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Cytochrome c oxidase in migrating larvae of Ascaris lumbricoides var. suum
EXPERIMENTAL
PARASITOLOGY
24, 32-36
(1969)
Cytochrome c Oxidase in Migrating Larvae Ascaris lombricoides var. suuml Stephen Johns Hopkins (Submitted
of
R. Sylk2
School of Hygiene, for publication,
Baltimore,
Maryland
13 September
21205
1968)
SYLX, S. R. 1969. Cytochrome c Oxidase in Migrating Larvae of Ascaris lumbricodes var. suum. Experimental Parasitology 24, 32-36. The metabolism of Ascaris eggs is known to be aerobic, while that of the adults is anaerobic. The developmental stage in which this metabolic transition occurs was investigated by comparing the cytochrome c oxidase activity of migrating larvae recovered from swine and guinea pig lungs with that of fourth-stage larvae recovered from the swine intestine, or produced by in uitro exsheathment of lung-stage larvae from guinea pigs. Cytochrome oxidase activity was observed in lung larvae from both hosts but not from fourthstage larvae, indicating that the transition from aerobic to anaerobic metabolism occurred after the emergence of the fourth-stage larvae. INDEX DESCRIPTORS: Ascaris Zumbricoides, cytochrome oxidase of; Cytochrome oxidase, in larvae of Ascaris lumbricoides; Exsheathment, of Ascaris lumbricoides larvae; Aerobic metabolism, of Ascaris larvae; Anaerobic metabolism, of Ascari.s larvae.
enzyme by direct assay. Kmetec, Beaver, and Bueding (1963) demonstrated cytochrome c oxidase activity in extracts of aerobically and anaerobically cultured eggs, and Costello, Oya, and Smith (1963) and Oya, Costello, and Smith ( 1963) showed an increase in enzymatic activity which could be correlated with egg development during embryonation. In recent studies, Saz et al. (1968) found that third-stage larvae were able to oxidize reduced cytochrome c. Thus, an active cytochrome system has been shown to exist through the third larval stage. Accordingly, these stages of the life cycle are aerobic ( Faure-Fremiet, 1912; Brown, 1928; Jaskoski, 1952; Passey and Fairbairn, 1955). On the other hand, the adult nematodes, which reside in an essentially anaerobic environment (Hobson, 1948) lack this enzyme. These findings indicate that a transition from the one type of metabolic process to the other might occur during some stage of larval development.
Studies on the metabolism of Ascaris lumbricoides var. suum have been conducted either on eggs, newly hatched larvae, or adult worms. Bueding and Charms (1952) demonstrated that adult ascarids possess no cytochrome oxidase activity in muscle or reproductive organs. This was confirmed by Rathbone ( 1955), Fairbairn ( 1957), Kmetec and Bueding ( 1961), and Chance and Parsons ( 1963). Passey and Fairbairn ( 1955) presented indirect evidence for the presence of cytochrome c oxidase in developing eggs, but were unable to identify the 1 This paper is a portion of a thesis submitted in partial fulfillment of the requirements for the degree of Doctor of Science to the Johns Hopkins School of Hygiene and Public Health, Baltimore, Maryland, 1967. This study was supported by USPHS Grant 5 TO1 A100020 and USPHS NIAID Grant AI 01508. a Present address: Department of Parasitology, University of Pennsylvania School of Veterinary Medicine, Philadelphia, Pennsylvania. 32
CYTOCHROME OXIDASE IN Ascaris lumbricoides METHODS AND MATERIALS
Source of Larvae Third-stage larvae were recovered from the lungs of experimentally infected guinea pigs on days 7-10 post-infection; and from experimentally-infected swine on days 7 and 8. Fourth-stage larvae were isolated from the small intestine of swine autopsied after infection on day 13. In order to obtain fourth-stage larvae from guinea pigs, methods of in vitro cultivation were employed. Previous studies (Sylk, 1967) have established that third-stage larvae recovered from the lungs of guinea pigs were capable of accomplishing the third ecdysis in culture. The technique was one previously described by Schiller (1965) for the cultivation of Hymenolepis diminuta. In all cases, larvae were frozen at -20°C prior to their preparation for enzymatic analysis. Preparation of Larvae for Biochemical Analysis ( Fig. 1) After thawing, a crude preparation was made by homogenizing the larvae in a teflon tissue grinder with 0.1 M PO4 buffer ( pH 7.4). A portion of this preparation was retained for biochemical analysis and the remainder was centrifuged at 1000 rpm for
Homogemzed
Centrifuged
(1000
The oxidation of reduced cytochrome c by larval homogenates was recorded spectrophotometrically at 550 mu at room temperature (Smith, 1955). Solutions of reduced cytochrome c were prepared by adding a few crystals of Na&O* to the oxidized form and excess reducing agent was destroyed by aeration. Protein determinations were carried out on all homogenates according to the procedure of Lowry et al. (1951). RESULTS
The results of a series of experiments designed to determine the activity of cytochrome c oxidase in larval preparations are presented in Table I. Homogenates of larvae recovered from the lungs of guinea
buffet
(pH 7.4)
I Sediment
112,000
rpm
for
(discarded)
20 minulesl
I Sediment
(Part~culote
c Oxidase Determinations
I rpm for 5 mtnutesl
I
Rehomogemzed
Cytochrome
homogenale)”
supernatea
Centrifuged
33
5 minutes. The sediment was discarded, and 0.1 ml of the supernate was saved for analysis. The remaining supernatant fluid was centrifuged at approximately 10,000 rpm for 20 minutes following which the sediment was rehomogenized in 0.01 M PO4 buffer ( pH 7.4). This represented the particulate fraction of the larvae and was analyzed for the presence of cytochrome oxidase.
in 0. I M PO, (Crude
var. suum
Supernate
(discarded)
I” 0.01 M POs buffer fraction)”
FIG. 1. Diagrammatic representation of the procedure for the preparation of larval homogenates. Q Indicates fractions which were assayed for the presence of cytochrome c.
34
SYLK
The Oxidation
of Reduced
Cytochrome
TABLE I c by Homogenates of Larvae Infected Animals
Recovered
from
Experimentally
Cytochrome c oxidized (m~moles/min/mg protein)
Lightly spun homogenate
Particulate fraction
By larvae cultured for an additional 3 days in 95% N,5% co, -
By in vivo
Larval
source
Day of larval recovery (postinfection)
Crude homogenate
larvae
Guinea pig (lungs)
Swine
( lungs )
Swine (sm. intestines)
7
4.5
-
44.5
8 9 10
4.5 5.3 3.5
5.9 4.2 -
21.8 20.1 19.8
7 8
4.5 -
-
14.7
-
13 13
-
-
0 0
-
pigs and swine 7-10 days post-infection were shown to be capable of oxidizing reduced cytochrome c in the presence of oxygen. Therefore, an aerobic metabolism was indicated for all stages of development to this point. In view of these findings, extracts of fourth-stage larvae recovered from the small intestines of swine on day 13 post-infection, and from in vitro cultures of lung-stage larvae also were examined for their ability to oxidize reduced cytochrome c. Neither the homogenates of in vivo nor in vitro molted larvae were able to catalyze the oxidation of reduced cytochrome c as shown in Table I. To examine the possibility that these results were due to the presence of an inhibitor in the homogenates, fourth-stage larvae developed both in vivo and in vitro were examined. If such an inhibitor were present, a noticeable reduction in the enzymatic activity of a positive control system consisting of a dilute liver mitrochondrial preparation plus reduced cytochrome c would be produced. As shown in Table II (Expts. 1, 2, 3), the addition of the re-
0 0 0
homogenized residue or the supernatant fluid from in vitro molted larvae resulted in no inhibition of activity. Similarly the addition of the supernate or the residue of larvae which had molted in vivo did not result in inhibition of cytochrome oxidase activity of the positive control system. DISCUSSION
In the guinea pig, an unnatural host for A. lumbricoides var. suum, development is arrested in the lungs, where larvae remain in the third-stage of development (Sylk, 1967). Factors which preclude development beyond the third larval stage in the guinea pig host are, as yet, not fully understood. However, in vitro studies, together with cytochrome oxidase determinations suggest that the gaseous environment of the lungs is suboptimal for advanced third-stage larvae. One explanation of this host-parasite incompatibility may be that the lungs of guinea pigs are not sufficiently anaerobic for third-stage larvae to complete the molting process which is essential for further growth and development. Larvae may become arrested in the lungs due to unsuitable
CYTOCHROME
OXIDASE
IN
Ascaris lumbricoides TABLE
The Absence
Experiment 1
2
of Inhibitors
4
Laroae Cytochrome c oxidized” (m~moles/min/ me protein)
-
-
-
0.10 0.10
555.6 669.4 563.1
0.10
-
Supernate (ml)
-0
9 10 -b 8
II c Oxiduse in Fourth-Stage
35
Rehomogenized residue (ml)
Day of recovery (postinfection)
--/I 3
of Cytochrome
var. suum
-
9 10
0.10 0.10
-11 13 13
-
-
0.08
0.08 -
0 Dilute suspension of liver mitochondria. b Control for each experiment (Liver preparation
oxygen and carbon dioxide tensions, and therefore, are unable to complete their migration to the small intestines where, in the natural swine host, the third ecdysis takes place. Prolongation of their residence in the lungs of guinea pigs may be of suflicient duration for the immunological defense mechanisms of the host to encapsulate and ultimately destroy them. Since fourthstage larvae reside in the same environment as do the adults, it is conceivable that their metabolic processes would be similar. Therefore, it was postulated that Ascaris larvae make the transition from an aerobic to an anaerobic metabolism at the time of the third molt. Such a transition might be essential for the subsequent larval stage to survive and continue the next phase of its development. To test this hypothesis, larval homogenates were assayed for the presence of cytochrome c oxidase, which is responsible for terminal oxidation in most aerobic tissues. It catalyzes the oxidation of reduced cytochrome c by molecular oxygen. In mammalian tissues, the presence of the complete cvtochrome svstem has been taken as an indication of ‘an aerobic metabolism. With
584.3 555.6 435.6 569.3 420.6
-
+ reduced cytochrome
114.5 106.2 125.3 c).
respect to the helminths, this has not been clearly established, as some are capable of utilizing molecular oxygen but possess no demonstrable cytochrome system. The presence of an active cytochrome system in third-stage larvae for 10 days in the guinea pig and 8 days in swine has been demonstrated. However, fourth-stage larvae were unable to oxidize reduced cytochrome c. On the basis of these results, two possibilities were proposed concerning the inability of molted larvae to demonstrate the presence of this enzyme: (a) That an inhibitor was present in the preparation at this stage which prevented enzymatic activity; or (b) that at this stage of development, the larvae no longer possessed any cytochrome oxidase. Since no inhibition of the positive control system occurred upon the addition of larval homogenates, the hypothesis concerning the presence of an inhibitor in fourth-stage larvae was rejected. These findings, therefore, strongly suggest that once larvae have undergone the third molt to become fourth-stage, they no longer possess cytochrome c oxidase. Furthermore, the results support the hypothesis
36
SYJX
that the transition from an aerobic to an anaerobic metabolism in migrating AX&S larvae occurs during the third molt, and the subsequent emergence of fourth-stage larvae. Although the specific terminal oxidase of fourth-stage larvae has not been identified, from these studies it may be concluded that they do not possess an active cytochrome oxidase. It appears likely that the terminal oxidase is a metal catalyzed flavoprotein enzyme similar to that found in the adult worm by Kmetec and Bueding (1961). Further studies, however, are necessary before any definitive conclusion can be reached regarding the mechanism of electron transport in larvae developed beyond that of the lung stage. ACKNOWLEDGMENTS I express my thanks to Drs. E. B. Bueding, H. J. Saz, and their technicians at the Johns Hopkins School of Hygiene for their valuable advice and assistance in the performance of the biochemical determinations; to Drs. A. 0. Foster, F. Douvres, F. Tromba, and their staff at the Beltsville Animal Parasite Laboratory of the U. S. Department of Agriculture, Beltsville, Maryland, for their kind cooperation and assistance with the studies involving swine hosts; to Dr. E. L. Schiller of the Johns Hopkins School of Hygiene for his overall guidance in this project.
REFERENCES BROWN, H. W. 1928. Further studies on the longevity of Ascaris eggs. Journal of Parasitology 15, 14-22. BUEDING, E., AND CHARMS, B. 1952. Cytochrome c, cytochrome oxidase, and succinoxidase activities of helminths. Journal of Biological Chemistry 196, 615-627. CHANCE, B., AND PARSONS, D. F. 1963. Gytochrome function in relation to inner membrane structure of mitochondria. Science 142, 1176-1179. COSTELLO, L. C., OYA, H., AND SMITH, W. 1963. The comparative biochemistry of developing Ascaris eggs. I. Substrate oxidation and the cytochrome system in embryonated and unembryonated eggs. Archives of Biochemistry and Biophysics 103, 345-351.
FAIHBAIRN, Ascaris. 554.
D. 1957. The Biochemistry of Experimental Parasitology 6, 491-
FAURE-FREMIET, E.
1912. Graisse et glycogene dans le developpement de I’dscaris megalocephala. &dletin de la Societe Zoologique de France 37, 233-234. HOBSON, A. D. 1948. The physiology and cultivation in artificial media of nematodes parasitic in the alimentary tract of animals. Parasitology 38, 183-227. JASKOSKI,B. J. 1952. The protein coat in development of Ascaris lumbriocoides eggs. Erperimental Parasitology 1, 291-302. KMETEC, E., BEAVER, P. C., AND BUEDING, E. 1963. Succinoxidase activities and cytochrome oxidase of extracts of Ascaris eggs. Comparative Biochemistry and Physiology 9, 115-120. KMETEC, E., AND BUEDING, E. 1961. Succinic and reduced diphoshopyridine nucleotide oxidase systems of Ascaris muscle. Journal of Biological Chemistry 236, 584-591. LOWRY, 0. H., ROSEBROUGH,N. J., FARR, A. L., AND RANDALL,R.J. 1951. Protein measurements with the Folin phenol reagent. Journal of Biological Chemistry 193, 265275. OYA, H., COSTELLO, L. C., AND SMITH, W. N. 1963. The comparative biochemistry of developing Ascaris eggs. II. Changes in cytochrome c oxidase activity during embryonation. Journal of Cellular and Comparative Physiology 62, 287-294. PASSEY, R. F., AND FAI~AIRN, D. 1955. The respiration of Ascaris lumbricoides eggs. Canadian Journal of Biochemistry and Physiology 33, 1033-1046. RATHBONE, L. 1955. Oxidative metabolism in Ascaris Eumbricoides from the pig. Biochemical Journal 61, 574-579. SAZ, H. J., LESCURE, 0. L., AND BUEDING, E. B. 1968. Biochemical observations of Ascaris suum lung-stage larvae. Journal of Parasitology 54, 457461. SCHILLEH, E. L. 1965. A simplified method for the in vitro cultivation of the rat tapeworm, Hymenolepis diminuta. Journal of Parasitology 51, 516-518. SMITH, L. 1955. Cytochrome a, ai, az, and a3. Methods in Enzymology, III, 732. SYLK, S. R. 1967. Studies on the growth and development of migrating larvae of Ascaris Zumbricoides var. suum. Sc.D. thesis. Johns Hopkins School of Hygiene and Public Health.
PARASITOLOGY
24, 32-36
(1969)
Cytochrome c Oxidase in Migrating Larvae Ascaris lombricoides var. suuml Stephen Johns Hopkins (Submitted
of
R. Sylk2
School of Hygiene, for publication,
Baltimore,
Maryland
13 September
21205
1968)
SYLX, S. R. 1969. Cytochrome c Oxidase in Migrating Larvae of Ascaris lumbricodes var. suum. Experimental Parasitology 24, 32-36. The metabolism of Ascaris eggs is known to be aerobic, while that of the adults is anaerobic. The developmental stage in which this metabolic transition occurs was investigated by comparing the cytochrome c oxidase activity of migrating larvae recovered from swine and guinea pig lungs with that of fourth-stage larvae recovered from the swine intestine, or produced by in uitro exsheathment of lung-stage larvae from guinea pigs. Cytochrome oxidase activity was observed in lung larvae from both hosts but not from fourthstage larvae, indicating that the transition from aerobic to anaerobic metabolism occurred after the emergence of the fourth-stage larvae. INDEX DESCRIPTORS: Ascaris Zumbricoides, cytochrome oxidase of; Cytochrome oxidase, in larvae of Ascaris lumbricoides; Exsheathment, of Ascaris lumbricoides larvae; Aerobic metabolism, of Ascaris larvae; Anaerobic metabolism, of Ascari.s larvae.
enzyme by direct assay. Kmetec, Beaver, and Bueding (1963) demonstrated cytochrome c oxidase activity in extracts of aerobically and anaerobically cultured eggs, and Costello, Oya, and Smith (1963) and Oya, Costello, and Smith ( 1963) showed an increase in enzymatic activity which could be correlated with egg development during embryonation. In recent studies, Saz et al. (1968) found that third-stage larvae were able to oxidize reduced cytochrome c. Thus, an active cytochrome system has been shown to exist through the third larval stage. Accordingly, these stages of the life cycle are aerobic ( Faure-Fremiet, 1912; Brown, 1928; Jaskoski, 1952; Passey and Fairbairn, 1955). On the other hand, the adult nematodes, which reside in an essentially anaerobic environment (Hobson, 1948) lack this enzyme. These findings indicate that a transition from the one type of metabolic process to the other might occur during some stage of larval development.
Studies on the metabolism of Ascaris lumbricoides var. suum have been conducted either on eggs, newly hatched larvae, or adult worms. Bueding and Charms (1952) demonstrated that adult ascarids possess no cytochrome oxidase activity in muscle or reproductive organs. This was confirmed by Rathbone ( 1955), Fairbairn ( 1957), Kmetec and Bueding ( 1961), and Chance and Parsons ( 1963). Passey and Fairbairn ( 1955) presented indirect evidence for the presence of cytochrome c oxidase in developing eggs, but were unable to identify the 1 This paper is a portion of a thesis submitted in partial fulfillment of the requirements for the degree of Doctor of Science to the Johns Hopkins School of Hygiene and Public Health, Baltimore, Maryland, 1967. This study was supported by USPHS Grant 5 TO1 A100020 and USPHS NIAID Grant AI 01508. a Present address: Department of Parasitology, University of Pennsylvania School of Veterinary Medicine, Philadelphia, Pennsylvania. 32
CYTOCHROME OXIDASE IN Ascaris lumbricoides METHODS AND MATERIALS
Source of Larvae Third-stage larvae were recovered from the lungs of experimentally infected guinea pigs on days 7-10 post-infection; and from experimentally-infected swine on days 7 and 8. Fourth-stage larvae were isolated from the small intestine of swine autopsied after infection on day 13. In order to obtain fourth-stage larvae from guinea pigs, methods of in vitro cultivation were employed. Previous studies (Sylk, 1967) have established that third-stage larvae recovered from the lungs of guinea pigs were capable of accomplishing the third ecdysis in culture. The technique was one previously described by Schiller (1965) for the cultivation of Hymenolepis diminuta. In all cases, larvae were frozen at -20°C prior to their preparation for enzymatic analysis. Preparation of Larvae for Biochemical Analysis ( Fig. 1) After thawing, a crude preparation was made by homogenizing the larvae in a teflon tissue grinder with 0.1 M PO4 buffer ( pH 7.4). A portion of this preparation was retained for biochemical analysis and the remainder was centrifuged at 1000 rpm for
Homogemzed
Centrifuged
(1000
The oxidation of reduced cytochrome c by larval homogenates was recorded spectrophotometrically at 550 mu at room temperature (Smith, 1955). Solutions of reduced cytochrome c were prepared by adding a few crystals of Na&O* to the oxidized form and excess reducing agent was destroyed by aeration. Protein determinations were carried out on all homogenates according to the procedure of Lowry et al. (1951). RESULTS
The results of a series of experiments designed to determine the activity of cytochrome c oxidase in larval preparations are presented in Table I. Homogenates of larvae recovered from the lungs of guinea
buffet
(pH 7.4)
I Sediment
112,000
rpm
for
(discarded)
20 minulesl
I Sediment
(Part~culote
c Oxidase Determinations
I rpm for 5 mtnutesl
I
Rehomogemzed
Cytochrome
homogenale)”
supernatea
Centrifuged
33
5 minutes. The sediment was discarded, and 0.1 ml of the supernate was saved for analysis. The remaining supernatant fluid was centrifuged at approximately 10,000 rpm for 20 minutes following which the sediment was rehomogenized in 0.01 M PO4 buffer ( pH 7.4). This represented the particulate fraction of the larvae and was analyzed for the presence of cytochrome oxidase.
in 0. I M PO, (Crude
var. suum
Supernate
(discarded)
I” 0.01 M POs buffer fraction)”
FIG. 1. Diagrammatic representation of the procedure for the preparation of larval homogenates. Q Indicates fractions which were assayed for the presence of cytochrome c.
34
SYLK
The Oxidation
of Reduced
Cytochrome
TABLE I c by Homogenates of Larvae Infected Animals
Recovered
from
Experimentally
Cytochrome c oxidized (m~moles/min/mg protein)
Lightly spun homogenate
Particulate fraction
By larvae cultured for an additional 3 days in 95% N,5% co, -
By in vivo
Larval
source
Day of larval recovery (postinfection)
Crude homogenate
larvae
Guinea pig (lungs)
Swine
( lungs )
Swine (sm. intestines)
7
4.5
-
44.5
8 9 10
4.5 5.3 3.5
5.9 4.2 -
21.8 20.1 19.8
7 8
4.5 -
-
14.7
-
13 13
-
-
0 0
-
pigs and swine 7-10 days post-infection were shown to be capable of oxidizing reduced cytochrome c in the presence of oxygen. Therefore, an aerobic metabolism was indicated for all stages of development to this point. In view of these findings, extracts of fourth-stage larvae recovered from the small intestines of swine on day 13 post-infection, and from in vitro cultures of lung-stage larvae also were examined for their ability to oxidize reduced cytochrome c. Neither the homogenates of in vivo nor in vitro molted larvae were able to catalyze the oxidation of reduced cytochrome c as shown in Table I. To examine the possibility that these results were due to the presence of an inhibitor in the homogenates, fourth-stage larvae developed both in vivo and in vitro were examined. If such an inhibitor were present, a noticeable reduction in the enzymatic activity of a positive control system consisting of a dilute liver mitrochondrial preparation plus reduced cytochrome c would be produced. As shown in Table II (Expts. 1, 2, 3), the addition of the re-
0 0 0
homogenized residue or the supernatant fluid from in vitro molted larvae resulted in no inhibition of activity. Similarly the addition of the supernate or the residue of larvae which had molted in vivo did not result in inhibition of cytochrome oxidase activity of the positive control system. DISCUSSION
In the guinea pig, an unnatural host for A. lumbricoides var. suum, development is arrested in the lungs, where larvae remain in the third-stage of development (Sylk, 1967). Factors which preclude development beyond the third larval stage in the guinea pig host are, as yet, not fully understood. However, in vitro studies, together with cytochrome oxidase determinations suggest that the gaseous environment of the lungs is suboptimal for advanced third-stage larvae. One explanation of this host-parasite incompatibility may be that the lungs of guinea pigs are not sufficiently anaerobic for third-stage larvae to complete the molting process which is essential for further growth and development. Larvae may become arrested in the lungs due to unsuitable
CYTOCHROME
OXIDASE
IN
Ascaris lumbricoides TABLE
The Absence
Experiment 1
2
of Inhibitors
4
Laroae Cytochrome c oxidized” (m~moles/min/ me protein)
-
-
-
0.10 0.10
555.6 669.4 563.1
0.10
-
Supernate (ml)
-0
9 10 -b 8
II c Oxiduse in Fourth-Stage
35
Rehomogenized residue (ml)
Day of recovery (postinfection)
--/I 3
of Cytochrome
var. suum
-
9 10
0.10 0.10
-11 13 13
-
-
0.08
0.08 -
0 Dilute suspension of liver mitochondria. b Control for each experiment (Liver preparation
oxygen and carbon dioxide tensions, and therefore, are unable to complete their migration to the small intestines where, in the natural swine host, the third ecdysis takes place. Prolongation of their residence in the lungs of guinea pigs may be of suflicient duration for the immunological defense mechanisms of the host to encapsulate and ultimately destroy them. Since fourthstage larvae reside in the same environment as do the adults, it is conceivable that their metabolic processes would be similar. Therefore, it was postulated that Ascaris larvae make the transition from an aerobic to an anaerobic metabolism at the time of the third molt. Such a transition might be essential for the subsequent larval stage to survive and continue the next phase of its development. To test this hypothesis, larval homogenates were assayed for the presence of cytochrome c oxidase, which is responsible for terminal oxidation in most aerobic tissues. It catalyzes the oxidation of reduced cytochrome c by molecular oxygen. In mammalian tissues, the presence of the complete cvtochrome svstem has been taken as an indication of ‘an aerobic metabolism. With
584.3 555.6 435.6 569.3 420.6
-
+ reduced cytochrome
114.5 106.2 125.3 c).
respect to the helminths, this has not been clearly established, as some are capable of utilizing molecular oxygen but possess no demonstrable cytochrome system. The presence of an active cytochrome system in third-stage larvae for 10 days in the guinea pig and 8 days in swine has been demonstrated. However, fourth-stage larvae were unable to oxidize reduced cytochrome c. On the basis of these results, two possibilities were proposed concerning the inability of molted larvae to demonstrate the presence of this enzyme: (a) That an inhibitor was present in the preparation at this stage which prevented enzymatic activity; or (b) that at this stage of development, the larvae no longer possessed any cytochrome oxidase. Since no inhibition of the positive control system occurred upon the addition of larval homogenates, the hypothesis concerning the presence of an inhibitor in fourth-stage larvae was rejected. These findings, therefore, strongly suggest that once larvae have undergone the third molt to become fourth-stage, they no longer possess cytochrome c oxidase. Furthermore, the results support the hypothesis
36
SYJX
that the transition from an aerobic to an anaerobic metabolism in migrating AX&S larvae occurs during the third molt, and the subsequent emergence of fourth-stage larvae. Although the specific terminal oxidase of fourth-stage larvae has not been identified, from these studies it may be concluded that they do not possess an active cytochrome oxidase. It appears likely that the terminal oxidase is a metal catalyzed flavoprotein enzyme similar to that found in the adult worm by Kmetec and Bueding (1961). Further studies, however, are necessary before any definitive conclusion can be reached regarding the mechanism of electron transport in larvae developed beyond that of the lung stage. ACKNOWLEDGMENTS I express my thanks to Drs. E. B. Bueding, H. J. Saz, and their technicians at the Johns Hopkins School of Hygiene for their valuable advice and assistance in the performance of the biochemical determinations; to Drs. A. 0. Foster, F. Douvres, F. Tromba, and their staff at the Beltsville Animal Parasite Laboratory of the U. S. Department of Agriculture, Beltsville, Maryland, for their kind cooperation and assistance with the studies involving swine hosts; to Dr. E. L. Schiller of the Johns Hopkins School of Hygiene for his overall guidance in this project.
REFERENCES BROWN, H. W. 1928. Further studies on the longevity of Ascaris eggs. Journal of Parasitology 15, 14-22. BUEDING, E., AND CHARMS, B. 1952. Cytochrome c, cytochrome oxidase, and succinoxidase activities of helminths. Journal of Biological Chemistry 196, 615-627. CHANCE, B., AND PARSONS, D. F. 1963. Gytochrome function in relation to inner membrane structure of mitochondria. Science 142, 1176-1179. COSTELLO, L. C., OYA, H., AND SMITH, W. 1963. The comparative biochemistry of developing Ascaris eggs. I. Substrate oxidation and the cytochrome system in embryonated and unembryonated eggs. Archives of Biochemistry and Biophysics 103, 345-351.
FAIHBAIRN, Ascaris. 554.
D. 1957. The Biochemistry of Experimental Parasitology 6, 491-
FAURE-FREMIET, E.
1912. Graisse et glycogene dans le developpement de I’dscaris megalocephala. &dletin de la Societe Zoologique de France 37, 233-234. HOBSON, A. D. 1948. The physiology and cultivation in artificial media of nematodes parasitic in the alimentary tract of animals. Parasitology 38, 183-227. JASKOSKI,B. J. 1952. The protein coat in development of Ascaris lumbriocoides eggs. Erperimental Parasitology 1, 291-302. KMETEC, E., BEAVER, P. C., AND BUEDING, E. 1963. Succinoxidase activities and cytochrome oxidase of extracts of Ascaris eggs. Comparative Biochemistry and Physiology 9, 115-120. KMETEC, E., AND BUEDING, E. 1961. Succinic and reduced diphoshopyridine nucleotide oxidase systems of Ascaris muscle. Journal of Biological Chemistry 236, 584-591. LOWRY, 0. H., ROSEBROUGH,N. J., FARR, A. L., AND RANDALL,R.J. 1951. Protein measurements with the Folin phenol reagent. Journal of Biological Chemistry 193, 265275. OYA, H., COSTELLO, L. C., AND SMITH, W. N. 1963. The comparative biochemistry of developing Ascaris eggs. II. Changes in cytochrome c oxidase activity during embryonation. Journal of Cellular and Comparative Physiology 62, 287-294. PASSEY, R. F., AND FAI~AIRN, D. 1955. The respiration of Ascaris lumbricoides eggs. Canadian Journal of Biochemistry and Physiology 33, 1033-1046. RATHBONE, L. 1955. Oxidative metabolism in Ascaris Eumbricoides from the pig. Biochemical Journal 61, 574-579. SAZ, H. J., LESCURE, 0. L., AND BUEDING, E. B. 1968. Biochemical observations of Ascaris suum lung-stage larvae. Journal of Parasitology 54, 457461. SCHILLEH, E. L. 1965. A simplified method for the in vitro cultivation of the rat tapeworm, Hymenolepis diminuta. Journal of Parasitology 51, 516-518. SMITH, L. 1955. Cytochrome a, ai, az, and a3. Methods in Enzymology, III, 732. SYLK, S. R. 1967. Studies on the growth and development of migrating larvae of Ascaris Zumbricoides var. suum. Sc.D. thesis. Johns Hopkins School of Hygiene and Public Health.