- Email: [email protected]
The tricarboxylic acid cycle enzymes in the adult dog heartworm, Dirofilaria immitis
Comp. Biochem.Physiol., 1971, Vol. 38B,pp. 493 to 500. PergamonPress. Printedin Great Britain
T H E TRICARBOXYLIC ACID CYCLE ENZYMES IN T H E A D U L T DOG HEARTWORM, D I R O F I L A R I A I M M I T I S * K. M. M c N E I L L t
and W. F. H U T C H I S O N
Department of Preventive Medicine, University of Mississippi School of Medicine, Jackson, Mississippi 39216 (Received 24 August 1970)
A b s t r a c t - - 1 . The specific activities and intracellular locations of the tri-
carboxylic acid cycle enzymes; citrate synthase, aconitase, isocitrate dehydrogenase, ct-ketoglutarate dehydrogenase, succinate dehydrogenase, fumarase, malate dehydrogenase and glutamate dehydrogenase were determined in the adult dog heartworm, Dirofilaria immitis. Coenzyme specificities were determined when applicable. 2. Although all of the enzymes of this cycle were demonstrated, low levels of aconitase and isocitrate dehydrogenase plus the apparent absence of malate dehydrogenase activity in the forward direction suggest that the complete cycle may be of questionable importance in D. immitis.
INTRODUCTION T~E EXISTENCE of a complete tricarboxylic acid cycle has been demonstrated in Ascaris lumbricoides (Oya et al., 1965), Echinococcus granulosus scolices (Agosin & Repetto, 1963), TrichineUa spiralis larvae (Goldberg, 1957) and Fasciola hepatica (Prichard & Schofield, 1968). In addition, all enzymes of this cycle except fumarase, which was not investigated, are known to be present in the unembryonated eggs of A . lumbricoides (Costello & Brown, 1962; Oya et al., 1963; Smith et al., 1963). A systematic study of each of the enzymes of the tricarboxylic acid cycle of the adult dog heartworm, Dirofilaria immitis, was made to determine if a complete cycle exists in this nematode parasite. T h e activities of these enzymes of D. immitis determined in the present work are compared to those as reported in F. hepatica by Prichard & Schofield (1968). T h e same assay procedures used in this previous study with F. hepatica were followed as closely as possible in our work with D. immitis allowing for a direct comparison of the characteristics of the tricarboxylic acid cycle of a filarial worm with that of a fluke. * This investigation was supported in part by National Institutes of Health General Research Support Grant 5-SO1-RRO5386 to the University of Mississippi School of Medicine. t National Institutes of Health Predoctoral Research Fellow, Fellowship No. 5-FO1GM-42, 727-02. 493
494
K . M . McNEILL AND W. F. HUTCHISON MATERIALS AND M E T HODS
Animals Immediately following the sacrifice of dogs, adult D. imrnitis were removed from the heart and pulmonary vessels and placed in physiological saline for transport to the laboratory. After repeated washings in physiological saline to free them of blood clots, the worms were immediately prepared for enzyme assay or kept alive overnight in medium 199 containing streptomycin and penicillin for assay the following day.
Homogenization and centrifugation T o prepare the soluble (extra mitochondrial) fraction, freshly washed worms were blotted dry on filter paper, minced with scissors and placed in a Potter-Elvehjem homogenizer along with approximately 10 vol. of ice-cold 0"15 M KC1 containing 8 ml of 0"2 M KHzCOs/1. to maintain the pH at 7"0. The worms to be used for the preparation of the mitochondrial fraction were handled in the same manner except they were homogenized in 0"25 M sucrose. The temperature of the extracts was maintained at 2-4°C throughout homogenization by keeping the tissue grinder immersed in an ice-bath. The soluble fraction was prepared by centrifugation of the homogenate at 12,000 g for 15 min. in a Servall RC-2 refrigerated centrifuge at 2-4°C. The supernatant was retained in an ice-bath for immediate enzyme assay. The mitochondrial suspension was prepared according to the method described by Prichard & Schofield (1968) and stored in an ice-bath for immediate enzyme assay.
Reagents All substrates and cofactors except potassium ferricyanide were obtained from Sigma Chemical Company. These include D,L-isocitrate, oxaloacetate, citrate, a-ketoglutarate, succinate, fumarate, malate, acetyl-CoA, ADP, NAD, NADP, N A D H , , N A D P H , and D T N B . * The potassium ferricyanide was obtained from Scientific Products. Substrates and cofactors were buffered before being added to the reaction mixture. Oxaloacetate and cysteine were brought to neutral p H immediately before use.
Enzyme assay procedures All enzymes were assayed spectrophotometrically at 24-26°C with a Hitachi Perkin Elmer Model 139 in silica cuvettes of 1 cm light path, temperature being regulated by a circulating water bath. Zero-order kinetics were observed in all assays. Citrate synthase (E.C. 4.1.3.7). The activity of this enzyme was measured by the method of Shepherd & Garland (1969). The assay contained: Tris-HC1 buffer, pH 8"0, 100 m M ; D T N B , 66"6/xM; oxaloacetate, 166.5/zM; acetyl-CoA, 33"3/zM; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of the enzyme preparation. The D T N B and acetyl-CoA must be added previously and in that order. The activity was determined from the increase in extinction at 412 m/z. Aconitase (E.C. 4.2.1.3). The assay, based on Anfinsen (1955), contained: phosphate buffer p H 7"6, 50 m M ; citrate, 33 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of citrate. The activity was determined from the increase in extinction at 240 m/z. Isocitrate dehydrogenase (E.C. 1.1.1.41). The assay was based on Goebell & Klingenberg (1964). The assay contained: phosphate buffer pH 7"6, 100 m M ; MgC12, 8 m M ; ADP, 2 m M ; NAD, 2 m M ; D,L-isocitrate, 1"6 m M ; enzyme preparation; water to 3.0 ml. The reaction was started by the addition of D,L-isocitrate. The activity was determined from the increase in extinction at 340 m/z. Isocitrate dehydrogenase (E.C. 1.1.1.42). The assay was based on Ochoa (1955b). The assay contained: glycylglycine pH 7"6, 25 m M ; MnC12, 0"8 m M ; NADP, 45/zM; D,Lisocitrate, 0"2 raM; enzyme preparation; water to 3'0 ml. The reaction was started by the *
5,5"-Dithiobis-(2-nitrobenzoic acid).
T R I C A R B O X Y L I C ACID CYCLE ENZYMES I N D I R O F I L A R I A I M M I T I S
495
addition of D,L-isocitrate. The activity was determined from the increase in extinction at 340 m/~. o~-Ketoglutarate dehydrogenase (E.C. 1.2.4.2). The assay was based on Massey (1960). The assay contained: phosphate buffer, pH 6"4, 50 m M ; KCN, 5 raM; KaFe(CN)s, 667 /zM; ct-ketoglutarate, 3"3 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of the enzyme preparation. The activity was determined from the decrease in extinction at 410 m/L. Succinate dekydrogenase (E.C. 1.3.99.1). Two assay procedures were used to assay this enzyme. The first assay, measuring the oxidation of succinate to fumarate in the presence of ferricyanide, was that of Slater & Bonner (1952). The assay contained: phosphate buffer, p H 7"24, 140 m M ; K3Fe(CN)6, 1 m M ; KCN, 10 raM; succinate, 26 raM; enzyme preparation ; water to 3"0 ml. The reaction was started by the addition of the enzyme preparation. The activity was determined from the decrease in extinction at 400 m/~. The other method, measuring the reduction of fumarate to succinate, was that of Prichard & Schofield (1968). The assay contained: phosphate buffer, pH 7"6, 42 m M ; CaCI~, 33/~M; MgCla, 1.7 m M ; NADH~, 57/~M or NADPH~, 42/LM; fumarate, 33 m M ; mitochondrial suspension; water to 3'0 ml. The reaction was started by the addition of fumarate. The activity was determined from the decrease in extinction at 340 m/~. Fumarase (E.C. 4.2.1.2.). The assay was based on Racker (1950). The assay contained: phosphate buffer, pH 7-6, 50 m M ; malate, 1"7 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of malate. The activity was determined from the increase in extinction at 240 m/L. Malate dehydrogenase (E.C. 1.1.1.37). The assay to determine the rate of oxidation of malate by this enzyme was based on Ochoa (1955a). The assay contained: Tris, pH 8"0, 50 m M ; NAD, 0"2mM; malate, 6-7 raM; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of malate. The activity was determined from the increase in extinction at 340 m/~. The reduction of oxaloacetate was assayed by the technique of Shonk & B.xer (1964). The assay contained : triethanolamine, p H 7"6, 54 m M ; ethylenediaminetetraacetate, 5 m M ; NADH~, 0"17 m M ; oxaloacetate, 0"33 raM; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of oxaloacetate. The activity was determined from the decrease in extinction at 340 m/~. Glutamate dehydrogenase (E.C. 1.4.1.2/1.4.1.3). The assay was based on Schmidt (1963). The assay contained: triethanolamine, pH 8"0, 36 m M ; ethylenediaminetetraacetate 2"5 raM; ammonium acetate, 100 m M ; NADH~, 167/~M or NADPHa, 84/~M; ct-ketoglutarate, 8"3 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of ct-ketoglutarate. The activity was determined from the decrease in extinction at 340 m/z. Protein determinations. Protein determinations were according to the biuret method of Gornall et al. (1949). Specific activities are expressed in terms of m/~mole product formed/min per mg protein. Assays for each enzyme were performed on two groups of worms and results are expressed as means. Activity was proportional to the amount of enzyme preparation used. RESULTS All e n z y m e s of the tricarboxylic acid cycle are p r e s e n t i n D. immitis. M a r k e d differences i n levels of activity, c o e n z y m e speciflcities a n d i n t r a c e l l u l a r location are n o t e d b e t w e e n t h e e n z y m e s of this parasite a n d those of F. hepatica as s h o w n i n T a b l e 1. Citrate s y n t h a s e occurs i n b o t h the s o l u b l e a n d m i t o c h o n d r i a l fractions of D. immitis as i n F. hepatica a l t h o u g h the activity is n o t e v e n l y d i s t r i b u t e d b e t w e e n these two fractions as i n the fluke.
D. immitis F. hepatica
0
95"0 0 208"0 0 0
--
25 "2 0 0 0 76"8 176"8
65"2 14"4 3"5 1"8 0 23" 1 70"6 0 1375"5 1 "8 0
Mitochondrial fraction
Soluble fraction
D. immitis
15 "4 4"8 0 17"2 0 23 1"5 61 111 2180 426 22
Soluble fraction
15"3 0 0 3"2 13"2 283 12 0 238 2010 1685 222
Mitochondrial fraction
F. hepatica*
The following abbreviations are used: suc., succinate; fum., fumarate; mal., malate; oaa., oxaloacetate. * Prichard & Schofield (1968).
Citrate synthase Aconitase Isocitrate dehydrogenase (NAD-specific) lsocitrate dehydrogenase (NADP-specific) ~-Ketoglutarate dehydrogenase Succinate (suc.-+fum.) dehydrogenase (fum.~suc.) Fumarase Malate (mal.-->oaa.) dehydrogenase (oaa.-+mal.) Glutamate ( N A D H 2-specific) dehydrogenase (NADPHz)
C O M P A R E D T O A C T I V I T Y LEVELS OF C O R R E S P O N D I N G
Enzyme activity (m/zmole product formed/min per mg protein)
ENZYMES IN
O F E N Z Y M E S OF T H E T R I C A R B O X Y L I C A C I D C Y C L E I N
Enzyme
TABLE 1--ACTIVITY
z
O
t" t~
Z
¢3
~o G~
TRICARBOXYLIC
ACID
CYCLE
ENZYMES
IN DIROF1LARIA IMMITIS
497
A higher level of aconitase is noted in D. immitis than in F. hepatica. In both organisms this enzyme is confined to the soluble fraction. In contrast to the liver fluke, both NAD- and NADP-speeific isocitrate dehydrogenase activity in D. immitis is confined to the soluble fraction. The enzyme specific for NADP is found in both fractions in F. hepatica but none specific for NAD occurs. A higher level of ~-ketoglutarate dehydrogenase activity is found in D. immitis than in F. hepatica. This enzyme is confined to the mitochondrial fraction in both organisms. i Succinate dehydrogenase activity catalyzing the forward reaction is high in D. immitis, comparing very closely to that found in F. hepatica. The mitochondrial fraction was assayed for activity of this enzyme in the reverse reaction. Although no measurable levels were detected in D. immitis, both fractions possess activity in this direction in F. hepatica. D. immitis shows a high level of fumarase activity in the mitoehondrial fraction while the liver fluke has none. Approximately equal levels are present in the soluble fractions of both organisms. D. immitis apparently lacks malate dehydrogenase activity in the forward direction although a very active reverse reaction occurs. Although F. hepatica does possess activity of this enzyme in both directions, the reverse reaction is likewise much more active. A very low level of only NADH2-speeific glutamate dehydrogenase occurs in D. immitis and is confined to the soluble fraction. In contrast F. hepatica possesses glutamate dehydrogenase reacting with both NADH 2 and NADPH 2 in significant amounts in both the soluble and mitochondrial fractions. DISCUSSION Although no enzymatic study of this type has been performed previously with D. immitis, electron micrographs of the intracellular structure of this worm indicate that only a few rudimentary mitochondria are present in this organism (Lee & Miller, 1967, 1969). Electron micrographs of unsuspended mitochondrial fractions used in the present study substantiate these previous findings. Although the importance of the tricarboxylic acid cycle would seem questionable in view of this information, the presence of all the enzymes of this cycle has been demonstrated in the present work. The presence of citrate synthase indicates that D. immitis is able to synthesize citrate, the first intermediate of the tricarboxylic acid cycle. Ball (1965) indicates that some of the citrate may be diverted into other systems such as the citratemalate cycle for lipogenesis, but the citrate cleavage enzyme is yet to be demonstrated in D. immitis. Studies of aconitase activity in normal and anaerobically grown yeast have shown that oxygen deficiency greatly reduces the activity of this enzyme (Hirsch, 1952). Likewise, isocitrate dehydrogenase activity is high in tissues which possess a predominant aerobic type of metabolism (Ochoa & Weisz-Tabori, 1948). The low
498
K.M. McNEILL ANDW. F. HUTCHISON
levels of these enzymes in D. immitis suggest that the tricarboxylic acid cycle may not be of major importance in this parasite. This correlates closely with the situation found in F. hepatica (Prichard & Schofield, 1968). Work with T. spiralis (Goldberg, 1957) and A. lumbricoides (Oya et al., 1963, 1965; Smith et al., 1963) indicates that these parasites possess only NADP-specific isocitrate dehydrogenase. Prichard & Schofield (1958) report similar findings in F. hepatica, However, the present work has demonstrated isocitrate dehydrogenase specific for both NAD and NADP in D. immitis although the activity with each coenzyme is very low. Agosin & Repetto (1963) also found isocitrate dehydrogenase specific for both NAD and NADP in E. granulosus scolices. Isocitrate dehydrogenase specific for NAD occurs almost solely in the mitochondria of rat liver, while the NADP-specific enzyme is more concentrated in the extra mitochondrial fraction (Ernster & Navazio, 1956; Prichard & Schofield, 1968). In contrast, isocitrate dehydrogenase specific for each coenzyme is present in measurable amounts only in the soluble fraction in D. immitis. D. immitis possesses a very active ~-ketoglutarate dehydrogenase. The possible importance of this enzyme is enhanced by low glutamate dehydrogenase activity, the enzyme which competes with the former for ~-ketoglutarate. It appears that the oxidative decarboxylation of ~-ketoglutarate to succinate by ~-ketoglutarate dehydrogenase predominates as the major reaction at this step in the tricarboxylic acid cycle in D. immitis. A very active glutamate dehydrogenase in F. hepatica tends to indicate a relative unimportance of ~-ketoglutarate dehydrogenase in the fluke (Prichard & Schofield, 1968). Only NAD-specific glutamate dehydrogenase is present in D. immitis. Dixon & Webb (1964) state that only liver glutamate dehydrogenase shows no coenzyme specificity while this enzyme in other animal tissues is specific for NAD. Glutamate dehydrogenase activity was found to proceed in the presence of NAD or NADP in the liver fluke (Prichard & Schofield, 1968). These authors postulate that a NAD-specific as well as a non-specific glutamate dehydrogenase exists in F. hepatica since a NADP-specific enzyme has not yet been reported in animal tissues. D. immitis has a high succinate dehydrogenase activity in the forward direction but the reverse reaction could not be detected in this study. Prichard & Schofield (1968) reported both the forward and reverse reactions in F. hepatica, but they found relatively low activity in the reverse reaction. Although this indicates an aerobictype of metabolism, neither they nor van Grembergen (1949) were able to show marked inhibition of succinate oxidation in F. hepatica by addition of malonate, an effect readily demonstrated in rat liver, a predominately aerobic tissue. Further studies may reveal a similar situation in D. immitis. Although Prichard & Schofield (1968) reported that fumarase activity is confined to the soluble fraction in F. hepatica, high activity of fumarase is present in both the soluble and mitochondrial fractions of D. immitis. The particulate fraction of A. lumbricoides also contains fumarase activity (Oya et al., 1965). Every attempt to demonstrate the forward reaction of malate dehydrogenase in D. immitis was unsuccessful although a highly active reverse reaction is present. If
T R I C A R B O X Y L I C ACID CYCLE ENZYMES I N D I R O F I L A R I A I M M 1 T I S
499
malate dehydrogenase activity in the forward direction is absent in this parasite, malate may possibly be converted to oxaloacetate by some alternate pathway. One such possible pathway could be the conversion of malate to pyruvate by the malic enzyme with pyruvate being carboxylated to form oxaloacetate by the action of pyruvate carboxylase. Obviously the synthesis of oxaloacetate by some mechanism is essential to allow for the formation of citrate in the initial step of the tricarboxylic acid cycle. D. immitis possesses a high activity of enolase which acts to form phosphoenolpyruvate but only a low level of pyruvate kinase activity (Hutchison & McNeill, 1970). Phosphoenolpyruvic acid carboxykinase may compete with pyruvate kinase for phosphoenolpyruvate providing another possible source of oxaloacetate. Future studies are planned to determine the actual fate of malate at this step in the tricarboxylic acid cycle and to determine the major source of oxaloacetate for citrate synthesis in this parasite. Prichard and Schofield (1968) found malate dehydrogenase activity in both directions in F. hepatica although the reverse reaction was far more active. They suggest that malate may be metabolized to succinate by a partial reversal of the tricarboxylic acid cycle which may then be converted to the propionic acid excreted by this fluke. The highly active reverse reaction of malate dehydrogenase in D. immitis indicates that a similar situation may occur in this parasite. Further exploration in this area is required before definite conclusions can be formed. Although D. immitis contains all of the enzymes of the tricarboxylic acid cycle, the low activity levels of aconitase and isocitrate dehydrogenase plus the apparent absence of malate dehydrogenase activity in the forward direction suggest that the complete cycle may be of relatively minor importance in the overall metabolism of this parasite. Similar conclusions have been reported concerning the importance of the complete cycle in A . lumbricoides (Oya et al., 1965) and in F. hepatica (Prichard & Schofield, 1968). Acknowledgements--The authors are indebted to Professors J. Neal Brown and Joseph L. Haining for the assistance and encouragement they have shown during the course of this work.
REFERENCES AOOSINM. & REPErrO Y. (1963) Studies on the metabolism of Echinococcus granulosus--VI I. Reactions of the tricarboxylic acid cycle in E. granulosus scolices. Comp. Biochem. Physiol. 8, 245-261. ANFINSENC. B. (1955) Aconitase from pig heart muscle. In Methods in Enzymology (Edited by COLOWmKS. P. & KaPL~ N. O.), Vol. 1, pp. 695-698. AcademicPress, New York. BALL E. G. (1965) Regulation of fatty acid synthesis in adipose tissue. In Advances in Enzyme Regulation (Edited by WEBERG.), Vol. 4, p. 11. Pergamon Press, Oxford. COSTELLOL. C. & BROWNH. (1962) Aerobic metabolism of unembryonated eggs of Ascaris lumbricoides. Expl Parasitol. 12, 33-40. DIXON M. & WEBBE. C. (1964) Enzymes, pp. 686--689. Longmans, Green, London. ERNSTERL. & NAVAZlOF. (1956) The cytoplasmic distribution of isocitric dehydrogenase. Expl Cell Res. 11, 483-486. GOEBEr.L H. & KLXNCENBERCM. (1964) DPN-spezifische isoCitratedehydrogenase der Mitochondrien--I. Kinetische Eigenschaften, Vorkommen und Funktion der DPNspezifischen isoCitratedehydrogenase. Biochem. Z. 340, 441-464.
500
K. M. McNEILL AND W. F. HUTCHISON
GOLDBERG E. (1957) Studies on the intermediary metabolism of Trichinella spiralis. Expl Parasitol. 6, 367-382. GORNALL A. G., BARDAWlLLC. J. & DAVID M. M. (1949) Determination of serum proteins by means of the biuret reaction. J. biol. Chem. 177, 751-766. HIRSCH H. M. (1952) A comparative study of aconitase, fumarase and DPN-linked isocitric dehydrogenase in normal and respiratory-deficient yeast. Biochim. biophys. Acta 9, 674-686. HUTCHISON W. F. & McNEILL K. M. (1970) Glycolysis in the adult dog heartworm, Dirofilaria immitis. Comp. Biochem. Physiol. 35, 721-727. LEE C. C. & MILLER J. H. (1967) Fine structure of the body-wall musculature of Dirofilaria immitis. Expl Parasitol. 20, 334-344. LEE C. C. & MILLER J. H. (1969) Fine structure of the intestinal epithelium of Dirofilaria immitis and changes occurring after vermicidal treatment with Caparsolate Sodium. J. Parasitol. 55, 1035-1045. MASSEY V. (1960) T h e composition of the ketoglutarate dehydrogenase complex. Bioehim. biophys. Acta 38, 447-460. OCHOA S. (1955a) Crystalline condensing enzyme from pig heart. In Methods in Enzymology (Edited by COLOWICK S. P. & KAPLAN N. O.), Vol. 1, pp. 685-694. Academic Press, New York. OcnoA S. (1955b) Isocitric dehydrogenase system (TPN) from pig heart. In Methods in Enzymology (Edited by COLOWICK S. P. & KAPLAN N. O.), Vol. 1, pp. 699-704. Academic Press, New York. OCHOA S. & WEIsz-TABoRI E. (1948) Biosynthesis of tricarboxylic acids by carbon dioxide fixation--II. Oxalosuccinic carboxylase. J. biol. Chem. 174, 123-132. OVA H., COSTELLOL. C. & SMITH W. (1963) Succinate and malate oxidation and cytochrome c reduction in embryonated and unembryonated Ascaris eggs. J. Parasitol. 49 (5, Sect. 2) 51. OYA H., KIKUCHI G., BANDO T. & HAYAS8I H. (1965) Muscle tricarboxylic acid cycle in Ascaris lumbricoides vat. suis. Expl Parasitol. 17, 229-240. PRICHARD R. K. & SC8OFIELDP. J. (1968) A comparative study of the tricarboxylic acid cycle enzymes in Fasciola hepatica and rat liver. Comp. Biochem. Physiol. 25, 1005-1019. RACKER E. (1950) Spectrophotometric measurements of the enzymatic formation of fumaric and cis-aconitic acids. Biochim. biophys. Acta 4, 211-214. SCHMIDT E. (1963) Glutamate dehydrogenase. In Methods of Enzymatic Analysis (Edited by BERGMEYER H. U.), pp. 752-756. Verlag Chemic, Academic Press, New York. SHEPHERD D. & GARLAND P. B. (1969) Citrate synthase from rat liver. In Methods in Enzymology (Edited by LGWENSTEIN J. M.), Vol. 13, pp. 11-16. Academic Press, New York. SHONK C. E. & BOXER G. E. (1964) Enzyme patterns in human tissues--I. Methods for the determination of glycolytic enzymes. Cancer Res. 24, 709-721. SLATER E. C. & BONNER W. D. (1952) The effect of fluoride on the succinic oxidase system. Biochem.ff. 52, 185-196. SMITH W., COSTELLO L. C. & OYA H. (1963) Dehydrogenase activity in unembryonated and embryonated Ascaris eggs. ft. Parasitol. 49, (5, Sect. 2), 51. VAN GREMBERGEN G. (1949) Le metabolisme respiratoire du trematode Fasciola hepatica Linn. Enzymologia 13, 241-257.
Key Word Index--Dirofilaria immitis; tricarboxylic acid cycle; citrate synthase; aconitase; isocitrate dehydrogenase; ~-ketoglutarate dehydrogenase; succinate dehydrogenase; fumarase; malate dehydrogenase; glutamate dehydrogenase.
T H E TRICARBOXYLIC ACID CYCLE ENZYMES IN T H E A D U L T DOG HEARTWORM, D I R O F I L A R I A I M M I T I S * K. M. M c N E I L L t
and W. F. H U T C H I S O N
Department of Preventive Medicine, University of Mississippi School of Medicine, Jackson, Mississippi 39216 (Received 24 August 1970)
A b s t r a c t - - 1 . The specific activities and intracellular locations of the tri-
carboxylic acid cycle enzymes; citrate synthase, aconitase, isocitrate dehydrogenase, ct-ketoglutarate dehydrogenase, succinate dehydrogenase, fumarase, malate dehydrogenase and glutamate dehydrogenase were determined in the adult dog heartworm, Dirofilaria immitis. Coenzyme specificities were determined when applicable. 2. Although all of the enzymes of this cycle were demonstrated, low levels of aconitase and isocitrate dehydrogenase plus the apparent absence of malate dehydrogenase activity in the forward direction suggest that the complete cycle may be of questionable importance in D. immitis.
INTRODUCTION T~E EXISTENCE of a complete tricarboxylic acid cycle has been demonstrated in Ascaris lumbricoides (Oya et al., 1965), Echinococcus granulosus scolices (Agosin & Repetto, 1963), TrichineUa spiralis larvae (Goldberg, 1957) and Fasciola hepatica (Prichard & Schofield, 1968). In addition, all enzymes of this cycle except fumarase, which was not investigated, are known to be present in the unembryonated eggs of A . lumbricoides (Costello & Brown, 1962; Oya et al., 1963; Smith et al., 1963). A systematic study of each of the enzymes of the tricarboxylic acid cycle of the adult dog heartworm, Dirofilaria immitis, was made to determine if a complete cycle exists in this nematode parasite. T h e activities of these enzymes of D. immitis determined in the present work are compared to those as reported in F. hepatica by Prichard & Schofield (1968). T h e same assay procedures used in this previous study with F. hepatica were followed as closely as possible in our work with D. immitis allowing for a direct comparison of the characteristics of the tricarboxylic acid cycle of a filarial worm with that of a fluke. * This investigation was supported in part by National Institutes of Health General Research Support Grant 5-SO1-RRO5386 to the University of Mississippi School of Medicine. t National Institutes of Health Predoctoral Research Fellow, Fellowship No. 5-FO1GM-42, 727-02. 493
494
K . M . McNEILL AND W. F. HUTCHISON MATERIALS AND M E T HODS
Animals Immediately following the sacrifice of dogs, adult D. imrnitis were removed from the heart and pulmonary vessels and placed in physiological saline for transport to the laboratory. After repeated washings in physiological saline to free them of blood clots, the worms were immediately prepared for enzyme assay or kept alive overnight in medium 199 containing streptomycin and penicillin for assay the following day.
Homogenization and centrifugation T o prepare the soluble (extra mitochondrial) fraction, freshly washed worms were blotted dry on filter paper, minced with scissors and placed in a Potter-Elvehjem homogenizer along with approximately 10 vol. of ice-cold 0"15 M KC1 containing 8 ml of 0"2 M KHzCOs/1. to maintain the pH at 7"0. The worms to be used for the preparation of the mitochondrial fraction were handled in the same manner except they were homogenized in 0"25 M sucrose. The temperature of the extracts was maintained at 2-4°C throughout homogenization by keeping the tissue grinder immersed in an ice-bath. The soluble fraction was prepared by centrifugation of the homogenate at 12,000 g for 15 min. in a Servall RC-2 refrigerated centrifuge at 2-4°C. The supernatant was retained in an ice-bath for immediate enzyme assay. The mitochondrial suspension was prepared according to the method described by Prichard & Schofield (1968) and stored in an ice-bath for immediate enzyme assay.
Reagents All substrates and cofactors except potassium ferricyanide were obtained from Sigma Chemical Company. These include D,L-isocitrate, oxaloacetate, citrate, a-ketoglutarate, succinate, fumarate, malate, acetyl-CoA, ADP, NAD, NADP, N A D H , , N A D P H , and D T N B . * The potassium ferricyanide was obtained from Scientific Products. Substrates and cofactors were buffered before being added to the reaction mixture. Oxaloacetate and cysteine were brought to neutral p H immediately before use.
Enzyme assay procedures All enzymes were assayed spectrophotometrically at 24-26°C with a Hitachi Perkin Elmer Model 139 in silica cuvettes of 1 cm light path, temperature being regulated by a circulating water bath. Zero-order kinetics were observed in all assays. Citrate synthase (E.C. 4.1.3.7). The activity of this enzyme was measured by the method of Shepherd & Garland (1969). The assay contained: Tris-HC1 buffer, pH 8"0, 100 m M ; D T N B , 66"6/xM; oxaloacetate, 166.5/zM; acetyl-CoA, 33"3/zM; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of the enzyme preparation. The D T N B and acetyl-CoA must be added previously and in that order. The activity was determined from the increase in extinction at 412 m/z. Aconitase (E.C. 4.2.1.3). The assay, based on Anfinsen (1955), contained: phosphate buffer p H 7"6, 50 m M ; citrate, 33 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of citrate. The activity was determined from the increase in extinction at 240 m/z. Isocitrate dehydrogenase (E.C. 1.1.1.41). The assay was based on Goebell & Klingenberg (1964). The assay contained: phosphate buffer pH 7"6, 100 m M ; MgC12, 8 m M ; ADP, 2 m M ; NAD, 2 m M ; D,L-isocitrate, 1"6 m M ; enzyme preparation; water to 3.0 ml. The reaction was started by the addition of D,L-isocitrate. The activity was determined from the increase in extinction at 340 m/z. Isocitrate dehydrogenase (E.C. 1.1.1.42). The assay was based on Ochoa (1955b). The assay contained: glycylglycine pH 7"6, 25 m M ; MnC12, 0"8 m M ; NADP, 45/zM; D,Lisocitrate, 0"2 raM; enzyme preparation; water to 3'0 ml. The reaction was started by the *
5,5"-Dithiobis-(2-nitrobenzoic acid).
T R I C A R B O X Y L I C ACID CYCLE ENZYMES I N D I R O F I L A R I A I M M I T I S
495
addition of D,L-isocitrate. The activity was determined from the increase in extinction at 340 m/~. o~-Ketoglutarate dehydrogenase (E.C. 1.2.4.2). The assay was based on Massey (1960). The assay contained: phosphate buffer, pH 6"4, 50 m M ; KCN, 5 raM; KaFe(CN)s, 667 /zM; ct-ketoglutarate, 3"3 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of the enzyme preparation. The activity was determined from the decrease in extinction at 410 m/L. Succinate dekydrogenase (E.C. 1.3.99.1). Two assay procedures were used to assay this enzyme. The first assay, measuring the oxidation of succinate to fumarate in the presence of ferricyanide, was that of Slater & Bonner (1952). The assay contained: phosphate buffer, p H 7"24, 140 m M ; K3Fe(CN)6, 1 m M ; KCN, 10 raM; succinate, 26 raM; enzyme preparation ; water to 3"0 ml. The reaction was started by the addition of the enzyme preparation. The activity was determined from the decrease in extinction at 400 m/~. The other method, measuring the reduction of fumarate to succinate, was that of Prichard & Schofield (1968). The assay contained: phosphate buffer, pH 7"6, 42 m M ; CaCI~, 33/~M; MgCla, 1.7 m M ; NADH~, 57/~M or NADPH~, 42/LM; fumarate, 33 m M ; mitochondrial suspension; water to 3'0 ml. The reaction was started by the addition of fumarate. The activity was determined from the decrease in extinction at 340 m/~. Fumarase (E.C. 4.2.1.2.). The assay was based on Racker (1950). The assay contained: phosphate buffer, pH 7-6, 50 m M ; malate, 1"7 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of malate. The activity was determined from the increase in extinction at 240 m/L. Malate dehydrogenase (E.C. 1.1.1.37). The assay to determine the rate of oxidation of malate by this enzyme was based on Ochoa (1955a). The assay contained: Tris, pH 8"0, 50 m M ; NAD, 0"2mM; malate, 6-7 raM; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of malate. The activity was determined from the increase in extinction at 340 m/~. The reduction of oxaloacetate was assayed by the technique of Shonk & B.xer (1964). The assay contained : triethanolamine, p H 7"6, 54 m M ; ethylenediaminetetraacetate, 5 m M ; NADH~, 0"17 m M ; oxaloacetate, 0"33 raM; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of oxaloacetate. The activity was determined from the decrease in extinction at 340 m/~. Glutamate dehydrogenase (E.C. 1.4.1.2/1.4.1.3). The assay was based on Schmidt (1963). The assay contained: triethanolamine, pH 8"0, 36 m M ; ethylenediaminetetraacetate 2"5 raM; ammonium acetate, 100 m M ; NADH~, 167/~M or NADPHa, 84/~M; ct-ketoglutarate, 8"3 m M ; enzyme preparation; water to 3"0 ml. The reaction was started by the addition of ct-ketoglutarate. The activity was determined from the decrease in extinction at 340 m/z. Protein determinations. Protein determinations were according to the biuret method of Gornall et al. (1949). Specific activities are expressed in terms of m/~mole product formed/min per mg protein. Assays for each enzyme were performed on two groups of worms and results are expressed as means. Activity was proportional to the amount of enzyme preparation used. RESULTS All e n z y m e s of the tricarboxylic acid cycle are p r e s e n t i n D. immitis. M a r k e d differences i n levels of activity, c o e n z y m e speciflcities a n d i n t r a c e l l u l a r location are n o t e d b e t w e e n t h e e n z y m e s of this parasite a n d those of F. hepatica as s h o w n i n T a b l e 1. Citrate s y n t h a s e occurs i n b o t h the s o l u b l e a n d m i t o c h o n d r i a l fractions of D. immitis as i n F. hepatica a l t h o u g h the activity is n o t e v e n l y d i s t r i b u t e d b e t w e e n these two fractions as i n the fluke.
D. immitis F. hepatica
0
95"0 0 208"0 0 0
--
25 "2 0 0 0 76"8 176"8
65"2 14"4 3"5 1"8 0 23" 1 70"6 0 1375"5 1 "8 0
Mitochondrial fraction
Soluble fraction
D. immitis
15 "4 4"8 0 17"2 0 23 1"5 61 111 2180 426 22
Soluble fraction
15"3 0 0 3"2 13"2 283 12 0 238 2010 1685 222
Mitochondrial fraction
F. hepatica*
The following abbreviations are used: suc., succinate; fum., fumarate; mal., malate; oaa., oxaloacetate. * Prichard & Schofield (1968).
Citrate synthase Aconitase Isocitrate dehydrogenase (NAD-specific) lsocitrate dehydrogenase (NADP-specific) ~-Ketoglutarate dehydrogenase Succinate (suc.-+fum.) dehydrogenase (fum.~suc.) Fumarase Malate (mal.-->oaa.) dehydrogenase (oaa.-+mal.) Glutamate ( N A D H 2-specific) dehydrogenase (NADPHz)
C O M P A R E D T O A C T I V I T Y LEVELS OF C O R R E S P O N D I N G
Enzyme activity (m/zmole product formed/min per mg protein)
ENZYMES IN
O F E N Z Y M E S OF T H E T R I C A R B O X Y L I C A C I D C Y C L E I N
Enzyme
TABLE 1--ACTIVITY
z
O
t" t~
Z
¢3
~o G~
TRICARBOXYLIC
ACID
CYCLE
ENZYMES
IN DIROF1LARIA IMMITIS
497
A higher level of aconitase is noted in D. immitis than in F. hepatica. In both organisms this enzyme is confined to the soluble fraction. In contrast to the liver fluke, both NAD- and NADP-speeific isocitrate dehydrogenase activity in D. immitis is confined to the soluble fraction. The enzyme specific for NADP is found in both fractions in F. hepatica but none specific for NAD occurs. A higher level of ~-ketoglutarate dehydrogenase activity is found in D. immitis than in F. hepatica. This enzyme is confined to the mitochondrial fraction in both organisms. i Succinate dehydrogenase activity catalyzing the forward reaction is high in D. immitis, comparing very closely to that found in F. hepatica. The mitochondrial fraction was assayed for activity of this enzyme in the reverse reaction. Although no measurable levels were detected in D. immitis, both fractions possess activity in this direction in F. hepatica. D. immitis shows a high level of fumarase activity in the mitoehondrial fraction while the liver fluke has none. Approximately equal levels are present in the soluble fractions of both organisms. D. immitis apparently lacks malate dehydrogenase activity in the forward direction although a very active reverse reaction occurs. Although F. hepatica does possess activity of this enzyme in both directions, the reverse reaction is likewise much more active. A very low level of only NADH2-speeific glutamate dehydrogenase occurs in D. immitis and is confined to the soluble fraction. In contrast F. hepatica possesses glutamate dehydrogenase reacting with both NADH 2 and NADPH 2 in significant amounts in both the soluble and mitochondrial fractions. DISCUSSION Although no enzymatic study of this type has been performed previously with D. immitis, electron micrographs of the intracellular structure of this worm indicate that only a few rudimentary mitochondria are present in this organism (Lee & Miller, 1967, 1969). Electron micrographs of unsuspended mitochondrial fractions used in the present study substantiate these previous findings. Although the importance of the tricarboxylic acid cycle would seem questionable in view of this information, the presence of all the enzymes of this cycle has been demonstrated in the present work. The presence of citrate synthase indicates that D. immitis is able to synthesize citrate, the first intermediate of the tricarboxylic acid cycle. Ball (1965) indicates that some of the citrate may be diverted into other systems such as the citratemalate cycle for lipogenesis, but the citrate cleavage enzyme is yet to be demonstrated in D. immitis. Studies of aconitase activity in normal and anaerobically grown yeast have shown that oxygen deficiency greatly reduces the activity of this enzyme (Hirsch, 1952). Likewise, isocitrate dehydrogenase activity is high in tissues which possess a predominant aerobic type of metabolism (Ochoa & Weisz-Tabori, 1948). The low
498
K.M. McNEILL ANDW. F. HUTCHISON
levels of these enzymes in D. immitis suggest that the tricarboxylic acid cycle may not be of major importance in this parasite. This correlates closely with the situation found in F. hepatica (Prichard & Schofield, 1968). Work with T. spiralis (Goldberg, 1957) and A. lumbricoides (Oya et al., 1963, 1965; Smith et al., 1963) indicates that these parasites possess only NADP-specific isocitrate dehydrogenase. Prichard & Schofield (1958) report similar findings in F. hepatica, However, the present work has demonstrated isocitrate dehydrogenase specific for both NAD and NADP in D. immitis although the activity with each coenzyme is very low. Agosin & Repetto (1963) also found isocitrate dehydrogenase specific for both NAD and NADP in E. granulosus scolices. Isocitrate dehydrogenase specific for NAD occurs almost solely in the mitochondria of rat liver, while the NADP-specific enzyme is more concentrated in the extra mitochondrial fraction (Ernster & Navazio, 1956; Prichard & Schofield, 1968). In contrast, isocitrate dehydrogenase specific for each coenzyme is present in measurable amounts only in the soluble fraction in D. immitis. D. immitis possesses a very active ~-ketoglutarate dehydrogenase. The possible importance of this enzyme is enhanced by low glutamate dehydrogenase activity, the enzyme which competes with the former for ~-ketoglutarate. It appears that the oxidative decarboxylation of ~-ketoglutarate to succinate by ~-ketoglutarate dehydrogenase predominates as the major reaction at this step in the tricarboxylic acid cycle in D. immitis. A very active glutamate dehydrogenase in F. hepatica tends to indicate a relative unimportance of ~-ketoglutarate dehydrogenase in the fluke (Prichard & Schofield, 1968). Only NAD-specific glutamate dehydrogenase is present in D. immitis. Dixon & Webb (1964) state that only liver glutamate dehydrogenase shows no coenzyme specificity while this enzyme in other animal tissues is specific for NAD. Glutamate dehydrogenase activity was found to proceed in the presence of NAD or NADP in the liver fluke (Prichard & Schofield, 1968). These authors postulate that a NAD-specific as well as a non-specific glutamate dehydrogenase exists in F. hepatica since a NADP-specific enzyme has not yet been reported in animal tissues. D. immitis has a high succinate dehydrogenase activity in the forward direction but the reverse reaction could not be detected in this study. Prichard & Schofield (1968) reported both the forward and reverse reactions in F. hepatica, but they found relatively low activity in the reverse reaction. Although this indicates an aerobictype of metabolism, neither they nor van Grembergen (1949) were able to show marked inhibition of succinate oxidation in F. hepatica by addition of malonate, an effect readily demonstrated in rat liver, a predominately aerobic tissue. Further studies may reveal a similar situation in D. immitis. Although Prichard & Schofield (1968) reported that fumarase activity is confined to the soluble fraction in F. hepatica, high activity of fumarase is present in both the soluble and mitochondrial fractions of D. immitis. The particulate fraction of A. lumbricoides also contains fumarase activity (Oya et al., 1965). Every attempt to demonstrate the forward reaction of malate dehydrogenase in D. immitis was unsuccessful although a highly active reverse reaction is present. If
T R I C A R B O X Y L I C ACID CYCLE ENZYMES I N D I R O F I L A R I A I M M 1 T I S
499
malate dehydrogenase activity in the forward direction is absent in this parasite, malate may possibly be converted to oxaloacetate by some alternate pathway. One such possible pathway could be the conversion of malate to pyruvate by the malic enzyme with pyruvate being carboxylated to form oxaloacetate by the action of pyruvate carboxylase. Obviously the synthesis of oxaloacetate by some mechanism is essential to allow for the formation of citrate in the initial step of the tricarboxylic acid cycle. D. immitis possesses a high activity of enolase which acts to form phosphoenolpyruvate but only a low level of pyruvate kinase activity (Hutchison & McNeill, 1970). Phosphoenolpyruvic acid carboxykinase may compete with pyruvate kinase for phosphoenolpyruvate providing another possible source of oxaloacetate. Future studies are planned to determine the actual fate of malate at this step in the tricarboxylic acid cycle and to determine the major source of oxaloacetate for citrate synthesis in this parasite. Prichard and Schofield (1968) found malate dehydrogenase activity in both directions in F. hepatica although the reverse reaction was far more active. They suggest that malate may be metabolized to succinate by a partial reversal of the tricarboxylic acid cycle which may then be converted to the propionic acid excreted by this fluke. The highly active reverse reaction of malate dehydrogenase in D. immitis indicates that a similar situation may occur in this parasite. Further exploration in this area is required before definite conclusions can be formed. Although D. immitis contains all of the enzymes of the tricarboxylic acid cycle, the low activity levels of aconitase and isocitrate dehydrogenase plus the apparent absence of malate dehydrogenase activity in the forward direction suggest that the complete cycle may be of relatively minor importance in the overall metabolism of this parasite. Similar conclusions have been reported concerning the importance of the complete cycle in A . lumbricoides (Oya et al., 1965) and in F. hepatica (Prichard & Schofield, 1968). Acknowledgements--The authors are indebted to Professors J. Neal Brown and Joseph L. Haining for the assistance and encouragement they have shown during the course of this work.
REFERENCES AOOSINM. & REPErrO Y. (1963) Studies on the metabolism of Echinococcus granulosus--VI I. Reactions of the tricarboxylic acid cycle in E. granulosus scolices. Comp. Biochem. Physiol. 8, 245-261. ANFINSENC. B. (1955) Aconitase from pig heart muscle. In Methods in Enzymology (Edited by COLOWmKS. P. & KaPL~ N. O.), Vol. 1, pp. 695-698. AcademicPress, New York. BALL E. G. (1965) Regulation of fatty acid synthesis in adipose tissue. In Advances in Enzyme Regulation (Edited by WEBERG.), Vol. 4, p. 11. Pergamon Press, Oxford. COSTELLOL. C. & BROWNH. (1962) Aerobic metabolism of unembryonated eggs of Ascaris lumbricoides. Expl Parasitol. 12, 33-40. DIXON M. & WEBBE. C. (1964) Enzymes, pp. 686--689. Longmans, Green, London. ERNSTERL. & NAVAZlOF. (1956) The cytoplasmic distribution of isocitric dehydrogenase. Expl Cell Res. 11, 483-486. GOEBEr.L H. & KLXNCENBERCM. (1964) DPN-spezifische isoCitratedehydrogenase der Mitochondrien--I. Kinetische Eigenschaften, Vorkommen und Funktion der DPNspezifischen isoCitratedehydrogenase. Biochem. Z. 340, 441-464.
500
K. M. McNEILL AND W. F. HUTCHISON
GOLDBERG E. (1957) Studies on the intermediary metabolism of Trichinella spiralis. Expl Parasitol. 6, 367-382. GORNALL A. G., BARDAWlLLC. J. & DAVID M. M. (1949) Determination of serum proteins by means of the biuret reaction. J. biol. Chem. 177, 751-766. HIRSCH H. M. (1952) A comparative study of aconitase, fumarase and DPN-linked isocitric dehydrogenase in normal and respiratory-deficient yeast. Biochim. biophys. Acta 9, 674-686. HUTCHISON W. F. & McNEILL K. M. (1970) Glycolysis in the adult dog heartworm, Dirofilaria immitis. Comp. Biochem. Physiol. 35, 721-727. LEE C. C. & MILLER J. H. (1967) Fine structure of the body-wall musculature of Dirofilaria immitis. Expl Parasitol. 20, 334-344. LEE C. C. & MILLER J. H. (1969) Fine structure of the intestinal epithelium of Dirofilaria immitis and changes occurring after vermicidal treatment with Caparsolate Sodium. J. Parasitol. 55, 1035-1045. MASSEY V. (1960) T h e composition of the ketoglutarate dehydrogenase complex. Bioehim. biophys. Acta 38, 447-460. OCHOA S. (1955a) Crystalline condensing enzyme from pig heart. In Methods in Enzymology (Edited by COLOWICK S. P. & KAPLAN N. O.), Vol. 1, pp. 685-694. Academic Press, New York. OcnoA S. (1955b) Isocitric dehydrogenase system (TPN) from pig heart. In Methods in Enzymology (Edited by COLOWICK S. P. & KAPLAN N. O.), Vol. 1, pp. 699-704. Academic Press, New York. OCHOA S. & WEIsz-TABoRI E. (1948) Biosynthesis of tricarboxylic acids by carbon dioxide fixation--II. Oxalosuccinic carboxylase. J. biol. Chem. 174, 123-132. OVA H., COSTELLOL. C. & SMITH W. (1963) Succinate and malate oxidation and cytochrome c reduction in embryonated and unembryonated Ascaris eggs. J. Parasitol. 49 (5, Sect. 2) 51. OYA H., KIKUCHI G., BANDO T. & HAYAS8I H. (1965) Muscle tricarboxylic acid cycle in Ascaris lumbricoides vat. suis. Expl Parasitol. 17, 229-240. PRICHARD R. K. & SC8OFIELDP. J. (1968) A comparative study of the tricarboxylic acid cycle enzymes in Fasciola hepatica and rat liver. Comp. Biochem. Physiol. 25, 1005-1019. RACKER E. (1950) Spectrophotometric measurements of the enzymatic formation of fumaric and cis-aconitic acids. Biochim. biophys. Acta 4, 211-214. SCHMIDT E. (1963) Glutamate dehydrogenase. In Methods of Enzymatic Analysis (Edited by BERGMEYER H. U.), pp. 752-756. Verlag Chemic, Academic Press, New York. SHEPHERD D. & GARLAND P. B. (1969) Citrate synthase from rat liver. In Methods in Enzymology (Edited by LGWENSTEIN J. M.), Vol. 13, pp. 11-16. Academic Press, New York. SHONK C. E. & BOXER G. E. (1964) Enzyme patterns in human tissues--I. Methods for the determination of glycolytic enzymes. Cancer Res. 24, 709-721. SLATER E. C. & BONNER W. D. (1952) The effect of fluoride on the succinic oxidase system. Biochem.ff. 52, 185-196. SMITH W., COSTELLO L. C. & OYA H. (1963) Dehydrogenase activity in unembryonated and embryonated Ascaris eggs. ft. Parasitol. 49, (5, Sect. 2), 51. VAN GREMBERGEN G. (1949) Le metabolisme respiratoire du trematode Fasciola hepatica Linn. Enzymologia 13, 241-257.
Key Word Index--Dirofilaria immitis; tricarboxylic acid cycle; citrate synthase; aconitase; isocitrate dehydrogenase; ~-ketoglutarate dehydrogenase; succinate dehydrogenase; fumarase; malate dehydrogenase; glutamate dehydrogenase.