Catalysis of Mitochondrial NADH:NAD

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Catalysis of Mitochondrial NADH→ NAD+ Transhydrogenation in Adult Ascaris suum (Nematoda)

Andrew Holowiecki

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 2009

Committee:

Carmen F. Fioravanti, Advisor

Daniel M. Pavuk

Jill Zeilstra-Ryalls

Andrew Holowiecki

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ABSTRACT

Carmen F. Fioravanti, Advisor

Adult Ascaris suum inhabits the small intestine of its swine host where oxygen tension is

low. Despite the lack of oxygen, A. suum generates mitochondrial ATP by an NADH-requiring,

inner membrane-associated (IM), electron transport-coupled fumarate reductase, producing succinate. Compelling data suggests that the “malic” enzyme resides in the mitochondrial intermembrane space (IMS), forming the NADH required for anaerobic phosphorylation. Thus, the transfer of reducing power from IMS NADH across the IM to matrix NAD+ would be needed

to form the NADH required for anaerobic ATP generation. An IM-associated NADH → NAD+

transhydrogenation reaction has been implicated in this transfer and is thought to be a catalytic

activity of lipoamide dehydrogenase in ascarid mitochondria. The purpose of this study was to ascertain whether the NADH → NAD+ transhydrogenation reaction in adult A. suum results from

more than one catalytic activity, viz., lipoamide dehydrogenase and NADH dehydrogenase.

Studies of the mitochondrial NADH → NAD+ transhydrogenation reaction, lipoamide

dehydrogenase, and NADH dehydrogenase were performed using disrupted adult A. suum mitochondria as the source of enzymes. Based on studies evaluating the effects of pH on the ascarid activities, and the thermal labilities of these reactions, it appears that the lipoamide dehydrogenase and NADH dehydrogenase catalyze an NADH → NAD+ transhydrogenation

reaction in adult A. suum mitochondria. These findings were supported further by

intramitochondrial localizations of the three activities as well as the effects of inhibitors on these

systems.

In light of these findings, it is concluded that the NADH → NAD+ transhydrogenation reaction in adult A. suum is the result of lipoamide dehydrogenase and NADH dehydrogenase systems. Presumably, these studies will aid in the ultimate development of specific chemotherapeutic strategies for development of anthelmintics.

I would like to dedicate this thesis to my parents, Stanley and Katharina Holowiecki for encouraging me to go to school, and to not quit. You have succeeded in giving me all the opportunities that you never had. Thank you.

I would also like to dedicate this thesis to Henry and Becky Beiro for helping me to keep things in perspective. A great man once said “all models are wrong, some models are useful.”

The model you have created for life is useful. Thank you.

Finally, and most importantly, this work is dedicated to my wife, Kristy. This achievement would have been impossible without you. Your love, support, tolerance, patience, respect, and understanding have allowed me to pursue my dreams. I love you, and I need you.

ACKNOWLEDGMENTS

First, I would like to thank my advisor, Dr. Fioravanti, for his guidance and patience.

Your ability to teach and inspire has undoubtedly helped me grow as a student, teacher, and person. You have set a great example of how to be a true professional, and a leader. You never taught me through intimidation, and I never lost my sense of dignity. Thank you.

Additionally, I would like to thank the other members of the Fioravanti Lab, viz., Kurt

Vandock, and Chris Drummond. Your assistance in helping me to learn and improve my lab techniques was essential to my success. Gratitude is expressed to Jeff Meyers at J.H. Routh

Packing for allowing us to collect Ascaris suum.

I would also like to thank my other committee members, Dr. Dan Pavuk and Dr. Jill

Zeilstra-Ryalls for their support and willingness to contribute to my education.

Finally, I would like to thank Dr. Kathryn Durham of Lorain County Community College.

You made biology interesting, and you taught me how to learn. I would also like to thank Dr.

James Beil of Lorain County Community College. Your patience teaching me as an undergraduate and your willingness to help me as a graduate student has helped me to achieve my goals.

This work was supported in part by research grant AI-15597 from the National Institutes of Health, awarded to Dr. Carmen F. Fioravanti. This research was also supported by a Grant- In-

Aid of Research from the National Academy of Sciences, administered by Sigma Xi, The

Scientific Research Society.

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TABLE OF CONTENTS

Page

INTRODUCTION...... 1

CHAPTER I. A REVIEW OF THE LITERATURE ...... 3

CHAPTER II. MITOCHONDRIAL NADH→NAD+ TRANSHYDROGENATION IN

ADULT ASCARIS SUUM (NEMATODA) ...... 12

Introduction ………………………………………………………………………. . 12

Materials and Methods……………………………………………………………. . 14

Results…………………………………………………………………………….. . 16

Discussion………………………………………………………………………… . 26

CONCLUSIONS AND FUTURE DIRECTIONS……………………………………….... . 31

REFERENCES……………………………………………………………………………. . 33

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LIST OF TABLES

Table Page

1 Mitochondrial localizations of NADH → NAD + transhydrogenation, lipoamide

dehydrogenase, and NADH dehydrogenase activities of A. suum mitochondria…… 23

2 Effects of inhibitors on activities of Ascaris suum mitochondria

assessed under acidic and basic conditions…………………………………………. 25

LIST OF FIGURES

Figure Page

1.1 The life cycle of Ascaris lumbricoides……………………………………………… 4

1.2 Pathway of carbohydrate dissimilation in Ascaris lumbricoides muscle……………. 7

1.3 Pathway describing intramitochondrial malate oxidation and the potential role of

IM-associated NADH→NAD+ transhydrogenation.……………………………… 9

2.1 The effects of pH on NADH→NAD+ transhydrogenation,

NADH dehydrogenase, and lipoamide dehydrogenase activities of adult

Ascaris suum mitochondria………………………………………………………… 17

2.2 Thermal lability profiles of the NADH→NAD+ transhydrogenation,

NADH dehydrogenase, and lipoamide dehydrogenase activities of

adult Ascaris suum mitochondria assessed under acidic conditions……………… .. 20

2.3 Thermal lability profiles of NADH→NAD+ transhydrogenation,

NADH dehydrogenase, and lipoamide dehydrogenase activities

of adult A. suum mitochondria assayed under basic conditions ...... 21

2.4 Proposed roles of lipoamide dehydrogenase (LD) and NADH dehydrogenase (ND)

as NADH→NAD+ transhydrogenation mechanisms in the mitochondrial

energetics of adult Ascaris suum. …………………………………………………... 29

INTRODUCTION

The adult intestinal helminth of swine, Ascaris suum (Nematoda), is predominately

anaerobic (Scheibel and Saz, 1966; Scheibel et al.,1968; Saz and Lescure, 1969) and produces organic end products as a result of carbohydrate catabolism (Saz and Bueding, 1966). Malate is

generated in the cytosol and enters the mitochondria where it serves as the mitochondrial

substrate for a dismutation reaction (Bueding and Saz, 1968; Saz and Lescure, 1969). The

oxidative branch of this reaction produces pyruvate and carbon dioxide, and subsequently yields

reducing power in the form of NADH (Rew and Saz, 1974). The reductive branch of this

dismutation reaction entails the reduction of fumarate to succinate resulting from the electron

transport-dependent, NADH requiring, fumarate reductase (Fioravanti and Saz, 1980). Fumarate reduction is accompanied by the concomitant, site I-coupled, formation of ATP. Reducing power in the form of NADH accumulates in the mitochondrial IMS via the NAD+-linked “malic”

enzyme (Rew and Saz, 1974). Subsequently, reducing power from NADH must cross the IM of

the mitochondria to reduce matrix NAD+ to drive the anaerobic, ATP generating, electron transport system. A membrane- associated NADH → NAD+ transhydrogenation reaction has

been implicated in this transmembrane movement of hydride ions (Fioravanti and Saz, 1976;

Kohler and Saz, 1976). Previous studies have implicated an association between this

transhydrogenation and lipoamide dehydrogenase in adult Ascaris (Komuniecki and Saz, 1979).

Conversely, in the intestinal parasite of rats, adult Hymenolepis diminuta (Cestoda), it has been

reported that the NADH → NAD+ transhydrogenation reaction may be the result of both

lipoamide dehydrogenase and NADH dehydrogenase (Walker and Fioravanti, 1995). Chapter 1

of this thesis entails a review of the literature relative to anaerobic energy generation in the

helminths, with particular consideration given to the mitochondrial NADH→ NAD+ 2

transhydrogenation reaction. Chapter 2 presents information addressing the question as to

whether the mitochondrial NADH → NAD+ transhydrogenation reaction of adult A. suum is the sum of a catalytic entity other than an NADH → NAD+ transhydrogenase and/or lipoamide

dehydrogenase system.

CHAPTER I. A REVIEW OF THE LITERATURE

The Biology of Ascaris suum

Ascaris suum, the adult intestinal roundworm (Nematode) of swine, is a member of the phylum Nematoda, the class Secernentea, the order Ascaridida, and the family Ascaridae

(Bogitsh et al., 2005). Adult female ascarids are much larger than their male counterparts and male ascarids are easily distinguished from females by their ventrally curved tail. Adult A. suum is visually indistinguishable from the corresponding intestinal roundworm of humans, Ascaris lumbricoides, but differs from the former in that its eggs do not readily infect humans (Dailey,

1996). A. lumbricoides, the most common nematode parasitizing humans, is thought to infect over 1 billion people worldwide (Bogitsh et al., 2005).

While the host specificity of A. suum and A. lumbricoides differs, their life cycles are alike. Both nematodes have a monoxenous life cycle, i.e., they are highly specific to only one host. Both A. suum and A. lumbricoides have four juvenile stages and undergo four molts in their development from egg to adult. Whereas the first two molts occur within the egg, the third and fourth molts occur within the host. The lifecycle of A. lumbricoides is presented in Figure 1.

1. Undeveloped eggs are passed in host feces and embryonate during incubation in an aerobic environment. Upon ingestion by a suitable host, the eggs hatch within the stomach, thereby releasing larvae which penetrate the intestines. From the intestines the larvae travel, via the venous blood stream to the liver and subsequently migrate to the lungs where they undergo a third molt. Thereafter the larvae move up the trachea where they are swallowed for a second time. The fourth and final molt occurs in the small intestine producing the adult egg-laying parasite. 4

FIGURE 1.1: The life cycle of Ascaris lumbricoides. Adult worms (1) live in the lumen of the small intestine. Female worms produce approximately 200,000 eggs per day, which are passed with the feces (2). Fertile eggs embryonate and become infective after approximately 18 days

(3). Upon ingesting infective eggs (4), the larvae hatch (5), invade the intestinal mucosa, and are carried to the liver by the venous blood stream, and then, via systemic circulation, to the lungs

(6). Further larval development occurs within the lungs (lasting 10 to 14 days) after which the larvae penetrate the alveolar walls, ascend the bronchial tree to the throat, and are swallowed (7).

Inside the small intestine, the worms further develop into the adult egg laying parasite (1).

The ability of infectious parasites like A. suum and A. lumbricoides to not only survive,

but to thrive in a variety of differing environments, is evident by their success in nature.

Certainly their ability to go from one environment to another, regardless of changes in O2

tension, host immune responses, and other environmental changes present difficulties in terms of

creating specific chemotherapies (Barret, 1981). Nonetheless, biochemical studies of these ascarids, and other helminths, have contributed substantially to our understanding of aerobic and anaerobic respiration. For example, cytochromes, which have a key role in oxidative phosphorylation, were first observed by the parasitologist David Keilin when studying the intestinal horse parasite Gastrophilus intestinalis (Keilin, 1966; Slater, 2003). Additionally,

Francesco Redi used A. lumbricoides to disprove the theory of spontaneous generation in the 17th

century (Read, 1972; Cox, 2002). A. lumbricoides also served as the first organism in which

physiologically functional anaerobic mitochondria were described (Bueding, 1949; Saz and

Weil, 1962).

Ascaris suum Metabolism

Adult A. suum is essentially anaerobic in terms of its energetics (Scheibel and Saz, 1966;

Scheibel et al., 1968; Saz and Lescure, 1969) and produces succinate in addition to a variety of

other fatty acids derived from succinate (Saz and Bueding, 1966). Although energy generation in

adult A. suum is predominantly anaerobic, possibilities for the role of O2 have been suggested.

One such report implicates that cuticle formation in A. suum is dependent on the O2-utilizing enzyme proline hydroxylase (Fujimoto and Prockop, 1969; Cain and Fairbairn, 1971). The role of O2 in other helminths has been questioned and it appears that O2 may play a role separate

from energy generation, such as egg development in Schistosoma mansoni (Schiller et al., 1975). 6

The adult intestinal cestode, Hymenolepis diminuta, also generates energy anaerobically

(Scheibel and Saz, 1966) and produces succinate, acetate and lactate as end products of glucose

catabolism, with succinate being the main end product (Fairbairn et al., 1961). Both A. suum and

H. diminuta lack a fully functioning tricarboxylic acid cycle (Kmetic and Bueding, 1961; Ward

and Fairbairn, 1970) and serve as models for anaerobic helminth energetics. Carbohydrate

dissimilation in adult Ascaris suum muscle is summarized in Figure 1.2. Malate is generated in

the cytosol and enters the mitochondria where it serves as the substrate for a dismutation reaction

(Bueding and Saz, 1968; Saz and Lescure, 1969). In contrast to reported enzyme distribution

studies in mammalian systems, ascarid fumarase and the NAD+ -linked "malic" enzyme are

essentially located within the IMS (Rew and Saz, 1974), thus suggesting that NADH, pyruvate,

and fumarate are formed in the IMS (Rew and Saz, 1974). Glycolysis occurs within the cytosol.

Glucose is broken down to phosphoenolpyruvate (PEP) (Saz, 1981), and in the absence of

significant amounts of pyruvate kinase (PK) in the cytoplasm, CO2 is fixed to PEP producing

oxalacetate (OAA) (Saz and Lescure, 1969). Cytoplasmic NAD+ is formed as a result of the

reduction of OAA to malate by malate dehydrogenase and cytoplasmic NADH (Saz, 1981).

Cytoplasmic malate enters into the mitochondria and serves as the mitochondrial substrate where

it undergoes a dismutation reaction (Kohler and Saz, 1976). In the oxidative branch of the

reaction, the NAD + -linked “malic” enzyme catalyzes the oxidation of malate to pyruvate and

CO2 thus regenerating reducing power in the form of NADH within the mitochondria (Rew and

Saz, 1974). The reductive branch of this dismutation reaction entails the reduction of fumarate to

succinate resulting from the electron transport-dependent, NADH requiring, fumarate reductase

(Fioravanti and Saz, 1980). Fumarate reduction is accompanied by the concomitant, site I coupled, formation of ATP. 7

Figure 1.2: Pathway of carbohydrate dissimilation in Ascaris lumbricoides muscle (after Rew and Saz, 1974). Malate, derived from glycolytic activity and CO2 fixation in the cytosol, enters the mitochondrion. Within the mitochondrion, malate undergoes a dismutation reaction. The oxidative branch of this reaction, as catalyzed by the “malic” enzyme (1.1.1.39 L-malate:NAD oxidoreductase [decarboxylating]), results in the formation of pyruvate and CO2.The reductive branch of this dismutation reaction entails malate oxidation by fumarase with subsequent fumarate reduction by the NADH-dependent fumarate reductase. The resulting reduction results in electron transport-dependent ATP generation.

In order for the dismutation reaction to be complete, the fumarate reductase reaction must occur, which presumably happens on the matrix side of the mitochondrial membrane. Reducing power, in the form of NADH, is apparently accumulated in the mitochondrial IMS by the action of the NAD+-linked “malic” enzyme (Rew and Saz, 1974). Although NADH is required to catalyze the fumarate reductase reaction, due to its apparent origin within the IMS, a mechanism is needed to describe the means of reducing power translocation. A membrane- associated

NADH → NAD+ transhydrogenation reaction has been implicated in this transmembrane movement of hydride ions (Kohler and Saz, 1976). Figure 1.3 depicts the proposed transmembrane transhydrogenation.

Localization studies by Rew and Saz are in contrast to Kohler (1977), and have shown fumarase to be located within the IMS rather than the matrix in Ascaris (1974). Kohler (1977) has indicated that fumarate is unable to penetrate the mitochondrial IM. This presents a dilemma, as fumarate serves as the final electron acceptor in Ascaris (Kmetec and Bueding, 1961; Saz and

Lescure, 1969; and Seidman and Entner, 1961) and other invertebrates (Saz and Bueding, 1966;

Hochachka and Mustafa, 1972). Thus, the reduction of fumarate to succinate would be expected to occur on the matrix side of the inner mitochondrial membrane. The discrepancy between the proposed location(s) of fumarase in Ascaris notwithstanding, further characterization of the proposed NADH→ NAD+ transhydrogenase was initiated.

Figure 1.3: Pathway describing intramitochondrial malate oxidation and the potential role of IM- associated NADH→NAD+ transhydrogenation (after Kohler and Saz, 1976). Abbreviations used are as follows: OM, outer membrane; IM, inner membrane. Fumarate reduction is indicated as a reaction occurring on the matrix surface of the IM.

The pyruvate dehydrogenase complex (PDH) catalyzes the oxidative decarboxylation of pyruvate to acetyl-CoA, CO2, and concomitant NADH accumulation (Matuda and Saheki, 1985;

Behal et al., 1993). This series of reactions is catalyzed by a 3 component enzyme: pyruvate dehydrogenase (E1), EC 1.2.4.1; dihydrolipoamide transacetylase (E2), EC 2.3.1.12; and dihydrolipoamide dehydrogenase (E3), EC 1.8.1.4. (lipoamide dehydrogenase). 10

As noted, in adult A. suum, NADH is generated within the IMS and hydride ions are

thought to cross the mitochondrial IM in order to begin the fumarate reduction reaction

(Komuniecki and Saz, 1979). Studies by Kohler and Saz (1976), and Fioravanti and Saz (1976) implicate this transfer of reducing power from the IMS into the matrix by the membrane bound

NADH → NAD+ transhydrogenase. This NADH → NAD+ transhydrogenase activity has been

reported to be primarily associated with the lipoyl or lipoamide dehydrogenase in A. suum

(Kommuniecki and Saz, 1979). Isolation of lipoamide dehydrogenase from pig heart has

revealed the presence of two differing forms of this enzyme, viz., a soluble and a membrane

bound form (Sakurai et al., 1970); however, studies with the ascarid system indicate that both

forms of the lipoamide dehydrogenase are identical (Komuniecki and Saz, 1979).

In the adult cestode Hymenolepis diminuta, a mitochondrial transhydrogenation reaction

between NADPH and NAD+ and between NADH and NAD+ occurs (Saz et al. 1972; Fioravanti

and Saz, 1976). The reversible NADPH → NAD+ reaction results from an IM-associated pyridine nucleotide transhydrogenase (Fioravanti and Saz, 1976; Fioravanti, 1981; Fioravanti and Kim, 1983; McKelvey and Fioravanti, 1985). The NADH → NAD+ transhydrogenation

reaction has been found to be associated with both lipoamide dehydrogenase, and possibly

NADH dehydrogenase (Walker and Fioravanti, 1995). The mitochondrial NADH dehydrogenase

is the flavin-containing first component of the IM-associated anaerobic electron transport system

as well as a component of the outer membrane rotenone-insensitive NADH cytochrome c

reductase (as reviewed by Fioravanti and Vandock, 2009). However, the NADH → NAD+

transhydrogenation in H. diminuta appear to originate from a source(s) other than the NADPH

→ NAD+ transhydrogenase (Fioravanti and Saz, 1976). 11

Purpose of Study

The purpose of the present study is to determine if lipoamide dehydrogenase is the sole enzymatic entity catalyzing NADH → NAD+ transhydrogenation in A. suum, or if another

system, viz., NADH dehydrogenase contributes to this activity? Published reports have

suggested an association of NADH → NAD+ transhydrogenase activity with lipoamide

dehydrogenase, and NADH dehydrogenase in adult H. diminuta (Walker and Fioravanti, 1995).

In mammalian systems, it has been proposed that the NADH dehydrogenase contains two closely

related active sites (Hatafi and Galante, 1977). One site would accommodate the

dehydrogenation of NAD(P)H while the other site would allow for a second nucleotide to bind

for transhydrogenation (Hatafi and Galante, 1977). This study was undertaken to assess the

possibility that adult ascarid mitochondrial NADH→NAD+ transhydrogenation activity is the

result of a catalytic activity or activities in addition to that of the lipoamide dehydrogenase.

Using disrupted Ascaris mitochondria as the source of the enzymes, three NADH utilizing activities were evaluated as indicated by the following reactions and designations:

1. NADH + NAD+ → NADH (NADH→NAD+ transhydrogenation)

2. NADH + H+ + ferricyanide → NAD+ + ferrocyanide (NADH dehydrogenase)

3. Lipoamide + NADH + H+→ Dihydrolipoamide + NAD+ (lipoamide dehydrogenase).

These evaluations consisted of comparative thermal lability studies, pH optima, insoluble

and soluble mitochondrial localization, and the effects of some inhibitors. The results of this

study are presented here. 12

CHAPTER II. MITOCHONDRIAL NADH → NAD+ TRANSHYDROGENATION IN

ADULT ASCARIS SUUM (NEMATODA)

Introduction

As an adult, Ascaris suum, the intestinal nematode of swine, is essentially anaerobic in terms of its energy generation (Bueding, 1949; Fairbairn, 1957, 1970; Saz and Bueding, 1966;

Saz and Lescure, 1969). Like a number of parasitic helminths (Saz, 1971), this nematode produces succinate (derived from malate) and a variety of fatty acid-end products as the result of carbohydrate catabolism (Saz and Bueding, 1966). Malate, formed in the cytosol by CO2 fixation into glycolytically formed oxalacetate, serves as the anaerobic mitochondrial substrate in A. suum muscle (Bueding and Saz, 1968; Saz and Lescure, 1969). Upon entering the mitochondrion, malate undergoes a dismutation reaction (Bueding and Saz, 1968; Saz and

Lescure, 1969). One arm of the dismutation is catalyzed by the “malic” enzyme resulting in the formation of pyruvate, CO2 and NADH. Malate is also converted to fumarate by fumarase that, in turn, is reduced to succinate by the NADH requiring, electron transport coupled, fumarate reductase (Fioravanti and Saz, 1980).

Studies by Rew and Saz (1974) indicated that both the “malic” enzyme and fumarase are localized in the mitochondrial IMS of A. suum. Thus, upon entering the mitochondrion, the oxidative decarboxylation of malate, by the NAD+-utilizing “malic” enzyme, would result in

NADH formation within this space. Conversion of malate to fumarate also would occur in the

IMS. Since, it is expected that the reduction of fumarate to succinate occurs on the matrix side of the mitochondrial IM, a translocation of reducing equivalents from NADH in the IMS to matrix 13

NAD+, producing the required NADH for electron transport, would be needed inasmuch as

NADH is impermeable to the eukaryotic mitochondrial IM (Lehninger, 1951). Consistent with the considerations of Rew and Saz (1974), Kohler and Saz (1976) presented evidence demonstrating that A. suum mitochondria are capable of translocating reducing equivalents from

NADH in the IMS to matrix NAD+. These data, therefore, support the occurrence of an IM- associated, hydride-translocating mechanism in the adult A. suum system. Both Fioravanti and

Saz (1976) and Kohler and Saz (1976) suggested that this hydride transfer is physiologically

accomplished by an IM-associated NADH→NAD+ transhydrogenation mechanism.

Subsequently, Kohler (1977) reported that fumarate does not penetrate the mitochondrial

IM of A. suum. If fumarate does not penetrate the IM of Ascaris, the IM localization of fumarase

reported by Rew and Saz (1974) versus the data of Kohler (1977) presents somewhat of a

dilemma. Nevertheless, the data concerning the ascarid IM-associated NADH→ NAD+

transhydrogenation (Rew and Saz, 1974; Fioravanti and Saz, 1976; Kohler and Saz, 1976)

remain consistent with the notion that a mechanism(s) exists whereby reducing equivalents

arising in the IM are translocated across the IM to matrix NAD+.

Komuniecki and Saz (1979) presented convincing data that the mitochondrial NADH→

NAD+ transhydrogenation in adult A. suum mitochondria appears to be associated predominately

with the lipoamide dehydrogenase system. For another parasitic helminth, viz., the adult cestode

Hymenolepis diminuta, Walker and Fioravanti (1995) presented evidence that the mitochondrial

NADH→ NAD+ transhydrogenation reaction can result from the catalytic activities of both

lipoamide dehydrogenase and NADH dehydrogenase. Given these latter findings, the potential

association of mitochondrial NADH→NAD+ transhydrogenation activity resulting from a

catalytic activity associated with NADH dehydrogenase as well as lipoamide dehydrogenase 14

became apparent. In the present study, data are presented indicating that an NADH→NAD+

transhydrogenation reaction in adult A. suum mitochondria is catalyzed by not only lipoamide dehydrogenase, but by NADH dehydrogenase as well.

Materials and Methods

Mitochondrial preparation

Adult Ascaris suum were obtained from J.H. Routh Packing in Sandusky, Ohio. Muscle was dissected from female ascarids and mitochondria were extracted essentially as described by

Fioravanti and Saz (1976). Muscle tissue was minced in mitochondrial medium (10 ml/g tissue)

consisting of 250 mM sucrose, 15 mM ethylenediaminetetracetate (EDTA), and 10 mM

Trishydroxymethylaminomethane-HCl (pH 7.5). Thereafter minced muscle was homogenized

using a motorized Potter-Elvehjem homogenizer equipped with a “Teflon” pestle. Removal of

cellular debris was accomplished via centrifugation of the homogenates at 482 x g for 10 min.

Mitochondria were obtained from the resulting supernatant fraction by centrifugation at 9770 x g

for 30 min. Extractions were performed at 4⁰ C. Isolated mitochondria were suspended in

mitochondrial medium (~ 1 ml) and stored frozen.

Enzyme Assays

For enzymatic assays, mitochondria were thawed and diluted (1:5 v/v) with

mitochondrial medium. Thereafter, the organelles were sonically disrupted (3-5 sec bursts with

30 sec cooling intervals) using a Branson sonifier, equipped with a microtip, at a power setting of

20W unless otherwise noted. When needed, mitochondria were separated into insoluble and

soluble fractions by centrifugation of disrupted organelles at 257,320 x g for 60 min. Isolated

fractions were suspended in 0.4 ml of mitochondrial medium and stored frozen. 15

NADH → NAD+ transhydrogenation was assessed by measuring acetylpyridine

(AcPyAD) reduction at 375 nm (Fioravanti, 1981). In addition to enzyme, the 1.0 ml assay volume contained the following in µmoles: NADH, 0.24; AcPyAD, 0.6; and either sodium acetate, sodium phosphate, or Tris-HCl, 100 at the pH indicated.

NADH dehydrogenase was evaluated by measuring the rate of ferricyanide reduction at

410 nm (Walker and Fioravanti, 1995). In addition to enzyme, the 1.0 ml assay volume contained the following in µmoles: NADH, 0.24; potassium ferricyanide, 0.6; and either sodium acetate, sodium phosphate, or Tris-HCl, 100 at the pH indicated.

Lipoamide dehydrogenase was assessed by measuring NADH oxidation at 340 nm as described by Komuniecki and Saz (1979). In addition to enzyme and 3.8 % ethanol the 1.0 ml assay volume contained the following in µmoles: NAD, 0.1; NADH, 0.24; lipoamide, 2.5;

EDTA, 2.0; and either sodium acetate, sodium phosphate, or Tris-HCl, 100 at the pH indicated.

Thermal Lability

Thermal lability profiles were performed using disrupted A. suum mitochondria as the source of enzyme activities. Mitochondria were suspended in approximately 550 - 750 μl of mitochondrial medium and were heated in 5 min. intervals from 25⁰ - 95⁰ C. Termination of heat treatment was accomplished by quenching the samples at 4⁰ C.

Protein determinations were performed according to the method of Bradford (1976) using crystalline bovine serum albumin as the standard. Spectrophotometric assays were performed at

25⁰ C using a Shimadzu UV-1700 series spectrophotometer. 16

Statistical evaluations were performed using JMP 8 software. Significant differences (P <

0.05) in the comparisons of enzyme activities were established by one-way ANOVA.

Differences in the mean were established by a student’s t test.

Results

An evaluation of the effects of pH on the catalysis of the NADH→NAD+

transhydrogenation, NADH dehydrogenase, and lipoamide dehydrogenase activities by disrupted

A. suum mitochondria was performed and the data obtained are presented in Figure 2.1.

Although differences in the activities of the NADH→NAD+ transhydrogenation versus NADH dehydrogenase were noted at pH 9.0 and 8.5, with a decrease in pH from 8.0 to 7.0 both the

NADH→NAD+ transhydrogenation and NADH dehydrogenase activities were essentially

unchanged and virtually equivalent. Lipoamide dehydrogenase activity was relatively low at pH

8.5 and 8.0, but gradually increased, rising to the levels of the other reactions at pH 7.0 (Fig.

2.1). With further medium acidification, lipoamide dehydrogenase activity increased and peaked

at pH 6.0. Thereafter, a decline in this activity was noted with nearly a complete loss of activity

at pH 4.5. Interestingly, between pH 7.0 and pH 6.0, NADH dehydrogenase activity decreased,

whereas NADH→NAD+ transhydrogenation activity essentially fell at a midpoint between

NADH and lipoamide dehydrogenase. All three reactions displayed similar activity levels at pH

5.5. Subsequently, transhydrogenation activity decreased in a fashion similar to the lipoamide

dehydrogenase, but not to the same extent, while NADH dehydrogenase activity increased

immensely, exhibiting a pH 4.5 peak before decreasing at pH 4.0 (Fig. 2.1).

4.5

4.0

3.5

3.0

2.5

Activity 2.0

mol/min/mg protein)mol/min/mg 1.5 μ ( 1.0

0.5

0.0 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 pH

Figure 2.1. The effects of pH on NADH→NAD+ transhydrogenation (NT), NADH

dehydrogenase (ND), and lipoamide dehydrogenase (LD) activities of adult Ascaris suum

mitochondria. Symbols used are:-•- NADH→NAD+ transhydrogenation; -□- NADH

dehydrogenase; -○- lipoamide dehydrogenase. Activity assessments at each pH were performed in triplicate and the means are presented here. A mean of 0.08 mg protein was employed for assays. Values designated for points in insert at given pH are as follows: pH 6.0, NT 0.85 ±

0.058; ND 0.53 ± 0.103; LD 1.20 ± 0.052; pH 6.5, NT 0.76 ± 0.045; ND 0.45 ± 0.108; LD 1.02 ±

0.023. Values in insert are significantly different from each other ± SE; N=3 for each point.

The comparative thermal labilities of the NADH→NAD+ transhydrogenation, NADH

dehydrogenase, and lipoamide dehydrogenase activities of disrupted A. suum mitochondria were

evaluated under acidic and basic assay conditions. Presented in Figure 2.2 are the data obtained

employing acidic conditions of assay. For these assessments the acidic conditions employed

were those yielding apparent maximal activities for the NADH→NAD+ transhydrogenation and lipoamide dehydrogenase, i.e., pH 5.5 and 6.0, respectively. Because NADH dehydrogenase activity proved somewhat difficult for routine measurements, activity was assessed at pH 5.0. At this pH linear measurements of activity were more amenable.

As presented in Figure 2.2, with some variation all three activities were essentially

similar in their lack of thermal lability up to a temperature of 55⁰ C when assessed under acidic

conditions. The most notable lability at 65⁰ C was that of the NADH→NAD+ transhydrogenation

and reflected a decline in activity that was made more apparent at 75⁰ C and then 85⁰ C, with

complete inactivation at 95⁰ C. Conversely a mild increase in the lipoamide dehydrogenase

activity was observed at 65⁰ C with a peak of activity at 75⁰ C. However, NADH dehydrogenase

activity began to decline at 75⁰ C, but differed from the other activities when compared to

controls. A greater degree of NADH dehydrogenase degradation under acidic conditions was

noted at 85⁰ C (Fig. 2.2). The latter degree of degradation was intermediate between that noted

for lipoamide dehydrogenase and the NADH→NAD+ transhydrogenation. As with the

NADH→NAD+ transhydrogenation both the NADH dehydrogenase and lipoamide

dehydrogenase activities were completely inactivated at 95⁰ C (Fig. 2.2).

The comparative thermal labilities of the NADH→NAD+ transhydrogenation, NADH

dehydrogenase, and lipoamide dehydrogenase activities of disrupted A. suum mitochondria also 19

were evaluated under basic conditions of assay, i.e., at pH 8.0 and the data are presented in

Figure 2.3. The NADH dehydrogenase and lipoamide dehydrogenase both displayed a degree of lability that was apparent up to 45⁰ C. Thereafter, a dramatic increase in lipoamide dehydrogenase activity was noted with a peak occurring at 65⁰ C. In contrast, NADH

dehydrogenase activity markedly declined at 55⁰ C, and continued with even a greater activity

loss from 65⁰ C - 85⁰ C before an almost complete loss of activity at 95⁰ C (Fig. 2.3). Whereas

the NADH→NAD+ transhydrogenation reaction displayed essentially no lability up to 45⁰ C,

this activity declined between 55⁰ C and 75⁰ C in a fashion that was intermediate between

lipoamide dehydrogenase and NADH dehydrogenase activities. Thereafter, the

transhydrogenation activity declined similarly to the decline noted for lipoamide dehydrogenase

before a complete loss of activity at 95⁰ C (Fig. 2.3).

Figure 2.2. Thermal lability profiles of the NADH→NAD+ transhydrogenation, NADH

dehydrogenase, and lipoamide dehydrogenase activities of adult Ascaris suum mitochondria

assessed under acidic conditions. Symbols used and the pH of the assay medium were:

-•- NADH→NAD+ transhydrogenation, pH 5.5; -□- NADH dehydrogenase, pH 5.0;

-○- lipoamide dehydrogenase, pH 6.0. Control (100%) activities in μmol/min/mg protein were:

NADH→NAD+ transhydrogenation, 1.5; NADH dehydrogenase, 2.9; and lipoamide

dehydrogenase, 1.4. Activity assessments at each temperature were performed in triplicate and

the means are presented here. A mean of 0.09 mg protein was employed for assays. Values designated for lipoamide dehydrogenase and NADH dehydrogenase differ significantly at 65⁰ C,

75⁰ C, and 85⁰C. Values at these temperatures are significantly different from each other ± SE;

N=4 for lipoamide dehydrogenase, N= 3 for NADH→NAD + transhydrogenation and NADH

dehydrogenase. 21

Figure 2.3. Thermal lability profiles of NADH→NAD+ transhydrogenation, NADH

dehydrogenase, and lipoamide dehydrogenase activities of adult A. suum mitochondria assayed

under basic conditions, i.e., pH 8.0. Symbols used were: -•- NADH→NAD+ transhydrogenation;

-□- NADH dehydrogenase; -○- lipoamide dehydrogenase. Control (100%) activities in

μmol/min/mg protein were: -•- NADH→NAD+ transhydrogenation, 0.81; -□- NADH dehydrogenase, 0.77; -○- lipoamide dehydrogenase, 0.35. For all assessments, N= 2-3 and the means are presented here. A mean of 0.12 mg protein was employed for assays.

Adult A. suum mitochondria were separated into insoluble (membranes) and soluble

fractions via differential centrifugation. Thereafter, assessments of the NADH→NAD+

transhydrogenation, lipoamide dehydrogenase, and NADH dehydrogenase activities were made

using these fractions. The data obtained are presented in Table 1. All three activities were

assessed under acidic and basic conditions. As given in Table 1, all three activities displayed a

predominate insoluble association regardless of the pH of assessment. Under either condition

tested (i.e., pH 5.5 or 7.5), the NADH→NAD+ transhydrogenation as well as the NADH

dehydrogenase activities were clearly more pronounced in the insoluble fraction, being in excess

of 70% of the recovered activities. However, the distribution of lipoamide dehydrogenase

activity differed from the distributions of the other two activities when assessments were

performed either under acidic or basic conditions with insoluble and soluble localizations being

64% and 36%, respectively under acidic conditions, and 67% and 33%, respectively under basic conditions (Table 1).

Table 1. Mitochondrial localizations of NADH → NAD + transhydrogenation, lipoamide dehydrogenase, and NADH dehydrogenase activities of A. suum mitochondria.

Total Units* % Recovered Activity

Reaction pH Insoluble fraction Soluble fraction Insoluble fraction Soluble fraction

NADH → NAD+ 5.5 1.9 ± 0.12† 0.6 ± 0.01 76 % 24 % transhydrogenation

7.5 0.9 ± 0.05 0.3 ± 0.003 75 % 25 %

Lipoamide 5.5 1.6 ± 0.07 0.9 ± 0.06 64% 36% dehydrogenase

7.5 0.4 ± 0.016 0.2 ± 0.004 67% 33%

NADH 5.5 4.4 ± 0.15 1.4 ± 0.07 76 % 24 % dehydrogenase

7.5 2.7 ± 0.01 0.7 ± 0.02 79 % 21%

* Units express total activity in μmol/min. †Values are means ± SE. Lipoamide dehydrogenase values reflect the means of five assessments at pH 5.5 and four assessments at pH 7.5 while all other values are the means of six assessments. 0.03-0.04 mg protein was used for membrane fraction assessments while 0.007 – 0.008 mg protein was used for soluble fraction assessments.

Both copper chloride (CuCl2) and cadmium chloride (CdCl2) are known inhibitors of A.

suum mitochondrial lipoamide dehydrogenase activity (Komuniecki and Saz, 1979). Within this

context, assessments of the effects of these inhibitors on the NADH→NAD+ transhydrogenation

and lipoamide dehydrogenase activities of disrupted A. suum mitochondria were performed and

these data are given in Table 2. Under either acidic or basic conditions of assessment, CuCl2

markedly inhibited both activities, when compared to corresponding controls, with the greater

inhibitions for both noted under acidic conditions. Under basic conditions of assay, CuCl2

exerted a greater inhibition on the lipoamide dehydrogenase activity, i.e., 96% versus 62%

inhibition. With the addition of EDTA to the assays, CuCl2 inhibition was clearly relieved for

both reactions with the greater relief being noted with the transhydrogenation reaction under

basic conditions (Table 2). Similarly, CdCl2 inhibited both activities under acidic and basic

conditions of assay with the greater inhibition noted in terms of the lipoamide dehydrogenase

activity. As with CuCl2, CdCl2 inhibition was relieved by inclusion of EDTA in the assay system

with the greater relief being observed under basic conditions (Table 2). Attempts to evaluate the effects of these inhibitors on the mitochondrial NADH dehydrogenase activity, as assessed by ferricyanide reduction, were unsuccessful, inasmuch as inhibitor additions resulted in a significant precipitation of ferricyanide.

Table 2. Effects of inhibitors on activities of Ascaris suum mitochondria assessed under acidic and basic conditions

Activity (µmol/min/mg)

NADH→NAD+ transhydrogenation Lipoamide dehydrogenase

Addition(s) a b a b

None 1.250 ± 0.062* (13)† 0.588 ± 0.024 (8) 1.269 ± 0.051 (14) 0.260 ± 0.012 (7)

CuCl2 0.040 ± 0.003 (5) 0.231 ± 0.024 (5) 0.014 ±0.006 (5) 0.011 (1) [97]‡ [62] [99] [96]

CuCl2 plus 1.113 ± 0.265 (3) 0.515 ±0.029 (4) 0.910 ± 0.090 (3) 0.283 ± 0.008 (3) EDTA [23] [7] [14] [+11]

CdCl2 0.512 ± 0.021 (8) 0.250 (2) 0.036 ± 0.005 (5) 0.014 ±0.003 (4) [63] [63] [97] [94]

CdCl2 plus 1.141 ± 0.118 (6) 0.65 (1) 1.038 ±0.117(5) 0.184 ± 0.004 (3) EDTA [14] [2] [22] [22]

*Values are means ± SE † Number of observations is in parentheses. ‡Percent inhibition compared to untreated sample is in brackets. a- designates acidic assay conditions, pH 5.5. b-designates basic assay conditions, pH 7.5. A mean protein concentration of 0.10 mg protein was used for assessments. Inhibitors were added such that the assay concentration was 0.4 mM while EDTA was 2.0 mM

Discussion

An evaluation of the effects of pH on the NADH→NAD+ transhydrogenation, lipoamide dehydrogenase, and NADH dehydrogenase activities of disrupted A. suum mitochondria is presented here. Significant differences in activities were observed for all three reactions at pH

6.0 and pH 6.5. Interestingly, at pH 6.0 and pH 6.5, the NADH→NAD+ transhydrogenation

reaction displayed an activity level in between that noted for NADH dehydrogenase and

lipoamide dehydrogenase. These differences in activity under acidic assay conditions suggest

that the NADH→NAD+ transhydrogenation reflects an activity that is catalyzed by both the

lipoamide dehydrogenase and NADH dehydrogenase systems. It was also noted that the NADH

dehydrogenase activity displays a marked increase in activity at pH 4.5 while the other two

activities significantly decline in activity at this pH. Rew and Saz (1974) demonstrated the

occurrence of two NADH cytochrome c reductase activities; one is associated with the IM and is

rotenone-sensitive while the other is associated with the OM and is rotenone-insensitive. In our

preliminary studies with the A. suum system, pH evaluations of both the rotenone-insensitive and

rotenone-sensitive NADH cytochrome c reductase activities indicated that the rotenone-

insensitive activity simulates the peak of activity noted for the NADH dehydrogenase activity.

Accordingly, it is suspected that the large peak in NADH dehydrogenase activity noted under

acidic conditions may reflect, at least in part, an NADH dehydrogenase component of the

rotenone-insensitive NADH cytochrome c reductase activity and warrants further investigation.

The thermal profiles of the NADH→NAD+ transhydrogenation, NADH dehydrogenase,

and lipoamide dehydrogenase reactions were determined. Because of the effects of pH observed

on these reactions, thermal lability assessments entailed evaluations under both acidic and basic

conditions. All three reactions ceased after prolonged exposure to heat, in keeping with an 27

enzymatic catalysis of these reactions. While under acidic conditions differences in activities

were not apparent until samples were incubated at 65⁰ C and 75⁰ C. At these latter temperatures, lipoamide dehydrogenase and NADH dehydrogenase activities differed from each other significantly. At 85⁰ C, lipoamide dehydrogenase activity differed significantly from both the

NADH dehydrogenase and NADH→NAD+ transhydrogenation activities. Furthermore, the corresponding experiments performed under basic conditions indicate marked differences in activities when samples were heated at 55⁰ C and above. Taken together, these data also support the notion that the NADH→NAD+ transhydrogenation is the product of two catalytic entities;

viz., NADH dehydrogenase and lipoamide dehydrogenase. Walker and Fioravanti (1995)

observed that both the mitochondrial lipoamide dehydrogenase and NADH dehydrogenase

systems of the adult, anaerobic cestode, Hymenolepis diminuta, are responsible for the catalysis

of the NADH→NAD+ transhydrogenation. In view of the present findings, and those reported by

Walker and Fioravanti (1995), it would appear that the catalysis of a mitochondrial

NADH→NAD+ transhydrogenation in the mitochondria of parasitic helminths may be a more wide spread phenomenon.

Employing both insoluble and soluble fractions derived from isolated A. suum

mitochondria, assessments of the NADH→NAD+ transhydrogenation, NADH dehydrogenase, and lipoamide dehydrogenase were performed. The data obtained demonstrated that all three activities have a predominant association with the mitochondrial insoluble fraction. However, despite this predominant association, and regardless of the pH of assessment, there was less

lipoamide dehydrogenase activity associated with the insoluble fraction, and more of this activity

in the soluble fraction, than noted for the other two activities. Indeed, the distributions of the

other two activities, i.e., NADH dehydrogenase and NADH→NAD+ transhydrogenation, were 28

essentially the same. These data again suggest that the NADH→NAD+ transhydrogenation is an

enzymatic activity that results from both the lipoamide dehydrogenase and NADH

dehydrogenase systems.

It has been reported that the lipoamide dehydrogenase and NADH→NAD+

transhydrogenation catalyzed by A. suum mitochondria are significantly inhibited by the presence of CuCl2 and CdCl2 (Komuniecki and Saz, 1979). In agreement with their findings,

both these salts were also found to be significantly inhibitory for the A. suum mitochondrial

NADH→NAD+ transhydrogenation and lipoamide dehydrogenase activities. Furthermore, the

inhibitions exerted by these salts were relieved by addition of the chelating agent EDTA to the assay system as observed by Komuniecki and Saz (1979). However, in the present study, CuCl2

seemed to be a less effective inhibitor on the NADH→NAD+ transhydrogenation reaction than

the lipoamide dehydrogenase reaction under basic conditions. Similarly, CdCl2 was a less

effective inhibitor of the NADH→NAD+ transhydrogenation reaction in comparison to

lipoamide dehydrogenase under both acidic and basic assay conditions. Interestingly, under

+ acidic conditions of assessment, CdCl2 did not inhibit the NADH→NAD transhydrogenation reaction as potently as CuCl2. These data suggest that there are differences between the

transhydrogenation and lipoamide dehydrogenase reactions in regards to the degree of inhibition

exerted by these salts. These differences are of note, inasmuch as one would expect that if the

NADH→ NAD+ transhydrogenation reaction is a result of lipoamide dehydrogenase, that both

activities would be inhibited to the same degree. Thus, there would appear to be another entity

responsible for the catalysis of the mitochondrial NADH→ NAD+ transhydrogenation reaction in

A. suum. 29

Based upon the collective findings obtained in the present study, a model is proposed

denoting the mitochondrial energetics of adult A. suum, and this model is given in Figure 2.4. As

indicated, Rew and Saz (1974) presented compelling data indicating that both the “malic

enzyme” and fumarase of adult A. suum mitochondria are predominantly localized in the

mitochondrial IMS.

Figure 2.4. Proposed roles of lipoamide dehydrogenase (LD) and NADH dehydrogenase (ND) as

NADH→NAD+ transhydrogenation mechanisms in the mitochondrial energetics of adult Ascaris

suum. Abbreviations used are as follows: OM, outer membrane; IM, inner membrane; IMS, intermembrane space; ME, malic enzyme; F, fumarase; FR, fumarate reductase.

These findings were supported both by fractionation of isolated A. suum mitochondria

coupled to sucrose gradient centrifugation, as well as by evaluating the release of enzyme

activities noted when increasing digitonin concentrations were applied to the isolated nematode

organelles (Rew and Saz, 1974). The findings of Rew and Saz (1974) were made emphatic by

assessments of biochemical markers for the OM, IMS, IM, and matrix fractions. Furthermore, the efficacy of fractionation by the techniques used here for the A. suum organelles was verified using mammalian (rat liver) mitochondria subjected to the same techniques and marker enzyme assessments (Rew and Saz, 1974). In contrast to these findings, Kohler et al. (1983), using digitonin treatment of isolated A. suum mitochondria and marker enzymes (although without a mammalian organelle control), reported that fumarase was chiefly localized in the mitochondrial matrix and that “malic” enzyme was equally distributed between the IMS and matrix.

Furthermore, Kohler (1977) indicated that fumarate did not traverse the ascarid IM. Regardless of these differences noted in a comparison of the Rew and Saz (1974) and Kohler et al. (1983) studies, the question of usage of “malic” enzyme dependent NADH formation by the ascarid system remains. The physiological utilization of NADH accumulated in the IMS by the ascarid electron transport system still necessitates a transmembrane transfer of reducing equivalents.

Thus, the need for a transhydrogenation(s) reaction remains, and our data suggest that this is accomplished via NADH dehydrogenase and lipoamide dehydrogenase activities as given in

Figure 2.4. In this regard it has been noted that IM associated, but externally oriented NADH dehydrogenase systems have been found in plants, animals, fungi, and protists, and also with respect to bacterial cellular membranes (Sotthibandhu and Palmer, 1975; Day et al., 1976;

Brailovskaya et al., 2003; Rasmusson et al., 2008). A number of anaerobic, succinate-forming parasitic helminths are known to display a mitochondrial NADH→NAD+ transhydrogenation 31

reaction; examples include, Spirometra mansoides (Cestoda), Hymenolepis microstoma

(Cestoda), Taenia crassiceps (Cestoda), and Setaria digitata (Nematoda), as reviewed by

Fioravanti and Vandock (2009). Thus, it would be of interest to determine the impact of lipoamide dehydrogenase and NADH dehydrogenase on transmembrane hydride translocation in these organisms.

Conclusions and Future Directions

The adult intestinal nematode of swine, Ascaris suum, exhibits a mitochondrial

NADH→NAD+ transhydrogenation reaction, which transfers reducing power in the form of

hydride ions across the IM to matrix NAD+, thus regenerating the NADH needed to drive the

NADH requiring, electron transport coupled, fumarate reductase (Fioravanti and Saz, 1976;

Kohler and Saz, 1976; Fioravanti and Saz, 1980). This transhydrogenation reaction has been demonstrated to be the result of lipoamide dehydrogenase in A. suum (Komuniecki and Saz,

1979). The mitochondrial NADH→NAD+ transhydrogenation reaction is also present in the

intestinal helminth of rats, Hymenolepis diminuta (Cestoda). However, the reaction in H.

diminuta has been shown to result from both the lipoamide and NADH dehydrogenase activities

(Walker and Fioravanti, 1995). Based on the findings in H. diminuta, studies to further assess the

origin(s) of the NADH→NAD+ transhydrogenation reaction in adult A. suum were undertaken.

The purpose of this study was to assess if the mitochondrial NADH→NAD+ transhydrogenation reaction in adult A. suum results from lipoamide dehydrogenase and NADH dehydrogenase, as it does in H. diminuta. Comparative evaluations of the NADH→NAD+

transhydrogenation, lipoamide dehydrogenase, and NADH dehydrogenase reactions were carried out using disrupted A. suum mitochondria as the source of enzymes. These three activities were 32 observed in regards to their response to varying pH levels, thermal labilities, and inhibitors.

Further comparisons with respect to their intramitochondrial localizations were also conducted.

The data obtained are consistent with the notion that the mitochondrial NADH→NAD+ transhydrogenation reaction in A. suum does indeed result from lipoamide dehydrogenase and

NADH dehydrogenase.

Future studies of the ascarid system could well entail a characterization of the relationships between these NADH-utilizing activities. Continued differentiation of these reactions can be pursued utilizing submitochondrial particles (SMP) derived from A. suum muscle. Using inverted SMP would permit a determination as to whether the oxidation of NADH results in the formation of a proton gradient within the vesicle. Gradient formation can be assessed employing an appropriate fluorescent probe. In addition, using purified lipoamide dehydrogenase, anti-lipoamide dehydrogenase antibodies (AB) can be generated. In turn, these

AB can be used to determine if the lipoamide dehydrogenase activity of SMP can be blocked. If lipoamide dehydrogenase activity is abolished/minimized in the presence of the AB, it would be of interest to determine if proton translocation into the vesicular space would still occur. These latter experiments would assist in further evaluations of the NADH dehydrogenase system as well. The reciprocal experiment using AB directed against the NADH dehydrogenase would also be of value. Certainly the experiments contained in this thesis as well as those given as future endeavors should permit an evaluation of these mitochondrial reactions within the context of potential sites for specific chemotherapeutic attacks by anthelmintics.

REFERENCES

Barret, J. 1981. Biochemistry of parasitic helminths. Baltimore, MD: University Press.

Behal, R.H., Buxton, D.B., Robertson, J.G., and M.S. Olson. 1993. Regulation of the pyruvate dehydrogenase multienzyme complex. Annual Reviews. 13: 497-520.

Bogitsh, Burton J., Clint E. Carter, and Thomas N. Oeltmann. 2005. Human Parasitology: Third Edition. Amsterdam: Elsevier Academic Press. Bradford, M.M. 1976. A rapid and sensitive method for quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72:248 254. Brailovskaya, I.V., Gamper, N.L., and M.V. Savina. 2003. External pathway of NADH oxidation in the liver of the river lamprey Lampetra fluviatilis. Journal of evolutionary biochemistry and physiology. 39: 261-265. Bueding, E. 1949. Studies on the metabolism of the filarial worm, Litomosoides carinii. Journal of Experimental Medicine. 89: 107-130. Bueding, E. and H.J. Saz. 1968. Pyruvate kinase and phosphoenolpyruvate carboxykinase activities of Ascaris muscle, Hymenolepis diminuta, and Schistosoma mansoni. Comparative Biochemistry and Physiology. 24: 511-518. Cain, G.D., and D. Fairbairn. 1971. Protocollagen proline hydroxyllase and collagen synthesis in developing eggs of Ascaris lumbricoides. Comparative Biochemistry and Physiology. 40B: 165-179. Cox, F.E.G., 2002. History of human parasitology. Clinical Microbiology Reviews. 15 : 595- 612. Dailey, M.D. 1996. Meyer, Olsen, and Schmidt’s Essentials of Parasitology 6th ed. Boston, MA: McGraw Hill. Day, D.A., Rayner, J.R., and J.T. Wiskich. 1976. Characteristics of external oxidation by beetroot mitochondria. Plant Physiology. 58: 38-42. Fairbairn, D. 1957. The Biochemistry of Ascaris. Experimental Parasitology 6: 491-554. Fairbairn, D. Wertheim, G., Harpur, R.P. and E.L. Schiller. 1961. Biochemistry of normal and irradiated strains of Hymenolepis diminuta. Experimental Parasitology. 11: 248-263. Fairbairn, D. 1970. Biochemical adaptation and loss of genetic capacity in helminth parasites. Biological Reviews 45: 29–72. Fioravanti, C.F., and H.J. Saz. 1976. Pyridine nucleotide transhydrogenases of parasitic helminths. Archives of Biochemistry and Biophysics 175: 21-30. 34

Fioravanti, C.F. and H.J. Saz. 1980. Energy metabolism of adult Hymenolepis diminuta. In: Biology of the tapeworm Hymenolepis diminuta. H.P. Arai, ed., Academic Press. New York, 463 – 504. Fioravanti, C.F. 1981. Coupling of mitochondrial NADPH: NAD transhydrogenase with electron transport in adult Hymenolepis diminuta. Journal of Parasitology 67: 823-831. Fiorvanti, C.F. and Y. Kim. 1983. Phospholipid dependence of the Hymenolepis diminuta mitochondrial NADPH:NAD transhydrogenase. Journal of Parasitology 69: 1048-1054. Fioravanti, C.F. and K.P. Vandock. 2009. Transhydrogenase and the anaerobic mitochondrial metabolism of adult Hymenolepis diminuta. Parasitology, 1-16. Fujimoto, D., and D.J. Prockop. 1969. Protocollagen proline hydroxilase form Ascaris lumbricoides. Journal of Biological Chemistry 244: 205-210. Hatafi, Y., and Y.M. Galante. 1977. Dehydrogenase and transhydrogenase properties of the soluble NADH dehydrogenase of bovine heart mitochondria. Proceedings of the National Academy of Science. USA. 74: 846-850. Hochachka, P.W. and T. Mustafa. 1972. Invertebrate facultative anaerobiosis. Science. 178:1056-1060. Keilin, D. 1966. The history of cell respiration and cytochrome. Cambridge, UK. Cambridge University Press. Kmetec, E. and E. Bueding. 1961. Succinate and reduced diphosphopyridine nucleotide systems of Ascaris muscle. Journal of Biological Chemistry. 236:584-591. Kohler, P. and H.J. Saz. 1976. Demonstration and possible function of NADH:NAD+ transhydrogenase from Ascaris muscle mitochondria. Journal of Biological Chemistry 251: 2217 – 2225. Kohler, P. 1977. The transport of dicarboxylates and some properties of fumarase in the muscle mitochondria of Ascaris suum. International Journal of Biochemistry. 8:141-147. Kohler, P., Gisler, J., Bachman, R., and P. Wild. 1983. The localization of fumarase and malic enzyme in muscle mitochondria of Ascaris Suum. Molecular and Biochemical Parasitology 9: 329-336. Komuniecki, R. and H.J. Saz. 1979. Purification of lipoamide dehydrogenase from ascaris muscle mitochondria and its relationship to NADH:NAD+ transhydrogenase activity. Archives of Biochemistry and Biophysics 96: 239-247. Lehninger, A.L. 1951. Phosphorylation coupled to oxidation of dihydrodiphosphpyridine nucleotide. Journal of Biological Chemistry. 190: 345-359.

Matuda, S. and T. Saheki. 1985. Immunochemical comparison of lipoamide dehydrogenases from various sources and reactivity of various lipoamide dehydrogenases with rat heart pyruvate dehydrogenase-subcomplex. Biochemical and Biophysical Research Communications. 129: 479-484.

Mckelvey, J.R. and C.F. Fioravanti. 1985. Intramitochondrial localization of fumarate reductase, NADPH→NAD transhydrogenase, ‘malic enzyme’ and fumarase in adult Hymenolepis diminuta. Molecular and Biochemical Parasitology 17: 253-263.

Rasmusson, A.G., Geisler, D.A., and I.M. Moller. 2008. The multiplicity of dehydrogenases in the electron transport chain of plant mitochondria. Mitochondrion 8: 47-60.

Read, C.P. 1972. Animal Parasitism. New Jersey: Prentice – Hall.

Rew, R.W. and H.J. Saz. 1974. Enzyme localization in the anaerobic mitochondria of Ascaris lumbricoides. Journal of Cell Biology 63:125-135.

Sakurai, Y., Fukuyoshi, Y., Hamada, M., Hayakawa, T., and M. Koike. 1970. Mammalian α-keto acid dehydrogenase complexes. Journal of Biological Chemistry. 245: 4453-4462.

Saz, H.J. and A. Weil. 1962. Pathway of formation of α-methylbutyrate by Ascaris lumbricoides. Journal of Biological Chemistry. 237: 2053-2056.

Saz, H.J. and E. Bueding. 1966. Relationships between anthelminthic effects and biochemical and physiological mechanisms. Pharmacology Review. 18: 871-894.

Saz, H.J. and O.L. Lescure. 1969. The functions of phosphoenolpyruvate carboxykinase and malic enzyme in the anaerobic formation of succinate by Ascaris lumbricoides. Comparative Biochemistry and Physiology. 30: 49-60.

Saz, H.J. 1971. Facultative anaerobiosis in the invertebrates: pathways and control systems. American Zoologist 11:125-135.

Saz, H.J., Berta, J., and J. Kowalski. 1972. Transhydrogenase and anaerobic phosphorylation in Hymenolepis diminuta mitochondria. Comparative Biochemistry and Physiology. 43B: 725-732.

Saz, H.J. 1981 Energy metabolisms of parasitic helminths: adaptations to parasitism. Annual Reviews Physiology 43: 323-341.

Scheibel, L.W. and H.J. Saz. 1966. The pathway for anaerobic phosphorylation in Hymenolepis diminuta mitochondria. Comparative Biochemistry and Physiology 18:151-162.

Scheibel, L.W., Saz, H.J., and E. Bueding. 1968. The anaerobic incorporation of 32P into adenosine triphosphate by Hymenolepis diminuta. 243: 2229-2235. 36

Schiller, E.L., Bueding, E., Turner, V.M., and J. Fisher. 1975. Aerobic and anaerobic carbohydrate metabolism and egg production of Schistosoma mansoni in vitro. Journal of Parasitology. 61: 385-389.

Seidman, I. and N. Entner. 1961. Oxidative enzymes and their role in phosphorylation in sarcosomes of adult Ascaris lumbricoides. Journal of Biological Chemistry. 236: 915- 919.

Slater, E.C. 2003. Keilin, cytochrome, and the respiratory chain. Journal of Biological Chemistry 278: 16455-16461.

Sotthibandhu, R. and J.M. Palmer. 1975. The activation of non-phosphorylating electron transport by adenine nucleotides in Jerusalem-artichoke. Journal of Biochemistry. 152: 637-645.

Walker, D.J. and C.F. Fioravanti. 1995. Mitochondrial NADH→NAD transhydrogenation in adult Hymenolepis diminuta. Journal of Parasitology 81: 350-353. Ward, C.W. and D. Fairbairn. 1970. Enzymes of beta-oxidation and the tricarboxylic acid cycle in adult Hymenolepis diminuta (Cestoda) and Ascaris lumbricoides (Nematoda). Journal of Parasitology 56: 1009-1012.