Comp. Biochem. Physiol., 1975, VoL 5011,pp. 287 to 290. Pergamon Press. Printed in Great Britain
ISOZYMIC PATTERNS OF LACTATE DEHYDROGENASE IN WHOLE EMBRYOS AND ADULT TISSUES OF THE MEXICAN AXOLOTL DELAINE GILCREASE* AND J. T. JUSTOSt:~ Department of Zoology, Arizona State University, Tempe, Arizona 85281, U.S.A. (Received 19 November 1973)
Abstract--1. The objective of this study was to determine the general ontogeny of lactate dehydrogenase isozymes in whole embryos and various adult tissues of the Mexican axolotl. 2. The LDH isozymes were resolved by polyacrylamide disc electrophoresis. 3. LDH from embryos showed predominantly anodal activity. 4. Skeletal muscle LDH patterns also showed chiefly anodal activity. This result differs substantially from previously reported studies. 5. We suggest that the patterns of LDH isozymes from skeletal muscle of this aquatic salamander may reflect a greater availability of oxygen to this tissue than in terrestrial animals of this type.
INTRODUCTION
DURING the past two decades, extensive research concerning lactate dehydrogenase (LDH) has provided a wealth of information concerning its enzymatic activity. Not only was L D H found to exist as an isozyme in tissues, but it also exhibited tissue specificity for the isozyme patterns (Markert, 1963). Five distinct L D H isozymes have been demonstrated electrophoretically in most tissues from birds and mammals (Calm et aL, 1962; Markert & Ursprung, 1962). These isozymic bands were arbitrarily numbered one to five, from anodal to cathodal ends of the gel. These uniform patterns found among mammals contrast conspicuously with the great diversity found among amphibia. Most examined amphibia have either more or fewer than the predicted five isozymes (Adams & Finnegan, 1965; Wright & Moyer, 1966; Balek & Snow, 1967; Chen, 1968; Moyer et al., 1968; Wright & Moyer, 1968; Claycomb & Villee, 1971). The predominant type of L D H found in tissues may be related to the availability of oxygen to that tissue. There may be a direct correlation between tissues subjected to anaerobic metabolism (e.g. skeletal muscle) and high concentrations of LDH-5 (Markert, 1963). On this basis, Markert (1963) suggested that the epigenetic control of isozymes * Recipient of a summer fellowship from the Arizona Heart Association. t Established investigator for the American Heart Association. This work was supported by NIH Grant HL 15054-02 to J. T. Justus. ~:Reprint requests should be sent to Dr. J. T. Justus.
involves the oxygen tension in the cell. This suggestion also extends to embryonic forms. Mammalian embryos have relatively poor supplies of oxygen and the embryonic tissues have large amounts of LDH-5 (Markert & Ursprung, 1962). On the other hand, Calm et al. (1962) observed a preponderance of LDH-1 isozymes in bird embryos in accordance with their relatively aerobic environment. The adult isozyme patterns are similar in birds and mammals, but this terminal condition is reached from opposite starting points, from LDH-5 in mammals and from LDH-1 in birds. This seems to demonstrate that as the availability of oxygen changes during development, there is a corresponding change in the L D H isozyme pattern. Furthermore, the results of Balek & Snow (1967) indicate the changes observed during amphibian development more closely resemble the changes found in chick embryos than those found in mammalian embryos. The objective of the present study was to determine the general ontogeny of L D H isozymes in whole embryos and various adult tissues of the Mexican axolotl, A m b y s t o m a mexicanum. Balek & Snow (1967) previously reported changes of L D H isozymes in whole embryos and adult tissues of this animal. Our studies of the changes in these isozymes during the early development of the axolotl are similar to those of Balek & Snow (1967); however, in adult tissues of this salamander, we found fundamental differences in the L D H patterns from those reported by the above-mentioned authors. MATERIALS AND METHODS Embryos were obtained from spawnings of Mexican axolotls. The embryos were maintained at 14°C to
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prolong each developmental stage. Assignment of developmental stages of the urodele embryos were determined by comparison with Harrison's tables for Arabystoma maeulatum (Hamburger, 1969). The following developmental stages were used in this study--unfertilized eggs, stage 33 (preheartbeat), stage 34 (initiation of heartbeat) and stage 41 (hatching). In addition, young larvae which had been feeding for 4 months and adult animals were used. Jelly coats from eggs and embryos at selected stages were removed using watchmaker's forceps. Eggs and whole embryos were sonicated in cellulose nitrate tubes for approximately 1 min with the small tip of a Heat Systems-Ultrasonic Model W140D Sonifier Cell Disrupter. The tubes were kept in an ice-bath during sonication to retard denaturation. One whole embryo or egg was sonicated in 100 p.1 of a sodium phosphate buffer (0.05 M, pH 7"5) containing 0.25 M sucrose. All juvenile and adult tissues were homogenized at a ratio of 1 : 20 (w/v) in the same buffer using a Potter-Elvehjem glass homogenizer equipped with a motor driven pestle. All homogenares were clarified for electrophoresis by centrifugation for 10 min at 30,000 g in a Sorvall RC2-B refrigerated centrifuge. The supernatants were used immediately for electrophoresis. Twenty p.1 of guinea-pig LDH standard (Sigma), a control for all electrophoretic separations, was diluted with 1"0 ml of sodium phosphate buffer (0'05 M, pH 7.5). The protein concentration of several crude extracts was determined by the procedure of Murphy & Kies (1960). Both Davis (1964) and Gabriel (1971) reported the total amount of protein in a crude extract applied to a polyacrylamide gel should not exceed 200/~g. The amount of protein applied to the gels in our experiments did not exceed this figure. The L D H isozymes of all homogenates were resolved by polyacrylamide disc electrophoresis according to the method of Davis (1964). Electrophoresis was carried out at 4°C at 4-5 m A per tube for approximately 1 hr, utilizing a Hoefer Model D E 102 Polyacrylamide Gel Unit connected to a Heathkit Model IP-I 7 power supply. The leads were connected such that the cathode was in the upper tray buffer and the anode in the lower tray buffer. Migration of the protein is from cathode to anode. Completion time was determined by a band of bromphenol blue reaching the bottom of the separating gel. To determine whether any enzyme migration was going into the upper tray buffer, the electrodes were reversed in some experiment s. A m m o n i u m persulfate has been reported as a possible source of artifacts because of its oxidation effect on protein sulfhydryl groups (Mitchell, 1967; Gabriel, 1971 ). Both of these authors suggest that pre-electrophoresis of
the separating gel for 2 h r at 4-5 mA per tube will essentially remove the ammonium persulfate. Comparison of resolution of the LDH isozymes on prewashed and non-prewashed gels showed no difference in electrophoretic separation except for mobility; subsequently, pre-electrophoresis was not performed. Following completion of electrophoresis, the gels were removed from the tubes and placed in small test-tubes in preparation for staining. The positive LDH stain, a slight modification of the method of Rowe (1971), was prepared in the dark immediately before staining. Control gels were run, but in the absence of the substrate, lactate, to detect any activity of "nothing dehydrogenase" (Shaw & Koen, 1965). Enzyme activity was present on control gels, incubated in stain lacking the substrate, corresponding to the bands of LDH activity localized with the positive stain. A negative stain was used to supplement these results and to rule out artifacts of the positive stain due to "nothing dehydrogenase". The negative stain was prepared according to Burke (unpublished) in the following way: Solution A--substrate solution: sodium pyruvate (Sigma), 100.0mg; N A D H (Sigma), 50'0rag: 0-1 M sodium phosphate (pH 7.5), 25.0 ml. Solution B--staining solution: nitro blue tetrazolium (NBT), 5-0mg; phenazine methosulfate (PMS), 0.3 ml: 0.1 M sodium phosphate buffer (pH 7-5), 25.0 ml. Solution B was protected from the light. The gels were first incubated with solution A at room temperature for 15 min, thoroughly rinsed with water and incubated in the dark for 15 rain with solution B. The LDH present in the gel will convert the components in solution A to lactic acid and N A D and at all other regions the pyruvate and N A D H will diffuse into the gel. When solution B is added, areas of LDH activity will show clear bands but the rest of the gel with NADH will be oxidized with the subsequent electron transfer through PMS to NBT which will precipitate as an insoluble purple formazan. At the end of all staining procedures, the gels were rinsed with tap water and stored in 7.0~ acetic acid in the dark. RESULTS T h e m a t e r n a l L D H b a n d s o f unfertilized eggs of A. mexicanum were c o m p a r e d with whole e m b r y o s f r o m stage 33 (preheartbeat), stage 34 (initiation of h e a r t b e a t ) a n d stage 41 ( h a t c h i n g ) (Fig. 1). This c o m p a r i s o n suggests t h a t the m a t e r n a l pattern persists at least until hatching. All three gels exhibit two darkly staining a n o d a l b a n d s a n d three lighter
Fig. 1. Electrophoresis pattern of L D H isozymes from unfertilized egg and developing embryos. EGG, Unfertilized egg; ST 33, stage 33 (preheartbeat); ST 34, stage 34 (initial heartbeat); HATC, stage 41 (hatching). Fig. 2. Electrophoretic patterns of LDH isozymes from skeletal muscle of juvenile and adult axolotls. SMj, Juvenile; SM A, adult. Fig. 3. Electrophoretic patterns of LDH isozymes from various axolotl tissues. H, Heart; SM, skeletal muscle; L, lung; SPL, spleen. Fig. 4. Electrophoretic patterns of LDH isozymes from various axolotl tissues. BR, Brain; Sk, skin; STOM, stomach; LIV, liver.
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LDH in embryonic and adult tissues of the axolotl bands toward the cathode. One of the lighter bands seems to be missing on gels of hatched embryos. Whether it is missing due to poor resolution or to a change in the pattern at this stage is not clear. The skeletal muscle patterns from juvenile and adult animals were compared (Fig. 2). Both juvenile and adult skeletal muscle showed a preponderance of anodal activity, atypical of the skeletal muscle pattern found in most vertebrate species. The adult isozyme patterns from the heart, skeletal muscle, lung, spleen, brain, skin, stomach and liver are shown (Figs. 3 and 4). All these tissues show a greater amount of anodal activity. Since Balek & Snow (1967) have reported that skeletal muscle of the axolotl seemed to show more cathodal activity than we observed, the electrodes of the gel unit were reversed to determine whether any protein migration into the upper tray buffer had occurred. Since the electrodes were reversed, any LDH originally migrating into the upper tray buffer should go into the gel. All of the above axolotl tissues were run with reversed leads and no activity was detected on the gel. Control gels stained without the substrate, lactate, showed activity corresponding to the bands of LDH activity seen on gels stained with positive stain. Even the electrophoretically pure guinea-pig heart LDH standard showed bands of activity when stained without lactate in the mixture. In an effort to determine if these bands of activity were LDH isozymes, a negative stain was used. A comparison of positive, negative and control gels without lactate are shown (Figs. 5-7). Other control gels stained with a mixture lacking both lactate and NAD showed no activity. Guinea-pig heart LDH standard was utilized as a control in all electrophoretic separations. Its isozyme pattern is typical of mammalian heart LDH pattern with predominantly anodal activity (Fig. 7). As a control for our methods of tissue disruption, homogenizing buffer and electrophoretic system, the same methods were employed in preparation and electrophoretic separation of tissues from a laboratory mouse. The LDH patterns from the mouse heart, skeletal muscle and kidney are shown (Fig. 8). Sub-banding was present in both heart and skeletal muscle, an observation commonly noted in the
289
literature (Costello & Kaplan, 1963; Koen & Shaw, 1965; Koen, 1967; Dudman & Zemer, 1969). Despite the sub-banding, the characteristic isozymic pattern was evident, with heart and kidney showing anodal activity and skeletal muscle showing cathodal activity, patterns similar to those found by Markert & Ursprung (1962). DISCUSSION
The ontogeny of LDH isozymes of amphibians and fish closely resemble changes found in the chick embryo, with the initial activity at the anode followed by a gradual increase of activity toward the cathode as development proceeds (Goldberg et al., 1961; Balek &Snow, 1967; Chen, 1968; Moyer et al., 1968; Claycomb & Villee, 1971). Similarly, our results on axolotl embryos showed greater activity at the anode, and in this regard are consistent with previously published observations. Five bands of LDH activity were resolved in the adult axolotl heart with most of the activity at the anode. This is a typical pattern found in the hearts of vertebrates. Isozyme patterns from other axolotl tissues, including lung, spleen, brain, skin, stomach and liver, showed a preponderance of anodal activity. The axolotl heart isozyme pattern was typical of vertebrate hearts; the skeletal muscle LDH pattern observed was quite atypical. Both juvenile and adult skeletal muscle patterns showed predominantly anodal activity, whereas most other vertebrate skeletal muscle patterns show predominantly cathoda1 activity. Balek & Snow (1967) reported axolotl skeletal muscle to have predominantly cathodal activity. They used starch gel electrophoresis with a Tris-borate buffer, pH 8.6. Our results using polyacrylamide gel electrophoresis differ from those reported by these authors. Several experimental conditions in our study suggest that the anodal activity is peculiar to axolotl skeletal muscle. (1) Skeletal muscle patterns from both neotenous and adult Ambystoma tigrinum, a urodele similar to A. mexicanum, showed isozyme patterns with the concentration of activity at the anode (unpublished). (2) In our studies, characteristic tissue patterns for the laboratory mouse were
Fig. 5. Comparison of electrophoretic patterns from axolotl heart on positively stained, negatively stained and control gels lacking substrate. POS, Positively stained; NEG, negatively stained; NO SUB, control gel lacking substrate in stain. Fig. 6. Comparison of electrophoresis patterns from axolotl skeletal muscle on positively stained, negatively stained and control gels lacking substrate. POS, Positively stained; NEG, negatively stained; NO SUB, control gel lacking substrate in stain. Fig. 7. Comparison of electrophoretic patterns from guinea-pig heart LDH standard on positively stained, negatively stained and control gels lacking substrate. POS, positively stained; NEG, negatively stained; NO SUB, control gel lacking substrate in stain. Fig. 8. Electrophoretic pattern of LDH isozymes from mouse heart, skeletal muscle and kidney. H, Heart; SM, sketleal muscle; KID, kidney.
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obtained by using the same experimental conditions as for amphibian tissue. The mouse tissue patterns showed the typical isozyme bands, with heart and kidney showing predominantly anodal activity and skeletal muscle with concentrated cathodal activity. (3) LDH activity was not lost into the upper tray buffer. (4) Furthermore, loss of enzyme activity due to heat inactivation should not have been a problem in the present study because electrophoresis was performed at 4°C. The results presented in this paper provide convincing evidence that the electrophoretic separation of skeletal muscle isozymes with the predominant LDH activity concentrated at the anode is peculiar to the Mexican axolotl. Low oxygen tension favors synthesis of slower migrating isozymes, the form best suited for anaerobic metabolism and found in high concentrations in skeletal muscle from most vertebrates (Dawson et al., 1964). This information suggests that skeletal muscle from the axolotl may have a greater availability of oxygen than skeletal muscle from other vertebrates. This does not seem unreasonable in the light of the observation that about half of the respiratory gas exchange takes place across the skin in A. mexicanum (Szarski, 1964). Perhaps the large amount of anodal activity seen in axolotl skeletal muscle reflects a well-oxygenated tissue exhibiting aerobic metabolism. Experiments are being designed, using oxygen microelectrodes, to test this possibility.
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D AWSON D., G~~DFRIEND T. L. & KAPLAN N. 0. (1964) Lactic dehydrogenases: functions of the two other types. Science, Wash. 28, 929-933. DUDMAN N. & ZERNER B. (1969) Simplifications of mouse lactate dehydrogenase electrophoretic patterns. Biochim. biophys. Acta 171, 195-197.
GABRIEL 0. (1971) Analytical disk gel electrophoresis. In Methods in Enzymology (Edited by COLOWICK S. P. & K APLAN N. 0. Vol. 22, pp. 565-578. Academic Press, New York. G OLDBERG E., CUERRIER J. P. & WARD J. C. (1969) Lactate dehydrogenase ontogeny, paternal gene activation, and tetramer assembly in embryos of brook trout, lake trout, and their hybrids. Biochem. Gen. 2, 335_~ 350.
H A M B U R G E R V. (1969) A Manual of Experimental Embrvoloav. University of Chicago Press. Chicago K OEN A. Lyi1967) Lactate dehydrogenase isozyme-&bbands: new bands derived from isolated sub-bands. Biochim. biophys. Acta 140, 496-502.
K OEN A. L. & SHAW C. R. (1965) Studies on lactate dehydrogenase tissue-specified isozymes: electrophoretie migration of isolated sub-bands. Biocbim. biophys. Acta 96, 231-236.
MARKERT C. L. (1963) Epigenetic control of specific protein synthesis in differentiating cells. In Cytodifferentiation and Macromolecular Synthesis (Edited by LOCKE M.), pp. 65-84. Academic Press, New York. MARKERT , C. L. & URSPRUNG H. (1962) The ontogeny of isozyme patterns of lactate dehydrogenase in the mouse. Devel. Biol. 5, 363-381.
M ITCHELL W. M. (1967) A potential source of electrophoretic artifacts in polyacrylamide gels. Biochim. biophys. Acta 147, 171-174.
Acknon~ledgements-The authors wish to express their gratitude to Drs. A.-Young Woody and R. W. McGaughey for critically reading the manuscript. REFERENCES ADAMS E. Kc FINNEGAN C. V. (1965) An investigation of lactate dehydrogenase activity in early amphibian developmeni. J.Exp. Zoology. 158, 241-252. B ALEK R. W. & SNOW J. (1967) Ontogenetic changes of lactate dehydrogenase isozimes in two species of Ambystoma. Lqk Sci. 6, 2587-2595.
B URKE W. F. (1971) Lactate dehydrogenase of Hymenolysis diminuta: isolation and characterization. Master’s thesis, North Texas State University. C AHN R. D., KAPLAN N. O., LEVINE L. & ZWILLING E. (1962) Nature and development of lactic dehydrogenases. Science, Wash. 136, 962-969. C HEN P. S. (1968) Patterns of soluble proteins and multiple forms of dehydrogenase in amphibian development. J. Exp. Zool. 168, 337-350. CLAYCOMB W. C. & VILLEE C. A. (1971) Lactate dehydrogenase isozymes of Xenopus laevis; factors affecting their appearance during early development. Devel. Biol. 24, 413427.
C OSTELLO L. A. & KAPLAN N. 0. (1963) Evidence for two forms of M-type lactate dehydrogenase in the mouse. Biochim. bioohvs. Acta 73, 658-660. DAVIS B. J. (1964) Di& eiectrophoresis-II. Method and application to human serum proteins. Ann. N. Y. Acad. Sci. 121. 404-427.
M OYER F. H., SPEAKER , C. B. & WRIGHT D. A. (1968) Characteristics of lactate dehydrogenase isozymes in amohibians. Ann. N. Y. Acad. Sci. 151. 650-669. MURPHY J. B. & KIES M. W. (1960) N&e on spectrophotometric determination of proteins in dilute solutions. Biochim. biophys. Acta 45, 383-384. ROWE J. J. (1971) Electrophoretic forms and distribution of bacterial NADH-dependent isocitric dehydrogenase. Purification and partial characterization of NADPdependent isocitric dehydrogenase from Pseudomonas aeraginosa. Master’s thesis, Arizona State University. S HAW C. R. & KOEN A. L. (1965) On the identity 01 “nothing dehydrogenase”. J. Histochem. C’ytochem. 13. 431-433.
SZARSKI H. (1964) The structure of respiratory organs in relation to body size in amphibia. Evolution 18, I18126. WRIGHT D. A. & MOYER F. H. (1966) Parental influences on lactate dehydrogenase in the early development of hybrid frogs in the genus Rana. J. exp. Zoo/. 163, 215230.
WRIGHT D. A. & MOYER F. H. (1968) Inheritance of frog lactate dehydrogenase patterns and the persistence of maternal isozymes during development. J. exp. Zoo/. 167,97-206. Key Word Index-Isozyme; LDH ; embryos; Mexican axolotl; Ambystoma mexicanum.
ISOZYMIC PATTERNS OF LACTATE DEHYDROGENASE IN WHOLE EMBRYOS AND ADULT TISSUES OF THE MEXICAN AXOLOTL DELAINE GILCREASE* AND J. T. JUSTOSt:~ Department of Zoology, Arizona State University, Tempe, Arizona 85281, U.S.A. (Received 19 November 1973)
Abstract--1. The objective of this study was to determine the general ontogeny of lactate dehydrogenase isozymes in whole embryos and various adult tissues of the Mexican axolotl. 2. The LDH isozymes were resolved by polyacrylamide disc electrophoresis. 3. LDH from embryos showed predominantly anodal activity. 4. Skeletal muscle LDH patterns also showed chiefly anodal activity. This result differs substantially from previously reported studies. 5. We suggest that the patterns of LDH isozymes from skeletal muscle of this aquatic salamander may reflect a greater availability of oxygen to this tissue than in terrestrial animals of this type.
INTRODUCTION
DURING the past two decades, extensive research concerning lactate dehydrogenase (LDH) has provided a wealth of information concerning its enzymatic activity. Not only was L D H found to exist as an isozyme in tissues, but it also exhibited tissue specificity for the isozyme patterns (Markert, 1963). Five distinct L D H isozymes have been demonstrated electrophoretically in most tissues from birds and mammals (Calm et aL, 1962; Markert & Ursprung, 1962). These isozymic bands were arbitrarily numbered one to five, from anodal to cathodal ends of the gel. These uniform patterns found among mammals contrast conspicuously with the great diversity found among amphibia. Most examined amphibia have either more or fewer than the predicted five isozymes (Adams & Finnegan, 1965; Wright & Moyer, 1966; Balek & Snow, 1967; Chen, 1968; Moyer et al., 1968; Wright & Moyer, 1968; Claycomb & Villee, 1971). The predominant type of L D H found in tissues may be related to the availability of oxygen to that tissue. There may be a direct correlation between tissues subjected to anaerobic metabolism (e.g. skeletal muscle) and high concentrations of LDH-5 (Markert, 1963). On this basis, Markert (1963) suggested that the epigenetic control of isozymes * Recipient of a summer fellowship from the Arizona Heart Association. t Established investigator for the American Heart Association. This work was supported by NIH Grant HL 15054-02 to J. T. Justus. ~:Reprint requests should be sent to Dr. J. T. Justus.
involves the oxygen tension in the cell. This suggestion also extends to embryonic forms. Mammalian embryos have relatively poor supplies of oxygen and the embryonic tissues have large amounts of LDH-5 (Markert & Ursprung, 1962). On the other hand, Calm et al. (1962) observed a preponderance of LDH-1 isozymes in bird embryos in accordance with their relatively aerobic environment. The adult isozyme patterns are similar in birds and mammals, but this terminal condition is reached from opposite starting points, from LDH-5 in mammals and from LDH-1 in birds. This seems to demonstrate that as the availability of oxygen changes during development, there is a corresponding change in the L D H isozyme pattern. Furthermore, the results of Balek & Snow (1967) indicate the changes observed during amphibian development more closely resemble the changes found in chick embryos than those found in mammalian embryos. The objective of the present study was to determine the general ontogeny of L D H isozymes in whole embryos and various adult tissues of the Mexican axolotl, A m b y s t o m a mexicanum. Balek & Snow (1967) previously reported changes of L D H isozymes in whole embryos and adult tissues of this animal. Our studies of the changes in these isozymes during the early development of the axolotl are similar to those of Balek & Snow (1967); however, in adult tissues of this salamander, we found fundamental differences in the L D H patterns from those reported by the above-mentioned authors. MATERIALS AND METHODS Embryos were obtained from spawnings of Mexican axolotls. The embryos were maintained at 14°C to
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prolong each developmental stage. Assignment of developmental stages of the urodele embryos were determined by comparison with Harrison's tables for Arabystoma maeulatum (Hamburger, 1969). The following developmental stages were used in this study--unfertilized eggs, stage 33 (preheartbeat), stage 34 (initiation of heartbeat) and stage 41 (hatching). In addition, young larvae which had been feeding for 4 months and adult animals were used. Jelly coats from eggs and embryos at selected stages were removed using watchmaker's forceps. Eggs and whole embryos were sonicated in cellulose nitrate tubes for approximately 1 min with the small tip of a Heat Systems-Ultrasonic Model W140D Sonifier Cell Disrupter. The tubes were kept in an ice-bath during sonication to retard denaturation. One whole embryo or egg was sonicated in 100 p.1 of a sodium phosphate buffer (0.05 M, pH 7"5) containing 0.25 M sucrose. All juvenile and adult tissues were homogenized at a ratio of 1 : 20 (w/v) in the same buffer using a Potter-Elvehjem glass homogenizer equipped with a motor driven pestle. All homogenares were clarified for electrophoresis by centrifugation for 10 min at 30,000 g in a Sorvall RC2-B refrigerated centrifuge. The supernatants were used immediately for electrophoresis. Twenty p.1 of guinea-pig LDH standard (Sigma), a control for all electrophoretic separations, was diluted with 1"0 ml of sodium phosphate buffer (0'05 M, pH 7.5). The protein concentration of several crude extracts was determined by the procedure of Murphy & Kies (1960). Both Davis (1964) and Gabriel (1971) reported the total amount of protein in a crude extract applied to a polyacrylamide gel should not exceed 200/~g. The amount of protein applied to the gels in our experiments did not exceed this figure. The L D H isozymes of all homogenates were resolved by polyacrylamide disc electrophoresis according to the method of Davis (1964). Electrophoresis was carried out at 4°C at 4-5 m A per tube for approximately 1 hr, utilizing a Hoefer Model D E 102 Polyacrylamide Gel Unit connected to a Heathkit Model IP-I 7 power supply. The leads were connected such that the cathode was in the upper tray buffer and the anode in the lower tray buffer. Migration of the protein is from cathode to anode. Completion time was determined by a band of bromphenol blue reaching the bottom of the separating gel. To determine whether any enzyme migration was going into the upper tray buffer, the electrodes were reversed in some experiment s. A m m o n i u m persulfate has been reported as a possible source of artifacts because of its oxidation effect on protein sulfhydryl groups (Mitchell, 1967; Gabriel, 1971 ). Both of these authors suggest that pre-electrophoresis of
the separating gel for 2 h r at 4-5 mA per tube will essentially remove the ammonium persulfate. Comparison of resolution of the LDH isozymes on prewashed and non-prewashed gels showed no difference in electrophoretic separation except for mobility; subsequently, pre-electrophoresis was not performed. Following completion of electrophoresis, the gels were removed from the tubes and placed in small test-tubes in preparation for staining. The positive LDH stain, a slight modification of the method of Rowe (1971), was prepared in the dark immediately before staining. Control gels were run, but in the absence of the substrate, lactate, to detect any activity of "nothing dehydrogenase" (Shaw & Koen, 1965). Enzyme activity was present on control gels, incubated in stain lacking the substrate, corresponding to the bands of LDH activity localized with the positive stain. A negative stain was used to supplement these results and to rule out artifacts of the positive stain due to "nothing dehydrogenase". The negative stain was prepared according to Burke (unpublished) in the following way: Solution A--substrate solution: sodium pyruvate (Sigma), 100.0mg; N A D H (Sigma), 50'0rag: 0-1 M sodium phosphate (pH 7.5), 25.0 ml. Solution B--staining solution: nitro blue tetrazolium (NBT), 5-0mg; phenazine methosulfate (PMS), 0.3 ml: 0.1 M sodium phosphate buffer (pH 7-5), 25.0 ml. Solution B was protected from the light. The gels were first incubated with solution A at room temperature for 15 min, thoroughly rinsed with water and incubated in the dark for 15 rain with solution B. The LDH present in the gel will convert the components in solution A to lactic acid and N A D and at all other regions the pyruvate and N A D H will diffuse into the gel. When solution B is added, areas of LDH activity will show clear bands but the rest of the gel with NADH will be oxidized with the subsequent electron transfer through PMS to NBT which will precipitate as an insoluble purple formazan. At the end of all staining procedures, the gels were rinsed with tap water and stored in 7.0~ acetic acid in the dark. RESULTS T h e m a t e r n a l L D H b a n d s o f unfertilized eggs of A. mexicanum were c o m p a r e d with whole e m b r y o s f r o m stage 33 (preheartbeat), stage 34 (initiation of h e a r t b e a t ) a n d stage 41 ( h a t c h i n g ) (Fig. 1). This c o m p a r i s o n suggests t h a t the m a t e r n a l pattern persists at least until hatching. All three gels exhibit two darkly staining a n o d a l b a n d s a n d three lighter
Fig. 1. Electrophoresis pattern of L D H isozymes from unfertilized egg and developing embryos. EGG, Unfertilized egg; ST 33, stage 33 (preheartbeat); ST 34, stage 34 (initial heartbeat); HATC, stage 41 (hatching). Fig. 2. Electrophoretic patterns of LDH isozymes from skeletal muscle of juvenile and adult axolotls. SMj, Juvenile; SM A, adult. Fig. 3. Electrophoretic patterns of LDH isozymes from various axolotl tissues. H, Heart; SM, skeletal muscle; L, lung; SPL, spleen. Fig. 4. Electrophoretic patterns of LDH isozymes from various axolotl tissues. BR, Brain; Sk, skin; STOM, stomach; LIV, liver.
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ST 33
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LDH in embryonic and adult tissues of the axolotl bands toward the cathode. One of the lighter bands seems to be missing on gels of hatched embryos. Whether it is missing due to poor resolution or to a change in the pattern at this stage is not clear. The skeletal muscle patterns from juvenile and adult animals were compared (Fig. 2). Both juvenile and adult skeletal muscle showed a preponderance of anodal activity, atypical of the skeletal muscle pattern found in most vertebrate species. The adult isozyme patterns from the heart, skeletal muscle, lung, spleen, brain, skin, stomach and liver are shown (Figs. 3 and 4). All these tissues show a greater amount of anodal activity. Since Balek & Snow (1967) have reported that skeletal muscle of the axolotl seemed to show more cathodal activity than we observed, the electrodes of the gel unit were reversed to determine whether any protein migration into the upper tray buffer had occurred. Since the electrodes were reversed, any LDH originally migrating into the upper tray buffer should go into the gel. All of the above axolotl tissues were run with reversed leads and no activity was detected on the gel. Control gels stained without the substrate, lactate, showed activity corresponding to the bands of LDH activity seen on gels stained with positive stain. Even the electrophoretically pure guinea-pig heart LDH standard showed bands of activity when stained without lactate in the mixture. In an effort to determine if these bands of activity were LDH isozymes, a negative stain was used. A comparison of positive, negative and control gels without lactate are shown (Figs. 5-7). Other control gels stained with a mixture lacking both lactate and NAD showed no activity. Guinea-pig heart LDH standard was utilized as a control in all electrophoretic separations. Its isozyme pattern is typical of mammalian heart LDH pattern with predominantly anodal activity (Fig. 7). As a control for our methods of tissue disruption, homogenizing buffer and electrophoretic system, the same methods were employed in preparation and electrophoretic separation of tissues from a laboratory mouse. The LDH patterns from the mouse heart, skeletal muscle and kidney are shown (Fig. 8). Sub-banding was present in both heart and skeletal muscle, an observation commonly noted in the
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literature (Costello & Kaplan, 1963; Koen & Shaw, 1965; Koen, 1967; Dudman & Zemer, 1969). Despite the sub-banding, the characteristic isozymic pattern was evident, with heart and kidney showing anodal activity and skeletal muscle showing cathodal activity, patterns similar to those found by Markert & Ursprung (1962). DISCUSSION
The ontogeny of LDH isozymes of amphibians and fish closely resemble changes found in the chick embryo, with the initial activity at the anode followed by a gradual increase of activity toward the cathode as development proceeds (Goldberg et al., 1961; Balek &Snow, 1967; Chen, 1968; Moyer et al., 1968; Claycomb & Villee, 1971). Similarly, our results on axolotl embryos showed greater activity at the anode, and in this regard are consistent with previously published observations. Five bands of LDH activity were resolved in the adult axolotl heart with most of the activity at the anode. This is a typical pattern found in the hearts of vertebrates. Isozyme patterns from other axolotl tissues, including lung, spleen, brain, skin, stomach and liver, showed a preponderance of anodal activity. The axolotl heart isozyme pattern was typical of vertebrate hearts; the skeletal muscle LDH pattern observed was quite atypical. Both juvenile and adult skeletal muscle patterns showed predominantly anodal activity, whereas most other vertebrate skeletal muscle patterns show predominantly cathoda1 activity. Balek & Snow (1967) reported axolotl skeletal muscle to have predominantly cathodal activity. They used starch gel electrophoresis with a Tris-borate buffer, pH 8.6. Our results using polyacrylamide gel electrophoresis differ from those reported by these authors. Several experimental conditions in our study suggest that the anodal activity is peculiar to axolotl skeletal muscle. (1) Skeletal muscle patterns from both neotenous and adult Ambystoma tigrinum, a urodele similar to A. mexicanum, showed isozyme patterns with the concentration of activity at the anode (unpublished). (2) In our studies, characteristic tissue patterns for the laboratory mouse were
Fig. 5. Comparison of electrophoretic patterns from axolotl heart on positively stained, negatively stained and control gels lacking substrate. POS, Positively stained; NEG, negatively stained; NO SUB, control gel lacking substrate in stain. Fig. 6. Comparison of electrophoresis patterns from axolotl skeletal muscle on positively stained, negatively stained and control gels lacking substrate. POS, Positively stained; NEG, negatively stained; NO SUB, control gel lacking substrate in stain. Fig. 7. Comparison of electrophoretic patterns from guinea-pig heart LDH standard on positively stained, negatively stained and control gels lacking substrate. POS, positively stained; NEG, negatively stained; NO SUB, control gel lacking substrate in stain. Fig. 8. Electrophoretic pattern of LDH isozymes from mouse heart, skeletal muscle and kidney. H, Heart; SM, sketleal muscle; KID, kidney.
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obtained by using the same experimental conditions as for amphibian tissue. The mouse tissue patterns showed the typical isozyme bands, with heart and kidney showing predominantly anodal activity and skeletal muscle with concentrated cathodal activity. (3) LDH activity was not lost into the upper tray buffer. (4) Furthermore, loss of enzyme activity due to heat inactivation should not have been a problem in the present study because electrophoresis was performed at 4°C. The results presented in this paper provide convincing evidence that the electrophoretic separation of skeletal muscle isozymes with the predominant LDH activity concentrated at the anode is peculiar to the Mexican axolotl. Low oxygen tension favors synthesis of slower migrating isozymes, the form best suited for anaerobic metabolism and found in high concentrations in skeletal muscle from most vertebrates (Dawson et al., 1964). This information suggests that skeletal muscle from the axolotl may have a greater availability of oxygen than skeletal muscle from other vertebrates. This does not seem unreasonable in the light of the observation that about half of the respiratory gas exchange takes place across the skin in A. mexicanum (Szarski, 1964). Perhaps the large amount of anodal activity seen in axolotl skeletal muscle reflects a well-oxygenated tissue exhibiting aerobic metabolism. Experiments are being designed, using oxygen microelectrodes, to test this possibility.
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J. T. JUSTUS
D AWSON D., G~~DFRIEND T. L. & KAPLAN N. 0. (1964) Lactic dehydrogenases: functions of the two other types. Science, Wash. 28, 929-933. DUDMAN N. & ZERNER B. (1969) Simplifications of mouse lactate dehydrogenase electrophoretic patterns. Biochim. biophys. Acta 171, 195-197.
GABRIEL 0. (1971) Analytical disk gel electrophoresis. In Methods in Enzymology (Edited by COLOWICK S. P. & K APLAN N. 0. Vol. 22, pp. 565-578. Academic Press, New York. G OLDBERG E., CUERRIER J. P. & WARD J. C. (1969) Lactate dehydrogenase ontogeny, paternal gene activation, and tetramer assembly in embryos of brook trout, lake trout, and their hybrids. Biochem. Gen. 2, 335_~ 350.
H A M B U R G E R V. (1969) A Manual of Experimental Embrvoloav. University of Chicago Press. Chicago K OEN A. Lyi1967) Lactate dehydrogenase isozyme-&bbands: new bands derived from isolated sub-bands. Biochim. biophys. Acta 140, 496-502.
K OEN A. L. & SHAW C. R. (1965) Studies on lactate dehydrogenase tissue-specified isozymes: electrophoretie migration of isolated sub-bands. Biocbim. biophys. Acta 96, 231-236.
MARKERT C. L. (1963) Epigenetic control of specific protein synthesis in differentiating cells. In Cytodifferentiation and Macromolecular Synthesis (Edited by LOCKE M.), pp. 65-84. Academic Press, New York. MARKERT , C. L. & URSPRUNG H. (1962) The ontogeny of isozyme patterns of lactate dehydrogenase in the mouse. Devel. Biol. 5, 363-381.
M ITCHELL W. M. (1967) A potential source of electrophoretic artifacts in polyacrylamide gels. Biochim. biophys. Acta 147, 171-174.
Acknon~ledgements-The authors wish to express their gratitude to Drs. A.-Young Woody and R. W. McGaughey for critically reading the manuscript. REFERENCES ADAMS E. Kc FINNEGAN C. V. (1965) An investigation of lactate dehydrogenase activity in early amphibian developmeni. J.Exp. Zoology. 158, 241-252. B ALEK R. W. & SNOW J. (1967) Ontogenetic changes of lactate dehydrogenase isozimes in two species of Ambystoma. Lqk Sci. 6, 2587-2595.
B URKE W. F. (1971) Lactate dehydrogenase of Hymenolysis diminuta: isolation and characterization. Master’s thesis, North Texas State University. C AHN R. D., KAPLAN N. O., LEVINE L. & ZWILLING E. (1962) Nature and development of lactic dehydrogenases. Science, Wash. 136, 962-969. C HEN P. S. (1968) Patterns of soluble proteins and multiple forms of dehydrogenase in amphibian development. J. Exp. Zool. 168, 337-350. CLAYCOMB W. C. & VILLEE C. A. (1971) Lactate dehydrogenase isozymes of Xenopus laevis; factors affecting their appearance during early development. Devel. Biol. 24, 413427.
C OSTELLO L. A. & KAPLAN N. 0. (1963) Evidence for two forms of M-type lactate dehydrogenase in the mouse. Biochim. bioohvs. Acta 73, 658-660. DAVIS B. J. (1964) Di& eiectrophoresis-II. Method and application to human serum proteins. Ann. N. Y. Acad. Sci. 121. 404-427.
M OYER F. H., SPEAKER , C. B. & WRIGHT D. A. (1968) Characteristics of lactate dehydrogenase isozymes in amohibians. Ann. N. Y. Acad. Sci. 151. 650-669. MURPHY J. B. & KIES M. W. (1960) N&e on spectrophotometric determination of proteins in dilute solutions. Biochim. biophys. Acta 45, 383-384. ROWE J. J. (1971) Electrophoretic forms and distribution of bacterial NADH-dependent isocitric dehydrogenase. Purification and partial characterization of NADPdependent isocitric dehydrogenase from Pseudomonas aeraginosa. Master’s thesis, Arizona State University. S HAW C. R. & KOEN A. L. (1965) On the identity 01 “nothing dehydrogenase”. J. Histochem. C’ytochem. 13. 431-433.
SZARSKI H. (1964) The structure of respiratory organs in relation to body size in amphibia. Evolution 18, I18126. WRIGHT D. A. & MOYER F. H. (1966) Parental influences on lactate dehydrogenase in the early development of hybrid frogs in the genus Rana. J. exp. Zoo/. 163, 215230.
WRIGHT D. A. & MOYER F. H. (1968) Inheritance of frog lactate dehydrogenase patterns and the persistence of maternal isozymes during development. J. exp. Zoo/. 167,97-206. Key Word Index-Isozyme; LDH ; embryos; Mexican axolotl; Ambystoma mexicanum.
