns were washed with approximately 500 ml of fresh buffer and equilibrated overnight at 4” prior to use. All chromatography was performed at 4” with flow rates of 1 ml/min maintained with a peristaltic pump. Fractions (15 ml) were automatically collected. Protein components were located in the effluent fractions by determining the absorbance at 280 nm. Enzyme assays. Glucoamylase solutions were diluted with 0.05 M acetate buffer, pH 4.8, to a concentration of 80-160 pg protein per ml. Activities were determined by incubating 0.5 ml aliquots of the diluted enzyme with 1.5 ml of 4% starch (Lintner soluble) solution in 0.05 M acetate buffer, pH 4.8, for 1 hr at 30”. The reaction w&s terminated by the addition of 5.0 ml of absolute ethanol, and the precipitated starch and protein were removed by centrifugation. When necessary, the supernatant solution was diluted with 70y0 ethanol to give 60-180 pg glucose per ml prior to analysis. Glucose was determined in the supernatant solutions using a n-glucose oxidase preparation (Glucostat). A portion (0.5 ml) of the starch hydrolyzate was incubated with 5.0 ml of the enzyme-chromophore solution for 30 min at 30”. The reaction was terminated by the addition of 2 drops of 4 M HCl, and the absorbance was determined at 400 nm. A standard curve was prepared using 20-100 pg glucose. One unit of glucoamylase activity was defined as the amount of enzyme required to liberate 1 Imole of glucose per minute under the reaction conditions defined above. Specific activity was expressed as units per mg protein, determined by the Lowry method (8).

AND

RASMUSSEN

Because the specific activity of the crude euzyme was the sum of more than one starch-hydrolyzing enzyme present in the starting material, the specific activity of glucoamylase I and II in the crude enzyme solution was determined in the following manner. The crude enzyme was fractionated by the stepwise elution procedure described below. Six fractions, which encompassed the entire elution volume, were collected from this chromatography, and their volumes were determined. These six fractions were composed of tubes 2&90, 91-115, 116-172, 173-189, 190-244, and 245275 in Fig. 1. Protein analysis of the fractions by the Lowry method (8) revealed that 87y0 of the protein placed on the column was recovered. Aliquots of these fractions were then heated in boiling water for 1 hr to inactivate the enzymes. To determine the specific activity of any fraction, the unheated fraction was mixed with the other heated aliquots in amounts proportionate to their elution volumes. The resulting assay solutions were composed of the same relative proportions of each protein found in the crude starting material. The resulting mixtures were assayed as described above for enzymatic activity and total protein. Thus, the specific activity of any reconstituted component reflects only the activity of one of the starch-hydrolyzing enzymes. The presence of other glycosidase activities in the fractions obtained from the gradient elution purification was assayed by incubating 0.5 ml of the enzyme solution, diluted 40-fold, with 2.0 ml of an approximately 2 mM aqueous solution of the appropriate p-nitrophenyl glycoside for 2 hr at 30”. The solution was then made alkaline by adding 2.0 ml of 0.2 M sodium carbonate and the absorbance determined at 420 nm. The substrates used were the a-n-glucoside, P-o-glucoside, WDgalactoside, o-n-galactoside, a-o-mannoside, and 2-acetamido-2-deoxy-@-n-glucoside derivatives of p-nitrophenol. The presence of glucosyl transferase activity was assayed by incubating 0.1 ml of the enzyme solution with 0.1 ml of 10% maltose in 0.1 M acetate buffer, pH 4.8, at room temperature. Aliquots of the solutions were spotted on Eaton-Dikeman 613 chromatography paper after 0, 1,2,4,8, and 24 hours of incubation. The papers were developed by three ascents of a 1-butanol:pyridine:water (6:4:3 v/v/v) solvent system (9), and the carbohydrate areas located by the silver nitrate dip reagent (10). The presence of isomaltose or panose was indicative of glucosyl transferaae activity. Enzyme purijkation. Takamine Diazyme from Aspergillus niger was the source of the glucoamylase. This enzyme product, prepared from a culture filtrate of d. niger by an alcohol precipitation

GLUCOAMYLASE

OF

procedure, was supplied as a tan powder which was t,he starting material. The glucoamylases were separated from other constituents of the Diasyme by chromatography on a DEAE-cellulose ion-exchange column using the stepwise elution procedure reported by Pazur and Ando (1). To insure adsorption of the glucoamylases under the starting conditions, the columns were prepared in 0.025 M citrate-phosphate buffer of the desired starting pH, and the enzyme was adsorbed onto the DEAE-cellulose in buffers of the same ionic strength and pH. After adsorption onto the DEAE-cellulose, the constituents of the crude enzyme preparation were eluted by the successive addition of approximately 500 ml of 0.025 M citrate-phosphate buffer, pH 8.0, 500 ml of 0.05 M citrate-phosphate buffer, pH 8.0, 750 ml of the same buffer, pH 6.0, and 1260 ml of the same buffer, pH 4.0. Each buffer was added when 5(tlOO ml of the previous buffer remained in the chromatographic column. The eluates that contained glucoamylase I were combined and rechromatographed (1). Glucoamylase I was adsorbed onto the DEAF,cellulose at pH 6.0 and was recovered by successive addition of 500 ml of 0.05 M citrate-phosphate buffer, pH 6.0, and sufficient amounts of the same buffer, pH 4.0, to elute the enzyme. The eluates containing glucoamylase II were combined and further purified by chromatography on a DEAEcellulose column. Adsorption of this enzyme was effected from 0.025 M citrate-phosphate buffer, pH 8.0, and elution was performed in a manner analogous to that described for glucoamylase I using 0.05 M citrate-phosphat,e buffers of pH 8.0 and 6.0. Prior t,o rechromatography, both glucoamylases were dialyzed against running tap water and adjusted to the desired ionic strength and pH using a concentrated (0.2 M) buffer solution. An alternative procedure for purification of the enzyme involved elution of the proteins from the DEAE-cellulose column with a linear gradient of decreasing pH. The crude enzyme was adsorbed onto the DEAE-cellulose at pH 8.0 in 0.025 M citrate-phosphate buffer, and the column was eluted wit,h 300 ml of 0.035 M buffer, pH 8.0. The proteins were then eluted with a linear gradient of decreasing pH using 2 liters of 0.05 M citrate-phosphate buffer, pH 8.0, in the mixing chamber and 2 liters of the same buffer, pH 3.2, in the reservoir. Rechromatography of glucoamylase I was performed using 1 liter of 0.05 M citrate-phosphate buffer, pH 6.0, in the mixing chamber and 1 liter of the same buffer, pH 3.0, in the reservoir. Glucoamylase II was rechromatographed using 1 liter of 0.05 M citrate-phosphate buffer, pH 8.0, in the mixing chamber and 1 liter of the same buffer, pH 5.0, in the reservoir.

ASPERGILLUS

NIGER

541

The protein concentration of the purified solutions was determined by the Lowry method (8) using bovine serum albumin as a standard. Electrophoresis. A Spinco Model IZ elect.rophoresis cell was used to determine electrophoretic migration of the crude and purified glucoamylases. Samples of 0.01+.03 ml (lo-30 pg protein) of the enzyme solutions were applied to the paper strips. Electrophoresis was performed in 0.1 M phosphate buffer, pH 7.6, for 18 hr at 120 V and a current of 16-15 mA. Starch-hydrolyzing enzymes were located on the paper strips in the manner reported by Pazur and Okada (11). After the strips were dried, the protein components were located by staining them with light green SF yellowish (12). Disc-gel electrophoresis was performed using the method of Davis (13) modified in the following manner. The gel system was that of Williams and Reisfeld (14) using a Tris-EDTA-boric acid buffer (15) (2.0 g:O.2 g:5.5 g, respectively, diluted to 1 liter), pH 7.0, in the electrode chambers. Gels were prepared with acrylamide concentrations of 4 and 7%. The polymerized running gels contained 10% glycerol, and glycerol was substituted for sucrose in the stacking and sample gels. Protein samples (50-100 pg) were polymerized into sample gels, and electrophoresis was performed for 90 min at 5-10” using a current of 3 mA per tube. The gels were removed from the tubes and fixed for 30 min in 12.5yc trichloroacetic acid. Protein components in the gels were stained with Coomassie brilliant blue for 1 hr using the procedure of Chrambach el al. (16). The relative intensities of the protein bands in the gels after staining were determined by measuring the absorbance of the gel at 570 nm using a Photovolt Densicord electrophoresis recording densitometer. Glucoamylase activity in the gels was determined by slicing gels, which had not been treated with trichloroacetic acid, into 0.3-0.5 mm sections and assaying the sections for starch-hydrolyzing activity. Each section was incubated in 1.0 ml of 0.05 M acetate buffer, pH 4.8, for 2 hr. Aliquots (0.1 ml) of the buffer solutions were then incubated with 0.3 ml of 4% starch for 1 hr at 30”. The reaction was terminated by the addition of 1.0 ml of absolute ethanol. Precipitated starch was removed by centrifugation, and aliquots of the supernatant solution were analyzed for glucose by the glucose oxidase procedure described previously. Purified samples of glucoamylase I and II (0.025 ml, 2550 rg of protein) were subjected to paper electrophoresis in 0.05 M citrate buffer at pH values of 3.0, 3.2, 3.4, 3.6, 3.8, 4.0, and 4.2. Electrophoresis was performed for 18 hr at, 120 V and a current of 2-4 mA. Protein components were located as previously described. Dyes K-150 and

542

LINEBACK,

RUSSELL.

K-160 were used as internal standards to determine the amount of electrodiffusion occurring during the experiments. Sedimentation velocity measurements. Purified enzyme solutions were concentrated to approximately O&lye protein by pervaporation at 4” and dialyzed against 0.1 M NaCl. Sedimentation velocity experiments were conducted on each of the purified enzymes at a constant temperature of 20” with a rotor speed of 59,780 rpm using a Spinco Model E analytical ultracentrifuge. Five photographs were obtained for each experiment at time intervals ranging from 16 to 100 min. The homogeneity of sedimentation of the purified enzyme solutions was evaluated by a detailed boundary analysis of the sedimentation velocity experiments. The apparent diffusion coefficient was calculated from each pict,ure obtained during the experiment by the height area method and plotted versus time (17). pH Optima. The glucoamylase activity of the purified enzymes was determined using 47, starch substrate at various pH values. The enzyme solution was adjusted to the same pH as the starch substrate prior to incubation. Starch solutions (4%) were prepared in 0.1 M glycine-HCI buffer (pH 2.2 and 2.6), 0.1 M citrate buffer (pH 3.4, 4.2, 4.6, 4.8, 5.2, and 5.8), and 0.1 M phosphate buffer (pH 6.5, 7.0, and 7.5). Starch-hydrolyzing activity was assayed at each pH value in the manner previously described. Temperature stability. The stabilities of glucoamylase I and II were determined at 30” and at elevated temperatures. The enzymes, in 0.05 M citrate-phosphate buffer, pH 4.8, were maintained at 30” and 0.5 ml aliquots were removed at 0, 20, 42, 90, 168, and 200 hr. These samples were incubated with 1.5 ml of 4% starch in 0.1 M citrate buffer, pH 4.8, at 30” for 1 hr. Duplicate aliquots of 0.2 ml were then removed and applied onto EatonDikeman 613 chromatography papers (8 X 11 in), and the glucose separated by two or three ascents of a 1-butanol:pyridine:water (6:4:3 v/v/v) solventsystem(9).Theglucosebandwaslocated byremoving l-cm strips from each edge of the individual papers and staining these by the silver nitrate dip method (10). The glucose band was then cut from the chromatographic sheet and eluted with water. A blank chromatographic paper was submitted to the same procedure to serve as a control for the analytical procedure. The aqueous eluate was analyzed for glucose content by the diphenylamine calorimetric procedure (18). The results were corrected for the amount of glucose removed in the l-cm strips. Aliquots (0.5 ml) of glucoamylase I and II were maintained at 30”, 40”, 50”, 60”, and 70” for 30 min,

AND

RASMUSSEN

rapidly cooled to 30”, and incubated with 1.5 ml of 4y0 starch in 0.1 M citrate buffer, pH 4.8, for 1 hr at 30’. The glucose was chromatographically separated as described above and analyzed by the diphenylamine calorimetric procedure. Hydrolysis of oligosaccharides. Glucoamylase I (0.12 ml, 0.23 mg protein/ml) aud II (0.12 ml, 0.25 mg protein’ml) were each incubated with 0.12 ml of 0.2 M solutions (in 0.1 M citrate buffer, pH 4.8) of maltose, maltotriose, and maltoheptaose for 2 hr at 35”. An identical procedure was performed with an isomaltose substrate using a 6-hr incubation. Duplicat,e aliquots (0.1 ml) were then applied to chromatography papers, and the glucose was chromatographically separated and analyzed as described previously. The action pattern of each glucoamylase was determined by incubating 0.1 ml of the enzyme with 0.1 ml of each 0.2 M oligosaccharide solution at room temperature. Samples of the solutions were spotted on chromatography papers after 0, 1, 2, 4, 8, and 24 hr of incubation. The papers were then developed by two ascents of a l-butanol:pyridine:water (6:4:3 v/v/v) solvent system (9), and the carbohydrate areas located by the silver nitrate dip reagent (10). Immunological properties4. Purified solutions of glucoamylase I (7.3 mg protein/ml) and II (10 mg protein/ml) were used as antigens for determining the immunological properties of the enzymes. Each enzyme antigen emulsified in Freund’s complete adjuvant (19) (0.5 ml enzyme antigen plus 0.5 ml adjuvant) was injected into the thigh muscle of individual rabbius weekly for three consecutive weeks. A total of 10.9 mg of glucoamylase I and 15 mg of glucoamylase II was given to the respective rabbits. Bleedings were made weekly by cardiac puncture in the third, fourth, and fifth week, at which time the rabbit was exsanguinated. The serum was prepared as follows. The blood, collected by cardiac puncture, was allowed to clot completely at room temperature for periods up to 2 hr. The vessel was then rimmed, when necessary, with an applicator stick to loosen the clot that adhered to the walls and heated at 37” for one-half hour. After refrigeration at 2-5” overnight, the liquid portion was pipetted from the clot and centrifuged at low speed to remove any remaining blood cells. The supernatant fluid was removed and used as the serum. The serum was divided into aliquots and stored at -20” until used. 4 The immunological studies were performed by Dr. E. M. Ball, Pathologist, Crops Research Division, Agricultural Research Service, Department of Agriculture, stationed in the Department of Plant Pathology.

GLUCOAMYLASE

OF ASPERGILLUS

The antigen-antibody reactions were then measured by the microprecipitin technique (20, 21) and by observing the zones of precipitation formed in agar after the antisera were adjusted by dilution and added to peripheral wells in agar diffusion tests. NHS-terminal amino acid. Purified solutions of glucoamylase I and II, containing approximately 30 mg of protein, were dialyzed against running tap water for 24 hr and freeze-dried. Dinitrophenyl (DNP) derivatives of the enzymes were formed using Procedure 1 of Fraenkel-Conrat et al. (22). The reaction mixture was extracted six to eight times with 5 ml of ether and, after evaporation of the ether, the residue was hydrolyzed with about 1 ml of constant boiling HCl (about 5.7 N) at 105” for 16 hr in a sealed tube. The hydrolyzate was diluted to 1 N HCl and extracted 5 times with 5 ml of ether; the ether was removed under reduced pressure. The residue was dissolved in a small amount of acetone. The acetone solution was subjected to descending chromatography using the tert-amyl alcohol: phthalate system of Blackburn and Lowther (23) and also a two dimensional system (24) employing toluene: chloroethanol :pyridine : aqueous ammonia (25) in the first dimension and concentrated (1.5 M) phosphate buffer (pH 6.0) in the second as described by Fraenkel-Conrat et al. (22). The aqueous phase was subjected to descending chromatography in the tert-amyl alcohol:phthalate system and also in a phthalate:propanol (30’$&) : cyclohexane system (25). DNP-enzyme was hydrolyzed in concentrated HCl at 105” for 4 hr in a sealed tube to determine the possible presence of DNP-glycine or DNPproline since these derivatives are destroyed under the conditions of hydrolysis cited above. The ether extract of the hydrolyzate was subjected to chromatography in the tert-amyl alcohol:phthalate system. Treatment of glucoamylase I with dissociating reagents. A purified solution of glucoamylase I was dialyzed against running tap water for 24 hr and concentrated to approximately 0.5-lyo protein by pervaporation at 4”. Attempts to dissociate the protein were performed as follows: (1) The enzyme w&s dialyzed against 0.1 M NaCl containing 1.5, 4.0, or 8.0 M urea for about 16 hr at room temperature. The urea had been twice recrystallized from 70yo ethanol as described by Steinhardt (26). (2) The enzyme was dialyzed against 0.1 M NaCl containing 8 M urea and 0.1 M p-mercaptoethanol for 16 hr at room temperature. (3) The enzyme was dialyzed against 0.1 M NaCl for 16 hr, then 1 N HCl was added rapidly to obtain a pH of 2 or 4. The acidified solution was allowed to

NIGER

543

stand at room temperature for 1 hr. Controls for these experiments consisted of enzyme which had been dialyzed against 0.1 M NaCl for the same lengths of time. Dissociation of the protein was tested by subjecting the sample to sedimentation velocity at a constant temperature of 20” and a rotor speed of 59,780 rpm, except for the experiment at pH 2 which was performed in a Kel-F cell at 50,740 rpm. A solution (1 ml, 2.3 mg protein) of glucoamylase I was mixed with 0.17 ml of 7 M urea to give a final urea concentration of 1 M. After 2 hr at room temperature, a 0.5 ml aliquot was layered on top of a glycerol gradient (5-4070 glycerol) and was centrifuged at 35,000 rpm for 28 hr in a refrigerated Beckman Model L preparative ultracentrifuge using a swinging-bucket SW39L rotor. The density-gradient tube was then fractionated into 0.3 ml fractions (27). Each fraction was analyzed for protein by the Lowry method (8), for csrbohydrate content by the orcinol-sulfuric acid procedure (28), and for relative enzyme activity by incubation with a 4yo starch substrate and visual estimation of the relative amount of glucose formed using glucose oxidase test strips. RESULTS

Enzyme purification. Two forms of the glucoamylase of Aspergillus niger were separable by chromatography on DEAEcellulose. A typical pattern containing three major protein components obtained from the stepwise elution of the enzyme is shown in Fig. 1. The first protein peak contained glucosyl transferase and p-nitrophenyl Q-Dglucoside-hydrolyzing (a-n-glucosidase) activity. The second component was glucoamylase II and the third \vas glucoamylase I. Glucoamylase II appeared as two peaks. However, these two peaks were shown to be the same enzyme by several methods. Paper electrophoresis of a sample from each glucoamylase II peak resulted in a single protein and enzymatically active component of identical mobility. The material from both peaks exhibited identical electrophoretic mobility. Disc-gel electrophoresis of a sample from both peaks on 4% and 7 % polyacrylamide gels revealed that the major component in both samples possessedthe same mobility. However, the sample from the second glucoamylase II peak also contained additional minor components. This behavior is not unexpected since the sharp

LINEBACK,

544

RUSSELL,

- a.4 - a.0 - 7.6 -72 -68 -6.4 -

PH

-6.0 - 5%

40

a0

I20

I60

200

TUBE

NUMBER

240

280

AND

RASMUSSEN

purification were incubated with the corresponding p-nitrophenyl glycosides. All three major protein peaks had activity towards p-nitrophenyl a-D-glucoside but, only the second and third peaks had activity to starch. Glucosyl transferase activity was also observed in the first protein component. Activity corresponding to an a-D-galactosidase was also observed in the first major protein component, while the 2-acetamido-2-deoxy-P-D-glucosidase activity was found in the region between glucoamylase I and II. The latter enzyme activity was almost, completely separated from the glucoamylase activities with only a very minor overlap of activities. Activities corresponding to an a-n-mannosidase or p-D-glucosidase were not observed in the crude enzyme under the assay conditions used, but low amounts of /3-n-galactosidase

I

I 8.4

FIG. 1. Chromatography of crude enzyme on DEAE-cellulose. (A) Absorbance at 280 nm; (0) pH value. The crude enzyme (5 g) was chromatographed on a column of DEAE-cellulose (3.5 X 45 cm) at a flow rate of 1 ml/min using the stepwise elution procedure described in Methods. The fractions labeled I and II refer to glucoamylase I and II, respectively.

a.0

-6.8 PH

-6.4

change in pH which occurred just prior to elution of the second peak might be expected to elute additional components. When

the material from the first glucoamylase II peak was collected and rechromatographed under the same conditions, the pattern of two peaks was again observed. These observations established the identity of the material in both peaks as glucoamylase II. When the crude enzyme was puriied on DEAE-cellulose using a linear pH gradient, the glucoamylases were eluted as single peaks as indicated in Fig. 2. Starch-hydrolyzing activity was observed to parallel the protein components which were assigned as glucoamylase I and II on the basis of their electrophoretic mobility. Activities corresponding to an cr-D-glucosidase, an cY-n-galactosidase, and a 2-acetamido-Z deoxy-@-D-glucosidasewere observed when fractions from the first chromatographic

\. I

200 TUBE

240

280

NUMBER

FIG. 2. Chromatography of crude enzyme on DEAE-cellulose. (A) Absorbance at 280 nm; (0) starch-hydrolyzing activity (absorbance at 400 nm) measured by the glucose oxidase procedure described in the text; (Cl) pH elution gradient. The crude enzyme (5 g) wa9 chromatographed on a column (3.5 X 45 cm) of DEAE-cellulose at a flow rate of 1 ml/min using the gradient procedure described in Methods. The fractions labeled I and II refer to glucoamylase I and II, respectively.

GLUCOAMYLASE

OF

activity were present in the crude enzyme and were not detected after the chromatography. Tests for heterogeneity. Paper electrophoresisof each of the purified glucoamylases at pH 7.6 yielded one enzymatically active band whose mobility coincided with the single protein band. Paper electrophoresis of each purified glucoamylase was also performed at pH values between 3.0 and 4.2 using dyes as internal standards. The protein component for both enzymes migrated as a single band at each pH value investigated. At pH 3.4 and 4.0, glucoamylase I and II, respectively, showed no electrophoretic migration indicating that these values represent the isoelectric points for the two enzymes at ionic strength 0.05. Migration of the protein component of glucoamylase I and II as a single band at pH values from 3.0 to 7.6 was indicative of a high degree of purity for both enzymes.

FIG. 3. Disc-gel electrophoresis of crude and purified glucoamylases. (A) Crude glucoamylase on 7% polyacrylamide gel; (B) crude glucoamylase on 4yc polyacrylamide gel; (C) purified glucoamylase II on 4y0 polyacrylamide gel; (D) purified glucoamylase I on 4y0 polyacrylamide gel. The proteins (50-199 pg) were electrophoresed for 90 min at 5-10” using a current of 3 mA/tube and stained with Coomassie blue under the conditions described in the text.

ASPERGILLUS

NIGER

545

Disc-gel electrophoresis on 7 % polyacrylamide gels established that the crude enzyme contained several protein components which were removed during the purification procedure (Fig. 3). Separation of glucoamylase I and II was not affected on 7 % polyacrylamide gels (Fig. 3A) but readily occurred when 4% gels were used (Fig. 3B). Starch-hydrolyzing activity was observed to correspond to the two major protein components in the crude enzyme and to the single protein component of each of the purified enzymes. The most rapidly migrating glucoamylase component on the 4 % gel corresponded to glucoamylase I and hence was analogous to the results obtained with paper electrophoresis (1). Absorbance patterns, obtained from scanning stained gels (Fig. 4) revealed that each of the purified enzyme solutions contained only one stainable component. The third starch-hydrolyzing component in Fig. 4B has not been identified. It appeared only in disc gels of the crude enzyme and may coincide with a weakly staining protein component. The efficiency of the purification procedure at each step is indicated in Table I using values from a typical purification procedure. Variations in specific activity were obtained with the stepwise elution procedure depending on the amount of each buffer used, the molar&y of the buffer, and the length of dialysis of the crude enzyme prior to purification. The gradient elution procedure yielded much smaller variations in specific activity. However, these variations did not significantly alter the reproducibility of the overall purification procedure, and tests for heterogeneity using the final product indicated high degrees of purity regardless of the purification method, employed. Protein analysis of the three major fractions obtained from the first column fractionation revealed that 65-70% of the protein in the crude enzyme was glucoamylase I and II. The purification procedure provided approximately a 2.5-fold increase in the activity of both glucoamylases. The extent of purification of each glucoamylase was determined from the specific activity of that enzyme in the crude starting material: The specific activity of the crude enzyme cannot

546

LINEBACK,

RUSSELL,

AND RASMUSSEN

A

FIG. 4. Densitometer scans of the polyacrylamide gels in Fig. 3. (--) Absorbance at 570 nm; (-----) starch-hydrolyzing activity (absorbance at 400 nm) measured by the glucose oxidase procedure described in the text. A, B, C, and D correspond to the gels in Fig. 3. The tops of the gels correspond to the left side of the scan.

be used as the base figure since more than one starch-hydrolyzing enzyme was present, making this a composite value. Rechromatography did not increase the specific ac-

tivity of the glucoamylases. The difference in purification factor and specific activity between Table I and the values reported for glucoamylase I by Pazur and Ando (1)

GLUCOAMYLASE

OF ASPERGILLUS TABLE

NIGER

I

PURIFICATIONOF GLUCOAMYLASE FROMAspergillus Enzyme Procedure

Crude enzyme Glucoamylase Glucoamylase Chromatography Glucoamylase Glucoamylase

Vol (ml)

COIlCllTotal

(units/ml)

166

547

137

units (X 10-Z)

137

niger

activity

Protein b&Ill)

11.6

I II

Cu~~s”i”n~

TZd0

12 5” 6”

100

13 15

31 42

Purification”

1.0 1.0

(DEAE-cellulose) I II

Subtotal Rechromatography of glucoamylase I (DEAE-cellulose) Glucoamylase I Rechromatography of glucoamylase II (DEAE-cellulose) Glucoamylase II a Activity determined on reconsituted b Reflects increase in specific activity

395 480

11 12

43 58

0.85 0.81

iii

2.6 2.5

73

366

10

30

0.83

12

22

2.4

568

10

57

0.62

16

42

2.7

crude preparation (See Methods). over crude enqyme.

cannot be completely reconciled at this time. The latter reported a 13.8-fold increase in activity between the crude enzyme (210 units glucoamylase activity per mg of nitrogen) and the purified glucoamylase I (2900 units per mg of nitrogen). The activity for the crude enzyme was reported per mg of nitrogen and was determined prior to the dialysis step. In our hands, a 25-40% loss of protein determined by the Lowry procedure (8) occurred during dialysis. The value for the crude enzyme in Table I was determined after dialysis. Prior to dialysis, a value of 8 units per mg protein was obtained. Conversion of the units of enzymatic activity used in the present work to those of Paaur and Ando (1) are difficult since a soluble starch substrate was used in the present assay whereas Pazur and Ando used methanol-extracted commercial corn starch. The different substrates could result in a considerable difference in the apparent enzymatic activity. Lack of knowledge concerning the exact nitrogen composition of the glucoamylases precludes the use of a conversion factor for converting protein into nitrogen with any degree of accuracy. However, all criteria of heterogeneity indicate that glucoamylase I and II in this investigation possessa very high degree of purity. On the basis that glucoamylase I and II

comprise 65-70 % of the protein in the crude enzyme preparation after dialysis, a 13.S fold increase in activity would be virtually impossible in this work. A typical sedimentation velocity ultracentrifuge pattern for glucoamylase I is illustrated in Fig. 5. When subjected to ultracentrifugal analysis (approximately 200,000 g) both enzymes sedimented as single symmetrical boundaries, suggesting that they were not significantly contaminated by moleculesof different sedimentation coefhcients. A detailed boundary analysis was performed on the purified enzyme solutions by calculating the apparent diffusion coefficient from each picture and plotting these values as a function of time (17). If a sample contains molecules of varying sedimentation coefficients, the apparent diffusion coefficient will increase with time in a linear fashion. An increase in the apparent diffusion coefficients of the enzymes was not obtained, indicating each was highly purified. Properties of the glucoamylases. The pH optima for the action of each enzyme on a starch substrate was determined in buffers of varying pH and are shown in Fig. 6. For both enzymes nearly identical pH curves were obtained with a broad region of maximum activity at 4.5-5.0 and with approximately 25 % inactivation at pH values

548

LINEBACK,

RUSSELL,

FIG. 5. Sedimentation velocity photographs NaCl at pH 4 (B); and 0.1 M NaCI-4 M urea rpm.

(C).

obtained with Sedimentation

c ?.46z 4 37: ? .2aI 4 g .Iad

.09I

I I.0

I 20

I 30

, 40 ti

I 50

1 60

I 70

AND

1 a0

FIG. 6. pH optima for action of glucoamylase I (0) and II (0) on starch. The enzymes (0.5 ml) were incubated with 4y0 starch (1.5 ml) at 30” for 1 hr, and the glucose produced was determined as described in the text.

RASMUSSEN

glucoamylase I in 0.1 M NaCl (A); 0.1 M was at 20” and a rotor speed of 59,780

below 3.0 or above 6.5. This agrees with the value of pH 4.8 reported for glucoamylase I with approximately 50 % inactivation at pH values below 3.0 or above 6.5 (1). The temperature stability of glucoamylase I and II was studied at elevated temperatures and at 30” for prolonged periods of time. The two enzymes gave nearly identical stability curves at temperatures ranging from 30” to 70” (Fig. 7). The enzymes were cprite stable up to 60” with essentially no loss of activity in 30 min but were inactivated (approximately SO%) between 6OO” and 70”. A difference in the relative stabilities of glucoamylase I and II were noted on storage of the enzyme at 30”, with glucoamylase II being somewhat less stable. Glucoamylase I showed essentially no change in activity after 168 hr at 30”, while glucoamylase II had lost approximately 19 % of its activity under the same conditions. These results contrast with those reported

GLUCOAMYLASE

93-

OF ASPERGZLLUS

NIGER

549

substrates at about the same rate. The sequence of production of hydrolytic products from maltoheptaose was determined by paper chromatography for each enzyme. During the course of the reaction, maltohexaose, maltopentaose, maltotetraose, maltotriose, maltose, and glucose were formed -h 5 6.5from maltoheptaose. The concentration of .tIn the higher oligosaccharides decreased with 2 time, as would be expected for a multichain > 5.6mechanism of action (1). 5 The two enzymes were used as immuniz5 4.6ing antigens in rabbits to determine their immunological properties and obtain in8 formation on possible differences in amino 2 37acid sequence at the antigenic centers of z the two glucoamylases. The reciprocal of s2 2.8the dilution endpoint as determined by the d microprecipitin technique (20, 21) was 4 for 1.8glucoamylase I as immunizing antigen against test antigen I and II, and for glucoamylase II as immunizing antigen was 0.916 for test antigen I and 32 for test antigen II. When the antisera were adjusted by 50 33 40 60 70 80 dilution and added to peripheral wells in TEMPERATURE (“C) agar diffusion tubes, no significant differFIG. 7. Temperature stability of glucoamylase ence was noted in the zones of precipitation I (0) and II (X ). The enzyme solutions were that formed in the agar. Both antisera reheated at the indicated temperatures for one-half solved themselves into two lines that fused hour, rapidly cooled to 30”, and the activity at each end or coalesced to form one line determined against 4% starch by the procedures with the enzyme antigen. described in the text. The NHr-terminal amino acid was determined by the 1-fluoro-2,4-dinitrobenzene by Pazur and Ando (1) that glucoamylase technique originally developed by Sanger II showed four times as much inactivation as glucoamylase I at 50” and 60” and was (29). Chromatography of the ether extracts less stable at 30’ (52 % inactivation at 140 of the hydrolyzed DNP-protein by means of a two-dimensional system (24) and a hr storage vs. none for glucoamylase I). The action of each glucoamylase on one-dimensional system revealed only one maltose, maltotriose, maltoheptaose, and spot in each DNP-enzyme hydrolyzate in isomaltose was investigated qualitatively and quantitatively in citrate buffer, pH 4.8. TABLE II The quantitative results are shown in Table HYDROLYSIS OF OLIGOSACCHARIDES BY II. The extent of hydrolysis was calculated GLUCOAMYLASE I AND II on the basis of equal protein concentration GlucoamvlaseI Glucoamvlase II for each enzyme. Both enzymes hydrolyzed Substrate mg glu/mg % HY- mg glu/mg % HYthe a-~-(1 -+ 4) linkage approximately 40 protein-hr drolysis protein-hr drolysis times more rapidly than the a-~-(1 + 6) 66 42 61 38 linkage. This agrees with the relative rates Maltose 84 35 86 36 of 30 reported by Pazur and Kleppe (5) for Maltotriose Maltoheptaose 160 29 183 33 glucoamylase I. Both enzymes were ob- Isomaltose 1.6 15 1.5 14 served to act against these oligosaccharide

550

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024

RUSSELL,

t

g 016a 8 0123 Q OOBoc%c

_

.y

7”” II

I

2

4

I1

/

6

1

8 TUBE

I

IO NUMBER

,/,I,,

12

14

16

FIG. 8. Concentration of protein and carbohydrate in fractions obtained from densitygradient centrifugation of glucoamylase I after treatment with urea (1 M) and centrifugation in a glycerol gradient (540%) at 35,000 rpm for 28 hr. (0) Absorbance at 280 nm for proteins; (0) absorbance at 505 nm for carbohydrate (orcinolsulfuric acid assay).

addition to the usual spots of dinitrophenol and dinitroaniline. This spot corresponded to DNP-alanine. No spots corresponding to possible NH%terminal amino acids were observed when the aqueous phase was chromatographed. DNP-proline or DNPglycine were not observed when samples from a short hydrolysis in concentrated HCl were chromatographed using the onedimensional svstem. Thus, the KHz-terminal amino acid in each glucoamylase appears to be alanine. Dissociation of glucoamylase I into subunits was attempted in 1, 1.5, 4, and 8 M urea and at pH 2 and 4. The attempted dissociation in 1 M urea was investigated by density-gradient ceutrifugation. After centrifugation in a glycerol gradient, the gradient was fractionated, and each fraction was analyzed for protein and carbohydrate content and for relative enzymatic activity. The relative concentrations of protein and carbohydrate were plotted against fraction number and are shown in Fig. 8. It can be seen that the protein and carbohydrate distribution parallel each other yielding only one peak. The glucoamylase activity, measured by a visual estimate of the glucose produced from a starch substrate using

AND

RASMUSSEN

glucose oxidase reagent strips, paralleled the protein and carbohydrate concentrations. The effects of higher concentrations of urea and acid were determined by sedimentation velocity experiments. The 8 M urea caused some masking of the boundary by giving a high baseline value, but it still appeared that only a single uniform boundary sedimented down the cell. This was not significantly changed in the presence of 0.1 M fl-mercaptoethanol. Only a single uniform boundary was observed in the 1.5 and 4 M urea solutions, and the estimated rate of sedimentation, compared to a control sample containing no urea, was insufficiently lower to be caused by any dissociation into subunits of smaller size. The slightly lower sedimentation rate in urea was probably due to the increased viscosity of these solutions. At pH 4, glucoamylase I exhibited the same sedimentation pattern as the control sample. At pH 2, only a diffuse boundary formed which sedimented much more rapidly than the control sample. This is probably due to aggregation of the protein in this strongly acid solution and cannot be attributed to any dissociation into subunits, which would sediment at a lower rate. The sedimentation velocity patterns in 4 M urea and at pH 4 are shown in Fig. 5. DISCUSSION

Chromatography of a crude preparation of the glucoamylase from A. niger on DEAEcellulose ion-exchange columns yields two forms of the glucoamylase. Both enzymes are separated from the a-amylase and glucosyl transferase present in the crude enzyme (1). The purified enzymes possess a high degree of purity SLSindicated by paper electrophoresis, sedimentation velocity, and disc-gel electrophoresis. Disc-gel electrophoresis, a highly resolving method, indicates that a number of protein components present in the crude starting material have been removed during the purification and the two forms of the glucoamylase have been separated completely. The densitometer scan of purified glucoamylase I and II on 4 % polyacrylamide gels clearly revealed that a single component was present. Discgel electrophoresis of the crude enzyme

GLUCOAMYLASE

OF

resolved the two glucoamylases and further indicated that both were present in the starting material. Thus, the two forms did not arise as artifacts of the isolation procedure. Elution of the enzymes from DEAEcellulose using a linear pH gradient produced results comparable to those obtained with the stepwise elution (1); however, the former has been found to be a simplified system for routine use. A major problem with the stepwise elution procedure is the necessity of adding buffers at various times during the elution. It thus becomes very difficult to add identical amounts of each buffer every time the purification procedure is performed. The use of different amounts of the various buffers resulted in slightly different elution patterns each time. The gradient procedure eliminated this problem, since a constant amount of buffer is used each time the purification is performed and it also was found to yield more consistent elution patterns. The two glucoamylases were observed to have nearly identical pH optima, temperature stability at elevated temperatures, action patterns on malto-oligosaccharides, and NHz-terminal amino acids indicating a considerable degree of similarity between the two. Since the reciprocals of the dilution endpoint as determined by the microprecipitin technique were the same for antigen glucoamylase I and differed only by a factor of two for antigen glucoamylase II, it was concluded that no major differences in structure at the antigenic sites of the twro enzymes were detectable by this technique. Production of the same zones of precipitation in agar by the antisera of both enzymes supported the idea that the differences were in concentration of antibodies, not in differences of antibodies produced by immunization with the two enzymes. Thus, the two enzymes produced antibodies in rabbits that were serologically similar, which may reflect similarity in the polypeptide structure at the antigenic centers of the enzymes or in the carbohydrate moieties, if they are the antigenic determinants. The close correspondence in physical properties and antigenic behavior indicated

ASPERGILLUS

NIGER

551

that the two enzymes must be very closely similar to each other. Among the properties thus far investigated, the two enzymes differ only in electrophoretic mobility, isoelectric point, and, to a slight extent, temperature stability over prolonged periods at 30’. Pazur, Kleppe, and Ball (6) have also reported that the two glucoamylases differ in carbohydrate content. However, the carbohydrate moieties in both enzymes appear to be O-glycosidically linked through mannose to the hydroxyl groups of serine and threonine in the polypeptide chain (7). Multimolecular or isoenzyme forms of enzymes have been demonstrated for more than 30 enzymes (30), the best known of which is lactate dehydrogenase (31, 32). Multimolecular forms of hydrolyzing enzymes are less common but have been observed for certain esterases(33), the glucoamylase of A. niger (1, 2), and the glucoamylase of A. oryzea (34). Four fractions of glucoamylase, differing chromatographically from one another, were isolated and purified from a culture of A. oryzae on steamed rice (35). These components resembled one another in their properties except for small differences in their electrophoretic mobilities, sedimentation coefficients, and pH stabilities (35). One of the components (Fraction A-3) was obtained as a homogeneous component and characterized as a glycoprotein (36). However, apparently not all fungi produce extracellular glucoamylases having multimolecular forms. A single glucoamylase has been purified and characterized from culture filtrates of Coniophora cerebella grown in starch-containing media (37) and from Rhizopus delemar (11). The latter glucoamylase has also been identified as a glycoprotein (11). Data from numerous experiments in which enzymes were subjected to dissociating agents and examined by ultracentrifugation have shown that many enzymes and isoenzymes are composed of subunits held together by electrostatic forces or disulfide bonds. Enzymes which have been shown to consist of subunits include rabbit-muscle aldolase (38), lactate dehydrogenase (39,40), yeast alcohol dehydrogenase (41)) malate

552

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dehydrogenase (42)) and rabbit-muscle enolase (43). Investigation of glucoamylase I by density-gradient centrifugation after treatment with 1 M urea indicated that the protein, carbohydrate, and enzymatic activity parallel one another. This is additional evidence that the carbohydrate residuesare covalently bonded to the protein in the two enzymes (6) and are not held by electrostatic bonds. The failure of additional boundaries to be noted in the sedimentation velocity experiments in the absence of dissociating agents also indicates that the carbohydrate is covalently bonded to the protein, unless the sedimentation rate of carbohydrate is identical to that of the protein. This appears to be unlikely. Failure to observe dissociation with the concentrations of urea, acid, and p-mercaptoethanol used indicate that glucoamylase I is not dissociated into subunits under any of these conditions. Under similar conditions, lactic acid dehydrogenase (40) and rabbit-muscle aldolase (33) have been dissociated into subunits. It is concluded that Aspergillus nigey produces a glucoamylase consisting of two forms which are very similar to each other. It appears that this glucoamylase is an example of an enzyme that exists in multimolecular forms, which are not dissociable into subunits. Further studies are being pursued to determine the molecular weights, carbohydrate contents, and fine structures of these two enzymes. ACKNOWLEDGMENTS This investigation was supported by a grant from the Agricultural Research Service, U.S. Department of Agriculture, grant No. 12-14-1909143 (71), administered by the Northern Utilization Research and Development Division, Peoria, Illinois. We express our appreciation to Mr. W. E. Baumann, Mr. R. L. Doty, Mr. G. E. Farley, and Mrs. S. Beygu Farber for their excellent technical assistance, to Dr. E. M. Ball for the immunological studies, and to Drs. H. W. Knoche and F. W. Wagner for many helpful discussions. REFERENCES 1. Pazv~, J. H., Ann ANDO, T., J. Biol. Chem. 234, 1966 (1959). 2. FLEMING, I.D., AND STONE, B. A., Biochem. J. 97, 13P (1965).

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E. C., Nature 203, 821 (1964). J. H., AND ANDO, T., J. Biol. Chem. 236, 297 (1960). 5. PAZUR, J. H., AND KLEPPE, K., J. Riol. Chem. 237, 1992 (1962). 6. PAZUR, J. H., KLEPPE, K., AND BALL, E. M., Arch. Biochem. Biophys. 103, 515 (1963). 7. LINEBACK, D. It., Carbohydrate Res. 7, 106 (1968). 8. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J., J. Biol. Chem. 198, 265 (1951). 9. FRENCH, D., AND WILD, G. M., J. Am. Chem. Sot. 76,2612 (1953). 10. MAYER, F. C., AND LBRNER, J., J. Am. Chem. Sot. 81, 188 (1959). 11. PAZUR, J. H., AND OKADA, S., Carbohydrate Res. 4, 371 (196’7). 12. PAYNE, L. C., AND MARSH, C. L., J. Nutrition 76, 151 (1962). 13. DAVIS, B. J., Ann. N. Y. Acad. Sci., 121, 404 (1964). 14. WILLIAMS, D. E., AND REISFELD, R. A., Ann. N. Y. Acad. Sci. 121, 373 (1964). 15. AKROYD, P., Zone Electrophoresis, In “Chromatographic and Electrophoretic Techniques” (I. Smith, ed.), Vol. II, p. 463. Interscience, New York (1968). 16. CHRAMBACH, A., REISFELD, R. A., WYCKOFF, M., AND ZACCARI, J., Anal. Biochem. 20, 150 (1967). 17. SCHACHMAN, H. K., In “Methods in Enzymology” (S. P. Colowick and N. 0. Kaplan, eds.), Vol. 4, p. 32. Academic Press, New York (1957), 18. PAZUR, J. H., J. Biol. Chem. 206,75 (1953). 19. FREUND, J., Ann. Rev. Microbial. 1, 291 (1947). 20. SLOGTEREN, D. H. M. VAN., Proc. Conf. Potato virus Dis. .%d, Lisse-Wageningen, 51 (1955). 21. SLOGTEREN, E. VAN, AND SLOGTEREN, D. H. M. VAN., Ann. Rev. Microbial. 11, 149 (1957). 22. FRAENKEL-CONRAT, H., HARRIS, J. I., AND LEVY, A. L., In “Methods of Biochemical Analysis” (D. Glick, ed.), Vol. II, p. 359. Interscience, New York (1955). 23. BLACKBURN, S., AND LOWTHER, A. G., Biothem. J. 48, 126 (1951). 24. LEVY, A. L., Nature 174, 126 (1954). 25. BISERTE, G., AND OSTEUX, R., Bull. Sot. Chim. Biol. 33, 50 (1951). 26. STEINHARDT, J., J. Biol. Chem. 123, 543 (1938). 27. PAZUR, J. H., KLEPPE, K., AND ANDERSON, J. S., Biochim. Biophys. Acta 66, 369 (1962). 28. WINZLER, R. J., In “Methods of Biochemical Analysis” (D. Glick, ed.), Vol. II, p. 279. Interscience, New York (1955). 29. SANGER, F., Biochem. J. 39, 507 (1945).

GLUCOAMYLASE

OF

30. GREGORY, K. F., Ann. N. Y. Acad. Sci. 94, 657 (1961). 31. MARKERT, C. L., AND M$LLER, F., Proc. Natl. Acad. Sci. U.S. 46,753 (1959). 32. VESELL, E. S., AND BEARN, A. G., Proc, Sot. Exptl. Biol. Med. 94, 96 (1957). 33. AUGUSTINSSON, K. B., Ann. N. Y. Acad. Sci. 94, 844 (1961). 34. MORITA, Y., SHIMIZU, K., OHGA, M., AND KORENAGA, T., Agr. Biol. Chem (Tokyo) 30, 114 (1966). 35. OHGA, M., SHIMIZU, K., AND MORITA, Y., Agr. Biol. Chem. (Tokyo) 30, 967 (1966). 36. MORITA, Y., OHGA, M., AND SHIMIZU, K., Mem.

ASPERGILLUS

37. 38. 39. 40. 41. 42. 43.

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553

Research Inst. Food Sci., Kyoto Univ. 29, 18 (1968). KING, N. J., Biochem. J. 106,577 (1967). STELLWAGEN, E., AND SCHACHMAN, H. K., Biochemistry 1, 1056 (1962). APPELLA, E., AND MARKERT, C. L., Biochem. Biophys. Res. Commun. 6, 171 (1961). FRITZ, P. J., BND JACOBSON, K. B., Science 140, 64 (1963). KAGI, J. H. R., AND VALLEE, B. L., J. Biol. Chem. 236, 3188 (1960). MUNKRES, K. 1)., Biochemistry 4, 2168 (1965). WINSTEAD, J. A., AND WOLD, F., Biochemistry 4, 2145 (1965).