EVOLUTIONARY AND ADAPTIYE ASPECTS OF LIPID AND KETONE
BODY METABOLISM IN GASTROPOD MOLLUSCS
A Thesis
Presented to
The Faculty of Graduate Studies
of
University of Guelph
by
Jefiey Alan Stuart
L in partial fulfillment of requirernents
for the degree of
Doctor of Philosophy
@ JeBey Alan Stuart, 1998 Nationai Library Bibliothèque nationale ûfCana& du Canada Acquisitions and Acquisitions et Bibliographie SeMces services bbiiographiques 395 wdrig(on Street 395, rue Wellington OaawaON K1AW OItawaON K1AW canada Canada
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The author retains ownership of the L'auteur conserve la propriété du copyright in this thesis. Neither the droit d'auteur qui protège cette these. thesis nor substantial extracts fiom it Ni la thèse ni des extraits substantiels may be printed or otherwise de celle-ci ne doivent être imprimés reproduced without the author's ou autrement reproduits sans son ~ermission. r - -- autorisation. Evolutionary and Adaptive Aspects of Lipid and Ketone Body Metabohm in Gasuopod
Molluscs
Jeffrey Alan Stuart advisor: University of Guelph James S. Ballantyne
Terrestrial gastropods of the order Stylomrnatophora (Pulmonata) have a unique organization of ketone body metabolism, charactenzed by two cytosolic isoforms of the typically mitochondriai en y me O-hydroxy butyrate dehy dmgenase (B HBDH), which interconverts the ketone bodies acetoacetate and û-hy droxybutyrate (BHB ). One of these cytosolic BHBDH isoforms is further unique in that it is specûic for the L- enantiom=i af
BHB, whereas alI other known forms of BHBDH oxidize exclusively D-BHB. The occurrence of the two cytosolic BHBDH isoforms is tissue specific. L-BHBDHis found in hepatopancreas and kidney, and D-BHBDH is found in hem and kidney. Enzyme activities suggest that L- and D-BHB may be synthesized through the incomplete ondation of fatty acids in the kidney, where the highest advities of enzymes of lipid cataboiism are found, and transported to peripheral tissues for oxidation. The modified BHBDH isofonns do not occur in the closely related puhonate order Basommatophora, indichg that they have evolved only a single tirne, in the stylommatophoran lineage which succesâilly invaded terrestrial mvironments.
The importance of these modified pathways of ketone body metabolism during estivation was investigated kough measurements of maximal activities of ketogenidketolytic enzymes and those of carbohydrate, lipid and amino acid rnetabolism. No changes in the activities of any of these mqmes were observed after six weeks of estivation. Only the activities of citrate synthase and cytochrome c oxidase (CCO), which catalyze pathways which are "centraln to aerobic metabolism, are reduced during estivation.
The suppression of CC0 activity may be mediated by changes in the phospholipid composition of mitochondrial membranes which occur during estivation. The content of cardiolipin, which is essential for maximal CC0 autivity is reduced by 83%. Mitochondrial membranes of estivating mails dso contain fewer total phospholipids, fewer n-3 fatty xids and more monoenes, modifications which are consistent with reduced membrane proton permeability. Estivation induces other changes in the phospholipid and faw acid composition of the mitochondria which are consistent with a more stable, less fluid membrane, suggesting reduced biological activity. Thus mitochondnal function is regulated during estivation and mitochondrial membranes appear to be an important target of this regdation. ACKNOWLEDGEMENTS
1 would not have enjoyed, or been motivated to do, this work without the
Company of my wife, Connie. She has been a constant source of support and
encouragement throughout this adventure, and has never wavered in her belief in me
and what 1 have been doing (even when 1 have). 1 thank Savannah and Bai, who 1 hope will read diis one day and reaiize how importaut they have both been to my mental hedth, my ability to integrate the study of life with the Living of it.
Our yean in the Bdantyne Iab have been important and Mforgettable ones for both Connie and 1. My advisor, Dr. James Ballantyne, has created an environment nch in freedoms, interest and excitement... a lab that has aiways been a fun place to be. 1 am indebted to Jim for this, for never saying no, and for direction and advice at important intes (always given fairly and generously).
In my early years here, I benefted tremendously from my relationship with
Héléne Hussé Glémet, who showed me that science and lab work were fun. 1 consider her my earliest mentor and a very positive influence on my desire to continue with science.
1 have been fortunate to observe the work ethic and talents of my old labmate
Matin Gerrits, who showed me just how high personal goals could be set. 1 remai. in awe of his ability and endurance.
Most of my dechhg years in the lab were spent working alongside my good friends Todd Gillis (geaus Gillichthyes) Kimby Barton (Kim, for short), Dug Fudge
(and the wife, Esta Spalding) and Paul (Wog) LeBlanc. We had many a fun time in the lab, in Tobermory, at the Huntsman, and at 72 Glasgow St. These memones will remain with me always, and Fm hoping many more are in store.
1 have aijoyed also getting to know (however briefly) the new contingent of
Ballantyne mderlings: Arash, Kamla and Jason. 1 hope they get as much out of being here as 1 dici.
1 thank Ei Lin Ooi for her invaluable assistance this past year and a half.
Also, thds ta Sharky McLeod and Amy Boums.
So much for the lab ...1 have met some incredible people and made some
lasting fnends in the Dept. of Zoology. I am lucky to have befnended John
Wilmshurst. John, and Traudi OBiia, have been such great friends to us, and left a large impression on our memones of Guelph. 1 have benefited from many a good coffee break with Fudgie McCann, whose presence has been missed during Ej last year and a haif here. 1 have enjoyed the fneidship of Steve Crawford, whose biggest single accomplishment I believe to be the institution of Thursday hockey. Thanks
Steve.
1 thank my parents, Heather and Garry Stuart for providing such outstanding
examples of, well, people I guess, and for instilling me with a love of leaming at an early age. Their belief and encouragement has been critical to me throughout my mmy years of school. Their teachings have beai essential to my arriving at this point. Thanks too, to Connie's prnaits, Pat and Ed Powers, for all of their help and support over this long stretch of hard work and penodic hanaai insolvency. TABLE OF CONTENTS
Chapter 1
Generai htroduction ...... l
Chapter 2
Correlation of environment and phylogeny nit&the expression of bhydroxybutyrate dehydrogenase in the Moilusa ...... 16
Absrnt ...... **...... -...... 17 Introduction ...... -18 Materials and Methods ...... 21 Results ...... 27
Discussion ...... -29
Chapter 3
Importance of ketone bodies to the intermediary metabolism of the terrestrial snaii, ArckcICotinu ventricosu: evidence fkom enzyme activities ...... 41
iii
Discussion ...... ,...... *...... 93
Chapter 6
Tissue-specific forms of 8-hydroxybutyrrte dehydrogenase ondize the D or
G enantiomers of bhydrorybutyrate in the terrestrial gastropod
Cepaea nemordis ...... 104 Introduction ...... 105 Materials and Methods ...... IO6 ResuIts ...... 109 Discussion ...... ,...... ,.,...... lll
Chapter 7
A preliminary kinetic characterizatioi of the cytosolic D-O-hydrosybutyrate dehydrogenue from the heart of the terrestriai snaii, Cepaea mtmralis ...... 117 introduction ...... 118
Materials and Methods ...... *...... 120
Resuits anci Discussion ...... 124
Chapter 8
D- a~~dLa-hydroxybutyrate dehydrogenase and the evolution ofketone body metabolism in gastropod moUuscs ...... 137
Ab~trac~dintroduction...... ,...... 138 Materials and Methais ...... ****.*...... *...... *...... 140 Resuits and Discussion ...... 142
Chapter 9
Manmal activities of enzymes of intermediary metaboüsm in the er tivating terrestriai snaii Cepaerr nemwaik ...... 151 Abstract ...... 152 InflOdudon ...... 153 Materiais and Methods ...... 155
Results ...... 160 Discussion ...... 162
Chapter 10
Remodeiing of phosphoüpid fatty acids in mitochoidriai membranes
Introduction ...... 177 Materiais and Methods ...... 179
Chapter 11 Compodtionai correlates of metabolie depression in mitochondrid membranes of an estivating mail, Cepaea nenorulis ...... 200
Abstract ...... 21 Introduction ...... -202 Materiais and Methods ...... A Results ...... 208 Discussion ...... * ...... 210
Chapter 12
Generai discussion ...... *...... *.*...... *....221 Conclusions and prospectus ...... W
Lierature ated ......
vii List of Tables
Chapter 2
Table 2.1. 8-hydroxybutyrate dehydrogeme activity under standard assay conditions using various homogenization buffers ...... -35
Table 2.2. Activity of O-hydroxybutyrate dehydrogenase in marine, freshwater and tenestria1 molluscs ...... *36 Table 2.3 Activities of enzymes of lipid and ketone body metabolisrn in
Cepaea nemoralis, Stagnicola elodes and Littorina littoria heptopancreas ...... 37
Table 2.4 Hepatopancreas enzyme activities standardued w ith citrate sy nthase activity , in hepatopancreas of Cepaeu nemoralis, Stagnicola elodes and Littorîna lzttorio ...... -38
CIiapter 3
Table 3.1 Maximal activities of selected enzymes in tissues of
Amhachutina veniricosa ...... -56 Table 3.2 Maximal enzyme activities indexed to citrate synthase activity in venaicle/heart of the tenesuial mail Archachatina ventricosa , the marine snail Busycon contrarium and the marine elasmobranch hja erinacea ...... -~.0e~e~.0~...~m*~*-~*a*m**-*--***57
Chaptv 4
Table 4.1 Maximal activities of enzymes in Cepaea nemoralis hepampncreas ...... -...78
Table 4.2 Recovery of whole homogenate enzyme activities in
viii summed rnitochondr ial and cytosolic fractions and dis tribu tion of
enzyme activities benveen mitoc hondriai and cytosolic fractions ...... 79
Cbapter S
Table 5.1 Kinetic constants of Cepaea nemoruiis hepatopancreas 0-hydroxybutyrate dehydrogeme ......
Table 5.2 Effects of various metabolites on V, for B-hydroxy-
butyrate dehydrogenase in directions of acetoacetate reduction
and L-8-hydroxybutyrate dehydrogenation ...... 98
Table 5.3 Effects of various metabolites on K, for acetoacetate
and L-8-hydroxybutyrate in directions of acetoacetate reduction and LB-hydroxybutyrate dehydrogenation ...... -99
Table 5.4 Apparent Y values of 0-hydrox ybutyrate dehydrogenase
fiom different sources ...... lm
Chapter 6
Table6.1 ActivitiesofcitratesynthaseandD-hydroxybutyrate
dehydrogenase in mitochondrial and cytosol ic compartments of
Cepeu nemoralis tissues ...... 115
Chapter 7
Table 7.1 Kinetic constants of Cepea nemoralis heart D-0- hy&oqbutyrate dehydrogenase ...... 128
Table 7.2 Apparent &, values of 0-hydroxybutyrate dehydrogenase f?om different wurces ...... ,...129 Table 7.3 Effects of various metabolites on V, for D-8-hydroxy-
butyrate dehy drogenase in directions of acetoacetate reduction and L-
8-hydroxybutyrate dehydrogemîion ...... 130
Table 7.4 Effecu of various metabolites on K,,,for acetoacetate and L-B-hydroxybutyrate in directions of acetoacetate reduction and
LLI-hydroxybutyrate dehydrogenation ...... 13 1
Chapter 8
Table 8.1 Activities of O-hydroxybutyrate dehydrogenase and
marker enzymes in mitochondrial and cytosolic cornpamnents of gastfopod hepatopancreas ...... 14%
Table 8.2 Higher classification of gastropod species ued for rneasurements of &hydroxybutyrate activity ...... Al9
Chapter 9
Table 9.1 Protein concentration of tissues and weights of tissues from active and estivating Cepa nemoralis ...... 170
Table 9.2 Maximal activities of enzymes in kidneys of estivating and active Cepueu nemorrils ...... 171
Table 9.3 Maximal enzyme activities in hem of active and
~vatmgCepaememoraiis..t...... -172
Table 9.4 Maximal enzyme activities in hepatopancreas of active . . ande~t~va~gCepememoruiis...... 173
Table 9.5 pH effects on activities of hepampanmeas enzymes ...... 174 Chapter 10
Table 10.1 Percentages of individual fatty acids in lemice...... 193
Table 10.2 Percentages of individual fatty acids in hepatopancreas
mitochondriai phospholipids from active and estivating Cepaeu nernoralis ...... 194
Table 10.3 Percentages of individuai fatty acids in cardiolipin of
hepatopancreas mitochondria fiom active and estivating Cepea nemomlis ..... -195
Table 10.4 Percentages of individual fatty acids in phosp hatidy lethanolamine
of hepatopancreas mitochondria from active and estivating Cepaea nemoralis ...... 196
Table 10.5 Percentages of individuai fatty acids in phosphatidy lchol ine of
hepatopancreas mitochondria from active and estivating Cepaea nemoralis ...... -197
Table 10.6 Percentages of individual fatty acids in phosphatidylserine of
hepatopancreas mitochondria from active and estivating Cepueu nemoralis ...... 198
Table 10.7 Percentages of individual fatty acids in p hosp hatid y linos itoi
of hepatopancreas mitochondria from active and estivating Cepeanemordis ...... -199
Chapter 11
Table 11.1 Morphological and biochemical parameters ftom
hepatopancreasofactiveandestivatingsnaiI~...... 218
Table 11.2 Proportions (mole 46) of phospholipid species in mitochondrial
membranes of hepatopancreas fkom active and estivating snails ...... 219
Table 11.3 Proportions (mole %) of individual fatty acids in mitochondrial phosphoiipids frorn hepatopancreas of active and estivating snails ...... 220 List of Figures
Chapter 1
Figure 1.1 Phylogeny of major moIIuscan classes ...... -14
Figure 1.2 Pathways of ketone body metabolism in mammals ...... 15
Chapter 2
Figure 2.1 Effects of salt concentration on the activity of O-hydroxy- butyrate dehydrogenase fiom (A) the terrestrial snail Cepoea nemoralis and (B) the fkeshwater snail Stagnicolu elodcs ...... -39
Figure 2.2 Expression of O-hydroxybutyrate dehydrogenase in
rnolluscan taxa from different environmeno ...... 40
Chapter 4
Figure 4.1 Proposed organization of aerobic metabol ism, such as
~uldoccur following a bout of anoxia, in Cepaea nemoralis hepatopancreas ...... 8 1
Figure 4.2 Proposed pathways for the generation of anaerobic end p roducts in Cepueanemoraiis hepatopancteas ...... -82
Chapter 5
Figure 5.1 Effect of pH on V, of B-hydroxybucyrate dehydrogenase in
(a) the direction of acetoacetate reduction and (b) L-8-hydroxy butyr ate dehydrogenation ...... 101 Figure 5.2 Effect of pH on (a) apparent K, (acetoacetate) and (b) apparent Y (GB-hydroxyburyrare) ...... 102 Figure 5.3 Calculated velocities of B-hydroxybutyraue dehydrogenase
xii in the directions of L-0-hydroxybutyrate synthesis (a) and dehydrogenation 0).
under control conditions and in the presence of effectors ...... -103
Chapter 6
Figure 6.1 Cellulose acetate gel of ventricle, hepatopancreas and kidney stained with DL-fi-hydroxybutyrate ...... 116
Chapter 7
Figure 7.1 Demy of D-B-hydroxybutyrte dehydrogenase activity over time in tissues hcmogenized in different buffers ...... -132
Figure 7.2 Inhibition of D-B-hydroxybutyrate oxidation in the presence of varying concentrations of L-0-hydroxybutyrate ...... 133
Figure 7.3 Dixon plot of the inhibition of D-O-hydroxybutyrate oxidation by L-O-hydroxybutyrate ...... 134 Figure 7.4 Effecu of pH on the (a) iQcetoacetate and
(b) IQhydroxybutyrate ...... 135
Figure 7.5 Effecu of pH on V, in the direction of (a) D-fi- hydroxybutyrate oxidation and @) acetoacetate reduction ...... 136
Chapter 8
Figure 8.1 Gastropod phylogeny and the occurrence of mitochondriai
D-8-hydroxybutyrate dehydrogenase and cytosolic L-O-hydroxybutyrate dehydrogenase ...... 150
Chapter 12
Figure 12.1 Subcellular organhtion of pathways of ketogenesis and
xiii ketolysis in (a) Vertebrata. (b) Basornmatophora (Gasuopoda: PuIrnonata) and(c) StyIommatophora(Gastropoda: hihonata) ...... -237
Figure 12.2 Tissue organizaton of ketone body metabolisrn in stylommatophoran gastropods. showing proposed usages of pathways ...... 238 Figure 12.3 Proposed mechanism for modifying mitochondrial membrane composition during estivation ...... m.e...... 239
xiv CHAPTER 1
GENERAL INTRODUCTION The metabolism of animals is largely a function of phylogenetic history and environmental adaptation. However, the complex interactions between present and historical environmental change, and phylogeny, present a challenge to the interpretation of patterns underlying physiological characteristics (Garland and Carter,
1994). which can be rneasured only in extant species.
The interactions between environment and metabolic biochemistry may be particular 1y tangible in those species w hich do not expend energy regulating their interna1 environment, and must therefore adapt protein and membrane properties so that cellular biochemical processes are maintained under a range of ambient conditions. In general, two suategies for coping with variation in any environmental parameter can be identified: regulation or conformation (Hochachka and Somero,
1984). Perhaps the most extreme example of the strategy of regulation is provided by mammals, which maintain reiatively constant body temperatures and plasma and tissue ion levels in the face of extreme variations in environmentai levels of these parameters. The other end of this physiological spectrum, that of conformation, is typically represented by invertebrate species. Molluscs, for example, allow body temperature, tissue osmolarity and oxygen tensions to Vary with environmenui fluctuations in these parameters (Little, 1990). This strategy has profound implications for the metabolism, and ' metabolic organization' of these anirnals. For example, intertidal marine bivalves cannot regulate hemolymph oxygen tensions upon emersion, and are thus periodically exposed to hypoxic conditions. These species possess modified anaerobic pathways which permit swival during hypoxia (De Zwann, 1983).
The Mollusca thus provide an excellent mode1 for snidies of the interactions between metabolic organization, environment, and evolution. The second largest animal phylum, it includes a number of classes (Fig. 1.1), some with representatives in marine, freshwater and terrestrial environrnents. Molluscs have an evolutionary history dating to the appearance of marine species in the Cambrian era. Due to the presence, in most classes, of a hard shell, this evolutionary history is described by a relative1y cornpiete foss il record (Pojeta et al., 1987). The Gastropoda (snails) have shown perhaps the rnost impressive adaptability of all classes within this phylum.
From a marine or ig i n. gastropods have invaded freshwater and terresuial environrnents, requiring nurnerous morphological and physiological re-organizations
(Little, 1990). Arguably the most notable adaptation of non-marine gastropods to life on land or in ephemeral freshwater bodies is the ability to estivate. Estivation occurs in response CO desiccating environmental conditions, and is essentially an extended period of starvation accompanied by a dramatic suppression of basal metabolic rate
(Storey and Storey, 1990).
The metabolism of molIuscs has not been studied from the perspective of environmenial adaptation and phylogenetic history. Amino acid metabolism, and its regulation in marine molluscs, has been a primary focus of research on the interrnediary metabolism of molluscs (Bishop et al., 1983; De Zwaan, 1983; Moyes et al., 1990). Carbohydrare metabolism also has been extensively investigated in molluscs, largely because of the role of glycogen fermentation in sustaining facultative anaerobiosis ( DeZwann, 1983; Livingstone, 1983; Storey , 1993). The organization of lipid rnetabolism in molluscs has received relatively Meattention, though for reasons detailed below this is an important area of study.
In mammals, starvation and lipid metabolism are inexnicably linked. Due to the inability of most animals to store substantial arnounts of carbohydrate or to synthesize it from Iipid, carbohydrates become limiting in the absence of food (see
Sugden et al., 1989 for review). As a result, tissues such as brain (Robinson and
Williamson, 1980), which rely heavily on carbohydrates, mut switch to other fuels.
In mammals, such tissues increase their reliance on ketone bodies as a form of oxidizable fuel when carbohydrate sources are limiting (Newsholme and Leech,
1983). To provide ketone bodies, stored lipid is transported in the blood to the liver where it is partially oxidized to ketone bodies which are, in turn, uansported to exvahepatic tissues for oxidation (Newsholme and Leech, 1983). The applicability of this model to other groups of organisms has not been examined.
Though the organization of lipid and ketone body metabolism of molluscs remains largely unexplored, terrestrial gasnopod molluscs represent an interesting model for the study of these pathways because of the important role periods of food deprivation play in their life history. In the face of environmental desiccation, the rypical gasrropod response of reueating into the sheil, secreting an egiphragm over the shell aperture, and remaining inert occurs at the price of cessation of feeding.
These snails must, therefore, periodically endure periods of food deprivation which, during drought conditions, can persist for months or, in some habitats, years (Schmidt-Nielsen et al., 1971). Their ability to do so is signifiicantly enhanced by the adaprive strategy of estivation, in which basal metabolic rate is reduced to only about
16% of normal (Hand and Hardewig, 1996).
Previous studies of intermediary metabolism during estivation have focused almost exclusively on the regulation of glycolysis, and a large body of lirerature now exists describing this (for reviews see Storey and Storey, 1990; Guppy et al., 1994;
Hand and Hardewig, 1996). Many glycolytic enzymes are modified upon enny into estivation by reversible phosphorylation. Enzyme phosphorylation has been shown to alter Km and K, values and thus affect rates of catalysis (Storey, 1993). However, information regarding the control and regulation of other aspects of intermediary metabolism is largely lacking, though a number of studies have demonstrated the disappearance of lipid stores throughout estivation (Heeg, 1977: Krupanidhi et al.,
1978; Rees and Hand, 1993). In this thesis, I address two poorly understwd areas of estivation metabol ism, rnechanisms for the conuol of mitochondrid oxidative metabolism, and the participation of pathways of lipid and ketone body metabolism in estivation.
Studies of the lipid metabolism of terresuial mails have been confined to measuremenu of stored lipid under different experirnental conditions (Heeg, 1977;
Krupanidhi et al., 1978; Umezurike, 1983; Rees and Hand, 1993). These show the presence of relatively large lipid stores, especially in hepatopancreas, which are substantially reduced during estivation. How lipid stored in hepatopancreas is accessed by peripheral tissues is poorly understood. Virnially no data on activities of enzymes of lipid metabolism are available and studies of lipid oxidation by mitochondria are not possible. Attempa to isolate well-coupled. respiring mitochondria from terrestrial snails have been without exception unsuccessful
(Vorhaben et al. 1980; M. Gupp y, personal communication; personal observations).
A single snidy represenu the state of knowledge on the ketone body metabolism of molluscs (Meyer et al., 1986). Ketone body metabolism has been studied primaril y in vertebrates, where pathways of ketogenes is and ketolysis are catalyzed by five enzymes (Fig. 1.2). Acetyl-CoA, derived primarily from the incornpiete oxidation of fatty acids, enters the ketogenic pathway at acetoaectyl-CoA thiolase (AAT) which catalyzes the production of acetoacetyl-CoA from two molecules of acetyl-CoA. Acetoacetyl-CoA enters sequential reactions catalyzed by hydroxymethylglutaryl-CoA synthase (HMS)and hydroxymethylglutaryl-CoA lyase
(HML)which result in formation of acetoacetate (acac). The enzyme B- hydroxybutyrate dehydrogenase (BHBDH)catalyzes the interconversion of acac and
BHB. There is some ambiguity as to whether molluscs are capable of synthesizing or oxidizing both ketone bodies, acac and O-hydroxybutyrate (BHB). BHBDH is undetectable in a marine gastropod (Beis et al., 1980), and mitochondria isolated from tissues of marine molluscs do not oxidize the ketone body BHB (Bailantyne et al., 1981; Ballantyne and Moyes 1987). BHBDH has, however, been demonsuatecl in a number of other gastropod species (Meyer et al., 1986; Rudolph and Burch, 1987:
Mimfoundi and Greer, 1989). Taken together, these studies suggested that the occurrence of BHBDH in mollusc species may be a Function of environmental adaptation andlor phylogenetic history. Thus, a fist step in characterizing ketone body metabolism in rnolluscs was to investigate the bais for the occurrence of
BHBDH in only some rnollusc species.
A comprehensive study of the occurrence of BHBDH in molluscs from different environments and phylogenetic positions is, therefore, the focus of chapter
2. Measurements of BHBDH activity were made in a number of mollusc species, and then analyzed using the software application MacClade (Maddison and Maddison,
1992). With this approach, character States are mapped ont0 an existing cladograrn and parsimony analysis used to identify phylogenetic and environmental patterns underiying the occurrence of the enzyme. I identified an environmentai component to the occurrence of BHBDH, which is found in al1 fieshwater and terrestrial mollusc species, apparently regardless of taxonomic group, but is absent from marine species.
To examine whether this was related to a difference in the importance of ketone body metabolism between marine and non-marine species, measurements of al1 five of the enzymes which catalyze steps in the ketogenic and ketolytic pathways were made in marine, freshwater and ierresuial species.
1 further investigated the organization of ketone body metabolism in terrestrial gastropods in Chapter 3. This study represented the initial step in identifying the organization and tissue disuibution of pathways of ketone body metabolism, and also the relative importance of these pathways compared to those of carbohydrate, amino acid and fatty acid metabolism in a non-aquatic mollusc. Measurements of al1 known ketogenic and ketolytic enzymes and of key enzymes from pathways of carbohydrate, amino acid and lipid metabolism, the pentose phosphate shunt and the tricarboxylic acid cycle, were made in foot muscle, mantle, heart, kidney and hepatopancreas. For this study, giant African terresaial snails Archachatina ventricosa were used, as they provided tissues of sufficient quantity for measurements of al1 of the necessary enzymes on each tissue. Interestingly, while completing this study, a unique subcellular organization of pathways of ketone body metabolism was found, which was further explored in chapters 4 to 8.
BHBDH has been studied primarily in mammals, where the enzyme is localized to the mitochondrial inner membrane and maintains an essential association w ith the phospholipid phosphatidylcholine (Sandermann et al., 1986). The apoenzyme, i.e. BHBDH removed from the membrane and delipidated, is inactive
(Sandermann et al., 1986). This mitochondrial isoform of BHBDH is found in vertebrates (Beis et al., 1980) and in many invertebrates (Meyer et al., 1986;
Gibbelato and Chamberlin, l994). However, 1 found that mitochondrial BHBDH is absent in hepatopancreas of terrestrial snails, but BHBDH activity appeared to occur in the cytosol. This contrasu with the mitochondrial location of dl previously known forrns of BHBDH, and is the only known instance of a cytosolic BHBDH in animals.
A similar cytosolic activity had been identified in the livers of ruminant mammals
(Koundakjian and Snoswell, 1970). though this finding was subsequently refuted
(Williamson and Kuenzel, 1971). Thus, in Chapter 4, 1 verified the subcellular local kat ion of hepatopancreas BHBD H activity and character ized the subcellular organization of intermediary metabolism in a tenesuial gastropod. For this study, I used Cepaea nemoraiis, a terrestrial snail which occurs ldlyand can be collected readily.
In Chapter 5, a detailed kinetic characterization of the cytosolic BHBDH from
C. nemoralis hepatopancreas was underraken to provide an initial description of this unique enzyme. An initial hypothesis was that the cytosolic BHBDH could be adaptive during periods of hypoxic stress, by facilitating the maintenance of redox balance in the cytosol. Thus, the hinction of the enzyme was studied under assay conditions designed to simulate the intiacellular conditions in which BHBDH operates during hypoxia and recovery from hypoxia. The ability of various metabolites and pH to rnodulate the apparent Km for substrates and V,, in borh forward and reverse reaction directions was inves tigated .
Subsequent studies of C. nemomiis led to the identification of a second unique cytosolic isoform of BHBDH, which differs in enantiomeric specifity, and occurs in other tissues. The presence of this second unique enzyme and aspects of its subcellular and tissue distribution are the focus of Chapter 6.
In Chapter 7, a detailed kinetic characterization of this second cytosolic isoforrn of D-BHBDH,similar to the approach taken in Chapter 5, was undertaken with the enzyme from C. nemoralis heart.
Though the typical mitochondrial BHBDH isoform is not found in tissues of
C. neniorah, it has been previously reported in tissues of the fieshwater pulrnonate snail Biomphaiaria glabrata (Meyer et ai., 1986). To understand the basis for the apparent occurrence of mitochondrial and cytosolic isoforms of BHBDH in different gastropod species, a survey of freshwater and terrestrial gastropod species, combined with an analysis of character evolution similar to that employed in Chapter 2 was undertaken in Chapter 8. This smdy indicated a relationship between the occurrence of cytosolic BHBDH isoforms and the success of species' invasions of terrestrial environments .
The invasion of terrestrial environrnents has required a number of adaptations in gastropods, and perhaps the most important of these is the ability to withstand prolonged desiccation by estivating. The presence of highly modified pathways of ketone body metabolism in the most successful terresuial snails suggesu that these substrates may be important during estivation. Studies of lipid and ketone body metabolism in chapters 2 and 3 indicate that most snail tissues have a very limited ability to oxidize fatty acids. The energy stored in lipid may thus be made available to tissues as ketone bodies, following the incomplete oxidation of Myacids in kidney. Lipid stores are urilized during long-term estivation (Heeg, 1977; KNpanidhi et al.. 1978; Rees and Hand, 1993). ar rates which suggest that their consumption accounts for at least a third of total energy usage.
The roles and regulation of lipid and ketone body metabolism during estivation were investigated in Chapter 9. Maximal enzyme activities in tissues of active snails were compared with those which had been estivating for six weeks.
During estivation, hem01 y mph and tissue pH drop by approximately 0.5 units (Rees et al., 199 l), and this acidificarion has been found to account for approxirnately 35 % of the overall metabolic depression in snail tissues (Pedler et al., 1996). Thus, 1 aiso investigated the abil ity of p hysiological reductions in pH to modifj maximal enzyme activities, to determine if a direct effect on these activities is involved in the observed metabolic depression. The experimental design described in Chapter 9 allowed four distinct hypotheses to be tested: (1) do the activities of enzymes of ketone body metabolism, or of other pathways, change in response to estivation; (2) are these responses tissue-specific; (3) are there differences in the relative changes of key enzymes that relate to differential responses of pathways of carbohydrate, arnino acid or fatty acid metabolism. and (4) is there a direct effect of a reduction in pH (equal to that which occurs during estivation) on activities of these enzymes?
These experiments provided no evidence that pathways of lipid and ketone body metabolism are emphasized during estivation. Rather, they indicated that suppression of catabolisrn in long-term estivation does not occur through the downregulation of activi ties of enzymes of carbohydrate, amino acid, lipid or ketone body metabolism. This suggesu that the regulation of catabolism in long-term estivation may occur in pathways which are "centraln to oxidative metabolism. These pathways (the electron transport chah (ETC)and the TCA cycle) are housed within the mitochondria. While the smdy of mitochondrial hinction in estivating snails has been limiied by the difficulty of isolating well-coupled, respiring mitochondria from tissues of these anirnals, other approaches are possible. Many mitochondrial pathways , including the ETC, are catalyzed b y enzymes and transporters embedded within the inner and outer mitochondrial membranes (Voet and Voet, 1990).
Membrane-bound proteins show a dependence on propemes of the bulk phospholipid bilayer, such as fluidity (Carruthers and Melchior, 1986). order (Hazel, 1995), or phase behaviour (Epand, 1990). Some proteins have specifc phospholipid requirernents for optimal function. For example, phosphatidy lcholine is a cofactor in the reaction catalyzed by the membrane-bound enzyme BHBDH (El Kebbaj and
Latruffe, 1986). Cytochrome c oxidase (CCO) requires the phospholipid cardiol ipin for activity (Robinson, 1993). Therefore. just as the activities of soluble proteins are suppressed through post-translational means such as reversible phosphorylation, the activities of mitochondrial membrane-bound proteins could be suppressed through alterations to the phospholipid composition of the i~erand outer membranes.
This rationale led to the studies described in chapters 10 and 11, where 1 inves tigated the potential for regulation of mitochondrial oxidative metabol isrn through the modification of membrane phospholipid composition. In Chapter 10, the phospholipid fatty acy l chah compositions of mitochondrial membranes frorn hepatopancreas of active and estivating snails were characterked.
The resulu of Chapter 10 suggested that a large loss of mitochondrial phospholipid occurs during estivation. In Chapter 11, measurements of whole hepatopancreas phospholipids in active and estivating snails were made to determine whether phospholipid lost fiom the mitochondria remains within the ce11 or is metabolised. Paralle1 measuremenu of the ETC enzyme CC0 were also made as an indirect test of the ability of altered membrane composition to modulate membrane- bound enzyme activities. These resulu suggest a phospholipid-mediated suategy for regulation of mitochondrial membrane function during estivation. Taken together, the studies of enzyme and membrane composition and adaptation described in this thesis serve to highlight the importance and unique organization of ketone body metabolism in the intermediary metabolism of pulmonate gastropods and connibute to the understanding of mechanisms which mediate the regulation of intermediary metabolism during estivation. Fig. 1.1. Phylogeny of the major molluscan classes (Pojeta, 1987) showing the inter- relationships of Aplacophora (wormlike moiluscs); Monoplwphora (limpeu); Polyplacophora (chitons); Cephalopoda (octopus, squid); Gastropoda (mils, slugs); Scaphopoda: Pel ycopoda (b ivaives) . hydmrryma~yfglutiryl-
AAT BHBDH 2 rcatyl4oA --t=-œLyl.coA \=-am- CUiydroxybutyn6b
Fig. 1.2. Pathways of ketone body metabolism in manmals. Five enzymes catalyze reactions in distinct ketogenic and ketolytic pathways. Reactions catalyzed by MT and BHBDH are fully reversible. 30AT is also reversible, but is believed to function primarily in the ketolytic direction, while ketogenesis is catalyzed by HMS and HML. Abbreviations: AAT = acetoaceryl-CoA thiolase; HMS = hydroxymediylglu~fyl-CoA synthase; HMt = hydroxymethylglutary1-CoA lyase; 30AT = 3-oxoacid CoA- transierase: BHBLW = O-hydroxybutyrate dehydrogenase. Correlation of environment and phylogeny with the expression of
0-hydroxybutyrate dehydrogenase in the MoUusca
published in Comparative Biochernistry and Physiology, 1996, 114B: 153-160 Abstract
The enzyme O-hydroxybutyrate catalyzes the interconversion of the ketone bodies, acetoacetate and O-hydroxybutyrate. While the activity of 8-hydroxybutyrate dehydrogenase is relative! y high in fieshwater and terrestrial molluscs, it is undetectable in al1 marine molluscs examined. We tested the effects of conditions of tissue preparation and ionic strength of assay media on the measurement of O- hydroxybutyrate dehydrogenase and conclude that the apparent absence of this enzyme in marine molluscs is not an artifact of conditions of measurement.
Measurements of other enzymes involved in ketone body and lipid metabolism in the hepatopancreas of the marine snail Littorina littoria, the freshwater snail Stagnicola elodes and the terrestrial snail Cepaea nenioralis, suggest that the differential expression of the enzyme may be related to differences in the metabolic organization of marine molluscs and their freshwater and terrestrial counterparts. Mapping of the presence/absence of B-hydroxybutyrate dehydrogenase ont0 a bivalve and gastropod phylogeny indicates Four independent occurrences of the enzyme. INTRODUCTION
The ketone bodies. acetoacetate (Acac) and 8-hydroxybutyrate (BHB), are important metabolic substrates in some organisms. Throughout the animal kingdom they are used variously in the specialized role of glucose replacement during starvation (Newsholme and Leech, 1983). as subsaates for the synthesis of lipids
(Williamson, 1985) or as energy subsaates in routine intermediary rnetabolisrn
(Meyer et al., L 986; Pontes et al., 1988). In mammals, ketone bodies are produced in the liver mainly from the incomplete oxidation of fatty acids and are exported to peripheral tissues (see Newsholme and Leech, 1983).
The enzyme O-hydroxybutyrate dehydrogenase (BHBDH)catalyzes the interconversion of Acac and BHB. The occurence of this enzyme in species representing various invertebrate and vertebrate phyla has been studied by Beis et al.
(1980). These authors demonstrated that, while virtually al1 animals have the capacity to produce Acac, many lower vertebrates and invertebrates lack BHBDH (Beis et ai.,
1980) and therefore cannot produce or consume BHB. BHB is a more reduced substrate than Acac, and thus better fulfilh the role of oxidative substrate in tissues.
The lack of BHBDH in many lower organisms has been termed 'primitive'
(Newsholme and Leech, 1983).
Beis et al. (1980) found BHBDH to be absent in a marine gastropod mollusc, crustaceans, insects, teleost fish and a reptile. BHBDH is present in chondrichthian fih, amphibians. birds and mamrnais (Beis et ai., 1980). More recentiy, however, the enzyme has been demonstrated in a reptile (Pontes et al., 1988) and severai teleosr fish (LeBlanc and Bal lantyne, 1993).
The occurence of BHBDH in the rnollusca has also been demonsuated (Meyer et al., 1986; Rudolph and Burch, 1987; Mimfoundi and Greer, 1989). There is evidence that BHBDH expression in molluscs may be related to habitat or phylogeny.
Meyer et al. (1986) rneasured signifiwit BHBDH activity in hepatopancreas of a freshwater gastropod, Bionzphaiaria glabrata. BHBDH has also been demonsuated in other freshwater gastropods (M imfoundi and Greer 1985; Rudolph and Burch 1987).
Marine molluscs apparently lack the enzyme (Beis et ai.. 1980; Ballantyne et al.,
198 1; Bal lantyne and Berges, 199 1).
To understand the factors responsible for the presence or absence of BHBDH in the Mollusca, we examined the occurrence of the enzyme in marine, freshwater and terrestrial representatives of two of the three major molluscan classes, gastropoda and bivalvia. Since the differences in intracellular ionic strength (see Burton, 1983) between marine versus freshwater and terrestrial molluscs may influence the function of BHBDH, it was assayed over a range of concentrations of sodium and potassium using gastropods representative of each environment. The extent to which the general organization of lipid and ketone body metabolism might be correlated with the presence or absence of BHBDH in molluscs was investigated through the measurement of maximal activities of selected enzymes in these metabolic pathways.
TO aid in our assessment of the influences of phylogenetic history and environmental adaptation, we mapped the presencehbsence of BHBDH ont0 a phylogeny of gastropods and bivalves using MacCIadem (Maddison and Maddison, 1992), to determine the most parsimonious explanation for iu occurence in different taxa. MATERIALS AND METHODS
Experimental nnim als
Marine molluscs were collected near St. Andrews, New Brunswick in the fdl of 1992 and 1993, with the exception of Cfione limacina, which was collected from
the Arctic Ocean at Igloolik Bay, Igloolik, NWT. AI1 marine molluscs were used
immediately or mainüiined at the University of Guelph at 10" C in a recirculating seawater system (32 ppt) for several months prior to experimentation. Herbivorous
marine gastropods were fed lettuce daily. Predatory marine gastropods fed on other
molluscs and herring. Marine bivalves were fed suspensions of herring and shrimp.
Terrestrial molluscs, except Archachatina ventricosa, were collected on and around the campus of the University of Guelph in July and August of 1992, and kept in terraria until used, during which time they were fed lettuce ad libitum. A. ventricosa were taken from our laboratory breeding colony, initiated by snails obtained from the
Toronto Meao Zoo. Freshwater molluscs were collected from several ponds near the
University of Guelph campus, with the exception of Vivipanrs georgiunus and
Pamacaea bridgesi which were obtained from a local supplier. These animals were kept in filtered freshwater aquaria at 20" C for up to one week before use and fed lettuce ad libitum.
Tissue Preparation
As heptatopancreas is considered the functional liver in molluscs, preparations of this tissue were used for eruyme measurements when possible.
Where possible, enzyme activities in other tissues were measured as well. In some cases it was necessary to use whole animals. These are indicated in table 2.2.
LeBlanc and Ballantyne (1993) demowtrated that BHBDH activity in teleosu
was best preserved with a homogenization buffer containing glyceroi. We, therefore,
examined the ability of five different tissue homogenization buffers to preserve
BHBDH activity in the hepatopancreas of Anodonta grandis (Table 2.1).
Ali tissue preparation procedures were carried out on ice. Molluscs were
deshelled, the tissue(s) removed and placed in 2 rnL of ice-cold homogenization
buffer.
Tissues were homogenized using three 10 s bursu at half power with a
Polyuon PT10 unit (Kinematica Gmbh., Luzurn, Switzerland). The homogenates
were centrifuged at 14,500 g for 10 min at 2" C. Aliquots of this supernatant were
used immediately in enzyme determinatiow. In instances where no BHBDH activity
was detected, aliquou of uncentrifuged homogenate also were assayed.
Isolation of mitochondria for carnitine acyihansferase measuremeats
Due CO interference from high cytoplasmic deacylase activiv, the activities of
car nitine palmitoyl tram ferase (CPT)and carnitine octanoyl transferase (COT)were
rneasured in mitochondria isolated from hepatopancreas. Molluscs were deshelled and the hepatopancreas removed and placed in 2 mL of ice-cold isolation medium A or 8, as detailed below. Tissues were homogenized with 3 passes of a Poner-
Elvehjem tissue homogenizer. Homogenates were centrifiiged at 500g for 10
minutes, the supernatant collected and centrifuged at 10,Oûûg for 10 minutes. The pellet was resuspended and "washed" with a second 10 minute, 10,000g centrifugation and resuspended in 2 mL of isolation medium containing 2% Triton X-
100. Aliquots of this suspension were used immediately in CPT and COT determinations. Isolation medium A, used with C. nernoralis and S. elodes, contained
20 mM N-[2-hydroxyethyi]piperazine-N'-[2-ethanesuIfonic acid] (HEPES) and 50 mM KCI, pH 7.5. The isolation medium B, used with L. littoria, contained 20 mM
HEPES, 500 mM sucrose and 200 mM KCI, pH 7.5. Both isolation media gave over
85 1 yield of mitochondria as tested using cytochrome C oxidase as a mitochondrial marker enzyme.
Enzyme assays
Maximal enzyme activities were determined using a Varian DMS 100 UV- visible spectrophotorneter (Varian Canada Inc., Georgetown, Ont., Canada) equipped with a thermostated cell changer maintained at required temperature with a Haake D8 circulating water bath (Haake Buchler Instruments Inc., Saddlebrook, NJ). Rates of reactions involving NADH were followed at 340 nm (rnillimolar extinction coefficient s, = 6.22). Reactions involving 5,s'-dithio-bis-(2-niaobenzoic acid) (DTNB)were followed at 412 nm (millimoiar extinction coefficient s4l2 = 13.6). Acetoacetyl-CoA awferase (AACT) . 3-oxoacid transferase (30AT)and hydroxymethylglutaryi-CoA synthase (HMS)were measured by following the accumulation or disappearance of acetoacetyl-CoA at 303 nm (millirnolar extinction coefficient E, = 13.8). Enzymes were measured at environmentai temperature, which was 20 OC for al1 terrestrial and freshwater molluscs and at 10 OCfor al1 marine molluscs. Enzyme activities were expressed as units per gram of wet tissue weight where 1 unit equals 1 pmol of substrate converted to product per minute. Enzyme activities measured in isolated mitochondria were also expressed on a per tissue weight basis. Activities of these mitochondrial enzymes were adjusted for 85 56 recovery of the mitochondrial fraction, as established in preliminary experiments. Activities of enzymes from fatty acid oxidation pathways in molluscs are stimulated by KCI (personal observations).
Optimal levels of KCI were included in the assay medium for these enzymes. The following optimal assay conditions were determined with respect to substrate and cofactor concentrations.
Lipid catabolism : carnitine palmitoyl transferase (CPT) (E.C. 2.3.1.21): 0.1 mM DTNB, 50 pM palmitoyl-CoA, LOO mM KCl, 5 mM L-carnitine (omitted for control), pH 8.0. carnitine octanoyl transferase (COT) (E.C. 2.3.1.21): 0.1 mM DTNB, 50 pM octanoyl-CoA, 200 mM KCI, 5 mM L-carnitine (omitted for control), pH 8.0.
3-hydroxyacyl-CoA dehydrogenase (HOAD) (E.C. 1.1.1.35): 0.2 mM NADH, 0.1 mM acetoacetyl-CoA (omitted for control), 300 mM KCl, pH 8.0. oridative metaboiism : citrate synthase (CS) (E.C.4.1.3.7): 0.1 mM DTNB, 0.3 mM acetyl-CoA, 0.5 mM oxaloacetate (omitted for control), pH 8.0. ketone body metabolism: acetoacetyl-CoA thiolase (AN) (E.C. 2.3.1.9): 10 mM MgCl*, 0.16 mM acetoacetyl-CoA, 0.2 mM CoA (omitted for control), pH 8.0. 3-oxoacid tramferase (30AT) (E.C. 2.8.3.5): 0.4 rnM succinyl-CoA, 5 mM MgCl,,
5 mM iodoacetamide, 10 mM acetoacetate (omitted for control). pH 8.0.
BHBDH (E.C. 1.1.1.30): 2 mM DïT, 2 mM NAD, 40 mM DL-O-hydroxybutyrate
(omitted for control), pH 8.2 hydroxymethylglutaryI-CoA synthase (HMS) (EX.4.1.3.5): 10 mM MgCI?, 2 mM
Dm,5 mM acetyl phosphate, 10 pM acetoacetyl-CoA, 100 pM acetyl-CoA and 10 units phosphotransacetylase (PTA) (both ornitted for control), pH 8.0. hydroxymethylgluraryI-CoA lyase (HML)(E.C. 4.1.3.4): 10 rnM MgCI,, 0.2 mM
NADH, 1 unit BHBDH. 0.2 mM HMG-CoA (omitted for control), pH 8.0.
50 mM imidazole buffer was used in al1 assays. For determinations of KCl and NaCl effects on BHBDH activity, the smndard BHBDH assay was used with the addition of the appropriate concentration of salt. Saturating substrate conditions were tested at the beginning, rnidpoint and end of salt ranges. Standard assay conditions were found to be saturating over the tested ranges of sait concentrations.
Chemicals
Al1 chemicals except oxaloacetate, KCl and MgCl, were obtained ftom the
Sigma Chernical Co. (St. Louis, MO). Oxaloacetate was obtained fiom Boehringer-
Manheirn Canada. KCI and MgCI, were obtained From Fisher Scientific (Ottawa,
Ontario). S tatistics
Homogenization buffers 1 through 5, as described above, were compared us ing ANOVA and activity means compared using Bonferroni's procedure on s Y STAT~~.
Phylogeny of BHBDH expression
The computer program MacCIade (Version 3.04, Maddison and Maddison
1992) was used to map the presence/absence of BHBDH ont0 a phylogeny of gastropod and bivalve evolution constructed using an amalgamation of the morphology-based bivalve phylogeny suggested by Pojeta (1987) and a modification of the gastropod molecular phylogeny of Tillier et al. (1994), which corresponds closely to the morphology-based gastropod phylogeny of Haszprunar (1988). RESULTS
A significant difference in the ability of homogenization buffer 1 through 5 CO preserve BHBDH activity was found using ANOVA (P = 0.002). Homogenization buffer 5 preserved significantly greater enzyme activity than buffers 1 and 2 (P =
0.002 and P = 0.019, respectively) (Table 2.1). No other signifiant differences between homogenization buffers were found. Homogenization buffer 5 was used in al1 tissue preparations for enzyme measurements in al1 molluscs. In marine molluscs,
BHBDH also was assayed in tissue prepared using hornogenization buffer 1.
BHBDH activity was undectable in any tissues from marine gastropods and bivalves under any conditions of tissue preparation or enzyme assay tested. The enzyme was present in al1 terresuial and freshwater gastropods and bivalves examined (Table 3.2). BHBDH activities in a11 freshwater and terrestrial gastropods was relatively high, compared with the values for bivalves.
BHBDH activity was maintained over a range of [KCl] and [NaCl] from O to
350 and O to 250 mM, respectively, in the hepatopancreas of the terrestrial mail C. nemoralis (Fig. 2.l(a)) and the freshwater snail S. elodes (Fig. 2.l(b)). These ranges of WC11 and [NaCl] are p hysiological in L. littoria. No BHBDH activity was detected in the hepatopancreas of L- littona over these sarne ranges of [KCl] and
[NaCl].
CPT and COT, enzymes invoived in the carnitine-dependent oxidation of fatty acids, are present in hepatopancreas of al1 three snails, with the exception thar COT was undetectable in L. littoria (Table 2.3). Interestingly, levels of HOAD activity were relatively high in L. littoria, suggesting a signifiant capacity for O-oxidation.
HMS and HML catalyze the synthesis of acetoacetate from acetoacetyl-CoA in mammalian liver mitochondria. Neither enzyme was detectable in any of the snails under any conditions of tissue preparation. 30AT and AAT catalyze the pathway for ketone body oxidation in mammals. These enzymes were present in hepatopancreas of al1 three snails.
CS, a relative index of oxidative metabolism, was measured in a11 three snails, with highest CS activity seen in L. littoria. Ratios of activities of enzymes of intermediary metabolism to CS activity within a tissue can provide a qualitative indication of potential for flux through respective rnetabolic pathways . Ratios of enzymes of lipid and ketone body metabolism to CS activity were highest in S. ebdes
(Table 2.4), suggesting high rates of fany acid and ketone body metabolism in the hepatopancreas of this snail. These ratios were also relatively high in C. nemoralis, but much Iower in L. littoria, indicating a decreased reliance on fatty acid and ketone body metabolism in the marine snail.
The most parsimonious reconstruction of BHBDH shows this enzyme appearing independently in four different lineages which invaded freshwater or terrestrial environments (Fig 2.2). Within the bivalve subclass Heterodonta, BHBDH is present in Dreissenu and absent in Mercenaria, indicating divergence with respect to BHBDH expression within the subclass. A parallel situation exists in the gastropod order Mesogastropoda. where BHBDH is present in Pomacea and Vivipulrus, but absent in Linorina sp., indicating divergence in BHBDH expression within this order. DISCUSSION
Although conditions of tissue isolation and enzyme assay have been shown
here (Table 2.1) and in LeBlanc and Ballantyne (1993) io influence BHBDH activity,
no conditions were found in the present study under which the enzyme activity was
measureable in marine moliuscs. Isolation media constituents which resulted in
higher measured BHBDH activity in A. grandis did not induce detectable activity in
marine mollusc tissues. Based on these observations, we believe that lability of the
marine mollusc enzyme is probably not a factor affecting BHBDH detecrion in this
study . This conclusion is supported b y studies of rnitochondrial oxidation where
BHB was not oxidized by intact, respiring mitochondria isolated from various marine .
mollusc tissues (see Moyes et al., 1990 for review).
Sirnilarly, our inability to detect BHBDH in marine molluscs is not because
assay conditions do not adequately simulate the ionic strength of the intracellular environment. Marine mol luscs are osmoconformers and typically maintain
hemolymph and inuacellular osrnolarity an order of magnitude higher than non-
marine molluscs (Burton. 1983). Conespondingly, ionic strength of the intracellular
milieu is tenfold higher in marine molluscs than in freshwater and terrestrial species
(Burton, 1983). Some enzymes, like CPT, require minimum concentrations of
potassium andfor sodium chloride to function optimaily (Wood, 1973). Enzymes
adapted to function in a higher ionic strength intracellular envuonment may require
these ions in an assay medium to function optimally (Hochachka and Somero, 1984).
We addressed these characteristics of enzyme function by investigating the interactions of assay medium KCI and NaCl concentrations with BHBDH activity in a marine, a freshwater and a terrestrial gastropod. Sustained activity of BHBDH is seen in the fkeshwater (Fig. 2.1 (b)) and terrestrial (Fig . 2.l(a)) snails under conditions of increasing ionic suength and these same conditions fail to stimulate
BHBDH activity in the marine snail. This supports Our contention that the enzyme is indeed absent in marine molluscs.
The expression of BHBDH in molluscs does not correlate strongly with phylogeny. Mapping of the presence of BHBDH activity onto a gastropod and bivalve phylogeny is useful in illustrating a number of independent occurences of BHBDH within different lineages. Without exception, BHBDH expression occurs within taxa which inhabit freshwater and terrestrial environrnents. Within the bivalve subclass
Heterodonta and the gastropod order Mesogasûopoda, BHBDH is absent in marine species but present in closely related freshwater taxa. This suggesa that environment is a stronger determinant of BHBDH expression than phylogeny in the Mollusca.
Differential BHB DH expression may be indicative of fundamental d ifferences in metabolic organization of marine molluscs relative to their frcshwater and terrestrial relatives.
HMS and HML were undetectable in the hepatopancreas of the three species examined. In an earlier study (Meyer et al., 1986). HMS and HML activities were reported for hepatopancreas of the freshwater snail B. gkabmta. The assay of HMS used by Meyer et al. (1986) did not account for acetoacetyl-CoA lost through the combineci activities of AAT and acetyl-CoA hydrolase. This methodology overestimates HMS activity (Quant et ai., 1989). The assay of Meyer et al. (1986) gave measureable, though faIse, activity when used with our preparations (results not shown). The assay system used in our measurements, adapted fkom Quant et al.
(1989), elirninates background AAT activity through the regeneration of acetyl-CoA from acetyl phosphate and CoA via phosphouansacetylase. Using this assay, no measureable HMS activity could be detected in these gasaopods. Similarly, we detected no HML activity in hepatopancreas of any of the three snails.
Thus, the organization of ketone body metabolism in the gastropod hepatopancreas differs substantially from the marnmalian model. In mammais, HMS and HML catalyze ketone body synthesis, whereas 30AT catalyzes their oxidation.
30AT is present in very low activity in mammalian liver, where it is thought to participate in control of ketogenic flux. This enzyme occurs in higher activity in tissues which oxidize ketone bodies. In the present snidy, significant 30AT activity was detected in hepatopancreas of al1 three snails. In mammals, high 30AT activity and low or absent HMS and HML activities suggest a ketoiytic tissue. However, a ketogenic role for 30AT has been suggested in teleost fish (Phifips and Hkd, 1977), which are similar in having higher hepatic 30AT activities.
Ketone bodies synthesized in the hepatopancreas may be important metabolic substrates in peripheral tissues of freshwater and terrestrial rnolluscs. Relative to CS activity, enzymes of fatty acid oxidation (CPT, COT, HOAD) and ketone body metabolism (AAT, 30AT, BHBDH) are high in C. nemomlis and S. elodes, suggesting a high capacity for producing ketone bodies €tom fatty acids. Most molluscs store significant reserves of lipid in the hepatopancreas. Lipid stores in freshwater prosobranchs (Catterjee and Ghose, 1973) and the terrestrial mail
Cryptoiona ligulata (Krupanidhi et al., 1978) are comparable to gl ycogen stores.
Whole body (Lambert and Dehnel, 1974) and hepatopancreas (Heeg, 1977) lipid content have been shown to decrease following cessation of feeding in gastropods.
There is, however, no evidence for significant hemolymph transport of lipid as either free fats, acid (FFA) or triglyceride (TG) in rnolluscs (Allen, 1977). Similarly, no hemolymph albumin analogue has been demonstrated. Electrophoresis of hemolymph proteins (Allen, L977) provide no evidence of a hemolymph lipid binding protein in the marine chiton Cryptochtton stolleri. Furthermore, most mollusc extrahepatic tissues do not oxidize appreciable amounts of lipid (see Moyes et al., 1990 for review). Fatty acids and their carnitine esters are pwr oxidative substrates for rnitochondria isolated from gills and hemof marine bivalves and gastropods
(Ballantyne and Storey, 1983; Burcham et al., 1983; Burcham et al., 1984; Chih and
El1 ington, 1987). In contrast. mollusc hepatopancreas readily oxidize palmitoyl- wnitine (Bailantyne and Storey, 1984; Ballantyne and Moon, 1985). Thus, lipid stored in the hepatopancreas may be metabolically interconverted within this tissue, but not mobilized as FFA or TG in appreciable quantities. The peripheral utilization of this stored lipid carbon probably depends upon its conversion to more soluble and readily transported subsuates, like ketone bodies. The freshwater gastropod B. giabrahis appears to use this strategy. Hemolymph levels of BHB and Acac are high and decrease with starvation in this organism (Meyer et al., 1986). The lack of BHBDH in marine molluscs and generally lower activities of enzymes of ketone body metabolism relative to CS in L. littona suggests that the formation of ketone bodies by the hepatopancreas for peripheral oxidation is less important in these organisms. The physiological basis for this difference may be related to differences in the roles of protein and amino acid metabolism in these organisms, due to their strategy of osmoconforrnity. High intracellular concentrations of free amino acids are used by marine mollusc cells as osmolytes to balance intracellular and hemolymph osmolarities. There is some evidence that marine molluscs draw on this large pool of oxidizable amino acids extensively in cheir routine energy rnetabolism (see Moyes et al., 1990 for review). A number of snidies have shown that mitochondria irom many marine mollusc tissues are capable of oxidizing various amino acids at high rates, usually greater than pyruvate, fatty acyl- carnitines and ketone bodies, though Acac can be oxidized (Moyes et al., 1990).
Similarl y, transport of amino acids between tissues is probably greater in marine molluscs, which have several-fold higher levels of free arnino acids in their hemolymph than do freshwater and terrestrial mollusa (Bishop et al., 1983). Thus the distribution of hepatopancreas lipid carbon to peripherai tissues in marine molluscs may be primarily through production, transport and peripherai oxidation of arnino acids, whereas ketone bodies may fulfill this role in freshwater and terrestriai snails.
There is considerable evidence that emphasis on amino acid oxidation in energy metabolism may preclude the use of BHB as an oxidizable substrate. A number of mammalian studies (see Thompson and Wu, 1991 for review) have demonstrated inhibition of muscle protein and amino acid metabolism by ketone bodies. Goldstein et al. (1982) showed that glutamine metabolism in isolated rat kidneys and renal mitochondria is inhibited by BHB but not Acac. The authors attribute this decrease in glutamine oxidation to inhibition of GDH through a reduction in the NAD'INADH ratio. A number of studies (Ballantyne and Moon,
1985; Moyes et al., 1985; Ballantyne and Moyes, 1987a, 1987û; Gilles, 1987) support a central role for GDH in the catabolism of arnino acids in molluscs. A
GDH-Iinked transdeamination pathway plays a major role in regulating the enuy and exit of arnino acids from the inmacellular pool in euryhaline molluscs (Bishop et al.,
1981). Thus, a substantial and sustained flux through GDH would result in decreased flux through BHBDH, to the extent that it is adaptively advantageous to eliminate the latter pathway. Table 2.1. BHBDH activity under standard assay conditions using various homogenization buffers. Values are means f standard error, in pmollmin/g wet weight. Values in parentheses are numbers of independent measurements taken. Homogenization Buffer BHBDH activity
(1) 50 mM imidazole, pH 8.2 0.07 f 0.01 (9)
(2) 50 mM triethanolamine. 1 mM EDTA, 2 mM MgCI,, 0.08 f 0.01 (9) pH 8.2
(3) 50 mM triethanolamine. i mM EDTA, 2 mM MgC12, 0.09 i 0.01 (9) 30 mM mercapethand, pH 8.2
(4) 50 mM triethanolamine, 1 rnM EDTA, 2 mM MgCI,, 0.10 f 0.01 (8) 40% (viv) glycerol, pH 8.2
(5) 50 mM miethanolamine, 1 mM EDTA, 2 rnM MgCl,, 0.11 i 0.01 (9) 30 mM mercaptoethanol, 40% (vlv) glycerol, pH 8.2 Table 2.2. Activity of BHBDH in marine, fieshwater and terrestrial mollucs O GASTROPODA BHBDH ac tivity terrestrial Retinella wheatleyi Succinea ovallis Arion subfiscus (whole animal) Archachatina ventricosa Cepaea nemoralis freshwater Helisomu anceps (whole animal) Vivipanrs georgianus Pomacea bridgesi Stagnicola elodes marine Littorina littoria N.D. (5) Littorina saxatilis (whole animal) N.D. (5) Littorina obtusara (whole animal) N.D. (5) Buccinum undatum N.D. (5) Amaea testudinalis (whole animai) N.D. (5) Clione limacina N.D. (3) freshwater Anodonta grandis 0.11 * 0.01 (9) Dreissena polymorpha (whole animal) 0.14 f 0.04 (5) marine Mercenaria mercenaria hepatopancreas N.D. (5) gill N.D. (5) adductor muscle N.D. (5) mantle N.D. (5) Mytilus edulis hepatopancreas N.D. (5) gill N.D. (5) Crassostrea virginica N.D. (5) Placopecten mage llanims N.D. (5) Modio lus modiolus N.D. (5) Table 2.3. Activiries of enzymes of lipid and ketone body metabolism in C. nernoralis, S. elodes and L. Iittona hepatopancreas. Values are means f standard error of measuremenu made on five individuals. Enzyme activities are in pmollmin/g wet tissue weight. Enzvmes C. nem oralis S. elodes L. fittoria d --- - - TCA cycle Lipid metabolism HOAD 4.37 & 1.74 17.69 k 1.15 13.01 I0.64 COT 0.04 $r 0.01 0.1 1 f 0.06 N.D. CPT 0.06 0.03 0.18 k 0.01 0.06 i 0.02 ketone body metsbolism AAT 1.91 k 0.20 3.07 f 0.43 2.36 * O. 17 30AT O. 18 * 0.03 0.78 * 0.04 0.06 & 0.02 HMS N.D. N.D. N.D. HML N.D. N.D. N.D. Table 2.4. Hepatopancreas enzyme activities standardid with citrate synthase activity, in hepatopancreas of a terrestrial snail, Cepaea nemomlis, a freshwater snail, Stagnicola- elodes and a marine snail, Littonna littoria. Ratio C. nem oralis S. elodes L. littoria HOADICS 2.11 10.22 3 .O3 COT/CS 0.02 O. 07 - CPTICS 0.03 0, 11 0.0 1 AAT/CS 0.92 1.78 0.55 30AT/CS 0.09 0-45 0.0 1 BHBDHICS 0.63 0.55 - Fig. 2.1. Efface of sait concentration on the anivity of 8-hydroxybutyrate dehydrogenase fiom (a) the terrestriai snaü Cepaea nemomlfs and @) the fieshwater saail Stagnicola elodes. Data points an means of 5 separste determimions k S.E. CHAPTER 3 Importance of ketone bodies to the intermediary metabolism of the terrestriai sn ail, Archachafinu vertîricosu: evidence from enzyme activities published in Comparative Biochemisuy and Physiology, 1997, 117B: 197-201 Abstract The maximal activities of enzymes of ketone body rnetabolism and selected enzymes of other pathways of intermediary metabolism were measured in hem, hepatopancreas , kidney , man tle, and foot muscle of the terrestr ial p ulmonate gastropod Archachatina ventricosa. These rneasurements indicate the importance of ketone bodies in the energy metabolism of several aerobic tissues in this snail. Three enzymes which catalyze steps in the pathway which converts ketone bodies to acetyl- CoA are particularly high in the ventricle, which suggests that 0-hydroxybutyrate and acetoacetate are important energy substrates in this tissue. The organization of ketone body metabolism in Archacharina ventricosa tissues appears io differ from that of mammals in that no hydroxymethyIglutaryI-CoA synthase activity is detected in any tissue. However, ketogenesis may occur through other pathways including the readily reversible reactions of the ketolytic pathway. Our data suggest that the kidney may be a ketogenic tissue in these mails. INTRODUCTION The study of intermediary metabolisrn in molluscs has focused largely on carbohydrate and amino acid metabolism (Bishop et al., 1983; Livingstone and de Zwaan, 1983; Rees and Hand, 1993). Studies of substrate oxidation by mitochondria isolated from marine molluscs have shown that amino acids and carbohydrates are the preferred oxidative substrates of many tissues (Moyes et al., 1990). Fatty acids are not oxidized by most extrahepatic tissues at appreciable rates (Moyes et al., 1990) and ketone body use is restricted to acetoacetate (Acac) as marine molluscs lack the enzyme 8-hydroxybutyrate dehydrogenase (BHBDH)(Stuart and Ballantyne, 1996) and there fore cannot use 8-h ydroxybutyrate (BHB) . The emphasis in the intermediary metabolism of marine molluscs on oxidation of amino acids rnay be related to the presence of large innacellular amino acid pools, as these substrates are accumulated as osmolytes to maintain the cells of these organisms isosmotic with their environment. Amino acids are not important as inuacellular osmolytes in non-marine molluscs. Hemolymph and intracellular concentrations of amino acids in freshwater and terrestrial snails are an order of magnitude lower than those of marine molluscs (Burton, 1983). Generaily smaller arnino acid pools in non-marine mollusc cells may affect a diminished role for these substrates in oxidative metabolism. Amino acids could be replaced in part by ketone bodies as oxidizable substrates in fres hwater and terres trial snails. The hepatopancreas of non-mar ine snails has the enzyme BHBDH and thus can synthesize BHB, a more reduced and thus energy-rich oxidizable subsuate than Acac, for export to peripheral tissues (Stuart and Ballantyne, 1996). This ability of non-marine molluscs to utilize both ketone bodies could indicate a greater reliance upon them as energy substrates. Here we have examined the metabolic organization of tissues of the terrestrial snail A rchachatina ventricosa. To es tabl is h the relative importance of ketone bodies with respect to arnino acids, carbohydrates and lipids, we have rneasured maximal activities of key enzymes from the tricarboxylic acid cycle, the electron transport chain, and pathways of carbohydrate, amino acid, fatty acid and ketone body metabol ism . MATERIALS AND METEIODS Experimental animals A. venîricosa were maintained in an environmental chamber (Controlled Environments, Winnipeg, Canada), with a 12 hour light: dark cycle and a temperature of 23" * 2" C. Within the chamber, approximately 10 snails were housed in a two 50 cm by 40 cm by 15 cm plastic containers with ventilation holes cut into the lid. The bottom of the container was lined with sphagnum moss which was maintained moist by the periodic addition of water (humidity 30% f 5%).Snails were fed lettuce, apples, bananas and trout chow 3 times per week, with uneaten food removed and replaced with fresh food at these times. Sphagnum moss was replaced every 3 to 4 weeks. Under these conditions, snails did not estivate and were active on a daily basis. Tissue preparation As A. ventricosa are typically active at night, al1 animais were sacrificed in late evening and measurements of enzyme activities were determined immediately using the fresh tissues. Adult snails, weighing between 61.0 g and 125.9 g, were rapidly deshelled, tissues excised, weighed and placed in 10 volumes of ice-cold imidazole buffer (50 mM; pH 7.5). Tissues were homogenized with three ten second bursu of a Polytron PT10 unit (Kinematica Gmbh., Luzurn, Swiaerland) set at half power. Homogenates were centrifuged for 10 min at 10 000 g. The supernatants were used directly in enzyme assays. Enzyme assays Optimal substrate and cofactor concentrations were as described in Stuart and Bdlantyne (1996b). Maximal enzyme activitiu were determineci using a Hewlett Packard HP8452 diode array spectrophotometer (Hewlen Packard, M ississauga, Ontario, Canada), equipped with a thermostated ce11 changer maintaineci at 23" C with a Haake D8 circulating water bath (Haake Buchler Instruments Inc., Saddlebrook, NJ). Rates of reactions involving NADH were followed at 340 nm (millimolar extinction coefficient % = 6.22). Reactions involving 5,s'-dithio-bis- (2-nitrobenzoic acid) (DTNB)were followed at 4 12 nm (~~1,= 13.6). Acetoacetyl- CoA thiolase, 3-oxoacid transferase and hydroxymethylglutaryl-CoA synthase were measured by following the accumulation or disappearance of acetoacetyl-CoA at 303 nm (b, = 13.8). Cytochrome C oxidase activity was measured by following the oxidation of hilly reduced cytochrome C at 550 nm (E, = 29.5). Enzyme activities are expressed as units per gram of wet tissue weight where 1 unit equals 1 pmol of substrate converted to product per minute. Al1 enzyme assays were buffered in 50 mM imidazole, adjusted CO appropriate pH with KOH or HC1. Assay conditions were as follows: Oxïdative metabolism cytochrorne c oxidase (CCO)(E. C. 1.9.3.1): 50 pM reduced cytochrome C (ornitted for conaol), pH 8.0. citrate synthase (CS) (E.C. 4.1.3.7): 0.1 mM DTNB, 0.3 mM acetyl-CoA, 0.5 mM oxaloacetate (omitted for conuol), pH 8.O. Ketooe body metabolism aceioucetyl-CoA thiotase (MT)(E.C. 2.3.1 3): 10 mM MgCI,, 0.16 mM acetoacetyi-CoA, 0.2 mM CoA (ornitted for control), pH 8.0. 3-oxoacid-CoA-transferase (30AT) (E.C.2.8.3 S): 0.4 mM succinyl-CoA, 5 mM MgCI,, 5 mM iodoacetamide, LO mM acetoacetate (omitted for control), pH 8.0. J-hydron/bbutyrate dehydrogenase (BHBDH)(E. C. 1.1.1.30): 2 mM Dm,2 mM NAD, 40 mM DL-a-hydroxybutyrate (omitted for control), pH 8.0. hydro~erhyigIutu~i-CoAsynthase (HMS)(E. C. 4.1.3.5) (modi fied from Quant et al., '89): 10 mM MgCl,, 2 mM Dm,5 mM acetyl phosphate, 10 pM acetoacetyl- CoA, 100 pM acetyl-CoA and 10 units phosphotransacetylase (PTA) (both omitted for control), pH 8.0. hydroxymethyIgt1rra~I-CoA[vase (HM L) (E.C. 4.1.3.4) (mod ified from Ballantyne and Berges '91): 10 mM MgCI,, 0.2 mM NADH, 1 unit BHBDH, 0.2 mM HMG- CoA (omitted for control), pH 8.0. Lipid metaboüsm 3-hydroqacyi-CoA dehydrogenase (HOAD) (EX. 1.1.1.35): 0.2 mM NADH, O. 1 mM acetoacetyl-CoA (omitted for control), pH 8.0. camitine palmitoyl transferase (CPT)(E.C. 2.3.1.21): 0.1 mM DTNB, 50pM palrnitoyl-CoA, 100 mM KCl, 5 mM L-carnitine (omitted for control), pH 8.0. camitine octanoyl transferase (COT) (E.C. 2.3.1.21): 0.1 mM DTNE, 50 pM octanoyl-CoA, LOO mM KCI, 5 mM L-carnitine (omitted for control), pH 8.0. Amino acid metabolism glutamate dehydrogenase (GDH) (E.C. 1.4.1.2): 250 mM ammonium acetate, 0.1 mM EDTA, 0.05 mM NADH, 1 mM ADP, 7 mM a-ketoglutarate (aKG) (omitted for control), pH 8.0. glutamate-pymvate transaminase (GPT)(E. C. 2.6.1 -2): 200 mM L-alanine, 0.2 mM NADH, 0.025 mM pyridoxal-5'-phosphate, 1 unit iactate dehydrogenase, 10.5 mM aKG (ornitted for control), pH 7.5. glutamate-oxaloacetate transaminase (GOT)(E.C. 2.6.1.1): 0.2 mM NADH, 0.25 mM pyridoxal-5'-phosphate, 30 rnM aspartate, 1 unit MDH, 3.5 mM aKG (omitted for control), pH 7.5. Carbohydrate metabolism lactate dehydrogenase (LDH)(E.C. 1.1.1.27): 0.4 mM NADH, 1 mM pyruvate (omitted for control), pH 7.5. pymvate kinase (PK) (E. C. 2.7.1.40): 10 mM MgCI,, 0.4 mM NADH, 1.3 rnM ADP, 2 units LDH, LO mM phosphoenolpyruvate (omitted for control), pH 7.5. hexokinase (HK)(E.C. 2.7.1.1): 10 mM MgCl,, 0.16 mM NADP, 1 mM ATP, 2 uniu glucose-6-phosphate dehydrogenase, 10 mM D-glucose (omitted for control), pH 7.5. glucose-6-phosphate dehydrogenase (G6PDH) (E.C. 1.1.1 -49): 7 mM MgCl,, 0.4 mM NADP, I mM glucose-6-phosphate (omitted for control), pH 7.5. RESUL,TS and DISCUSSION Maximal enzyme activities in tissues of A. ventricosa (Table 3.1) provide a qualitative view of the metabolic organization of this terrestrial snail. Our main finding is that although carbohydrates play a major role in the intermediary metabolism of terrestrial gastropods (Table 3.1; Rees and Hand, 1990; Rees and Hand, 1993), ketone bodies also appear to be important subtrates in the rnetabolism of certain tissues. Ventricle The ventricle is highly aerobic, based upon high activities of CS and CCO, relative to other tissues. Chih and Ellington (1987) measured the maximal activities of enzymes of intermediary metabolism in the marine whelk Busycon coniranum at environmentai temperature and physiological pH values. Our measuremenu have been made under similar conditions, and thus a cornparison of enzyme activities from B. contrarium with those of A. ventricosa ventricle cm provide an indication of the relative importance of metabolic pathways in the two species. Citrate synthase (CS) activity in A. ventncosa ventricle is approximately three times greater than that measured in the marine gastropod Bicsycon contrarium (Chih and Ellington, 1987), suggesting that the former is a more metabolically active tissue. To compensate for this difference, we have made comparisons of enzyme activities between these organisms relative to CS activities (Table 3.2). These comparisons suggest that amino acids are less important as subsuates in the oxidative metabolism of the ventricle of the terresnial snail than in the marine gasrropod. The ratios of GDHKS, GPTKS and GOT/CS for the ventricle of the terrestrial snail A. ventricosa are as much as six- fold lower than those from the marine gastropod B. contrarium (Table 3.2). This apparent decreased reliance upon amino acids may be balanceci by an increased use of ketone bodies in the terrestrial snail ventricle. Enzymes of ketone body metabolism were not measured in the marine gastropod, but high activities of BHBDH, AAT and 30AT in A. ventricosa ventricle suggest a tissue that is poised to utilize ketone bodies at high rates. Similarly, activities of AAT and 30AT in A. ventricosa hem are greater than those found in other tissues (Beis et al., 1980) which have been shown to have a substantial reliance on ketone bodies as energy subsuates, such as the hearts of elasmobranch fishes (Moyes et al., 1990). BHBDH activity in A. ventricosa ventricle is more chan an order of magnitude greater than that measured in elasmobranch hem, and several times greater than values fkom mammalian hearts (Beis et al., 1980). Given the lower metabolic rate of terrestrial snails compared with mammals, ketone bodies. especially BHB, are Iikely very important energy substrates in A. ventricosa ventricle. The ventricle of this terrestrial gastropod is thus unique in the animal kingdom in its highly developed capacity for ketone body utilization. The high capacity for oxidation of ketone bodies in the ventricle contrast with the apparent low capacity for fatty acid oxidation in this tissue. A. ventricosa ventricle appears similar to that of marine molluscs (8allantyne et al., 1981; Bailantyne and Storey, 1983) in that enzyme activities suggest that the utilization of fatty acids as energy subsuates is minimal. Mitochondria isolated from the marine gastropod, B. contrarium , venuicle do not oxidize pdmitoy l-L-carnitine, palmitoyl- CoA or sodium oleate (Chih and Ellington, 1987). In A. ventricosri, HOAD activity is relatively low, COT activity is near the limits of detection and CPT activity is below detectable levels . Wh ile activities of membrane-bound enzymes 1ike COT and CPT may be underestimated due to loss from the homogenate in membrane fragments pelleted during centrifugation, our recovery of substantial activity of CC0 can be taken as an indication of a significant recovery of mitochondrial membrane. An examination of COT and CPT activities relative to CC0 activity in various tissues reveals that these activities are low in ventricle compared to kidney. This suggests that car ni tine-dependendent pathways of long- and medium-chah fatty acid catabolisrn are not likely to be important in the energy metabolism of the ventricle. While CS activity in the ventricle is approximately five-fold greater than in kidney, HOAD activity is less than double that of kidney. This also suggests that a relatively low proportion of the aerobic metabolism of ventricle is fuelled by fatty ac ids . High activities of enzymes of carbohydrate rnetabolism (Table 3.1) indicate that carbohydrates are also important energy substrates in the ventricle of this terresuial snail. The ratios HKfCS and PKfCS fiom A. ventricosa ventricle are sirnilar to those from the marine snail B. contrariun (Table 3.2), which utilizes both carbohydrates and arnino acids at high rates, based on studies of isolated mitochondria (Chih and El1 ington, 1987). Hepatopancreas Gastropods store significant amounts of lipid in the hepatopancreas (see Lambert and Dehnel , 1974; Heeg, L 977; Umezur ike and Iheanacho, 1983). Indeed. particularly high G6PDH activities in A. ventncosa hepatopancreas, relative to other tissues (Table 3. l), indicate a well-developed mechanism for the generation of cytosolic NADPH for lipogenesis. However, the metabolic fate of this stored lipid carbon is not clear. It may be transported intact to other tissues, such as the kidney which, based on enzyme activities, has a greater capacity for fatty acid oxidation. Whiie lipid transport proteins have been identified in hemolymph of a variety of mol luscs, includ ing gas tropods (Gar in and Pollero, 1995) their role in intermediary metabolism has not been studied. Though ketone bodies may be synthesized from either lipid or amino acid precursors, in mammals ketone bodies are produced primarily from the incornplete oxidation of fany acids in the liver (Robinson, 1980). Low or undectable COT and CPT activities in the hepatopancreas of A. ventricosa suggest a 1imited capacity for the carnitine-dependent oxidation of fatty acids to ketone bodies. Carnitine- independent pathways for fatty acid oxidation have been demonstrated in other invertebrates (Stevenson, L 968; Chamber lin, 1987) and may contribute to ketogenesis in the gastropod hepatopancreas. However, relatively low HOAD activities suggest that this may also be of only limited importance. The ratio of activities HOADlCS in A. ventricosa hepatopancreas is more than an order of magnitude lower than the corresponding values from livers of elasmobranch fsh (Moon and Mommsen, 1987). which have a high capacity for ketogenesis (Moyes et al., 1990). Enzymes of carbohydrate rnetabolism are present in high activity relative to enzymes of rnost other pathways in this tissue. Relative to CS. the activities of PK and LDH are higher in hepatopancreas than in ventricle or kidney. An active carbohydrate metabolism in the hepatopancreas of terrestrial snails has been well established (see Livingstone and DeZwaan, 1983). Kidney Based on maximal enzyme activities. A. ventncosa kidney has the best developed ability to utilize fatty acids of any tissue examined. Relatively high COT and undetectable CPT (Table 3.1) activities suggest a preference for medium-chain fatty acid oxidization through a car nithe-dependent pathway . This tissue may play a role in ketogenesis through the incomplete oxidation of medium-chain fatty acids, which could also fuel a significant proportion of energy rnetabolism in kidney. The kidney also has the highest levels of HML activity observed (Table 3.1). Though HMS and HML are known to catalyze ketogenesis in vertebrates, HMS is undetectable in A. ventricosa tissues and we have failed to detect this enzyme in other mails (Stuart and Ballantyne, 1996). However, HMG-CoA can be syntheswd Lhrough other pathways, including one leading from the amino acid leucine. Thus, kidney could synthesize acetoacetate through HML if HMG-CoA is produced from some arnino acid precursors. Ketone bodies could also be synthesized through the reversible reactions catalyzed by AAT, 30AT and BHBDH. In generai, these characteristics of the rnetabolic organization of kidney suggest a ketogenic tissue, though further studies are required to identify the origin of ketone bodies and the fate of lipid stores in A. ventricosa. Based on CS activities, kidney is a highly aerobic tissue. The oxidation of fatty acids may have a sparing effect upon carbohydrate oxidation, as HK, PK and LDH are lower relative to CS in this tissue than in hepatopancreas or ventricle. Mantle and Foot muscle Less aerobic tissues of A. ventncosa, including mantle and foot muscle, appear to utilize relatively low amounts of ketone bodies, based on lower activities of enzymes of ketone body metabolism relative to enzymes of other rnetabolic pathways. Foot muscle, in particular, shows little ability to utilize ketone bodies. Oxidation of fatty acids is also probably minimal as CPT and HOAD levels are low in both tissues. Intermediary metabolism in rnantle and foot muscle appears to be primarily supported by carbohydrates and am ino acids. Summary and Concl~isions The most suiking findings of the present study are the exceptionally high levels of enzymes of ketone body metabolism, especially BHBDH, in A. venzricosa vennicle. These activities are similar to those found in animals with much higher overall metabolic rates (Beis et al.. 1980). The apparent use of BHB as an important oxidative substrate in tissues of this terrestrial snail contrasts with the absence of BHBDH from marine snail tissues (Stuart and Ballantyne, 1996) and the inability of marine mollusc tissues to oxidize BHB (Ballantyne et al., 1981; Moyes et al., 1990). Based upon the present study and the presence of BHBDH in tissues of al1 terrestrial and freshwater molluscs examined to date (Stuart and Ballantyne, 1996). ketone body metabolism appears to contribute substantially to the energy metabolism of non- marine mo I luscs . Table 3.1 . Maximal activities of selected enzymes in tissues of Archachatina = hepatopancreas. ~eetext for abbreviations of enzyme names. heart he~ato kidney made foot oxidative metabolism CS CC0 lipid catabolism COT CPT HOAD ketone body metaboiism HMS HML AAT 30AT BHBDH amino acid metaboiism GPT GOT GDH carbohydrate metabolism HK 3.16 t 0.17 PIS 29.20 i- 4.71 LDH 24.42 I7-72 G6PDI-i 4.50 * 0.30 Table 3.2. Maximal enzyme activities indexed to CS activity in ventriclelheart of the terrestrial snail Archachatina ventricosa, the marine snail Busycon contrarïum and the marine elasmobranch Raja ennacea. See text for enzyme abbreviations. ------Ratio A, ventricosa B. contrarium ' R erinaced ' Data from Chih and Ellington (1987). ' Data from Moon and Mommsen (1987). Subceîiular organization of intermediary metaboüsm in the hepatoprncreas of the terrestrial saail, Cepaeu nemorolis: a cytosoüc B-hydroxybutyrate dehydrogenase published in The Journal of Experimental Zoology, 1996, 274:291-299 Abstract The subcellular distributions of key enzymes of the triwboxylic acid cycle, electron transport chain, ketone body, amino acid and carbohydrate rnetabolism were studied in the hepatopancreas of the terrestrial snail Cepaeu nernoralis. The presence of mitochondrial carnitine octanoyl transferase, carnitine palmitoyl transferase and 3- hydroxyacyl-CoA dehydrogenase indicate an active lipid catabolic pathway in this tissue. Activities of enzymes of ketone body metabolism are similar in magnitude to those of carbohydrate metabolism, suggesting an important metabolic role for ketone bodies in C. nemoralis. Two enzymes which, in mammals, catalyze the synthesis of ketone bodies, hydroxymethylglutaryl-CoA synthase and hydroxymethylglutaryl-CoA lyase were not detectable in Cepea hepatopancreas. The activity of 3-oxoacid-CoA transferase and a portion of acetoacetyl-CoA thiolase activity was found in the mitochondria. i3- hydroxybutyrate dehydrogenase is a cytosolic enzyme in this tissue, and preferentially oxidizes the L-isomer of B-hydroxybutyrate. A portion of glutamate dehydrogenase activity is also cytosolic. The subcellular organization of interrnediary rnetabolism is discussed with respect to adaptation to periodic anoxia. INTRODUCTION The study of intermediary metabolism in gastropod molluscs generally has been biaseâ toward carbohydrate-based metabolism in marine species (see Livingstone and de Zwann 1983 for review). Relatively Me is known about the participation of other energy substrates in the intermediary metabolism of gastropods, especially in non-marine species. In particular, the roles of lipids and ketone bodies have not been examined in detail. A single study by Meyer et al. (1 986) examined the metabolism of ketone bodies, acetoacetate (Acac) and 8-hydroxy butyrate (B HB) , in the fres hwater gas uopod Biomphaloria glabrata. High activities of enzymes of ketone body metabol ism in various tissues and hemolymph levels of Acac and BHB suggest a signifiant roie for ketone bodies in routine energy metabolism in B. glabrata. In the hepatopancreas of this freshwater mail, the enzymes of ketone body metabolism, including O-hydroxybutyrate dehydrogenase (BHBDH), are localized within the mitochondria (Meyer et al., 1986). In our studies of ketone body metabolism in the terrestrial snai! Cepea nemoralis, we found chat BHBDH, which in virtuaily al1 animals exists bound to the inner membrane of the mitochondrion, is cytosolic in hepatopancreas. We dius further examined the subcellular distribution of enzymes of ketone body metabolism in this tissue. To determine the relative importance of ketone body metabolism in the intermediary metabolism of this terrestrial gastropod, we also have measured the activities and subcellular distributions of enzymes of the tricarboxylic acid cycle, the elecaon transport chain, and carbohydrate, amino acid, lipid metabol ism in hepatopancreas. MATERIALS AND METEODS Expenmental animals C. nemoralis were collecteci on the University of Guelph campus in the spring of 1994. Snails were kept in a laboratory terrarium at 20' * 2' C for up to one week prior to experimentation and fed lettuce ad libitum. Humidity within the terrarium was kept between 50% and 100% with moist sphagnum rnoss. Tissue preparation Snails were deshelled, the hepatopancreas excised, intestine removed, and placed immediately in 3 mL of ice cold mitochondrial isolation buffer containing 20 mM N-12- hydroxyethyllpiperazine-Nt-[2-ethanesulfoicacid] (HEPES) and 100 rnM sucrose, pH 7.5. AI1 tissue preparation procedures were carried out on ice. Hepatopancreas was homogenized with 3 passes of a Potter-Elvejhern homogenizer with a teflon pestle attached to a drill press, operating at < 100 revolutions per minute. The homogenate was centrifuged at 200 g for 10 minutes to remove cell debris and nuclei. A 1 mL sample of this supernatant, the "wholehomogenate", was taken and aeated as described below. The remaining homogenate was centrifuged at 10,000 g for 10 minutes. The resultant pellet, the "initochondriai fraction" was resuspended in 1 mL of mitochondrial isolation buffer and treateâ as described below .This fraction contained virtuall y al1 rnitochondr ia, based on activities of cytochrome C oxidase, and citrate synthase. The supernatant frorn this step was centrifuged at 40,000 g for 10 minutes to remove non-soluble materials and yieldexi a pellet too smdl to use for enzyme determinations. The resultant supernatant was wnsidered the "cytoplasrnic Fractionw. A 1 mL sample of this fraction was collected. Al1 Fraction sarnples were sonicated with 5 bursts of 5 seconds at 60% power output, 25 watts, on a Vibra-Cell sonicator (Sonics & Materials Inc., Danbury, CT, USA) and subsequently used in enzyme assays. Enzyme assays Optimal substrate and cofactor concentrations were determined for al1 inzymes assayed with the supernatant of a crude homogenate of C. nernoruIis hepatopancreas. For these determinations, hepatopancreas of deshelled snails were excised and placed in 2 mL of ice-cold 50 mM imidazole and homogenized with 3 passes of a Polyuon PT10 tissue homogenizer (Kinematica Gmbh., Luzurn, Switzerland) and centrifuged at 10,000 g Maximal enzyme activities were deterrnined using a Hewlen Packard HP8452 diode array spectrophotometer (Hewlett Packard, Mississauga, Ontario, Canada), equipped with a thermostated ce11 changer maintained at 20" C with a Haake D8 circulating water bath (Haake Buchler Instruments Inc., Saddlebrook, NI). Rates of reactions involving NADH were followed at 340 nm (millimolar extinction coefficient % = 6.22). Reactions involving 5,s'-dithio-bis-(2-nitrobenzoic acid) (DTNB)were followed at 4 12 nm (millimolar extinction coefficient s4,, = 13.6). Acetoacetyl-CoA thiolase, 3-oxoacid tramferase and hydroxyrnethylgIutaryI-CoA synthase were rneasured by following the accumulation or disappearance of acetoacetyl-CoA at 303 nm (millimolar extinction coefficient = 13.8). Enzyme activities are expressed as uniu per gram of wet tissue wet where 1 unit equals 1 pmol of subsuate converted to product per minute. For convenience of cornparison, enzyme activities measured in isolated mitochondria also were expressed as activities per tissue wet weight. All enzyme assays were buffered in 50 mM imidazole, adjusted to appropriate pH with KOH and HCI. The following optimal assay conditions were determined with respect to substrate and cofactor concentrations: TCA cycle malate dehydrogenase (MDH) (E.C. 1.1.1.37): 0.15 mM NADH, 5 mM oxaloacetate (omitted for control), Ph 7.5. citrate synthase (CS) (E.C.4.1.3.7): 0.1 mM DTNB, 0.3 rnM acetyl-CoA, 0.5 mM oxaloacetate (omitted for control), pH 8.0. Electron transport chah cytochrome c oxidase (CCO)(E.C. 1.9.3.1): 50 pM reduced cytochrome C (omitted for control), pH 8.0. Lipid metabolism 3-hydroxyacyl-CoA dehydrogenase (HOAD)(E.C. 1.1.1.35): 0.2 mM NADH, 0.1 mM acetoacetyl-CoA (omitted for control), pH 8.0. carnitine palmitoyl transferase (CPT) (E.C. 2.3.1.21): 0.1 mM DTNZ, 50pM palmitoyl-CoA, 100 mM KCI, 5 mM L-carnitine (omitted for control), pH 8.0. carnitine octanoyl tramferase (COT) (E.C.2.3.1.2 1): 0.1 mM DTNB,50 pMoctanoy l- CoA, 100 mM KCI, 5 mM L-carnitine (omined for control), pH 8.0. malic enzyme (ME) (E.C. 1.1.1.40): 1.O mM MgCl,, 0.4 mM NADP, 1 mM of either D- or L- malate (individually) (omitted for control), pH 7.5. ATP-citrate lyase (ATPCL) (E.C. 4.2.3.8): 20 rnM MgCl,, 0.1 mM NADH, 1 mM D'TT, 0.4 mM CoA, 10 rnM ATP, 25 units MDH. 20 mM citrate (omitted for control), pH 8.0. isocitrate dehydrogenase (ICDH)(E.C. 1.1.1.42): 5 mM MgCI,, 0.4 mM nicotinamide adenine diphosphate (NADP), 0.5 mM threo-D,L,isocitrate (om itted for connol), pH 8.0. Ketone body metabolism acetoacetyl-CoA thiolase (AAT) (E.C. 2.3.1.9): lOmM MgCl,, 0.16 mM acetoacetyl- CoA, 0.2 mM CoA (omitted for control), pH 8.0. 3-oxoacid-Co A-transferase 0.4 mM succinyl-CoA, 5 mM Mg Cl,, 5 mM iodoacetamide, 10 rnM acetoacetate (omitted for control), pH 8.O. BHBDH (E.C. 1.1.1.30): 2 mM DTT, 2 mM NAD, 40 mM DL-O-hydroxybutyrate (omitted for control), pH 8.0. hydroxymethylgIutitry1-CoA synthase (HMS)(E.C. 4.1.3.5) (modified from Quant et al., 1989): 10 mM MgCI,, 2 mM DTï, 5 mM acetyl phosphate, 10 pM acetoacetyl-CoA, 100 pM acetyl-CoA and 10 units phosphotransacetylase (PTA) (both omitted for control), pH 8.0. hydroxymethylglutary 1-CoA lyase (HML)(E. C. 4.1 -3.4) (modified from Ballantyne and Berges 1991): 10 mM MgCI,, 0.2 mM NADH, 1 unit BHBDH, 0.2 mM HMG-CoA (omitted for control), pH 8.0. Amino acid metabolism glutamate dehydrogenase (GDH)(E.C. 1.4.1.2): 250 mM ammonium acetate, 0.1 mM EDTA, 0.05 mM NADH, 1 mM ADP, 7 mM a-ketoglutarate (aKG) (omitted for control), pH 8.0. glutamate-pyruvate transaminase (GPT) (E.C. 2.6.1.2): 200 mM L-alanine, 0.2 mM NADH,0.025 mM pyridoxal-5'-phosphate, 1 unit lactate dehydrogenase, 10.5 mM aKG (omitted for control), pH 7.5. glutamate-oxaloacetate transaminase (GOT) (E.C.2.6.1.1): 0.2 mM NADH, 0.25 mM pyridoxai-S'-phosphate, 30 mM aspartate, 1 unit MDH, 3.5 mM aKG (or;.i:%d for control), pH 7.5. Carbohydrate metabolism lactate dehydrogenase (LDH) (E.C. 1.1.1.27): 0.4 rnM NADH, 1 mM pyruvate (omitted for control), pH 7.5. pyruvate kinase (PK) (E.C.2.7.1.40): 10 mM MgCI,, 0.4 mM NADH, 1.3 mM ADP, 2 uni@ LDH, 10 mM phosphoenolpyruvate (omitted for control), pH 7.5. hexokinase (HK) (E.C.2.7.1.1): 10 mM MgCI,, 0.16 mM NADP, 1 mM ATP, 2 units glucose-dphosphate dehydrogenase, 10 rnM D-glucose (omitted for control), pH 7.5. glucose-6-phosphate dehydrogenase (GQDH) (E.C. 1.1.1.49): 7 mM MgCl?, 0.4 mM NADP, 1 mM glucose-6-phosphate (omitted for control), pH 7.5. a-glycerophosphate dehydrogenase (aGPDH) (E.C. 1.1.1 A): 0.2 mM NADH,0.4 mM dihydroxyacetone phosphate (omitted for control), pH 8 .O. The maximal activities of BHBDH with the D- or L- stereoisomers were determined. BHBDH activity with each isomer is the mean of 6 measurernents, determined on homogenates prepared as described above. RESULTS The levels of enzymes of various metabolic pathways and their proportional distribution between mitochondria and cytosol provide a qualitative view of the rnetabolic organization of C. nemoralis hepatopancreas. The activities of ail enzymes measured in the hepatopancreas are presented in table 4.1. The percentage of this activity recovered in the summed mitochondrial and cytosolic fractions, and the percentage of this amount appearing in either the mitochondrial or cytosolic fractions individually, are presented in table 4.2. TCA cycle CS ad MDH activities are detectable in the mitochondrial fraction of the hepatopancreas. Though CS appears to be exclusiveIy mitochondr ial (9 1% in mitochondrial fraction), a substantial proportion of MDH activity is cytosolic (82%). Electron transport chah CC0 activity is exclusively rnitochondrial, with 98% of its activity in this fraction. Lipid metabolism The enzymes involved in the transport and 8-oxidation of fatty acids are substantially mitochondrial. CPT and COT, whîch catalyze the carnitine-deyendant transport of long and medium chah fatîy acids, respectively, into the mitochondrion, are not detectable in the cytosolic fraction. HOAD catalyzes the 8-oxidation of fatty acids. HOAD is primarily a mitochondrial activity, with 16.6 % of recovered activity appearing in the cytosolic fraction. Enzymes which can provide substrate and reducing equivalents (NADPH) to fuel cytoplasmic 1ipid synthesis were examined. Malic enzyme, w hich catal yzes the formation of pyruvate fiom malate with the concomitant generation of NADPH, is undetectable in al1 cellular fractions. NADP+-dependantICDH also provides NADPH for cytosolic lipid synthesis. The relatively high ICDH activity was primarily cytosolic, with a small proportion of total activity detected in the mitochondrial fraction. ATPCL catalyzes what is believed to be an important step in lipid synthesis in some biological systems (Newsholme and Leech 1983), the production of acetyl-CoA from citrate produced in and exported frorn the mitochondria. ATPCL was present in approximately equal proportions in the rnitochondrial and the cytosolic fractions. The summed activities in these two fractions is about twice that measured in the whoie homogenate. The bais for this is not known. Ketone body metabolism Activities of BHBDH, AAT and 30AT are of the same order of magnitude as those of key enzymes (PK and HK) of carbohydrate metabolism. BHBDH, normally a rnitochondriai membrane-bound enzyme, was exclusively cytosolic in C. nemarulis hepatopancreas, with no detectable mitochondrial activity. 30AT, which catalyzes the reversible oxidation of acetoacetate, was rnitochondrial, with less than IO % appearing in the cytosolic fraction. AAT produces acetoacetyl-CoA from acetyl-CoA. AAT activity is Iargely cytosolic, with about 30% of total activity appearing in the mitochondrd fraction. HMS and HML catalyze the synthesis of acetoacetate from acetoacetyl-CoA in mammals, through the transient production of hydroxymerhylglutary1- CoA. Neither of these enzymes was detectable in any cellular fraction. The validity of the assays was verified using rat and mouse liver (data not shown). Amino acid metabolism The amino acid transaminases, GPT and GOT, have high proportions of cytoplasmic activity, with lesser activities appearing in the mitochondrial fraction. Activity of GOT is greater than GPT. GDH, which catalyzes the oxidative deamination of glutamate to a-ketoglutarate, is approximately equal l y disaibuted between mitochondrial and cytosolic fractions. Carbohyârate metabolism HK, which catalyzes the first reaction in the intracellular metabolism of glucose, is present in the cytosol, with minor contamination of the mitochondrial fraction. PK, which catalyzes what is considered to be one of the rate-limiting steps in glycolysis, the production of pyruvate from phosphoenolpyruvate, is detectable exclusivel y in the cytosol. Virtually no LDH activity is seen in the mitochondrial fraction. All LDH activity measured in the whole hornogenate is recovered in the cytosol. aGPDH is involved in the cytoplasmic amof the a-glycerophosphate shuttle, which panicipates in balancing mitochondrial and cytoplasmic redox. The activity of this enzyme is also vhally exclusively cytosolic. G6PDH catalyzes the fust committed reaction of the pentose phosphate pathway, and generates NADPH which can be used to drive lipid synthesis. G6PDH is entirely cytoplasmic in C. nemoralis hepatopancreas. Stereospecificity of BHBDH for D or L-BEîB Maximal BHBDH activity with L-BHB was optimized at a L-BHBconcentration of 20 mM. The mean activity with the L- isomer was 1.758 f 0.235 pmoI/min/g wet tissue weight. Optimal BHBDH activity with D-BHB was achieved with 40 mM D- BHB. Mean activity with the D- isomer was 0.515 f 0.039 pmollminlg wet tissue weight. DISCUSSION As the major storage site for lipid (Voogt, 1983). the hepatopancreas plays an important role in the energy metabolism of terrestrial snails (Heeg, 1977; Krupanidhi, 1978). Although the tissue itself is capable of lipid catabolism as we have demonstrated, it is unclear if lipid stored in the hepatopancreas can be transported to other tissues. The most metabolically active transport form of lipid in many organisms are non-esterified fatty acids (NEFAs). nie very Iow solubility of NEFAs in aqueous solutions requires that they be transported bound to a carrier. There is no evidence of a fatty acid binding protein in molluscs suggesting that the transport of fatty acids in hemolymph may not conaibute substantially to mollusc energy rnetabolism (Allen, 1977). Lipid carbon stored in hepatopancreas may thus be transported in another, more soluble, form. Partial oxidation of lipid in the hepatopancreas and conversion to more freely soluble metabolic inter mediates, 1ike ketone bodies, which can be readil y exported to peripheral tissues may be an important strategy for distribution of stored Iipid reserves in Cepaea. This strategy appears to be employed by elasmobranch fish (Zamrnit and Newsholrii~,1979) which use ketone bodies extensively, and also lack NEFA carrier proteins (Lauter et al., 1968). Based on enzyme activities, the metabolism of ketone bodies in the hepatopancreas is substantial. In Cepea hepatopancreas, activities of enzymes of ketone body metabolism are of the same order of magnitude as those of carbohydrate metabolism. In most other animals these activities are only about 10% of activities of key enzymes of carbohydrate metabolism (see Newsholme and Leech, 1983). Similady, the concentration of BHB in fed B. gïabmta hemoiymph (0.6 mM) (Meyer et al., 1986) is comparable to that of glucose (0.62 mM) (see Livingstone and de Zwaan, 1983). This suggests that ketone bodies are possibly as important as carbohydrates in the intermediary metabolism of freshwater and terrestrial gastropods. The data of Meyer et al. (1986) support a role for ketone bodies in routine energy metabolism in B. glubruta, as opposai to their more specialwd role in starvation in most other animals (see Newsholme and Leech, 1983). The organization of the pathways of ketone body metabolism in C. nernoralis differs from that of mammals in several respects. In most mammals, ketogenesis occurs primarily through reactions catalyzed by HMS and HML, whereas oxidation occurs through 30AT (Newsholme and Leech, 1983). HMS and HML are undetectable in both the mitochondr ial and cytoplasmic fractions of Cepaea hepatopancreas indicating that this pathway of ketone body synthesis is not present. The lack of HMS and HML in Cepaea hepatopancreas contrasu with the results of Meyer et al. (1986) who measured HMS and HML activities in the hepatopancreas of the freshwater snail Biomphalana glabrafus. We assayed HMS using the method of Quant et al. (1989) which effectively prevents the loss of acetoacetyl-CoA via the combined actions of acetyl-CoA hydrolase and AAT, by vinually eliminating the accumulatio~of CoA through an acetyl-CoA regenerating system. The assay used by Meyer et al. (1986) did not account for acetoacetyl-CoA lost through the latter pathway and therefore may have substantially overestimated the activity of HMS in B. glabruta. Ketone body synthesis and degradation do not depend upon the presence of HMS and HML. Acac can also be synthesized through the reversal of the reaction catalyzed by 30AT (Fig. 4.1). Correspondingly, the levels of 30AT are 3-fold higher in Cepaea hepatopancreas than in rat liver (0.21 prnollmidg) (Zammit and Newsholme, 1979). High 30AT activity in Cepea hepatopancreas may be related to a major role for this enzyme in ketogenesis. A ketogenic role for 30AT has been suggested in fish (Philips and Hird, 1977), where activity of this enzyme is also quite high, for exûziple 12 pmollminlg in bass (Zammit and Newsholme, 1979). There is some evidence that 30AT is involved in ketogenesis in elasmobranch fish. In starved elasmobranchs, an increase in blood ketone body levels is parallelled by a threefold increase in the activity of Iiver 30AT (Zarnm i t and News holme, 1979). Thus, ketogenesis in Cepeu hepatopancreas must be catalyzed by AAT, 30AT and BHBDH (Fig. 4.1) without the involvement of HMS or HML Another important finding of the present study is the subcellular distribution of BHBDH. BHBDH activity is exclusively cytosolic in C. nemoralis hepatopancreas (Fig. 44, whereas in most animals it is a mitochondrial enzyme, found associated with the inner mitochondrial membrane (Wang et al., 1988). A cytosolic BHBDH is, however, found also in ruminant Iiver and kidney (see Zamrnit, 1990 for review). The physiologicai basis for the shift of BHBDH frorn mitochondria to cytosol in ruminant liver and kidney has not been thoroughly investigated. The only rationale for the cytoplasmic BHBDH has been provided by Zarnmit (1990) who suggests that cymsolic BHBDH may promote gluconeogenesis from pyruvate under ketogenic conditions. Gluconeogenesis is the main source of glucose and glycogen in ruminants, as virtually no glucose is produced by ruminant gut (Heitmann et al., 1987). Zammit (1990) speculates that since Acac is a better exchange ion for pynivate (Paradies and Papa, 1975), export of Acac, rather than BHB, from the mitochondria would maxirnize pyruvate import. Furthermore, the shifi of BHBDH from rnitochondria to cytosol results in a more reduced intramitochondrial and less reduced cytosolic NADH: NAD' ratio under ketogenic conditions (ammit, 1990). This would promote the export of malate, thus lowering the concentration of intramitochondrial oxaloacetate and minimizing its use for the synthesis of aspartate and phosphoenolpyruvate. Cytosolic regeneration of malate and its subsequent oxidation by MDH lads to a more highly reduced NAn:NADH couple in the cytosol serving to drive gluconeogenesis. Depletion of rnitochondr ial oxaloacetate would limit citrate synthesis resulting in an accumulation of acetyl-CoA. A buildup of acetyl-CoA would promote ketone body production. Gluconeogenesis in Cepaea hepatopancreas, in the fed state, is probably less important than in ruminants. Gastropods possess a variety of carbohydrases, and transport of sugars from the gut has been demonstrated (Livingstone and de Zwaan, 1983), suggesting that, uniike ruminants, gut is an important source of glucose in gastropods in the fed state. Gluconeogenesis is, however, very important during recovery from anoxia when glycogen reserves are depleted (Livingstone and DeZwaan, 1983). Cytosolic BHBDH may play a role in promoting gluconeogenesis in these situations. Most terresnial snails have a well developed anaerobic metabolism (Livingstone and DeZwaan, 1983), adapted to a life-history which includes prolonged anoxia during hibernation (Oudejans and van der Horst, 1974), when burrowing snails can become immersed in water or ice (Livingstone and DeZwaan, 1983). Cyclic hypoxia can occur during estivation, with intermittent bouts of aerobic and anaerobic metabolism (Barnhart and McMahon, 1987). During per iods of anoxia, terrestrial gasuopods deplete the ir endogenous glycogen reserves and accumulate alanine and large amounts of lactate in die hepatopancreas (Churchill and Storey , 1989). Upon resumption of aerobic metabolisrn, both anaerobic end products are reoxidized to pyruvate, which then may be used to regenerate glucose and glycogen (Fig . 4.1). Ketogenesis in Cepuea hepatopancreas fiom stored 1ipid (Fig. 4.1) occurring in parallel with the reincorporation of pyruvate into carbohydrates would stimulate this process in two ways. First, b y providing intramitochondrial Acac for export wh ich would stimulate pyruvate uptake through the rnitochondrial monocarboxylate wrier as described above. Secondly, cytosolic reduction of Acac to BHB would aid in maintenance of cytosolic redox through reoxidation of NADH generated by reformation of pyruvate from lactate, via LDH, and from alanine, through the combined actions of AlAT and cytosolic GDH (Fig. 4.1). Cytoplasmic redox might otherwise become too highly reduced following anaerobic bouts, as hepatopancreas rnust reoxidize endogenous pools of alanine and lactate, as well as pools of these substrates transported from other tissues via the hemolymph (Livingstone and DeZwann, 1983: Churchill and Storey, 1989) (Fig 4.1). Cytosolic BHBDH may thus aid in the transition from anaerobic to aerobic metabolism in Cepaea hepatopancreas, Interestingly, a signifiant capacity for anaerobic lipid synthesis has been demonstrated in C. nemoralis (Oudejans and van der Horst, 1974). Sorne authors (Van der Horst, 1974; Zs-Nagy, 1979, DeZwaan, 1983) have suggested that fatty acids may function as terminal electron acceptors in mollusc hepatopancreas dur ing anaerob iosis. In hepatopancreas of estivating giant Afr ican snails Achatino, free fatty acid content fluctuates over 24 days of estivation, showing periods of increase and decrease (Umezerike and Iheanacho, 1983). These data support a biphasic role for lipids in estivation, with anaerobic synthesis providing an electron sink and aerobic partial oxidation ieading to the sy nthes is of ketone bodies. The cytoplasmic BHBDH of C. nemoralis hepatopancreas also is similar to that of ruminant liver in that it preferentially oxidizes the L- stereoisomer of BHB (Williamson and Kuenzel, 1971). The significance of this is not clear. Other molluscan enzymes display unusual stereospecificities. Molluscs possess a D-lactate specific LDH, whereas most LDH's are specific for L-lactate. Long (1976) suggests that there is no adaptive advantage to using either the D- or L- isorner of lactate and that a mutative change to a single arnino acid may result in a change in stereospecificity cf LDH. Similarly, L-BHB may not be physiologically significant in the energy metabolism of C. nemomlis, but rather the result of non-specifity of the cytosolic BHBDH. No studies have investigateû the occurence of L-BHB in ruminant blood. A necessary step to clarify the physiological importance of this phenornenon is the measurement of hemolymph levels of both stereoisomers of BHB. Other aspects of the subcellular organization of C. nernoraiis hepatopancreas suggesting adaptation to episodic anoxic conditions include the signifiant proportion of GDH activity which occurs in the cytosol. The predominantly anaerobic rat tapeworm, Hylnenolepis diminuta (Cestoda), has an exclusively cytosolic GDH (Mustafa et al., 1978) with lactate and alanine among the accumulated anaerobic end products (Mustafa et al., 1W8). These authors have suggested that the shift of GDH from mitochondria to cytosol may facilitate the maintenance of cytosolic redox in H. diminuta. The utility of a cytosolic GDH may be best understd through comparison of terrestrial snail anaerobic processes with those of marine molluscs. In marine bivalves, cytosolic redox is maintained through coupling the generation of NADH from glycolysis to: (1) NADH oxidation during the reduction of pyruvate to alanopine and (2) NADH oxidation during the aspartate-linked reduction of oxaloacetate to malate (Hochachka, 1980). Large intracellular aspartate pools (12 pmollg wet weight in Busycon contrarium ventricle) (Ellington, 1981) allow this strategy in marine gastropods and bivalves (DeZwann, 1983). However, terrestrial snails must use alternative strategies, as they lack substantial aspartate pools (1.72 and 0.85 pmollg wet weight in O. Iactea foot and hepatopancreas, respective1y) (Churchill and Storey , 1989). Anaerob ic 0. lucieu hepatopancreas appear to qu ickly use up aspartate reserves , but continue to accumulate alanine and lactate throughout 14 h of anoxia (Churchill and Storey, 1989). The subcellular distribution of enzymes in Cepaea hepatopancreas suggest that thp halance of cytosolic redox in this tissue is achieved through the reduction of pyruvate to lactate via LDH and of pyruvate to alanine, via a cytosolic AlAT and GDH couple (Fig. 4.2). This couple would not deplete glutamate or aKG pools, since it provides cyclic fluxes of these metabolites. This pathway resuln in the net consumption of NH, (Walsh and Henry, 1991), which may be of further benefit in providing a proton sink, reducing the rate of acidification of the cytosol. The role of aspartate reduction in redox balance is probably restricted to early stages of anoxia. This organization is further suggested by the high cytosolic activity of AlAT relative to mitochondrial activity as is seen in other good facultative anaerobes (see Hochachka, 1980). A large proportion of AlAT activity is cytosolic also in 0. lactea hepatopancreas (Sollock et al., 1979). Thus, some terrestrial snails rnay have replaced the aspartate couple used by marine molluscs with a cytosolic AIATIGDH couple (Fig. 4.1), which maintains redor balance over longer periods of anoxia than might be possible through reduction of a limited aspartate pool. In summary, the organization of ketone body metabolism in the hepatopancreas of the terrestrial snail C. nemoralis differs From that of freshwater snails and most mammals, and is similar in several respects to that of ruminants. Based on our observations we suggest there are links between ketone body metabolism and gluconeogenesis and amino acid metabolism in this tissue. The subcellular distribution of enzymes of intermediary metabolism in this tissue are consistent with adaptations to periodic anoxia, which typify terrestrial snail life-history . Further stud y of the tissue distribution of these enzymes and hemolymph concentrations of substrates are requùed to define the roles of ketone bodies in the intermediary metabolism of these organisms. Table 4.1. Maximal activities of enzymes in C. nemorulis hepatopancreas . Activities (pmol/min/g wet tissue weight) are means of 6 separate determinations, f SE. N.D. = no t detected . Enzyme w hole homogenate act iv i ty TCA cycle CS ICDH MDH oxidative metabolism CC0 2,676 * 0.432 ketone body metabolism HMS N.D. HML N.D. AAT 3.909 * 0.810 30AT 0.795 f 0.202 BHBDH 0.825 * 0.096 iipid metabolism COT 0.032 * 0.020 CPT 0,149 * 0.043 HOAD 0.296 * 0.051 ATPL 0.067 * 0.025 ME N.D. amino acid metabolism GDH GPT GOT cubohydi.ate metabolism PK HK LDH G6PDH a-GPDH Table 4.2. Recovery of whole homogenate enzyme activities in sumrned mitochondrial and cytosolic fractions and distribution of enzyme activities between mitochondrial and cytosol ic fractions. Values represent average % recovery of 6 separate determinations. Enzyme % activity recovered in % of recovered activity % of recovered summed fiactions in mitochondr id activity in cytosolic fraction fraction TCA cycle CS 104.9 f 9.1 90.6 k 2.1 9.4 i2.1 ICDH 87.8 * 9.2 22.9 I 0.9 77.1 & 0.9 MDH 76.2 * 11.9 17.7 & 1.0 82.3 I 1.0 oridative metabolism CC0 98.9 * 6.6 98,2 i 6.6 1.8 * 0.3 ketone body metabolism HMS N.D. N.D. HML N.D. ND. AAT 133.7 * 17.6 33.3 i 6.5 66.7 * 6.5 30AT 108.7 * 11.9 92.8 * 1.4 7.2 & 1.4 BHBDH 150.6 * 37.0 0.0 100.0 lipid metabolism cm 100.0 100.0 0.0 CPT 57.0 * 18.0 100.0 0.0 HOAD 107.8 f 29.0 83.4 * 8.0 16.6 & 8.0 ATPL 200.4 * 65.1 74.3 I11.5 25.7 * 11.5 ME N,D. N.D. N.D. amino acid metsbolism GDH 129.2 f 30.7 59.4 f 8.4 40.6 * 8.4 GPT 100.1 * 3.8 28.6 & 3.1 71.4 * 3.1 GOT 96.3 * 7.8 23.1 f 1.3 76.9 * 1.3 carbohydrate metaboiism PIC 117.0 * 10.9 0.0 100.0 HK 84.9 * 14.6 8.8 * 4.2 90.2 * 4.8 LDH 103.5 * 15.4 3.4 I0.9 96.6 k 0.9 G6PDH 92.4 * 27.5 10.5 f 0.3 89.5 * 0.3 aGPDH 88.4 I7.2 2.1 * 0.6 97.9 I0.6 Abbreviations used in figures: AAT = acetoacetyl-CoA thiolase aKG = alpha-ketoglutarate Acac = acetoacetate BHB = 8-hydroxybutyrate BHBDH = fi-hydroxybutyrate dehydrogenase CPT = carnitine paimitoyl transferase GDH = glutamate dehydrogenase GOT = glutamate-oxaloacetate transaminase GPT = g lutamate-pyruvate transaminase HOAD = L-3-hydroxyacyl-CoA dehydrogenase LDH = lactate dehydrogenase MDH = malate dehydrogenase 30AT = 3-oxoacid-CoA transferase Fig . 4.1. Proposed organization of aerobic metabolisrn, such as could occur iuiiow ing a bout of anoxia, in C. nemoralis hepatopancreas (for claritiy, some enzymes and intermediates have been omitted). Note particularly, anaerobic endproducts lactate and alanine are reoxidized by LDH and a GPTfGDH couple, respectively. NADH generated in these processes is reoxidized by cytosolic BHBDH and gluconeogenesis and redox balance is maintained. Pyruvate incorporation into glucose and/or glycogen is maximized by: (1) excbange with Acac; (2) maintenance of high mitochondrial redox, promoting rnaiate expon; (3) low mitochondrial concentrations of oxaloace tate prorno te syntizes is of Acac for export €tom fatty acids. Fig. 4.2 Proposed pathways involvexi in the generation of anaerobic endproducts in C. nemoralis hepatopancreas (for clarity, some enzymes and intermediates have been omitted). Note particularly cyclic fluxes of glutamate and aKG, and redox coupling. NADH produ& through a~erobicglycolysis is reoxidized via LDH and via the cytosolic GPWGDH couple. Kinetic characterizatioo of a cytosolic L-O-hydroxybutyrate dehydrogenrse from hepatopancreas of a terrestriai snaii, Cepaea neltylcalis published in The Journal of Experimental Zoology, 1997, 278: 140-146 Abstract The hepatopancreas of the terres trial gastropod Cepaea nemomlis has a cy top lasm ic fi-hydroxybutyrate dehydrogenase specific for L-8-hydroxybutyrate. No dehydrogenation of D-8-hydroxybutyrate by the enzyme was detected under Our experimrntal conditions. The apparent K, for L-0-hydroxybutyrate is similar to K, values for D-8-hydroxybutyrate determined for the rnitochondrial D-0- hydroxybutyrate dehydrogenase from other sources. The apparent Km for acetoacetate of the cytoplasmic L-B-hydroxybutyrate dehydrogenase is an order of magnitude greater than that of the mitochondrial enzyme. The former enzyme is rnarkedly sensitive to pH, with opposite effects on V,, in the forward and reverse directions. L-O-hydroxybutyrate dehydrogenase kinetics are also affected by adenosine phosphates and acetoacetyl-CoA. The kinetic propenies of the enzyme suggest that, while L-8-hydroxybutyrate dehydrogenation appears to be favoured , it could ~2ta1 yze the production of L-O-hydroxybutyrate from acetoacetate in the hepatopancreas of estivating terresaial snails. JNTRODUCTION 0-hydroxybutyrate dehydrogenase (BHBDH)catalyzes the reversible conversion of D-8-hydroxybutyrate (D-BHB)to acetoacetate (Acac), with the concornittant reduction of NAD. In most mammals, BHBDH is localized to the inner mitochondrial membrane and maintains essential associations with specific membrane phospholipids (Isaacson et al. 1979). It is highly specific for the D- stereoisomer of BHB (Lehninger et al. 1960). Another form of BHBDH occurs in liver and kidney of ruminant mammals. This form is predominantiy cytosolic (Koundakjian and Snoswell 1970) and oxidizes the L stereoisomer of BHB preferentially (Williamson and Kuenzel 1971). We have demonstrated a similar cytosolic form of BHBDH found in high activity in the hepatopancreas of the terrestrial snail, Cepuea nemoralis. Here, BHBDH occurs exclusively in the cytosol, with no mitochondrial activity, and L-BHB is the preferred enantiomer (Stuart and Ballantyne 1996). In mammals and elasmobranch fish, ketone bodies are known to be particularly important as energy subsnates during starvation (News holme and Leech 1983). Terresaial snails endure prolongeci starvation through estivating, and thus studying BHBDH function under conditions associateci with estivation may be important in elucidating the enzyme's physiological role. Estivation is characterized by irregular heart rate (Kratochvil 1976) and cyclic apnea and hypercapnia (Barnhart 1986). The resultant cycling of hemolymph oxygen and carbon dioxide content (Barnhart and McMahon 1987) is paralleled by fluctuations in intracellular pH, and a general decrease of pH values by approximately 0.5 uni& (Barnhart and McMahon 1988; Barnhart 1989; Rees et al. 1991). To characterize the kinetic functions of the cytosolic L-BHBDH of Cepea hepatopancreas, we have measured the substrate preferences and kinetic parameters of the enzyme, and measured these kinetic parameters in both directions of czdysis over a range of pH values similar to those found in active and estivating snails. We also have assessed the ab il ity of various metabolites whose concentrations change during anoxia to modulate BHBDH function. MATERIALS AND METHODS Experimental animals C nemoralis were collected from fields near the University of Guelph, Guelph, Ontario, in the late summer. Snails were kept in the laboratory at 22 & 2' C, in a terrarium with a bottom covering of sphagnum moss which was rnaintained moist by the addition of water several times ;week. The snails were fed an amount of lettuce which could be consumed in a 24-hour period twice a week, and trout chow twice monthly. The snails were kept under these conditions for between one and four rnonths during which time none was observed to estivate (as identified by the secretion of a calcareous epiphragm). Snails used for enzyme characterization averaged approximately 3 g and 1.5 cm shell diameter. Tissue Preparation To avoid the possible influence of diurnal cycles upon BHBDH activities, al1 animals were sacrificed in late afternoon and measurements taken immediately thereafter. Snails were removed fkom the terrarium approxirnately one hour btfore being sacrificed and placed upon moist paper towels with a small amount of lettuce. This stimulated activity, and crawling snails were decapitated and deshelled. The hepatopancreas was quickiy removed, separated from intestinal tissue, blotted dry, weighed and immersed in 10 volumes of ice-cold 50 mM imidazole buffer, pH 7.5. Hepatopancreas from two snails were pooled in each case. Al1 subsequent procedures were carried out at 4°C. Hepatopancreas were homogenized by three bursts of 10s each of a Polytron PT10 unit (Kinematica Gmbh., Lunirn, Switzerland). Homogenates were then centrifuged at 14 500 g for ten minutes. The supernatanu were collected and passed through 5 mL Sephadex G-25 columns centrifuged at 2000 g for 2 minutes. The eluants were used directly in determinations of BHBDH activities. Since the mitochondrial form of the enzyme is absent in Cepaea hepatopancreas (Stuart and Ballantyne l996), th is preparation contains on1 y cytosol ic L-BHBDW. BHBDH assay The activity of BHBDH was measured at 22*C, as the oxidation of NADH in the presence of Acac, or the reduction of NAD in the presence of BHB. The accumulation or disappearance of NADH was monitored at 340 nm (E, = 6.22) using a Hewlett Packard HP8452 diode array spectrophotometer equipped with a thermostatted ceil changer (Mississauga, Ontario, Canada) attached to a Haake D8 circulating water bath (Haake Buchler Instruments Inc., Saddlebrook, NJ). Imidazole solutions of various pH's were prepared using 50 rnM imidazole adjusted to the desired pH with KOH or HCI. The standard assay conditions were (pH 7.6, unless otherwise stated): dehydrogenation direction: 10 mM L- or D-BHB and 2 mM NAD or NADP; reduction direction: 50 mM Acac and 0.2 mM NADH or NADPH. Determinations of apparent K, (hereafter referred to as K,J and V,, for L-BHB,D-BHB, Acac, NAD, NADP. NADH and NADPH, and the effecu of pH and metabolites on these parameters were made by holding one reaction substrate constant at saturating levels while varying the other. For K, L-BHB or K, D-BHB determination, the L-BHB or D-BHB concentrations were 0.05, 0.07, 0.09, 0.14, 0.25, 0.5 and 5.0 mM. For K, Acac determination, the Acac concentrations were 0.5, 0.7, 0.9, 1.4, 2.5, 5.0 and 50.0 mM. For Y, NAD or K, NADP determination, the NAD or NADP concentrations were 0.02, 0.028, 0.036, 0.056, 0.1, 0.2, 2.0 mM. For rneasurement of inhibition by D-BHB,the dehydrogenation of 0.6 rnM L-BHB in the presence of 0.25, 0.5, 2.5 and 5.0 mM D-BHB was measured. For determination of the effecu of metabolites on kinetic parameters, one of O. 1 rnM acetoacetyl-CoA, 0. L mM CoA, 0.1 mM acetoacetyl-CoA, 3.0 mM L-lactate or 3.0 mM L-alanine were included in the assay medium. Adenosine phosphate effecu were measured at 0.03, 0.3 and 3.0 rnM AMP, ADP or ATP. Kinetic constants were estimated using a cornputer-aided iterative procedure to fit the data to the rectangular hyperbola. Chernicals Al1 chernicals were obtained from the Sigma Chernical Co. (St. Louis, MO) and were of the highest purity available. D-BHB was 99% pure, with less than 0.4 % L-BHB contamination. Statis tical analyses Effects of metabolites on kinetic constants of BHBDH were tested using ANOVA and means of BHBDH activity in the presence of metabolites were contrasteâ with control means using SYSTAT? RESULTS Subsbate specificity and kinetic constants The cytosolic BHBDH from C. nernorulis hepatopancreas is highly specific for L-BHB,and L-BHB dehydrogenation is not inhibited by concentrations of D-BHB up to 5 mM. NAD is the preferred coenzyme of the cytosolic L-BHBDH, and activity with NADP is less than 10% of that with NAD (Table 5.1). NADP fails to stimulate D-BHB dehydrogenation, measured under standard conditions at pH 7.6. At the same pH, V, for L-BHBDH in the direction of Acac reduction is approximately half of V,, V,, in the direction of L-BHB dehydrogenation. LAcac is approximately i~;ûrder of magnitude greater than &L-BHB. pH effects on kinetic constants L-BHBDH from C. nernorulis hepatopancreas is markedly sensitive to pH changes in the physiological range. The enzyme has a pH optimum for L-BHB dehydrogenation near 8.0 (results not shown), which is similar to optima from bacteria (Delafield and Doudoroff 1969; Krebs et al. 1969) but more sharply defined. Changes in pH have opposite effects on V,, of L-BHBDH in the directions of Acac reduction and L-BHB dehydrogenation in vitro (Fig 5.1). The V, for Acac reduction shows a trend of increasing as pH decreases. V, of Acac reduction is almost five times greater at pH 6.9 than at pH 8.1 (Fig. 5. la). In contrast, the V, for L-BHB dehydrogenation shows a trend of increase at higher pH values, with a V,, V,, L-BHB dehydrogenation three times greater at pH 8.1 than at pH 6.9 (Fig. 5. lb). Over the physiological range of pH 6.9 to pH 7.6 no changes in GL-BHB or IQAcac are statistically significant. However, &L-BHB does show a trend of increase with increasing pH. The LL-BHB (Fig. 5.2b) and LAcac (Fig. 5.2a) show statistidly signifiant differences from values at pH 7.6 only at pH 8.1, where &L- BHB is almost doubled and K,Acac is almost halved, compared with values at pH 7.6. Reaction velocities in both directions, at pH 6.9 and 7.6, were calculated using the experimentally derived values for kinetic constanu with the Michaeiis- Menten equation (Fig . 5.3). Metabolite effects No effects on K, or V,, values in either fonvard or reverse directions were observed foi CoA, acetyl-CoA, L-lactate, or L-alanine. Similarly, adenosine phosphates had no effect on these parameters at concentrations of 0.03 and 0.3 mM. At concentrations of 3 mM, however, the adenosine phosphates were strong activators of the V,, of reduction of Acac by L-BHBDH (Table 5.2). At these concentrations ATP also inhibited V,, of L-BHB dehydrogenation by 17.516, though no effecu of AMP or ADP were observed. Effecu of adenosine phosphates on K,Acac and QL-BHB of C. nemoralis BHBDH were also observed only at concentrations of 3 mM (Table 5.3). At 3mM, AMP, ADP and ATP cause 73.656, 97.7 56 and 6 1.4 56 increases in K,,,Acac respectively, indicating a decreased afiYinity for Acac under these conditions. Of the other metabolites examined, only acetoacetyl-CoA exerted effects on V- values of BHBDH and no effects on K, values were observed. Acetoacetyl-CoA caused a 70.5 % stimulation of V,, of Acac reduction and an 18.8 % stimulation of V,, V,, of L-BHB dehydrogenation (Table 5.2). Experimental conditions controlled for interference from L-3-hydroxyacyl-CoA dehydrogenase, which is also present in hornogenates of hepatopancreas (Stuart and Ballantyne 19%). Effects of acetoacetyl-CoA and 3mM ATP on reaction velocities in both directions of catal ysis were calculated using experimentall y derived values for kinetic constants with the Michaelis-Menten equation (Fig . 5.3). DISCUSSION The BHBDH found in the hepatopancreas of the terrestrial snail C. nernoralis differs froni the enzyme found in most mammals in that it is not membrane-bound and occurs in the cytosol. A unique fea~eof the snail enzyme is that it exclusively oxidizes L-BHB.We have used highly purified L- and D-BHB preparations to demonstrate that D-BHB is not oxidized and does not effect L-BHB oxidation. A low level of activity with D-BHB previously reported (Stuart and Ballantyne 1996) appears to have resulted from the presence of L-BHB, as impurity in the D-BHB preparation. Though the presence of D-BHB has been demonstrated in the hemolymph of a fkeshwater gaseopod (Meyer et al. 1986), the use of L-BHB as a substrate iii iiie energy metabolism of terrestrial snails, or other animals, has not been studied. Possible advantages of using L-BHB instead of D-BHB are not immediately obvious. The preference of C. nernorah hepatopancreas BHBDH for L-BHB may result from random mutation and subsequent evolutionary divergence from an ancestral D- BHBDH. This evolutionary event must have occurred within the pulmonate gastropods, as D-BHBDH has been demonstrated in the hepatopancreas of the freshwater pulmonate, B. glabmta. Certain kinetic constants of the enzyme also differ from values measured in other animals and bacteria. Cornparison of the kinetic constants of BHBDH from snail hepatopancreas with those of the enzyme fiom other sources may be used to infer the physiological roles of the snail enzyme. BHBDH from C. nernoralis hepatopancreas is similar to the cytosolic enzyme measured in ruminant liver and kidney in its specificity for L-BHB (Williamson and Kuenzel 197 1). However, the &L-BHB and KmAcac of C. nemomlis L-BHBDHare low (Table 5.4) compared with those observed with the sheep enzyme (Williamson and Kuenzel 197 1). In this respect, the snail enzyme appears to be more functionally similar to BHBDH €rom other sources. LL-BHB for L-BHBDH is similar to &D-BHB for D-BHBDH purifieci from severai species of bacteria and from the beef hem mitochondrial enzyme (Table 5.4). The snail enzyme is also similar to BHBDH from the bacterium Azospidlum brasilense in the inhibition of BHB dehydrogenation by adenosine phosphates. The V,, of D-BHB dehydrogenation in these bacteria is inhibited by 121, 1156 and 30% by 3 mM AMP, ADP and ATP, respectively (Ta1 et al. 1990). With both the snail and bacterial enzymes, no effects of adenosine phosphates are seen below these concentrations. BHBDH from C. nemomlis hepatopancreas differs ftom that of A. brasilense, however, in that YAcac is approximately an order of magnitude greater for the snail enzyme. This high LAcac is likely of considerable importance in determining the funcrional direction of the enzyme in vivo, as detailed below , Measured at a pH value of 7.6, typical of intracellular pH in active snails (Rees et al. 1991), L-BHBDHappears to be poised toward the net utilization of L- BHB, and not its synthesis. The high LAcac indicates a high [Acac] is necessary to saturate the forward reanion. At subsanirating [Acac] , under control conditions, the velocity of L-BHB dehydrogenation appears to greatly exceed that of Acac reduction (Fig. 5.3). Given these kinetic properties, it is likely that there is littie BHB formation by hepatopancreas. If hepatopancreas is in fact ketogenic, primari:? Acac would be produced under the intracellular conditions charactexistic of active snails. pH and metabolite effects on kinetic constants One change observed in estivating snails is a decrease in intracellular pH of approximately 0.5 uni@ (Rees et al. 1951). Lowering pH would decrease flux in the direction of L-BHB dehydrogenation, at al1 [LSHB] and increase flux in the direction of Acac reduction, at al1 [Acac], relative to control values (Fig. 5.3). This indicates that BHBDH kinetics become more favourable for L-BHB production at the low pH's characteristic of estivation. Effects on forward and /or reverse reactions are also observed in the presence of AMP, ADP or ATP (Fig. 5.3), but only at concentrations of 3 mM. BHBDH appears equally sensitive to al1 three adeny lates examined, and thus the relevant parameter which coufd affect the enzyme's function is the size of the total intracellular adenosine phosphate pool, which has been measured in terrestr iaI gasuopods at 1.6 mM (Churchill and Storey 1989) and 1.1 mM (Rees and Hand 1991). As the size of this pool is below 3 mM, and unaffected by estivation (Rees and Hand 199 1) adenosine phosphates appear not to be physiologicall y important modulators of BHBDH kinetics. Hepatopancreas BHBDH is also sensitive to acetoacetyl-CoA at concentrations of 0.1 mM. Acetoacetyl-CoA is a substrate in the reversible reaction catalyzed by 3- oxoacid-CoA transferase, w hic h synthesizes acetoacetate and thus immediately precedes BHBDH in the ketogenic direction. The ability of this effector to increase the rate of Acac reduction relative to L-BHB dehydrogenation suggests a means by which BHBDH activity is responsive to increased flux through this pathway. An increase in the supply of acetoacetyl-CoA for ketogenesis in the hepatopancreas could increase the rate of supply of Acac to BHBDH and, in response, the rate of BHBDH catalysis in the direction of L-BHB synthesis. In summary, the ability of certain effectors to modulate kinetic constants of hepatopancreas BHBDH indicates that this enzyme is responsive to changes in physiological state. Kinetic characteristics of L-BHBDH suggest that the enzyme bars some functional resemblance to D-BHBDH characterized fiom other sources, despite an absolute specificity for L-BHB and a relatively high LAcac. The high KAcac is an important factor in determining the functional direction of the reaction. In, active mails, the hepatopancreas is likely to be either ketolytic, or produce primarily Acac, as L-BHBDH kinetics do not appear to favour L-BHB production. At the depressed pH's typical of estivation, however , L-BHBDH kinetics are more favourable for L-BHB synthesis. Table 5.1. Kinetic constants of Cepea nemoralis hepatopancreas BHBDH. Values are means * S.E. of 6 observations, nad. = not determined S ubstrate KI,(mM) V,, (pmol/min/g wet wt.) L-BHB 0.61 k 0.07 D-BHB n.d. NAD O. 17 & 0.01 NADP f .O1 * 0.03 Acac 6.35 I0.64 NADH 0.063 * 0.016 Table 5.2. Effects of various metabolites on V,, for BHBDH in directions of Acac reduction and L-BHB dehydrogenation. Data are presented as % of control value (no effector) and are means f S.E. (n = 4). "*" denotes significant metabolite effects (P < 0.05). "n.mW= not measufed. pp - - - - - Assay Direction Treatment Acac to BHB L-BHB to Acac control 100 3 mM AMP 254.3 f 28.0 ' 3 mM ADP 279.2 & 42.6 ' 3 mM ATP 249.6 & 41.1 ' 0.1 mM AACoA 170.5 f 24.8 ' Table 5.3. Effects of various metabolites on K, for Acac and L-BHB in directions of Acac reduction and L-BHB dehydrogenation. Data are presented as % of control value and are means f S.E. (n = 4). "*" denotes significant metabolite effects compared to control (P < 0.06). %.m." = not measured. Treatment Acac to BHB L-BHB to Acac conttol 100 100 3 mM AMP 173.6 i 32.7 ' 69.3 I07.9 ' 3 mM ADP 197.7 i 40.5 ' 119.2 * 12.2 3 mM ATP 161.4 i 31.1 ' 81.9 * 6.4 0.1 mM AACoA 111.5 i: 17.5 103.3 f 10.3 Table 5.4. Apparent K, values of BHBDH from different sources. enzyme source Apparent K, (mM) Pseudomonas lemoigneil (bacterium) (D-BHB) 0.6 (Acac) 0.9 Rhodopseudomonas spheroide3 (bacterium) (D-BHB) 0.4 (Acac) 0.3 Azospirilhm brasilonse3 (bacter ium) (D-BHB) 1.0 Beef hart submitochondrial vesicles4 (D-BHB) 0.8 Rat liver submitochondrial vesicles4 (D-BHB) 1.3 Sheep kidney cytoplasmic fractiod (L-BHB) 6.2 (Acac) 25 .O C. nenorulis hepatopancreas (L-BHB)0.61 (Acac) 6.4 Delafield and Doudoroff (1969). Krebs et al. (1969). Tal et al. (1990). McIntyre et al. (1988). ' Williamson and Kuenzel (1971). Fig 5.1. Effect of pH on V, of 0-hydronybutyrate dehydrogenase in (a) the direction of acetoacetate reduction (n = 5, '*' = significantly different fiom control at P < O.O4), and (b) L-BHB dehydrogenation (n = 6, '*' = significantly different fkom control at P < 0.02). Data presented as 96 of V, at pH 7.6 (control) and an meam & S.E. Fig 5.2. Effect of pH on (a) apparent &(acetoacemte) (n = 5, '*' = significantly differem ftom control at P < 0.04) and @) apparent &(L-0-hydroxybutyrate) (n=6, '*' = significantly different fiom control at P < 0.002) of 8-hydroxybutyrate dehydrogenase. Data points are means f S.E. Control = & at pH 7.6. Fig . 5.3. Calculated velocities of 6-hydroxy butyrate dehydrogenase reaction @mollmin/g wet W.) in the directions of L-8-hydroxybutyrate synthesis (a) and dehydrogenation (b), under control conditions @H 7.6, no effectors) and in the presence of effectors. Velocities were calculateci using experimentally determined kinetic parameters (V, and K,,J applied c the Michaelis-Menten equation. Insets are expandeci representations of reaction velocity at low subsaate concentratioas (shown as boxed in areas on primary graph). CHAPTER 6 Tissucspecific forms of Il-bydroxybutyrate dehydrogenase oridize the D or L- enantiomers of 8-hydroxybutyrate in the terrestrial gastropod Cepaca ~mralis published as a Rapid Communication in The Journal of Experirnental Zoology, 1997, 278: 115-1 18 INTRODUCTION 8-hydroxybutyrate dehydrogenase (BHBDH) catalyzes the interconversion of the ketone bodies , ace toacetate (Acac) and Dd-hydroxybutyrate (BHB) . In vir tua11 y al1 anirnals, including marnmals and fish (Newsholme and Leech, 1983) and freshwater molluscs (Meyer et al. 198Q, BHBDH exists within the mitochondria. The mammalian BHBDH is a popuiar mode1 for the study of the kinetics of membrane-bound enzymes, since it has an obligate requirement for certain phospholipid species (Isaacson et al. 1979). We have recently described a different and unique form of BHBDH found in the terrestrial mail Cepaea nemoralis (Stuart and Ballantyne 1996) which differs in two respects from the enzyme described above. This enzyme occurs in the cytosol of hepatopancreas cells and oxidizes exclusively the L enantiomer of BHB. This form of BHBDH does not occur in al1 tissues of C. nemoralis. Here we describe the presence in other tissues of another cytosolic form of BHBDH with an enantiomeric specificity for the D-stereoisomer, and demonstrate that it is a separate protein from the L-BHBDH. MATERiALS AND ME3"I'ODS C. nemoralis were collected from fields near the University of Guelph campus. A colony of these snails was maintained in the laboratory, in terrariums kept near 30% ambient hurnidity with periodically moistened sphagnum moss. Snails were fed lettuce ad libitum, Tissue preparation for enzyme assays was essentially as described in Stuart and Ballantyne (1996). Active (withdrawn from shell) adult snails approximately 1.5 cm diameter were decapitated and deshelled. Tissues were prepared for the enzyme assays by placing the excised tissue in 2 mL of ice-cold mitochondrial isolation medium (20 mM N-[2- hydroxyethyllpiperazine-N'-[2-ethanesulfo acid] (HEPES) and 100 mM sucrose, pH 7.5) and homogenizing by five passes of a Potter-Elvejhem homogenizer with a tefion pestle attached to a drill press operated at < 100 revolutions per minute. Al1 subsequent procedures were carried out at 5" C. Two separate centrifugation protocols were applied to homogenates. Initially, we used a 10 OOOg, ten minute centrifugation to separate tissues into mitochondrial and cytosotic fractions and verifid the absence of BHBDH activity in the 10 OOOg pellet. Subsequently, homogenates were centrifuged at 200g for 10 minutes and the resultant pellet discarded. This supernatant was centrifbged at 10 000 g for 10 minutes and resultant supernatanu decanted. The remaining mitochondrial pellets were resuspended in 2 mL of mitochondrial isolation medium. The supernatant was centrifuged at 30 ûOg for 10 minutes, the pellet discarded and the resultant supernatant considered the cytosolic fraction. Both tissue fractions were then sonicated with a 15 s burst at 80% output, 50 Watts, on a Vibra-Ce11 sonicator (Sonics & Materials Inc., Danbury, CT). Aliquots of these fractions were used directly in enzyme assays. Citrate synthase (CS) and BHBDH activities were measured as described by Stuart and Ballantyne ( 1996) with the following exception: the BHBDH assay medium contained 2 mM NAD and either 400 mM DL-BHB or 200 rnM D- or L- BHB in 50 mM imidazole, pH 8.0. Al1 chernicals for enzyme assays were purchased fiom Sigma Chernical Co. (St. Louis, MO) and were of the highest purity available. Tissues were prepared for electrophoresis on cellulose-acetate gels by adding about 20 mg of tissue to 200 CL (heart), 100 pL (kidney) and 250 pL (hepatopancreas) of nis-glycine gel buffer (25 mM Tris, 200 mM glycine, pH 9.0) in eppendorff tubes and hornogenizing with a tight-fitting plastic pestle. Homogenates were clarified with a 5 minute, 10 Oûûg centrifugation. A sample of each supernatant was applied using a Super Z Applicator (Helena Laboratories. Beaumont, Texas) to a 76 mm by 76 mm Titan III cellulose-acemte plate (Helena Laboratories) which had been presoaked in gel buffer. Gels were placed in a plexiglass electrophoresis tank, with gel buffer used as electrode buffer. A 300 V differential was applied, using a Heathkit Regulat-A Power Supply (mode1 IP-2717A), for two hours at 5" C. Gels were removed and stained with a solution containing 0.3 mg M'iT (3-[4,5-Dimethylthiazoi-2-YI]-2,s- diphenylteaazoliurn bromide), 0.07 mg phenazine methosulfate, 1.O mg NAD, 33.6 mg DL- or D- or L- BHB in 3 mL aisglycine (pH 9.0) added to 2 mL agarose gel maintained at 60" C. The staining solution was removed fiom gels after approximately I hour and gels were photographed while still moist. RESULTS We used CS as an indicator of the extent of leakage of mitochondrial rnatrix enzymes from mitochondria damaged in the tissue fractionation process (Table 6.1). The appearance of CS in the cytosolic fractions of kidney and hepatopancreas was 11 % and 13 % of summed mitochondrial and cytosol ic activities, respectively , indicating minimal leakage frorn the rnitochondrial matrices in these tissues. In the ventricle, cytosolic CS accounted for 45 96 of total activity, suggesting greater leakage of mitochondrial contents in this tissue. This greater proportion of damaged mitochondria in the fractionation of ventricle tissue did not affect the interprpthn of results. Virtually no rnitochondrial BHBDH activity was detected in any tissue. Cross-contamination of the mitochondrial Fraction with cytosolic enzyme was 3.4 96, 1.0% and 1.4% of total BHBDH activity in venuicle, kidney and hepatopancreas, respectively . Homogenates of ventricle tissue oxidize dmost exclusively D-BHB,whereas hepatopancreas is specific for L-BHB and kidney oxidizes both stereoisomers (Table 6.1). Some apparent non-specificity of BHBDH activity in ventricle and hepatopancreas was likely the result of small amounts of the other stereoisomer present as impurity in the commercial BHB preparations. Oxidation of L-BHB in ventricle occurred at OS% the rate of D-BHB oxidation. The commercial L-BHB preparation was 97 1pure and containecl approximately 0.4 % D-BHB.This contamination likely acwunts for the low activity of LBHBDH in heart. Similady, oxidation of D-BHB in hepatopancreas occurred at 4.5% the rate of L-BHB oxidation. The commercial D-BHB tion was 98 % pure with less than 2% L- BHB contamination. Thus, contaminating L-BHB likely accounts for the low rate of oxidation of D-BHB in this tissue, nie oxidation of D-BHB by D-BHBDH appears to be inhibited by L-BHB.In kidney, activity with DL-BHB as substrate was less than would be expected by summing the independant D- and L activities (Table 6.1). Similarly, BHBDH activity in ventricle is greatly decreased when assayed with DL-BHB compared with D-BHB. Staining of elecrophoretic gels with DL-BHB clearly demonsuates that two separate enzymes are involved in the oxidation of the enantiomers of BHB in the tissues examined. Two distinct bands which migrate different distances from the origin are visible (Fig. 6.1). The venaicle band (D-BHBDH)occurs equidistant from the origin to the top kidney band, indicating that this form occurs in both tissues. Similarly, the hepatopancreas band (L-BHBDH) matches the bottom kidney band. The kidney thus contains both forms of the enzyme (L-BHBDHand D-BHBDH). Separate staining with D- or L- BHB corroborated the identification of excIusively L- BHBDH in hepatopancreas and exclusively D-BHBDH in ventricle (not shown). DISCUSSION Our results indicate that two distinct forms of BHBDH exist in C. nernoralis tissues. Both enzymes are cytosolic and each is specific for one stereoisomer of BHB. Subcellular location of BHBDH in newralis tissues There is no evidence for the existence of the typical mitochondrial membrane- bound form of BHBDH in hem, kidney or hepatopancreas of the terrestrial snail, C. nemoralis. In al1 of these tissues, BHBDH is localized to the cytosol. Very low activities of the enzyme in the mitochondrial compartments of these tissues are consistent with minor contamination from the cytosol during the tissue fractionation procedure. The subcellular location of Cepaea BHBDH contrasts with that of freshwater gastropods. where BHBDH is rnitochondrial (Meyer et al. 1986; persona1 observations). Stereospecifity of BEBDH C. nemoraiis have the ability to utilize both enantiorners of BHB, each catalyzed by a different protein as indicated by cellulose acetate electrophoresis. While ventricle and hepatopancreas oxidize exclusively D-BHB or L-BHB, respectively, the kidney can oxidize both subsuates. The demonstration of two separate proteins involved in the utilization of D-and L-BHB rules out another possible mechanism which could account for the pattern of L- and D- BHB utilization observeci in Table 6.1. Racemisation of one enantiomer to another, e.g. L-BHB to D- BHB, could precede oxidation by D-BHBDH. This would require two separate enzymes, a racemase and a BHBDH. However, these proteins would likely have been separateci by the gel matrix. Thus where L or D- BHB was directly oxidized, a single clear band would have been visible. Where racemization was coupled to oxidation, a blurred band would have occurred equidistant from the origin to the clear band, indicating that both the racemase and dehydrogenase had migrated simiiar distances. Alternatively, if the racernase and dehydrogenase migrated very different distances, no band would have been visible. Thus, as two distinct bands were clearly visible at different distances from the origin, two separate enzymes must be responsible for the oxidation of the stereoisomers of BHB. Though many enzymes exist as multiple isoforms, with individual isozymes often Iocalized to specific tissues, these isoforms typicaily differ from one another in their kinetic propenies, including Michaelis constants and sensitivity to cofac~rs. The differential occurrence, within tissues of a single organism, of two distinct isoforms of an enzyme which are specific for different enantiomers of a substrate is, to Our knowledge, unique to this terresnial snail. We are not aware of other examples of this phenornenon in the animal kingdom. Lactate dehydrogenase (LDH)occurs as either D-LDH or L-LDH throughout the animal kingdom (Long 1976). However, no organism has been shown to have both D- and L-LDH (Long 1976). Some rnolluscs use both the D- and L- enantiomers of certain arnino acids (Ballantyne and Chamberlin 1994). Alanine, in panicular , occurs as both D- and L- stereoisomers in high concentrations in tissues of some marine bivalves (Yarnada and Matsushima 19%). In these molluscs, however , there is no evidence of a D- and L- alanine aminotransferase (Hayashi 1993). Instead, a D- amino acid oxidase or a racernase have been implicated in the oxidation of intracellular D-alanine (Matsushima and Hayashi 1992; Ballantyne and Chamberlin 1994). These authors suggest that this may be signifiant in rnaintaining a role for D- alanine as an intracellular osmolyte while L-alanine is accessible to the oxidizable substrate pool. Though the mechanism of D- and L-BHB metabolism in Cepaea differs from that of D- and L- alanine in some bivalve molluscs, the designs are similar in that in both cases a subsnate is made unavailable for oxidation by a tissue. In Cepaea. hemolymph L-BHB could not serve as an energy subsaate for the ventricle, nor could D-BHB be used by hepatopancreas. This effectively creates a partitioning of BHB. Such a design would allow BHB to be directed to specific tissues, or to specific ce11 types within a tissue, e.g. kidney, where D-BHBDH and L-BHBDH could occur in different ce11 types. Achizving this extra level of control over the metabolism of BHB mav be an important adaptation of terrestrial snails, where relatively high activities of BHBDH and other enzymes of ketone body metabolism suggest that ketone bodies play a prominent role in energy metabolism (Smand Bailantyne, 1997). As fatty acids do not appear to be oxidized substantially by peripheral tissues (Stuart and Ballantyne, 1997) ketone bodies may be an important means of distributing lipid carbon fiom cenual stores to peripheral tissues. Thus, the controlled synthesis of one or the other enantiomer of BHB may allow stored lipid carbon to be directed to a certain tissue or tissues for oxidation. In summary, terrestrial snails possess a unique organization of ketone body metabolism, which differs fiom the mammalian mode1 in the subcellular compartrnentation and stereospecificity of BHBDH. The present snidy reports the first demonstration of a cytosolic D- BHBDH in animal tissues. Similarly, the presence in different tissues of two distinct forms of BHBDH, each specific for a single enantiomer of BHB is, we believe, unique. This organization of ketone body metabolisrn results in a partitioning of BHB availability between tissues, and the physiological implications of this phenornenon should be investigated. As both D- and L-BHBDH are present in substantiai activities, it is likely that they play important roles in the intermediar y metabol ism of terrestrial mollusc tissues. Finally, this description of BHBDH isoforms in C. nemoralis tissues should be noted by population geneticists, as gel staining with DL-BHBDH could give misleading results if not interpretted on the basis outlined above. Table 6.1. Activities of citrate synthase (CS) and fl-hydroxybutyrate dehydrogenase (BHBDH)in mitochondrial and cytosolic cornpartmenu of Cepaea nemoralis tissues (n = 5). Enzyme(s) Cornpartment Ventricle Kidnev Hepatopancreas CS rnitochondria 2.82 f 0.34 1.87 =t 0.55 1.N f 029 CS cyioplasm 3.40 k 0.90 0.23 I0.07 O. 18 * 0.06 DL-BHBDH mitochondria 0.35 * 0.22 0.03 f: 0*02 O 02 I: 0.01 DL-BHBDH cytoplasm 10-12 * 1.33 2.79 =f= 0.37 1.30 * 0.24 D-BHBDH cytoplasm 36.36 * 4.55 3.99 * 0.65 0.06 * 0.02 L-BHBDH cytopiasm 0.17 * 0.08 1.61 I 0.31 1.37 * 0.25 hepatopancreas kidnty Fig. 6.1 Cellulose acetate gel of ventricle (lane l), hepatopancreas (lane 2) and kidney (iane 3), stained with DL-BHB (see text). A Preliminary Kinetic Characterization of the Cytosoüc D-B-Hydrorybutyrate Dehydrogenase from the Heart of the Terrestrial Snail, Cepaeo nenioralis The highest activities of the en yme l3-hy droxy butyrate dehy drogenase (BHBDH) in the animal kingdom are found in the hearts of terrestrial puhonate snaîis. BHBDH interconverts the ketone bodies, acetoacetate (acac) and 0- hydroxybutyrate (BHB). These substrates appear to be particulatly important to the intermediary rnetabolism of terrestrial pulmonates (Stuart and Bdmtyne, 1997a). In vertebraîes and all other animals where BHBDH activity is fouud, this enzyme is localized to the mitochondriai inner membrane, where it maintains an essential association with the phospholipid phosphatidylcholine (El Kebbaj and Latruffe, 1986). Terrestrial puimonate mails, vimially aU of which belong to the order Stylommatophora, have evolved a unique organization of ketone body metabolism which is not found in the closely related freshwater pulmonates (order Bassommatophora) (Stuart et al., 1998) or elsewhere in the animal kingdom. Specifically, tissues of Stylommatophoran mails lack the typicai mitochondrial form of BHBDH which was previously thought to be the only form in which BHBDH exists, and which is found in dl other animals which show BHBDH activity, including the closely related Bassomatophora Instead, Stylommatophorans have two novel isoforms of BHBDH. Both cWer fiom the "typical",mitochondrial BHBDH in that they occur in the cytosol (Stuart and Baüantyne, 199%). One BHBDF isoform is speafic for the L-aiantiomer of BHB, and is present in hepatopanaeas and kidney tissue (Stuart and Ballantyne, 1997%). Another, BHBDH isoform oxidues exclusively D-BHB and is found in kidney and ventride. We have characterized aspects of the kmetic properties of the L-BHBDHfound in the hepatopancreas (Stuart and Bdantyne, 1997~).This enyme has a near-absolute requirement for NADH, is highly speafic for L-BHB, adactivity is not inhibited by D-BHB. This enzyme is bighly seisitive to changes in intracellular pH. Kinetic parameters of this enzyme are also affected by adenosine phosphates, at concentrations of 3 mM, and by acetoacetyl-CoA at 0.1 mM. Here, we describe some functional characteristics of the cytosolic D-BBDH from the hearts of the stylommatophoran siail Ceposa nemondis, discuss how these Wer from the mitochondrial enzyme and the L-BHBDH from hepatopancreas, and formulate several hypothesa regarding the possible adaptive benefits which could * have selected for these modifications, which appear to have occurred sometime after the Bassommatophora and Stylommatophora evolved from a common ancestor. MATERIALS AND MEIXIODS Exprimenîai dmds C. nemomlis were coilected from fields near the University of Guelph. Guelph, Ontario, in the late surnmer. Snails were kept in the laboratory at 22 k 2O C, in a terrarium with a bonom covering of sphagnum moss which was maintained moist by the addition of water several times a week. The snails were fed an amount of lemice which could be consumed in a 24-hour period twice a week. The mails were kept under these conditions for up to one month, during which time none was observed to estivate (as idmtified by the secretion of a calcareous epiphragm). Snds used for enzyme characterization averaged approximately 3 g and 1.5 cm diameter. Tissue hparation To avoid the possible influence of dimal cycles upon BHBDH activities, ail animals were sadced in late aftemoon and measurements taken immediately thereafter. Snails were removed from the terrarium approximately one hour before being sacrificeci and placed upon moist paper towels with a small amount of lettuce. This stimulated activity, and crawling mails were decapitated and deshelIed Ventrides were removed and placed on ice. As ventricle weight averaged about 2 mg, it was necessary to pool tissue from 8 mails to acquire eiough homogenate for repeated measurements Pooled tissues were weïghed and immersed in 1 mL of ice- coId buffer (see below for descriptions of bdfers used). AU subsequent pro cedures were canied out at 4°C. Tissues were homogaized by three 10 s bursts of a Polytron PT10 unit (Kinematica Grnbh., Luzum, Switzedand). Homogenates were then centrifuged at 10 000 g for ten minutes. The supematants were either used duectly in assays. In some experimeits where protocols which minimized loss of activity were assessed, the supernatant was next passed through 5 rnL Sephadex G-25 columns centrifuged at 2000 g for 2 minutes, and the eluants used in subsequent determinations. D-BHBDH is highly labile, so we tested the ability of three buffers and two tissue preparation procedures to maintain enzyme activity for a sufficieut period of time to measure kinetic parameters. Buf'fer 1 consisted of 50 mM imidazole, pH 7.5. Buf'fer 2 was identical to bis, but with the addition of 2 mM Dm.Buffer 3 was identical to the first buffer, but also contained 40% (vh) glycerol. Buffer 3 was able to maintain D-BHBDH activity near 100% over a period of 130 min (Fig. 7.1) and at 83% after 395 min. Attempts to remove low molecular weight substances fouowing the initial extraction, using a Sephadex G-25 column. resulted in a rapid loss of activity following elution. Therefore, the original supernatant in imidazole-glycerol buffer was used for aU subsequent measurements. The final tissue dilution within the cuvette was approximately 4 OOOX. Thus, low molecular weight compounds other than glycerol were substantially diluted BEiBDH assay The activity of BHBDH was measured at 22°C- as the oxidation of NADH (of NADPH) in the presence of Acac, or the reduction of NAD (of NADP) in the presence of D-BHB. The accumulation or disappearance of NADH was monitored at 340 nm (E, = 6.22) using a Hewlett Packard HP8452 diode array spectrophotometer equipped with a thsrmostatted ceil changer (Mississauga, Ontario, Canada) attached to a Haake D8 circulating water bath (Haake Buchler Instruments Inc., Saddlebrook, NJ). Imidazole solutions of various pHs used in D-BHBDH assays were prepared using 50 mM imidazole, adjusted to desired pH with KOH or HCl. The standard assay conditions were (pH 7.6, unless otherwise stated): dehydrogenation direction: 1O mM L- or D-BHB and 2 mM NAD or NADP; reduction direction: I mM Acac and 0.2 mM NADH or NADPH. Determinations of apparent K, (hereafter referred to as Y) aud V,, for L-BKB, D-BHB,Acac, NAD, NADP, NADH and NADPH, and the effects of pH and metabolites on these parameters were made by holding one reaction constituent constant at saturathg levels while varying the other. For K,,, L-BHB or &, D-BHB determination, the L- BHB or D-BHB concentrations were 0.1, O. 14, 0.18, 0.28, 0.5, 1.O and 1 0.0 mM. For &, Acac determination, the Acac concentrations were 0.01, 0.014, 0.018, 0.028, 0.05, 0.1 and 1.0 mM. For y NAD or K,,,NADP determination, the NAD or NADP concentr&ons were 0.02, 0.028, 0.036, 0.056, 0.1, 0.2, 2.0 mM, and for K, NADH determination, NADH concentrations were 0.002, 0.0028, 0.0036, 0.0056, 0.01, 0.02, 0.2 mM. For measurement of inhibition by L-BHB,the dehydrogenation of 0.6 mM D-BHB in the preseice of 0.1, 0.3, 0.6, 1.0, 2.0 and 4.0 mM L-BHB was measured For determination of the &ects of adenosine phosphates on kinetic parameters, 3.0 mM AMi?, ADP or AT'were added to the assay . Duplicate measuremaits were made on a single preparation of eight pooled ventticles. Kinetic constants were estimated for each set of measurements using a cornputer-aided iterative procedure to fit the data to the rectangular hyperbola. Duplicate determinations of kinetic parameters thus determined were averaged Chernicals AU chernicals were obtained fiom the Sigma Chemical Co. (St. Louis, MO) and were of the highest purity avaiiable. D-BHB was 98% pure, with less than 2.0% L-BHg contamination. L-BHB was 99% pure, with less than 0.4% D-BHB contrimination. RESULTS AND DISCUSSION Ventricle D-BHBDH was highly labile. Approximately 50% of initiai activity was lost by 30 minutes post-homogenbtion, when tissues are homogenized in imidazole baer alone, or in the presence of ditbiothreitol (DTT) (Fig. 7.1). Stabilization with a giycerol containhg medium, however, prevents this loss of activity. When stabilized this way, activity rernahs at or above 100% of initial after 130 min and at 83% of initial after 395 min (not show). This property of ventricle D-BHBDH dif'fers dramatically from the L-BHBDH from hepatopancreas, which was stable in imidazole buffer for greater thau 24 hrs (Stuart and Bdantyne, 1997). The ventricle enzyme also diffezs from the hepatopancreas isoform in its recognition of the non-oxidized enantiomer of BHB. Oxidation of L-BHB by L- BHBDH is not inhibited in the presence of D-BHB.However, D-BHB oxidation by heart D-BHBDH is strongly inhibited by L-BHB (Fig. 7.2). We used a Dixon plot (llv vs m) (Roberts, 1977) to determine the nature of this inhibition, and KI, graphically (Fig. 7.3). This gives a straight line (? = 0.99) with a y-intercept (KJ of 0.11. This indicates a linear cornpetitive inhibition of D-BHB oxidation by L-BHB, where both L-BHB and D-BHB are competing for the active site on D-BHBDH.The possible physiological ~i~canceof biis is unknown, and depends on whether L- BHB synthesized in other tissues can be transported into the ventricle. It indicates, however, that as the rate of D-BHB oxidation could be strongly aEected by the presence of L-BHB,tissue measurements of both D and L-BHB are required for estimates of reaction velo*. D-BHBDH was highly specific for NAD, and no activity was detectable when this was repiaced by NADP (Table 7.1). Cornparison of D-BHBDH kiaetic constants reveals that the YD-BHB for the ventricle enqme is similar to K,,,L-BHB for the hepatopancreas enzyme and to &,D-BHB from most animal tissues and bacteria (Table 7.2). In contrast, KAcac for the vaiaicle enyme was more than two orders of mapitude lower than that for hepatopancreas L-BHBDH, and an order of magnitude lower than D-BHBDH from several bactena (Table 7.2). This dramatic Merence betweai the kinetic properties of mitochondnal and cytosolic D-BHBDHs suggests that the shift in compartmentaiization of D-BHBDHfrom mitochondrion to cytosol has been 'accompanied by a change in the physiologicd roles of BHB metabolism. It is also likely, given the dramatic differences in kinetic charactenstics between D- and L-BHBDH,that these two enzymes have different fimctions in hem and hepatop ancreas, respectively . The low K, pyruvate relative to D-BHB resembles the kinetic characteristics of terminal dehy drogenases. Marine molluscs are kno wn to possess a number of unique anaerobic pathways. Molluscm Iactate dehydrogenase, octopine dehydrogaiase, aianopine dehydrogaiase aud strombine dehydrogenase all have similarly low y'spyruvate relative to Csfor lactate, octopine, danopine and strombine, respectively (Gade and Grieshaber, 1986). This appears to be important in ailowing intraceliular concentrations of these anaerobic aidproducts to accumulate during anolaa Similady, D-BHB could serve as an anaerobic endproduct in tenestria1 snail vmtricle, if the carbon source of acac under auoxic conditions is not lipid An important feature of cytosolic D-BHBDH kinetics in this regard is that, while &,,Acac appeared insensitive to pH changes in the physiological range (Fig. 7.4a). &, BHB increased by aimost 25% as pH is lowered fkom 7.1 to 6.5 (Fig. 7.4b). This characteristic suggests that DBKBDH kinetics increasingly favour D-BHB accumulatioii as intracellular pH drops during hypoxia (Rees et al., 199 1). Accumulation of D-BHB during hypoxia has been demonstrated in squirrels (ITAlecy et al., 1990), where D-BHBDH is mitochondrial, indicating that, hypoxia cm alter mitochondrial redox state to favour D-BHB.It is equaily Iikely that cytosolic redox bdance would favour D-BHB production in anoxic siail hearts. The possibility that cytosolic D-BHBDH serves as a taminal dehydrogenase in anoxic conditions, regmerating NAD* in the cytosol for giycolysis, should be investigated. The cytosolic D-BHBDH had a pH optimum of 7.6 for V,, in the direction of BHB oxidation (Fig. 7.5a). V,, in the direction of BHB production was opkd at pH 6.5 (Fig. 7.5b). The enzyme appean to be designed for D-BHB oxidation in active mails. At pH 7.6, which is typical of tissues of active terrestrial mails (Barnhart and McMahon, 1988; Rees et ai., 1991) the ratio Vfl, (the ratio of V,, in the fonvard, D-BHB producing direction to V,, in the backward, D-BHB oxidizing direction) was 0.14, indicating that D-BHB oxldation could proceed at seven times the rate of its production if substrate concentrations are saturating. This suggests that the ventride enzyme strongly favours D-BHB oxidation mder innacellular conditions typical of active mails. At the lower pH values which occur during estivation or anoxia (Rees et al., 1991). this ratio was increased to 0.25, indicating that kinetics becorne more favourable for BK6 production. In general howevq the cytosolic D- BHBDH was less pH-sensitive than the cytosolic L-BHBDHfrom hepatopancreas. In cornparison, VPrat pH 7.6 for L-BHBDHis 0.52 (Stuar& and Ballantyne 1997c), and lowering pH to values typical of estivating mails changed this ratio to 1.40, suggesting that the forward reaction direction of the L-BHBDHreaction cm be favoured under these conditions. Kinetic parameters of the cytosolic D-BHBDH appeared to be relatively insensitive to adenylates at concentrations of 3 mM (Tables 7.3 and 74,though statistical comp~sonswere not made. In contrast, the cytosolic L-BHBDH is moduiated by these concentrations of adenylates (Stuart aud Ballantyne, 1997~). In summary, the mail heart DBHBDH appears to have been functionaiiy modified from the presurnably ancestral mitochondrial enzyme. Perhaps most siWcmt is the large reduction in &, Acac, rendering the enyme functiondy simiiar to terminal dehydrogenases that catalyze reactions designed to accumulate anaerobic endproducts during anoxiaThis hypothesis shodd be tested in fume studies through measurements of D-BHB in heart tissue during anorcia. It wil! dso be interesting to determine, assuming that the cytosolic D-BHBDHis a modified copy of the mitochondnal -me, what cornpositional changes have occurred to cause this altered function. Table 7.1. Kinetic constants of Cepueu nem omlis heart D-BHBDH. Values are means of two measurements on eight pooled hem. n.d = not deterxnined Substrate % (m-) V,, (pznol/min/g wet W.) L-BHB n,m n.d. D-BHB 0.73 20.22 NAD 0.13 20.22 NADP am. n.d Acac 0.039 3.65 NADH 0.03 1 3.65 Table 7.2. Apparent K,,, values of BHBDH From different sources. Pseudom onas lem oigne? (bacteriun) (D-BHB) 0.6 (Acac) 0.9 Rhodopseudom onas spheroide? (bactenum) (D-BHB)0.4 (Acac) 0.3 A zospirilhm b rasilense' (bacterium) (D-BHB)1.0 Beef heart submitochondrid vesicles4 (D-BHB) 0.8 Rat liver sïbmitochond~ialvesicles4 (D-BHB) 1.3 Sheep kidney cytopiasmic fraction5 (L-BHB) 6.2 (Acac) 25.0 (L-BHB)0.61 (Acac) 6.4 C. nemorolis ventricle (D-BHB) 0.73 (Acac) 0.039 ' Delfield and Doudoroff (1969). ' Krebs et al. (1969). ' Ta1 et ai. (1990). ' McIntyre et al. (1988). ' Williamson and Kuenzel(l971). Stuart and Ballantyne ( 1997~). Table 7.3. EfYects of variom metabolites on V,, for BHBDH in directions of Acac reduction and L-BHB dehydrogenation. Data are presented as % of control value (no effector) and are means of duplicate measurements on a preparation of eight hearts. Assay Direction Treatment Acac to BHB L-BHB to Acac control 3 mM AMP 3 mM ADP 3 mM ATP Table 7.4. Effects of various metabolites on K,,,for Acac and L-BHB in directions of Acac reduction and L-BHB dehydrogenation. Data are presented as % of control value and are meaus of duplicate measurements of a preparation of eight hearts. Assay Direction Treatment Acac to BHB L-BHB to Acac control 3 mM AMP 3 mM ADP 3 mM ATP O control 2 rnM DTT time (min) Fig. 7.1. Decay of D-BHBDH activity over time (under standard assay conditions) in tissues homogenized in different buffers. Fig. 7.2. Inhibition of D-BHB oxldation in the presence of varying concentrations of L-BHB . Fig. 7.3. Dixon plot of the inhibition of D-BHB oxidation by L-BHB. Fig. 7.4. Eects of pH on the (a) &Acac and @) &BHB. us Fig. 7.5 Effens of pH on V,, in the direction of (a) D-BHB oxïation and (b) D-BHB production. CHAPTER 8 D and L-D-hydroxybutyrate debydrogenases and the evolution of ketone body metabolism in gastropod moiiuscs accepted for publication as a Research Note in The Biological Bulletin Abstract and Introduction In vertebrate animals, ketone bodies, synthesized primarily from stored lipid, are important metabol ic substrates (Newsholme and Leec h , 1983) dur ing starvation, ketone bodies, acetoacetate (Acac) and O-hydroxybutyrate (BHB) , are oxidized by some extrahepatic tissues at high rates, and thus perform the important hinction of sparing limited glycogen stores (Newsholme and Leech, 1983; Robinson and Williamson, 1980). The enzyme O-hydroxybutyrate dehydrogenase (BHBDH),which catalyzes the interconversion of the ketone bodies is found in ail mammals and most vertebrates, but is absent in most invertebrates (Beis et al., 1980), including marine molluscs (Stuart and Ballantyne, W6a). The highest measured BHBDH activities in the animal kingdom, however, are found in the hearts of terrestrial gastropod molluscs (Stuart and Ballantyne 1997a and b). We recently have demonsuated that, in tissues of the terrestrial gastropod, Cepaea nemordis, two unique and previously unknown isoforms of BHBDH occur (Stuart and Ballantyne, L997b). The isoforms differ from the wefl-characterized mitochondrial membrane-bound D-BHBDH found in al1 other animals (Lehninger et al., 1960) in that they are cytosolic, and one isoform is specific for the L enantiomer of BHB. Here we identio patterns in the evolution of these enzyme isoforms in the Gastropoda. BHBDH activities, stereospecifity and subcellular compamnentalization were measured in gastropod species representing four major groups with fres hwater and terrestrial representation, Ner itomorpha (primitive gilled gastropods) , Architaeniogl ossa (more advanced gilled gastropods), Basommatophora (freshwater pulmonates) and Stylomrnatophora (terresuial pulmonates) . Mapping of these data ont0 a phylogeny of the Gasuopoda (Harsewych et al., 1997) indicates that cytosolic D- and LBHBDH have arisen a single time, in an ancestral stylornmatophoran. Al1 gasaopods of the order Stylommatophora possess this unique organization of ketone body metabolism. which is found nowhere else in the animal kingdom. MATERIALS AND METHODS Most gastropods were collected from fields and ponds near the University of Guelph, in Guelph, Ontario, Canada, or purchased (Pontcrcea, Campelorna) hm local aquarium stores. The terrestrial prosobranchs, Helicinu orbiculata, were collected near Jacksonville, Florida, USA. The giant African snails, Archachutina ventricosa, were from our laboratory population established with animals provided by the Toronto Metro Zoo. Al1 snails and slugs were kept in terraria or aquaria at room temperature (22" f 2' C) and fed lettuce. We observed no effect of duration of tirne spent under these conditions on BHBDH compamentation or enantiomeric specificity. Taxonomie classification of gastropods was as in Haszprunar (1988) (Table 8.2). A. ventricosa tissues were not fiactionated. They were prepared as in Stuart and Ballantyne (1997a). For enzyme measurements in al1 other gastropods, excised tissues were placed in approxirnately twenty volumes of sucrose buffer (100 mM sucrose, 20 mM N-[2-hydroxyethyl]piperazine-N'-[2-ethanesulfonic acid] (HEPES) , pH 7.5). Cells were disrupted by five passes of a Potter-Elvejhem homogenizer operated at low speed (< 100 rpm). This homogenate was centrifuged at 10 000 g for 10 min to separate it into mitochondrial (pellet) and cytosolic (supernatant) fia~tions. The mitochondriai pellet was resuspended in a volume of buffer equai to the original buffer addition. thus maintaining equal the dilution of both fractions. Both fractions were sonicated with three five second bursts of a Vibra-Cell sonicator (Sonics & Materiais Inc., Danbury, CT) set to 8056 output, 50 Watts. These preFaratiohs were used directly in the measurement of enzymes. We used CC0 to mark the mitochondrial membrane, CS to mark the leakage of rnatrix enzymes from mitochondria damaged in the fractionation process, and LDH to evaluate the contamination of the mitochondrial fraction with cytosolic enzymes. CCO, CS and LDH were measured as in Stuart and Ballantyne (1997b). The assays for D-,L- and DL-BHBDH contained 2 mM NAD and either 200 mM D- or L-BHB or 400 mM DL-BHB,respectively (BHB ornitted for control) in 50 mM imidazole, pH 8.0. Al1 chemicals were purchased from Sigma Chernical Co. (St. Louis, MO). RESULTS NYD DISCUSSION Activities and subcellular distributions of D- and L-BHBDH were measured in hepatopancreas and, in some cases, heart and kidney of gastropods by fractionating tissues into mitochondrial and cytosol ic fractions, and assay ing enzyme activi ties in each fraction (Table 8.1). Recovery of mitochondria in the * mitochondrial fraction" was 79% or greater for hepatopancreas from al1 species, based on the distribution of activity of cytochrome C oxidase (CCO), an exclusively mitochondrial, membrane- bound enzyme (Table 8.2). A large proportion of thwe mitochondria maintained structural integrity, based on leakage of the matrix enzyme citrate synthase (CS), of less than 23 % in all cases. The subcellular distribution of D-BHBDH approxirnated those of CC0 and CS in the hepatopancreas of ail prosobranchs and basomrnatophora with less than 28% of total activiv occurring in the cytosolic fraction. In these species, L-BHBDH activity was either not detected or was less than 6% of D- BHBDH activity. Low L-BHBDHactivities detected in Helisoma and Physa were likely the result of contaminating D-BHB,present as impurity, in the commercial L- BHB preparation (the L-BHB preparation contained up to 0.4% D-BHB). Altematively, it is possible that these low activities are due to trace levels of L- BHBDH, indicating a transitional state where both mitochondrial and cytosolic isoforms are present. The contamination of the mitochondrial fraction of the stylommatophoran Bradybcrena hepatopancreas with cytosolic lactate dehydrogenase (LDH) is low (7 1). The low L-BHBDHactivity observed in the mitochondrial fraction thus can be attributed to cytosolic contamination. Low D-BHBDH activities in Bradybuena, Arion and Archochatina hepatopancreas are Iikely due to contaminating L-BHB in the commercial D-BHB preparation (L-BHB contamination was up to 2%).As we used a [D-BHB]of 20 mM in the assays, as much as 0.4 mM L-BHB could have been present . Hean and kidney were fractionated as was hepatopancreas. Fractionation of heart tissue resulted in a lower recovery of mitochondria and greater Ieakage into the cytosol of mitochondrial matrix enzymes (Table 8.I), perhaps due to the srnaIl size of these tissues (Stagnicola hearts averaged 0.0008 g ; Pomocea hem averaged 0.01 g). Nonetheless, the distribution of BHBDH activity approximated those of CC0 and CS in these tissues, indicating that BHBDH activity is localized to the mitochondria in basommatophoran heart and prosobranch hart and kidney. In al1 of these tissues, BHBDH was specific for D-BHB,with no oxidation of L-BHB observed. The pattern of D- and L-BHBDH distribution in Archachatina (Table 8.1) parallels that seen in Cepm. In C. nemoralis tissues, no mitochondrial form of BHBDH is found (Stuart and Ballantyne, 1997b). Instead, a cytosolic L-BHBDH is present in hepatopancreas and kidney, and a cytosolic D-BHBDH is found in heart and kidney. Similarly, D-BHB is oxidized by Archachatina hem and kidney homogenates and L-BHB is oxidized by kidney and hepatopancreas. This configuration of BHBDH organization appears to be characteristic of the Stylommatophora. We investigated the evolution of cymolic BHBDH isoforms by mapping the occurrence of the mitochondrial D-BHBDH and cytosolic L-BHBDH of gasaopod hepatopancreas ont0 a gasuopod phylogeny (Harasewych et ai., 1997) using MacClade (Maddison and Maddison, 1992) (Fig. 8.1). Mollusc phylogenies based upon morphological (Haszprunar, 1988) and molecular (Harasewych et al., 1997; Winnepenninckx et al., 1994; Tillier et al., 1994; Rosenberg et al., 1994) data support the monophyly of Gastropoda, within which pulmonates are monophyletic. Within Pulmomta, the Stylommatophora and Basommatophora are each also monophyletic. For this analysis, we have considered the presence of mitochondrial D- BHBDH to be the ancestral condition. Though BHBDH activity is undetectable in marine gasnopods, it is found in al1 freshwater and terrestrial gastropods which we have snidied. Thus, the enzyme appears to have first arisen in a gastropod ancestral to these groups. Our phylogenetic analysis suggests that the cytosolic L-BHBDH isoforrn evolved a single tirne in an ancestral stylommatophoran. Gastropods of this order are almost exclusively terrestrial and comprise the vast majority of terresaial snails and slugs (Solem, 1985). To test whether the presence of L-BHBDH correlates with terresaiality, we included the distantly related terrestrial gilled gastropod Helicina (Neritomorpha) in our analysis. The absence of L-BHBDH in Helicina, however, suggests that the presence of this isoform correlates exciusively with phylogenetic position. A general upregulation of BHBDH activities appears to have occurred in tissues of pulmonate gastropods. In pulmonate hem, exceptiondly high activities of D-BHBDH (approximately two orders of magnitude greater than in Pomacea hm) (Table 8.1) suggest that D-BHB is particularly important as a metabolic substrate in these tissues. High activities of al1 enzymes of ketone body metabolism in Stagnicola elodes, C. nemoralis and A. ventricosu indicate a substantial flux through this pathway in ail pulmonates. D-BHB levels in hemolymph are as high as those of glucose in the basommatophoran pulmonate Biomphalaria glabratu (Meyer et al., 1986). These snails, and tissues isolated fiom them, actively oxidize ketone bodies. BHB appears, therefore, to be an important metabolic substrate in pulmonate snails. Unlike in mammals, however, D-BHB levels in the hemolymph decrease during starvation, suggesting ketone bodies may be used routinely by pulmonates as energy substrates, as opposed to their more specialized role during starvation in mammals. This may be related to a decreased emphasis on arnino acid metabolism and a low capacity for extrahepatic oxidation of fatty acids in pulmonates. The metabolic organization of tissues of these organisms suggests that they use ketone bodies, which are, unlike fatty acids, freely soluble, as a means of transporting lipid carbon from central stores to peripheral tissues for oxidation. In stylommatophoran pulmonates, upregulation of BHBDH activity has been followed by the elaboration of new isofonns of the enzyme. This may have occurred through an initiai loss of the trammembrane amino acid sequence from the membrane-bound mitochondriai BHBDH, to allow the enzyme to function in the cytosol. In stylommatophoran hepatopancreas and kidney, this enzyme may have been Mermodifed to act upon L-BHB, rather than D-BHB. However, this scenario for the occurrence of cytosolic BHBDH isoforms assumes divergence from the ancestral mitochondrial D-BHBDH.Alternatively, the cytosolic enzymes could derive from other proteins, unrelated to the mitochondrial D-BHBDH,and have achieved lunctionai similarity through evolutionary convergence. This appears to have occurred in the evolution of D- and L-LDH's in bacteria (Kaplan and Ciotti, 1961). However, divergence of cytosolic L-BHBDH from cytosolic D-BHBDH is suggested by the difficulty of separation of these isoforms when elecuophoresed together on a gel which separates proteins on the bases of sue and charge (Stuart and Ballantyne, 1997b). Analyses of primary structures are necessary to determine the relatedness of the three BHBDH isoforrns. The use of both enantiomers of a single metabolic substrate in routine energy metabolism is unusual in the animal kingdom, though other examples of this phenomemon exist among molluscs. Both D- and L-alanine are found in tissues of some marine bivalves. The occurrence of D-alanine appears to be related to a role in osmoregulation (Kaplan and Ciotti, 1961). Both D- and L- specific isoforms of LDH also occur within individual cephalopods (Mulcahy et al., 1997). However, the physiological significance of this has not been established. The advantages of the stylommatophoran cytosolic BHBDH isoforrns are also not immediately obvious. The existence of both D- and L- BHBDH may allow the metaboiic panitioning of BHB between specific tissues. Enzyme activities indicate that ketone bodies could be synthesized in the kidney from fatty acids under normal conditions. Both D- and L- BHBDH are present in stylommamphoran kidneys, which are thus able to produce both D- and L- BHB. Each of these enantiomers of BHB may be speciftcally targetteci to a tissue, with D- and L- BHB being oxidized by hart and hepatopancreas, respectively. This adaptation could be related to the apparently greater role for ketone bodies in the intermediary metabolism of molluscs. i.e. a refining of a much used pathway. The phylogenetic pattern of hepatopancreas BHBDH stereospecificity and subcellular distribution in gaseopods suggesu that L-BHBDH,and perhaps also O- BHBDH, could be valuable characters for assessing phylogenetic relationships within the Gasaopoda. Both enzymes can be rapidly and inexpensively assayed. The presence of hepatopancreas L-BHBDH rnay be a useful defining characteristic of the Stylornmatophora. As such, it will be of interest especially to identi€y which isoforms of BHBDH are present in tissues of the Archiopuimonata, a group of much-debated phylogenetic position. Certainly, the presence of L-BHBDH in stylommatophoran gastropods should be noted by population geneticists, as staining of electrophoretic gels of gastropod tissues with racemic DL-BHB mixtures will give results which are a hinction, in part, of the phylogenetic position and tissue of the mail examined. Table 8.1. Activities of O-hydroxybutyrate dehydrogenase (BHBDH)and marker enzymes in mitochondrial and cytosolic compartments of gastropod hepatopancreas. DL-, D- and L- BHBDH = activity measured with racemic mixture of DL-B- hydroxybutyrate, high purity D-8-hydroxybutyrate and high purity L-O-hydroxybutyrate, respectively. CC0 = cytochrome C oxidase, CS = citrate synthase, LDH = lactate dehydrogenase. "tissue" = whole tissue homogenate; "miton = mitochondrial fraction; "cytosol" = cytosolic fraction. n.d. = not detected. n. m. = not measured. Uni& are pmollminlg wet wt. Fraction CC0 CS DL-BHBDH D-BHBDH L-BHBDH LDH Genus-. Sty lonmutophora A~chachutirÜ~ hepatopancreas tissue hart tissue kidney tissue Brady bue M mito cytosol Arion mit0 cylowl Bw~nwuopIiora Physa iiiito cytosol tIdisoma niito cyt0st)l Sîagnicola hepatopancreas niito cytosol niito cytosal nii to cytosol niito cy tosol niito cytosol tidta c y tom1 kidney nii to 2.97 + 1.83 Table 8.2. Higher classification of gastropod species used for measurements of BHBDH activity. Gastro poda Orthogastropoda Neritimorpha Hel icinoidea Helicina orbicufata (terrestrid) Apogastropoda Architaenioglossa V iv iparoidea Campelorna decisurn (fteshwater) Pomacea bridgesi (fres hwater ) Heterobranchia Euthyneura Pulmonata Basommatophora Physa gyrina (freshwater) Helisoma trivolvis (freshwater) Stagnicoh elodes (freshwater) Sty lommatophora Archachatina ventricosa (terrestrial) Bradybaena similaris (terrestr ial) Arion subfuscus (terresuial) Figure 1. Gastropod phylogeny and the occurrence of mitochondrial D-8-hydroxybutyrate dehydrogenaîe (unfilled lines) and cytosolic L-0-hydroxybutyrate dehydrogenase (fîiled lines) in hepatopancreas. Hatched line = equivocal occurrence, denoting that the cytosolic enzyme arose at an undetermineci point dong this lineage. Cepaea nemoruiïs data are from Stuart and Ballantyne (1997b) Biomphutaria glabruta data are from Meyer et ai (1986). CHAPTER 9 Maximal activities of enzymes of intermediary metabolism in the estivalng terrestrial snd Cepueta nemwaiis accepted for publication in Comparative B iochemistry and Ph y siology B Abstract The effecu of estivation on maximal activities of enzymes of intermediary metabolism were studied in the terrestriai snail Cepaea nemoralis. With the exception of a 40 % decrease in citrate synthase activity in hepatopancreas, maximal activities of key enzymes of aerobic, carbohydrate, lipid, ketone body and amino acid metabolism in heart and hepatopancreas of the terrestrial snail, Cepaea nemoralis were unaltered following 6 weeks of estivation. Activities of enzymes of lipid, ketone body and carbohydrate metabotkm were reduced in kidney when expressed on a per gram wet tissue weight basis. However, estivation resulted in a near-doubling of kidney weight, though protein content of this tissue was decreased by 27%. When enzyme activities were expressed as units per mg protein, no changes were observed with estivation in kidney. We suggest that the changes in kidney weight are due to the storage of metabolic byproducts, perhaps uric acid. No change in weight or protein content of hean or hepatopancreas were observed. Our resulu suggest that the profound metabolic depression experienced during long-term estivation is rnediated by mechanisms other than down-regulation of enzyme activities. INTRODUCTION Terrestrial snails respond to conditions of environmental desiccation by withdrawing into their shells, secreting a protective epiphragm at the shell aperture. and estivating. During estivation, basal metabolic rate is depressed to values near 15% of normal (Hand and Hardewig, 1996). This profound metabolic depression allows the snails to extend the length of tirne that they can survive on stored metabolic fuels to periods of months or, in some species, years (Schmidt-Nielsen, 1971). Although metabolism in active pulmonate snails is carbohydrate based, other substrates contribute to intermediary metabol ism. This is especiall y apparent as the level of aerobic metabolism increases, as indicated by the lower RQ values of active compared with resting snails (Rees and Hand, 1990). Metabolism in estivating terresaial snails is aerobic (Rees and Hand, 1990). In medium-term estivation (up to two months), approxirnately 251 of cellular ATP is provided by lipid catabolism, based on calculations of ATP derived from measurements of consumption of stored lipid (calculatecl from (Rees and Hand, 1993)). Ketone bodies may be the form in which this stored lipid is catabolized by peripheral tissues in estivating mails. Highly aerobic tissues Iike hart have a limited capacity to oxidize fatty acids (Stuart and Ballantyne, L997a) and ketone bodies appear to be important metabolic substrates in active pulmonate mails. B- hydroxybut~~ate(BHB) and acetoacetate (Acac) are present in the hemolymph of some pulmonate snails at levels similar to glucose concentrations (Meyer et al., 1986), and high activities of enymes of ketone body metabolism are found in highly aerob ic tissues, like hepatopancreas, kidney and heart (Stuart and Bailantyne, 1997a). Here we have studied Iipid, ketone body, carbohydrate and amino acid pathways of intermediary metabolism and enzymes of the tricarboxylic acid cycle and glucose phosphate shunt in tissues of active and estivating snails. These measurements allow an assessrnent of alterations in fuel use which may occur during estivation, as reflected by changes in activities of enzymes in specific pathways relative to those in other pathways. We also have used maximal activities of key enzymes to compare the importance of ketone body metabolism in active and estivating states, and to zxamine the ability of C. nemoralis to regulate intermediary metabolism during long-term estivation through up-or down-regulation of enzyme activities. As one of the most striking changes to the intracellular environment during estivation is lowered pH (Rees et al., 1991) and since pH is a strong modifier of enzyme activities (Hochachka and Somero, 1984), we also have examined the effects of low pH to directly modulate activities of these enzymes and thus conaol flux through these pathways. MATERIALS AND METaODS Experimental Animals Cepoea nemorah were collected from fields near the University of Guelph, Guelph, Ontario in October and acclimated to laboratory conditions for approximately 5 weeks. Snails were kept in a terrarium lined with sphagnum moss which was moistened periodically to maintain arnbient hurnidity at Ievels which stimulated periodic activity. Lettuce was provided two to three times weekly, and we ensured that al1 snails were eating regularly by removing them from the top and sides of the terrarium and placing them on the lettuce provided. Lettuce uneaten after one day was removed. Under these conditions no snails were observed to estivate. A number of these snails were removed to an adjacent dry terrarium which did not contain sphagnum moss. These snails were not fed for a period of 6 weeks, during which time, the control group of snails continued to feed regularly. Al1 food deprived snails had initiated estivation within the first week, as indicated by the presence of epiphragma. Tissue preparation For determinations of maximal enzyme activities in active and estivating snails, an equai number of active and estivating individuals were removed after 6 weeks, sacrificed and deshelled. Hean, kidney and hepatopancreas were excised, puickly weighed and immediately fiozen in liquid nitrogen. After severai darj ihese tissue samples were removed from liquid nitrogen and stored at -80" C for several months. These tissues were then thawed and homogenized (in 1 mi, of buffer containing 50 mM imidazole, 5 % glycerol and O. I % Triton X-LOO, pH 7.5.) using a single 10 s burst of a Polytron Pt10 unit (Kinematica Gmbh., Luzurn, Swiaerland) at hdf speed. Homogenates were centrifuged at 200 g for ten minutes and the supernatants used directly for enzyme measurements. pH effects on maximal enzyme activities of cytosolic and mitochondrial hepatopancreas enzymes were determined on tissues taken from active snails. Excised hepatopancreas were homogenized in 2 mL of mitochondrial isolation buffer (100 mM sucrose, 20 mM HEPES, pH 7.5) with a Potter-Elvejhern homogenizer atmched to a drill press operated at low speed. Mitochondria were isolated by cenuifuging the homogenate at L0,ûûû g for 10 min. The resultant mitochondrial pellet was resuspended in 2 mL isolation buffer and this fraction and the supernatant were subjected to a 10 s sonication at 60% power output, 25 W. on a Vibra-Ce11 sonicator (Sonics & Materials Inc., Danbury, CT). Measurements of enzyme activities were made directly on these two fractions. Enzyme measurements Enzyme activities were determined using a Hewlen Packard HP8452 diode array spectrophotorneter (Hew len Packard, Miss issauga, Ontario, Canada), equipped with a thermostatted ceIl changer maintained at 20" C with a Haake D8 circulatig water bath (Haake Buchler Instruments Inc., Saddlebrwk, NJ). Rates of reactions involving NADC or NADH were followed at 340 nrn (millimolar extinction coefficient % = 6.22). Reactions involving 5-S 'dithio-bis-(2-nitrobewic acid) (Dm)were followed at 412 nrn (~~1,= 13 A). Acetoacetyl-CoA thiolase, 3- oxoacid CoA-transferase and succinyl-CoA ketotransferase were measured by follow ing the accumulation or disappearance of acetoacety 1-CoA at 303 nrn (~,,d. AI1 enzyme assays were buffered in 50 mM imidazole, adjusted to appropri~tepH with KOH and HCI. Specif'ic assay conditions were as follows: TCA cycle citrate synthase (CS) (E.C. 4.1.3.7): 0.1 mM DTNB, 0.3 mM acetyl-CoA, 0.5 mM oxaloacetate (omitted for control) . Electron transport chah cytochrome C oxidase (CCO) (E.C. 1.9.3.1): 50 pM reduced cytochrorne C (ornitted for conuol). Cytochrome C was prepared by titration with dithionite until fuliy reduced. Carbohydrate metabolism pyruvate kinase (PK) (E.C.3.7.1.40): 10 mM MgCl,, 0.4 mM NADH, 1.3 mM ADP, 2 uniu LDH, 10 mM phophoenolpyruvate (omitted for control). hexokinase (HK) (E.C. 2.7.1.1): 10 mM MgCI,, 0.16 mM NADP, 1 mM ATP, 2 units G6PDH, 10 mM D-glucose (omined for control). lactate dehydrogenase (LDH)(EX. 1.1.1.27): 0.4 mM NADH, 1 mM pyruvate (omitted for control). Lipid meta bolism 3-hydroxyacyl-CoA dehydrogenase (HOAD)(E.C. 1.1.1.35): 0.2 mM NADH, 0.1 mM acetoacetyl-CoA (ornitted for control). carnitine palmitoyl ûansferase (CPT)(E.C. 2.3.1.21): 0.1 mM Dm,50 pM 157 palmitoyl-CoA, 100 mM KCl, 5 mM L-carnitine (omitted for conuol). carnitine octanoyl transferase (COQ (E.C. 2.3.1.21): 0.1 mM DTNB, 50 pM octanoyl-CoA, 100 mM KCl, 5 mM L-carnitine (omitted for control). Ketoae body metaboüsm acetoacetyl-CoA thiolase (AAT) (E.C. 2.3.1.9): 10 mM MgCl,, 0.16 mM acetoacetyl-CoA, 0.2 mM COA (omitted for control). 3-oxoacid-CoA transferase (30AT)(E.C. 2.8.3 S): 5 mM MgCl?, 0.4 mM succinyl- CoA, 5 mM iodoaceiamide, 10 mM acetoacetate (omitted for control). succinyl-CoA ketotransferase (SKT) (E.C.2.8.3.5): 5 mM MgCL, 0.1 mM acetoacetyl-CoA, 1 mM succinate (omitted for control). D-8-hydroxybutyrate dehydrogenase (D-BHBDH)(EX. 1.1.1.30): 2 mM NAD+, 10 mM D-8-hydroxybutyrate (omitted for conuol). L-8-hydroxybutyrate dehydrogenase (L-BHBDH):2 mM NAD', 20 mM L- Bhydroxybutyrate (omitkd for control) . Amino acid metabolism glutamate dehydrogenase (GDH) (E.C. 1.4.1.2): 0.05 mM NADH, 250 mM ammonium acetate, 0.1 mM EDTA, 1 mM ADP, 7 mM a-ketoglutarate (omitted for control) . Pentose phosphate shunt glucose-&phosphate dehydrogenase (G6PDH) (E.C. 1.1.1.49): 7 mM MgCl?, 0.4 mM NADP +,1 mM glucose-6-phosphate (ornitted for control). For mwurements of maximal enzyme activities in tissues of active and estivating snails, PIC, LDH and G6PDH were measured at pH 7.1 and al1 other enzymes at pH 8.0. For determinations of pH effects on enzyme activities. cytosolic enzymes (Stuart and Ballantyne, 1996) were measured at pH 7.6 (intracellular pH in active snails) and 7.1 (typical intracellular pH in estivating snails). It is not known whether the rnitochondrial pH gradient is maintained in estivating snails. Mitochondrial pH is typically 0.5-1 .O units greater than cytosolic (Newsholme and Leech, 1983). If this gradient is maintained, then mitochondrial pH would be expected to drop to 7.6 in estivators. But, if the pH gradient is dissipated, mitochondrial pH could approach cytosolic values, or about 7.1. Thus, for mitochondrial enzymes, measurernents of activities were taken at pH 8.1, 7.6 and 7.1. Protein measurements Total tissue protein was measured in al1 tissue homogenates using the BioRad (Bio-Rad Laboratories, Hercules, CA) protein assay, standardized with bovine serum albumin. Chemicals Al1 chernicals were purchased from Sigma chemicals, Inc. (Sr. Louis. MO, USA). Statistics Statistical cornparisons of experimental groups were made using the snidents t- test. RESULTS For al1 tissues, we measured flux through the reversible reaction catalyzed by 30ATlSKT in both directions. Activity of 30ATlSKT (acetoacetate degradatiodacetoacetate production) in hmand hepatopancreas of active snails , was equal to 1.85 and 3.14, respectively. In kidney, this ratio was 0.74. No significant differences were observed in tissue weights or protein contents of hart or hepatopancreas following 6 weeks of estivation. However, in kidney a 97% increase in mean kidney weight and a 63 % decrease in protein concentration per unit tissue wet weight was observed (Tabie 9.1). This is equivalent to a statistically significant (P < 0.05) 27% decrease in kidney protein content of estivating snails when expressed as arnount of protein per whole tissue (calculation not shown). Changes in activites of HOAD, 30AT, D-BHBDH, L-BHBDH,PK and LDH were observed in kidneys of estivating smils when activities were calculated on a per gram wet weight basis (Table 9.2). However, when calculated on a per unit protein basis, no s ignificant differences in enzyme activities were apparent between active and estivating snails. No significant differences between the maximal enzyme activities of active and estivating snail hearu were observed (Table 9.3). In hepatopancreas, oniy CS activity decreased in estivators (Table 9.4). Al1 other enzyme activities were unchanged in estivating snails. Enzyme activities in hepatopancreas are affecteci by pH in vitro. When enzyme activities are measured holding assay conditions constant, while lowering pH to levels seen in estivating snail tissues (Rees et al., 1991), statistically significant differences in activity are observed for CS, SKT, PK, LDH, GDH and G6PDH (Table 9.5). In general, assay medium acidification enhances activities of enzymes of carbohydrate metabolism and suppresses activities of enzymes from pathways of ketone body and amino acid metabolism, and the pentose phosphate shunt. At pH 7.1, an 18.456 decrease in CS activity was observed. DISCUSSION Organization of ketone body metaboüsm in active snd tissues Some tissues of pulmonate gastropods have a high reliance on ketone bodies, based on the presence of high activities of ketolytic enzymes (Stuart and Ballantyne, 1997b). high levels of BHB and Acac in hemolymph (Meyer et al., 1986) and oxidation of ketone bodies both in vivo and in isolated tissues (Meyer et al., 1986). We have proposed that, in terrestriai pulrnonates, ketone bodies consumed by highly aerobic tissues, like heart, may be synthesized in the kidney (Stuart and Ballantyne, 1997b). This suggestion is based on a number of observations. BHBDH, the enzyme which interconverts Acac and BHB, occurs as two unique cytosolic isoforms in terrestrial pulmonates, each of which recognizes only one of the D- and L- enantiomers of BHB (Stuart and Ballantyne, 1997b). Kidney is the only tissue in which both isoforms of BHBDH (D-and L- BHB specific) are found, and is thus the only tissue capable of synthesizing both D- and L- BHB. In addition, ketone bodies are synthesized in ketogenic tissues primarily fiom the incomplete oxidation of fatty acids (Newsholrne and Leech, 1983), and kidney has relatively high activities of enzymes of lipid catabolism (Stuart and Ballantyne, 1997a). We have further proposed that ketogenesis in terrestrial snail tissues may occur through a reversa1 of the catabolic reaction catalyzed by 30AT (Phillips and Hird, 1977), as al1 mail tissues lack the enzyme hydroxymethylglutaryl-CoA synthase (Stuan and Ballantyne, 199%) which catalyzes an alternative ketogenic pathway. Our mode1 of ketogenesis in kidney, through the 30AT reaction, is supported by the present results. We measured activity of 30AT both in the ketolytic direction (30AT) and in the reverse, ketogenic, direction (SKT). Measurements of activity in both directions of catdysis show that kinetic characteristics of the enzyme Fdvour ketogenesis in the kidney, while in heart and hepatopancreas, kinetics favour ketolysis. This indicates that tissue-specific isoforms of the enzyme may exist. The higher ratio of SKT130AT in kidney, compared with hart and hepatopancreas, suggests that an isoform with kinetic characteristics more favourable for ketogenesis is found in kidney. Estivation effects on tissue morpbometrics Absolute amounts of protein are maintained in Cepuea nemoralis heart and hepatopancreas during six weeks of estivation in spite of the dramatic suppression of metabolic rate w hich characterizes this phys iological state. This suggests that a stabilization of proteins occurs in estivating snails, allowing the maintenance of protein levels at reduced metabolic cost. This could be achieved through parallel reductions in rates of protein synthesis and degradation, as has been demonsuated in other hypometabolic animals, Iike Artemia (Anchordoguy et al., 1993; Hofmann and Hand, 1994). However, a study of polysome profiles in active and estivating terrestrial snails provides no evidence for modified rates of protein synthesis in snails estivating for 2 months (Hobbs et al., 1994). Thus, the mechanisms whereby protein levels are maintained in estivating s~ilsare presently not understood, and it is not known if rates of synthesis and degradation are reduced. In contrast to hean and hepatopancreas, a signifiant decrease in protein concentrahm was observed in kidney Ussue, which was accompanied by a doubling of tissue weight. We speculate that increased kidney weights may have been due to the deposition of nitrogenous waste producu, like uric acid and potassium urates in this tissue, as has been reported in other estivating terrestrial snails (McNabb, 1985; Kaloyianni and Lazaridou-Dimitriadou, 1989). The reason for decreased kidney protein concentration is less clear. Al1 terrestrial snail tissues are known to undergo "tissue degrowth " when estivation is extended for very long periods of time and cellular protein stores begin to be catabolised (Rees and Hand, 1993). It is possible that this process begins earlier in kidney, perhaps as a result of the metabolic demands of niaogen metabolism placed upon this tissue. Marimal emyme activities in active and estivating snails The stable protein concentrations observed during long-term rnetabolic depression in hem and hepatopancreas are paralleled by stable activities of almost al1 enzymes. Control of metabolic flux can be achieved through modulation of maximal activities of key rate-limiting enzymes, as during exercise and starvation (Newsholme and Leech, 1983). However, the absence of modifications to activities of virnially al1 of the key enzymes of intermediary metabolism rneasured here suggests that suppressed rates of metabolic fuel use during long-term estivation are not achieved through a coordinated downregulation of enzyme activities. As maximal enzyme activities are maintahed constant in estivators, the decreased flux through pathways of intermediary metabolism mut, therefore, be achieved through ailosteric effects or other mechanisms which modulate enzyme funMion without affecting maximal activities. Two such mechanisms, which conaibüte to flux suppression in many glycolytic enzymes (Storey, 1993) are enzyme phosphorylationldephosphorylation and reversible association of enzymes with each other in the cellular particulate fraction. However, these saategies for control of metabolic flux appear to be associated primarily with the early stages of estivation (Brooks and Storey, 1990). After two months of estivation, kinetic parameters of pyruvate kinase fiom snail ventricle, which are aitered in early estivation due to enzyme phosphorylation, return to normal values (Michaelidis and Pardalidis, 1994). Similarly, glycolytic enzyme binding to the cellular particulate fraction decreases in the fwst days of estivation, but returns to control levels after 3 weeks of estivation. Thus, the participation of different mechanisms in suppressing metabolism during short- and long-term estivation is likely. Although most enzyme activities were unchanged during estivation, hepatopancreas CS activities decreased by 40 8. In accordance with this, in a related study, we have observed an 84% reduction in CC0 activity in hepatopancreas (Stuart et al., in press). The fact that CS activity declined only in hepatopancreas suggesis that the extent of metabolic depression may be tissue-specific. The hepatopancreas is both the site of metabol ite storage and metabolic interconversions , and extracellular digestion. Because digestion would be expected to case completely in estivating snails, the metabolic demands on this tissue during estivation may be particularly scaied down, perhaps resufting in a greater rnetabolic depression in hepatopancreas. Thar different tissues may undergo different degrees of metabolic depressioo &ring estivation is suggested by the fact that metabolic rate of rnantle tissue isolated nom estivating snails is depressed by only 48 56 (Pedler et ai., 1996). cornpared w?t whole organism metabolic rates, which are depressed by 84%. The focus of enzyme downregulation in hepatopancreas on CS and CC0 (Stuart et al., in press) activities indicates that mitochondria are likely important sites of metabolic regulation during estivation at least in this tissue. This 'boaom up' control of oxidative metabolism, where aerobic flux is controlled largely at the levels of the tricarboxylic acid cycle and the electron transport chain, could be a central strategy for suppression of aerobic metabolism in some tissues of estivating snails. A controlled suppression of pathway fluxes may also be mediated by changes in intracellular conditions. Entry into estivation is characterized by a sharp and rapid decrease in hemolymph and intracellular pH (Rees et ai., 1991), brought about by periodic apnea and the resultant hypercapnia (Barnhart and McMahon, 1987; Barnhan and McMahon, 1988). We investigated the effect of this acidification on enzyme activities. A decrease in pH of 0.5-1.0 units, which is typical of estivating snail tissues (Rees et al., 1991) depresses maximal acitivities in vitro of a number of enzymes, with the notable exceptions of LDH and PK, which are stimulated. The contribution of pH acidification alone, to the metabolic depression of isolated mantle tissue of the terrestrial snail Heli* aspersa, has been estimated at approximately 40% of the total depression of tissue metabolic rate (Pedler et al., 1996). Our data provide qualitative evidence that this rnight be achieved in piut through direct effecu of pH on activities of some enzymes, especially if the mitochondrial pH gradient is diminished and the pH of the matrix approaches cytosolic values. Metabolic fuel use in active and estivating snails Metabolic reorganization is a common response of anirnals faced with new metabolic demands associated with a change of physiological state (Newsholme and Leech, 1983). Reorganization of intermedias, metabolism typically manifests itself in part as modified activities of key, rate-limiting enzymes (Newsholme and Leech, 1983). However, measurements of activities of enzymes of lipid, ketone body, carbohydrate and amino acid metabolism and the pentose phosphate shunt provide no evidence for up or down-regulation of specifc pathways in any tissue. The differential rûponses CO lowered pH of enzymes of lipid, ketone body and amino acid metabolism, relative CO glycolytic enzymes, suggests an active Pasteur effect, with acidification of the cytosol serving to shift metabolism toward glycolysis. This is countered in short term estivation by reversible enzyme modifications, but many of these modifications do not appear to persist during longer term estivation (Brooks and Storey, 1990; Michaelidis and Pardalidis, 1994). Though this would suggest that metabolism becomes increasingly carbohydrate-based as estivation progresses, evidence from other studies indicates this is not the case. Respiratory quotients (RQ's) from terresnial snails indicate that, during short- term estivation, the proportional utilization of non-carbohydrate fuels is intermediate between that of active and resting nonestivating snails (Rees and Hand, 1990). Inactive, but non-estivating, snails consume aimost exclusively carbohydrates (RQ = 0.992). Activity increases the consumption of non-carbohydrate fuels (RQ= 0.932) . Assuming negligible protein catabolism, this RQ value (0.932) indicates that lip ids account for approxirnately 23 96 of metabolic fuel used. In short-term estivation, intermediary RQ values (0.953) suggest that proportional combustion of lipid is intermediary between active and inactive non-estivating snails. However, measurements of the use of stored fiel during 2 months of estivation indicate that approximately 25% of ATP production in snails estivating two months is derived from lipid catabolism (calculated based on data fkom (Rees and Hand, 1993)), which is similar to the proportional contribution of lipids observed in active snails. This suggests that lipid catabolism is increased over time during the fust two months of estivation. However, the rnechanisms by which this is achieved do not appear to include a relative upregulation of enzymes in lipid catabolic pathways or pH-mediated effects on these enzymes. Summrry and Conclusions The means by which snails supress their metabolic rates remain incompletely understood, though the participation of a number of rnechanisms is evident frorn data demonsaating the independent effects of pH (present smdy, Pedler et al.. lF%), oxygen limitation (Pedler et el., l9%), enzyme phosphorylation (Storey, 1993), reversible dissociation of enzymes from the cellular paniculate fraction (Brooks and Storey, 1990), and down-reguiation of activities of specific enzymes in some tissues (present study). It is, however, known that the metabolic depression is profound (near 85 4% of basal metabolic rate) (Hand and Hardewig, 1996), can occur quite rapidly @ours) (Barnhart and McMahon, 1987; l988), and is readily and rapidly (minutes to hours) reversible (Herreid, 19n) when environmentai conditions improve. The maintenance of most enzyme activities at normal levels may be an important component of a strategy which allows rapid emergence from estivation, and the concorninant rehirn to activity and feeding, upon a retm of favourable but possibly tram ient environmental conditions. t ndeed, a unifying characteristic of many of the biochemical phenomena associated with estivation appears to be their rapid reversibility. Table 9.1. Protein concentration of tissues (top values, n = 6 or 9) (mghg wet tissue weight) and weights of tissues (bottom values, n = 17 or 18) (mg) from active and estivating Cepaea nemoralis. Values are means f SE. ' *' = significantly different from active values (P < 0.01). The larger "n" for tissue weights reflecu the inclusion of weights from tissues used in another experiment. n.s. = not signifiant. Active Estivating % change Hem 0.25 I0.01 0.28 I0.02 n.s. 1.5 & 0.1 1.3 k 0.1 n.s. Kidney 0.083 f 0.005 0.031 f 0.002' -63% 18.5 * 1.6 36.4 * 6.3' +97% Hepatopancreas 0.046 f 0.003 0.052 * 0.002 n.s. 109.7 I 6.2 99.1 & 6.4 n.s. Table 9.2. Maximal activities of enzymes in kidneys of estivating and active C. nemoralis. Top values are pnollmin/g tissue wet weight (n = 9). Bottom values are pmollminlg tissue protein (n = 6). Values are means f SE. '*' = significantly different from active values (P < 0.025). active estivating aerobic metabol ism CS lipid metabolism CPT COT ketone body metabolism 30AT SKT D-BHBDH Carbohydrate metabolism PK LDH amino acid rnetabotism GDH pentose phosphate shunt G6PDH Table 9.3. Enzyme activities (pmol/min/g tissue weight) in hmof active and estivating Cepea nemordis. Values are means f SE (n = 9). "n.d. " = not detectable. active estivating aerobic metabol ism CS lipid metabolism CPT COT HOAD ketone body metabol ism 30AT SKT D-BHBDH carbohydrate metabol ism PK LDH arnino acid metabolisrn GDH pentose phosphate shunt Table 9.4. Maximal enzyme activities @mol 1 min 1 g tissue weight) in hepatopancreas of active and estivating mils. Values are means I SE (n = 6). ' *' = significantly different fiom active values (P = 0.0017). active (II=@ estivating (n =6) aerobic rnetabolism CS lipid metabolism CPT COT HOAD ketone body metabol km 30AT SKT AAT L-BHBDH carbohydrate metabol km PK LDH arnino acid metabolism GDH pentose phosphate shunt G6PDH Table 9.5. pH effects on activities of hepatopancreas enzymes. Values are presented as percentage of control activity (pH 8.1 for mitochondrial enzymes; pH 7.6 for cytosolic enzymes) (see text) and represent the means f SE (n = 5). Determinations of activity at different assay pH values were made on each tissue preparation. n.m. = not measured. '*' = significantly different fiom control (P < 0.02). aerob ic metabol isrn lipid metabolism COT CPT ketone body mctaboi ism SKT car bohydrate metabolism LDH amino acid metabol ism GDH pentose phwphate shunt CHAPTER 10 Remodeiing of phospholipid fatty acids in mitochondrial membranes of estivating snaiis accepted for publication in Lipids Abstract The effects of estivation on the phospholipid composition of mitochondrial membranes in the hepatopancreas of the terrestrial snail Cepaea nemoralis were investigated. Phospholipid composition was significantly altered in snails es tivating for six weeks, indicating that substantial remodeling occurs. The most profound changes occurred in cardiolipin. Cardiolipin of estivating snails was 13-fold more saturated, containeci 9-fold more monoenes, and had 45% fewer polyenes than in active snails. These differences were due, in part, to a reduction in linoleic acid (18:2n6) content of cardiolipin from estivators. As in marnmals, cardiolipin of active mils appas to preferentially incorporate 18:2n6, which accounu for 60% of the acyl chah in this phospholipid. This proportion was reduced by 50 1 in estivators. Changes in the fatty acyl content of other phospholipids of esrivating snails inciuded increased monoenes in phosphatidylethanolamine and phosphatidylinosirol, reduced ratios of n31n6 polyenes in phosphatidylethanolamine and phosphatidylcholiio, and an increased n3h6 ratio in phosphatidylserine. Arachadonic acid (20:4n6) levels were reduced in phosphatidylserine but increased in cardiolipin and phosphatidy lcholine. Taken together, these alterations a fatty acid composition are consistent with decreased biologicai activity of membrane-related processes which occur in conjunction with the reduction of mitochondrial aerobic metabolism observed during estivation. INTRODUCTION Depression of basal metabolic rate is a common physiological strategy, ailowing organisms to prolong the duration of their tolerance of sub-optimal environmental conditions (Guppy et al., 1994; Hand and Hardewig, 1996). Profound metabolic rate depress ion occurs during hibernation, torpor and estivation. As the sites of oxidative metabolism, suppression of mirochondrial respiration is a key component of strategies to reduce basal metabolic rate. Indeed, in hibernating ground squirrels, electron transfer is inhibited by 70% to 80%, at the site of ubiquinol:cytochrome c,, and adjustments of mitochondrial membrane composition are implicated in the mediation of this reduction (Brustovetsky et al., 1990). However, hibernation and torpor are characterized by reductions in both metabolic rate and body temperature, and as such it is diffcult to amibute changes of membrane phospholipid composition to one or the other of these parameters. Several authors have attempted to overcome the confounding effects of temperature and metabolic rate reduction by comparing membrane compositional changes active and quiescent overwintering animais (Farkas et al., 1984; Pruitt, 1988), thus conrrolling for responscs induced by lowered temperature. Nonetheless, interspecies differences su11 remain. Estivation, however, provides a superior mode1 for studies of this nature. In estivating organisms the temperature and species variables are circumvented, as metabolic rate reduction occurs in the absence of temperature change, and within the same otganism. And metabolic depression in estivators is at least as great as in hibernators or torpid animals. Estivating terresaial snails, for example, undergo a metabolic rate reduction of typically 851 (Hand and Hardewig, 1996). Thus, the relationship between profound metabolic rate reduction and membrane composition can be studied directly in estivators. As the functional mileu of the electron transport chain and many of the transporters and channels associated with oxidative p hosp hor y lation, the mitochondrial membranes represent a potentially powerful site for the regulation of metabol ism. M itochondrial membranes are composed principal1y of proteins and phospholipids, and the hinction of many of the proteins associated with the membranes are responsive to, and in many cases dependent upon, surrounding phospholipids (Hoch, 1992; Carruthers, 1986). Altering the composition of associated phospholipids can effect large changes in protein function (Schlame et al., 1990; Paradies et al., 1991; Hoch, 1992; Brown, 1994). Modifications to the phospholipid environment include substitution of one phospholipid species for another, and/or modification of the acyl chains of particular phospholipids. Here, we have tested the hypothesis that phospholipid remodeling of mitochondrial membranes is associated with the metabolic depression which occurs in estivating snails. We report that profound changes of the fatry acyl composition of specific mitochondrial membrane phospholipids occur during estivation, which are consistent with signifcantly reduced mitochondrial function associated with metabolic depression. MATERIALS AND METHODS Experimental animals Cepaea nenorulis, were collected in early summer, kept in terraria in the laboratory for approximately 6 weeks and fed a diet of lettuce. A group of these snails was removed to a dry terrarium and food was withheld, to induce estivation, which lasted for 6 weeks. Mitochondrial isolation Hepetopancreas from active or estivating snails were excised , and immersed in 10 volumes of mitochondrial isolation buffer (100 mM sucrose, 20 mM N-[2- Hydroxyethyllpiperazine-Nt-[2-ethanesulfoicacid] (Hepes), 0.5 % bovine serum albumin (BSA), pH 7.3, and homogenized by three passes with a Potter-Elvejhem homogenizer. Homogenates were centrifuged at 150g for 10 min and the pellet discarded. The remaining supernatant was centrifuged at 50gfor 10 min, and the pellet 'washed' by repeating this step rwice. This procedure consistently gave a recovery of approximately 85 % of mitochondria (Stuart and Ballantyne, 1996b). Purity of the mitochondrial preparation and recovery of mitochondrk -id not differ significantiy (P > 0.05) between active and estivating snails, based on the distribution of cytochrome C oxidase (mitochondrid mar ker) . peroxidase (peroxisomai marker) and proportionai content of the phospholip id sphingomyelin (used to mark non-rnitochondriai membranes, including nuclear membrane, endoplasmic reticulum, and lysosomal and plasma membranes). Extraction and analysis of mitochondnai phospholipids Total mitochondrial lipids were extracted by the method of Bligh and Dyer (1959). Phospholipids were separated from neutral lipids and each other by thin-layer chromatography. and methylated, as in Holub and Skeaff (1987). Bands were visualized using dichlorofluorescein. There were no band overlaps. An internai standard (17:O) was added to the phospholipid fatty acids prior to methylation. Individual fatty acids fiom each phospholipid fraction were separated and identified using gas chromatography, using a reversed phase DB-225 hsed silica column (J&W Scientific, FoIsom, CA) as described by Glémet and Ballantyne (1995). Total amounts of each phospholipid were determined from the summed arnounu of fatty acids in each phosphol ipid. A srna11 sarnple (1 g) of die diet (lettuce) was homogenized in 50 mM imidazole buffer using 3 X 10s bursu with a Polytron PT10 unit (Kinematica Gmbh., Lunirn, Switzerland). Lipid extraction of the homogenate was as above. Individual fatty acids were separated and quantified as above. Chemicals Chemicals were obtained from the Sigma Chernical Co. (St. Louis, MO) or Fisher Scientific Ltd. (Whitby, Ontario) and were of the highest purity available. The fatty acid standard was obtained from NU Chek Prep. Inc. (Elysian, MN) and was augmenteci Ûy the addition of a menhaden extract. Statistical un~lysis Proportionai phospholipid fatty acid compositional data were arcsine transformed and compared using students t-tests. P-values were adjusted (based on the number of tests made and degrees of fkeedorn) to compensate for the use of multiple t-tests in comparing the fatty acid compositions of each phosphoiipid from active and estivating snails (Hochberg and Tamhane, 1987). RESULTS The snail diet (lettuce) was composed prirnarily of three fatty acids, 16:0, 18:2n-6 and 18:3n-3, which combine to account for 83% of total fatty acids (Table 10.1). l8:2n-6 was the most abundant fatty acid in lethice, at almost 42 % of total. Lettuce contained a large proportion (64%)of 18-carbon fatty acids, a lower proportion (29%)of 16-carbon fatty acids, and trace amounts of 14, 20, 22 and 24- carbon fatty acids. Phosphol ipids fiom hepatopancreas mitochondr ia of active snails contained lower proportions (47%) of 18-carbon fatty acids than dietary lettuce, and greater proportions of their elongation products, especially 20-carbon fatty acids (37 %). Some of the most common fatty acids in hepaiopancreas mitochondrial phosp holipids of active snails are 18:2n-6 (18%), 18:3n-3 (82) and 20:4n-6 (18%) (Table 10.2). The fatty acid composition of mitochondr ial phosphol ip ids was al tered dramatically in estivating snails (Table 10.2). The proportion of 18:2n-6, 18:3n-3 and 18:4n-3 decreased by 60 % , 6 1% and 63 % , respective1y, in hepatopancreas mitochondria of estivating snails. 18-carbon fatty acids constituted only 30% of total fatty acids in estivators, compareci with 47% in active snails (see above). These changes were offset by greater proportions of shorter chain fatty acids (14 aiii 16- carbons) and longer chain (22-carbons) fatty acids, which increased from 10% to 18 % and fkom 5 1 to 9 96, respectively. The phospholipid fatty acids of estivating snails contained significantly more monoenes and fewer polyenes. The reduced polyene content of mitochondrial membranes from estivators resulted primarily from a Lower content of n-3 fatty acids, the proportional occurrence of which was reduced by 51%. The greatest differences between the fatty acid composition of active and estivating snail mitochondria occur in the major constituent phospholipids of the mitochondrial membranes (PC, PE and CL, which constitute 37.7 1, 35.2% and 11.4 96 of total phospholipids in mitochondria from the hepatopancreas of active snails (Stuart, Gillis and Ballantyne, in press). Perhaps the most profound alterations of fatty acid composition occurred in cardiolipin (Table 10.3). A 50 % reduction in 18:2n6 content was observed in cardiolipin which, in combination with a 57% decrease in 18:3n3, accounted for a 55%decrease in the total polyene content of cardiolipin fiom estivating, compared to active, snails. While polyene levels decreased, saturated fatty acids increased 13-fold, and monoenes increased 9-fold. This resulted in a 33 % reduction in cardiolipin unsaturation index. Unsaturation index was not altered during estivation in any other phospholipid species. However, in PE, the proportion of monoenes increased 48 %, due in part to an almost 4-fold increase in 16:1 content (Table 10.4). Membranes of estivators contained 43% fewer n-3 polyenes, largely due to significant reductions in 18-wbon n-3 fatty acids. This resulted in a lowering, by 43516, of the n-3111-6 ratio in esiivating snail mitochondria. NO statistically signifiant changes to proportions of saturates, monoenes, or polyenes were observed in any other phospholipids. However, PC showed a similar decrease (51%) in n-3 fany acid content in estivators (Table 10.5). This occurred primarily &ough significant reductions to 18%-3, 18An-3 and 20:3n-3 contents during estivation. The n-3111-6 ratio of PC from estivating snails was thus decreased by 56%. PC was the only phosphoiipid to show a change in average fatty acid chain length between experimental groups. PC fiom mitochondria of estivating snails contained longer fatty acyl chains on average than in active snails. The n-31n-6 polyene ratio appeared to be 4.5-fold greater in PS from estivating snails (Table 10.6), though a reiatively large standard error made this statistically insignifiant at a = 0.0034. The altered value of the n-3111-6 ratk was Iargely due to a statistically significant decrease of 50% in 20:4n6 and a concomitant, but not significant, 2.6-fold increase in 18:3n3. In PI, ody 20:l and 169 contents were altered in estivators (Table 10.7). Proportions of both increased, which was reflected in a doubling of the proportion of monoenes in PI, though this was aiso not significant at a=0.0034. Arachidonic acid (20:4 n-6) was present in trace arnounts in lettuce (Table 10. l), but occurred as a major constituent fatty acid of al1 phospholipids, except for CL, where it accounted for less than 4% of total fatty acids in active snails. 20:4n6 was enriched 70% in PC during estivation, while PS 20:4n-6 content decreased by 52%. DISCUSSION The phosphol ipid composition of mitochondr ial membranes from C. nemomlis hepatopancreas is ciramaticaily altered during estivation. May aspects of this remodeling indicate a reduced biological activity of the mitochondrion during estivation, when metabolic rate is reduced to about 15 56 of normal (Guppy et al., 1994; Hand and Hardewig, 1996). These changes are consistent with the inhibition of certain mitochondrial membrane-bound proteins, a stabilizing of the bilayer and a reduced propensity toward hexagonal phase for mat ion. Perhaps the most drarnatic changes to constituent fatty acids occurred in CL, a unique phospholipid which, in invertebrate animals is found exclusive1y in the mitochondrial inner membrane. CL is found in close association with certain mitodiondrid proteins, including the mono- di- and tri- carboxylate carriers. carnitine- palmitoyl translocase, cytochrome c oxidase, ADPlATP exchanger and Pi- transporter (Hoch, 1992). Many of these proteins do not function, or function sub- maximally, in its absence. Membrane content of CL is also correlated with metabolic rate. Decreased membrane CL is associated with decreased aerobic metabolism in hypothyroidism (Hoch, 1992), aging (Shineneaga et al., 1994) and atrophic muscle (Wicks and Hood, 1991). Similarly, we have observed a 27 % decrease in the proportion of CL in C. nernoralis hepatopancreas mitochondria following s Ut weeks of estivation (Stuart, Gillis and Ballantyne, in press). This well established correlation between mitochondrial membrane CL content and aerobic metabolic rate appears to be due to direct interactions of this phospholipid with proteins embedded within the inner membrane (reviewed in Hoch, 1992). In many cases, the ability of CL to stimulate protein function depends in part upon its fatty acyl composition (Hoch, 1992; Yamaoka-Koseki et al., 1991; Schlarne et al., 1990). For example, several studies have demonstrated a specific requirement of the respiratory chain enzyme cytochrome c oxidase for CL with 18:2n6 acyl chains. In rats fed dieu deficient in 18:2n6, which results in CL 18:2n6 being replaced with other fatty acids (Yamaoka et al., 1990). mitochondrial function (Yamaoka et al., 1988) and cytochrome c oxidase activity (Yamaoka-Koseki et al, 1991) are both decreased by as much as 50%.This loss of activity has been shown to result directly from the presence of 18:2n-6 deficient CL, as activity can be fully recovered when del ipidated cytochrome c oxidase is reconstituted with 18: 2n-6/l8:h- 6 CL (14). In estivating Cepaea, cytochrome c oxidase activity is reduced by 85% (Stuart, Gillis and Ballantyne, in press), and this could be mediated in part by the reduced proportion of CL and the significant reduction of CL 18:2n6 content. As in mammals, Cepea CL is particularly enriched in 18:2n6, which accounts for 60% of al1 fatty acids in CL of active snails. The 18:2n6 content of CL from estivating Cepaea decreased by 50 56. A similar decrease in CL 18:2n6 content in rats fed an 18:2n6-deficient diet resulted in a 26% decrease in cytochrome c oxidase activity (Yamaoki-Koseki et al., 1991). This suggests that both the decreased content, during estivation, of CL 18:2n6 may participate in the mediation the observed reduction of cytochrome c oxidase activity. The activities of other CL-requiring mitochondrial membrane proteins that have specific interactions with 18:2n6 acyl chahs (Hoch, 1992; Yamaoka-Koseki et al., 1991; Schlame et al., 1990) may be similarly reduced. The altered fatty acyl composition of CL also changes the molecular geometry of this phospholipid, and thus its functional properties in the membrane (Seddon et al., 1983; Sankaram et ai., 1989). CL of estivating snails is 13-fold more saturated, contains 9-fold more monoenes, and has 45% fewer polyenes than that of active snails. Thus, the unsaturation index of CL from estivating snails is 33 % reduced fkom control values. The more highly saturated CL found in estivating snails effecrs changes in the molecular geometry of CL which decrease its propensity to form non- lamellar structures, like inverse hexagonal (Hd phases (Seddon et al., 1983; Sankaram et al., 1989). H, phase-favouring phospholipids affect the structui~ond physical properties of membranes (Carruthers and Melchoir, 1986; Cullis et al., 1986; Williams et al., 1993; Epand et al., 1991). These, in turn, can modulate the hinction of specifc membrane proteins (Epand, 1990), an interaction illustrated by the requuement of a minimal proportion of hexagonal phase-preferring lipids for proper function of membrane-bound proteins, like rhodopsin in the visual system (Brown, 1994). The mitochondrial membranes of estivating snails are characterized by more highly saturated CL and an increased proportion of monoenes in PE. They also wntain lower proportions of H, phase-prefering phospholipids, PE and CL (Stuart, Gillis and Ballantyne, in press). The other major constituent of mitochondrial membranes, PC, shows an increased average acyl chah length. Al1 of these changes are in a direction consistent with the adoption of a more stable lamellar phase in the phospholipid bilayer of estivators. H, phase-prefering phospholipids are known to be important in a number of biological functions which are key to anabolic and catabolic processes, including traficking of membrane fragments and proteins, fusion, and mitochondrial contact sites (Carruthers and Melchoir, 1986; EIIens et al., 1989; Yeagle and Bentz, 1989; Ardail et al., 1990; Epand, 1990; Massari et al., 199 1). Compositional changes which lower the tendency for formation of non-b ilayer structures in the membrane appear to be related to the depressed metabolism, and therefore reduced biological activity , of estivating Cepaeo. To achieve the profound reductions in metabolic rate which characterize this state, anabolic and catabolic processes must be suppressed in concert during estivation. Such a coordinated suppression could be mediated by direct modification of membrane phospholipid composition to stabilize the bilayer and thereby reduce the rates of trans-membrane processes. It is also possible that a greater incorporation of lamellar phase-preferring phospholipids could also be a response to acidification of the cytosol, by about 0.5 units, which occurs during estivation (Rees et al., 1991; Pedler et al., 1996). Low pH is known to promote the adoption of non-lamellar phases in CL and some other phospholipids (Seddon et al., 1983; Massari et al., 1991), and modifications of membrane composition may be required to counteract this. However, the pH reduction observed in estivating snails may not be great enough to stimulate the responses found under other experimental conditions, where pH is typically iowered by 4 to 5 units from neutrality (Seddon et al., 1983; Massari et al., 1991). The ability of pH changes in the order of 0.5 units to alter phase behaviour of phospholipids, however , has not been demo wtrated. Other aspects of the phospholipid compositions of mitochondria from estivators snails are typical of those observed in association with reduced metabolic rates. The proportional content of monoenes is greater in estivating snails. This appears to be due to the replacement of polyenes, like 18:2n6 and 18:3n3, with 16: 1 and 18: 1 in CL, PE and PI. Greater incorporation of monoenes can induce looser packing arrangements in mode1 phospholipid systems (Applegate and Glomset, 1991). However , this has been demonstrated only by modeling diacylgl ycerols with a saturated acyl chain in the sn-1 position. Studies of di-rnonoenoic phospholipids give different results (Will iarns et al.. 1993). Di-monoenoic phosphol ipids are better able to form highly ordered phases than di-pol yenoic phospholipids . Thus, the neighbouring fatty acyl chain will determine how monounsaturation affects propenies of a phospholipid. The impact of increased monoene incorporation in PE, PI and CL on membrane bilayer stability is therefore uncertain without positional information. However , iricreased proportions of phosp holipid monoenes are strong1 y correlated with reduced metabolic rate in mammals an reptiles (Brand et al., 1994; Porter et al., 1996). Similarly, the higher levels of monoenoic acyl chahs in mitochondrial phospholipids may be associated with depression of metabolism during estivation. However, the mechanism underlying this correlation, rernains unknown. The n-3/n-6 ratio of membrane phospholipid fatty acids is reduced in estivating snails, primarily due to decreased proportions of 18:3n3 and 18:4n3 in PE and PC. Similarly, lower proportional contents of n-3 polyenes are characteristic of the mitochondrial membranes of anirnals with lower metabolic rates (Brand et al., 1994; Porter et al., 1996). This has been demonstrated allometrically in mammals (Porter et ai., 1996), and through comparisons of reptiles with mammals (Brand et al., 1994). In contrast, higher levels of n-3 PUFA are associated with superior recovery from post-ischemia reperfusion in rat heart (Damaison et al., 1994), when oxygen levels rnay be expected to be abnorrnally high. Thus, the significance of phospholipid n-3 content may be related to the cellular oxygen concentrations which characterize different metabolic rates and physiological states. Lower relative levels of n-3 polyenes in estivating snails thus appear to be related to the metabolic rate reduction in these animals. Arachadonic acid accounts for high proportions of the fatty acids of FE, PC, PS and PI. These levels are altered in some phospholipids of estivating snails, with the arachidonic acid content of PC and CL being increased, but reduced in PS. Notably, levels are unchanged with estivation in PE and PI. This is consistent with other studies (Vossen et al., 1993) which have found that levels of arachidonic acid incorporation into PI are defended, while they are allowed to Vary considerably in other phospholipids. This has been related to the role of arachidonic acid in signal transduction and the associated need to maintain relatively constant levels in PI for rapid cellular response to stimuli. Though the participation of mitochondrial PI in signal transduction has not bem specifidly explored, it is known that PI in mitachondria is wnfined primarily to the outer membrane (Daum, 1985), where it could be in communication with the cytosol and therefore able participate in second messenger cascades. Changes in membrane phospholipid composition induced by estivation cannot be attributed directly to the effecu of elongation or desaturation. We have observed that a significant reduction in total mitochondrial phospholipid occurs during estivation, indicating a reduction in membrane surface area (Stuart, Gillis and Ballantyne, in press). This modification is also consistent with reduced metabolic rate (Brand et al., 1992; Brand et al., 1994). However, preferential removal of specific phospholipid or fatty acid species €rom the membrane provides a mechanism for altering proportional composition which is independent of elongating and desaturating processes. The numerous changes in phospholipid fatty acid composition which we have quantifieci are consistent with a strategy of modify ing mitochondrial membrane properties in concert with metabolic rate reduction during estivation. The present results suggest that estivation provides a valuable model for study ing the rel~t innship behkreen mitochondrial membrane composition and metabolic rate. During estivation, profound reductions in basal metabolic rate, similar to or in excess of those observed in hibernation, occur in the absence of changes in body temperature. Thus, estivation appears to be a superior model to hibernation for studies of this nature, as it elirninates the confounding variable of temperature fiom the interpretation of experimental resulu. The results of the present study should be extended to vertebrate estivators, and to investigations of the functional propenies of mitochondrial and cellular membranes during estivation. Table 10.1. Percentages of individual fatty acids in lettuce (average of two Fatfy acid Amount (mole %) 24: 1 0.73 Total 100 Total Saturates 30.21 Total Monoenes 7.75 Total Polyenes 62.76 n-3 Polyenes 19.26 n-6 Polyenes 43.50 n-3/n-6 0.44 Mononeneslpolyenes O. 12 Unsaturation Index 157.33 Chain Length 17.00 Table 10.2. Percentages of individual fatty acids in hepatopancreas mitochondrial phospholipids from active and estivating C. nemordis. Active Estivating (n = 7) (n = 8) - Fatty acid 14:O 0.97 * 0.40 2.41 * 0.62 24: 1 n.d. n.d. Total 100 100 Total saturates 18.46 i1.00 23.01 * 2.00 Total Monoenes 14.52 * 0.75 19.85 * 1.35' Total Polyenes 67.02 * 0.91 56.96 * 3.30' a-3 Polyeiies 15.68 I0.81 7.99 I0.62' n-6 Polyenes 51.33 & 0.88 48.96 * 2.74 n-3111-6 0.31 * 0.02 O. 16 * 0.01' Monoenes/ Polyenes 0.22 f 0.01 0.37 * 0.05 Unsaturation index' 222.45 I 3.86 213.78 * 10.21 Chain lengthZ 18.36 fi).05 18.52 f 0.16 Values are ~resentedas means + SE. n.d. = not detectable. ' = ~ignififantl~different fiom-active values, a = 0.0034. = Unsaturation Index = Zrn) 9;where miis the mole percentage and n, is the number of C-C double bonds in fany acid ' i' . = Mean Chain Length = Zf- T~;where fi is the mole fraction and ci is the number of carbon atoms in fany acid 5' . Table 10.3. Percentages of individual fa acids in cardiolipin of hepatopancreas mitochondria €tom active and estivatinp, Fnemoralis. Active Estivating - Fatty acid 14:O 0.27 * O. 14 5.26 & 2.35 14: 1 0.49 f 0.43 2.60 f 0.87 16:O 0.86 * 0.23 9.07 & 2.46 16: 1 0.78 f 0.42 12.18* 3.64 18:O 0.60 * 0.31 5.96 & 2.41 18: 1 1.32 f 0.47 5.59 * 2.06 18:2n-6 60.19 * 1.67 29.68 * 5.84' 18:3n-3 20.58I 0.87 8.85 f 2.04' 18:4n-3 8.57 I1.45 4.07 f 1.64 20:O n.d. n.d. 20: 1 0.08 * 0.04 2.70 f 0.65' 20: 211-6 0.85 & 0.08 1.31 & 0.29 20: 3n-6 0.71 I0.13 1.25 f 0.17 20:4n-6 3.67 i0.27 5.11 f 0.98' 20: 311-3 0.63 f 0.08 n.d. 20:4n-3 n.d. n.d. 20: 511-3 0.18 f: 0.07 1.14 f 0.89 22:O n.d. n.d. 22: 1 nad. n.d. 22:2n-6 n.d. 0.11 * 0.11 23:O O. 11 * 0.05 3.33 f 1.47' 22:4n-6 n.d. n.d. 22:Sn-6 n.d. n.d. 22: 51-3 n.d. 0.19 f 0.19 22:6n-3 n.d. 0.34 * 0.20 24:O n.d. n.d. 24: 1 n.d. nad. Totai 100 100 Total saturates 1.85 I0.48 24.75 * 6.29' Total Monoenes 2.67 * 0.64 23.07 I4.69' Total Polyenes 95.48 & 1.11 52.18 * 9.40' n-3 Polyenes 29.99 f 1.43 14-58 i: 3.98 n-6 Polyenes 65.49 f 1.85 37-60 * 6.34' n-3/n-6 0.46 f: 0.03 0.37 f 0.09 MonoeneslPol yenes 0.03 f 0.01 0.81 * 0.31 Unsaturation Index1 240.76 * 2.91 161.64 f 22.52' Chain Length2 17.23 * 0.04 17.16 * 0.20 Values are presented as means f SE, ad. = not detectable. = Significantly different fiom active values, a = 0.0034. = Unsaturation Index = Lq a;where m, is the mole percentage and n, i~ the number of C-C double bonds in fatty acid 'i'. = Mean Chain Length = T~;where f, is the mole fraction and ci is the number of carbon atorns in fatty acid 'i' . Table 10.4. Percen es of individual fatty acids in phos hatidylethanolamine of hepatopancreas mitoc3 ondria from active and estivating 8. nomoraiis. Active Estivating (n = 7) (n = 8) Fatty acid 14:O 1.74 * 0.76 2.37 * 0.71 14: 1 1.30 * 0.36 2.00 * 0.64 16:O 2.56 f 0.20 4.70 * 0.71 16: 1 1.67 * 0.34 6.25 * 0.84' 18:O 15.28 * 0.90 9.36 f 0.58' 18: 1 4.76 * 0.24 3.84 * 0.62 18:2n-6 10,41 * 0.79 5.10 f 0.40' 18:3n-3 4.06 f 0.22 1.24 i 0.23' 18:4n-3 1.26 f 0.19 0.43 f 0.07' 20:O 0.45 * O, 11 0.57 * 0.09 20: 1 3.26 * O. 15 4.20 * 0.23 20: 2n-6 7.91 * 0.17 10.14 * 0.58 20:3n-6 3.60 & 1.21 5.68 I0.77 20:4n-6 27.59 * 1.87 28.49 I 1.71 20:3n-3 3.42 * 0.63 1.10 f 0.30 20:4n-3 0.08 * 0.08 0.74 I0.27 20:s n-3 2.34 =f= 0.28 2.20 k 0.42 22:O n.d. n.d. 22: 1 n.d. n.d. 22:2n-6 n.d. n.d. 23:O 0.82 * 0.09 1.37 f 0.16' 22:4n-6 2.12 f 0.07 3.09 f 0.18' 2251-6 4.58 * 0.29 5.99 * 0.17 22:5n-3 0.31 * 0.08 0.29 * O, 13 22:6n-3 0.45 * 0.02 0.85 * 0.14 24:O n.d. n.d. 24: 1 n.d. n.d. Total 100 100 Total saturates 20.85 f 1.62 18.37 I 1.45 Total Monoenes 10.99 * 0.21 16.29 f 1.51' Total Polyenes 68.16 f 1.49 65.34 * 2.71 n-3 Polyenes 11.93 f 0.87 6.84 f 0.49' n-6 Polyenes 56.22 f 1.26 58.50 f 2.33 n-3/n-6 0.21 & 0.02 0.12 * 0.01' MonoenedPoI yenes 0.16 * 0.01 0.26 f 0.04' Unsaturation Index1 243.85 & 6.90 247.84 * 8.22 Chain Length2 18.91 * 0.07 18.97 I 0.11 Values are presented as means * SE, n.d. = not detectable. = Significantly different nom active values, a = 0.0034. = Unsaturation Index = ZmJ q;where mi is the mole percentage and 4 is the number of C-C double bonds in fattv acid 3'. = Mean Chain Length = Zfi 7;bhere f, is the mole fraction and ci is the number of carbon atoms in fatty acid ' i'. Table 10.5. Percentages of individual fatty acids in phos hatidylcholine of hepatopancreas mitochondria From active and estivating 2' nemoralis. Active Estivating (n = 7) (n = 8) Fatty acid 14:O 0.45 * O. 14 1.18 & 0.63 14: 1 1.95 2 1.25 0.98 * 0.45 16:O 7.32 * 0.38 6.64 * 0.28 16: 1 1.45 * 0.56 2.57 & 0.68 18:O 2.38 1: 0.12 4.06 f 0.40' 18: 1 13.93 f 0.88 11.22 * 1.14 l8:2n-6 19.69 * 1.05 9.09 f 0.62' 18:3n-3 8-14 * 0.44 2.37 * 0.50' 18:4n-3 2.83 f 0.47 1.00 f 0.15' 20:O 0.32 * 0.03 0.50 * O. 10 20: 1 3.37 i: 0.39 4.60 * 0.40 20: 211-6 12.62 f 0.42 16.11 f 0.64' 20: 3n-6 2.53 & 0.44 4.58 * 0.67 20:4n-6 12.36 f 0.59 21.01 * 0.91' 20: 3n-3 3.77 * 0.44 1.70 & O. 12' 20:4n-3 n.d. O. 12 * 0.09 20:Sn-3 1.35 * 0.20 0.90 * 0.15 22:O n.d. n.d. 22: 1 n.d. n.d. 22:2n-6 n.d. n.d. 23:O 0.21 f 0.18 0.10 * 0.05 22 :4n-6 0.39 * 0.03 0.60 * 0.04 2251-6 3.65 f 0.22 8.24 * 0.66' 22:Sn-3 n.d. n.d. 22:6n-3 1.10 * 0.16 2.35 f: 0.58 24:O n.d. n.d. 24: 1 n.d. n.d. To ta1 100 100 Total saturates 10.77 * 0.44 12.48 * 1.13 Total Monoenes 20.70 & 1.30 19.38 I1.50 Total Polyenes 68.53 + 1.64 68.14 I2.06 n-3 Polyenes 17.28 * 1.16 8.50 f 0.67' n-6 Polyenes 51.26 * 1.59 59.64 * 2.22 n-3/n-6 0.34 * 0.03 0.15 f 0.02' MonoenedPolyenes 0.31 * 0.03 0.29 & 0.03 Unsaturation Index1 222.82 I3-90 246.51 I6.74 Chain LenM 18.33 *0.08 19.03 O. 17' Values are ~resentedas mean1 = Signifiktly different from-active values, a = 0.0034. l = Unsaturation Index = Zmj 3;where miis the mole percentage and q is the number of C-Cdouble bonds in fatty acid 'i'. = Mean Chain Length = Zfi T~;where 4 is the mole fraction and ci is the number of carbon atoms in fatty acid 'i' . Table 10.6. Percentaees of individual fat& acids in phosphatidylserine of hepatopancreas rnitoc%ondria from active -ad es tivahg C nemoralis . . Active Estivating Fatty acid 14:O 0.45 0.24 3.07 f: 1.54 24: 1 n.d. n.d. Total 100 100 Total saturates 34.60 f 0.89 33.91 f: 5.58 Total Monoenes 20.10 f 1.60 20.81 * 4.62 Totai Polyenes 45.30 f 1.34 45.29 I9.87 n-3 Polyenes 12.64 f 0.85 27.13 * 9.94 n-6 Polyenes 32.67 * 1.64 18.16 I2.31' n-3 111-6 0.40 * 0.04 1.84 * 0.70 MonoenesPolyenes 0.45 * 0.04 0.75 * 0.20 Unsaturation Index' 181.18 * 3.97 167.04 * 23.87 Chah Length2 18.24 * 0.09 16.93 & 0.42 Values are oresented as means f SE. n.d. = not detectable. = Signifiktly different fiom-active values, a = 0.0034. l = Unsaturation Index = Zmj q;where miis the mole percentage and ni is the nurnber of C-C double bonds in fatty acid 'i'. = Mean Chain Length = Z$-Iî; where f, is the mole fraction and ci is the nurnber of carbon atoms in fatty acid i . Table 10.7. Percenta es of individual fatty acids in phos hatidylinositol of hepatopancreas mitocfi ondria hmactive and estivating 8. nemordis. Active Estivating (n = 7) (n = 8) Fatty acid 14:O 0.80 * 0.45 4.68 * 1.65 ' 14: 1 0.62 * 0.32 1.39 * 0.70 16:O 24.07 f 0.94 19.05 I: 2.36 16: 1 2.61 * 0.58 7.89 f 1.51' 18:O 26.45 * 0.89 23.67 * 1.18 18: 1 4.95 1.00 7.46 * 1.05 18:2n-6 2.32 & 0.30 1.05 k 0.35 18:3n-3 4.21 I0.26 2.67 i0.41 18:4n-3 1.12 * 0.16 0.94 * 0.45 20:O 1.16 f 0.10 0.93 * 0.20 20: 1 2-60 =t O. 15 5.36 i 0.73' 20:2n-6 1-63 I0.24 1.57 i: 0.20 20: 3n-6 2.52 * 0.49 2.48 * 0.57 20:4n-6 22.59 & 0.77 18.03 I2.68 20: 3n-3 1.01 I; 0.26 0.51 * 0.27 20:4n-3 n.d. n.d. 20511-3 0.67 I0.11 0.43 * 0.17 22:O n.d. n.d. 22: 1 n.d. n.d. 22:2n-6 n.d. n.d. 23:O 0.20 & 0.10 0.19 O. 16 22:4n-6 0.04 * 0.04 0.14 * 0.6 22:Sn-6 0.32 f 0.13 1.27 * 0.42 22:Sn-3 n.d. n.d. 22: 6n-3 0.09 * 0.06 0.30 * 0.12 24: O n.d. n.d. 24: L n.d. n.d. Total 100 100 Total saturates 52.68 f 1.64 48.52 I2.89 Total Monoenes 10.78 f 1.41 22.10 I2.31 Totai Polyenes 36.54 f 1.58 29.38 I: 4.05 n-3 Polyenes 7.10 f 0.37 4.85 f 0.88 n-6 Polyenes 29.44 f 1.31 24.54 * 3.42 n-3/n-6 0.24 f 0.01 0.20 f 0.04 MonoenesPol yenes 0.30 1: 0.04 0.98 * 0.29 Unsaturation Index1 142.42 * 4.74 131.04 * 14.64 Chain Length2 17.86 * 0.05 17.62 * 0.25 Values are presented as means * SE, n.d. = not detectable. ' = Significantiy different from active values, a = 0.0034. ' = Unsaturation Index = ZmI 3;where miis the mole percemage and ni is the number of C-C double bonds in fatty acid 'i'. = Mean Chain Length = Zf -cg; where f, is the mole fraction and ci is the number of carbon atorns in fatty acid 'L'. CHAPTER II Compositional correlates of metaboüc depression in mitochondrial membranes of an estivating mail, Cepaea nemmalis accepted for publication in The American Journal of Physiology (to be published before Chapter 10) Abstract The phospholipid and protein compositions of mitochondrial membranes from hepatopancreas of active and estivating terrestrial snails, Cepaea nenorulis, were compared. Mitochondria from estivating snails contained 82.7% less cardiolipin and this was associated with an 83.9 56 reduction in cytochrome c oxidase activity. Substantial changes also occurred in the proportional amounts of other individual phospholipid classes and theu consituent fatty acids, including a 72% loss of total mitochondrial phospholipids, a 37% increase in monoenes and 4956 fewer n-3 fatty acids in membranes of estivating snails. These changes are consistent with those correlated with lowered metabolic rate, and lower rates of proton le&, in other animai models. Estivating snail hepatopancreas showed no change in total phospholipid content, indicating that the phospholipids lost from mitochondrial membranes may be sequestered elsewhere within the cell. We suggest that estivating snails rernodel mitochondrial membranes as part of a coordinated, reversible suppression of mitochondriai membrane-associated processes, which may include a concornmitant reduction in rates of proton pumping and leaking. INTRODUCTION Slowing of metabolic rate is a defining characteristic of such physiological States as fiacultative anaerobiosis, hibernation, torpor, dormancy and estivation (Guppy et al., 1994). Estivation occurs in a number of fish and amphibian species, but is perhaps best characterized in terrestriai snails, where it occurs in response to desiccating environmental conditions. In estivating land snails, basal metabolic rate (BMR) is depressed to about 16 96 of standard resting rates (Hand and Hardewig, 1996). In this state, the snails can survive for periods of rnonths or years until a return of favourable environmental conditions (Schmidt-Nielsen, 197 1). Metabolism in estivating snails is aerobic, with minimal recruitment of anaerobic pathways (S torey, 1993). Thus, mitochondrial oxidative metabolism continues, but at a significantly reduced rate. Metabolic rate reduction in estivating snails must, therefore, involve a controlled suppression of mitochondrial aerobic metabolism, though this has not been studied directly . The function of the mitochondrion is largely organized within and around the mitochondrial membranes, which provide the milieu within which the elecaon transport chain (ETC), and related enzymes and transporters, operate. Specific membrane phospholipids, like cardiolipin, are required for the optimal function of a wide variety of mitochondrial membrane-bound transporters and enzymes (Yeagie, 1992; Hoch, 1992). Physical properties of the bulk bilayer also directly affect the function of many membrane-bound proteins (Carruthers and Melchior, 1986; Epand, 1990; Gordon and Mobley, 1985). Modulation and control of the activities of mitochondrial membrane-bound proteins may thus be achieved through modification of the membrane phospholipid composition. Estivation offers a unique opportunity to study such compositional modifications, as they relate to metabolic depression. Unlike during hibernation and torpor, the depression of BMR in estivators occurs in the absence of temperature change. Unlüce in facultative anaerobes, anaerobic pathways do not appear to play a signifiant role in maintaining ATP levels (Storey, 1993). We have tested the hypothesis that mitochondrial function and phospholipid composition are altered during estivation in the terrestrial mail Cepaeu nemoralis in a manner consistent with a reduction in mitochondrial oxidative metabolism and the observed lowering of whole animal BMR (Herreid, 1977; Rees and Hand, 1990; Storey, 1993; Guppy et al., 1994; Hand and Hardewig, 1996). MATEIUAES AND MIETHODS Cepaea nemoralis, were collecteci in early summer, kept in terraria in the laboratory for approximately 6 weeks and fed a diet of lettuce. A group of these snails was removed to a dry terrarium and food was withheld, to induce estivation. Estivation, identified by the presence of a calcareous epiphragm at the shell aperture, lasted for 6 weeks. Hepatopancreas fiom active or estivating snails were excised, weighed, and immersed in 10 volumes of mitochondrial isolation buffer (100 mM sucrose, 20 mM N-[2-Hydroxyethyl1piperazine-N' -[2-ethanulonic acid] (Hepes), 0.5% bovine serum albumin (BSA), pH 7.5), and homogenized by three passes with a Potter-Elvejhem homogenizer . Homogenates were centrifuged at 150g for 10 min and the pellet discarded. The remaining supernatant was centrifuged at 5000g for 10 min, and the pellet 'washed' by repeating this step twice. The mitochondrial pellets were immediately frozen at -20°C.They were thawed prior to lipid extraction approximately 24 h later. Purity of the mitocbondriai preparation This protocol for isolation of intact mitochondria is similar to those used in other studies of C. nemoralis (SM and Ballantyne, 1996b). This procedure consistentiy gives a recovery of approximately 85% of mitochondria. Equal recovery of mitochondria fiom hepatopancreas of active snails and estivators is indicated by similar recoveries of cytochrome c oxidase activity (82.4 i 5.1 1and 88.8 f 4.0 % , respectively, of whole homogenate activity ; n = 4; P > 0.05). Peroxisomal contamination was assessed using measwments of peroxidase activity. Only 3.5 * 1.0 % of homogenate peroxidase activity was recovered in the mitochondrial fraction from active snails, and 3.8 f 0.8% in estivating snails. These values were not significantly different (P > 0.05: n = 4). The amount of sphingomyelin in the mitochondrial Fraction cm thus be us4 to indicate the presence of non-rn itwhmdrial membrane Fragments, since mitochondria contain only trace amounts of sphingomyelin, which is localized largely in the outer membrane (Daum, 1985). Nuclear membranes, endoplasmic reticulum, and lysosomal and plasma membranes al1 have much higher proportional contents of sphingomyel in (Daum, 1985). The proportional content of sphingomyelin in the mitochondrial fraction was 2.4 I 1.S % in active snails and 4.2 f 1.5% in estivators. These values were not significantly different (P > 0.05; n = 7 and 8). Lipid extraction and analysis Total mitochondriai lipids were extracted by the method of Bligh and Dyer (1959). Phospholipids were separated from neutral lipids and each other by thin-layer chromatography as in Holub and Skeaff (1987). Fatty acids were methylated by scraping individual phospholipid bands into glas kimex tubes containing 2 mL of 6% H2S04, in methanol and 10 pg of heptadecanoic acid. The tubes were tightly sealed with Teflon-lined caps and vortexed for 60 s, then maintained at 80' C for 2 hrs. Petroleum ether (2 mL) was then added to the cooled tubes and each tube vortexed for 60 S. Double-distilled H,O (1 mL) was added and the tubes vortexed again for 30 S. The upper petroleum ether phases, were aansferred to glas mini vials. Individual fatty acids nom each phospholipid fraction were separated by gas chromatography as described by Glémet and Ballantyne (Glérnet and Ballantyne, 1995). Petroleum ether samples containing fatty acid methyl esters were dried under oxygen-free nitrogen gas and redissolved in 2581 carbon disulfide (CS&. CS, samples (2 pl) were injected into a gas chromatograph (Hewlett-Packard, HP5890 series II) fitted with a flame ionization detector and an automatic injector (Hewlett-Packard, 7673A). Fatty acid methyl esters were analyzed on DB 225 rnegabore fused silica column (Chrornatographic Speciaities Inc., Brockville, Ontario, Canada) at 2 10°C for 30 min, which included an initial ramping from 150 to 210°C over the first minute. Fatty acids were identified by comparison of retention times fiom a known standard containing al1 fatty acids of interest. Chain Iengths shorter than C: 14 were not resolved under these conditions and therefore not reponed. Total amouna of each phospholipid were determined from the summed arnounts of fatty acids in each phospholipid. Absolute arnounts of individual fatty acids were determined by comparison with a known concentration of an interna1 standard, heptadecanoic acid (17:0), added to the samples before the methylation process. Measurements of enzyme activities and protein Cytochrome c oxidase (CCO) was measured as described by Stuart and Ballantyne (Stuart and Ballantyne, 1996b). Briefly, oxidation of fully reduced cytochrome c (50 pM) by homogenate in 50 mM imidazole (pH 8.0) buffer, was followed at 550 nm using a Hewlett Packard HP8452 diode array spectrophotometer (Hewlett Packard, Mississauga, Ontario, Canada), equipped with a thermostatted ce11 changer maintaineci ai 20°C with a Haake D8 cuculating water bath (Haake Buchler Instruments Inc., Saddlebrook, NI). Peroxidase activity was measured at 240 nm, after addition of 30 mM peroxide (H,OJ to diluted homogenate in 50 rnM imidazole (pH 7.0). Protein was determined on aliquots of thawed samples using the BioRad (Bio- Rad Laboratories, Hercules, CA) protein microassay. Determinaiion of total hepatopancrers phosphoiipids Total hepatopancreas membrane phosphol ipids in active and estivating individuals were measured in a second group of mils kept under the same &matory conditions as above. For these snails, whole hepatopancreas were homogenùed in 2 mL of 50 mM imidazole, pH 7.5, homogenized with three ten second bursts of a Polytron PT10 unit (Kinematica Gmbh., Luzurn, Switzerland). Phospholipid extïaction was as described above, with the exception that the phospholipid fraction was separateci oniy into cardiolipin and al1 other phospholipids. Further analysis was mied out by gas chromatography as in the fist experiment. S tatisticai anaiyses Absolute data values were compared using Student ' s t-tests. Proportional phospholipid and fatty acid compositional data were arcsine transformed and then compared using Student 's t-tests. RESULTS No sig nificant differences were observed between active and estivating snails in hepatopancreas weight or protein content (T'able 11.1). Similarly, the protein content of mitochondrial fraction isolated from hepatopancreas tissue was unchanged following SU weeks of estivation. Though the lack of change in protein content suggests that the absolute amounts of most enzymes may have remained constant in estivating snails, the activity of the respiratory chain enzyme, cytochrome c oxidase, was reduced by 83.9%. In contrast to the unchanged protein levels, mitochondrial membrane phospholipid composition was altered significantly in estivators. Perhaps the most dramatic changes were a 71.7% decrease in tord mitochondrial phospholipid content of hepatopancreas from estivating Cepaea and an 82.7 % reduction in cardi01 ipin content (Table 11.1). Despite the loss of phospholipid from the mitochondrial fraction, there were no changes in the total phospholipid content of whole hepatopancreas tissue with estivation (Table 11.1). Similarly, no significant difference between groups was observed for whole tissue cardiolipin content. Relative proportions of mitochondrial phospholipids were also altered in hepatopancreas of estivators (Table 11.2). Reductions occurred in proportional content of the hexagonal p hase-preferr ing p hospholipids, cardiolipin and phosphatidylethanolamine, of 35 56 and 14% respectively. These were accompanied by an 89% increase in the proportion of phosphatidylinositol in the mitochondrial membranes (Table 11.2). The fatty acid composition of mitochondrial phospholipids (pooled) was also altered in estivators (Table 11.3). Mitochondriai membranes of estivating snails contained greater proportions of monoenes and fewer polyenes. This difference was reflected in the monoenes/polyenes ratio, which increased by 68 56. The proportional content of n-3 polyenes was reduced in estivating snails, which lowered the n-3111-6 ratio by half. No signifiant differences between active and estivating snails were observeci for proportions of saturated fatty acids, unsaniration index or average fatty acid chain length. DICUSSION These data indicate that substantial changes to mitochondrial structure and hinction occur in estivating C. nemordis. The general suppression of oxidative metabolism which accompanies estivation in terrestrial snails is reflected by a significant reduction in CC0 activity. The changes in membrane phospholipid composition are also consistent with a reduced mitochondrial function. Reductions in CC0 activity in hepatopancreas of estivating snails could be explained by a reduction in the number of mitochondria in hepatopancreas. However, both whole tissue and miiochondrial protein levels were similar in active and estivating snails. Thus, while the protein content of mitochondria appears io be unchanged during estivation, substantial reductions ofcur in mitochondrial phospholipid content. This suggests ihat the downregulation of metabolic flux in mitochondria does not occur through wholesale reductions in number of mitochondria or enzyme concentrations. In estivating snails, post-translation modifications, such as reversible phosphorylation, have been shown to be a particularl y common means of regulating activities of soluble enzymes during metabolic depression (Guppy et al., 1994; Hand and Hardewig, 1996; Storey, 1993). However, as many of the enzymes involved in oxidative metabolisrn are not soluble, but membrane-bound, an alternative regulatory strategy is available for their regulation. Modulation of the lipid environment of membrane-bound proteins may be one rnechanism for their regulation during rnetabolic depression. The activities of many membrane-bound proteins are responsive to changes in their phospholipid environment (Yeagle, 1992; Hoch, 1992; Robinson, 1993). Cardiolipin, in particular , is found in close association with a nurnber of mitochondrial inner membrane proteins, and in some cases has been shown to directly affect activity (Hoch, 1992). For exarnple, cytochrome c oxidase, the mono, di- and tri-carboxy 1ic acid transporters, carnitine acy l-tram ferases, ATPase, ADPl ATP exchanger and Pi transporter dl have absolute or partial requiremenu for cardiolipin to achieve maximal activities (Hoch, 1992). The drarnatic reduction in cardiolipin content of estivating Cepoeu mitochondria (82.7%) which occurs in conjunction with a nearly identical (83.9%) reduction in cytochrome c oxidase activity, and corresponds closely to the approximately 84% reduction in BMR observed in several species of estivating pulmonate snails (Guppy et al., 1994; Hand and Hardewig, 1996; Herreid, 1977; Rees and Hand, 1990; Storey, 1993), is therefore an interesting result. Activities of many enzymes and transporters in the mitochondrial imer membrane could be reduced in a cwrdinated fashion simply by rernoving cardiolipin hiii the membrane. For instance, the observed reduction in CC0 activity in estivating snails could be mediated through the removal of cardiolipin from the mitochondrial inner membrane. A number of studies have dernonstrated the direct dependence of CC0 on cardiolipin for activity (Robinson, 1993, reviewed in Hoch, 1992). Inhibition of ETC activity through modifications of phospholipid composition is also suggested b y studies of mitochondr ia fiom hibernating squirrels (Brustovetsky et al., 1990). Electron transport in hibernating squirrels is inhibited by 70-8046 at ubiquinol:cytochrome. This inhibition is reversible through hypoosmotic swelling, which activates phospholipase &. However , inhibition of phosphol ipase A, dur ing swelling prevents the recovery of electron transfer. In the absence of membrane compositional data, the authors proposed that general adjustments to membrane fluidity may be involved in mediating the reduction of electron transfer. Our results indicate some of the specific compositional changes which may occur to accomplish this. In addition to the specific effects of cardiolipin on protein activities, properties of the bulk membrane, such as phase behaviour and fluidity, also have been shown to modulate protein function. The mitochondrial membranes of es tivating Cepaea contain lower proportions of the hexagonal phase-preferring phospholipids, cardioiipin and phosphatidylethanolarnine, and unchanged or greater proportions of more bilayer-stable phospholipids, including phosphatidylcholine, phosphatidy lserine and phosphatidylinositol. Hexagonal phase-preferring lip ids are associated with a number of active processes (reviewed in Epand, IWO), including aans-membrane trafficking, mitochondrial contact sites and stimulation of some enzyme activities (Carruthers and Melchior, 1986). Thus, the lower proportion of these phospholipids suggests reductions in these processes dur ing estivaiion. The observed changes in mitochondrial phospholipid composition may also be related to reductions in the permeability of the imer membrane to protons. As mitochondria frorn al1 animals are semi-permeable to protons, a futile cycle of outward proton pumping and inward proton leak exists across the imer membrane (Brand et ai., 1994). Leakage of protons fiom the cytosol to the mitochondriai matryt accounts for a considerable energy expenditure in animal cells. In mammals, the cost of this proton leak is estimated at between 20% (Rolfe and Brown, 1997) and 25 % (Porter et al., 1996) of BMR, and this proportion appears to be similar in ectotherms, based on similarities in the properties of lizard and mammalian hepatocytes (Brand et al., 1994). Estivating land snails consistently achieve BMR reductions of 84%. which occur in the absence of anaerobic metabolism (Rees and Hand, 1990). Thus, assuming that mitochondrial proton leak incurs a similar cost as a proportion of BMR in snails as in other ectotherms, for BMR to be lowered to 16 % of normal during estivation, a concommitant and significant reduction in the rate of proton leak must occur. - The mechanism of proton leak in most tissues rernains unknown. It does not occur direcdy through a protein channel (Brand et al., 1994), nor across the phospholipid bilayers of liposomes which contain no protein (Brookes et al., 1997). Nonetheless, reductions in proton leak correlate well with some mitochondrial phospholip id compos itional parameters, suggesting a role for specific p hosphol ip id fatty acids in determining proton permeability of the intact membrane. Specifically, two properties of the imer membrane, a decrease in the total arnount of phosphol ipids , and therefore surface am, and reduced inner membrane leakiness per unit membrane surface ara, are correlates of less proton permeable mitochondrial membranes (Brand et al., 1991; Brand et ai., 1992; Brand et al., 1994; Porzr et al., 1996). The modifications to fatty acid compositional of mitochondr iai p hospholip ids of estivating mails are consistent with both of these mechanisms of leak reduction. Decreased mitochondrial inner membrane surface area correlates with reduced proton leak in hypothyroidism (Brand et al., 1992). Sirnilarly, in allometric studies of marnmalian mitochondria, differences in inner membrane surface ara explain two- thirds of the large differences in proton leak between mitochondria from mammals of different sizes (Porter et al., 1996). Other studies (Brand et al., 1992) have demonstrated that total phospholipid content is approximately proportional to membrane surface area. As the convoluted inner membrane accounts for a rnuch greater proportion of totai mitochondrial phospholipids than the outer membrane, the reduction in total phospholipids in Cepaea mitochondria should reflect a decrease in the surface area of the imer membrane. Thus, the observed 72% reduction in total mitochondrial phospholip ids in estivating Cepaea may reflect a similar reduction in membrane surface are and thus a corresponding reduction in proton leak. Two phospholipid fatty acid parameters are also modifed in manners which suggest decreased proton leak per unit membrane surface. The proportion of monounsaturateci fatty acids, which tends to be higher in organisms with lower rnetabolic rates and less leaky membranes (Brand et al., 199 1; Brand et al., MM), increases 37 56 in mitochondrial membrane phospholipids of estivating snails relative to active ones. Similarly, the proportion of n-3 polyunsaturates correlates positively with membrane leakiness and metabolic rate (Brand et al., 1991; Porter et al., 1996), and a 49% decrease in the proportion of n-3 fatty acids in mitochondrial membranes is observed in estivating snails. These observations are both consistent with a decreased leakiness per unit surface area of membranes fiom estivators. Thus, both the reduction in mitochondrial membrane surface area and an altered fatty acyl composition of the remaining membrane phospholipids are consistent with a reduction of ion Ieak in these membranes. The substantial reduction observed in CC0 activity, and the adoption of membrane characteristics associated with reduced proton leak, suggest a coordinated reduction in proton pumping and leaking occurs in estivating Cepaea, which is likely a key component of the total energy savings necessary to depress BMR by 84%. Such coordinated reductions would be similar in principle to the 'channel arrest' mechanism by which Na4 and K+ gradients are maintained across the plasma membranes of rnetabolically depressed organisms (Buck and Hochachka, 1993; Couture and Hulbert, 1995; Flanigan et al., 1993; Perez-Pinzon et ai., 1992). Reductions in Na+ le* across plasma membranes cm be achieved through the downregulation of specific ion charnels (Perez-Pinzon, 1992). Sirnilarly, it may be possible to reduce mitochondrial proton leak through removal of membrane components which are associated with increased leak. In this context, our finding that the total phospholipid content of hepatopancreas tissue does not change during estivation is interesting. This suggests that those phospholipids which are lost from mitochondrial membranes remain within the cell. A possible mechanism for sequestering these phospholipids is that which has been suggested to reduce plasma membrane Na+ leak, i.e. the 'blebbing off' of channel-containing membrane fragments, which are then stored elsewhere in the ce11 (Pereze-Pinzon et al., 1992; Buck and Hochachka, 1993). Such reversible reductions in membrane surface area have been demonstrated in the alga, Dunafiella solina (Einspahr et al., 1988), in which exposure to hyperosmotic conditions results in the transfer of membrane phospholipids from mitochondria and other organelles to the endoplasmic reticulum, which serves as a temporary reservoir for membrane until a return to isosmotic conditions. Such a strategy could allow snails emerging from metabolically depressed states to restore a rnitochondrial phospholipid compwition typical of active metabolism through fusion with vesicles of stored phospholipid, so that phosphopid-mediad reductions in BMR might be rapidly reversed. Snails, typical of many estivators, enter into and emerge from metabolically depressed states quite rapidly (i.e. minutes to hours) (Herreid, 1977). We have demonstrated that dramatic alterations, consistent with a coordinated downregulation of metabolic processes, occur in mitochondr ial membranes of C. nemoralis during estivation. We suggest that estivating snails are a valuable animal mode1 for fuwe studies of the relationship between mitochondrial processes and metabolic rate. In particular, studies of mitochondrial proton leak in active and estivating snails should provide opportunities to confm and extend relationships identified in studies of hypo- and hyperthyroidism (Brand et al., 1992), comparisons of endo- and ectotherms (Brand et al., 1991), and allometric trends of marnmalian mitochondria (Porter et al., 1996). Estivation offers some advantages over these systems, in that the confounding effects of interspecies differences and temperature, which limit between-species comparisons and hibernation, respectively, can be circumvented. Similarly. surgical or pharmacological interventions which may be required to induce hypo- or hyperthyroidism are unnecessary. Table 11.1. Morphological and biochemical parameters from hepatopancreas of active and estivating snails. Morpholog ical and B iochemical Active Estivating % Change Measurements hepatopancreas weight 275 * 26 238 * 5 NS. (mg) mitochondrial protein 8.9 * 1.0 8.1 * 0.3 NS. (mglg hepatopancreas) total hepatopancreas protein 26.4 * 2.2 25.6 & 1.3 N.S. (mg/g hepatopancreas) mitochondrial phospholipid 89.1 * 8.0 25.2 f 2.7' -71.7% (nmol/mg mitochondrial protein) mitochondrial phospholipid 30.0 * 2.7 8.0 * 0.9' -73.3% (nmollmg cellular protein) total hepatopancreas phospholipid 68.5 * 7.7 59.3 * 6.4 NS. (nmol/mg cellular protein) cytochrorne C oxidase activity 1.9 f 0.2 0.3 f 0.1' -83.9 % (pmol-min-'-g wet tissue wt.) mitochondrial cardiolipin content 3.4 f 0.1 0.6 f 0.1' -82.7% (nrnollmg cellular protein) tissue cardiolipin content 3.8 * 0.7 2.6 * 0.4 NoS. (nmol /mg cellular protein) Values are means f SEM of 7 or 8 measurements. '*' = signifiicantly different from active snails (P < 0.05). N.S. = not significant. Table 1 1.2. Proportions (mole 5%) of phospholipid species in mitochondrial membranes of hepatopancreas ftom active and estivating snails. Phospholipid Act ive Estivating % change (mole %) (mole %) ------Cardiol ipin 11.4 * 0.1 7.4 & 1.4' - 35% Phosphatidylcholine 37.7 * 2.7 39.6 & 1.9 N.S. Phosp hatidy Iethanolam ine 35.2 f 1.5 30.1 f 0.7' Phosphatidylinositol 8.0 f 0.6 15.1 f 1.8' Phosphatidylserine 7.7 * 0.3 7.9 * 1.7 N.S. Values are means f SEM of 8 individual rneasurements. '*' = significantly differentiy from active snails (P < 0.05). N.S. = not significant. Table 11.3. Proportions (mole %) of individual fatty acids in mitochondrial phospholipids from hepatopancreas of active and estivating snails. Values are means * SEM of 8 measurements. '*' = significantiy different from active snails (P < 0.05). Parame ter Act ive Estivating % change (mole %) (mole %) Total Saturates 18.5 i 1.0 23.0 f 2.0 N.S. To ta1 Monoenes 14.5 * 0.8 19.8 * 1.4' + 37% Total PUFA 67.0 k 0.9 56.9 5 3.3' - 15% n6 PUFA 51.3 I0.9 48.9* 2.7 NS. Unsaturation Index' 222.5 f 3.9 213.8 * 10.2 N.S. Chain length2 18.4 I0.1 18.5 * 0.2 N.S. - Unsaairation Index = Xmiq; where miis the mole percentage and ni 1s the number of C-C double bonds of the fatty acid i. = Mean Chain Length = Zf$, ; where fi is the mole fraction and ci is the number of carbon atoms of the fatty acid i. GENERAL DISCUSSION The examination of ketone body metabolism in this study clearly illustrates that the metabolic organization of tissues of gastropod molluscs is a function of both phylogenetic history and environmental adaptation. BHBDH activity is present only in freshwater and terrestrial mollusc species, and not in any marine mollusc. This finding is confirmed by the failure of mitochondria isolated from marine molluscs to oxidize BHB (Ballantyne et al., 198 1; Ballantyne and Moyes, 1987). Examination of the activities of other ketone body enzymes indicates that the lack of BHBDH in marine molluscs is part of a general de-emphasis of ketone body metabolism. Because gasaopods lack the HMG-CoA pathway of ketogenesis, Acac must be both synihesized and oxidized by 30AT. When 30AT activities in tissues of marine and non-marine molluscs are standardized to general oxidative metabol ism (us ing CS activity) and compared, the adjusted activities are substantially lower in a marine species, relative to a freshwater and a terrestrial species (Chapter 2). This suggests that ketone body metabolism has a less important role in the intermediary metabolisrn of marine gastropods, especially cornpared to freshwater and terrestrial pulmonate gastropods. Indeed, activities of other enzymes suggest a greater emphasis on amino acid metabc!ism in marine species. This is similarly illustrated by the cornparison of CS-standardized activities of enzymes of amino acid metabolism in tissues of a marine gastropod with a non-marine gastropod (Chapter 3). The standardized activities are NO- to six-fold higher in marine species. These differences in metabolic organization between marine and non-marine molluscs may be related to the strategy of osmoregulation used. Marine gastropods are osmoconformers, using intracellular amino acid pools to maintain the osmotic pressure of their tissues near equilibrium with the surrounding medium. Total intracellular amino acid concentrations can be as high as 400 mM (Bishop et al., 1983). This large amino acid pool is virnially absent from tissues of freshwater and terrestrial molluscs (Bishop et al., 1983). With the loss of these pools in non-marine species, the need for high levels of enzymes to interconvert and oxidize amino acids would be reduced. The use of amino acids as energy substrates in intermediary metabolism may have similarly been reduced. Mile amino acid metabolism appears to have been downregulated in conjunction with the phylogenetic expansion of gastropods into non-marine habitats, pathways of ketone body metabolism have taken on greater importance. This is illustrated by the appearance of BHBDH activity in al1 freshwater and terrestrial gastropod species, and by the large upregulation (25-fold) of BHBDH activity in the pulmonate clade (Chapter 8). Similarly, pulmonate tissues oxidize acac and BHB at relatively high rates (Meyer et al., 1986). Tissue activities of ketone body enzymes are comparable CO values for glycolytic enzymes (Chapter 3), suggesting that the pathways of ketogenesis/ketolysis have assumed an importance similar to that of glycolysis in pulmonate gastropods. The subcellular organhtion of ketone body me tabolism in gas tropods differs from that of vertebrate animals (Fig. 12.1). In vertebrates, five mitochondrial enzymes catafyze ketone body metabolism, and the pathways of ketogenesis and ketolysis are different (Fig . 12.1a). Gastropods lack hydroxymethylg Iutary1-CoA synthase activity, and thus lack the main vertebrate ketogenic pathway (Fig. 12. lb). Thus, in molluscs, both ketogenesis and ketolysis must occur through the same pathway. This organization is Mermodifted in terresnial gasuopods (stylommatophoran pulmonates) through the localization of BHBDH isoforms to the cytosol (Fig . 12. lc) . The phylogenetic history of the Pulmonata includes a branching into two clades - Basommatophora and Stylomrnatophora (Chapter 8). The Stylommatophora, which have been by far the most successful invaden of terrestrial environmenu, and comprise the majority of al1 terrestrial snails (Little, l99O), have evolved unique isoforms of BHBDH, and thus modified pathways of ketone body metabolism. An important feature of both BHBDH isoforms is that they occur in the cytosol. All other known forms of BHBDH are compartrnentalized within the mitochondria. A second unique feature of one of the isoforms is its specifity for L-BHB.Al1 other known forms of the enzyme oxidue D-BHB exclusively. The cytosolic isoforms are also tissue-specific. L-BHBDH is found in hepatopancreas and kidney, and D- BHBDH is found in venuicle and kidney . Electmphoretic investigations have demonstratexi that the L and D-BHBDH activities are each the result of a single distinct enzyme. The resulu of Chapters 3 to 8 allow a mode1 for the design of ketone body metabolism in stylornmatophoran snails to be proposed (Fig. 12.2). Most terresuial mollusc tissues have a limited ability to oxidize fatty acids, but have well-developed ketolytic pathways. In active snails, ketogenesis could occur in the kidney, where relatively high activities of the enzymes which catalyze the 8-oxidation of fatty acids are found and 30AT kinetics appear to favour ketogenesis (Chapter 9). Kidney also contains both D- and L-BHBDH and can thus synthesize both D- and L-BHP Thus, kidney may conven the energy stored in fatty acids to ketone bodies, with L-BHB exported to hepatopancreas and D-BHB to heart and other tissues, allowing a partitioning of BHB between different tissues through differential production of the L- and D- enantiomers. This organization of ketone body metabolism is unique in the animal kingdom. Iu signifcance may lie in the additional level of control it provides over substrate traficking, beyond the regulatory effects of hormones, second messengers and allosteric effectors. In general, the ability to use both enantiomers of any substrate in intermediary metabolism is quite rare, though some evidence suggests that it may be a characteristic of molluscs. Several species of marine mollusc accumulate both L- and D-alanine in thei. tissues (Matsushima and Hayashi, 1992). Presumably, the advantage of this organization is that D-alanine can serve as a metabolically inert osmolyte, while L-alanine participates in energy metabolism. Thus, a partitioning of alanine into two distinct roles is achieved. Similarly , the enantiomer-specific organization of BHB metabolism may ailow for a partitioning of BHB into distinct rnetabolic roles. The specifity of tissues for either D- or L-BHB suggests that the metabolic finctions of BHB oxidation may differ in, for instance, hean and hepatopancreas. The partitioning of D- and L-alanine, outlined above, is achieved duough the combined actions of alanine amino tramferase atid a racemase (Matsuchima and Hayashi, 1992). This is fundamentally different from the organhtion of BHB metabolism, where D- and L-BHBDH are distinct, individual enzymes. Similarly, several cephaiopod species have both D- and L lactate dehydrogenases, which also appear to show some tissue specificity (Mulcahy et al., 1997). The physiologicai significance of this in cephalopods has not been established however. In general. elucidation of the functional significance of the enantiomer- specific organization of ketone body metabolism, and other metabol ic pathways, will require hirther research. Tissue specificity for substrate enantiomers may represent a means of metabolic regulation which has evolved in certain animai lineages, similar to other protein regulatory mechanisms, such as allosteric interactions, and reversible enzyme phosphorylation and subunit association. 1s there an adaptive advantage to the cytosolic cornpartmentation of BHBDH isoforms in terrestrial (stylommatophoran) snails? As outlined in Chapter 3, ~wving the hepatopancreas BHBDH reaction from mitochondrial to cytosolic compartments could be a strategy to promote gluconeogenesis. Snails rely in part on glycogen stores during estivation and anoxia (Storey and Storey, 1990). In both States, these stores become substantially depleted. Thus, ketogenesis may be particularly important following bouts of anoxia and estivation, when glycogen is resynthesized f'rom accumulated anaerobic endproducts. The utility of this during estivation is questionable given that this state has been shown to be fully aerobic (Rees and Hand, 1990). However , some experimental evidence suggests that anaerobic endproducts may accumulate transientiy, but are re-oxidized regularly following breaths. This is consistent with other chatacteristics of gastropod estivation physiology. During estivation, kart rate (Kratochvil, 1976). oxygen consurnption (Kratochvil, l976), car bon dioxide release (Barnhart and McMahon, 1987), and beat dissipation (Rees and Hand, 1990) al1 show cyclic change. Similarly, relatively large standard errors are associated with measurements of metabolic intermediates (Rees and Hand, 1991) and anaerobic endproducts (Churchill and Storey, Mg), consistent with large changes in concentrations during the period of the experirnent. Thus, it is possible that tissue iqpoxia and the concornittant accumulation of anaerobic endproducts occur transiently during estivation, between periods of gas exchange. Such a saategy would be promoted by the modified organization of BHB metabolism (as detailed in Chapter 3). Glycogen fermentation to lactate could provide ATP when, in the prolonged periods betweeen breaths, tissues could become transiently hypoxic. Immediatel y following gas exchange, lactate could be oxidized to pyruvate and reincorporated into glycogen. Ketogenesis from stored lipid at this tirne, and with the shifi of the BHBDH reaction to the cytosol, would promote gluconeogenesis through the altered intrarnitochondrial redox balance as outiined by Zammit (1990) and in Chapr: 4. The ketone body metabolism of terrestrial gastropods also differs from that of most vertebrates in that these pathways appear to be important even in the fed condition, based on the exceptionaily high enzyme activities in tissues of active snails (Chapter 3). This conclusion is supported by the observation that, in freshwater pulmonates, Ussues excised from active snails oxidize acac and BHB at high rates (Meyer et al., 1986). The roles of ketone body metabolism during estivation are less certain, however. Evidence from these and other studies suggests that the flux through ketogenic/ketolytic pathways, relative to that through other pathways, may be unchanged during estivation. Ketone body metabolism probably continues at reduced rates, as other catabolic processes are also reduced. This is suggested by the unchanged activities of ketone body enzymes following six weeks of estivation (Chapter 9). Similarly, as discussed in Chapter 5, mean respiratory quotients of active snails are approxirnately equal to those of snails entering estivation (Rees and Hand, 1990). No such data is available over longer-term estivation, but the rate of use of lipid and wbohydrate stores over two months of estivation (Rees and Hand, 1993) allows an estimation of this value, as no net consumption of protein occurs over this time (Chapter 9). The respiratory quotient thus estimated is approximately equal to that observed early in estivation. Thus, lipid catabolism appears to be important throughout estivation and ketone bodies are probably the preferred form of interorgan lipid transport in both active and estivating snails. Although my stuciies do not yet allow specific physiological roles of kerone body metabolism during estivation to be definitively identified, they provide some insight into mechanisms through which rnetabolic depression is achieved in snails. Enzyme ph~sphory lation and reversible binding of gl ycolytic enzymes to the cellular particulate fraction, are employed at the initiation of estivation (Storey and Storey, 1990). but may not be persistent over longer-term (weeks to months) estivation (Brooks and Storey, 1990; Michaelidis and Pardalidis, 1994). It had been suggested that, in long term estivation, suppression of enzyme activities may be achieved in part through the downregulation of enzyme concentrations, or metabolic reorganizations (Brooks and Storey, 1990; Michaelidis and Pardalidis, 1994). The results of Chapter 9, however , clearl y demonstrate that neither phenomenon occurs, at least in heart, kidney or hepatopancreas. Activities of virtually al1 of the enzymes of intermediary metabolism in these tissues are unchanged following six weeks of estivation. Similady, no signifiant difference between the tissue protein coc!c~sof active and estivating snails is observed, indicating that concentrations of enzymes of intermediary rnetabolism are unchanged. pH modulation of enzyme activity is also unlikely to be directly responsible for suppression of enzyme activities. Simulated intracellular acidification affects maximal enzyme activities in a number of metabolic pathways differentially (Chapter 9), but these effects do not appear to be subsrantial enough to account for the observed suppression of rnetabolic rate. Thus, suppression of intermediary metabolism in these tissues during long-term estivation appears to occur via some other mechanism, or combination of mechanisms. It is also possible that the extent of metabolic depression is tissuedependent and that heart, kidney and hepatopancreas, arguably the most highly aerobic gasaopod tissues, respond differently to prolonged estivation than less aerobic tissues like mantle and foot muscle. This possibility should be further explored. In hepatopancreas, the reductions in citrate synthase (Chapter 9) and CC0 (Chapter 11) activities indicate that control over mitochondrial oxidative metabolism may be exercised at more central levels, Le. the TCA cycle and FM3. Evidence that catabolic pathways are 'turned down' proportionally includes the lack of differential response of the maximal activities of enzymes from different pathways, and the similar rates of usage of different metabolic reserves (carbohydrate and lipid) in short and long-term estivation, which suggests that the net ratio of carbohydrate to lipid merabolised is constant over months of estivation. Thus, although glycolysis rnay be the immediate target of metabolic suppression upon entry into estivation, regulation of oxidative metabolism in longer-term estivation appears to focus on rnitochondrial function. The studies of mitochondrial membrane composition described in Chapters 10 and 11 suggat that mitochondrial structure and function are altered during estivation. The compositional changes observed in mitochondr ial membrane phospholipids are consistent with a suppression of membrane-bound enzyme and transporter activities. The dramatic reduction in mitochondrial membrane cardiolipin content parallels (and virtually equals) the almost 85% suppression of cytochrome c oxidase activity during estivation. This suggests a strong correlation of the two parameters, and perhaps a causai relationship. Because CC0 has an absolute requirement for cardiolipin to maintain activity, suppression of CC0 could be achieved directiy through removal of cardiolipin from the mitochondrial imer membrane. As so many mitochondr2 membrane-bound enzymes and transporters are aiso dependent upon interactions with cardiolipin (Hoch, 1992), reducing the membrane content of this phospholipid could provide a general mechanism for downregulating activities of these proteins. Other aspects of the phospholipid compos itional changes which occur dur ing estivation are consistent with a reduction of mitochondrial proton leak (Brand et al., 1994, Porter et al., 1996). This suggests that one means of achieving the substantial energy savings associateâ with metabolic depression is reducing the energy expended counteracting the proton leak. This mechanism, which 1 have termed 'leak arrest', is conceptually similar to the 'channel arrest' which occurs in some metabolically depressed states. Participation of 'channel arrest' in reducing energy requirements in physiological states involving a dramatic metabolic depression has been the focus of much research (Hochachka, 1986). Concerted ion pumping and gradient collapse occun across most cellular membranes. The energy €'rom the gradients created is harnessed for the transmernbrane transport of substrates and other ions through specific protein channels (Berne and Levy, 1988). Na' and KC gradients are maintaineci across the plasma membranes of virtually dl cells. Intracellular Na+ is pumped out of cells by membrane-bound Naf/K+ ATPase, in exchange for plasma KC,thus rnaintaining and inward flowing Na' gradient and outward flowing K* gradient. The Na+ gradient thus created drives the symport and antiport of various substrates (Berne and Levy, 1988). For example, plasma glucose and amino acids enter cells in some tissues via a Na+-linked transport mechanism (Berne and Levy, 1988). At reduced metabolic rates, the rates of these and other transport processes would be expected to be suppressed, as the demand for import of metabolic substrates (for exarnplz) into cells would be reduced. Channel arrest is the mechanism proposed for the suppression of transport without disrupting gradients. This occurs through concerted dgwnregulation of ion pumping and transport. Na+ channel arrest has been shown to occur in anoxic turties, which aiso undergo signifiant metabolic depression (Perez-Pinton et al., 1992; Buck and Hochachka, 1993). Evidence for arrested channel-mediated ion transport has also been provided by studies of estivating frogs (Flanigan et al., 1993). The data presented in Chapter 1 1 suggest that mitochondrial le* arrest, a reduction in rates of proton pumping and leaking, could occur in the mitochondrial inner membrane during estivation. This proposed Ieak arrest is analogous to channel arrest. In both cases the processes involved in the creation and collapse of a gradient are reduced in concert. My resulu further suggest that the mechanism by which leak arrest could occur (Fig. 12.3), may be similar to that suggested for achieving channel arrest in hypoxic turtles (Perez-Pinton et al., 1992), where channel-containing pieces of membrane are 'blebbed off' and stored. In the case of leak arrest, the total amount of phospholipid appears to contribute to proton leak, and by removing p hosphol ipid from the membrane the surface area across which the proton leak occurs is decreased. Complete reduction of proton leak could result in an energy savings of up to 20% (Rolfe and Brown, 1997). Na+ channel arrest has similarly been estimated to represent an energy savings of 5-10% (Guppy et al., 1994). Thus, it is apparent that 'globalized' channel and leak arrests, where the rates of pumping and leaking in al1 such systems are reduced, could realize considerable energy savings in estivators. This process could potentially be one of the largest contributors to metabolic depression. The proposed mechanism of leak arrest is consistent with one of the unifying themes of adaptations to metabolic depression in the terrestrial snail - that al1 modifications appear to be rapidly reversible. There is no evidence for overall reductions in the concentrations of enzymes, or the number of mitochondria. Rather, all of the metabolic 'machinery' seems to remain in place even over extended periods of profoundly reduced metabolic rate so that, upon a return to amenable environmental conditions, metabolic processes can be quickly resumed. Hobbs et al. (1994) provided evidence that the rate of protein synthesis remains constant in snails, even over long-term estivation. Indeed, apparently full recovery from weeks or months of estivation can often be observed in less than an hour (Herreid, 1977). This may be readily reconciled with the life-history of terrestrial snails. Favourable environmenial conditions in some habitats rnay be transient and a rapid response is probably necessary to capitalize on these. CONCLUSIONS AND PROSPECTUS Terrestrial snails (of the order Stylommatophora) have evolved an organization of ketone body metabolism found nowhere else in the animal kingdom. The modified ketogenicketolytic pathway appears to be important to the intermediary metabolism of gastropods, but the physiological roles of ketone body metabolism remain to be fully defined. It is possible that the cytosolic BHBDH's are adaptive to periodic hypoxia and may also be important during estivation. Investigations of ketone body metabolism in both of these physiological States should be an important focus for future studies. The unique D- and L-BHB specific isoforms rnay also provide an interesting mode1 for studying the relationship between the evolution of protein structure and function. It is apparent that an ancestor of the Stylommatophora had a typical mitochondrial BHBDH. An initial hypothesis regarding the evolution of cytosol ic isofons might assume divergence of these from the ancestral, mitochondrial isoform. This would necessitate a number of discrete steps: (1) the mitochondrial membrane-bound isoform has a large trammembrane region (Sandermann. 1986) and a functional requirement for phosphatidylcholine. Thus, an initial step rnay have been the loss of both characteristics, allowing the enzyme to Function as a soluble protein in the mitochondrial matrix; (2) the compartmentation of the enzyme was altered, ftom mitochondrial matrix to cytosol, and (3) an isoform evolved with modified stereospecifity for BHB, and was expressed in some tissues. Testing this hypothesis will requue identification and sequencing of the genes which encode al1 isoforms, and rnay providt valuable insight into the nature of protein evolution. For example, cornparison of the primary sequence of mitochondrial D-BHBDH From basommatophoran snails with the cytosolic D-BHBDH from stylommatophoran snails rnay allow the general arnino acid sequence which interacts with phosphatidylcholine in the membrane-bound isoform to be identified. Similarly, determination of the primary structures of cytosolic D- and L-BHBDHs rnay suggest what modifications result in altered stereospecifity. It is possible that only a few substitutions, additions or deletions rnay be sufficient to change the orientation of the active site. This is certainly suggested by the difficulty of separating the two isoforms on cellulose- acetate gels (see chapter 6) which separate on the basis of protein size and charge. In general, few natural models for studies of this nature exist, and the mitoch~ndrial and cytosolic snail BHBDHs should prove invaluable for studying the relationship between prirnary structure and the enantiomeric specificity of the active site. The cytosolic BHBDHs have a strong phylogenetic basis. This characteristic suggests that they will provide a valuable twl for discerning phylogenetic relationships amongst the pulmonate gastropods. In particular, a srnall number of species are sometimes classified under the order Systellommatophora (Vaught, 1989), which bear some afinities to pulmonates and also to opisthobranchs. It will be straightforward, and possibly quite informative to investigate the BHBDH activities of these species. Presence of L-BHBDH in tissues of systellommatophorans would argue strongly for a close relationship to stylommatophorans. Certainly , the function of m itochondria with respect to the reg ulation of oxidative metabolism should be t'urther studied. The profound membrane compositional changes observed in estivators suggest that mitochondrial function is duectly suppressed during estivation and that membranes play an important role in mediating this metabolic depression. Future studies should investigate the relationship between these membrane compositional changes and functional characteristics of the mitochondrial membranes, and the mitochondrion as a whole, including proton leak. In this Iight, the ability of CC0 activity to be modulated during estivation by altered membrane cardiolipin composition should also be investigated. For exarnple, does adding cardiolipin to the native membrane (containing CCO) fiom an estivating mail restore activity? This question can be directly addressed through in vitro manipulation of mitochondriai composition. The fate of phospholipids which are removed from the mitochondria during estivation should also be studied. An important aspect of the proposed strategy of le& arrest, where phospholipids are temporarily removed, but not oxidized, is its rapid reversibility. Thus, it will be necessary to establish a time-line for phospholipid modification which parallels that for general metabolic depression and recovery. Delineation of the details of this proposed strategy for modulating mitochondrial function promises to provide important insights into the regulation of metabohm during metabolic depression. It may also lead to the identification of global channel and leak arrest as a major component of the overall metabolic depression. Finally , it is possible that such a strategy is practised by other estivators, and by animals in other hypometabolic States, including hibernation, torpor? dormancy and facükitive anaerobiosis. Thus, extension of these studies to such systems should lead to the establishment of general principles of membrane adaptation during metabolic depression. (a) Vsrtsbtat. I (b) ûmsommatophora AC&- AACoA (c) Stylommatophora L Fig. 12.1. Subcellular organization of pathways of ketogenesis and ketolysis in (a) Venebrata, (b) Basommatophora (Gastropoda: Pulmonata) and (c) Stylommatophora (Gasuopoda: Pulmonata) . ACAC = acetoacetate; BHB = B-h ydroxybutyrate; AAT = acetoacetyl-CoA thiolase; HMS = hydroxymethylglutiiryl-CoA synthase; HML = hydroxymethylglutaryl-CoA lyase; 30AT = 3-oxoacid CoA transferase; BHBDH = 8-hydroxybutyrate dehydrogenase. ADP ATP Kidney u oxidation ACAC i L-BH B \ ADP ATP \ L-BH B 1 Fig. 12.2. Tissue organization of ketone body metabolkm in stylommatophoran gasuopods, showing proposed usages of pathways. D- and L BHB are synthesized in kidney from fauy acids and transported to hean and hepatopancreas, respectively, where they are oxidzed to produce ATP. ACoA = acetyl-CoA; AACoA = acetoac~l-CoA;8-oxid = the fatty acid B-oxidation spiral. Other acronyms as in Fig. 12.1 0 cardiolipin head group othw phospholipid head groups 2 unsaturateâ fatty acy chain 1 saturated faty acy chain Active Animal Cell œ Metaboiic 1 Depression1 r Estivating Animal Cell Fig. 12.3. Proposed mechanism for modifying mitochondrial membrane composition during estivation. Mitochondria of active snails contain more wdiolipin with more highly unsaturated faay acyl chahs. These are selectively removed from the membrane, dong with other phospholipids, and stored within the cell, presumably to be replaced upon a remto normal activity. These modifications to membrane composition are consistent with aitered function and reduced biological activity. In this proposed scheme, cardiolipin and other phospholipids are stored in the endoplasmic reticulum. LITERATURE CLTED Allen, W.V. 1977. Interorgan transport of lipids in the blood of the gumboot chiton Cryptochiton steellen' (Middendorff). Comp. Biochem. Physiol. 57A: 4 1-46. Aloia, R.C. 1981. 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