Biochem. J. (1976) 153, 621-625 Printed in Great Britain

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Physical and Catalytic Properties of a-Amylase from Tenebrio molitor L. Larvae By VINCENZO BUONOCORE,*t ELIA POERIO,* VITrORIO SILANOt and MAURIZIO TOMASIt

*Istituto di Chimica Organica, Universitta di Napoli, 80134 Napoli, and tLaboratori di Chimica Biologica, Istituto Superiore di Sanit&, 00136 Roma, Italy (Received 15 September 1975) The amylase from Tenebrio molitor L. larvae (yellow mealworm) was characterized according to a number of its molecular and catalytic properties. The insect amylase is a single polypeptide chain with mol.wt. 68000, an isoelectric point of 4.0 and a very low content of sulphur-containing amino acids. The enzyme is a Ca2+-protein and behaves as an a-amylase. Removal of Ca2+ by exhaustive dialysis against water causes the irreversible inactivation of the enzyme. Moreover, the enzyme is activated by the presence in the assay mixture of Cl-, or some other inorganic anions that are less effective than Cl-, and is inhibited by F-. Optimal conditions of pH and temperature for the enzymic activity are 5.8 and 37°C. The insect amylase exhibits an identical kinetic behaviour toward starch, amylose and amylopectin; the enzyme hydrolyses glycogen with a higher affinity constant. Compared with the non-insect a-amylases described in the literature, Tenebrio molitor amylase has a lower affinity for starch. A number of a-amylases (1,4-a-D-glucan glucanohydrolase, EC 3.2.1.1) from different origins have been purified and characterized according to their physical and kinetic properties (Fisher & Stein, 1960; Takagi et al., 1971; Thoma et al., 1971), but as far as we know only one insect amylase (that from Callosobruchus chinensis) has been extensively studied (Podoler & Applebaum, 1971a,b). As many insect species depend on the effectiveness of their amylases for survival, characterization studies of insect amylases are not only of interest for comparative investigations, but they can also contribute to clarifying the compatibility of some natural diets with insect development. This is particularly true for an insect species such as Tenebrio molitor L. (yellow mealworm) that feeds on wheat flour during both larval and adult life. With the characterization study of T. molitor larval amylase reported in the present paper, we also wished to gain some of the basic knowledge needed to investigate the inhibition of this enzyme by protein inhibitors from wheat flour. This inhibition has been shown to occur not only in vitro (Applebaum, 1964; Shainkin & Birk, 1970; Silano et al., 1973), but also in vivo (Applebaum, 1964). The characterization of T. molitor larval amylase has been made possible by the availability of the purified enzyme from the affinity method of Buonocore et al. (1975), which allows the one-step purification of a number of amylases inhibited by wheat albumins. Preliminary characterization data t To whom reprint requests Vol. 153

should be addressed.

about crude T. molitor larval amylase preparations have been reported (Applebaum et al., 1961; Applebaum, 1964).

Experimental Materials Starch was purchased from Connaught Laboratories Ltd., Toronto, Ont., Canada. Rabbit liver glycogen, potato starch amylopectin and amylose were supplied by BDH Chemicals Ltd., Poole, Dorset, U.K. Also from this source were a-amylase-free barley,B-amylase, acrylamide and bisacrylamide. Carrier ampholytes (Ampholine) were obtained from LKB, Stockholm, Sweden. Bovine serum albumin, chymotrypsin and cytochrome c were from Sigma Chemical Co., St. Louis, MO, U.S.A. Bio-Gel P-100 was from Bio-Rad Laboratories, Richmond, CA, U.S.A. T. molitor larval amylase was purified from insect larvae by the method of Buonocore et al. (1975), involving filtering a salt extract from larvae through a Sepharose-wheat albumin column and eluting the retained enzyme with maltose. Gel electrophoresis and electrofocusing Disc electrophoresis was carried out in 0.05MTris/0.383M-glycine buffer, pH8.5, as described by Davis (1964). Slab electrophoresis in polyacrylamide gel was carried out at pH4.3 as described by Nimmo

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V. BUONOCORE, E. POERIO, V. SILANO AND M. TOMASI

(1963) in 0.35M-fl-alanine/0.13M-acetic acid buffer. Electrofocusing was performed on a polyacrylamide-gel slab in a Multiphor apparatus from LKB by following the manufacturer's instructions. Electrofocusing in the pH range 2.5-10.0 was carried out with a mixture of Ampholines containing Ampholine pH2.5-4.0 (0.8ml), Ampholine pH4.0-6.0 (0.2ml), Ampholine pH5.0-7.0 (0.2ml) and Ampholine pH3.5-10.0 (2.4ml). Preformed gel plates (Ampholine PAGplate) from LKB were used for electrofocusing in the pH range 2.5-6.5. The pH gradient in the gel was measured with an Ingold (Zurich, Switzerland) surfaced pH electrode, type 403-30-M8.

et al.

Molecular-weight studies Polyacrylamide-gel electrophoresis in sodium dodecyl sulphate was carried out as described by Weber & Osborn (1969). The filtration on Bio-Gel P-100 was carried out on a column (1.2cmx 110cm) equilibrated with 0.02M-sodium cacodylate/HCl buffer, pH5.8. Reference proteins for both electrophoresis and gel filtration were bovine serum albumin, chymotrypsinogen and cytochrome c, with mol.wts. 68000, 25700 and 12500 respectively. Sedimentation-equilibrium runs were performed in a Beckman E analytical ultracentrifuge for 72h at 4°C and 20000rev./min in 0.05M-cacodylate/HCI buffer containing 0.01M-NaCl, pH5.8; the amylase concentration was 0.3 mg of protein/ml. Optical methods The u.v.-absorption spectrum was studied in 0.1 MKCI with a Cary 15 spectrophotometer. The fluorescence spectrum was deternined with an AmincoBowman 4-8202 spectrofluorimeter and the circulardichroism measurements were made with a Cary 60 spectropolarimeter equipped with a 6002 CD accessory. Amino acid analyses Protein samples (0.6mg) were hydrolysed in vacuo at 1 10°C for 24 and 72hin 6M-HCI. Hydrolysates were analysed with a Beckman 116 amino acid analyser as described by Spackman et al. (1958). Appropriate corrections were made for the slow release of some amino acids. Tryptophan was calculated from the tyrosine:tryptophan ratio determined optically by the method of Bencze & Schmid (1957). The partial specific volume was calculated from the amino acid composition as described by Schachman (1957).

Activity assay and action pattern The amylase activity was assayed by Nelson's (1944) colorimetric method as described by Robyt & Whelan

(1968). Unless otherwise stated the reaction mixture for T. molitor amylase activity contained, in a final volume of 1 ml, 0.05M-sodium cacodylate/HCl buffer (pH 5.8), 0.01 M-NaCl, 0.1 mM-CaCl2, 2mg of starch and 0.02-0.2 unit of amylase. The reaction was started by the addition of the enzyme and was carried out at 37°C for 10min. When the effect of anions on the enzymic activity was tested, the salts were added to a reaction mixture containing 0.05M-sodium cacodylate/cacodylic acid buffer, pH5.8, and 2mg of starch dialysed against water. One unit of amylase is the amount of enzyme that produces 1 umol ofmaltose in 1 min at 370C under our experimental conditions. The assay of barley fi-amylase was performed by the same procedure in a 0.05 M-sodium acetate/acetic acid buffer, pH4.8. The action pattern of T. molitor larval amylase was checked by the method of Dr4mmond et al. (1971) by using oxidized amylose as substrate. To prepare this oxidized substrate, 100(mg of amylose was dissolved in 24ml of boiling water; the solution was cooled and then added to 1 ml of 0.1 M-NaIO4. After I h in the dark at room temperature (20°C) the oxidized amylose was dialysed against three changes (6h and 21itres each) of water. Protein determination

Protein concentration was determined by the Lowry method as modified by Hartree (1972), with bovine serum albumin as standard. Results Physical properties Only one band was detected when purified T. molitor larval amylase was submitted to polyacrylamide-gel electrophoresis in two buffer systems (pHT8.5 and 4.3) or to gel electrofocusing in the pH ranges 2.5-10.0 or 2.5-6.5. The enzyme migrated towards the anode in both the alkaline and acidic buffer systems. At pH4.3 the enzyme moved only slightly from the application slot, whereas at pH 8.5 its electrophoretic mobility was about 0.95 of that of Bromophenol Blue. In good agreement with such electrophoretic behaviour, the amylase focused in the region of the gel with pH4.,0. One band was also detected by submitting the amylase to gel electrophoresis in a buffer system containing 1 % (w/v) sodium dodecyl sulphate. In this system the amylase mobility was identical with that of bovine serum albumin. The u.v.-absorption spectrum of T. molitor amylase showed two poorly resolved maxima at 274 and 280nm; the ElNcm at these two wavelengths was 8.1. The fluorescence spectrum of the enzyme was similar to that exhibited by a number of proteins, with a 1976

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a-AMYLASE FROM TENEBRIO MOLITOR maximum at 340nm. The circular-dichroism spectra of the amylase in the far- and near-u.v. regions were also determined. In the region of peptide-bond absorption, two negative bands at 208 and 224nm and a crossover, near 200nm, to a relatively intense positive band at 195nm were evident. The intensity of the spectrum near 208nm suggests that T. molitor larval amylase has a low content of a-helical structure. In the region of aromatic side-chain absorption, three positive bands at 278, 286 and 293nm, with a shoulder at 269nm, were evident; this spectrum is very different from those reported by Ogasahara et al. (1970) for the a-amylases from Bacillus subtilis and Bacillus stearothermophilus. T. molitor larval amylase and bovine serum albumin exhibited identical equilibrium-sedimentation patterns and identical elution patterns from a Bio-Gel P-100 column. It appears that, within the error range of the methods used, the molecular weight of T. molitor larval amylase can be considered identical with that

of bovine serum albumin, i.e. 68000. -The amino acid molar proportions determined in hydrolysates of T. molitor larval amylase were as follows: Lys 2.8; His 2.1; Arg 4.2; Asp 16.3; Thr 4.4; Ser 7.1; Glu 9.3; Pro 4.2; Gly 12.3; Ala 6.7; half-Cys trace; Val 6.7; Met 2.1; Ile 4.3; Leu 6.5; Tyr 3.7; Phe 4.5; Trp 2.8. The partial specific volume calculated from this amino acid composition was 0.716.

Parameters influenctng enzymic activity and'stability The pH-activity profile ofpurified T. molitor larval amnylase is a bell-shaped cuwve with the maximum at pH5.8, similar to that reported for a crude enzyme preparation from the samesourec by Applebaumeatal. (1961). At pH 5.8 the enzyme exhibited maximal activity at 50°C, but at such a temperature the timecourse of starch hydrolysis was linear only for a few minutes. To obtain a time-course linear for at least 30min, a working temperature of 37°C was chosen. Stability of T. molitor larval amylase to pH was studied by preincubating the enzyme in solutions of different pH values for 30min and 24h at both 4°C and 37°C before the enzymic assay. At the lower temperature the amylase could be incubated for 24h without any detectable loss of activity at any pH in the range 5.8-8.5, whereas at pH4.0 or 12.0 even the 30min preincubation caused the complete denaturation of the enzyme. As shown in Fig, l(a) at 370C the amylase exhibited a narrow range of stability to pH. At temperatures higher than 370C the enzyme showed poor stability, even at the pH of maximal activity (Fig. lb). The crude T. molitor larval amylase preparation studied by Applebaum et a!. (1961) was irreversibly inactivated after 30min incubation at 350C, showing a much lower stability, than the purified amylase. This difference might be ascribed to the presence of proteinase contants in the amylase preparation of Applebaum et al, (1961).

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6

8

0

30

40

50

60

Temperature (°C) pH Fig. 1. Effect of(a)pHand (b) temperature on the stability ofT. molitor larvalamylase (a) T. molitor amylase was preincubated at 37°C for 30min (0) or 24h (0) at the pH values indicated and then assayed under optimal conditions for activity. The buffers used were: 0.02M-sodium acetate/acetic acid (pH3.5-5.5); 0.02Msodium cacodylate/HCI (pH5.0-7.0); 0.02M-sodium barbital/HCI (pH7.0-9.0); 0.02M-glycine/NaOH (pH8.5-11.0). All buffers were added witb 0.01 M-NaCl and 0.1 mM-CaCI2. (b) T. molitor amylase was preincubated in 0.05 mSsOdium cacodylate/UCI buffer, pH 5.8, for 5 (0), 20 (A) or 60 (0) min at the temperatures indicated, then assayed under the optimal conditions for activity.

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V. BUONOCORE, E. POERIOj V. SILANO AND M. TOMASI

Table 1. Effect of common inorganic anions on the activity of T. molitor larval amylase T. molitor larval amylase after dialysis against O.1miCaSO4 was tested in 0.05M-sodium cacodylate/cacodylic acid, pH5.8. Dialysed starch was used as substrate. The anions were added as 10mM sodium salts. Relative activity (%) Anion-added 100 Chloride 79 Bromide 51 Iodide 25 Fluoride 72 Nitrate 62 Chlorate 56 Sulphate 52 None

Purified T. molitor larval amylase was fully and irreversibly inactivated by exhaustive dialysis at 4°C against water or O.lmM-Na2SO4, but it retained its activity when the dialysis was carried out against 0.1 mM-CaSO4 thus indicating its Ca2+-protein nature. We have also observed that the activity of the enzyme dialysed against CaSO4 is not sensitive to the addition of Ca2+ to the assay mixture up to 10mM. The effect of several organic and inorganic anions on the amylase activity was studied by testing the enzyme in 0.05M-sodium cacodylate/cacodylic acid buffer (pH5.8) added with the sodium salt of the anion studied. T. molitor amylase activity was not affected by acetate or oxalate up to 10mM, whereas the addition of Cl- ions increased the enzymic activity dose-dependently; maximal activation was observed at 10mM(Table 1). Of several other inorganic anions tested, most were less effective than C1- in activating the amylase, and F- strongly inhibited the enzyme (Table 1). Catalytic and kinetic patterns The enzyme showed an identical degrading activity on native and oxidized amylose, whereas barley f6-amylase, used as a control, was able to hydrolyse native amylose, but showed a very poor degrading activity on oxidized amylose (Fig. 2). These results indicate that T. molitor larval amylase has a typical endoamylolytic nature. An identical conclusion was reached by Applebaum (1964) from the analysis of the oligosaccharides produced by the action of crude T. molitor larval amylase on starch. Kinetic behaviour of T. mwlitor amylase toward starch, amylose and amylopectin was not significantly different. As calculated from LineweaverBurk plots, the Km for these three substrates was about 1 .8mg/ml and the Vma.. was 1600pmol of maltose/min per mg of protein. The amylase was also able to hydrolyse glycogen with a Km of 13.3mg/ml and a

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60

120

180

Time (min) Fig. 2. Action patterns of T. molitor amylase and barley fi-amylase on native and oxidized amylose T. molitor amylase was incubated at 37°C with native (0) or oxidized (A) amylose as substrate; at the times indicated samples were withdrawn and reducing sugars were determined. Degradation patterns of barley fi-amylase on native (0) and oxidized (A) amylose are given for comparison. The maximal activity of T. molitor amylase and barley I8-amylase with native amylose was taken as 100.

Vmax. of 1520,umol of maltose/min per mg. Some of the anions listed in Table 1 (Cl-, Br- and NO3-) have been tested for their ability to affect the maximum velocity and the affinity of the enzyme for starch. In agreement with the results reported by Levitzki & Steer (1974) for pig pancreas a-amylase, the Vmax. of the enzyme, but not the Ki, was dependent on the nature ofthe anion present.

Discussion We have shown that the amylase from T. molitor L. larvae is a typical Ca2+-endoamylase consisting, according to polyacrylamide-gel electrophoresis in sodium dodecyl sulphate, of a single polypeptide chain; the enzyme has mol.wt. 68000, a relatively high content of acidic amino acids and a low content of sulphur-containing amino acids. These physicochemical properties are common to most a-amylases described in the literature. Quite peculiar in this respect are the oc-amylases from pig pancreas (Robyt et al., 1971) and B. subtilis (Fisher & Stein, 1960), which are not formed of a single polypeptide chain, and the a-amylase from B. stearothermophilus of Manning et al. (1961), which has a lower molecular weight. However, the existence of pig pancreas subunits has been questioned by Steer et al. (1974) and conflicting data have been reported for the molecular weight of the B. stearothermophilus 1976

a-AMYLASE FROM TENEBRIO MOLITOR enzyme (Pfueller & Elliott, 1969; Ogasahara et al., 1970; Yutani, 1973). With respect to the strength of Ca2+ binding, T. molitor larval amylase appears related to plant a-amylases, rather than to mammalian, bacterial and mould a-amylases, which do not release Ca2+ on dialysis against a Ca2+-free solution (Thoma et al., 1971). On the other hand the dependence of T. molitor larval amylase activity on different anions is very similar to that exhibited by mammalian a-amylases (Fisher & Stein, 1960; Thoma et al., 1971; Levitzki & Steer, 1974). In contrast with the similarity of the molecular aspects, a large variability has been observed for the catalytic properties of the a-amylases. The optimal temperature and pH ranges for both activity and stability of amylases vary remarkably, reaching extreme values for enzymes of acid-stable and thermostable bacteria. The catalytic properties of T. molitor larval amylase appear related to those of C. chinensis a-amylase, which is the only other insect amylase extensively characterized up to now (Podoler & Applebaum, 1971b). The two insect amylases exhibit optimal activity at similar pH and temperature ranges. Moreover, the affinity constant of T. molitor amylase for gelatinized starch is close to that of C. chinensis (2.3 mg/ml). These values are significantly higher than those reported in the literature for non-insect a-amylases, which are in the range 0.1-0.8mg/ml. Further investigations are needed to establish if a higher affinity constant for gelatinized starch is a peculiar feature of insect a-amylases as compared with the a-amylases from other origins. This reseach was supported in part by a grant from the Ministero dell'Agricoltura e Foreste.

References Applebaum, S. W. (1964) J. Insect Physiol. 10, 897-906 Applebaum, S. W., Jankovic, M. & Birk, Y. (1961) J. Insect Physiol. 7, 100-108

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625 Bencze, W. L. & Schmid, K. (1957) Anal. Chem. 29, 1193-1196 Buonocore, V., Poerio, E., Gramenzi, F. & Silano, V. (1975)J. Chromatog. 114, 109-114 Davis, B. J. (1964) Ann. N. Y. Acad. Sci. 121,404-427 Drummond, G. S., Smith, E. E. & Whelan, W. J. (1971) FEBS Lett. 15, 302-304 Fisher, E. H. & Stein, E. A. (1960) Enzymes, 2nd edn., 4, 313-343 Hartree, E. F. (1972) Anal. Biochem. 48, 422-427 Levitzki, A. & Steer, M. L. (1974) Eur. J. Biochem. 41, 171-180 Manning, G. B., Campbell, L. L. & Foster, R. J. (1961) J. Biol. Chem. 236, 2958-2961 Nelson, N. (1944) J. Biol. Chem. 153, 375-380 Nimmo, C. C., O'Sullivan, M., Mohammad, A. & Pence, J. W. (1963) Cereal Chem. 40, 390-398 Ogasahara, K., Imanishi, A. & Isemura, T. (1970) J. Biochem. (Tokyo) 67, 65-75 Pfueller, S. L. & Elliott, W. H. (1969) J. Biol. Chem. 244, 48-54 Podoler, H. & Applebaum, S. W. (1971a) Biochem. J. 121, 317-320 Podoler, H. & Applebaum, S. W. (1971b) Biochem. J. 121, 321-325 Robyt, J. F. & Whelan, W. J. (1968) in Starch and its Derivatives (Radley, J. A., ed.), pp. 430-476, Chapman and Hall, London Robyt, J. F., Chittenden, C. G. & Lee, C. T. (1971) Arch. Biochem. Biophys. 144,160-167 Schachman, H. K. (1957) Methods Enzymol. 4, 32103 Shainkin, R. & Birk, Y. (1970) Biochim. Biophys. Acta 221, 502-513 Silano, V., Pocchiari, F. & Kasarda, D. D. (1973) Biochim. Biophys. Acta 317, 139-148 Spackman, D. H., Stein, W. H. & Moore, S. (1958) Anal. Chem. 30, 1190-1206 Steer, M. L., Tal, N. & Levitzki, A. (1974) Biochim. Biophys. Acta 334, 389-397 Takagi, T., Toda, H. & Isemura, T. (1971) Enzymes, 3rd edn., 5, 235-271 Thoma, J. A., Spradlin, J. E. & Dygert, S. (1971) Enzymes, 3rd edn., 5, 115-189 Weber, K. & Osborn, M. (1969) J. Biol. Chem. 244, 4406-4412 Yutani, K. (1973) J. Biochem. (Tokyo) 74, 581-586