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Regulation of intracellular free calcium and protein kinase C in the motoneuron cell line NSC-19 Hasham, Mohammed Iqbal 1995

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R E G U L A T I O N OF I N T R A C E L L U L A R FREE C A L C I U M A N D PROTEIN KINASE C IN THE M O T O N E U R O N C E L L LINE NSC-19 by M O H A M M E D IQBAL H A S H A M B.Sc , The University of British Columbia, 1989 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF T H E REQUIREMENTS FOR THE D E G R E E OF M A S T E R OF SCIENCE in THE F A C U L T Y OF G R A D U A T E STUDIES (Department of Medicine, Neurology) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH C O L U M B I A August 1995 © Mohammed Iqbal Hasham, 1995 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of \ A e."b\C(iO \5 The University of British Columbia Vancouver, Canada Date ^TeMJiee- 2 ^ , t ^ S " DE-6 (2/88) Abstract Changes in the concentrations of intracellular free calcium ([Ca2+]j) were determined in response to the metabolic inhibitors, Amytal and carbonyl cyanide /w-chlorophenylhydrazone (CCCP), and the relationship between [Ca 2 +] s and protein kinase C (PKC) activation was investigated in the motoneuron cell line NSC-19. Amytal (5 mM) produced transient elevations of [Ca2+]j of approximately 80 nM. CCCP (10 uM) produced sustained elevations of [Ca 2 +] t of about 280 nM. These responses were reduced by 32% and 47%, respectively, when cells were studied in Ca2+-free solutions. In spite of the sustained elevation of [Ca 2 +] ; induced by CCCP, no reduction in cell viability was seen after 48 hours when compared to controls. These results indicate that exposure of NSC-19 cells to Amytal and CCCP produce Ca 2 + increments by release from internal stores, as well as by transmembrane influx, and demonstrate that small increments in [Ca2+]j due to metabolic inhibitors or other toxins may not be associated with immediate cell death. In Ca2+-containing solutions, inhibition of Na7Ca 2 + exchange led to increased [Ca 2 +] i 5 as did blockade of Ca2 +-ATPase, suggesting that these membrane transporters are functional in NSC-19. [Ca2+]j in these cells was modified by changes in extracellular Ca 2 + concentrations ([Ca2 +]0). Exposure of cells to increased [Ca 2 +] 0 of up to 10 mM resulted in sustained elevations of [Ca 2 +] i 5 which maintained steady state levels for at least 10 min. P K C activity in cytosolic and membrane extracts from cells exposed to varying [Ca 2 +] 0 was assessed by the ability of the enzyme to phosphorylate histone HI in the presence or absence of Ca 2 + , diolein, and phosphatidylserine. Extracts from cultures incubated in 1.3 mM [Ca 2 +] 0 (control, [Ca2+]; was 83 ± 17 nM) contained ii P K C activity predominantly in the cytosol fractions. A 10 min exposure of the cells to 2.5 mM [Ca z +] 0 ([Ca2+]; was 140 ± 8 nM) resulted in the partial translocation of cytosolic P K C activity to the membrane, and exposure to 5 mM [Ca 2 +] 0 ([Ca2+]; was 232 ± 24 nM) resulted in almost complete translocation. Cells exposed to 10 mM [Ca 2 +] 0 ([Ca2+]j was 365 ± 13 nM) showed a two-fold increase in cytosolic PKC activity and an eleven-fold increase in membrane-associated P K C activity, suggesting that the increased activation was translocation-independent. Total P K C activity decreased in extracts from cells exposed to 25 mM and 50 mM [Ca 2 +] c. [Ca2 +]; rose transiently to over 600 nM and 900 nM, respectively, and then returned to steady state values of 202 ± 1 4 nM and 122 ± 6 nM. Under these conditions, cytosolic and membrane-bound protein kinase M (PKM) activity rose approximately eight-fold and ten-fold, respectively, for both groups. Thus, three modes of P K C activation by increased [Ca2+]j were observed: (i) translocation of P K C activity from the cytosol to the membrane at [Ca 2 +] ; between 140 nM and 230 nM; (ii) apparent translocation-independent activation of P K C at [Ca 2 +] ; values around 365 nM; and (iii) proteolytic cleavage of PKC to P K M following a Ca 2 + transient of greater than 600 nM. These three distinct modes of P K C activation may have unique physiological consequences that depend on the amplitude and duration of the initiating [Ca 2 +] ; signal. iii Table of Contents Abstract i i Table of Contents iv List of Figures vi List of Tables vii List of Abbreviations viii Acknowledgements x Chapter 1 Introduction 1.1 Mechanisms of calcium entry into cells 1 1.1.1 Voltage-dependent Ca 2 + channels 2 1.1.2 Receptor-operated channels . . ! 4 1.1.3 Ionotropic glutamate receptor ion channels 4 1.1.4 Other Ca2+-permeable channels 5 1.1.5 Receptors linked to guanine nucleotide binding proteins 5 1.2 Extracellular E A A levels in central nervous system damage 6 1.3 EAA-mediated elevation of [Ca2+]j .7 1.4 Role of Ca 2 + in cell death . 10 1.5 Characteristics of raised [Ca2 +]; 13 1.6 Involvement of Ca2+-dependent processes in cell death 15 1.6.1 Activation of PKC 16 1.7 Objectives 20 Chapter 2 Materials and Methods 2.1 Materials 22 2.2 Cell culture 22 2.3 Fluorescence studies 23 2.3.1 Cell loading and buffers 23 2.3.2 Calcium imaging 24 2.3.3 Calibration of the imaging system 25 2.3.4 Determination of cell viability 27 iv 2.4 Biochemical analyses 28 2.4.1 Preparation of cell extracts 28 2.4.2 Determination of protein content 28 2.4.3 Partial purification of P K C and P K M 29 2.4.4 Protein kinase assays 29 2.4.5 Determination of cytosolic adenine nucleotides 30 2.5 Western blot analyses 31 2.5.1 Anti-PKC antibodies 32 2.6 Statistics 33 Chapter 3 Results 3.1 Fura-2 calibration and basal [Ca2 +]; in NSC-19 cells 34 3.2 Effect of changing [Ca 2 +] 0 on [Ca 2 +] ; 37 3.3 Role of Ca 2 + ion channels in changing [Ca2+]j 38 3.4 Role of other Ca 2 + transporters in changing [Ca2+]; 41 3.5 Effect of metabolic inhibition on NSC-19 cells 46 3.5.1 Inhibition by amobarbital (Amytal) 46 3.5.2 Inhibition by carbonyl cyanide/w-chlorophenylhydrazone (CCCP) 51 3.6 Effect of changing [Ca 2 +]D on the activity of P K C 56 v L i s t of Figures Fig. 1.1 Routes of Ca 2 + entry into cells 3 Fig. 1.2 EAA-mediated elevation of [Ca2+]j 9 Fig. 2.1 Experimental setup of imaging system 26 Fig. 3.1 Calibration of fura-2 imaging 35 Fig. 3.2 Distribution of basal [Ca 2 +] ; in NSC-19 cells 36 Fig. 3.3 Effect of changing [Ca 2 +] 0 on [Ca 2 +] { 39 Fig. 3.4 Effect of increasing [Ca 2 +] 0 on [ C a 2 ! 40 Fig. 3.5 Effect of vanadate on [Ca2+]; 43 Fig. 3.6 Effect of blocking Na7Ca 2 + exchange on [ C a 2 l 44 Fig. 3.7 [Ca 2 +] ; responses to Amytal 50 Fig. 3.8 [Ca 2 +] ; responses to CCCP 54 Fig. 3.9 MonoQ chromatography of cell extracts 58 Fig. 3.10 Western blot analysis of MonoQ peak I and II fractions 60 Fig. 3.11 Effect of [Ca 2 +] 0 on activation of P K C and P K M 61 Fig. 3.12 Western blots of crude extracts from NSC-19 cells 63 vi L i s t of Tables Table 1.1 Patterns of expression of selected P K C subspecies 17 Table 3.1 Effects of Amytal and CCCP on ATP, ADP, A M P , and energy charge 48 Table 3.2 Effects of Amytal and CCCP on basal, peak, and recovery [Ca 2 +] ; 49 vii List of Abbreviations A 5 9 5 Spectrophotometric absorbance at 595 nm. ACPD 1S-3R (trans)-l-amino-cyclopentane-l,3-dicarboxylate ADP Adenosine diphosphate A M P Adenosine monophosphate A M P A DL-a-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid Amytal Amobarbital sodium ATP Adenosine triphosphate B S A Bovine serum albumin BSS Balanced salt solution CBP Calcium binding proteins CCCP Carbonyl cyanide m-chlorophenylhydrazone CNS Central nervous system cpm Counts per minute C R A C C a 2 + release-activated C a 2 + (channels) D A G Diacylglycerol DMSO Dimethyl sulfoxide D N A Deoxyribonucleic acid DO Diolein DTT Dithiothreitol E A A Excitatory amino acid FCCP Carbonyl cyanide /?-fluoromethoxyphenylhydrazone Fmax(380nm) Maximum fluorescence intensity at 380 nm (under saturating [Ca Fmin (380nm) Minimum fluorescence intensity at 380 nm (under zero [Ca 2 +]) FPLC Fast protein liquid chromatography Fura-2/AM Fura-2 acetoxymethyl ester Fura-2/K5 Fura-2 pentapotassium salt HBSS HEPES buffered Hank's balanced salt solution viii HRP Horseradish peroxidase IgG Immunoglobulin G IL-x Interleukin-x (example IL-2 = Interleukin-2) IPs Inositol 1,4,5-trisphosphate K d Dissociation constant M E M Minimal essential medium mRNA Messenger ribonucleic acid N A Numerical aperature N A D H Nicotinamide adenine dinucleotide (reduced) N M D A N-methyl-D-aspartate PAF Platelet activating factor P K C Protein kinase C PKLP Cyclic AMP-dependent protein kinase inhibitor peptide P K M Protein kinase M PLC Phospholipase C PMSF Phenylmethyl sulfonic acid PPTase Protein phosphatase PS Phosphatidylserine R Ratio -^ max Maximum ratio •^ min Minimum ratio rpm Revolutions per minute S.D. Standard deviation SDS Sodium dodecyl sulfate S E M Standard error of the mean TPA 12-o-tetradecanoylphorbol 13-acetate (phorbol ester) V D C C Voltage-dependent calcium ion channel x g Force times gravity ix Acknowledgements I would like to thank my supervisor, Dr. Charles Krieger, for his enthusiasm and encouragement. Without his guidance, this study would not have been possible. I also wish to thank Dr. Steven L. Pelech for his support and constructive critisicm of my work, and for the use of his laboratory and equipment at Kinetek Biotechnology Corporation. I appreciate Dr. H . Koide for his help with kinase assays and Western blots, Dr. S. U . Kim for allowing me to use his tissue culture facilities, G. Ritchie for performing the ATP measurements following metabolic inhibition, D. Naumann for performing preliminary measurements on both [Ca2+]j and ATP in NSC-19 cells, Dr. N : R. Cashman for providing the cell line, and the members of Dr. Pelech's laboratory for their suggestions and assistance. My deepest gratitude goes to my family whose love and support is ever present, and to whom I will always be indebted. Chapter 1 Introduction Calcium ions (Ca 2 +) are one of the major second messengers in neurons. Hormones, neurotransmitters, and electrical activity produce rapid increases in intracellular free Ca 2 + concentration ([Ca2+];) which are used as signals to regulate cell functions. To use free Ca 2 + in controlling cellular processes with a high degree of spatial and temporal precision, cells must have tight control over the internal Ca 2 + concentration. Cells in the nervous system, as well as in other parts of the body, maintain the cytosolic concentration of free Ca 2 + within the range of 50 to 200 nM, approximately four orders of magnitude lower than in the extracellular space (about 1.3 mM). The steep concentration gradient is maintained largely by two surface membrane proteins, a Na7Ca 2 + exchanger protein and an ATP-dependent Ca 2 + pump (Ca2+-ATPase), as well as a distinct intracellular Ca 2 +-ATPase which sequesters Ca 2 + into intracellular storage pools, primarily endoplasmic reticulum (Di Polo and Beauge, 1979; Blaustein et al, 1991). In addition, Ca 2 + is bound to Ca2+-binding proteins which may become activated by binding Ca 2 + or which may act to buffer Ca 2 + . In addition to controlling the level of cytoplasmic [Ca 2 +] ;, these mechanisms can maintain the free Ca 2 + concentration in the intracellular pools and in the cell nucleus independently of each other and of cytoplasmic [Ca 2 +] ; (Williams et al, 1985). Alterations in the mechanisms which control the maintenance of the Ca 2 + gradient may lead to a pathological loss of Ca 2 + homeostasis. 1.1 Mechanisms of calcium entry into cells Several pathways exist for Ca 2 + influx into cells. The principal routes of Ca 2 + entry include influx through voltage-dependent Ca 2 + channels (VDCC) and ligand-gated ion channels such as 1 the excitatory amino acid (EAA) receptor-operated ion channels (Fig. 1.1). Additional Ca 2 + influx has been shown to occur through the Na7Ca 2 + exchanger (Stys et al, 1991), through L a 3 + -sensitive Ca 2 + leak channels (Demirel et al, 1993), and through non-specific membrane leak. Al l these entry pathways have been demonstrated to cause increases in intracellular free calcium ([Ca2+];) in neuronal cells in culture. [Ca2+]j can also increase in neurons by Ca 2 + release from internal stores, such as the endoplasmic reticulum, mitochondria, or Ca2+-buflfering proteins. Alternatively, elevations of [Ca 2 +] ; have been demonstrated to occur following blockade of the energy requiring Ca 2 +-ATPase (Fig. 1.1) (Dehlingen-Kremer et al, 1991), and disruption of mitochondrial energy metabolism (Villalba et al, 1994). 1.1.1 Voltage-dependent Ca 2 + channels The V D C C comprise a group of multi-subunit membrane spanning proteins which allow Ca 2 + influx when activated by membrane depolarization. This group of channels has been divided into the L- , N - , T-, P-, and Q-type channels, as well as other channel types based upon their pharmacological sensitivities and activation/inactivation characteristics. The functional classification is only partially correlated with channel types defined by cloning studies (Snutch et al, 1990; Swandulla et al, 1991), likely reflecting the presence of as yet pharmacologically indistinct channel types. Individual cells possess a unique complement of V D C C types, which results in markedly different patterns of Ca 2 + influx between different cell classes. 2 Fig. 1.1 Routes of Ca 2 + entry into cells Some mechanisms of Ca 2 + entry into cells. Ca 2 + can enter through voltage-dependent Ca 2 + channels (VDCC), ionotropic excitatory amino acid (glutamate) receptor-operated cation channels, reverse operation of the Na7Ca 2 + exchanger, decreased operation of Ca2+-ATPases of the plasma membrane or endoplasmic reticulum, or Ca 2 + leak. [Ca 2 +] ; can also increase by release from calcium binding proteins (CBP) or by the metabotropic glutamate receptor-stimulated, phospholipase C (PLC)- and inositol 1,4,5-trisphosphate (rP3)-mediated release from internal stores. 3 1.1.2 Receptor-operated channels Receptor-operated ion channels, like VDCC, are multimeric proteins which span the plasma membrane or intracellular membranes. Their activation depends upon the binding of a specific ligand to the receptor complex, rather than upon the intrinsic 'voltage sensing' property of the VDCC. Examples of receptor-operated channels include the E A A (or glutamate) receptor-gated ion channels on the plasma membrane, and the intracellular inositol 1,4,5- trisphosphate (IP3) receptor channels which allow Ca 2 + flux upon binding of the appropriate agonist to the receptor. 1.1.3 Ionotropic glutamate receptor ion channels At least 16 different subunits of the ionotropic glutamate receptor-operated cation channels have been described to date (see Seeburg, 1993). Receptor/channel complexes are formed by a combination of five subunits, and the Ca 2 + permeability or agonist sensitivity of the resultant receptor ion channels is dependent upon the complement of subunits which make up the channel. For instance, the presence of at least one GluR2 subunits within the protein complex results in loss of Ca 2 + permeability of the channel. The N-methyl-D-aspartate (NMD A) subclass of ionotropic glutamate receptor-operated channels are several fold more permeable to Ca 2 + than the channels sensitive to kainate or DL-a-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA), the latter two being primarily permeable to Na + or K + ions. This difference in permeabilities may be due to the presence of an asparagine residue in the second transmembrane domains of the NMDA-sensitive subunits, which allows for Ca 2 + flux. The presence of a glutamine residue instead of asparagine in K A I and K A 2 subunits, may confer some Ca 2 + 4 permeability, but substitution of this residue to arginine such as in the GluR2 subunit, markedly decreases the Ca 2 + permeability of the channel. 1.1.4 Other Ca2+-permeable channels The IP 3 receptor-operated ion channels are tetrameric protein complexes which contain an IP3 binding site and an ion channel which allows for Ca 2 + flux. Molecular diversity of the receptor subunits arises from both alternative splicing and the existence of separate genes, as with the glutamate receptors (Sudhof et al, 1991; Ross et al, 1992). The membrane-spanning domains of the C-terminal regions of four receptor subunits combine to form the functional Ca 2 + channel. This channel is activated upon binding of IP3 to a cytoplasmic site in the N-terminal region of the protein. The ryanodine receptor shares considerable structural and molecular homologies with the IP3 receptor (Henzi and MacDermott, 1992; Tsien and Tsien, 1990) which may account for many functional similarities between the two intracellular receptors. Activation of the ryanodine receptor ion channel is thought to occur following plasma membrane depolarization and entry of a small amount of'trigger' Ca 2 + . This Ca 2 + then stimulates Ca 2 + flux through the channel by a positive feedback loop, known as Ca2+-induced Ca 2 + release. The presence of high [Ca 2 +] ;, however, inhibits the channel operation (see Berridge, 1993), providing for negative feedback. 1.1.5 Receptors linked to guanine nucleotide binding proteins The G-protein-linked receptors operate by the direct interaction of the receptor with guanine-nucleotide binding proteins (G-proteins). Receptors consist of single polypeptides which 5 traverse the membrane seven times, giving them the name 'serpentine receptors'. The metabotropic subclass of glutamate receptors, which belong to this group, may be coupled to secondary effector enzymes, such as phospholipase C (PLC) or adenylate cyclase (AC), through G-protein interaction. At least seven different forms of the metabotropic glutamate receptor have been identified. These cloned receptors, named mGluRj to mGluR7, are derived by alternative splicing and may be coupled through G-proteins to PLC or A C or both. The relationship of the cloned receptors to endogenously expressed metabotropic glutamate receptors is as yet unclear since only one selective agonist, 1S-3R (frans)-l-amino-cyclopentane-l,3-dicarboxylate (ACPD), has been found to date (Schoepp et al, 1990). This agonist, though selective for metabotropic glutamate receptors, appears to have effects on at least six clones. The lack of type-specific agonists, antagonists or radioligands for this class of receptor has prevented the detailed study of the different forms of metabotropic glutamate receptors, only some of which may be coupled to intracellular Ca 2 + release mechanisms. 1.2 Extracellular E A A levels in central nervous system damage Glutamate is the primary neurotransmitter at the majority of excitatory synapses in the central nervous system (CNS). Microdialysis experiments during experimental ischemia of rat hippocampus have demonstrated increases in extracellular glutamate concentration (Beneviste et al, 1984) and corresponding decreases in the glutamate contents of intracellular compartments (Ffagberg et al, 1985). Increased glutamate concentrations in the extracellular spaces can occur as a result of glutamate release from nerve terminals into synaptic clefts under periods of sustained electrical activity (for example, during seizures), or as a result of glutamate leakage 6 from mechanically injured cells. Alternatively, extracellular glutamate may increase as a result of decreased uptake by energy-dependent mechanisms under conditions of low cellular energy states. For example, depletion of adenosine triphosphate (ATP) stores under hypoxic or hypoglycemic conditions may result in ionic imbalance due to the decreased activity of ATP-dependent ion transporters. This in turn may lead to the failure of ion-coupled glutamate uptake systems, primarily found within glial cell populations (see Szatkowski and Attwell, 1994). Neuronal responses to increased extracellular glutamate will depend upon the cellular complement of glutamate receptor subtypes which are activated, and the Ca2+-buffering capabilities of individual cells. 1.3 EAA-mediated elevation of [Ca2+]; Increases in extracellular glutamate concentration can produce elevations in [Ca 2 +] ; by several distinct mechanisms (Fig. 1.2) (see Choi, 1988). First, activation of N M D A receptors can result in Ca 2 + influx through associated cation channels (Fig. 1.2, "A") (Tymianski, 1993a). Influx of Ca 2 + has been demonstrated following activation of some kainate and A M P A receptor-operated channels (Fig. 1.2, "B4") (Gibbons et al, 1993; Ogura et al, 1992), depending upon the type of ionotropic glutamate receptor subunits which form the pentameric receptor/channel complex (see Seeburg, 1993). Second, membrane depolarization (Fig. 1.2, "Bl") caused by Na + influx through the glutamate receptor-gated channels may increase VDCC-mediated Ca 2 + influx (Siesjo, 1990; Nicholls and Attwell, 1990). Third, Na + influx with a subsequent increase in intracellular Na + concentration, may also lead to Ca 2 + influx through reverse operation of the Na7Ca 2 + exchanger (Stys etal, 1991) (Fig. 1.2, "B2"), which normally exchanges intracellular Ca 2 + for extracellular Na + . Fourth, Ca 2 + may enter the cell by Ca 2 + leak through the plasma membrane (Fig. 1.2, "B3") as a result of Na+-influx-associated cell swelling. In addition to increased Ca 2 + influx by the above mechanisms, [Ca 2 +] ; may increase as a result of Ca 2 + release from internal stores such as endoplasmic reticulum and mitochondria. Glutamate binding to serpentine metabotropic receptors can activate inositol phospholipid metabolism by direct interaction with G-proteins (Gasic and Ffollmann, 1992). Activation of P L C by G-protein interaction leads to increased levels of intracellular EP3 (Fig. 1.2, "C"), which in turn stimulate mobilization of intracellular Ca 2 + (see Berridge, 1993). PLC activation also results in the production of diacylglycerol (DAG) from membrane phospholipids. D A G , along with phosphatidylserine and Ca 2 + , has been shown to activate the Ca 2 + - and phospholipid-dependent protein kinase (protein kinase C, PKC) (see Nishizuka, 1992). Application of ACPD, the glutamate receptor agonist selective for the metabotropic subtype of receptors, has been shown to potentiate increases in [Ca2 +]4 evoked by N M D A, kainate or A M P A in cultured neurons (Bleakman et al, 1992). The potentiation of Ca 2 + currents is likely due to the secondary activation of PKC, which has also been shown to increase NMDA-activated Ca 2 + currents (Chen and Huang, 1991). The effect of P K C on NMDA-stimulated Ca 2 + currents is reversed by the simultaneous application of P K C inhibitory peptide (Chen and Huang, 1991) derived from the N -terminal regulatory domain of PKC. Increases in NMDA-activated Ca 2 + currents by activated PKC may result from phosphorylation of the receptor and subsequent removal of the M g 2 + block of the ion channel (Chen and Huang, 1992). 8 Fig. 1.2 EAA-mediated elevation of [Ca 2 + ] ; Possible mechanisms of C a 2 + entry into cells under pathological conditions. See text for details. SEIZURES HYPOXIA/ ISCHEMIA Glutamate Release HYPOGLYCEMIA MECHANICAL TRAUMA •5 Glutamate Uptake tExtracellular Glutamate Glutamate Leakage © NMDA-R Stir, timulation non-NMDA-R Stimulation — -^ | Na+ Influx ^ -•'""Membrane s \Depolarization ) © V Metabotropic-R — , Stimulation f ^ V,^^ Swelling ./ t Reverse Na + /Ca 2 + Exchanger Operation t VDCC Conductance } I t Non-specific C a 2 + Leakage t Ca2+ Influx [Ca2+]i A Ca2+ T Release fI P 3 DAG PKC Calpain Cell Damage Other C a 2 + activated processes 9 1.4 Role o fCa 2 + i n cell death The cause of neuron death following ischemia or in neurodegenerative disorders of the CNS is unclear. Elevated [Ca 2 +] ; was suggested to mediate cell death, and Ca 2 + influx into cells was termed "the final common pathway of cell death" (Schanne et al, 1979). Since then, the "calcium overload hypothesis", has been based upon numerous studies of [Ca2+]; responses of neurons in vivo and in tissue culture models of hypoxia- and EAA-induced cell death. Evidence supporting the role of altered [Ca 2 +] ; homeostasis as a mediator of cell death in experimental models includes observations of Ca 2 + accumulation in nervous tissue in cerebral ischemia (Siesjo, 1990) and a strong correlation between cell death and increased 4 5 C a 2 + uptake in cultured cells following exposure to N M D A or glutamate (Eimerl and Schramm, 1994). Several studies have shown that increases in [Ca2+]j due to Ca 2 + influx through NMDA-gated ion channels are correlated with increased death of cells in culture. Furthermore, inhibition of Ca 2 + efflux through the Na7Ca 2 + exchanger enhances glutamate-associated cell death in cerebellar granule cell cultures (Andreeva et al, 1991). Pharmacological interventions which block NMDA-induced Ca 2 + influx or intracellular Ca 2 + release are protective against the death of cultured neurons (see Choi, 1988). For example, blockade of N M D A receptor ligand-binding sites or the N M D A -operated ion channel itself diminishes the loss of cultured cortical and hippocampal neurons in response to E A A application (Michaels and Rothman, 1990; Tymianski et al, 1993a). Also, blockade of the NMDA-operated ion channels, but not of NMDA-receptor interaction, attenuates the magnitude of the EAA-induced Ca 2 + currents (Michaels and Rothman, 1990). Other supporting evidence for a role of Ca 2 + in mediating cell death comes from studies which demonstrate that removal of extracellular Ca 2 + prior to the application of N M D A or other toxins 10 results in decreased EAA-associated neurotoxicity (Schanne et al, 1979; Choi, 1987; Hartley and Choi, 1989; Mattson etal, 1991; Goldberg and Choi, 1993). Under zero [Ca 2 +] 0 conditions, neither glutamate exposure nor anoxia of cultured hippocampal neurons resulted in increased [Ca2+]j (Friedman and Haddad, 1993; Tymianski et al, 1993a). Application of kainate has been demonstrated to produce a [Ca2 +]; increase which is associated with substantial cell death. However, the elevation of [Ca2 +]4 by kainate, as well as the associated toxicity, was partially reduced by the application of MK-801, an N M D A receptor antagonist. This suggests that a component of the Ca 2 + influx may have occurred through N M D A-gated ion channels, secondary to activation of kainate receptor-associated ion channels. Although a considerable amount of evidence supports the role of Ca 2 + in mediating cell death, not all reports have demonstrated an association between increased [Ca 2 +] ; and cell death under ischemic or other cytotoxic conditions and some studies provide evidence which suggests that elevated [Ca2 +]; does not mediate cell death. For instance, anoxia of neonatal rat CA1 neurons does not result in significant cell death as compared to adult neurons, even though both groups demonstrate similar magnitudes of [Ca 2 +] ; elevation (Friedman and Haddad, 1993). However, the observation that the latency to peak [Ca2+]j was much greater in neonatal cultures than in adult cells suggests that the nature of the Ca 2 + rise may be an important determinant of cell fate. Furthermore, the anoxia-induced [Ca 2 +] ; changes in the neonatal CA1 cultures were significantly different from their Ca 2 + responses to glutamate, whereas the responses of the adult neurons to anoxia or glutamate challenge were similar, indicating that susceptibility to either anoxia or glutamate challenge may be dependent upon the developmental stage of the cells. Other age and species related differences in the effects of E A A and toxins on [Ca 2 +] i 5 and in Ca 2 + -11 handling by cells have also been noted (for review see Peterson, 1992), and these may affect cell survival. Thus, the sensitivity of neurons for E A A - or [Ca2+] rrelated cell death, at least in culture, may be age- or species-dependent. Human embryonic neurons showed no sensitivity to glutamate toxicity until after 30 days in culture (Mattson et al, 1991). However, Ca 2 + responses to glutamate developed several weeks prior to sensitivity to glutamate-induced cell death, suggesting that the neurotoxic process(es) triggered by glutamate and possibly elevated [Ca2+];, had not developed even though cells responded to glutamate. In contrast, rodent neurons developed Ca 2 + responses to glutamate on day 4 in culture and were susceptible to glutamate toxicity by day 5, suggesting that neurotoxic processes developed in parallel with the development of responses to glutamate (Mattson et al, 1991). The Ca 2 + responses to the Ca2+-ionophore A23187 were sustained for longer periods of time in the rat neurons than in the human cultures following equivalent exposure of the ionophore to the two cultures. The sustained Ca 2 + responses of rodent neurons to A23187 suggests that the rat cells were not as effective at buffering Ca 2 + loads as were the human neurons, and may have resulted in prolonged activation of Ca2+-dependent processes in the rat cultures. Seemingly contradictory evidence for the role of Ca 2 + in mediating cell death also comes from studies which show that increases in [Ca 2 +] ; due to the activation of kainate or A M P A receptors or VDCCs, and/or due to mitochondrial inhibition of cultured neurons do not significantly affect neuronal survival (Dubinsky and Rothman, 1991; Tymianski et al, 1993a). In one study where [Ca 2 +] ; was made to rise transiently and to approximately equal values by activation of either VDCCs or NMDA-gated ion channels, the death of cultured spinal neurons was greater following Ca 2 + influx through NMDA-operated mechanisms than with influx through 12 VDCCs (Tymianski et al, 1993a). It is possible that the toxicity of raised [Ca 2 +] ; is dependent upon the co-activation of other processes, perhaps Na + influx, metabotropic receptor stimulation, or other Ca2+-dependent processes which may co-localize with N M D A receptors. It is evident that not all elevations of [Ca2+]j are associated with cell death, presumably reflecting differences in the manner in which a Ca 2 + load is handled by any given cell. Since Ca 2 + is an important second messenger, it is not surprising that increases in [Ca 2 +] ; are seen during several normal developmental processes such as development of growth factor-independent survival and neurite extension (Koike et al, 1992; Eichler et al, 1992; Mattson and Kater, 1987; Cohan et al, 1987). Thus, the responses to increased [Ca 2 +] ; may differ with the cell type, the period of cellular development, and or the source and nature of the Ca 2 + influx. Also, the ability of any given cell to compensate for a glutamate-induced increase in [Ca 2 +] ; may be compromised by decreased energy stores during periods of ischemia (Beal, 1992). In cultured hippocampal neurons, glutamate-induced elevations of [Ca2+]j were greatly amplified by the addition of sodium cyanide (NaCN), as was the extent of cell death (Dubinsky and Rothman, 1991). Although the application of NaCN alone resulted in a large increase in [Ca 2 +] ;, little or no cell death was observed. The latency to peak [Ca 2 +] ; was significantly longer with NaCN alone than with glutamate (alone or in combination with NaCN), suggesting that the rate of rise of the [Ca 2 +] ; may influence cell survival. 1.5 Characteristics of raised [Ca 2 +] ; It is not known what property of the [Ca2+]i rise might be responsible for neurotoxicity. The amplitude of the [Ca 2 +] ; rise, the spatial or temporal profile of the Ca 2 + response, or a 13 combination of these factors may be relevant. In addition, it is not known how or whether elevated [Ca 2 +] ; mediates cell death. Application of glutamate to cultured neurons results in a biphasic response in [Ca 2 +] ;. Levels of [Ca2+]; increase transiently to values ranging from less than 500 nM to greater than 1 u M (see Dubinsky, 1993b). Following this transient, [Ca 2 +] ; returns to near baseline levels for a variable period, usually several hours, before increasing a second time (Dubinsky, 1993a; Tymianski et al, 1993a; Glaum et al, 1990). The secondary rise in [Ca 2 +] i } which does not always occur, is followed by an increased permeability to vital dyes, indicating a compromise in plasma membrane integrity and imminent cell death (Tymianski et al, 1993b). Removal of extracellular Ca 2 + following glutamate-evoked transient elevations of [Ca2+];, prevents subsequent increases in [Ca 2 +] b but does not always prevent cell death (Michaels and Rothman, 1990). The lack of a secondary rise in [Ca 2 +] ; prior to cell death in some studies suggests that this Ca 2 + influx is not directly responsible for mediating cell death, but rather, that it is indicative of impending death of cells in culture. Also, evidence which suggests that the Ca 2 + ions per se are not cytotoxic include: (i) the observation that [Ca 2 +] ; is only transiently elevated in EAA-induced toxicity, and (ii) that Ca 2 + need not be present following the initial Ca 2 + transient for cell death to occur . This notion is not surprising since M g 2 + , a divalent ion like Ca 2 + , has not been demonstrated to be toxic. The intracellular free magnesium ion concentration ([Mg2+]j), like [Ca 2 +] ;, has been shown to increase, in the presence of extracellular Ca 2 + , from 0.5 mM at rest, to between 1 mM and 11 mM with application of glutamate (Brocard et al, 1993) or mitochondrial uncouplers (Li etal, 1993), such as carbonyl cyanide /w-chlorophenylhydrazone (CCCP), a protonophore at the mitochondrial membrane. This increase in [Mg 2 +] s has not been associated 14 with cell death and may in part be due to a lack of evidence for Mg2+-induced stimulation of intracellular enzymatic activity, as has been demonstrated for Ca 2 + . It is likely that [Ca 2 +] ; elevations following EAA-application to cultured neurons activates downstream processes which may then mediate the cytotoxic effect by a Ca2+-independent mechanism. 1.6 Involvement of Ca2+-dependent processes in cell death Ca 2 + can stimulate the activity of a variety of intracellular signalling enzymes including protein kinases, protein phosphatases, and proteases (Mattson, 1991; Fukunaga etal, 1992). Alterations in the activities of several Ca2+-dependent enzymes have been demonstrated following EAA-mediated elevations of [Ca 2 +] ; and these may be associated with toxicity (Maiese et al, 1993). The activation of Ca2+-dependent proteases as a result of raised [Ca 2 +] ; or metabotropic glutamate receptor stimulation may be important in mediating neurotoxic events following glutamate application to cultured neurons. The actions of calpain on cytoskeletal proteins (Siman and Noszek, 1988) may compromise membrane integrity and result bleb formation and cell swelling. Activation of the calpain family of proteases has been reported following ischemia and in EAA-induced toxicity (Siman and Noszek, 1988; Kuwaki et al, 1989; Siman et al, 1989; Lee etal, 1991). Inhibition of calpain activity by calpain inhibitor I and leupeptin has been shown to be neuroprotective in tissue culture and in vivo models of hypoxia (Arai et al, 1991; Lee et al, 1991). It is not clear whether the role of calpain in mediating cell death is primarily due to its actions on structural components or on other regulatory processes in cells such as the conversion of P K C to the P K M , blockade of which is also neuroprotective. 15 1.6.1 Activation of P K C P K C is a family of closely related enzymes responsible for the phosphorylation of several intracellular, cytoskeletal, nuclear and plasma membrane-associated proteins (Nishizuka, 1992). P K C participates in the regulation of a wide variety of physiological functions, including the gating of ion channels, induction of gene expression, exocytosis, desensitization of membrane receptors, and induction of long-term potentiation in the hippocampus (see Huang, 1990). It has recently been shown that P K C can phosphorylate some glutamate receptor subtypes, thereby potentially allowing for an activity-dependent regulation of E A A receptors (Tan et al, 1994). PKC appears to be ubiquitously expressed, with several isoforms present in brain (Table 1.1). The PKC isoforms were determined by cloning studies to be products of distinct genes as well as of alternatively spliced mRNA transcripts and can be classified into two major groups based on their structural similarities and Ca2+-dependence. Group A P K C isoforms such as P K C - a , -P, and -y, are dependent upon the presence of Ca 2 + as well as membrane phospholipids and D A G (Kikkawa and Nishizuka, 1986) for their activation. Activation of these and other P K C isoforms by phorbol esters can lead to changes in the Ca2+-dependency of the kinase (Pelech et al, 1991). Prolonged phorbol ester exposure leads to the down-regulation of PKC, presumably as a result of enhanced degradation (Tsukuda et al, 1988). Group B isoforms are distinct from those of group A in that they lack a region of conserved amino acids which are thought to confer the Ca 2 + -dependence of the group A P K C isoforms. Expression of group B P K C isoforms in COS cells by transfection with their respective cDNAs has demonstrated that these isoforms are independent of Ca 2 + for activity (see Huang, 1990). Thus, it is possible that the group B P K C isoforms are not involved in Ca2+-mediated cell death. 16 Table 1.1 Patterns of expression of selected P K C subspecies" Tissue-specific patterns of some P K C isoforms in group A and B. List is not inclusive of all forms of P K C described to date. Tissue localization was determined by in-situ hybridization and subcellular localization was performed by immunohistochemistry in most cases. P K C Tissueb Cellular Compartment Group A a brain, other0 except liver perikaryon, perinuclear area pr brain, liver, other except kidney perikaryon adjacent to plasma membrane PIP brain, liver, other except kidney cytoplasm around Golgi complex Y brain only cytoplasm, nucleus Group B 6 brain, other particulate fractions extracted with non-ionic detergent e brain, other except liver soluble fractions of cell homogenates c brain, other N D d skin, lungs, brain, spleen N D e skeletal muscle, other N D a Compiled from: Huang, 1989; Tanaka and Saito, 1992; and Hug and Sarre, 1993. b Sorted in order of levels of isoform detected (greatest to least). 0 Other includes lungs, spleen, thymus, and skin. d N D not determined. e pi and pil are derived by alternative splicing. 17 Perhaps the best evidence that Ca2+-dependent processes mediate cell death comes from studies which show that P K C activity is increased following NMDA-stimulated Ca 2 + influx. Also, P K C activity appears to be altered in some chronic neurodegenerative diseases (Saitoh et al, 1991). P K C may participate in the neurotoxicity associated with E A A exposure. Studies of cultured neurons demonstrate that NMDA-associated neuronal death can be attenuated by addition of inhibitors of PKC (Maiese etal, 1993; Felipo etal, 1993). Furthermore, the survival of cultured neurons increases if P K C is down-regulated prior to N M D A application (Favaron et al, 1990). However, the literature is conflicting as to whether P K C mediates or protects against cell death in animal models of global and focal cerebral ischemia (Hara et al, 1990; Aranowski et al, 1992; Madden et al, 1991; Zivin et al, 1990). Ambiguity also exists over the effects of phorbol ester-induced PKC activation on cell proliferation or differentiation in the macrophage cell line HL-60 (Clemens et al, 1992). In this instance, the conflict may likely be reconciled by the observation that cells proliferate in response to low concentrations of phorbol ester, but terminally differentiate into a phenotype characteristic of macrophages at high phorbol ester concentrations (Clemens et al, 1992). Slight differences between PKC isoforms in the requirements for activation, subcellular localization, and substrate specificities may be important variables for producing the different dose-dependent effects of phorbol ester. It is likely that the physiological effects of P K C activation in different cells is determined by differences in the level of P K C activation, the specific P K C isoforms present, or activated in a cell, and/or the mechanism by which P K C is activated. The relation between alteration of Ca 2 + homeostatic mechanisms and PKC activation has not been clearly defined. It appears likely that PLC-mediated phosphatidylinositol 18 4,5-bisphosphate (PrP 2 ) hydrolysis is not the only mechanism responsible for P K C activation, as was once thought (Ha and Exton, 1993; Huang et al, 1993; Dell and Severson, 1989). Several models for P K C activation have been proposed based on the biophysical properties of the enzymes and their interactions with membrane phospholipids, D A G , and Ca 2 + . One model for P K C activation involves a Ca2+-promoted interaction of the Ca2+-dependent PKCs with membrane phospholipids to form a pre-activated form of the enzymes that becomes activated upon interaction with PIP2-derived D A G . Thus, the activation of P K C can involve its translocation from cytosol to membrane (Huang, 1989), but this translocation does not necessarily result in enzyme activation (ie in the absence of D A G interaction). Furthermore, membrane-associated P K C can either be inserted into the membrane, or bound to another membrane protein. Activation of P K C may also occur independently of membrane translocation (Pelech et al, 1990a; Lu et al, 1994), perhaps as a result of changes in the phosphorylation state of the protein, or DAG-stimulated activation of pre-existing membrane-associated PKC. Finally, proteolysis of P K C can result in the generation of the Ca2+-independent kinase, protein kinase M (PKM), through cleavage by the Ca2+-activated proteases, calpains I and TJ (Melloni etal, 1985; Kishimoto et al, 1989; Savart et al, 1992). Studies which show increased P K C activity under cytotoxic conditions have generally focused only on one of the possible mechanisms of P K C activation. In addition, studies have not addressed the Ca2+-dependencies of the possible mechanisms of P K C activation, and there is controversy as to which mechanisms are important in mediating toxicity. Since PKC has also been implicated in several other cellular processes including long-term potentiation and cell development, it is possible that combinations of these mechanisms might be important under different circumstances. 19 1.7 Objectives To understand the role of Ca 2 + in cell death, we must first attempt to define the effects of increases in [Ca 2 +] ; on the activation of processes downstream of Ca 2 + influx or release. In addition, the system chosen must have relevance to the disease process under investigation. For studies relevant to motor system degeneration in man, experiments on single neuronal cells, using a Ca2+-indicator such as fura-2, would be helpful in determining the relationship between Ca 2 + and motoneuron death. However, it has been technically difficult to evaluate Ca 2 + homeostasis in primary cultures of mammalian motoneurons due to the limited ability of these neurons to survive in culture (Krieger and Kim, 1991). Also, the limited purity of primary motoneuron cultures might complicate the analysis of Ca 2 + responses to various stimuli. To simplify the evaluation of cultured single cells with motoneuron properties, we have used the motoneuron cell line, NSC-19. NSC-19 cells were derived from cell fusion between motoneuron-enriched primary embryonic spinal cord cells and N18TG2 neuroblastoma cells (Cashman et al, 1992). This cell line was used to evaluate Ca 2 + responses in a homogenous population of cells under conditions which mimic hypoxia, and which do not involve the effects of non-neuronal cells. Due to the presence of EAA-releasing cells in primary cell cultures, studies which demonstrate effects of metabolic inhibition on cell survival have not been able to rule out the possibility of E A A involvement. It was the purpose of the first part of this study to define the characteristics of alterations in [Ca2+]j seen with changes in [Ca 2 +] 0 and to determine whether metabolic inhibition (chemical hypoxia) was accompanied by changes in [Ca 2 +] ; in this motoneuron cell line. Additionally, we evaluated whether the changes in [Ca2 +]; with metabolic inhibition were due to internal release or 20 transmembrane influx of Ca 2 + . We evaluated [Ca2 +]; in this motoneuron cell line in response to two metabolic inhibitors, amobarbital and carbonyl cyanide /w-chlorophenylhydrazone. Amobarbital (Amytal) is an inhibitor of the NADH-coenzyme Q reductase complex (site I of the mitochondrial respiratory chain) (Slater, 1967). Carbonyl cyanide 7w-chlorophenylhydrazone (CCCP) transports protons across phospholipid membranes and uncouples oxidative phosphorylation in mitochondria, resulting in a reduction of the mitochondrial membrane potential (Gunther and Pfeiffer, 1990) and lowered ATP concentrations (Burkhardt and Argon, 1989). CCCP has also been shown to increase Ca 2 + permeability through the mitochondrial membrane (Gunther and Pfeiffer, 1990). We also examined the relationship between [Ca 2 +] ; and cell viability with metabolic inhibition. In the second part of this study, we investigated the effects of changing [Ca 2 +] 0 on the activation of P K C in NSC-19. The purpose of this part of the study was to determine whether modest but sustained increases of [Ca2+]; would result in changes in the activity of P K C and whether transient versus sustained Ca 2 + responses resulted in different patterns of PKC activity. In addition, we examined whether changes in [Ca2+]j would result in the generation of P K M , the constitutively active enzyme which contains the catalytic subunit of PKC. Generation of P K M could have important physiological consequences. 21 Chapter 2 Materials and Methods 2.1 Materials Amytal was obtained from Eli Lilly Co. (Scarborough, ON), CCCP from Sigma (St. Louis, MO), and vanadate was from Alomone Labs (Jerusalem, Israel). Fura-2 was purchased from Molecular Probes (Eugene, OR). Radioactive [y- 3 2P]ATP was obtained from Dupont (Mississagua, ON). Antibodies were purchased from Upstate Biotechnology (Lake Placid, NY) , Transduction Laboratories (Lexington, K Y ) , or Santa Cruz Biotechnology (Santa Cruz, CA). Protein kinase A inhibitor peptide (PKLP, amino acids 5-24) (Scott et al, 1986; Cheng et al, 1986) and all other chemicals and materials were obtained from Sigma or Fisher Scientific (Vancouver, B.C.). Amytal, bepridil, vanadate and ruthenium red (RR) were prepared by dissolving in FfBSS. CCCP and amiloride were added to buffer solutions from 1 mM dimethylsulfoxide (DMSO) stock solutions (the final concentration of DMSO did not exceed 0.5%). Phosphatidylserine (PS) and diolein (DO) stock solutions were prepared by dissolving each in PKC assay buffer (10 mM MOPS, 25 mM p-glycerophosphate, 2 mM EGTA, 2 mM EDTA, 0.25 mM DTT, pH 7.2) after evaporation of chloroform under N 2 (gas) to concentrations of 300 ug/ml (PS) and 30 ug/ml (DO) for P K C assays. 2.2 Cell culture Monolayer cultures of the motoneuron cell line, NSC-19, were used for all experiments. NSC-19 is a clonal hybrid cell line derived from the fusion of the aminopterin-sensitive neuroblastoma N18TG2 with primary dissociated spinal cord cells enriched for motoneurons 22 (Cashman et al, 1992). Spinal cord cells were obtained from CD1 mice aged embryonic day 12 to 14. Unlike its neuroblastoma parent, NSC-19 expresses properties observed in motoneurons including: generation of action potentials, expression of choline acetyltransferase and neurofilament triplet proteins, induction and twitching in cocultured myotubes and synthesis, storage and depolarization-evoked release of acetylcholine (Cashman et al, 1992). Cells were grown in 75 cm 2 culture flasks until confluent (approx 6-8 days) in Minimum Essential Medium (MEM, Stem Cell Technologies, Vancouver, B.C.), supplemented with 10% heat-inactivated fetal calf serum (Gibco, Grand Island, N Y ) at 5% C 0 2 - 95% air at 37°C. Half the medium in culture dishes was replaced three times weekly. One week prior to calcium imaging experiments, cells were transferred onto poly-D-lysine coated glass coverslips in six well plates (Corning, NY) . 2.3 Fluorescence studies 2.3.1 Cell loading and buffers NSC-19 cell cultures grown on 25 mm glass coverslips were studied when almost confluent. Cells were transferred to a HEPES-buffered Hanks' balanced salt solution (HBSS, pH 7.2) containing (in mM): NaCl, 137; KC1, 5.4; Na 2 HP0 4 , 0.16; NaHC0 3 , 4.22; MgS0 4 , 0.41; MgCl 2 , 0.49; HEPES, 20; glucose, 5.5; CaCl 2 , 1.26. Exceptions to this composition were as follows: glucose-free, glucose was omitted; Na+-free, Na + was replaced with equimolar amounts of L i + ; Ca2+-free, Ca 2 + was omitted and 50 u M E G T A was added. In buffer solutions with higher Ca 2 + concentrations, Na + was replaced iso-osmotically for Ca 2 + . The pH was checked and adjusted with NaOH after any additions and before the start of each experiment, except for Na + -free buffers which were adjusted with LiOH. 23 [Ca2+]j was measured using the fluorescent Ca 2 + indicator fura-2 acetoxymethyl ester (fura-2/AM). The fluorescent dye was dissolved in DMSO and solubilized with Pluronic F-127 (0.03%, Molecular Probes) in HBSS containing 0.02% bovine serum albumin (BSA). The final concentration of DMSO in this loading solution did not exceed 0.5%. Cells were incubated in 5 u M fura-2/AM solution for 30 min at room temperature (22 °C) during which time cytosolic esterases hydrolyze the fura-2/AM to fura-2, the membrane-impermeant polycarboxylate anion. Cells were washed twice with HBSS and allowed to incubate for at least an additional 30 min in HBSS to ensure complete hydrolysis of the fur a-2/AM to fura-2. Coverslips containing cells loaded with fura-2 were mounted in a perfusion chamber with a working volume of 0.2 ml (Warner Instrument Corp., Hamden, CT), and fitted onto the stage of an inverted microscope. Cells were perfused with HBSS (at 22 °C) at a flow rate of approximately 1 ml/min. The buffers used in most experiments were supplemented with glucose (5.5 mM), since this appeared to increase cell viability over extended periods. Cells incubated in the absence of glucose tended to lose their fluorescence signals more rapidly than those in glucose although this did not result in any significant differences with respect to [Ca 2 +] ; in treated cells. 2.3.2 Calcium imaging Fluorescence measurements were made using a Nikon 40x fluorite (1.3 NA) epifluorescent objective fitted to a Nikon Diaphot T M D inverted microscope, equipped with a 100 Watt xenon light source (Osram, Germany). During imaging, light was selectively filtered at 340 and 380 nm by a computer controlled filterwheel. Emitted fluorescence was detected by a Silicone Intensified Target (SIT) video camera (Hamamatsu C2400-08) and fed to an 80386 24 based desktop computer, equipped with an image detection software package and an analog-to-digital video digitizer board (Image-l/Fl, Universal Imaging Corp., West Chester, PA) (Fig. 2.1). The software also controlled the shutter/filterwheel and displayed images taken at the two wavelengths, after background subtraction, as well as an image of the 340/380 nm ratio on a RGB colour monitor. Ratios were obtained from 8-frame averages of pixel intensities (ranging from 0 to 255) at each of 340 nm and 380 nm, once every 5 seconds. Ratios were then converted to calcium concentrations using the method described by Grynkiewicz et al. (1985). SEM's were calculated for each data point, but were only plotted (as error bars) at a few points for clarity. 2.3.3 Calibration of the imaging system The system was calibrated using the fura-2 pentapotassium salt (fura-2/K5, 5 uM) with a series of calibration buffers containing: KC1100 mM, MOPS 10 mM and Ca 2 + -EGTA and E G T A in ratios to give Ca 2 + concentrations between 0 and 39.8 uM (pH 7.2, 22°C, Molecular Probes). Using ratio values obtained from the calibration buffer solutions, a K d for fura-2/K5 was determined according to the formula [Ca 2 +] ; = K d x p x (R,^ - R) / (R - R ^ J ; where p is the ratio of fluorescence intensities at 380 nm under zero and saturating Ca 2 + concentrations. This K d value was assumed to be similar to the K d for the intracellular fura-2 anion (see Williams et al, 1985), and was used in the calculation of all [Ca 2 +] ; concentrations according to the ratioing method described by Grynkiewicz et al. (1985). 25 Fig. 2.1 Experimental setup of imaging system A schematic diagram of the digital imaging microscope used to measure cytosolic [Ca 2 +] in fura-2 loaded cells. [Adapted from Moore et al, 1990] Fura-2 Loaded Cell N I , Shutter ^ - l e n g t h Xenon(UV) Light Source Nikon 40x Objective Filter Dichroic Mirror (410 nm) Emmision C Camera Filter (510 nm) ecu Microscope pontrol Logic Computer 5 i I Analog-to-digital board Hard Disk c —^ f \ L J o o o o Computer Monitor Video Monitor 26 Maximum ratios ( R ^ and 380 nm fluorescence intensities (F^^g^) were obtained by the addition of the Ca2+-ionophore 4-bromo-A23187 (10 uM, Molecular Probes) to the cells in HBSS containing 1.3 mM Ca 2 + . Minimum values (R,,^ and F ^ ^ o ^ ) were obtained by adding E G T A (typically 5 mM) to cells in the bath. Al l measurements were made on single, phase-bright cells with evident processes and recordings were made only of cells whose initial baseline ratios were stable for at least 3 minutes. Each experiment was performed at least three times. 2.3.4 Determination of cell viability Intravital staining with fluorescein diacetate and propidium iodide was performed as described elsewhere (Favaron et al., 1988). Briefly, coverslips containing cells were removed from M E M , washed in HBSS, and incubated for 3 min at 22 °C with a mixture of fluorescein diacetate (15 ug/ml) and propidium iodide (4.6 ug/ml). Fluorescein diacetate is a non-polar ester which crosses cell membranes. Once inside living cells, the ester is hydrolyzed by cellular esterases to produce a fluorescent green colour. Propidium iodide will not penetrate cells unless membrane permeability is compromised. Entry into cells results in the interaction of this dye with D N A leading to the formation of a bright red fluorescent complex. Cells were treated with CCCP (or HBSS) for 60-70 s, and incubated in 95% air-5% C 0 2 for various times before staining. The labelled cells were examined immediately with a fluorescence microscope equipped with a fluorescein / rhodamine filter set. Since cells inevitably float off the coverslips, even in HBSS-treated (control) cultures, images of the fields were stored on computer prior to treatment and after staining to facilitate quantification. Cell death was taken as the percent of live cells in three randomly chosen fields on each coverslip. These percentages were normalized to controls. 27 2.4 Biochemical analyses 2.4.1 Preparation of cell extracts NSC-19 cells were grown in M E M with 10% heat-inactivated fetal bovine serum for a period of 7-10 days or until confluent (approximately 107 cells). Cells were washed once in phosphate buffered saline for 1 min and then treated in HBSS containing various Ca 2 + concentrations at room temperature. EfBSS was then replaced with 3 ml ice cold lysis buffer containing: 10 mM MOPS (pH 7.2), 25 mM p-glycerophosphate, 2 mM EGTA, 2 mM EDTA, 0.25 mM dithiothreitol (DTT), 1 ug/ml leupeptin, 1 ug/ml pepstatin, 1 mMPMSF, and 1 mM benzamidine. Each treatment was performed in triplicate in 75 cm 2 flasks and the cells were scraped and pooled into 1.5 ml Eppendorf microcentrifuge tubes, on ice, and centrifuged at 1000 rpm (Biofuge) for 5 min at 4 °C. The lysis buffer was then aspirated and replaced with 0.6 ml of fresh ice cold buffer. Cells were then sonicated for 6 s, 3 times (at the 40 setting of a Sonics Vibra-Cell) on ice and clarified by centrifiigation (10,000 x g, 15 min) at 4°C. The supernatant was then subjected to further centrifiigation (200,000 x g, 25 min) at 4°C in a Beckman TL-100 ultracentrifuge and the cytosolic cell extract (supernatant) was immediately aliquoted and stored at -70 °C. The pellet was sonicated as before in 0.5 ml of lysis buffer containing 1% Triton X -100. The detergent-solubilized (membrane) extract was obtained after ultracentrifugation (200,000 x g, 25 min, 4°C) and was also aliquoted and stored at -70°C. 2.4.2 Determination of protein content The protein content of cytosolic (soluble) and membrane (particulate) extracts was determined using the method of Bradford (1976). Briefly, 5 and 10 ul of cell extract were made 28 up to 100 ul with buffer and added to 2.5 ml of Bradford reagent which contained (in 1 litre of aqueous solution): 100 mg Coomassie Blue G250, 50 ml 95% ethanol, and 100 ml 85% phosphoric acid. The absorbance at 595 nm ( A ^ ) was measured using a spectrophotometer after 15 min incubation at room temperature. The concentration of protein per sample was then calculated by comparison of the Ajgj from the extracts to the A 5 9 5 obtained from serial dilutions of a 1 mg/ml standard solution of BSA. The protein concentrations obtained from duplicate determinations were averaged and taken to be the concentration of protein in the cell extracts. On average, cytosolic and membrane extracts contained 2 mg and 1 mg total protein per ml, respectively. 2.4.3 Partial purification of PKC and P K M Extracts were rapidly thawed and 0.8 mg of cytosolic or Triton X-100 solubilized membrane extracts were fractionated on a 1 ml MonoQ (5/5) anion exchange column connected to a fast protein liquid chromatography (FPLC) system (Pharmacia). Fractions (0.25 ml) were eluted at a flow rate of 0.8 ml/min using a linear gradient of 0-0.8 M NaCl in 10 ml MonoQ buffer containing: 10 mM MOPS (pH 7.2), 25 mM p-glycerophosphate, 2 mM EGTA, 5 mM EDTA, 2 mM N a 3 V 0 4 , and 0.25 mM DTT. 2.4.4 Protein kinase assays MonoQ column fractions and crude extracts were assayed for the presence of P K C and P K M activity by measuring the extent of 3 2 P transfer from [y 3 2-P]ATP to histone HI (Sigma Type HIS). Column fractions or crude extracts (5 ul) were assayed for 10 min at 30°C in an assay tube 29 (final volume of 25.5 ul) which contained: histone HI (1 mg/ml), 50 u M [y 3 2-P] ATP (1250 cpm/pmol), 25 mM p-glycerophosphate, 10 mM MOPS (pH 7.2), 10 mM MgCl 2 , 2 mM EDTA, 2 mM EGTA, 1 mM N a 3 V 0 4 , 0.25 mM DTT, and 0.5 u.M cAMP-dependent protein kinase inhibitor peptide (PKIP). Incubations containing PKC activators also included: 60 ug/ml PS; 6 ug/ml DO; and 4.5 mM CaCl 2 (estimated at a free Ca 2 + concentration of 0.8 mM by the method of Fabiato and Fabiato, 1979). Reactions were terminated by spotting a 20 ul aliquot of the reaction mixture onto 2 x 2 cm pieces of Whatman P81 phosphocellulose paper which were dried for 30 s, and washed several times with 1% (v/v) orthophosphoric acid. The filter papers were then transferred to scintillation vials with 0.5 ml of Ecolume (ICN) scintillation fluid and analyzed for radioactivity using a Beckman LS5000 scintillation counter. Additionally, the radioactivity of four 5 ul of 250 u M [y 3 2-P]ATP samples was measured and averaged with each experimental run in order to account for the day-to-day and lot-to-lot variations in ATP radioactivity. 2.4.5 Determination of cytosolic adenine nucleotides Determination of cytosolic adenine nucleotides was performed by D. Naumann and G. Ritchie on confluent cultures of NSC-19 cells. Culture media was replaced with a balanced salt solution (BSS) which contained (in mM): NaCl, 147.2; KC1, 2.2; CaCl 2 , 1.8; MgS0 4 , 0.8; K H 2 P 0 4 , 0.4; Na2HP0 4, 0.3; HEPES, 5.0 (pH 7.4); with or without glucose (5 mM). BSS with or without 5 mM Amytal and 10 u M CCCP were added to individual wells and 200 ul of 0.3 M perchloric acid were added at specified time intervals to halt cellular enzymatic activity. Cells were scraped off with a plastic cell scraper, collected in a 1.5 ml Eppendorf tube and centrifuged at 15,000 rpm for 3 min. The supernatant was transferred and the protein pellet dissolved with 30 200 ul of 5% SDS in 0.2 M NaOH. The protein content was determined on 25 ul aliquots using a Lowry assay with B S A as a standard. Six ul of 10 M K O H were added to neutralize the supernatant which was centrifiiged again at 15,000 rpm for 3 min. The resulting supernatant was filtered into another Eppendorf tube and stored at -20 °C, to minimize ATP breakdown, until high performance liquid chromatography (HPLC) analysis. Sequential analysis of solutions showed no significant deterioration during storage. ATP, ADP and A M P were quantified by HPLC analysis using a hydrophobic silica gel column (Millipore uBondapak, 3.9 x 150 mm) eluted at 2 ml/min with 100 mM K 2 P 0 4 (pH 5.5) via a Millipore-Waters 600 Multisolvent Delivery System. The eluent was analyzed by measuring absorbance at 254 nm with a spectrophotometer (Millipore-Waters Lambda-Max Model 481) and the resulting signal was fed into an integrator/recorder to calculate the areas under the peaks. The areas were then compared to pure ATP, ADP, and A M P standards purchased from Sigma. Adenine nucleotide concentrations were expressed in nmoles/mg protein. 2.5 Western blot analyses Crude extracts or MonoQ fractions were resolved by SDS-PAGE on 11% gels. Samples were boiled for 3 min following addition of 5x Sample buffer containing: 10% SDS, 0.5 M Tris HC1 (pH 6.0), glycerol, concentrated 2-mercaptoethanol, and 0.1% bromophenol blue. Electrophoresis was performed overnight at 8 mA per gel. Proteins were then transferred (3 h, 0.3 A) to nitrocellulose membranes and probed for the presence of P K C and P K M using antibodies directed against the carboxyl-terminal (C-terminal) catalytic domains of P K C and P K M (as described below). Visualization was performed using a kit which employed enhanced 31 chemiluminescence (Amersham) following incubation with horseradish peroxidase (HRP)-conjugated anti-rabbit (1:5000) or anti-mouse (1:3000) IgG (Transduction Laboratories). Films were then scanned (Abaton Scan 300 Colour / Adobe Photoshop) into TIFF format and band densities were quantified using the NUT Image plotter program (version 1.57). A mixture of a-, P-, and y-PKC was purified from rat brain as described elsewhere (Kitano et al, 1986) and used as a positive control for antibody specificity. 2.5.1 Anti-PKC antibodies Antibodies directed against P K C were obtained from various sources and were directed against different regions of the C-terminal portions of various P K C isoforms as follows: i) UBI: • Anti-Pan PKC, rabbit (polyclonal): Synthetic peptide corresponding to C-terminal sequences of rabbit protein kinase type pH: TRHPP VLTPPDQE VIJAMDQ SEFEGFSF V N S E F L K P E V K S . ii) Santa Cruz Biotechnology: • cPKC a (C-20), rabbit (polyclonal): Epitope corresponding to amino acids 651-672 mapping at C-terminus of rabbit cPKCcc. • P K C (C4), rabbit (polyclonal): Epitope corresponding to amino acid residues 528-537 mapping within conserved C4 catalytic domain of human protein kinase C. iii) Transduction Laboratories: 32 • Anti-PKC a, mouse (monoclonal): Immunogen was an 18 kDa fragment of rat brain P K C a corresponding to amino acids 270-427. • Anti-PKC y, mouse (monoclonal): Immunogen was an 22.3 kDa C-terminal fragment of P K C y corresponding to amino acids 499-697. 2.6 Statistics Data are presented as mean ± S E M unless otherwise noted. Statistical comparisons between groups were made using Student's t-tests. Individual imaging experiments were performed on at least three different cultures (coverslips) and the "n's" indicated in the results refer to the number of cells studied. Each experiment (imaging or biochemical) was performed at least three times using cultures of comparable passage number. 33 Chapter 3 Results 3.1 Fura-2 calibration and basal [Ca 2 +] ; in NSC-19 cells The [Ca2+]j was calculated as described in Materials and Methods. Fluorescence measurements were made on cells which were pre-loaded using a solution containing 5 u M fura-2 / A M as this was the lowest fluorochrome concentration which allowed cells to load uniformly and without evidence of dye compartmentalization. The apparent dissociation constant for fura-2 binding (K d) was determined to be 213 ± 6 nM (Fig. 3.1) and the ratio of fluorescence at 380 nm under zero Ca 2 + and saturating Ca 2 + conditions, 'P' (see Section 2.3.3 in Materials and Methods) was determined to be 6.5 ± 0.7. The basal [Ca 2 +] ; of cells in FIBSS with or without Ca 2 + approximated normal distributions (Fig. 3.2). The mean [Ca 2 +] ; value of cells incubated for up to 1.5 h in 1.3 mM Ca2+-containing FfBSS was 83 ± 17 nM (mean ± S.D., n = 506 cells, Fig. 3.2a) and was significantly different (p < 0.001) from cells in Ca2+-free FfBSS (51 ± 14 nM, mean ± S.D., n = 447 cells, Fig. 3.2b). 34 Fig. 3.1 Calibration of fura-2 imaging Relationship between fura-2 fluorescence ratio and [Ca 2 +]. Fura-2 pentapotassium salt (5 uM) was added to small volumes of buffer containing known concentrations of C a 2 + as described in Materials and Methods. Ratios (n=10 for each point) were determined under the digital imaging microscope from fluorescence intensities at 340 nm and 380 nm excitation wavelengths and corrected for background fluorescence. The P value was determined from the 380 nm fluorescence intensities under zero C a 2 + and saturating C a 2 + conditions. o oo <n TJ-+1 & c c S o c o O E o 0-^ 1 + .3 g •1+ -2-4--3 T • J. • • + P = 6.5 ± 0.7 K d = 213 + 6 nM H ' 1 --8 -5 log [Ca 2 + ] -2 35 Fig. 3.2 Distribution of basal [Ca 2 +]i in NSC-19 cells Distribution of basal [Ca 2 + ] ; in NSC-19 cells loaded with Fura-2 as described in Materials and Methods and incubated for 1.5 h in FfBSS with (A) 1.3 mM [Ca 2 + ] 0 and (B) zero [Ca 2 + ] 0 . 180-160-140-# 120-1 g 100-0 | 80-S 1 60-40-20-0-180-160-140-«» 120-1 !=l 2 100-o I 80-60-40-20-0-B 15 1 I Mean [ C a 2 ^ = 83 ± 17 nM n = 506 cells i YZX. ~~\ 1 1 1 r — i 1 1 1 — C P — r 25 35 45 55 65 75 85 95 105 115 125 135 145 [Ca 2 + ] i (nM) 25 35 45 T 55 Mean \ c £ + \ = 51 ± 14 nM n = 447 cells I i i i T 65 T 75 85 r * — i " I — r — 95 105 115 125 [Ca 2 + ] i (nM) 36 3.2 Effect of changing [Ca 2 +] 0 on [Ca 2 +] ; Changing the [Ca2 +]„ resulted in either sustained (Fig. 3.3a) or transient (Fig. 3.3b) changes in [Ca 2 ^. Removal of [Ca 2 +] 0 resulted in a rapid reduction of [Ca2 +]; which reached a mean steady state level after 112 ± 11 s. The steady state mean [Ca 2 +] ; achieved (30 ± 8 nM, Fig 3.3a, "0 mM") was not significantly different from the basal [Ca 2 +] ; of cells in Ca2+-free HBSS described above, indicating that [Ca2+]i is rapidly altered by changes in [Ca 2 +] 0. When [Ca 2 +] 0 was reduced from 1.3 mM to 0.7 mM, [Ca2 +]; decreased to 48 ± 6 nM (Fig 3.3a, "0.7 mM") which was intermediate between the [Ca 2 +] ; levels for cells in 1.3 mM Ca2+-containing HBSS and Ca 2 + -free HBSS. The mean time for the cells to reach a constant [Ca 2 +] ; after reducing the [Ca 2 +] 0 from 1.3 mM to 0.7 mM was 115 ± 12 s, which was similar to the time for equilibration for cells transferred to Ca2+-free HBSS. When [Ca2 +]„ was elevated from 1.3 to 2.5 m M Ca 2 + (Fig 3.3a, "2.5 mM"), [Ca 2 +] ; rose to a steady state value of 140 ± 8 nM which was maintained for at least 10 min. Changing [Ca 2 +] 0 from 1.3 to 5 and 10 mM (Fig 3.3a, "5 mM" and "10 mM") also resulted in sustained increases in [Ca2 +]; to 232 ± 24 nM and 365 ± 1 3 nM, respectively, which were also maintained for at least 10 min. The rate of rise of [Ca2+]; was greater when [Ca 2 +] 0 was changed to 5 and 10 mM than when [Ca 2 +] 0 was changed to 2.5 mM. When NSC-19 cells were exposed to HBSS containing 25 and 50 mM Ca 2 + (Fig 3.3b, "25 mM" and "50 mM"), [Ca2 +]; rose transiently within 2.5 min to values greater than 600 nM and 900 nM, respectively. [Ca 2 +] ; then returned to 202 ± 1 4 nM and 122 ± 6 nM, respectively, and remained at that level for at least 10 min in the presence of 25 and 50 mM [Ca 2 +] 0. The transient nature of the profiles observed with 25 and 50 mM [Ca 2 +] 0, compared with the steady increase after exposure to 2.5 mM, or 5 and 10 mM [Ca 2 +] 0, suggests that different mechanisms likely mediate the changes in [Ca2+]i in NSC-19 37 cells depending on [Ca 2 +] 0. When 1.3 m M Ca 2 + was added to cells which had been in Ca2+-free HBSS for 1.5 h, [Ca2 +] increased to 97 ± 11 nM (Fig 3.4). The [Ca2 +]; level reached corresponds to the mean [Ca 2 +] ; of cells which had been incubated for an extended period (1.5 h) in 1.3 mM Ca2+-containing HBSS. The time for the [Ca 2 +] } to plateau (255 ± 9 s) was similar to the time for the rise in [Ca 2 +] { seen when the [Ca 2 +]D was raised from 1.3 to 2.5 mM (Fig. 3.3a, "2.5 mM"). The mean time for equilibration of [Ca 2 +] ; when [Ca 2 +] 0 is increased differs from that when it is decreased, suggesting that different mechanisms likely also mediate the changes in [Ca2 +]; under these two conditions. 3.3 Role of Ca 2 + ion channels in changing [Ca2 +]; To evaluate whether NSC-19 cells had voltage-dependent Ca 2 + channels (VDCCs) or excitatory amino acid (EAA) receptor-gated channels, bath application of KC1 (50-100 mM) and glutamate (500 uM) was performed, but did not evoke increases in [Ca2+]j in 35/36 cells and 61/63 cells, respectively. In addition, the calcium channel blockers verapamil (25 uM) or Lanthanum (La 3 + , 5 u M or 2 mM) did not affect the [Ca2 +]; responses of the cells to changes in [Ca 2 +] 0 when compared to untreated controls (data not shown), indicating that these are not the primary mechanisms by which changes in [Ca 2 +] 0 causes altered [Ca2+]j. 38 Fig. 3.3 Effect of changing [Ca 2 + ] Q on [ C a 2 ! (A) Cells were incubated in FIBSS containing 1.3 m M C a 2 + for 1.5 h before changing bathing solution to FIBSS with 10 mM, 5 mM, 2.5 mM, 1.3 mM, 0.7 mM, and 0 mM C a 2 + (top to bottom) in separate experiments. (B) Cells were exposed to 25 mM and 50 mM C a 2 + following incubation in 1.3 m M Ca 2 + . Traces indicate mean values for [Ca 2 +] ;; error bars indicate S E M for given mean values. For clarity, error bars are indicated at only a few points. 500 10 mM 5 mM 2 . 5 m M 1 .3mM 0.7 mM 0 mM 2 5 m M 5 0 mM 6 7 8 Time (min) 39 Fig. 3.4 Effect of increasing [Ca 2 + ] 0 on [Ca 2 + ] ; Cells were incubated in Ca2+-ffee FIBSS for 1.5 h before changing bathing solution to HBSS with 1.3 mM Ca 2 + . Traces indicate mean values for [Ca 2 +j;; error bars indicate S E M for given mean values. For clarity, error bars are indicated at only a few points. 150 40 3.4 Role of other Ca 2 + transporters in changing [Ca2+]j Transmembrane flux of Ca 2 + has also been associated with the action of the Ca 2 +-ATPase pump (Di Polo and Beauge, 1979) and the Na7Ca 2 + exchanger (Blaustein et al, 1991). To understand the possible contributions of these Ca 2 +-ion transporters in controlling the level of [Ca 2 +] ; in NSC-19, we employed drugs known to block their actions. Vanadate was used to block the action of the plasma membrane Ca 2 +-ATPase (Dehlingen-Kremer et al, 1991). Increasing [Ca 2 +] 0 in the absence of vanadate produced responses similar to those described above (see Fig. 3.3 a). The addition of vanadate (15 uM) to cells in Ca2+-free HBSS resulted in no significant change in [Ca 2 +] ;, but when 1.3 mM Ca 2 + was added to the external solution, the [Ca2+]; increased from 47 ± 4 nM to 271 ± 11 nM over a mean time of 11.8 ± 0.2 min before reaching a constant level (Fig. 3.5a). This increase was significantly greater than that observed in the absence of vanadate. The effect of vanadate on [Ca 2 +] ; in cells incubated in 1.3 mM Ca2+-containing HBSS was similar to the response shown in Fig. 3.5a. When 15 u M vanadate was added to cells in 1.3 mM Ca 2 + (Fig. 3.5b), the [Ca 2 +] ; increased rapidly until the Ca 2 + was removed from the external solution. Upon removal of the extracellular Ca 2 + , [Ca 2 +] ; did not increase further even in the presence of vanadate. These results suggest that the plasma membrane Ca 2 + - ATPase contributes to Ca 2 + efflux in the presence of 1.3 mM [Ca 2 +] 0. To determine if the Na7Ca 2 + exchanger is involved in the maintenance of [Ca 2 +] ; levels, the buffer solution was replaced with Na+-free HBSS so as to block the exchanger from working in the forward direction (Ca 2 + efflux) (Blaustein et al, 1991). The initial response in Fig. 3.6a shows the change in [Ca 2 +] ; to changes in [Ca 2 +] 0 when Na + is present in the bath. Replacement of Na + with L i + had little effect on the magnitude of the [Ca 2 +] ; response to increased [Ca 2 +] 0 compared to 41 control. Removal of extracellular Ca 2 + resulted in only a partial return of [Ca2+]j to baseline values indicating a more substantial effect on Ca 2 + efflux from NSC-19. A subsequent cycle of solution exchanges, varying [Ca 2 +] 0 between 0 and 1.3 mM, in the presence of N a + produced a response which was similar to control. We also employed amiloride, a blocker of the Na7Ca 2 + exchanger, which can also inhibit the Na7H + antiporter (Kleyman and Cragoe, 1988). The addition of amiloride (100 uM) to the bath resulted in a slight increase of [Ca2 +]; under Ca2+-free conditions (Fig. 3.6b). However, there was a significant increase in mean [Ca2 +]; with the addition of 1.3 mM [Ca2 +]„ (from 47 ± 3 nM to 204 ± 4 nM over a mean time of 13.3 ± 0.1 min before reaching a constant level). Reducing the [Ca 2 +] 0 back to 0 mM did not affect the [Ca 2 +] ; level of the cells (not shown). Bepridil is also an antagonist of Na7Ca 2 + exchange but not of the Na7H + antiporter and has a different mechanism of action than amiloride (Slaughter et al, 1988; Garcia et al, 1988). Addition of 100 u M bepridil to NSC-19 in 1.3 mM [Ca 2 +] 0 resulted in an increase in [Ca 2 +] ; from 6 9 ± 3 n M t o l l 3 ± 5 n M over a mean time of 220 ± 17 s (Fig. 3.6c). Washing the cells with 1.3 mM Ca2+-containing HBSS resulted in no significant decrease in [Ca 2 +] ;. The results obtained by using blockers of the Na7Ca 2 + exchanger, as well as those obtained in Na+-free media, indicate that the exchanger also plays a part in the efflux of Ca 2 + in the presence of external Ca 2 + . 42 Fig. 3.5 Effect of vanadate on [Ca2 +]j (A) [Ca 2 +]j responses of NSC-19 cells to changes in [Ca 2 + ] Q in the absence and presence of 15 u M vanadate (n=31). (B) [Ca 2 + ] ; responses to the addition of 15 u M vanadate to cells in 1.3 m M Ca 2 + -HBSS (n=32). Rules indicate media composition and bar indicates time of drug addition. 300-250 4 200-^ 150-1 p—i + C M u " 100-504 0-Van (15 uM) 1.3 mM 0 mM I 1 0 mM 4 8 250--i 1 1 1 > 1 1 j 1 r -12 16 20 24 28 Time (min) 32 + u 100-B Van (15 uM) 1.3 mM I 0 mM • I T—1—'—I— •—i—i—|—.— •> 1 1 1 • • 10 12 Time (min) 43 Fig. 3.6 Effect of blocking Na7Ca 2 + exchange on [Ca 2 + ] ; (A) Indicates [Ca 2 +]; responses to changes in [Ca 2 + ] 0 before, during and after replacement of N a + by equimolar concentrations of L i + (n=34). (B) [Ca 2 +]i responses to changes in [Ca 2 + ] 0 in the presence of 100 u M amiloride (n=38). (C) [Ca 2 + ] ; responses to addition of 100 bepridil in 1.3 mM Ca 2 + -HBSS (n=63). - A 100-f t s 7 5 " T 4 5 0 -u J -r NaMreeHBSS 2 5 - 1.3 mM 1.3 mM 1.3 mM | | 0 m M | I 0 mM I I 0 mM 0 - 1 1 1 1 1 1 1 — i 1 1 1 1 1 i 1 1 1 — 10 15 20 25 Time (min) 30 35 40 45 44 45 3.5 Effect of metabolic inhibition on NSC-19 cells 3.5.1 Inhibition by amobarbital (Amytal) The effects of exposure to Amytal (5 mM) for 70-80 s on the concentrations of adenine nucleotides in NSC-19 cells are presented in Table 3.1. Because the energy charge [(ATP + lA ADP)/(ATP + ADP + AMP)] of a cell has been related to metabolic function (Spragg et al, 1985), this parameter was also included in the table. Exposure of NSC-19 cells to Amytal in a glucose-free medium did not result in a significant change in ATP concentration or energy charge. The effect of Amytal exposure on [Ca 2 +] ; is shown in Fig. 3.7a. In the presence of 1.3 mM [Ca 2 +] 0 a transient increase in [Ca2+]; was observed when cells were exposed to 5 mM Amytal for 70-80 s. The mean [Ca 2 +] ; increased from a baseline level of 80 ± 8 nM to 156 ± 18 nM (Table 3.2 and Fig. 3.7a curve labelled "1.3 mM"). The mean time between the start of drug application and the peak [Ca 2 +] ; was 79 ± 13 s. After reaching maximum, the [Ca 2 +] ; returned to baseline levels over a (mean) period of 102 ± 18 s. In Ca2+-free HBSS, the mean peak [ C a 2 ! observed in response to Amytal (105 ± 9 nM, Table 3.2 and Fig. 3.7a, "0 mM") was significantly smaller (p < 0.001) than in 1.3 mM [Ca 2 +] 0. Since an increase in [Ca 2 +] ; is observed in cells in Ca2+-free HBSS, Ca 2 + must be released from intracellular stores, presumably from mitochondria. Also, the change in [Ca 2 +] t in the Ca2+-free condition is significantly smaller than that seen when extracellular Ca 2 + is present. This suggests that a proportion of the Amytal-induced increase in [Ca 2 +] ; in the presence of external Ca 2 + was due to a transmembrane influx of Ca 2 + . The mean time to peak was significantly faster in the Ca2+-free solution than in HBSS with Ca 2 + (54 ± 12 s compared to 79 ± 13 s, Table 3.2) suggesting that Ca 2 + release from intracellular stores may have occurred prior to transmembrane 46 influx. Following exposure to Amytal, the [Ca 2 +] ; returned to basal levels as under conditions where Ca 2 + was present extracellularly. To confirm that transmembrane influx of Ca 2 + occurs after Amytal exposure we added M n 2 + (100 uM), to the external solutions prior to Amytal (Fig. 3.7b). A small transmembrane influx of M n 2 + caused quenching of the fura-2 fluorescence when cells were excited at both 340 and 380 nm wavelengths. After Amytal exposure, the rate of fluorescence quenching rapidly increased, indicating an increase in the rate of transmembrane influx of divalent cations. 47 Table 3.1 Effects of Amytal and CCCP on ATP, ADP, A M P , and energy charge* Effects of Amytal and CCCP on nucleotide concentrations and on total charge. Cells were exposed to inhibitors for 1.2 min (70-80 sec) as described in text. Nucleotide concentrations are expressed in nmoles nucleotide/mg protein. Data was derived from 3 experiments each for Amytal and CCCP. Time refers to time after beginning of exposure to inhibitors. The 1.2 min time point was immediately after drug wash-off. AMYTAL CCCP Time (min) A T P A D P A M P Energy Charge A T P A D P A M P Energy Charge 0 22.7 ±2.2 7.7 ±1.5 4.0 ±1.3 0.77 19.5 ±2.8 12.9 ±2.0 3.6 ±0.7 0.72 1.2 23.1 ±0.5 8.5 ±0.5 2.9 ±0.8 0.79 12.8 ±1.4t 24.0 ±3.4 7.1 ±0.7 0.57T 2.5 19.9 ±2.3 8.2 ±0.9 3.9 ±0.7 0.75 17.0 ±2.3 18.2 ±2.5 5.2 ±0.6 0.65 5 20.4 ±0.5 7.9 ±0.8 4.2 ±0.6 0.75 16.1 ±1.3 14.6 ±2.5 3.5 ±0.2 0.68 7.5 21.7 ±3.2 8.0 ±1.6 4.6 ±1.1 0.75 17.6 ±1.3 12.5 ±2.5 2.3 ±0.6 0.74 10 22.2 ±1.9 7.4 ±0.4 4.8 ±0.4 0.75 16.9 ±1.8 13.7 ±3.3 2.3 ±0.4 0.6 NOTE: Values for ATP, ADP, and A M P given are means ± S.D. in nmol/mg protein. n=3 experiments performed for each of Amytal and CCCP. f different from control values p^0.05. * Energy Charge = [(ATP + ,/ 2ADP)/(ATP + ADP + AMP)] (Spragg et al, 1985). 48 Table 3.2 Effects of Amytal and CCCP on basal, peak, and recovery [Ca 2 +] ; Effects of Amytal and CCCP on basal, peak and recovery [Ca2+];. Time to peak refers to time between onset of drug exposure and peak [Ca2+];. Time to recover refers to time between peak [Ca2+]j and return to stable baseline values. Columns entitled 'with calcium' indicate [Ca 2 +] ; values obtained from cells in 1.3 mM [Ca 2 +] 0. Columns entitled 'without calcium' indicate [Ca 2 +] s values from cells in Ca2+-free media. A M Y T A L CCCP CCCP + ruthenium red With Calcium (n=43) Without Calcium (n=49) With Calcium (n=48) Without Calcium (n=53) With Calcium (n=32) Without Calcium (n=24) [Ca2+]i(nM) Resting 80 ± 8f 54 ± 6 Maximum 156±18j-* 105 ±9* Recovery 60 ± 7y* 42 ± 6* Time Course (seconds) To peak 79±13t 54 ±12 65 ± 6f 43 ± 2 343 ±7f* 190 ±3* 75±5f 90 ±10 70 ± 4t 47 ± 2 288 ±7t* 164 ±5* 110 ± 15J 100±10 To recover 102±18f 40 ±12 NOTE: Values shown are mean ± SEM. Time to peak refers to time between onset of drug exposure and peak [Ca 2 +] ;. Time to recover (Amytal-treated cells) refers to time between peak [Ca2+]j and return to baseline values. * Significantly different from resting calcium concentrations, p<0.001. f Significantly different from calcium-free condition, p<0.001. J Significantly different from calcium-free condition, p<0.01. 49 Fig. 3.7 [Ca ]; responses to Amytal (A) Mean [Ca 2 +]i of cells exposed to Amytal (5 mM) for 70 s in FfBSS containing 1.3 mM [Ca 2 + ] 0 ("1.3 mM", n=43) and in Ca2+-free HBSS ("0 mM", n=49) in separate experiments. (B) Effect of 5 mM Amytal on fura-2 fluorescence in the presence of M n 2 + (100 uM). The fluorescence emission of fura-2 loaded NSC-19 cells was recorded simultaneously at the two excitation wavelengths: 340 nm (bottom trace) and 380 nm (top trace). Error bars indicate S E M at given times. Time of drug application is indicated by bars. 50 3.5.2 Inhibition by carbonyl cyanide fw-chlorophenylhydrazone (CCCP) Exposure of NSC-19 cells to CCCP in glucose-free media produced a significant (p < 0.05) reduction in ATP concentration of about 33% which partially returned towards control values after drug wash off (Table 3.1). Energy charge was also transiently decreased by CCCP administration. When NSC-19 cells in 1.3 mM Ca2+-containing HBSS were exposed to CCCP (10 uM, in 0.5% DMSO) for 70-80 s, the mean [ C a 2 ! increased from 65 ± 6 nM to 343 ± 7 nM (Fig. 3.8a, curve labelled "1.3 mM"). Exposure to DMSO (0.5%) alone had no effect on [ C a 2 ! (data not shown). The mean time between CCCP application and observation of peak [ C a 2 ! w a s 75 ± 5 s, which was similar to the time observed until peak [ C a 2 ! a n"er Amytal exposure (Table 3.2). Unlike the response to Amytal, the CCCP-evoked rise in [ C a 2 ! did not return to baseline after removal of CCCP. The [ C a 2 ! l e v e l remained constant for at least 30 min under continuous perfusion with HBSS. The apparent lack of recovery suggests that the mitochondria were unable to take up cytosolic Ca 2 + ions following CCCP treatment. Although [Ca 2 +] ; did not return to baseline following CCCP exposure, this did not affect cell viability for up to 48 h after CCCP exposure as measured using the vital dyes fluorescein diacetate and propidium iodide. The percentages of viable cells after 0 h (100% ± 2 %, n = 885 cells), 2 h (98.1% ± 0.6 %, n = 275 cells), 5 h (100.0% ± 0.5 %, n - 358 cells), 24 h (99.8% ± 0.6 %, n = 446 cells) and 48 h (93.2% ± 0.9%, n - 227 cells) were not significantly lower than untreated controls. In contrast, exposure of cells to 10 mM glutamate resulted in 87 ± 8 % (n = 342 cells) cell death by 48 h. In Ca2+-free HBSS, exposure of cells to CCCP induced a mean rise in [Ca 2 +] ; from 43 ± 2 nM to 190 ± 3 nM (Fig. 3.8a, "0 mM"). This increase was significantly smaller than that observed 51 in the presence of extracellular Ca 2 + . As well, the mean time to reach peak [Ca 2 +] ; was 90 ± 10 s, which was significantly longer than the time to peak for cells in Ca 2 + -HBSS. No recovery of [Ca2+]j was seen after CCCP exposure, as with the cells in 1.3 mM [Ca 2 +] 0. CCCP, like Amytal, induced a smaller increase in [Ca2+]j under Ca2+-free conditions than when external Ca 2 + was present, indicating both release of intracellular (probably mitochondrial) Ca 2 + stores, and transmembrane influx of Ca 2 + ions. The presence of glucose in the medium may have had some influence on the changes observed in [Ca2+]j with CCCP treatment. We therefore omitted the glucose from the HBSS in some experiments (2 experiments each for 0 mM and 1.3 mM [Ca2 +]0). In glucose-free HBSS containing 1.3 mM Ca 2 + , the [Ca 2 +] ; increased from 85 ± 5 nM to 335 ± 9 nM (n=21) with CCCP. In glucose-free and Ca2+-free HBSS, CCCP caused an increase in [Ca2 +]; from 41 ± 4 nM to 192 ± 9 nM (n=47). These values were not significantly different from responses observed when glucose was present. To further characterize the increase in [Ca 2 +] t with CCCP, we employed ruthenium red (50 uM), an inhibitor of mitochondrial Ca 2 + uptake (Gunther and Pfeiffer, 1990), before CCCP exposure. The responses to CCCP were similar to those observed in the absence of the inhibitor except that the rise in [Ca 2 +] ; was smaller and the time to peak was longer in the presence of ruthenium red (Fig. 3.8b, Table 3.2). This suggests that mitochondria do not buffer or sequester the released Ca 2 + after CCCP treatment. The slight reduction in peak [Ca 2 +] ; observed in response to CCCP, when ruthenium red was applied prior to CCCP, might indicate that ruthenium red was also blocking release from a different calcium store. In the absence of ruthenium red, this second internal store might otherwise have contributed to the increase in [Ca 2 +] ;, either directly or 52 indirectly, upon exposure to CCCP. In experiments performed in Ca2+-ffee HBSS, cells were incubated in Ca2+-ffee buffer for 1.5 h prior to the start of recording (time = 0 in figures). During this time, mitochondrial Ca 2 + stores may have been depleted, resulting in the lower peak [Ca2+]j observed in comparison to that seen when 1.3 mM Ca2+-containing HBSS was used. To investigate the possibility that calcium depletion occurred during the incubation in Ca2+-ffee HBSS, we removed the extracellular Ca 2 + just prior to CCCP exposure (CCCP was applied less than 1 min after removal of extracellular Ca 2 +). Fig. 3.8c illustrates that the responses seen under these conditions are similar to that observed after extended incubation under Ca2+-ffee conditions. Both the peak [Ca 2 +] ; (220 ± 3 nM) and time to reach a constant level (96 ± 5 s) were comparable to the values observed after extended Ca2+-ffee conditions (Table 3.2), indicating that extended periods of zero Ca 2 + do not lead to reductions in [Ca 2 +] ; as a result of depletion of internal stores. The lack of depletion of mitochondrial stores further supports the view that a portion of the [Ca2 +]; response seen was due to transmembrane influx of Ca 2 + ions in the presence of extracellular Ca 2 + . In addition, treatment of NSC-19 with 10 u M CCCP in the presence of 100 u M MnCl 2 led to an increased rate of fluorescence quenching than before the addition of CCCP (data not shown). 53 Fig. 3.8 [Ca 2 +]i responses to CCCP (A) Mean [Ca 2 +]i of cells exposed to 10 u M CCCP for 70 s in HBSS containing 1.3 mM [Ca 2 + ] c ("1.3 mM, n=48) and in Ca2+-free HBSS ("0 mM", n=53) in separate experiments. In Figs. 3.8A and 3.8B, CCCP responses were obtained after cells had incubated in the indicated buffer for 1.5 h. (B) CCCP responses obtained following pretreatment with 50 u M ruthenium red (RR) (1.3 mM, n=32; 0 mM, n=24). (C) CCCP responses were obtained within 1 min of transferring NSC-19 cells to Ca2+-free HBSS (n=64). Error bars indicate S E M at given times. Time of drug application is indicated by bars and [Ca 2 + ] 0 is indicated by continuous rule at bottom. 54 55 3.6 Effect of changing [Ca 2 +] 0 on the activity of P K C Since NSC-19 cells are able to maintain reproducible levels of free [Ca2 +]; upon exposure to different levels of [Ca 2 +] 0, we investigated the effect of the different [Ca2 +]; on the activation of PKC. To determine whether sustained or transient increases in [Ca2 +]; altered the activity of PKC, we performed histone HI phosphorylation assays on cell extracts which had been exposed for 10 min to buffers containing 1.3, 2.5, 5, 10, 25, and 50 mM Ca 2 + . After exposure to buffers with different [Ca 2 + ] 0 , NSC-19 cell extracts were separated into soluble (cytosol) and particulate (membrane) fractions by ultracentrifugation. Extracts were then subjected to MonoQ anion exchange chromatography and column fractions were assayed for histone HI phosphotransferase activity either in the presence or in the absence of Ca 2 + , PS, and DO as described in Materials and Methods. Fig. 3.9a-c show representative MonoQ profiles of cytosolic and membrane fractions of NSC-19 cell extracts exposed to 1.3, 10, and 50 mM [Ca 2 +] 0, respectively. These profiles demonstrate the presence of two activity peaks (Peaks I and II), which eluted at approximately 0.3 M and 0.5 M NaCl. Peak I phosphorylating activity was determined to be Ca 2 + - and PS/DO-dependent as indicated by the differences between the profiles with Ca 2 + and PS/DO (closed symbols) and without activator (with EGTA, open symbols). Peak II activity was independent of Ca 2 + and PS/DO. The elution positions and activator dependencies of the two peaks are consistent with their respective identification as P K C and P K M as demonstrated elsewhere (Pelech al, 1990a;Pelechetal, 1990b;Pelechetal, 1991; Salariera/., 1990). The identification of the kinases in the two peaks was confirmed by immunoblot analysis (Fig. 3.10) of the MonoQ fractions using polyclonal antibodies specific for the C-terminal, catalytic domains, of P K C isoforms which are also retained in the catalytic fragment P K M . As shown in Fig. 3.10, 56 fractions 24-27 from MonoQ chromatography contained a =80 kDa band (PKC), whereas fractions 33-36 contained a =50 kDa protein corresponding to P K M (Nishizuka, 1984). The activities of P K C and P K M at each [Ca 2 +] 0 were determined by measuring the areas under peaks I and II, respectively, in MonoQ profiles determined by histone HI kinase activity assays. As shown in Fig. 3.11a, when NSC-19 cells were exposed to 2.5 mM [Ca 2 +]Q for 10 min, cytosolic P K C activity decreased to approximately 60% of control values. In addition, membrane associated P K C activity increased approximately five-fold, suggesting that cytosolic P K C had partially translocated to the membrane. Changing [Ca 2 +] 0 from 1.3 mM to 5 mM for 10 min resulted in an almost complete loss of cytosolic P K C activity with a corresponding five-fold increase in membrane-associated activity. Thus, P K C completely translocated to the membrane at a [Ca 2 +] ; of approximately 250 nM. Cells studied after 10 min incubation in 7.5 mM [Ca 2 +] 0 demonstrated levels of membrane-associated P K C activity which were comparable to cells exposed to 5 mM [Ca 2 +] 0. In addition, there was an increased level of PKC activity in the cytosolic fractions of cells exposed to 7.5 m M Ca 2 + which exceeded that detected under nonstimulated conditions. With 10 mM [Ca 2 +] 0, P K C activity was greater than that observed at 7.5 mM in both membrane and cytosol fractions. However, the increased activity observed at 10 mM [Ca 2 +] 0 was not accounted for by an increase in the amount of P K C protein (see Fig. 3.12), indicating that the specific activity of P K C was increased when [Ca 2 +] ; was elevated to approximately 350 nM. 57 Fig. 3.9 MonoQ chromatography of cell extracts MonoQ fast protein liquid chromatography (fplc) profile of cytosolic (left panels) and Triton X -100-solubilized particulate (right panels) fractions of NSC-19 cells exposed for 10 min to (A) 1.3 mM, (B) 10 mM, or (C) 50 mM Ca2+-containing HBSS. Cytosolic and particulate protein (0.8 mg) were separately loaded onto a 1-ml MonoQ (Pharmacia) column and eluted with a 10-ml linear gradient of 0 to 0.8 M NaCl in MonoQ buffer (dashed lines) as described in Materials and Methods. The 0.25-ml column fractions were assayed for phosphorylating activity toward histone HI in the presence of CaCl 2 , PS, and DO (closed symbols) or in the absence of these PKC activators (open symbols). Each profile is representative of data obtained in at least three separate experiments. 58 Cytosol Fractions A [Ca 2 4 1 0 = 1.3 mM Membrane Fractions 400 c o 03 b ~ 3001 o E l e 200H S | g l 100-1 2 co s 0.8 200-T — • — r 10 20 30 40 50 MonoQ fraction number 0.0 0-4* / 4* 4* 4* / * 4* 4* / 4* 4* / Peak II 4* 4 4* — r -\ - i — i — i — | — i — h0.6 0.8 0.0 10 20 30 40 50 MonoQ fraction number B [ C a 2 + | 0 = 10.0 mM 2000- 0.8 2000-r-10 20 30 40 50 MonoQ fraction number 0.6 1500-0.4 1000-h0.2 500-0.0 0 Peak 1 0.8 - i — | — i — | — . — | — i -10 20 30 40 50 MonoQ fraction number h0.6 0.4 cr 1-0.2 " 0.0 C [Ca^+lo = 50.0 mM 800-c o 5 £ Cu c oo . 3 5 S CU'— 53 g <u Cu c w o co s 0-Peak II / 9 / 4* m 4* • 4* * j ' a / a Peak 1 / f 4* I 4* | / I 3 \ \ \ \ • 0.8 400 0.0 0 10 20 30 40 50 MonoQ fraction number 59 l — 1 — I — ' — I — ~ 10 20 30 40 50 MonoQ fraction number Fig. 3.10 Western blot analysis of MonoQ peak I and II fractions Western irnmunoblot of peaks I and II from MonoQ profiles obtained from extracts of cells incubated in (A) 1.3 mM [Ca 2 +] 0, or (B) 50 mM [Ca 2 +]G for 10 min. A 200 ul portion of MonoQ peak tube fractions was loaded per lane and transferred to nitrocellulose after SDS-PAGE. Immunoblotting was performed with anti-Pan PKC polyclonal antibodies as described in Materials and Methods. A mixture of native a-, P-, and y-PKC isoforms, purified to near homogeneity from rat brain, was used as positive control (not shown). PKC-Soluble fractions from 1.3 mM [ C a 2 + ] 0 1-97 Mono Q fraction number (Peak I) Soluble fractions from 50 mM [ C a ^ + ] 0 PKC PKM - 9 7 - 4 9 L OJ 00 CO op c6 co co CO I uo CO co op co Mono Q fraction number (Peak II) 60 Fig. 3.11 Effect of [Ca 2 + ] 0 on activation of P K C and P K M NSC-19 cells were exposed to buffers with different [Ca 2 + ] 0 for 10 min prior to harvesting. Extracts containing 0.8-mg protein were loaded onto a 1-ml MonoQ column and 0.25-ml fractions were collected as described in Materials and Methods. Areas under the histone HI phosphorylating MonoQ profiles were determined and plotted as (A) P K C activity: area under peak I obtained in the presence of Ca 2 + , PS, and DO (see • , Fig. 3.9) minus area under same peak obtained in the absence of P K C activators (see • , Fig. 3.9); and (B) P K M activity: area under peak II in the absence of P K C activators (see 0 3 Fig. 3.9). Values are the means ± S.D. of at least three independent experiments for each [Ca 2 + ] 0 . In several groups, S.D.'s were too small to be distinguished from the means. 2500 2000 -J > ai Q 1 1 1500-| S 1000-*-H O x a ai & § 500-*-» X A PKC (Peak I) U T -r- ! TT I T I 2500 2000 H 1500 J 1000-^ 500-^ 1.3 2.5 5 7.5 10 25 [Ca2+]0 (mM) 50 1.3 2.5 5 7.5 10 25 50 [Ca 2 +] 0 (mM) E2 cytosohc MonoQ fractions membrane MonoQ fractions 61 At a [Ca 2 +] 0 of 25 mM, cytosolic P K C activity decreased by approximately 80%, and membrane-associated activity increased one- to two-fold compared to control. Exposure to buffer containing 50 mM Ca 2 + , resulted in decreased P K C activity in both cytosol and membrane fractions to approximately 12% and 36% of control values, respectively. These levels of P K C activity were substantially lower than those observed at 10 mM [Ca 2 +] 0. The decrease in P K C activity suggests that P K C was degraded. Consistent with this mechanism was the observation of a large increase in P K M activity (Fig. 3.1 lb) at 25 and 50 mM [Ca 2 +] 0 in both cytosol and membrane fractions compared to control. Additionally, immunoreactive P K M protein levels increased at 25 mM [Ca 2 +] 0 (Fig. 3.12b). The level of P K M activity (Fig. 3.1 lb) did not change significantly when [Ca 2 +]Q was changed from 1.3 mM to 2.5, 5, or 10 mM for 10 min. At 25 mM and 50 mM [Ca 2 +] 0, P K M activity in the cytosolic fractions for both groups increased seven- to eight-fold. Similarly, membrane-associated P K M activity increased nine- to eleven-fold over controls. The decrease in P K C activity and corresponding increase in P K M activity at 25 and 50 mM supports the possibility that P K C was proteolytically cleaved to P K M , possibly through the actions of calpains (Kishimoto et al, 1989; Melloni et al, 1985; Savart et al, 1992), which are believed to act preferentially on activated PKC. Data supporting this mechanism of P K C activation is shown in Fig. 3.12b. Incubation of cells to 2.5, 5 or 10 mM [Ca 2 +]G did not reveal any changes in immunoreactive P K M protein levels over control. However, in extracts from cells exposed for 10 min to 25 mM [Ca 2 +] 0, P K M levels were much higher than in control extracts, indicating that P K M was generated from PKC. 62 Fig. 3.12 Western blots of crude extracts fromNSC-19 cells (A) Western immunoblots of crude cytosolic and membrane extracts from NSC-19 cells exposed to buffers containing different [Ca 2 +] 0 for 10 min. Aliquots containing 150 ug of protein were loaded onto each well and transferred to nitrocellulose following SDS-PAGE. Anti-PKC a polyclonal antibody was used to determine the amount of P K C in each of the cell extracts. The [Ca 2 +] 0 at which cells were incubated is shown above the lanes. A mixture of native a-, P-, and y-PKC was used as positive control (lane 1). Visualization was performed by E C L and lane densities were estimated as described Materials and Methods. (B) Immunoblot of crude cytosolic extracts from NSC-19 exposed to different [Ca 2 +]Q. Membrane was probed for immunoreactivity to P K M using anti-PKC a polyclonal antibody. The bottom panel shows the densities of the bands of immunoreactive soluble (•) and particulate ( • ) P K C and soluble P K M (•)from at least three experiments (mean areas ± S.D.). 63 Soluble Extracts PKC J [ C a 2 + ] 0 ( m M ) PKC 1 -. _ 1 . •••<*>•<•<•>•• r.. — 95 — 69 Particulate Extracts (+) 1.3 2.5 5 10 25 50 ^ [ C a 2 + ] 0 ( m M ) VW^WTC"** --Vrf&iti&tfii&k* >'3&&£fi£fi&at- -'A&?$$i5$&ft'-~' / . — 95 — 69 8 Soluble Extracts 1.3 2.5 5 PKM [Ca 2 +] 0 (mM) 100 6 4 Chapter 4 Discussion Measurements of [Ca2+]i of NSC-19 cells were obtained using microspectrofluorometry with fura-2. Determination of [Ca 2 +] ; by this method permits accurate measurement of the free [Ca 2 +] ; in the cytoplasic compartment while allowing the determination of some of the temporal features of the Ca 2 + responses to changes in [Ca 2 +]O J metabolic inhibition, and other stimuli in NSC-19 cells. In contrast, other methods such as flux studies employing radioactive Ca 2 + ( 4 5Ca 2 +) do not distinguish between cytosolic Ca 2 + and Ca 2 + in internal reservoirs or buffers. Microscopic imaging of single cells, rather than a spectrophotometry analysis of cell suspensions, permitted the study of cells whose initial [Ca 2 +] ; levels were stable for at least 3 min before the start of recording. Cells which did not exhibit uniform dye loading, or whose initial basal [Ca 2 +] ; were not stable, were not counted. The use of NSC-19 also allowed us to study the Ca 2 + responses to metabolic insults in a homogenous cell type while excluding the effects of non-neuronal cells. Furthermore, the absence of an appreciable Ca 2 + response to glutamate in NSC-19 presumably simplifies the analysis of effects due to metabolic inhibition compared to experiments with primary neuron cultures. 4.1 Calibration of imaging system The apparent K d and P value determined for fura-2 in our system was similar to values reported elsewhere for imaging cells with a microscope (Uto et al, 1991). The signal-to noise ratio was increased by (i) using a microscope objective which has been reported to transmit 340 nm and 380 nm light with approximately equal intensities (Moore et al, 1990), (ii) recording multiple video frames which were sequentially added to memory and averaged before obtaining 65 ratios, and (iii) employing the use of neutral density filters which minimize the effects of photobleaching of the dye. The error in [Ca2 +]; was also minimized by subtracting the background current of the camera and the background fluorescence in the optical path from the images on a pixel-by-pixel basis. Finally, a fluorescence intensity threshold was set before obtaining the ratio in order to exclude non-cellular regions from affecting the ratio images. The in vitro calibraton curve obtained using a series of buffers with known Ca 2 + concentration was similar to in vitro and in situ curves reported elsewhere using a similar imaging system (Williams et al, 1985) and demonstrates that the ratio exhibited a steep (ten-fold) Ca 2 + dependence at the physiological range of [Ca 2 +] ; values (ratio of approximately 0.3 to 3 for [Ca2 +]; ranging from 0.05 to 2 uM). Although there are many pitfalls in using fura-2 as a Ca 2 + indicator, our system demonstrated reasonable initial [Ca2+]i values, and excluded non-cytoplasmic fura-2. In addition, the mean [Ca2+]j values determined under a variety of conditions demonstrated consistency and reproducibility. 4.2 [ C a 2 l in NSC-19 cells NSC-19 cells had resting [Ca 2 +] ; values which followed a normal distribution, with a mean value similar to previously published [Ca2 +]; in rat hippocampal neurons (Glaum et al, 1990; Dubinsky and Rothman, 1991; Tymianski et al, 1993a). In Ca2+-free media, the mean [Ca 2 +] ; in nonstimulated cells was significantly lower than with external Ca 2 + present, indicating that [Ca 2 +] ; was dependent on extracellular Ca 2 + . A similar dependence of [Ca2 +]; on [Ca 2 +] 0 has been observed previously for mammalian lymphocytes and thymocytes, where the addition of Ca 2 + to cells in Ca2+-free solutions resulted in a rise in [Ca 2 +] ; to steady state values over 3 min. Return of 66 thymocytes to a nominally Ca2+-ffee solution produced a rapid fall in [Ca 2 +] } (Tsien et al, 1982). The dependence of [Ca 2 +] ; on [Ca 2 +] 0 has also been reported for keratinocytes (Kruszewski et al, 1991) , rat hippocampal neurons (Tymianski etal, 1993a), and cultured rat cortical neurons (Villalba et al, 1994), and is also evident in the responses of the pancreatic cell line AR4-2J to substance P, bombesin or muscarinic receptor stimulation (Bird et al, 1992). The variation of [Ca 2 +] ; with [Ca 2 +] 0 indicates the operation of transmembrane Ca 2 + flux, either by channels or transporters. There was no evidence that transmembrane flux was mediated by EAA-gated channels or depolarization-dependent Ca 2 + channels, as Ca 2 + responses were not detected and blockade with application of known antagonists did not alter the change of [Ca 2 +] ; with [Ca 2 +] 0. However, it is possible that under the conditions studied, voltage-dependent Ca 2 + ion channels may be inactivated, as has been reported for a different NSC clone (NSC-34, Cashman et al, 1992) studied under similar initial conditions. In addition, blockade of "Ca2+-leak channels" by 2 mM L a 3 + (Demirel et al, 1993) did not alter the changes in [Ca2 +]; observed in the absence of L a 3 + , indicating that these channels, if present, are not involved in producing the response. Variations in [Ca2 +]; in NSC-19 cells with changes in [Ca 2 +] 0 are likely due to the actions of the Ca 2 +-ATPase (Mg2+-dependent) pump (Di Polo and Beauge, 1979) and of the Na7Ca 2 + exchanger (Blaustein et al, 1991). Both transporters appear to function in the maintenance of steady state [Ca2+]j levels in NSC-19 cells as inhibition of either of the two transporters resulted in increased [Ca2+]j levels in Ca2+-containing media. The Na7Ca 2 + exchanger may play a role in increasing [Ca 2 +] ; as addition of extracellular Ca 2 + in Na+-ff ee media decreased the rate of rise of [Ca 2 +] ;. Additionally, when [Ca 2 +] 0 was decreased to zero in Na+-free media, the rate of Ca2+efflux was lower and the resultant [Ca 2 +] ; level was greater than control. L i + did not alter the level of 67 [Ca 2 +] ; obtained in 1.3 mM Ca2+-containing media. Since cells were able to maintain initial basal [Ca2+]i levels in Li+-containing media, the Na7Ca 2 + exchanger in NSC-19 cells is likely not completely inhibited by L i + . Incomplete inhibition of the Na7Ca 2 + exchange system by L i + has also been noted for other cell types (Wacholz et al, 1993). The maximum level of [Ca 2 +] ; achieved with inhibition of the Ca 2 +-ATPase or the Na7Ca 2 + exchanger was 300 nM. This indicates that there likely exist other mechanisms for Ca 2 + buffering, sequestering, or efflux in NSC-19 cells. These processes, which were not investigated further, may not be active at [Ca 2 +] ; below about 300 nM. When NSC-19 cells were exposed to buffers containing 2.5 mM, 5 mM, or 10 mM Ca 2 + , [Ca2+]j increased steadily to levels which were maintained for at least 10 min. In contrast, changing [Ca 2 +] 0 from 1.3 mM to 25 or 50 mM resulted in large, transient increases in [Ca 2 +] ; which then returned to steady state values that approximated those obtained with 5 mM and 2.5 mM [Ca 2 +] 0, respectively. These steady state [Ca 2 +] ; values were lower than steady state [Ca2+]j seen with 5 and 10 mM [Ca 2 +]G. It follows that exposure of cells to 25 mM and 50 mM [Ca 2 +] 0 likely resulted in the activation of Ca 2 + extrusion or buffering processes which were not activated when the cells were exposed to lower [Ca 2 +] 0. This is consistent with observations made for [Ca 2 +] ; levels with inhibition of the Ca 2 + - ATPase and Na7Ca 2 + exchanger. The mechanisms of the Ca 2 + extrusion or buffering seen with exposure to 25 and 50 mM [Ca 2 +] 0 were not determined in this study. The large transient Ca 2 + responses observed following exposure to 25 and 50 m M Ca 2 + , may also have resulted in the activation of some Ca2+-dependent processes which require high levels of [Ca 2 +] ; for activation and were not seen with exposure to lower [Ca 2 +] 0. For instance, increased [Ca2+]i has been shown to activate calmodulin, a Ca2+-binding protein which 68 can increase the affinity of the plasma membrane Ca 2 +-ATPase for C a 2 + by twenty- to thirty-fold (Miller, 1991). The Ca 2 +-ATPase can also be activated by phosphorylation or mild proteolytic digestion (Miller, 1991), suggesting that activation of Ca2+-dependent kinases or proteases by raised [Ca2 +]j might increase the efficiency of the transporter. It has been difficult to precisely define the nature of the C a 2 + changes since the mechanism by which alterations in [Ca 2 +]i occurred in response to changes in [Ca 2 + ] 0 could not be blocked by the addition of V D C C antagonists or L a 3 + , and could not be attributed entirely to the actions of the plasma membrane Ca 2 +-ATPase or Na + /Ca 2 + exchanger. Since the [Ca 2 + ] ; measured by fura-2 does not indicate the route of C a 2 + entry, flux studies with 4 5 C a 2 + or electrophysiological recordings may be required to define the mechanism of C a 2 + entry in these cells. The differences in the kinetics between C a 2 + influx and efflux when [Ca 2 + ] 0 is changed further suggests that C a 2 + entry does not occur only by a C a 2 + leak mechanism. Although not investigated, it is possible that elevation of [Ca 2 + ] 0 resulted in the activation of capacitive C a 2 + entry or of Ca2+release-activated C a 2 + (CRAC) channels. Recently, Zweifach and Lewis (1995) demonstrated the presence of inward C a 2 + currents in Jurkat cells following elevation of [Ca 2 + ] 0 to 22 mM. These C R A C currents were partially dependent upon the level of depletion of intracellular stores, and inactivate with increases in the rate of C a 2 + influx (Zweifach and Lewis 1995) or PKC activation (Parekh and Penner, 1995). Observations of C a 2 + influx in the present study, both sustained and transient, with increased [Ca 2 + ] 0 in NSC-19 cells are consistent with the presence of C R A C channels. 4.3 Metabolic inhibition of NSC-19 cells Exposure of NSC-19 cells to Amytal (5 mM) did not result in a significant change in ATP 69 concentration. With CCCP, ATP was transiently reduced by 33%, which is similar to the effect of CCCP exposure (10-100 uM) on cellular ATP concentrations in hamster kidney cells (30-40 % of control; Burkhardt and Argon, 1989) and opossum kidney cells (20-30 % of control; L i et al, 1993). It is possible that significant reductions in ATP concentration might have been seen had 2-deoxy-D-glucose, a non-utilizable analog of glucose, been added to the media at the time of Amytal exposure. Cultured cells, especially cell lines, appear to be fairly resistant to metabolic inhibition due to a shift to glycolytic metabolism with greater glycogen stores (Freshney, 1987). Thus, the addition of a glycolytic inhibitor to this cell line may decrease ATP concentrations in response to metabolic inhibition by a greater amount than was observed here. Both Amytal and CCCP produced statistically significant increases of [Ca 2 + ] ; from resting levels, both in the presence and absence of external Ca 2 + . In response to Amytal, transient increases in [Ca 2 +] ; were obtained, whereas CCCP treatment resulted in elevated [Ca 2 + ] ; which did not recover over at least 30 min. The mean [Ca2 +]j responses produced by CCCP or Amytal were reduced by 30-50% when the experiments were repeated in Ca2+-free solutions indicating that the rise in [Ca 2 + ] ; was due both to internal release of C a 2 + as well as to C a 2 + influx. CCCP caused cellular ATP concentrations to fall within 80 seconds after the application of the drugs, while the maximal increase in [Ca 2 + ] ; was seen between 40 and 100 seconds after the drugs were applied, suggesting a possible causal relation between the decrease of ATP concentration and the increase in [Ca 2 +] ;. It was clear, however, that the reduction in ATP concentration did not parallel the rise in [Ca 2 +] ;, as CCCP transiently decreased ATP concentrations by 33%, whereas it increased [Ca 2 +] ; for a prolonged period. CCCP has been observed to cause the release of C a 2 + from mitochondria (Miller, 1991), 70 and FCCP (carbonyl cyanide p-fluoromethoxyphenylhydrazone) has been used to measure the size of the mitochondrial C a 2 + pool (Fulceri et al, 1991; Villalba et al, 1994). Although we did not directly determine the site of CCCP-induced intracellular C a 2 + release in NSC-19, mitochondria appear to be the likely source. The specific mechanism underlying the release of internal C a 2 + in response to Amytal was not investigated further but, like CCCP, was possibly due to release of sequestered [Ca 2 + ] ; in mitochondria (Fulceri etal, 1991). CCCP-evoked [Ca 2 +]i showed no return to baseline and application of ruthenium red made little difference, suggesting that mitochondrial re-uptake of [Ca 2 +]i likely did not occur after exposure to the protonophore. Ruthenium red pretreatment reduced the amplitude of the [Ca 2 + ] ; response to CCCP, suggesting that ruthenium red may partially prevent C a 2 + release from other internal stores, possibly by blocking ryanodine receptors (Gunther and Pfeiffer, 1990; Kapaus et al, 1990; Meissner, 1986). Application of ruthenium red alone did not cause any change in [ C a 2 ^ , indicating that the CCCP-sensitive internal C a 2 + store is insensitive to ruthenium red in NSC-19 cells. Stores which are sensitive or insensitive to release by ruthenium red have been described elsewhere (Rigoni and Deana, 1986; Fiskum and Cockrell, 1985). The results obtained on the effects of mitochondrial inhibitors in the present experiments are not unique, as Tsien et al (1982) have observed a two-fold increase in [Ca2 +]j after exposure of porcine lymphocytes to FCCP. The [Ca2 +]j response was considerably attenuated when cells were studied in Ca2+-free media, suggesting that either C a 2 + influx was eliminated or that C a 2 + stores were depleted (also see Villalba et al, 1994). In contrast, FCCP produced no measurable rise in [Ca 2 +]; in mouse thymocytes, indicating that these cells have no immediately releasable mitochondrial C a 2 + stores. In porcine lymphocytes, [Ca2 +]j returned to baseline values following 71 FCCP, probably due to the extrusion of Ca by the Ca pump, which can maintain reasonable levels of ATP by glycolytic activity (Tsien et al, 1982). In opossum kidney cells, CCCP produced a rapid [Ca 2 +]; response to over 500 nM within 30 s and returned to baseline by 200 s (Li etal, 1993). Although cells did not recover to baseline [Ca 2 + ] ; values, this did not affect cell viability for up to 48 h, as determined by fluorescent vital probes. The lack of cell death observed in these cultures is consistent with a previous study of metabolic inhibition in rat hippocampal neurons caused by 3 m M sodium cyanide (NaCN) (Dubinsky and Rothman, 1991). NaCN exposure resulted in micromolar levels of [Ca 2 +] ;. However, even after a 30 min exposure to NaCN, cell death in treated cultures assessed at 18 h was not different from controls. As with the resistance to decreased ATP concentrations, NSC-19 cells may be resistant to CCCP-induced cell death due to enhanced glycolytic metabolism. Although not determined, the addition of a glycolytic inhibitor may increase NSC-19 sensitivity to CCCP. In the neuroblastoma cell line SK-N-SH, addition of 2-deoxy-D-glucose was necessary for the observation of cell death induced by 2.5 mM NaCN (Johnson et al, 1994). The observation that 10 mM glutamate caused significant cell death may be accounted for by the actions of glutamate on receptor-operated channels which are not Ca2+-permeable or on metabotropic glutamate receptors. However, this was not investigated. In some cells the release of C a 2 + from mitochondria by CCCP appears to require an initial loading of the mitochondria with Ca 2 + , as will normally occur during a series of action potentials in neuronal cells (Miller, 1991). For instance, in neonatal rat dorsal root ganglion neurons, superfusion of CCCP (10 uM) produces virtually no change in resting [Ca 2 +] i 3 whereas application of the protonophore during a depolarization-induced [Ca 2 + ] ; transient will prolong the 72 duration of the transient (Thayer and Miller, 1990). However, in other cell types, the mitochondria appear to contain enough C a 2 + at rest to significantly increase [Ca 2 + ] ; with CCCP (Fulceri et al, 1991) or FCCP (Villalba et al, 1994), and this appears to be the case for NSC-19. A portion of the increase in [Ca 2 + ] ; which develops following CCCP exposure represents transmembrane influx, as this component disappears in Ca2+-free media (Villalba et al, 1994). The mechanism underlying this C a 2 + influx is unclear and does not appear to be due to either voltage-dependent or EAA-gated C a 2 + channels. Although increases in [Ca 2 + ] ; due to opening of voltage-dependent C a 2 + channels have been detected in glomus cells from the newborn rabbit carotid body in response to anoxia and cyanide (Sato et al, 1991), depolarization with KC1 did not result in increased [Ca 2 +]; in the present experiments. Ligand-gated C a 2 + channels have also been proposed to have an important role in the increase in [Ca2*]; seen in "chemical hypoxia" in hippocampal neurons in culture (Dubinsky and Rothman, 1991), yet glutamate exposure induced no appreciable change in [Ca 2 + ] ; in NSC-19 cells. Although not directly evaluated, the plasma membrane Na + /Ca 2 + exchanger can function in the reverse mode, such that C a 2 + influx and N a + efflux might occur. Operation in this manner is enhanced by increased intracellular N a + secondary to membrane depolarization, which is usually present during chemical hypoxia (Stys et al, 1991). The present experiments have demonstrated the presence of Na + /Ca 2 + exchanger in NSC-19 cells. However, we have been unable to confirm the involvement of this transporter in metabolic inhibition. 4.4 PKC activity with changes in [Ca 2 + ] 0 The relatively stable levels of [Ca 2 +] ; in response to changes in [Ca 2 + ] 0 were used as a 73 means of evaluating the relationship between P K C and [Ca 2 +] ;. The decrease in cytosolic P K C activity and corresponding increase in P K C activity in the membrane fraction with changes in [Ca 2 + ] 0 from 1.3 mM to 2.5 or 5 mM suggests that PKC translocates to the membrane, and presumably is activated, at relatively low levels of [Ca 2 +] i 3 ranging from 150 to 250 nM. The translocation of P K C from the cytosol to the membrane is conventionally accepted as the primary mechanism for the activation of P K C (for review see Huang, 1989). It should be noted, however, that translocation of PKC activity to the membrane does not necessarily reflect its activation. To our knowledge, there are no other reports which define a threshold level of [Ca 2 + ] ; in situ for P K C activation. Studies of P K C activation by C a 2 + in vitro have reported that P K C can be half maximally activated (A 1 / 2 ) by 200 to 600 nM C a 2 + in phospholipid/detergent mixed micelles assays (Huang, 1990) and that P K C binds to isolated neutrophil or platelet membranes at assay C a 2 + concentrations of 2 to 5 u M (Melloni et al, 1985). It is possible that the lower threshold value for P K C translocation observed in the intact NSC-19 cells is due to easier accessibility of the kinase to the membrane within the closed system. Although translocation of P K C activity at 150 to 250 nM was observed, this was not confirmed in the immunoblot analysis. This is likely due to a lack of sensitivity of the antibody in detecting small changes in P K C protein levels. The antibodies may not be detecting those isoforms of PKC whose activities are affected by the changes in [Ca 2 +] i 5 or they may be detecting other isoforms or proteins whose signals are masking the changes. In this regard, we have tried antibodies from several sources which claim that the antibodies are specific for particular P K C isoforms, but which have failed to demonstrate isotype specificity (results were not shown) when checked against pure P K C isoforms. With incubation at [Ca 2 + ] Q values of 7.5 mM and 10 mM corresponding to a [Ca2 +]j of 74 about 350 nM, P K C activity increased two-fold and eleven-fold in cytosol and membrane fractions, respectively. P K C protein was not increased, however, suggesting that the specific activity of P K C increases at approximately this [Ca 2 +] ;. In the mast cell/megakaryocyte cell line R6-XE.4 activated by 15 min exposure to IL-3, a six-fold increase in particulate-derived P K C activity was observed with no corresponding reduction in cytosolic P K C activity (Pelech et al, 1990b). Two- to four-fold increases in both cytosolic and particulate-derived P K C activities have also been observed for rabbit platelets exposed to platelet-activating factor (PAF) for 1 to 20 min. In contrast, a 5 min exposure of platelets tol2-0-tetradecanoylphorbol 13-acetate (TP A), resulted in the translocation of more than 95% of cytosolic P K C to the membrane (Pelech et al, 1990a; Salari et al, 1990; Pelech et al, 1991), indicating that both translocation-dependent and translocation-independent activation of P K C can occur in the same cell system under different conditions. The [Ca 2 +] ;, however, was not investigated in these experiments. Pelech et al. (1991) suggested that the apparent translocation-independent increases in P K C activity seen following PAF stimulation of platelets may be due to the phosphorylation of P K C either through autophosphorylation or by another kinase. Our observations at 10 mM [Ca 2 + ] 0 are consistent with covalent modification of P K C as immunoblot analysis of crude extracts did not reveal increased PKC protein in either cytosolic or membrane fractions compared to controls, as measured using anti-PKC antibodies obtained from different sources. As well, the increased P K C activity was stable to MonoQ chromatographic separation of cell extracts. It is interesting to note that Begum et al. (1991) have found that a sustained increase in [Ca 2 + ] ; to approximately 300 n M in rat adipocytes by membrane depolarization resulted in an inhibition of cytosolic and particulate protein phosphatase (PPTase) activity which corresponded to increased phosphorylation states of 75 two cellular proteins. It is thus possible that at a [Ca ] 0 of 10 mM ([Ca ] ; - 350 nM), dephosphorylation of P K C is also inhibited. The inhibition of dephosphorylation of P K C and/or the stimulation of PKC phosphorylation, presumably would have the effect of generating high levels of P K C activity in the absence of translocation. We cannot eliminate the possibility that the increase in cytosolic P K C activity at 10 m M C a 2 + was due to the disruption of loosely-bound membrane P K C and its release into the cytosol during sample preparation. Bazzi and Nelsestuen (1988) have demonstrated the presence of a pool of membrane-inserted P K C in isolated membranes which can be distinguished from a second loosely-bound membrane-associated pool of PKC. By the addition of Ca2+-chelators (eg. EGTA) to the homogenizing buffer, the latter is released into the cytosol fraction during cell disruption. However, if the increase in cytosolic P K C activity at 10 mM [Ca 2 + ] 0 in our study was due to the dissociation of loosely-bound P K C in the membrane by EGTA, then we would have expected to see an increase in the amount of P K C protein in the 10 mM sample. Lu et al. (1994) have reported an IL-2 stimulated activation of P K C which they claim to be translocation-independent and due to the activation of a pool of pre-existing, loosely-bound, membrane-associated PKC. It is possible that the increased P K C activity observed with 7.5 to 10 mM [Ca 2 + ] 0 is due to the activation, other than by phosphorylation, of a previously inactive pool of P K C which requires a threshold level of [Ca 2 + ] ; (-350 nM) for its activation, and that neither translocation nor covalent modification of P K C is the major mechanism resulting in increased P K C activity at this [Ca 2 +]j range. Whatever the mechanism of P K C activation at 10 mM [Ca 2 + ] 0 , the increases in both cytosolic and membrane-associated P K C activity cannot be explained by translocation alone, and likely involves the activation of pre-existing membrane-associated PKC. 76 At 25 mM [Ca 2 + ] 0 , the observed decrease in cytosolic P K C activity and slight increase in particulate P K C activity compared to control cultures may be indicative of a translocation of P K C from the cytosol to the membrane. This mechanism of P K C activation may be similar to that seen at 5 mM [Ca 2 + ] G as steady state [Ca 2 + ] ; values for these two groups were similar. However, the total P K C activity after exposure to 25 mM C a 2 + was less than that observed at control levels and considerably less than that observed at [Ca 2 + ] 0 of 10 mM. The decrease in total P K C activity at 25 mM and concomitant increases in cytosolic and particulate P K M activity suggest that P K C was proteolytically cleaved to generate P K M , possibly through the actions of calpain (Melloni et al, 1985; Kishimoto et al, 1989; Savart et al, 1992). Calpain activation is known to be C a 2 + -dependent with the two major isoforms of calpain differing in the threshold value of [Ca 2 + ] ; required for activation. Analysis of [Ca 2 +] ; at 25 and 50 mM revealed a biphasic response. Immediately after exposure of cells to 25 or 50 mM Ca 2 + , a transient increase in [Ca 2 + ] ; was seen followed by return to much lower steady state [Ca 2 + ] ; values which was comparable in magnitude to [Ca 2 +]; obtained with lower [Ca 2 + ] 0 . Since P K M activity in cultures incubated with [Ca 2 + ] 0 lower than 25 mM was substantially less than that seen at 25 and 50 mM, the presence of increased P K M activity in the latter two [Ca 2 + ] 0 is likely related to the C a 2 + transient rather than to the steady state level of [Ca2 +]j. We did not determine the magnitude of the C a 2 + transient necessary for P K M activation. However, since a [Ca 2 +]j value of close to 400 nM did not generate much of an increase in P K M activity whereas a C a 2 + transient to 600 n M did produce a considerable increase in P K M activity, we expect that PKC activation by cleavage to P K M must occur at approximately 500 - 600 nM. The activation of calpains due to the transient [Ca2 +]j increase may not have occurred with the 77 slow rise to steady state [Ca \ seen with the lower [Ca ] 0 treatments. These results are in agreement with those of Melloni et al. (1986), who showed that in neutrophils, phorbol ester-stimulated P K C was converted to P K M at micromolar concentrations of C a 2 + . Calpain translocation to the membrane has been shown to occur at the same C a 2 + concentrations as translocation of P K C in isolated neutrophil membranes (Melloni et al, 1985). Studies with platelets have shown that exposure to TPA (Tapely and Murray, 1984), or to the Ca2+-ionophore A23187 (Hoshijima et al, 1986) results in generation of P K M which is inhibitable by the simultaneous application of a calpain inhibitor. Although [Ca 2 + ] ; was not measured, P K M generation has also been reported for a neuroblastoma cell line following exposure to A23187 (Shea et al, 1994), with a time course consistent with the time course of P K M generation in our experiments. Thus in NSC-19 cells, the conversion of P K C to the constitutively active catalytic fragment P K M at [Ca2 +]j greater than 600 nM may represent a third mechanism for kinase activation. Increases in P K M activity may lead to phosphorylation of cytosolic and nuclear proteins which might otherwise not be accessible to PKC. In turn, this could result in significant changes in the activities of metabolic pathways or in gene expression. 78 Chapter 5 Conclusions The present observations indicate that exposure of NSC-19 cells to Amytal, or metabolic inhibition with CCCP is associated with elevations in [Ca2+];. This [Ca2+]j derives from both the release of internal stores and transmembrane influx. Raised [Ca2 +]; associated with metabolic inhibition or other stimuli can be much more sustained than the period of exposure to the stimulus and may not necessarily lead to decreased cell viability, at least over short periods of time (days). The observations with NSC-19 suggest that metabolic inhibition may result in elevated [Ca 2 +] ; in motoneurons and could potentially lead to cell dysfunction with time. This dysfunction could be mediated by prolonged activation of Ca2+-dependent processes such as the activation of PKC. The level and possibly the rate of change of [Ca 2 +] ; activates P K C in three distinct ways. At low levels of [Ca 2 +] ; above baseline, corresponding to approximately 150 nM in NSC-19, P K C activity translocates from the cytosol to the membrane, with the [Ca2 +]; value of approximately 250 to 300 nM producing maximal P K C translocation. At higher levels of [Ca 2 +] ; (approximately 350 nM), P K C also appears to be activated independently of translocation, possibly as a result of an increase in the specific activity of P K C by enhanced phosphorylation. At [Ca 2 +] ; levels around 600 nM, possibly in association with a rapidly changing Ca 2 + transient, P K M activity is prominent. We did not evaluate the time course of P K C or P K M persistence after activation, or the physiological consequences of PKC activation, nor did we evaluate the effects of metabolic inhibition with CCCP on P K C activation. Given that changes in [Ca 2 +] ; have been implicated as critical steps in neuronal death, it is possible that [Ca 2 +] ; may exert its neurotoxic effects through P K C activation. The present study makes it clear that the consequences of elevation of [Ca2 +]; can be very different depending on the 79 amplitude of [Ca 2 +] ; obtained, the rate of rise of [Ca 2 +] ;, and the presence of a Ca 2 + transient. EAA-induced elevations of [Ca 2 +] ; are typically rapid and associated with a prominent Ca 2 + transient, often followed by a normalization of [Ca 2 +] ; for a variable period of time prior to cell death (Dubinsky, 1993a; Tymianski et al, 1993a). This pattern of [Ca2 +]; change is similar to that seen in our experiments with exposure of cells to 25 and 50 mM [Ca 2 +] 0, and thus would suggest that EAA-induced [Ca 2 +] ; increases might be associated with P K M production. The generation of P K M , a constitutively active kinase, free from the regulatory constraints of PKC, could have major physiological consequences as this kinase would have access to the cytosol, and could phosphorylate a larger group of substrates than the membrane-bound P K C (Tapely and Murray, 1984; Nixon, 1989; Sauvage et al, 1991). This mechanism of P K C activation could lead to enhanced or prolonged phosphorylation of cytoskeletal components, possibly leading to a compromise in cell membrane integrity, and may have relevance to neuronal death following an increase in E A A release thought to be associated with ischemia (see Choi, 1990). Small increases in [Ca 2 +] i 3 which in the present study were not associated with Ca 2 + transients, may result in a pattern of kinase activation completely different from that seen with higher [Ca 2 +] ;. Observations that low amplitude, long duration increases in [Ca 2 +] ; following application of metabolic inhibitors to cells in culture do not result in significant cell death (Dubinsky and Rothman, 1991, Johnson et al, 1994) suggest that different cellular processes are activated compared to those which follow E A A application. It is possible that the different physiological consequences of E A A application and metabolic inhibition are due to differences in the nature of the Ca 2 + responses and [Ca 2 +] ; levels generated under the two conditions. [Ca 2 +] ; levels seen following metabolic inhibition in NSC-19 cells are similar to those observed with 80 exposure to [Ca 2 +] 0 of less than 25 mM and may be derived from release of internal stores, transmembrane influx, or a combination of sources. Also, the raised [Ca 2 +] ; associated with metabolic inhibition can be sustained much longer than the period of exposure to the inhibitor (eg. CCCP) and could potentially lead to motoneuron dysfunction if prolonged. It is possible that the mechanism of P K C activation following metabolic inhibition is similar to the activation observed with exposure of NSC-19 neurons to low [Ca 2 +] 0 (less than 25 mM). This mechanism of P K C activation appears to be distinct from that observed when NSC-19 cells are exposed to [Ca 2 +] 0 equal to or greater than 25 mM. 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