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Bovine metalloprotease characterization and in vitro connective tissue degradation¹

A. L. Alderton^*, W. J. Means^,2, N. Kalchayanand, R. J. McCormick and K. W. Miller

^* Department of Animal and Food Sciences, University of Kentucky, Lexington 40546 and and Department of Animal Science and and Department of Molecular Biology, University of Wyoming, Laramie 82071

Abstract

Metalloproteases that selectively hydrolyze connective tissue proteins may tenderize meat without creating texture problems associated with myofibrillar protein degradation. Our objective was to characterize the activity of bovine placental proteases to determine whether they can improve meat tenderness through disruption of the connective tissue matrix. Enzymes were extracted, crudely purified, and proteolytic activity was assessed against gelatin and collagen under varying pH and temperature conditions using both SDS-PAGE and zymography. Gelatin zymography revealed proteolysis between 57 and 63 kDa, with decreased activity as buffer pH decreased from pH 7.4 to 5.4 (37°C). Proteolytic activity was pronounced at 37°C, moderate at 25°C, and absent at 4°C following 48-h incubation (pH 7.4). Placental enzymes were metalloproteases inhibited by excess EDTA. Maximum proteolysis was achieved in the presence of Ca²⁺, with or without Mg²⁺ and Zn²⁺. Absence of Ca²⁺ decreased proteolytic activity. Complete degradation of both the 125- and 120-kDa proteins of the -chains of gelatin was achieved following enzyme incubation for 6 h at 37°C or 24 h at 25°C. No degradation was observed following enzyme incubation with native Type I collagen. Given the marked decrease in enzyme activity at pH 5.4 and 4°C (standard industry conditions), bovine placental metalloproteases would not be expected to contribute to connective tissue degradation or improve meat tenderness.

Key Words: Bovine • Collagen • Collagenase • Enzyme • Meat Tenderness • Metalloprotease

Introduction

Tenderness is one of the most important factors affecting consumer perception of beef (Morgan et al., 1991; Brooks et al., 2000) with estimated annual economic losses of $217 million (Smith et al., 1995). Meat texture is dictated by myofibrillar and connective tissue properties (Bailey, 1972), with the former receiving greater research focus (Penny, 1980; Goll et al., 1983; Koohmaraie et al., 1988; Koohmaraie, 1994). Collagen plays a significant role in meat texture and is thought to contribute a fixed amount of background toughness to meat (Bailey and Light, 1989). Plant (bromelin, ficin, and papain; Kang and Rice, 1970; Stanton and Light, 1987) and bacterial (collagenase; Foegeding and Larick, 1986; Berry and Shuttleworth, 1988) enzymes have been evaluated for use in postmortem meat tenderization, but neither is extremely effective.

No clear evidence for collagen degradation during aging exists; however, recent research identified structural weakening of perimysium and endomysium during prolonged aging (Takahashi, 1996; Nishimura et al., 1998). In an effort to elucidate a method to disrupt the connective tissue matrix and subsequently improve tenderness, Phillips et al. (2000) purified a bovine placental protease capable of gelatin and Type I acid-soluble collagen degradation; actin and myosin heavy chain remained intact. In vitro, the enzymes degrade the collagenous placental cotyledon-caruncle connection (Woolley and Evanson, 1980). The concept that meat tenderness could be improved by application of collagen-specific proteases deserves further study. Our objective, therefore, was to characterize bovine placental protease activity to determine whether study in meat systems was warranted. More specifically, our goals were to 1) determine whether bovine placental proteases were metalloproteases requiring metal ions for activity and 2) further characterize proteolytic activity of the proteases against gelatin and collagen across varying pH and temperature conditions.

Materials and Methods

Materials
All chemicals were purchased from Sigma Chemical Co. (St. Louis, MO) unless stated otherwise. Bovine placentas (n = 6) were obtained within 3 h of clearance (Sas Dairy, Laporte, CO; University of Wyoming Beef Unit, Laramie), stored on ice, and transported to the laboratory.

Enzyme Extraction and Purification
Extraction procedures of Phillips et al. (2000) were followed, with modified procedures of Dean and Woessner (1984). Briefly, placentas were washed, cotyledons removed, cleared of any visible vascular or connective tissues, and homogenized. The homogenate was subsequently heated (60°C) to release membrane-bound enzymes and cooled, and the supernatant collected via centrifugation (20 min, 10,000 x g, 4°C). Enzymes were collected using a two-stage ammonium sulfate (Fisher Scientific, Fair Lawn, NJ) precipitation (20 and 70% saturation), and resuspended in buffer (25 mM Tris•OH, 10 mM CaCl₂•2H₂O, 0.2 M NaCl, 0.05% Brij-35, and 1 mM phenyl methylsulfonyl fluoride; pH 7.4; 4°C). Enzymes collected from each extraction were pooled, and protein content was determined using the bicinchoninic acid protein assay procedure (Smith et al., 1985).

Enzyme Activation
Enzymes were concentrated using a Speed Vac Concentrator (Savant Instruments, Inc., Farmingdale, NY) to contain equal amounts of protein (4.3 mg/mL). To achieve maximal activity, latent enzyme forms were subsequently activated by addition of p-aminophenylmercuric acetate to a final concentration of 0.5 mM followed by a 4-h incubation at 37°C (Cawston and Murphy, 1981; Murphy et al., 1981).

SDS-PAGE and Zymography
Discontinuous electrophoresis procedures were followed as described by Laemmli (1970) using 7.5% polyacrylamide separating gels and 4% polyacrylamide stacking gels (1.0 mm x 8.0 cm x 7.0 cm; Mighty Small SE245 Dual Gel Caster; Hoefer/Pharmacia, Piscataway, NJ). Activated enzyme samples used for zymography were quenched with a final concentration of 2 mM EDTA, diluted 1:1 (vol/vol) with nonreducing sample buffer (0.125 M Tris•HCl, 5% SDS, and 20% glycerol; pH 6.8), and incubated for 30 min (25°C) before loading on the gels. Collagenase Type L from Clostridium histolyticum was loaded on the gels to serve as a positive control. Enzyme fractions were also assayed by electrophoretic analysis in polyacrylamide gels containing SDS and copolymerized gelatin (zymography). Modified procedures of Heussen and Dowdle (1980) were used to prepare 0.05% gelatin zymograms by substituting the appropriate amount of gelatin (Type A from porcine skin, solubilized in deionized water) for the water component of the gel, all gels were run at a constant current (14 mA for stacking gel increased to 20 mA for separating gel) with cold (10 ± 2°C) water circulation using a Hoefer electrophoresis unit (SE 250; Hoefer/Pharmacia, Piscataway, NJ). Following electrophoretic separation, enzymes were renatured as previously described, and recovered activity was identified by the presence of a clear zone where the incorporated substrate was hydrolyzed and subsequently rinsed from the gel (Lacks and Springhorn, 1980; Gabriel and Gersten, 1992). Gels were fixed and stained using Coomassie blue R-250, and, to provide more contrast to the clear bands of enzyme activity, zymograms were not destained. Approximate molecular weights (MW) were calculated relative to the high MW standard (SDS-6H) using relative mobility (R_f) values. When appropriate, a GS700 imaging densitometer (Bio-Rad, Rockland, CA) was used to compare SDS-PAGE band intensity. However, due to variation in the background color of gels and zymograms, densitometer results were complicated and prevented the quantification of data.

pH Treatment
To determine enzymatic activity as a function of pH following electrophoresis, SDS was washed from incorporated gels (n = 3 for each pH assayed), and enzymes present were allowed to renature and incubate (Lacks and Springhorn, 1980). Sodium dodecyl sulfate was removed by gently shaking the gel for 2 h in buffer (50 mM Tris•OH, 5 mM CaCl₂•2H₂O, 5 mM MgCl₂, 1 µM ZnCl₂, and 2.5% [vol/vol] Triton X-100; 25°C) adjusted to a pH of 5.4, 6.4, or 7.4. Gels were then rinsed in deionized water (25°C) for 5 min and subsequently allowed to incubate in the appropriate pH buffer for 2 h at 37°C.

Temperature Treatment
Following electrophoresis, SDS was removed by gently shaking the gels (n = 3 for each temperature assayed) for 2 h in buffer (50 mM Tris•OH, 5 mM CaCl₂•2H₂O, 5 mM MgCl₂, 1 µM ZnCl₂, and 2.5% (vol/vol) Triton X-100; pH 7.4) adjusted to 4, 25, or 37°C to determine enzymatic activity at varying temperatures. Gels were then rinsed in deionized water (25°C) for 5 min, and subsequently allowed to incubate in pH 7.4 buffer for the following times: 4°C for 48 h; 25°C for 24 h; and 37°C for 2 h.

Metalloprotease Identification
In order to broadly characterize the enzymes as metalloproteases, it was necessary to confirm that metal ions are a prerequisite for activity (Stricklin and Hibbs, 1988). Gelatin-incorporated gels (n = 3 for each buffer assayed) were cast as previously described. Following electrophoresis, the gels were washed by shaking gently for 2 h in buffer (50 mM Tris•OH, and 2.5% [vol/vol] Triton X-100; pH 7.4; 25°C) and rinsed for 5 min in deionized water (25°C). Gels were then exposed to one of the following five treatment buffers and shaken gently at 37°C for 2 h to allow for detection of enzymatic activities:

1. 50 mM Tris•OH, 1.0% Triton X-100, 10 mM CaCl₂, 5 mM MgCl₂, and 1 µM ZnCl₂ (pH 7.4)
   
2. 50 mM Tris•OH, 1.0% Triton X-100, 10 mM CaCl₂, 5 mM MgCl₂, 1 µM ZnCl₂, and 5 mM EDTA (pH 7.4)
   
3. 50 mM Tris•OH, 1.0% Triton X-100, and 10 mM CaCl₂ (pH 7.4)
   
4. 50 mM Tris•OH, 1.0% Triton X-100, and 5 mM MgCl₂ (pH 7.4)
   
5. 50 mM Tris•OH, 1.0% Triton X-100, and 1 µM ZnCl₂ (pH 7.4)
   

Gelatin and Type I Collagen Degradation
Our enzyme extract was incubated with gelatin or Type I collagen before SDS-PAGE (7.5%; n = 3 for each substrate assayed). Equal proportions (15 µg) of activated enzyme and substrate (gelatin or Type I collagen) were incubated in buffer (50 mM Tris•OH, 5 mM CaCl₂•2H₂O, 5 mM MgCl₂, 1 µM ZnCl₂, and 1.0% [vol/vol] Triton X-100; pH 7.4) at either 25°C (gelatin and Type I collagen) or 37°C (gelatin) for 0, 1, 6, 12, or 24 h before quenching with EDTA (2 mM final concentration). Gels were fixed and stained with Coomassie blue R-250.

Results and Discussion

Zymograms, which contain substrate in the gel matrix, allowed for background staining and did not appear to alter protein migration. When gelatin-incorporated SDS-PAGE of the enzymes were performed and gels subsequently incubated at either a pH of 5.4, 6.4, or 7.4 (37°C, 2 h), clear zones were revealed in a MW range of 57 to 65 kDa (Figure 1). Multiple zones are clearly evident in Lanes 3 and 5 of Figure 1, indicating the presence of multiple enzymes, or at least different isoforms of the same enzyme. The MW range of activity was consistent with that reported by Phillips et al. (2000). The activity observed was most pronounced in the gel incubated at pH 7.4 and least pronounced in the gel incubated at pH 5.4, with moderate activity observed in the gel incubated at pH 6.4. Visual assessment was similar to that of imaging densitometry results. Bacterial collagenase, used as a positive control to confirm enzyme renaturation system effectiveness, showed similar results (Figure 1). Additional gelatin-incorporated gels were electrophoresed and subsequently incubated (pH 7.4) at 4, 25, or 37°C for 48, 24, or 2 h, respectively. Again, clear zones were revealed in MW ranges comparable to those previously reported (Phillips et al., 2000). Visual and densitometer evaluation of the gel incubated at 4°C showed no apparent enzymatic activity following 48 h of incubation (Figure 2). However, the gels incubated at 25°C for 24 h revealed pronounced enzymatic activity, similar to that seen following incubation at 37°C for 2 h (Figure 2).

View larger version (55K): [in this window] [in a new window]   Figure 1. The SDS-PAGE (7.5%) of 0.05% gelatin zymogram. Lanes 1, 3, and 5 contained 26 µg of activated placental enzyme incubated at 37°C for 2 h in pH 5.4, 6.4, and 7.4 buffer, respectively. Lanes 2, 4, and 6 contained 10 µg of bacterial collagenase incubated at 37°C for 2 h in pH 5.4, 6.4, and 7.4 buffer, respectively. The arrow indicates the 57- to 65-kDa region of enzymatic activity.

View larger version (78K): [in this window] [in a new window]   Figure 2. The SDS-PAGE (7.5%) of 0.05% gelatin zymogram. Lanes 1, 3, and 4 contained 26 µg of activated placental enzyme incubated at 4, 25, or 37°C for 48, 24, or 2 h, respectively. Lane 2 contained 10 µg of bacterial collagenase incubated at 4°C for 48 h. All lanes were incubated at pH 7.4. The arrow indicates the 57- to 65-kDa region of enzymatic activity.

To classify an enzyme in the broad class of metalloproteases, a requirement for metal ions, such as Ca²⁺, Mg²⁺, or Zn²⁺, for activity must be demonstrated (Stricklin and Hibbs, 1988). In addition, it should be shown that chelating agents that bind metal ions, such as EDTA, inhibit hydrolytic activity (Cawston and Murphy, 1981; Stricklin and Hibbs, 1988). Gelatin-incorporated gels incubated with limited concentrations of metal ions were used to characterize the enzymes as metalloproteases. Figure 3 confirms gelatinolytic activity of the enzymes when incubated with excess CaCl₂, MgCl₂, and ZnCl₂. With the addition of sufficient EDTA to decrease cation concentrations, the majority of enzymatic activity was inhibited and only faint enzyme activity was observed. When gels were incubated in buffer with excess CaCl₂, intense activity was observed. When only MgCl₂ was present in the incubation buffer, enzyme activity was dramatically reduced. Activity was least evident when only ZnCl₂ was present in the incubation buffer (Figure 3).

View larger version (63K): [in this window] [in a new window]   Figure 3. The SDS-PAGE (7.5%) of 0.05% gelatin zymogram. All gels were incubated at 37°C, pH 7.4 for 2 h. Lane 1 contained 5 µL of high-molecular weight (MW) standard; Lane 2 contained 26 µg of activated placental enzyme incubated in a buffer containing CaCl2, MgCl2, and ZnCl2; Lane 3 contained 26 µg of activated placental enzyme incubated in a buffer containing CaCl2, MgCl2, and ZnCl2 and sufficient EDTA to bind metal ions; Lane 4 contained 26 µg of activated placental enzyme incubated in a buffer containing only CaCl2; Lane 5 contained 26 µg of activated placental enzyme incubated in a buffer containing only MgCl2; and Lane 6 contained 26 µg of activated placental enzyme incubated in a buffer containing only ZnCl2. The arrow indicates the 57- to 65-kDa region of enzymatic activity.

The MW range where hydrolytic activity occurred corresponds with reported values for other mammalian metalloproteases. Stricklin and Hibbs (1988) described rat uterus collagenases with MW of 60 and 64 kDa. They also identified human fibroblast collagenases with MW of 55 and 60 kDa, and Type IV collagenases with MW of 62 and 68 kDa. Because of the remarkably conserved nature of collagenases derived from the diverse sources previously mentioned (Stricklin and Hibbs, 1988), it is likely that enzyme extracts used in this research belong to the same broad family of mammalian matrix metalloproteases.

Metalloproteases can be generally classified on the basis of their substrate specificity: 1) stromelysins that preferentially degrade proteoglycans and basement membrane components; 2) gelatinases that preferentially degrade gelatin; and 3) collagenases that preferentially degrade collagen (Okada et al., 1986). Following incubation (at 25 and 37°C) of extracted enzymes with gelatin substrate, SDS-PAGE revealed the gradual disappearance of 125- and 120-kDa protein bands over time (Figure 4). Incubation of the enzymes with gelatin at 25°C for 6 h resulted in an approximately 50% decrease in the 125- and 120-kDa bands. After 24 h of incubation, the bands almost completely disappeared. In comparison, gelatin incubated with extracted enzymes at 37°C for 1 h revealed a dramatic decrease of intensity in the 125- and 120-kDa bands. In addition, after 6 h of incubation, the bands disappeared. This gel supports results from gelatin zymograms indicating that enzyme fractions contain gelatinolytic enzymes. For control purposes, SDS-PAGE of gelatin incubated at 25 or 37°C for 24 h (Lanes 3 and 14, respectively; Figure 4) revealed no visible changes in protein bands indicating no hydrolysis from contaminant protease sources.

View larger version (70K): [in this window] [in a new window]   Figure 4. The SDS-PAGE (7.5%) of gelatin incubated with placental enzyme at both 25 and 37°C. Lane 1 contained 5 µL of high–molecular weight (MW) standard; Lane 2 contained 15 µg of activated placental enzyme; and Lane 3 contained 15 µg of gelatin incubated at 25°C for 24 h. Lanes 4 through 8 contained 15 µg of activated placental enzyme and 15 µg of gelatin incubated at 25°C for 0, 1, 6, 12, or 24 h, respectively, whereas Lanes 9 to 13 contained 15 µg of activated placental enzyme and 15 µg of gelatin incubated at 37°C for 0, 1, 6, 12, or 24 h, respectively. Lane 14 contained 15 µg of gelatin incubated at 37°C for 24 h. The arrow pair indicates the 125- and 120-kDa protein bands.

View larger version (65K): [in this window] [in a new window]   Figure 5. The SDS-PAGE (7.5%) of Type I collagen incubated at 25°C with placental enzyme. Lane 1 contained 5 µL of high–molecular weight (MW) standard; Lane 2 contained 15 µg of activated placental enzyme; Lane 3 contained 15 µg of Type I collagen; and Lanes 4 through 8 contained 15 µg of activated placental enzyme and 15 µg of Type I collagen following 0, 1, 6, 12, and 24 h incubation at 25°C, respectively. The arrow pair indicates the 125- and 120-kDa protein bands.

On the other hand, Phillips et al. (2000) demonstrated that bovine placental crude extracts were able to degrade substrate in Type I collagen-incorporated gels. We suspect that the variance in results may be due to the physical state of collagen in these gels. Possibilities include partial denaturation of collagen substrate to gelatin at 37°C incubation temperatures, causing a decrease in specificity during our initial study (Woolley and Evanson, 1980; Welgus et al., 1981; Bailey and Light, 1989); possible tertiary structure effects of the free radical-generating environment on the substrate during polyacrylamide polymerization (Garfin, 1990); or effects of the SDS induced negative charges on substrate degradation.

Implications

Additional means of improving tenderness would prove beneficial to the meat industry, especially if collagen could be preferentially degraded. The placental proteases studied in the present experiment did not effectively hydrolyze connective tissue under conditions similar to those found in meat during storage. As a result, it is unlikely that the placental enzymes would be useful in improving meat toughness attributed to connective tissue proteins.

Footnotes

¹ This project was supported by the Univ. of Wyoming Agric. Exp. Stn. In addition, we thank Sas Dairy of LaPorte, CO, and the Univ. of Wyoming Beef Unit for the generous collection of bovine placentas during this research.

² Correspondence: Box 3684 (phone: 307-766-5283; fax: 307-766-2355; e-mail: means{at}uwyo.edu).

Received for publication August 26, 2003. Accepted for publication January 30, 2004.

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