Fish Physiology and Biochemistry19: 257–267, 1998.© 1998Kluwer Academic Publishers. Printed in the Netherlands.

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Characterization and functional properties of digestive proteases in twosparids; gilthead seabream (Sparus aurata) and common dentex (Dentexdentex)

F.J. Alarcon, M. Dıaz, F.J. Moyano∗ and E. Abellan1

Departamento Biolog´ıa Aplicada, E.P.S. Univ Almer´ıa, 04120, Almer´ıa, Spain;1Instituto Español Oceanograf´ıa,Mazarron, Murcia, Spain;∗Author for correspondence (Phone: 34.50.215294; Fax: 34.50.215476; E-mail:fjmoyano@ualm.es)

Accepted: November 10, 1997

Key words:digestive enzymes, proteases, stomach, intestine,Sparus aurata, Dentex dentex, SDS-PAGE

Abstract

Digestive proteases present in two sparids, seabream (Sparus aurata) and common dentex (Dentex dentex), havebeen characterized using both biochemical and electrophoretic techniques. Although optimum pH and temperaturefor maximum activity of both acid and alkaline proteases were similar in the two species, important differencesin total activity, as well as in thermal and pH stability were found. Specific inhibitors and SDS-PAGE zymogramswere used to clarify such differences. Evidences support the existence of a more active and complex protease set indentex. Results are discussed from the perspective of their application to the formulation of feeds for each species.

Introduction

In recent years a number of studies designed to char-acterize digestive enzymes of aquatic organisms havebeen performed. A detailed knowledge of the prote-olytic enzyme activity that exists in captive speciesis useful, both to ascertain the maximum periods fortheir storage to avoid autolysis (Connell 1980), andto develop practical applications for fish proteases inthe food industry (Raa 1990; Haard 1992). In culturedfish, this approach may be helpful in the selectionof feed ingredients (Lan and Pan 1993), particularlyfor newly cultured species, such as common dentex(Dentex dentex), a sparid which is an alternative in thediversification of Mediterranean fish farming. In thepresent work, several techniques, including the com-bination of specific inhibitors and SDS-PAGE, wereused to obtain information about digestive proteasespresent in seabream (Sparus aurata) and common den-tex. Characterization of the main enzymes involved inthe digestive process of both species revealed differ-ences that may affect nutritional strategies utilized inrearing each of them.

Materials and methods

Enzymes, substrate, inhibitors and general reagentsused were: trypsins (Type IX from porcine pancreasand; Type XX-S fromGadus morhua), chymotrypsin(Type II from bovine pancreas), pancreatin (fromporcine pancreas), Nα-tosil-L-arginin methyl ester(TAME), benzoyl-DL-arginin-p-nitroanilide(BAPNA),N-benzoyl-L-tyrosine ethyl ester (BTEE), phenyl-methylsulfonyl fluoride (PMSF), soybean trypsin in-hibitor (SBTI), N-tosyl-L-phenyl-chloromethyl keto-ne (TPCK), N-CBZ-L-phenylchlo-romethyl ketone(ZPCK), Nα-p-tosyl-L-lysin chloro-methyl ketone(TLCK), hemoglobin, tris(hydroxyme-thyl)aminome-thane base, ethylendiamine tetra acetate (EDTA) andethylene glycol bis(β-aminoethyl ether) N,N′-tetra ac-etate (EGTA) from Sigma Química (Madrid, Spain).Hammerstein grade casein, pepstatin A and chymosta-tin (CHYM) were purchased from ICN Biomedicals,Inc. (Costa Mesa, CA). Electrophoresis chambers andreagents from Bio-Rad (Richmond, CA). Molecularmass protein standard (MWM) for electrophoresis wasobtained from Pharmacia Biotech (Uppsala, Sweden).

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Live specimens of seabream (bodyweight 25 to40 g) were provided by a local fish farmer (FRAMARS.L.; Almería, Spain). Common dentex (bodyweight25 to 50 g) were obtained from the facilities of the In-stituto Español de Oceanografía in Mazarrón (Murcia,Spain). Seabream were fed on a commercial diet (45%protein) whereas the dentex were fed on a mixed dietincluding moist feed and raw fish.

After sacrificing specimens by submersing themin cold water (4◦C), the digestive tract was removedand dissected into the stomach and pyloric caecumregion. Samples of stomach were homogenized in dis-tilled water (1:10 w/v) and portions containing pyloriccaeca and proximal gut were homogenized (100 mgml−1) in cold 50 mM Tris-HCl buffer, pH 7.5. Super-natants obtained after centrifugation (16,000× g for30 min at 4◦C) were stored at –20◦C being furtherutilized for enzyme analysis. Concentration of solubleprotein in pooled samples was determined by Bradfordmethod using bovine serum albumin (1 mg ml−1) as astandard.

Alkaline proteinase activity of the extracts wasmeasured using the casein method of Kunitz (1947)as modified by Walter (1984), using substrate ca-sein (0.5%) in 50 mM Tris/HCl buffer, pH 9.0.Acid proteinase activity was evaluated according toAnson (1938) using 0.5% haemoglobin in 0.1 mMglycine/HCl, pH 2.0. The mixtures were incubatedfor 30 min at 25◦C and the reaction was stopped byadding 0.5 ml of 20% TCA. The absorbance of thesoluble TCA peptides was measured at 280 nm. Com-mercial enzymes used as reference were prepared as1 mg ml−1 solutions with the exception of cod trypsin(5 mg ml−1) and diluted to reach a linear1ABS280nmof nearly 0.5 after 30 min. One unit of enzyme activitywas defined as 1µg of tyrosine released per min, usingthe extinction coefficient for tyrosine= 0.005 mlµg−1

cm−1. All measurements were carried out in triplicate.The amidase activity of alkaline proteases (trypsin

amidase activity) was assayed at 37◦C accordingto Erlanger et al. (1961) using BAPNA as substrate(10mM in DMSO) and 50 mM Tris-HCl buffer, pH8.2, containing 10 mM CaCl2,. The reaction wasstopped by adding acetic acid. One unit of enzymeactivity was defined as 1µmol of p-nitroaniline re-leased per min, using an extinction coefficient of8800 cm−1 M−1. The esterase activity of the ex-tracts was evaluated at 25◦C as described by Hum-mel (1959) using 1mM TAME in 46 mM Tris-HClbuffer pH 8.1, containing 11 mM CaCl2. One unitof enzyme activity was defined as 1µmol of TAME

hydrolyzed per min, using an extinction coefficientof 540 cm−1 M−1. Spectrophotometric determinationof chymotrypsin activity in extracts was performedaccording to Ásgeirsson and Bjarnason (1991) mon-itoring the hydrolysis of BTEE at 256 nm (5mM in44.4 mM Tris with 55.5 mM CaCl2, pH 7.8 at 25◦C).One unit of enzyme activity was defined as 1µmol ofBTEE hydrolyzed per min, using an extinction coeffi-cient of 964 cm−1 M−1. All assays were performed intriplicate.

The optimal pH for protease activities was deter-mined using Universal Buffer (Stauffer 1989) rangingfrom 1.0 to 5.5 (acid proteases) and from 2 to 11.5(alkaline proteases). The effect of pH on stability ofproteases was determined by preincubation of extractsat different pH for 60 min, prior to assaying the resid-ual proteinase activity using casein/haemoglobin asthe substrate. The optimal temperatures for acid andalkaline proteases were determined by incubating en-zyme extracts (30 min in 50 mM Tris Hcl buffer, pH9.0) with either hemoglobin or casein pre-equilibratedat temperatures ranging from 10–70◦C. The effect oftemperature on the stability of protease activity wastested by preincubation of extracts at different temper-atures for 60 min followed by measurement of residualactivity as previously described.

The Michaelis-Menten constant (Kmapp) and max-imal velocity (Vapp

max) were determined in pyloriccaeca extracts of seabream, using specific substratesfor trypsin and chymotrypsin (hemoglobin for acidproteases). The ‘physiological efficiency’ was cal-culated as defined by Pollock (1965) and Fullbrook(1983), using Vmax/Km. The activation energy (Ea)was obtained from slopes in Arrhenius plots (slope =-Ea/R), which were constructed measuring activity at5 ◦C intervals from 25 to 45◦C for trypsin and chy-motrypsin activities of pyloric caeca extracts and 10 to45 ◦C for acid activity in stomach extracts.

The protease classes were determined using stan-dard inhibitors, following the methods described byDunn (1989) and García-Carreño (1992). The enzymeextract (20µl) was mixed with 0.5 ml of 100 mMTris-HCl, 10 mM CaCl2 buffer, pH 9 and 10µl ofdifferent inhibitors and incubated for 60 min at 25◦C.The mixture was assayed for protease activity as de-tailed previously. The assay included internal controlsfor inhibition of solvents and enzyme. Control en-zymes were assayed using 50 mM Tris-HCl, 20 mMCaCl2 buffer, pH 7.5; the percentage inhibition wasestablished based on activity without the inhibitor.

SDS-PAGE of the proteins in the enzyme prepara-

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tions was carried out according to Laemmli (1970),using 11% polyacrylamide and 8× 10 × 0.075 cmgels. Fiveµl of molecular weight marker (MWM)were loaded on each plate. Preparation of samples andzymograms of proteinase activities were done accord-ing to García-Carreño et al. (1993). The amount ofprotein sample applied was 10µg in all cases. Elec-trophoresis was carried out for 60 min at a constantvoltage of 100 V per gel at 5◦C. After electrophore-sis, gels were washed and incubated for 30 min at5 ◦C in 0,5% casein Hammerstein, pH 9.0. Then, theywere transferred for 90 min to the same solution at25 ◦C without agitation. Finally, gels were washedand fixed in 12% TCA, prior to staining with 0.1%Coomassie brilliant blue (R-250) in methanol-aceticacid-water (50:20:50). Destaining was carried out inmethanol-acetic acid-water (35:10:55).

Characterization of protease types in zymogramsfollowing SDS-PAGE separation by using specific in-hibitors was done according to García-Carreño andHaard (1993). Fourtyµl of the extracts were mixedwith 10 µl of inhibitor stock solutions (Table 5) andincubated for 1 h at 25◦C, mixed (1:1) with buffer, and25µl of the final solution was loaded on SDS-PAGEplates. Extracts incubated without inhibitor, commer-cial pure trypsins (bovine, cod) and chymotrypsin(bovine), were used as controls. Electrophoresis andzymograms were carried out as described above. Af-ter electrophoresis, gels were washed for 15 min with50 mM Tris-HCl buffer, pH 9.0, at room temperature,before incubation with the substrate (casein).

Electrophoresis of fish aspartic (acid) proteinaseswas carried out by non-dissociating discontinuousPAGE. The standard method consisted of a stack-ing gel at 4% polyacrylamide (PAA) in 0.1 MTris/phosphate, pH 5,5, a resolving gel at 13% PAAin 0.07 M Tris/HCl, pH 7.5, and 5 mM Tris, 0.62 mglycine electrode buffer, pH 7.0. Samples were pre-pared by mixing crude extracts with the same volumeof sample buffer (stacking buffers diluted with water(1:2) containing 10% glycerol and 0.01% methyleneblue). Electrophoresis was carried at constant voltage(100 V per gel) and 4◦C. After electrophoresis gelswere soaked in: 1) 0.1 M HCl for 15 min; 2) 0.2%haemoglobin in 0.1 M Gly/HCl, pH 2.0 for 30 min at4 ◦C, and for 90 min at 37◦C, and 3) distilled wa-ter. Staining was carried out as previously described.Characterization of protease type in zymograms usingspecific inhibitors was done as detailed previously.

Table 1. Protease activity measured in stomach andpyloric caeca extracts from seabream and dentex.Specific activities of commercial enzymes are in-cluded for comparison. Data are the mean of threedeterminations± SD

Enzyme Activity

(units mg−1)

Porcine pepsin 3168± 110

Seabream stomach extract 740± 80

Dentex stomach extract 894± 59

Porcine trypsin 1426± 78

Bovine chymotrypsin 561± 59

Porcine pancreatin 549± 50

Cod trypsin 60± 3

Seabream pyloric caeca extract 40± 6

Dentex pyloric caeca extract 57± 11

Results

Total proteolytic activities measured in stomach andpyloric caeca extracts of both sparids, as well as indifferent purified animal proteases are shown in Table1. They were high (particularly in the case of acidproteases) when compared to those obtained usingcommercial fish enzymes, considering that determina-tions were carried out using non purified extracts. Pro-tease activity measured in dentex extracts was greaterthan that found in seabream preparations (17 and 42%higher, respectively).

The optimum pH range for protease activities inboth species is shown in Fig. 1. Optimum activity ofacid proteases was measured at pH 2–2.5, decreasedat a higher pH and almost disappeared at pH 5.0.Alkaline protease activity in both sparids was main-tained over a wider pH range (5–11) and showed awell-defined optimum at pH 10.0. Determination ofresidual activity of proteases after incubation at dif-ferent pH was used as a measure of their stability(Table 2a). These results indicated a greater resistanceof acid proteases in dentex to alkalization, since theyretained 80% activity after 1 h of incubation at pH9.0, whereas the activity of seabream acid proteaseswas reduced to a half under such conditions. ThepH-stability of alkaline proteases in both sparids wassimilar, which retained 90–100% activity over a 5–12 pH range, but they were decreased in a more acidenvironment (pH 2.0).

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Figure 1. Effect of pH on protease activity in stomach and pyloric ceca extracts ofSparus aurataandDentex dentex. Data are mean of triplicatedeterminations.

Figure 2. Effect of temperature on protease activity in stomach and pyloric ceca extracts ofSparus aurataandDentex dentex. Data are meanof triplicate determinations.

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Table 2a.Percentage of protease activity in stomach and py-loric ceca extracts of seabream and dentex retained after 60 minincubation at different pH

S. aurata D. dentex

pH Stomach Pyloric ceca Stomach Pyloric ceca

2 100 20 110 20

5 100 90 90 100

7 100 90 90 100

9 55 100 80 100

12 10 100 0 100

Table 2b. Percentage of protease activity in stomach and pyloricceca extracts of seabream and dentex retained after 60 min incuba-tion at different temperatures. Activity measured at 25◦C withoutpreincubation was used as a reference

S. aurata D. dentex

T (◦C) Stomach Pyloric ceca Stomach Pyloric ceca

30 100 110 100 100

40 100 100 80 75

50 90 45 40 45

60 50 3 0 2

Optimum temperatures for acid and alkaline pro-tease activity in both species were 40◦C and 50–55 ◦C, respectively (Figure 2). Seabream acid pro-teases proved to be highly resistant to heating, re-taining 90% of their activity after 60 min incuba-tion at 50 ◦C (Table 2b). In contrast, the activityof dentex acid proteases was significantly affectedby incubation at temperatures over 40◦C. Alkalineproteases, present in both species, were more heatsensitivity, retaining less than 40% of activity afterincubation at 50◦C. Assays performed using seabreamextracts showed marked differences in thermal stabil-ity of the two main intestinal proteases in this species.Chymotrypsin-like proteases were reduced in activ-ity by 25% at 40◦C and denatured at 50◦C, whiletrypsin-like proteases remained completely active atthis temperature, even retaining a 10% of activity at60 ◦C (Figure 3). Kinetic parameters and theoreti-cally calculated energy of activation for both types ofproteases are summarized in Table 3.

Zymograms, performed using stomach extracts ofboth sparids, are shown in Figure 4. Two bands, rep-resenting the major protease activity, (Rf = 0.811 andRf = 0.666), were evident in extracts obtained fromboth species. The first band was better defined in

Figure 3. Effect of temperature on the stability of trypsin andchymotrypsin activities in seabream pyloric ceca extracts.

seabream and the second in dentex. Additional bands,showing different electromobility in each species,were also identified. Zymograms performed on intesti-nal extracts are shown in Fig. 5. More proteases werefound in dentex as seen in the greater number of ca-seinolytic bands. In seabream extracts, 5 active bandswere observed in the range of 24.5–90 kDa, while 8active bands, ranging from 24.5 to 69.5 kDa, were seenin dentex extracts.

Acid protease classes present in stomach extractsof both sparids following the use of several specificinhibitors are shown in Table 4. Both sparids showed

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Figure 4. Substrate – PAGE of acid proteinases in stomach ho-mogenates.Sparus aurata: lanes 1, 3 and 4 = crude extract; lane 2= inhibition with pepstatin A.Dentex dentex: lanes 5 and 6 = crudeextracts; lane 7 = inhibition with pepstatin A. Lanes 1 to 3 in 12%polyacrylamide (PAA) gels. Lanes 4 to 7 in 15% PAA.gels.

Table 3. Summary of apparent kinetics parameters calculated foralkaline proteases in seabream. Data are the mean of three determi-nations± SD

Vmax Km Vmax Km−1 Ea

Substrate (U mg−1) (mM) (Kcal mol−1)

Hemoglobin 820 0.031 26451 9.13

(Pepsin) (12.9) (0.002)

TAME 0.962 0.132 7.3 10.9

(Trypsin) (0.0322) (0.02)

BAPNA 0.060 0.125 0.48 –

(Trypsin) (0.002) (0.013)

BTEE 3.062 0.140 22 9.23

(Chym.) (0.134) (0.025)

Casein 40.160

Alk. protease (6.330) – – –

Table 4. Characterization of acid proteases present in stomachextracts of sea bream and dentex, using specific inhibitors

Percentage of inhibition

Enzyme control

Inhibitor Seabream Dentex Porcine

pepsin

PMSF 7.0± 0.2 9.2± 4.3 5.0± 0.1

SBTI 12.0± 0.3 7.0± 0.5 10.0± 0.5

Pepstatin A 99.0± 1.0 99.0± 1.0 99.2± 0.5

a similar response to inhibitors compared with thatof porcine pepsin, being almost completely inhibitedby pepstatin A (specific inhibitor for aspartic pro-teases). This inhibition was also confirmed by thedisappearance of bands in zymograms (Figure 4). Thecharacterization of alkaline proteases by the use ofinhibitors is summarized in Table 5. Protease activ-ity was found to be mainly due to serine proteases inboth species (50% approx.), but trypsin (sensitive toTLCK) was better represented in seabream extracts,whereas chymotrypsin (sensitive to TPCK and ZPCK)was mainly evident in dentex extracts. The presence ofchelant-sensitive proteases was also detected in bothspecies. Zymograms of alkaline proteases obtained us-ing the same inhibitors are shown in Figure 6. Twomajor groups of caseinolytic bands were identifiedin both species based on their response to inhibition.Proteases (28–42 kDa) were sensitive to TLCK (chy-motrypsin), whereas higher molecular weight forms(42 kDa) were sensitive to ZPCK (trypsin). Generallyspeaking, inhibition of proteases detected in dentexextracts was less specific than that found in seabreamextracts, since a high residual activity in bands of 69.5,61.5 and 56 kDa was found. On the other hand, higherdentex sensitivity to proteases (mainly of trypsin type)to inhibition by EDTA was found when compared toseabream (Figure 6, lane 8).

Discussion

Fish proteases have been the main objective of manyof studies, but it is difficult to compare the results ob-tained in different species because the data are affectedby the use of many different methodologies (Anson1938; Barret 1972; Clark et al. 1986), the state offeeding of experimental animals (fed fish; Kawai andIkeda 1971; Eshel et al. 1993; starved fish; Glass etal. 1989; Dimes et al. 1994), and the type of enzymepreparation (intestinal tissues alone; Benitez and Tiro1982; Jonás et al. 1983; including in the extracts theintestinal contents; Takii et al. 1985).

Currently, there is only one published study deal-ing with digestive proteases in the seabream (Munilla-Morán and Saborido 1996), and there are none aboutdentex. Measures on total protease activity, sensitivityof the different proteases to variations in pH and tem-perature, and characterization of such activities by theuse of several techniques was considered a preliminarystep in order to designin vitro tests for the assessmentof protein digestibility in those species.

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Figure 5. Substrate – SDS – PAGE (10% PAA) of alkaline proteinases from pyloric caeca extracts and enzyme controls. Lane 1 = bovinechymotrypsin, lane 2 = porcine trypsin, lane 3 = cod trypsin, lane 4 =Dentex dentex, lane 5=Sparus aurataand lane 6 = Molecular weightmarkers (MWM): (a) phosphorylaseb (94,000); (b) bovine albumin (67,000); (c) egg albumin (43,000); (d) carbonic anhydrase (30,000).

Table 5. Characterization of alkaline proteases present in seabream and dentex digestive extracts and control enzymes, using specificinhibitors and casein as the substrate. Data shown are mean of 10 determinations± SD. Enzymatic inhibition due to solvents neverexceeded 5%

Percentage of inhibition

Enzyme control

Inhibitor Class Stock Solvent Seabream Dentex Cod Porcine Bovine Thermolisin

proteases concentration trypsin trypsin chymotrypsin

PMSF serin- 100 mM Ethanol 24.3± 4.8 44.5± 9.9 54± 2.8 93± 4.8 100 11.7± 3.3

SBTI serin- 250µM Water 48.9± 6.3 41.3± 8.9 50.5± 3.5 100 97.5± 0.7 2± 1.4

TLCK trypsin-l 10 mM HCl 1mM 16.8± 2.7 6.0± 4.7 2.2± 1.7 99± 1.4 0 8.5± 9.3

CHYM chymot.-l 5 mM DMSO 45.4± 6.7 40.5± 7.5 55.5± 4.9 54± 17 99.0± 1.4 6.5± 4.9

TPCK chymot-l 5 mM Ethanol 19.5± 7.3 26.2± 8.9 31± 21.2 2± 2.8 97.5± 0.7 2

ZPCK chymot-l 10 mM DMSO 26.2± 5.4 36.1± 7.5 50± 17 3.5± 4.9 99.5± 0.7 14± 7.1

EDTA metallo- 0.5 M Water 33.6± 5.1 29.8± 7.7 39.5± 6.4 30 99.5± 0.7 100

EGTA metallo- 0.5 M Water 6.5± 0.5 17.0± 5.6 14± 2.8 13 7.0± 0.1 100

Acid proteolytic activity, carried out by an aspar-tic protease method closely resembling porcine pepsin(Davies 1990), revealed, in zymograms, two bandsthat could be identified as pepsin I (Rf = 0.666) andpepsin II (Rf = 0.811). Such a polymorphism hasbeen previously described in other fish pepsins (Gild-berg 1988). The pH profile of proteases present instomach extracts, showing an optimum activity at pH2–3, closely resembled that previously reported infreshwater fish such as rainbow trout (Oncorhynchusmykiss) (Kitamikado and Tachido 1960); catfish (Silu-

ris glanis) (Jonas et al. 1983), tilapia (Oreochromissp.) (Yamada et al. 1993), sole (Solea solea), turbot(Scophtalmus maximus), halibut (Hippoglossus hippo-glossus) (Glass et al. 1989), or seabass (Dicentrarchuslabrex) (Eshel et al. 1993). In both sparids, acid pro-teases retained 100% activity after a 1 h incubation atpH 5.0 or 7.0. This unusual stability of fish acid pro-teases may be related to their amino acid pattern, witha high proportion of basic residues compared withthose of mammals. Considering both the increases instomach pH resulting from food and water ingestion

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Figure 6. Substrate – SDS – PAGE (10% PAA) of pyloric caeca extracts of A)Sparus aurataand B)Dentex dentex, treated with differentspecific inhibitors. MWM is the same detailed in the previous figure. Extracts were mixed with the inhibitors indicated in the lanes. Lane 1 =MWM, lane 2 = control without inhibitor, lane 3 = PMSF, lane 4 = SBTI, lane 5 = TLCK, lane 6 = TPCK, lane 7 = ZPCK and lane 8 = EDTA.

determined in different fish species (4.3 in seabass(Eshel et al. 1993), 5.2 in eel (Anguilla japonica)(Takii et al. 1985), or around 4.0 in yellowtail (Serioladumerilli) (Caruso and Genovese 1992), and the needof a suitable pH for the activity of acid proteases, itmay be concluded that acid proteolysis will not occuruntil secretion of acid is sufficient to decrease stom-ach pH. The pH stability of seabream or dentex acidproteases is a good adaptation permitting completeprotein digestion.

High alkaline protease activity was detected inboth sparid species when compared with activitiesof cod trypsin (Table 1). The pH of these pro-

teases showed an optimal resembling that of otherfish species, such as rainbow trout (Kristjansson andNielsen 1992; Kitamikado and Tachino 1960), seabass(Eshel et al. 1993), milkfish (Chanos chanos) (Benitezand Tiro 1982), mackerel (Scomber japonicas) (Pyeunand Kim 1986), or flatfish (Glass et al. 1989). Thepresence of two pH peaks (7.0 and 10.0) suggests theexistence of at least two major alkaline proteases in theextract.

Contrary to the main feature of acid proteases, al-kaline proteases in the two species were sensitive toacid, and their activities were markedly reduced, al-most to zero at pH 3.0. Munilla-Moran and Saborido

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(1996) remark on the existence of a clear peak of acidactivity in the intestine, but the authors suggest thatthis may be an artifact. The authors suggest that thepyloric caeca may be related to the need to retain feedfor the neutralization of acid secretion. However, theysuggest that the pyloric caeca are associated with thestorage of alkaline proteases until the acid bolus isneutralized by other pancreatic secretions.

The effect of temperature on digestive protease ac-tivity in seabream and dentex is similar to that forother fish species, such as cod (Boreogadus saida)(Arunchalan and Haard 1985), tilapia (Oreochromisniloticus) (Yamada et al. 1993), dogfish (Scyliorhi-nus canicula) (Guerard and Le Gal 1987), and rain-bow trout (Torrisen 1984). A 10◦C difference wasobserved between maximum activities for acid andalkaline protease activities (45 vs. 55◦C). The mostrelevant result, derived from curves in Figure 2, isthe greater specific activity of both acid and alkalineproteases determined in dentex extracts, which is 1.5to 2.5 times that measured in seabream extracts. Addi-tionally, at 25◦C, (a common temperature in Mediter-ranean fish farms) the differences were more marked,particularly for alkaline proteases. Under such condi-tions, activity determined in dentex extracts was about100 Umg−1 (one third of maximum possible activity)but only 30 Umg−1 in seabream extracts (one sixth ofits maximum activity).

The optimum temperature for enzyme maximumactivity may be interesting for comparative studies,but such data offer little about enzyme activity un-der physiological conditions. Thus, enzyme resistanceto denaturation after incubation at different tempera-tures may be more useful. Thermal stability of acidproteases was greater than that observed in alkalineproteases. After preincubation of stomach proteasesat 60 ◦C for 1 h, 50% of total activity was retained,whereas alkaline proteases was reduced to almostzero. The incubation of seabream extracts in the pres-ence of specific substrates showed clear differences inthermal stability between chymotrypsin and trypsin;after preheating at 50◦C 100% of trypsin activitywas still present but no chymotrypsin were measured.Thus, both alkaline proteases differ not only in theirsubstrate specificity but also in their structure. Froma biological viewpoint it is difficult to deduce any ad-vantage for this fish in possessing proteases showingdifferent resistances to heating, since normal temper-ature of water rarely exceed 28◦C, (a temperaturewhere both proteases show maximum activity). Never-theless, from a biotechnological perspective, it may be

interesting to have information about active and easilydenaturalizable proteases potentially useful in the feedindustry (Haard 1992).

The calculated values of activation energy (appar-ent) (Ea) for acid and alkaline proteases of seabreamranged from 9 to 11 kcal mol−1. The Ea for seabreamintestinal proteases given by Munilla-Morán and Sa-borido (1996) were similar (12.9 kcal mol−1) to thoseseen here but they suggest a ‘break-point’ at 20◦C forstomach protease, suggesting a lower ability to digestprotein in waters below this temperature (18.68 kcalmol−1 vs. 3.04 kcal mol−1). We disagree with sucha conclusion since the former value of Ea determinedfor stomach proteases is not congruent with the habitatof the species, which is in the range of values de-scribed for cold water fish like polar cod (Boreogadussaida), living in water at 1.5 to 2◦C (2.9–3.2 kcalmol−1; Arunchalam and Haard 1985) or salmon (3.5kcal M−1; Norris and Elam 1940). The existence ofsuch a ‘break point’ should imply strange modifica-tions in tertiary structure of the protease molecule ornot very probable alterations in the ionization states ofimportant groups.

Seabream alkaline proteases were studied by mea-suring variations in total activity in relation to larvaldevelopment (Moyano et al. 1996) and assessing therole of endogenous (self produced) and exogenous(present in zooplankton) proteases in larval ability todigest food (Díaz et al. 1997). Comparison of results tothose obtained in the present paper, allows us to con-firm the relative importance in adult fish of trypsin-likeproteases (50% of total activity) previously detected inlarval seabream extracts. There is a clear distinctionbetween the mechanisms of catalysis of mammalianand fish proteases. Cod trypsin was not inhibited bya specific enzyme inhibitor, TLCK, but a 50% reduc-tion in activity was detected in the presence of ZPCK,specific for chymotrypsin. Since a similar result wasobtained when crude extracts of seabream or dentexwere tested, it is suggested that only enzymes obtainedfrom taxonomically related groups should be used asreference.

Although the type of proteases reported in the pa-per of Munilla-Morán and Saborido (1996) are similarto the results shown here, levels of inhibition ob-tained for alkaline proteases in the present study, usingboth biochemical assays and zymograms, were alwayslower than those obtained by the aforementioned au-thors. For example, a high inhibition by EDTA ofboth stomach and intestinal proteases (85 to 93%) wasfound. This may be true for intestinal proteases but it

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is more difficult to understand in the case of pepsin, anaspartic protease typically characterized by an actionmechanism independent of cations, and it is one ofthe main reasons why we suggest that the combina-tion of both specific inhibition assays and zymogramsare used. In dentex, the significant effect of EDTA ontrypsin proteases revealed a clear dependence of suchactivity on divalent cations, being more relevant whencompared with seabream proteases. Three differentaction mechanisms for dentex proteases are possible,instead of the two found in seabream proteases, in-cluding a lower susceptibility of dentex proteases tothe different inhibitors existing in raw materials uti-lized as feed ingredients.

Acknowledgements

This research was supported by the C.I.C.Y.T., ProjectMAR 95-1943-CO3-03. We are also grateful to Mr.Francisco Ruiz, from FRAMAR S.A., and to thetechnicians working in the I.E.O. Center of Mazarrón(Murcia), for their kind provision of adult fish.

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