PHOSPHOLIPID DEPENDENCY OF MEMBRANE-ASSOCIATED PYRIDINE NUCLEOTIDE-UTILIZING AND SUCCINATE DEHYDROGENASE ACTIVITIES OF ADULT HYMENOLEPIS DIMINUTA (CESTODA) AND ASCARIS SUUM (NEMATODA)
Carl Breidenbach
A Thesis
Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of
MASTER OF SCIENCE
December 2012
Committee
Carmen Fioravanti, Advisor
Jill Zeilstra-Ryalls
Raymond Larsen
© 2012
Carl Breidenbach
All Rights Reserved iii
ABSTRACT
Carmen Fioravanti, Advisor
The adult intestinal cestode, Hymenolepis diminuta, is essentially anaerobic in its metabolism and generates ATP without the need for oxygen. H. diminuta relies upon a mitochondrial NADPH→NAD transhydrogenase to link the NADPH produced by the pyruvate- forming arm of the malate dismutation reaction, catalyzed by the mitochondrial “malic enzyme”, with the NADH-requiring, anaerobic electron transport system. The electron transport-coupled fumarate reductase serves to reduce fumarate, the terminal electron acceptor, to succinate. A phospholipid dependency was established previously with respect to the transhydrogenase, fumarate reductase, and the lesser NADH oxidase. Of the phospholipids assessed, the transhydrogenase exhibited a phosphatidylcholine preference.
The present study expands on prior findings by using phospholipase A1, A2, C and D, organic solvent, and ammonium sulfate treatments of H. diminuta mitochondrial membranes.
Other reduced pyridine nucleotide-utilizing systems viz., NAD(P)H cytochrome c reductase,
NADH→NAD transhydrogenation, NAD(P)H-, and lipoamide dehydrogenase activities as well as succinate dehydrogenase were evaluated. A comparative study also was undertaken by treatment with the phospholipases of isolated mitochondrial membranes from the anaerobic intestinal nematode, Ascaris suum.
The data presented indicate a phospholipid dependence not only of the previously reported systems, but of membrane-associated mitochondrial systems in H. diminuta and A. suum. H. diminuta NADH cytochrome c reductase displayed phospholipid dependence based on phospholipase A2 and C treatments, and a neutral lipid dependence based on organic solvent iv
treatments. Ammonium sulfate fractionation had little effect. Succinate dehydrogenase activity
displayed phospholipid dependence based on phospholipase C and organic solvent treatments.
Ammonium sulfate fractionation decreased succinate dehydrogenase activity, but
phosphatidylcholine supplementation further diminished activity.
A. suum NADH cytochrome c reductase, NADH oxidase and fumarate reductase systems exhibited phospholipid dependence based on phospholipase A2 and C treatments. Interestingly,
A. suum succinate dehydrogenase appeared more resistant to phospholipase treatment than the
corresponding H. diminuta system. v
ACKNOWLEDGMENTS
I would like to thank Dr. Carmen Fioravanti for his help and support while researching
and writing this thesis. Also, special thanks for Elizabeth Shuler for helping me start my work in
the laboratory and her continued aid while doing my thesis research.
I would also like to thank my family, David, Sandra, Kurt and Dirk Breidenbach as well
as my girlfriend Brittany Murphy, for their support, especially during the writing process of this
thesis.
I would like to thank the Department of Biological Sciences at Bowling Green State
University for funding my research and a special thanks to Sigma Xi, The Scientific Research
Society for their Grant in Aid of Research. vi
TABLE OF CONTENTS
Page
CHAPTER I. LITERATURE REVIEW ......
Hymenolepis diminuta ...... 1
Ascaris suum ...... 4
Phospholipid Dependence ...... 7
Purpose ...... 9
CHAPTER II. THE STUDY
INTRODUCTION …………………………………………………………………….12
MATERIALS AND METHODS ...... 14
RESULTS
Treatment of H. diminuta mitochondrial membranes with phospholipases .. 19
Extraction of H. diminuta mitochondrial membranes with organic solvents 21
Effect of phospholipid addition to partially lipid-depleted H. diminuta
mitochondrial membrane preparations ...... 21
Treatment of Ascaris suum mitochondrial membranes with phospholipases 36
DISCUSSION ...... 47
LITERATURE CITED ...... 55
APPENDIX I: INSTITUTIONAL ANIMAL CARE AND USE COMMITTEE (IACUC)
Approval Information ...... 59 vii
LIST OF TABLES
Table Page
1 Effect of solvent extraction on reduced pyridine nucleotide-utilizing and succinate
dehydrogenase activities of adult Hymenolepis diminuta mitochondrial membranes 34
2 Effect of 30-55 ammonium sulfate fractionation and addition of
phosphatidylcholine on reduced pyridine nucleotide-utilizing and succinate
dehydrogenase activities in adult Hymenolepis diminuta mitochondrial membranes 35
3 Titration of phospholipase A2 on Hymenolepis diminuta mitochondrial membrane-
associated NADPH→NAD transhydrogenase ...... 53
viii
LIST OF FIGURES
Figure Page
1 Life cycle of Hymenolepis diminuta ...... 2
2 Pathway of primary carbohydrate utilization in Hymenolepis diminuta ...... 3
3 Life Cycle of Ascaris suum ...... 6
4 Cleavage of phospholipids by phospholipases ...... 10
5 Effects of phospholipases on percent differences in NADH cytochrome c reductase
activity of adult Hymenolepis diminuta mitochondrial membranes ...... 24
6 Effects of phospholipases on percent differences in NADPH cytochrome c reductase
activity of adult Hymenolepis diminuta mitochondrial membranes ...... 25
7 Effects of phospholipases on percent differences in NADH→NAD transhydrogenation
activity of adult Hymenolepis diminuta mitochondrial membranes ...... 26
8 Effects of phospholipases on percent differences in NADH dehydrogenase activity of
adult Hymenolepis diminuta mitochondrial membranes...... 27
9 Effects of phospholipases on percent differences in NADPH dehydrogenase activity of
adult Hymenolepis diminuta mitochondrial membranes...... 28
10 Effects of phospholipases on percent differences in succinate dehydrogenase activity of
adult Hymenolepis diminuta mitochondrial membranes...... 29
11 Effects of phospholipases on percent differences in NADH oxidase activity of adult
Hymenolepis diminuta mitochondrial membranes...... 30
12 Effects of phospholipases on percent differences in lipoamide dehydrogenase activity of
adult Hymenolepis diminuta mitochondrial membranes...... 31
ix
13 Effects of phospholipases on percent differences in fumarate reductase activity of adult
Hymenolepis diminuta mitochondrial membranes...... 32
14 Effects of phospholipases on percent differences in NADPH→NAD transhydrogenase
activity of adult Hymenolepis diminuta mitochondrial membranes ...... 33
15 Effects of phospholipases on percent differences in NADH cytochrome c reductase
activity of adult Ascaris suum muscle mitochondrial membranes ...... 38
16 Effects of phospholipases on percent differences in NADPH cytochrome c reductase
activity of adult Ascaris suum muscle mitochondrial membranes ...... 39
17 Effects of phospholipases on percent differences in NADH→NAD transhydrogenation
activity of adult Ascaris suum muscle mitochondrial membranes ...... 40
18 Effects of phospholipases on percent differences in NADH dehydrogenase activity of
adult Ascaris suum muscle mitochondrial membranes ...... 41
19 Effects of phospholipases on percent differences in NADPH dehydrogenase activity of
adult Ascaris suum muscle mitochondrial membranes ...... 42
20 Effects of phospholipases on percent differences in succinate dehydrogenase activity of
adult Ascaris suum muscle mitochondrial membranes ...... 43
21 Effects of phospholipases on percent differences in NADH oxidase activity of adult
Ascaris suum muscle mitochondrial membranes ...... 44
22 Effects of phospholipases on percent differences in lipoamide dehydrogenase activity of
adult Ascaris suum muscle mitochondrial membranes ...... 45
23 Effects of phospholipases on percent differences in fumarate reductase activity of adult
Ascaris suum muscle mitochondrial membranes ...... 46
1 CHAPER 1: LITERATURE REVIEW
Hymenolepis diminuta
Hymenolepis diminuta is an intestinal helminth primarily found in rodents, but it can also
infect man. It belongs to the phylum Platyhelminthes, the class Cestoda, the order
Cyclophyllidia, and the family Hymenolepididae. The life cycle of H. diminuta requires both an
intermediate and definitive host (Fig. 1). The infected mammalian hosts begin shedding terminal
proglottids, containing around 70,000 eggs, per day, at about 20 days post infection. Eggs are
ingested by an arthropod host, typically of the genus Tribolium, in which they develop into
cysticerci within the haemocoel. Cysticerci are the infective stage to mammals. Mammals then ingest the arthropod host and the cysticerci excyst and attach themselves onto the wall of the
small intestine where they mature into adults, and thus, continue the life cycle (Smyth, 1976).
2
Figure 1: Life cycle of Hymenolepis diminuta. From CDC website on
Hymenolepiasis (CDC 2009) (1) Eggs are passed from the host in feces. (2) The
arthropod intermediate host ingests eggs. (3) Eggs hatch and larvae penetrate
intestinal wall. (4) Cysticerci mature in the haemocoel, and are now able to infect
the mammalian host. (5) Cysticerci attach to the small intestine via the scolex. (6)
Cysticerci mature into adult tapeworms. (7) The sexually mature tapeworms
release gravid proglottids into feces.
Adult H. diminuta is essentially energetically anaerobic. Its primary metabolic pathway utilizes glucose as the initial energy source (Fig 2). Glucose is oxidized to phosphoenolpyruvate
(PEP) via the glycolytic sequence. CO2 is fixed into PEP to form oxaloacetate, which is reduced to form malate. Malate is utilized by the mitochondria wherein the “malic enzyme” catalyzes one arm of a dismutation reaction resulting in the formation of pyruvate, CO2, and NADPH. Malate
is also converted to fumarate (Fioravanti and Saz, 1980). Fumarate serves as the final acceptor
of the electron transport system in H. diminuta thereby resulting in ATP generation and the
accumulation of succinate as the major end product, while lesser amounts of lactate and acetate 3 are released (Watts and Fairbairn, 1974). H. diminuta utilizes an inner membrane-associated and
reversible NADPH→NAD transhydrogenase that plays a vital role in this anaerobic system in
that it joins the “malic enzyme’s” NADPH-forming activity with the NADH-requiring, electron transport-coupled fumarate reductase system (Saz et. al., 1972; Fioravanti and Saz, 1976). Also present in H. diminuta mitochondria is a less active NADH oxidase system (Fioravanti, 1982).
This system appears to be rotenone-sensitive and apparently requires a tightly bound manganous ion (Fioravanti, 1981; Fioravanti and Reisig, 1990).
Figure 2: Pathway of primary carbohydrate utilization in Hymenolepis
diminuta (Fioravanti and Saz, 1980; Fioravanti and Saz, 1976; Saz and Lescure,
1969).
Independent of the crucial NADPH→NAD transhydrogenase linking the malate
dismutation reaction to the energy-forming electron transport system, there is an NADH→NAD 4 transhydrogenation reaction in H. diminuta mitochondria (Fioravanti and Saz, 1976). This
NADH→NAD transhydrogenation seems to be the result of the mitochondrial membrane- associated lipoamide dehydrogenase and NADH dehydrogenase systems (Walker and Fioravanti,
1995; Walker et. al., 1997). It was has been established that a mitochondrial NADH→NAD transhydrogenation is associated with the lipoamide dehydrogenase in Ascaris suum mitochondria. Furthermore, it has been proposed that the NADH→NAD transhydrogenation is responsible for transmembrane transport of reducing power across the inner membrane (Köhler and Saz, 1976; Komuniecki and Saz, 1979).
H. diminuta mitochondria exhibit an NAD(P)H cytochrome c reductase, with NADH being the favored reductant. Rotenone-sensitive cytochrome c reductase is inner membrane associated and an outer membrane, rotenone-insensitive activity is noted. H. diminuta does not appear to have catalase, glutathione peroxidase or NAD(P)H peroxidase activities, but does have peroxide forming superoxide dismutase activity (Paul and Barret, 1980; Barret and Beis, 1983;
McKelvey and Fioravanti, 1986). Based on the finding that H. diminuta possesses a cytochrome c peroxidase activity, it has been suggested that the cytochrome c oxidase and reductase systems may serve to remove peroxides accumulated via the peroxide forming NADH oxidase system that has been noted in H. diminuta (Kim and Fioravanti, 1985).
Ascaris suum
Ascaris suum, a variant of Ascaris lumbricoides, is a helminth belonging to the phylum
Nemadota, class Secernentea, order Ascaridida, and family Ascaridae. It is capable of infecting both man, and very commonly pigs; being found throughout the world. There appear to be two different subspecies of A. lumbricoides, viz., Ascaris lumbricoides lumbricoides which is the prevalent type in human infection, and Ascaris lumbricoides suum; the common type found in 5 swine (Smyth, 1976). There has been some debate as to whether or not A. suum and A.
lumbricoides are the same species. Recent molecular biological and paleoparasitological evidence indicates that they are in fact the same species (Leles, 2012). For consistency, the A.
suum designation will be used for the nematode from swine. The life cycle of A. suum differs
from that of H. diminuta in that the ascarids require no intermediate host and undergo four molts
in progressing from egg to adult (i.e. a direct life cycle) (Fig. 2). Two of these molts occur while
the nematode is still in the egg, the third molt occurs when the larval stage is transitioning from
the alimentary tract to the lungs of the host. The fourth molt occurs during the transition
between juvenile and adult, after the larvae have left the lungs and entered the small intestine.
Adult female worms shed up to 200,000 eggs per day, but the eggs do not become infective until
they have embryonated over a period of 18 days to three weeks outside of the host. After
embryonated eggs are ingested, the larvae hatch and make their way through the intestinal
mucosa, into the portal and ultimately the systemic circulatory system, and arrive in the lungs of
the host. Here they mature for roughly two weeks before penetrating the alveolar walls and
migrate up the throat where they are swallowed. The larvae then enter the small intestine where
they develop into adults (Smyth, 1976). 6
Figure 3: Life cycle of Ascaris suum. From CDC website on Ascariasis. (1) Adult
female releases eggs that were fertilized by males. (2) Fertilized eggs begin developing
into infective larvae. (3) Larvae still in the egg are ingested by the host (4). (5) Larvae
exit the egg, pass through small intestine into liver and migrate via the portal system (6)
into the lungs where they develop further. (7) Now mature larvae are aspirated by the
host and swallowed. Larvae enter the small intestine where they mature into the adult
round worm.
A. suum displays essentially the same carbohydrate utilization pathway as described for
H. diminuta with the exception that it does not display an NADPH→NAD transhydrogenase, instead, the “malic enzyme” catalyzed arm of the mitochondrial dismutation results in NADH formation instead of NADPH, and other end products are formed from succinate (Saz and
Vidrine, 1959; Saz and Lescure, 1969).
7 Phospholipid Dependence
Previous studies demonstrated that multiple mitochondrial, membrane-associated,
pyridine nucleotide-utilizing systems in mammalian, free-living invertebrate and parasite models
display a phospholipid dependence. When the phospholipid components of the mitochondrial
membrane are disrupted, activities are diminished. This was demonstrated by Fioravanti and
Kim (1983) in H. diminuta in which the mitochondrial NADPH→NAD transhydrogenase, fumarate reductase, and NADH oxidase all showed a significant reduction in activity with
phospholipid alteration via the action of phospholipases A2 and C, with phospholipid extraction
using increasing aqueous acetone following hexane treatment, and with phospholipid removal via a 30-55 ammonium sulfate fractionation of mitochondrial membranes. Activity was restored in the 30-55 fraction by the addition of phosphatidylcholine, whereas addition of phosphatidylethanolaime and phosphatidylserine was without a significant effect on depleted membranes (Fioravanti and Kim, 1983).
A similar study done by Vandock et. al. (2011) demonstrated a phospholipid dependence of the mitochondrial NADPH→NAD transhydrogenase in the tobacco hornworm Manduca sexta. Again, activity of the system was diminished with phospholipase A2 and C treatments, phospholipid depletion using increasing aqueous acetone following hexane treatment, and with ammonium sulfate fractionation of mitochondrial membranes. Of the phospholipids examined, the only phospholipid shown to restore any significant activity to the ammonium sulfate fractionated preparation was phosphatidylcholine.
The effects of phospholipid depletion on membrane-associated bovine heart mitochondrial NADPH→NAD transhydrogenase were examined by Rydström et. al. (1976).
Their study revealed that phospholipase A and C treatments diminished the activity of the transhydrogenase, with A having a three-fold greater reduction in activity over C. Furthermore, 8 increasing aqueous acetone extraction of the phospholipids substantially reduced the activity of
the transhydrogenase. Supplementation with phospholipids of the 30-55 fraction restored
activity of the preparation, with cardiolipin being the single best phospholipid in restoring
activity. A mixture of mitochondrial lipids had an even greater reactivation of activity.
Rat liver mitochondrial cytochrome c oxidase showed a significant reduction in activity in comparison of young rats versus aged rats, with the younger rats showing the higher activity.
While both Km values and content of the cytochrome c oxidase for both aged and young rats
remained nearly constant, the maximal velocity was decreased by almost 30% from young to old.
A change in the amount of cardiolipin present in the inner membranes of the rats was noted; aged
rats had decreased levels of cardiolipin in comparison to younger rats. This decrease of the
cytochrome c oxidase activity was attributed to the phospholipids retention of or contribution to
the electrostatic and conformational properties of the cytochrome c oxidase (Paradies et. al.,
1993).
A study of the microsomal, membrane associated UDP-glucuronyltransferase of guinea pig liver demonstrated a phospholipid dependence (Singh, 1982). In this study phospholipids were removed from the purified enzyme using hydroxyapatite column chromatography (Singh,
1980). The treated enzyme displayed a significant reduction in activity, and increasing phospholipid removal caused a reduction in activity, up to a maximum of about 66%. Further evaluation indicated that the secondary structure of the enzyme changed from between 16-27%
α-helix to only 2-5% α-helix composition. This indicated that the helical structure of the enzyme depended on interactions with the phospholipids in which it was embedded for conformation retention.
9 Purpose
In the present study, a comparison was performed to evaluate how a number of pyridine-
nucleotide coupled, membrane-associated mitochondrial systems as well as the membrane- associated succinate dehydrogenase of both H. diminuta and A. suum responded to phospholipid disruption or depletion. The effects of phospholipid disruption using phospholipases A1, A2, C
and D were investigated. As given in Figure 4, phospholipase activities act as follows: A1 and A2
remove the acyl- group from the glycerol backbone at C1 and C2, respectively. Phospholipase C
removes the phosphate and polar group from C3 while D removes only the polar group
(Djordjevic, 2010).
10
Figure 4: Cleavage of phospholipids by phospholipases. Phospholipase A1
cleaves the fatty acid at the sn1 (C1) position while phospholipase A2 cleaves the
fatty acid at the sn2 (C2) position. The phosphodiester bond nearer the glycerol
backbone is cleaved by phospholipase C yielding diacylglycerol. Phospholipase
D cleaves the phosphodiester bond nearer the polar head group yielding
phosphatidic acid and the polar head group.
In addition to evaluating the effects of phospholipases, the removal of both membrane neutral lipids through hexane extraction and removal of phospholipids through aqueous acetone extraction following hexane extraction of lyophilized membrane preparations was investigated.
A 30-55 ammonium sulfate precipitation of protein was also employed to deplete membranes of
phospholipids. Restoration of the activities of these partially lipid depleted membranes using 11 phosphatidylcholine was evaluated. The following systems in both H. diminuta and A. suum were examined: NAD(P)H cytochrome c reductase, NAD(P)H dehydrogenase, NADH-fumarate reductase, lipoamide dehydrogenase, NADH oxidase, NADH → NAD transhydrogenation and
succinate dehydrogenase, and the NADPH → NAD transhydrogenase in H. diminuta
mitochondrial membranes. To date, there are little data on the phospholipid components of
invertebrate mitochondria generally and parasitic helminth mitochondria specifically. With the
exception of the study by Fioravanti and Kim (1983) on the cestode H. diminuta, little is known
of any other mitochondrial membrane associated enzymes in helminth systems with respect to
phospholipids. Accordingly, an investigation of what are the important phospholipid or neutral
lipid constituents for retention of activity in these membrane-associated systems is warranted.
Moreover, nothing is known of the phospholipid dependence of other helminth mitochondrial
systems, and for this reason the study was expanded to include the nematode A. suum. Given the
impact of their anaerobic energy metabolism and potential phospholipid dependencies, one
would think that investigation of these enzyme systems could serve in the development of
chemotherapeutic agents and in an understanding of the biochemistry of membrane systems in
general. This study extends to a wider array of mitochondrial systems in cestodes and nematodes, and provides a broader view of the importance of phospholipids generally. 12 CHAPTER 2: THE STUDY
INTRODUCTION
The adult intestinal cestode, Hymenolepis diminuta, is essentially energetically anaerobic
and its mitochondria generate ATP without the need for oxygen (Fioravanti and Saz, 1980). H.
diminuta uses malate as the mitochondrial substrate. Upon entering the mitochondrion, the
NADP-specific “malic enzyme” acts to catalyze one arm of a dismutation reaction thereby
forming pyruvate, CO2 and reducing power in the form of NADPH. However, NADH is
required for anaerobic electron transport. Therefore, H. diminuta relies upon an inner
membrane-associated and reversible NADPH→NAD transhydrogenase that plays a vital role in
this anaerobic system in that it joins the “malic enzyme’s” NADPH-forming activity with the
NADH-requiring electron transport system (Scheibel and Saz, 1966; Saz et. al. 1972; Fioravanti and Saz, 1980). Fumarate serves as the terminal acceptor of the mitochondrial anaerobic electron transport system in H. diminuta, resulting in ATP generation and succinate accumulation as the major end product (Scheibel and Saz, 1966; Watts and Fairbairn, 1974). H. diminuta also possesses a lesser peroxide forming NADH oxidase, though it seems to be of limited energetic importance (Fioravanti, 1981).
A phospholipid dependency of the H. diminuta mitochondrial NADPH→NAD transhydrogenase has been demonstrated. In addition, a phospholipid dependency was also apparent in terms of the NADH-utilizing fumarate reductase and NADH oxidase systems
(Fioravanti and Kim, 1983).
A broader study of the necessity of phospholipids in H. diminuta mitochondria by examining the effects of phospholipases A1, A2, C, and D, hexane/acetone extraction, and 30-55
ammonium sulfate fractionation on a number of membrane-associated, pyridine nucleotide-
utilizing and succinate dehydrogenase systems in the cestode H. diminuta was performed. The 13 adult intestinal nematode, Ascaris suum, also displays a similar energetic pathway as that of H. diminuta with the exception that it lacks an apparent NADPH→NAD transhydrogenase; its
“malic enzyme” yields NADH instead of NADPH. Additionally, A. suum produces a number of volatile fatty acids as end products (Saz and Vidrine, 1959; Saz and Lescure, 1969). Therefore, a comparative study of the effects of phospholipases on pyridine nucleotide-utilizing and succinate dehydrogenase activities was performed in the nematode A. suum. 14 MATERIALS AND METHODS
Male or female Sprague-Dawley rats were infected with 10 cysticercoids each by intubation. The beetle, Tribolium confusum, was employed as the intermediate host. Adult
Hymenolepis diminuta were obtained from infected rats after 21 or more days post-infection and helminth tissue was prepared for mitochondrial extraction using methods essentially as described by Fioravanti and Saz (1976).
Adult female A. suum were obtained from Routh Packing in Sandusky, Ohio. Nematode muscle tissue was isolated using techniques essentially as described by Saz and Lescure (1969).
Tissue was minced in mitochondrial medium comprised of 0.24M sucrose, 0.15% bovine serum albumin (BSA), and 0.01M Tris (hydroxymethylaminomethane)-HCl (pH 7.5) then homogenized using a Wheaton glass homogenization tube equipped with a Teflon pestle.
Homogenized tissue was subjected to centrifugation at 482 g for 10 min using a Sorvall RC2-B centrifuge. Mitochondria were isolated from the resulting supernatant via centrifugation at 9770 g for 30 min. Mitochondria were washed once using mitochondrial medium and mitochondria were obtained following centrifugation at 9770 g for 30 min (Fioravanti, 1981). Isolated mitochondria were sonicated using five successive 15-sec bursts with 30-sec cooling intervals using a Branson Sonifier equipped with a micro-tip using a power setting of 20 W.
Mitochondrial membranes were isolated via ultracentrifugation of disrupted organelles at
269,000 g for 60 minutes. Isolated membranes were then suspended in mitochondrial medium.
Helminth recoveries as well as mitochondrial and membrane isolations were performed at 4º C.
Treatments of H. diminuta mitochondrial membranes with phospholipases and solvent extraction of phospholipids were conducted as described previously (Rydström et. al., 1976;
Fioravanti and Kim, 1983). Membrane aliquots were incubated at 37 º C for 15 min in a treatment volume of 0.95 ml. Aside from mitochondrial membranes, equivalent to 1.8 mg 15 protein, the incubation contained the following: 5.0 mg BSA, 100 µmol Tris-HCl (pH 7.5), 2.5
µmol CaCl2 and 5 units phospholipase (A1, A2, C or D). Incubation was terminated by the
addition of 0.10 ml (10 µmol) ethylendiaminetetracetate (EDTA, pH 7.5). Two controls were
used, one in which 10 µmol EDTA was added prior to incubation and the other where no
phospholipase was added to the incubation volume.
Preparation of mitochondrial membranes for solvent extraction was accomplished by first
freezing the membrane suspension, equivalent to 10.0 mg protein, using liquid nitrogen and
thereafter lyophilizing the preparation for 1 hr with the aid of a Thermovac Freeze Dryer.
Lyophilized membranes were suspended in 0.4M Tris-HCl (pH 7.5) or extracted with organic
solvents. In order to extract neutral lipids, membranes were suspended in 6.0 ml of hexane and
incubated for 15 min at 4º C. The suspension was subjected to centrifugation at 12,000 g for 20 min and the pellet was extracted again with hexane. The pellet was recovered following a
12,000 g centrifugation for 20 min, dried under a stream of nitrogen and suspended in 0.6 ml
0.4M Tris-HCl (pH 7.5). To extract phospholipids, a subsequent 2x extraction of the hexane extracted pellet with 99% aqueous acetone was employed, following the same procedures employed for hexane extraction followed by drying of the 12,000-g pellet under nitrogen and suspension in 0.6 ml 0.4M Tris-HCl (pH 7.5).
The effects of phospholipid addition to lipid depleted membranes were investigated using techniques essentially as described by Rydström (1977). A partially lipid-depleted fraction, designated the 30-55 fraction, was obtained following precipitation of protein from isolated membranes using between 30% and 55% ammonium sulfate saturation. Mitochondrial membranes equivalent to 18.0 mg protein were suspended with 0.4% lysophosphatidylcholine in a 9.0 ml volume and incubated at 4 C for 15 min. After incubation, the preparation was subjected to centrifugation at 100,000 g for 30 min. The supernatant was supplemented with 16 0.5% sodium cholate and incubated for 10 min at 4º C. Ammonium sulfate equivalent to 30%
saturation was added to the preparation and samples were incubated for 10 min at 4º C followed
by centrifugation for 30 min at 100,000 g. The resulting supernatant was brought to 55%
saturation with ammonium sulfate and incubated for 10 min at 4 C. Preparations were subjected
to centrifugation at 100,000 g for 30 min. The resulting pellet was suspended in 0.4M Tris-HCl
(pH 7.5) and assessed for activity.
Addition of phospholipids to the partially lipid-depleted 30-55 fraction was carried out as
described by Anderson and Fisher (1978) and Fioravanti and Kim (1983). 0.5 µmol of the
indicated phospholipid in chloroform was placed in a glass tube, dried under a stream of nitrogen
and suspended in 0.25 ml of 0.4M Tris-HCl pH 7.5 containing 0.10% sodium cholate.
Phospholipids were solublized by sonication for 5 min under nitrogen using a Laboratory
Supplies Co. bath-type sonicator. An aliquot, equivalent to 0.63 mg protein, was taken from the
30-55 fraction and added to the sonicated lipid preparation, for a final volume of 0.5 ml. This sample was sonicated further for 1 min at 4º C. Samples of this preparation were assayed for
activity.
NADH oxidase and NADH-coupled fumarate reductase activities were assessed
spectrophotometrically by measuring reduced pyridine dinucleotide utilization at 340 nm as
described by Fioravanti (1981). Aside from protein and 5.0 mg BSA, the NADH oxidase assay
contained the following in a 1.0 ml reading volume: 0.1 mmol Tris-HCl (pH 6.5), 0.24 µmol
NADH. Aside from protein and 5.0 mg BSA, the fumarate reductase assay contained the
following in a 1.0 ml reading volume: 0.1 mmol Tris-HCl (pH 7.5), 0.6 µmol fumarate and 0.24
µmol NADH. NADPH→NAD transhydrogenase and the NADH→NAD transhydrogenation
activity were assessed by measuring the accumulation of reduced acytylpyridine adenine
dinucleotide (AcPyAD) at 375 nm. In addition to protein and 5.0 mg BSA, the NADPH→NAD 17 transhydrogenase assay contained the following in a 1.0 ml reading volume: 0.1 mmol Tris-HCl
(pH 6.5), 0.24 µmol NADPH, and 0.6 µmol AcPyAD. Aside from protein and 5.0 mg BSA, the
NADH→NAD transhydrogenation assay contained the following in a 1.0 ml reading volume: 0.1 mmol phosphate (pH 7.5), 0.24 µmol NADH, and 0.6 µmol AcPyAD.
NAD(P)H cytochrome c reductase activity was assessed by measuring the accumulation of reduced cytochrome c at 550 nm as described by Mahler (1955) and McKelevy and Fioravanti
(1985). Aside from protein and 5.0 mg BSA, The NAD(P)H cytochrome c reductase assay contained the following in a 1.0 ml reading volume: 0.1 mmol Tris-HCl (pH 6.5), 0.24 µmol
NADPH, and 0.3 mg cytochrome c. NAD(P)H and succinate dehydrogenase activities were assessed by measuring the reduction of ferricyanide at 410nm as described by Fioravanti (1981) and Walker and Fioravanti (1995). Aside from protein and 5.0 mg BSA, both the NAD(P)H and succinate dehydrogenase assays contained the following in a 1.0 ml reading volume: 0.1 mmol
Tris-HCl (pH 7.0 for NAD(P)H dehydrogenase, pH 7.5 for succinate dehydrogenase), 0.6 µmol ferricyanide, and 0.24 µmol NADH or succinate, respectively. Lipoamide dehydrogenase was assayed as described by Komuniecki and Saz (1979) by measuring the lipoamide-dependant oxidation of NADH. Aside from protein and 5.0 mg BSA, the lipoamide dehydrogenase assay contained the following in a 1.0 ml reading volume: 0.1 mmol phosphate (pH 7.5), 0.24 µmol
NADH, 2.0 µmol EDTA (pH 7.5), 0.1 µmol nicotinamide adenine dinucleotide (NAD), and 2.5
µmol lipoamide in 95% ethanol.
The protein content of mitochondrial membranes was assessed by the method of
Bradford (1976) using BSA as the standard. All assays were carried out at 25° C with the aid of a Shimadzu UV-1700 spectrophotometer.
Reduced pyridine nucleotides, acetylpyridine NAD, NAD, cytochrome c (horse heart), lipoamide, ferricyanide, fumaric acid, succinic acid, EDTA, phospholipases A1 (T. lanuginosus), 18 A2 (bee venom), C (Type X, C. perfringes) and D (Type II, peanut), L-α-
lysophosphatidylcholine, and sodium cholate were purchased from Sigma-Aldrich Chemical
Company, St. Louis, MO. Crystalline BSA was purchased from ICN Biochemicals, Inc, Irvine,
CA. Acetone was obtained from Pharmco-Aaper, Brookefield, CT. Hexane, sucrose and sodium
phosphate, monobasic were purchased from Fisher Scientific, Pittsburgh, PA. Bio-Rad protein
assay reagent was purchased from Bio-Rad Laboratories, Richmond, CA. Sodium phosphate,
dibasic and Tris were purchased from EMD Chemicals, Inc., Gibbstown, NJ and Amresco,
Solon, OH, respectively.
Statistical analyses through one-way ANOVA and Tukey’s Test were accomplished using JMP 10 software. Significance was determined when p < 0.05. 19 RESULTS
Treatment of H. diminuta mitochondrial membranes with phospholipases.
Isolated mitochondrial membranes from adult Hymenolepis diminuta were subjected to phospholipase treatments using phospholipases A1, A2, C, and D to evaluate the phospholipid
requirement of a number of mitochondrial pyridine nucleotide-utilizing systems and succinate
dehydrogenase. The figures dealing with phospholipase treatments are based on comparisons
made between activities of untreated samples (incubated with all components except
phospholipase), samples in which EDTA was added with phospholipases to inhibit the Ca++ -
dependent phospholipase activity, and treated samples (incubated with all components including
phospholipase). As presented in Figures 5 and 6, small but statistically significant decreases were
noted in NAD(P)H cytochrome c reductase and NADH dehydrogenase activities when membranes were incubated with phospholipase A1 plus EDTA as compared to untreated and
treated samples. A statistically significant decrease in NADPH cytochrome c reductase activity was noted in untreated samples in comparison to the EDTA inhibited and phospholipase A1
treated samples (Fig. 7). The NADH→NAD transhydrogenation exhibited a decreased activity in the EDTA inhibited sample as compared to the untreated preparations, whereas the phospholipase A1 treated sample had a significantly increased activity level relative to the
untreated sample (Fig. 7). There were no significant differences noted with the other enzymes
investigated (viz., NADPH-, succinate- and lipoamide dehydrogenases, NADH oxidase,
fumarate reductase, and NADPH→NAD transhydrogenation) (Figs. 8-12).
Phospholipase A2 treatment had a significant inhibitory effect on a number of the systems
evaluated. As reported previously (Fioravanti and Kim, 1983), H. diminuta NADH oxidase,
fumarate reductase, and NADPH→NAD transhydrogenase activities were significantly (50% or
more) decreased (Fig. 11-14). In comparisons of NADH oxidase and fumarate reductase 20 activities of untreated and EDTA inhibited preparations with treated samples, significant
differences were apparent with phospholipase A2 treatment. Interestingly, the NADPH→NAD
transhydrogenase system appeared more sensitive to incubation with phospholipase A2 than the
other activities. Only in the absence of phospholipase A2 was the activity maintained (Fig. 14).
NADH cytochrome c reductase activity diminished when subjected to phospholipase A2
treatment (Fig. 5). The other systems subjected to phospholipase A2 treatment did not exhibit
significant changes in activity (Figs. 6-10).
The effects of phospholipase C on the cestode systems were like those noted by
Fioravanti and Kim (1983) with respect to diminishment of NADH oxidase and fumarate reductase activities (Figs. 11 and 13). An inhibitory effect of phospholipase C treatment on the
NADPH→NAD transhydrogenase was suggested but was not found to be statistically significant
(Fig. 14). Moreover, both NADH cytochrome c reductase and succinate dehydrogenase
activities exhibited in excess of 50% activity reduction in comparisons of untreated samples and
those inhibited by EDTA with those treated with phospholipase C (Fig. 5). NADPH cytochrome c reductase and NADH dehydrogenase both displayed a statistically significant increase in activity with EDTA inhibited samples, although stimulation was not apparent with the phospholipase C treated sample (Figs. 6, 8).
Phospholipase D treatment of mitochondrial membranes caused a statistically significant reduction of NADPH cytochrome c reductase activity as compared to untreated and EDTA inhibited incubations (Fig 6). Inhibition following phospholipase D treatment was also noted with fumarate reductase and NADPH→NAD transhydrogenase activities as compared to the untreated samples as given in Figures 13 and 14. With EDTA inhibited incubations, apparent increases in NADH→NAD transhydrogenation and NADH dehydrogenase activities were noted, although these increases were not apparent in the treated samples (Figs. 7, 8). 21
Extraction of H.diminuta mitochondrial membranes with organic solvents.
H. diminuta mitochondrial membranes were subjected to lyophilization and subsequent
extraction with hexane or hexane extraction followed by 99% aqueous acetone. The results
obtained are given in Table I. As noted, lyophilization of H. diminuta mitochondrial membranes
had a stimulatory effect on a number of the enzyme systems investigated. Thus, NADH→NAD
transhydrogenation, NADH-, succinate-, and lipoamide dehydrogenase, and NADPH→NAD
transhydrogenase activities were significantly increased. NAD(P)H cytochrome c reductase and
NADH oxidase exhibited no significant change in activities, whereas the fumarate reductase
system exhibited nearly a 50% decrease in activity following lyophilization. When subjected to hexane extraction, the NAD(P)H cytochrome c reductase and fumarate reductase systems showed a marked reduction in activity while the other enzymatic systems investigated were
unaffected (Table I).
Extraction of hexane-treated membranes with 99% aqueous acetone had a diminishing effect on the activities of a number of the systems (Table I). NADPH→NAD transhydrogenase,
NADH oxidase, and succinate dehydrogenase activities were markedly decreased, and the
NADPH→NAD transhydrogenase system lost over 80% of its activity. The remaining systems investigated showed no significant change in activity with acetone treatment. It is of note that the NADPH dehydrogenase activity was unchanged when subjected to any of the treatments
(Table I).
Effect of phospholipid addition to partially lipid-depleted H. diminuta mitochondrial membrane preparations.
H. diminuta mitochondrial membranes were subjected to a 30-55 percent ammonium sulfate fractionation to affect partial phospholipid depletion, and the resulting protein fraction 22 was assessed for activities with and without phospholipid incorporation (Table II). As
established previously (Fioravanti and Kim, 1983), a phospholipid dependency of the
NADPH→NAD transhydrogenase was evidenced by a diminished activity in the partially
phospholipid depleted 30-55 fraction. When phosphatidylcholine was added to the depleted
membranes, activity was increased to nearly double that of untreated membrane preparations
(Table II). NAD(P)H cytochrome c reductase activities exhibited no significant change in
activity when subjected to 30-55 fractionation, nor was there a significant change when
phosphatidylcholine was added to the partially depleted membranes. The NADH→NAD
transhydrogenation, NADH-and lipoamide dehydrogenase systems exhibited a substantial
increase in activity with 30-55 fractionation. The NADH→NAD transhydrogenation and NADH
dehydrogenase doubled in activity while the lipoamide dehydrogenase activity was 150% of
untreated preparations (Table II). Following phosphatidylcholine supplementation,
NADH→NAD transhydrogenation activity was diminished by nearly 25% while the NADH
dehydrogenase had no significant change in activity. Lipoamide dehydrogenase activity of
supplemented samples was nearly double that of the 30-55 fraction level. While the NADPH
dehydrogenase showed no significant change in activity when membranes were subjected to
ammonium sulfate fractionation, this activity was nearly doubled when comparing the untreated
preparation to the preparation supplemented with phosphatidylcholine (Table II).
NADH oxidase showed no significant change in activity when subjected to 30-55 fractionation, and addition of phosphatidylcholine to the preparation caused activity to decrease by around 70% from the untreated level. Succinate dehydrogenase and fumarate reductase both showed diminished levels of activity when phospholipids were partially depleted, with the succinate dehydrogenase activity dropping by nearly half and the fumarate reductase by almost
90%. Addition of phosphatidylcholine to the 30-55 preparation had no significant effect on 23 fumarate reductase activity and appeared to nearly completely eliminate the succinate dehydrogenase activity (Table II). 24 25 26 27 28 29 30 31 32 33 34
35 36 Treatment of Ascaris suum muscle mitochondrial membranes with phospholipases.
A comparative study of the effects of phospholipases A1, A2, C, and D on Ascaris suum
muscle mitochondrial membranes was conducted. When treated with phospholipase A1, the
NADH dehydrogenase and fumarate reductase showed a small but significant increase in activity
(Figs. 18, 23). The NADH→NAD transhydrogenation activity statistically significantly increased in comparison of untreated samples with phospholipase A1 treated samples, as
presented in Figure 17. Conversely, an apparent decrease in the activity of the succinate
dehydrogenase was noted when comparing the untreated samples with samples treated with
phospholipase and phospholipase plus EDTA (Fig. 20). No significant effects of phospholipase
A1 treatments were apparent with respect to membrane-associated NAD(P)H cytochrome c
reductase, NADH→NAD transhydrogenation, NADPH dehydrogenase, NADH oxidase, and
lipoamide dehydrogenase activities (Figs. 16-22).
Ascarid NADH cytochrome c reductase, NADH oxidase and fumarate reductase activities
decreased by at least 50% with phospholipid disruption by phospholipase A2 treatment as
presented in Figures 15, 21, 23. NADH dehydrogenase activity increased when treated with
phospholipase A2 plus EDTA, and an even larger increase was noted when subjected to
phospholipase A2 treatment alone, while the other systems appeared to be insignificantly
impacted (Figs. 16-20, 22).
Phospholipase C treatment of A. suum mitochondrial membranes resulted in diminished
activity (around 50% less) of the NADH cytochrome c reductase, NADH oxidase and fumarate
reductase systems when comparing treated samples to untreated and EDTA inhibited samples
(Figs 15, 21, 23). While a decrease in activity was noted in the NADH cytochrome c reductase,
there was stimulation of activity of the NADPH cytochrome c reductase system when subjected
to phospholipase C treatment as compared to the untreated preparations (Fig. 16). Also, both the 37 NADH→NAD transhydrogenation and the NADPH dehydrogenase showed an apparent stimulation in activity when incubated with phospholipase C plus EDTA, though this stimulation was not noted when incubated with phospholipase C alone (Figs. 17, 19).
NADH cytochrome c reductase activity was stimulated when nematode membranes were incubated with phospholipase D plus EDTA, though this effect was not apparent when incubated with phospholipase D alone (Fig. 15). None of the other systems investigated showed a significant change in activity with respect to phospholipase D (Figs. 16-23).
38
39 40 41 42 43 44 45 46 47 DISCUSSION
Previous research by Fioravanti and Kim (1983) demonstrated a phospholipid
dependence of the H. diminuta mitochondrial, membrane-associated NADPH→NAD
transhydrogenase. This dependency was evidenced via disruption of membrane phospholipids
using phospholipases A2 or C and by extraction of phospholipids with organic solvent. A
phospholipid dependence was further established by phospholipid restoration of
transhydrogenase activity by phosphatidylcholine supplementation of an ammonium sulfate
fraction derived from mitochondrial membranes. A phospholipid dependence of the H. diminuta
mitochondrial membrane-associated, NADH-utilizing fumarate reductase and lesser NADH
oxidase also was demonstrated via disruption of phospholipids using phospholipases A2 or C as
well as by extraction of phospholipids using organic solvents.
Evidence of a phospholipid dependence of mitochondrial NADPH→NAD
transhydrogenase was established by Rydström et. al.(1976) for bovine heart and by Vandock et. al. (2011) for the tobacco hornworm, M. sexta. With both systems, the transhydrogenase
displayed a diminished activity both when phospholipids were disrupted by phospholipases as
well as when phospholipids were extracted with organic solvents. Activity of the
transhydrogenase was best restored through addition of cardiolipin or a mixture of phospholipids
in the bovine system and by phosphatidylcholine in M. sexta.
In the present study, phospholipid dependence of the H. diminuta mitochondrial
NADPH→NAD transhydrogenase, fumarate reductase and NADH oxidase was supported in that a clear decrease in these activities was apparent with phospholipase A2 treatment as well as by
extraction of phospholipids with aqueous acetone. The transhydrogenase also was restored when
a partially phospholipid-depleted preparation was supplemented with phosphatidylcholine.
Though both the fumarate reductase and NADH oxidase displayed a disruption of activity when 48 treated with phospholipase C, the NADPH→NAD transhydrogenase did not display a
statistically significant decrease in activity, although activities measured in the phospholipase C
treated preparations were lower than those found in both the EDTA control and untreated
preparations.
The Fioravanti and Kim (1983) study was expanded to include the effects of ammonium sulfate fractionation and subsequent phosphatidylcholine supplementation on the cestode fumarate reductase and NADH oxidase activities. The study was also expanded to examine the effects of phospholipase A1, A2, C and D treatments on NAD(P)H cytochrome c reductase,
NADH→NAD transhydrogenation, NAD(P)H dehydrogenase, succinate dehydrogenase, and
lipoamide dehydrogenase and succinate dehydrogenase activities. Additionally, phospholipid
extraction using organic solvents, and restoration of activities of a 30-55 membrane fraction with
phosphatidylcholine were studied.
With respect to incubation with phospholipase A1 treatments, the NAD(P)H cytochrome c
reductase, NADH→NAD transhydrogenation, and NADH dehydrogenase activities increased
with phospholipase A1 plus EDTA incubation, although this increase was not seen with the
phospholipase A1 treated preparations. The other activities were not significantly affected by
cleavage of the sn1 fatty acid.
Treatments with phospholipase A2, thereby removing the sn2 fatty acid of phospholipids,
caused a significant decrease in activity in a number of the enzyme systems. In accord with
Fioravanti and Kim’s (1983) findings, the NADPH→NAD transhydrogenase, fumarate reductase
and NADH oxidase all had diminished activities. Moreover, the NADH cytochrome c reductase
activity also was diminished significantly with phospholipase A2 treatment.
Phospholipase C treatment yielded similar results to those with phospholipase A2. Thus,
fumarate reductase, NADH oxidase and cytochrome c reductase activities decreased when the 49 polar head group plus phosphate group was cleaved. Treatment with phospholipase C had a decreasing effect on succinate dehydrogenase activity as well. Although the NADPH→NAD transhydrogenase did not show a statistically significant change in activity, it too displayed lower activity with phospholipase C treatment. As was found with phospholipase A1 treatment,
incubation with the combination of EDTA and phospholipase had an altering effect on the
activity of the NADPH cytochrome c reductase and the NADH dehydrogenase. In the case of
phospholipase C, the effect of incubation with EDTA and phospholipase C seemingly decreased
activity, although again the effect was not seen with incubation with only phospholipase.
When the polar head group alone was cleaved by phospholipase D, similar effects as
found with phospholipase A1 and C were noted. The activity of the NADH→NAD
transhydrogenation and the NADH dehydrogenase increased when incubated with EDTA plus
phospholipase, while the effect was not seen when only incubated with phospholipase. An
overall decrease in activity was noted in NADPH cytochrome c reductase activity with
phospholipase D treatment. A decrease in the activities of the NADPH→NAD transhydrogenase
and the fumarate reductase was noted, but this was only relative to the untreated samples and the
phospholipase D treated samples.
Membrane lyophilization markedly stimulated NADH→NAD transhydrogenation,
NADH dehydrogenase, succinate dehydrogenase, and NADPH→NAD transhydrogenase
activities. The NAD(P)H cytochrome c reductase, NADH oxidase, and lipoamide
dehydrogenase activities were not significantly affected, and the fumarate reductase activity
decreased following lyophilization. While these changes following lyophilization are of interest
and warrant investigation, they were not pursued further.
Based on hexane treatment of lyophilized membranes, the cestode NAD(P)H cytochrome
c reductase and fumarate reductase systems required neutral lipids for optimal activity. Indeed, 50 these activities did not show a significant decrease in activity following hexane/acetone
extraction thereby indicating a greater dependence on neutral lipids rather than phospholipids.
The NADH oxidase and NADPH→NAD transhydrogenase were the only systems displaying a
significant activity loss in comparisons of hexane extractions versus hexane/acetone extractions.
Accordingly, these latter systems appear more dependent on phospholipids than the others for
retention of their activities.
H. diminuta mitochondrial membranes were also subjected to ammonium sulfate
fractionation resulting in partially phospholipid-depleted membranes as demonstrated by
Rydström (1977). Interestingly, phospholipid depletion of membranes via ammonium sulfate
fractionation resulted in a stimulatory effect on the NADH→NAD transhydrogenation and
NADH dehydrogenase systems. This treatment had a diminishing effect on the activities of the
succinate dehydrogenase, fumarate reductase, and NADPH→NAD transhydrogenase, coinciding with their previously established phospholipid dependence found with phospholipase treatments.
Upon supplementation of the depleted membranes with phosphatidylcholine, the
NADPH→NAD transhydrogenase activity surpassed the untreated level, thereby supporting the result reported by Fioravanti and Kim (1983). Phosphatidylcholine addition had the opposite effect on the NADH→NAD transhydrogenation, the succinate dehydrogenase, and the NADH oxidase activities which were markedly decreased. The lipoamide dehydrogenase, however, showed a substantial increase in activity. It will be of interest to evaluate the effects of other phospholipids in supplementation of the cestode 30-55 fraction depleted membrane preparations.
The established phospholipid dependency of the NADPH→NAD transhydrogenase , fumarate reductase, and NADH oxidase reported by Fioravanti and Kim (1983) was confirmed.
In addition, a phospholipid dependence was observed with respect to the NADH cytochrome c reductase and succinate dehydrogenase systems. The NAD(P)H cytochrome c reductase 51 exhibited an apparent neutral lipid dependence. The data indicate that the sn2 fatty acid and
phosphate group attached to the glycerol backbone are of greater import in retention of activity
than the sn1 fatty acid chain or polar head group. Of note were the findings that despite the
phospholipid dependency noted with respect to NADH cytochrome c reductase, this activity was
not significantly affected by either 30-55 fractionation or by phospholipid supplementation.
Further evaluations as to why this occurred are required.
Interesting effects of phospholipase plus EDTA incubation in the NADPH cytochrome c
reductase, NADH→NAD transhydrogenation, and NADH dehydrogenase systems were
apparent, causing an increase in activity in some preparations while decreasing activity in others.
Though the cause of these effects is unclear, it is noteworthy that this effect is seen with both
NADH→NAD transhydrogenation and the NADH dehydrogenase activities. Previous research
suggested that the NADH→NAD transhydrogenation may be due to a component of the H.
diminuta NADH dehydrogenase as well as the lipoamide dehydrogenase systems (Walker and
Fioravanti, 1995; Walker et. al., 1997) and these finding may contribute to what was noted in the
present study.
Comparative studies were done on the effects of phospholipases A1, A2, C, and D on the
corresponding mitochondrial, membrane-associated activities of the intestinal nematode A. suum.
When treated with phospholipase A1, A. suum NADH dehydrogenase and fumarate reductase
exhibited a small, but statistically significant, increase in activity. In agreement with the data
derived from H. diminuta, the cleavage of the sn2 fatty acid of phospholipids by phospholipase
A2 caused a decrease in activity of the NADH cytochrome c reductase, NADH oxidase and the
fumarate reductase systems. There also was a significant increase in NADH dehydrogenase
activity, as compared to untreated samples, when incubated with the phospholipase as well as when incubated with phospholipase plus EDTA. 52 Further evidence of phospholipid dependence was evident with phospholipase C
treatment. As observed with H. diminuta, the NADH cytochrome c reductase, NADH oxidase
and fumarate reductase activities were diminished by the cleavage of the polar head plus
phosphate group. In further similarity with H. diminuta when treated with phospholipase C, an
apparent stimulation was noted with the NADH→NAD transhydrogenation and NADH
dehydrogenase when incubated with EDTA plus phospholipase, but not with phospholipase
alone.
Seemingly, the cleavage of the polar group from the phospholipid by the action
phospholipase D had no significant effect on the ascarid systems investigated with the exception
that the NADH cytochrome c reductase showed a slight stimulation in activity when incubated
with phospholipase D plus EDTA. No effect was seen with phospholipase incubation alone.
The comparative study with H. diminuta and A. suum revealed for the most part similar
effects of phospholipases on the two systems. The NADH cytochrome c, NADH oxidase and
fumarate reductase systems all displayed similar susceptibly to disruption with phospholipases
A2 and C. It is noteworthy that A. suum mitochondrial succinate dehydrogenase seemed more resistant to phospholipid disruption than the corresponding H. diminuta enzyme. Further investigation using organic solvent extraction and lipid depletion would be of interest.
Investigation into the phospholipid used in reconstitution of lipid depleted membranes will also be of interest, as phosphatidylcholine had a limited effect in restoring the activity of any system except the NADPH→NAD transhydrogenase in H. diminuta. Whether or not phospholipids in the helminth systems play a role in the conformational organization of mitochondrial, membrane-associated proteins, as was noted for mammalian glucoronyltransferase (Singh,
1982), remains to be evaluated. 53 The data presented here expanded on the prior study with H. diminuta (Fioravanti and
Kim, 1983) and extended to an evaluation of the effects of phospholipases A1, A2, C, and D on the A. suum systems. To my knowledge, this study and that of Fioravanti and Kim (1983) are the only studies demonstrating a need for phospholipids by a number of mitochondrial membrane- associated enzyme systems in the parasitic helminths. Aside from some indications in mammalian systems (Rydström et. al., 1976) and more recently in the tobacco hornworm, M. sexta (Vandock et. al., 2011) little data are available as to the phospholipid needs of mitochondrial enzyme systems.
The NADPH→NAD transhydrogenase was found to be particularly sensitive to phospholipase A2 treatments. With a titration of phospholipase A2 content, a clear effect of the phospholipase was observed as compared to phospholipase plus EDTA (Table III). These findings suggest a difference in the phospholipase A2 preparation used here versus that used previously from the same supplier in as much as Fioravanti and Kim (1983) and Vandock et. al.,
(2011) apparently did not observe this greater sensitivity.
A few caveats with respect to the present study are worthy of mention. Contaminations in phospholipase preparations are possible; the phospholipases were isolated from varying sources and were presumed to be purified to the extent noted by the suppliers. Additionally, in the process of isolating helminth mitochondrial membranes by differential centrifugation, the 54 occurrence of lysosomes/peroxisomes is possible in the membrane preparations. Sonication used
to disrupt mitochondria for membrane isolation results inverted membrane vesicles, which may vary from preparation to preparation. However, despite these caveats, prior studies support the use of the methods employed in the present study as sufficient to assess the membrane-associated activities that were investigated.
In the present study, not only were new systems identified that required neutral and/or phospholipids for optimal activity, but enzyme activities were identified that had seemingly little phospholipid dependence and/or neutral lipid dependence. A framework for further investigation into phospholipid/neutral lipid necessity of these and other enzyme systems was established. Certainly isolation and determination of phospholipid content of mitochondrial membranes and further studies as to which phospholipids, or combinations of phospholipids, best restores activities when reincorporated into phospholipid and neutral lipid-depleted membranes would be of value.
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59 APPENDIX I: INSTITUTIONAL ANIMAL CARE AND USE COMMITTEE (IACUC)
APPROVAL INFORMATION
Protocol ID: 10-011 “Anaerobic Energy Metabolism of Parasitic Helminths” approved July 22,
2010, approved and renewed May 5, 2011.