Ion Transport and Membrane Potential in CNS Myelinated Axons II. Effects of Metabolic Inhibition

Lisa Leppanen and Peter K. Stys

Loeb Research Institute, Ottawa Civic Hospital, University of Ottawa, Ottawa, Ontario K1Y 4E9, Canada

    ABSTRACT
Abstract
Introduction
Methods
Results
Discussion
References

Leppanen, Lisa and Peter K. Stys. Ion transport and membrane potential in CNS myelinated axons. II. Effects of metabolic inhibition. J. Neurophysiol. 78: 2095-2107, 1997. Compound resting membrane potential was recorded by the grease gap technique (37°C) during glycolytic inhibition and chemical anoxia in myelinated axons of rat optic nerve. The average potential recorded under control conditions (no inhibitors) was -47 ± 3 (SD) mV and was stable for 2-3 h. Zero glucose (replacement with sucrose) depolarized the nerve in a monotonic fashion to 55 ± 10% of control after 60 min. In contrast, glycolytic inhibition with deoxyglucose (10 mM, glucose omitted) or iodoacetate (1 mM) evoked a characteristic voltage trajectory consisting of four distinct phases. A distinct early hyperpolarizing response (phase 1) was followed by a rapid depolarization (phase 2). Phase 2 was interrupted by a second late hyperpolarizing response (phase 3), which led to an abrupt reduction in the rate of potential change, causing nerves to then depolarize gradually (phase 4) to 75 ± 9% and 55 ± 6% of control after 60 min, in deoxyglucose and iodoacetate, respectively. Pyruvate (10 mM) completely prevented iodoacetate-induced depolarization. Effects of glycolytic inhibitors were delayed by 20-30 min, possibly due to continued, temporary oxidative phosphorylation using alternate substrates through the tricarboxylic acid cycle. Chemical anoxia (CN- 2 mM) immediately depolarized nerves, and phase 1 was never observed. However a small inflection in the voltage trajectory was typical after approx 10 min. This was followed by a slow depolarization to 34 ± 4% of control resting potential after 60 min of CN-. Addition of ouabain (1 mM) to CN--treated nerves caused an additional depolarization, indicating a minor glycolytic contribution to the Na+-K+-ATPase, which is fueled preferentially by ATP derived from oxidative phosphorylation. Phases 1 and 3 during iodoacetate exposure were diminished under nominally zero Ca2+ conditions and abolished with the addition of the Ca2+ chelator ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA; 5 mM). Tetraethylammonium chloride (20 mM) also reduced phase 1 and eliminated phase 3. The inflection observed with CN- was eliminated during exposure to zero-Ca2+/EGTA. A Ca2+-activated K+ conductance may be responsible for the observed hyperpolarizing inflections. Block of Na+ channels with tetrodotoxin (TTX; 1 µM) or replacement of Na+ with the impermeant cation choline significantly reduced depolarization during glycolytic inhibition with iodoacetate or chemical anoxia. The potential-sparing effects of TTX were less than those of choline-substituted perfusate, suggesting additional, TTX-insensitive Na+ influx pathways in metabolically compromised axons. The local anesthetics, procaine (1 mM) and QX-314 (300 µM), had similar effects to TTX. Taken together, the rate and extent of depolarization of metabolically compromised axons is dependent on external Na+. The Ca2+-dependent hyperpolarizing phases and reduction in rate of depolarization at later times may reflect intrinsic mechanisms designed to limit axonal injury during anoxia/ischemia.

    INTRODUCTION
Abstract
Introduction
Methods
Results
Discussion
References

Mammalian CNS axons are susceptible to anoxic injury and sustain irreversible damage from an increase in [Ca2+]i (LoPachin and Stys 1995; Ransom et al. 1994; Stys et al. 1990; Waxman et al. 1991). Ca2+ homeostasis becomes disrupted as a result of energy depletion and the subsequent loss of ion gradients. Under normal physiological conditions, the low [Ca2+]i is maintained by two membrane proteins, the Na+/Ca2+ exchanger and Ca2+-ATPase (Blaustein 1988), as well as buffering by intracellular stores and Ca2+ binding proteins (Kostyuk and Verkhratsky 1994). The Na+/Ca2+ exchanger normally extrudes 1 Ca2+ in exchange for 3 Na+ by using the free energy available through the Na+ gradient (Steffensen and Stys 1996). Factors influencing the rate and direction of exchanger operation include the transmembrane Na+ gradient and membrane potential. The exchanger will mediate Ca2+ influx when the transmembrane Na+ gradient is reduced and/or the membrane depolarizes. The exchanger is sensitive to membrane potential because its ion transport is electrogenic, transferring one net charge per cycle (Rasgado-Flores and Blaustein 1987). For these reasons, given that the exchanger is the main pathway for anoxic Ca2+ overload in optic nerve axons (Stys et al. 1992b; Waxman et al. 1992), the rate and extent of depolarization during metabolic stress will influence the degree of exchanger-mediated Ca2+ entry and thus ultimate cellular injury. We therefore examined the ions and channels involved in mediating membrane depolarization during glycolytic inhibition and chemical anoxia in optic nerve. Preliminary results have been published in abstract form (Leppanen and Stys 1996).

    METHODS
Abstract
Introduction
Methods
Results
Discussion
References

Detailed methods and recording techniques are described in the companion paper (Leppanen and Stys 1997). The optic nerves were dissected from Long-Evans male rats (150-175 g) anesthetized with 80% CO2-20% O2 and decapitated. The recorded axonal compound resting membrane potential, Vg, was measured in vitro with the grease gap technique (Stys et al. 1993). The middle segment of one nerve was inserted into a slit silastic tube filled with petroleum jelly. One end was perfused at 37.0°C with oxygenated artificial cerebrospinal fluid (aCSF) or test solutions. The opposite end was depolarized using an isotonic K+ solution (NaCl replaced with equimolar KCl) containing 0.5 mM CaCl2. Solutions were gassed with 95% O2-5% CO2. The central nerve segmentwas cooled to improve recording stability. Vg was recorded with 3 M KCl/agar bridge electrodes. Junction potentials (typically <= 1-3 mV), which drifted linearly over time, were determined at the beginning and end of each experiment, and Vg was corrected by subtracting interpolated values of these potentials. The nerve studied immediately after the dissection was denoted nerve A. The second nerve (B) was placed in an oxygenated chamber (95%O2-5% CO2) containing aCSF at room temperature for later recording. Although the grease gap technique provided stable, long-term recordings, it is unable to discern between different axonal populations that may exhibit different responses to metabolic inhibition (Stys and LoPachin 1996). Our results therefore, represent a mean population response, biased in favor of larger axons (see companion paper Leppanen and Stys 1997). Because of the origin of the recorded potential, we cannot exclude the possibility that some drugs affected the short circuit factor and thus affected the recorded potentials. Finally, given the heterogeneous structure of myelinated axons, with the majority of the axolemma covered by tight myelin wraps, diffusion barriers into this region may have influenced the effects of some agents active at internodal loci.

Composition of aCSF and solutions of 2-deoxyglucose (DG; Sigma), KCN (Fisher Scientific), tetraethylammonium chloride (TEA; Sigma), zero-glucose, and zero-Na+/choline [choline Cl (BDH)] are listed in Table 1. NaCN (BDH), iodoacetate-Na+ salt (IAA; Sigma), pyruvate (Sigma), ouabain (Sigma), ethyleneglycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA;Sigma), procaine (Sigma), and QX-314 (a generous gift from Astra Pharma) were dissolved in aCSF. Tetrodotoxin (TTX; Sigma) was diluted from stock solution in distilled water.

 
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TABLE 1. Composition of solutions

Statistical differences were calculated using analysis of variance with Dunnett's test.

    RESULTS
Abstract
Introduction
Methods
Results
Discussion
References

Glycolytic inhibition

After nerve insertion in normal aCSF, Vg typically stabilized within 90 min and ranged from -40 to -55 mV [mean -47 ± 3 (SD) mV; Table 2]. We examined several methods of inducing glycolytic inhibition in the rat optic nerve (RON), a preparation that is essentially 100% myelinated (Foster et al. 1982). In Fig. 1A, the effect of zero-glucose (sucrose substitution) on resting membrane potential is shown. A delay of 28 ± 6 min preceded the onset of a monotonic, gradual depolarization proceeding at 1 ± 0.4 mV/min (Table 4), to 55 ± 10% of control after 60 min (Fig. 1A, arrow, Table 2). We compared ratios of potentials over time, normalized to the baseline reading obtained at time 0 (defined as 90 min after nerve insertion and stabilization). Using -80 mV as an estimate of true optic axon membrane potential (Stys et al. 1997), these ratios were used to estimate absolute membrane potentials over time (Tables 7 and 8 in DISCUSSION). In contrast to simple omission of glucose, substitution of glucose with DG [a competitive antagonist of the glycolytic enzyme hexokinase (Devlin 1992)], induced an initial hyperpolarizing response in five of seven nerves (Fig. 1B). This early hyperpolarization, termed phase 1, together with the delay before hyperpolarizing lasted for 38 ± 8 min after application of DG before nerves began to gradually depolarize (phase 2) at 2 ± 1 mV/min (Table 4). Again, in contrast to zero-glucose alone, the gradual depolarization was interrupted by a second late hyperpolarizing response in four of seven nerves studied (phase 3), finally terminating in a slow depolarization (phase 4) to 75 ± 9% of control after 60 min of application (Table 2). Because energy production still could occur from residual glucose in the extracellular space or from glycogen stores in astrocytes (Cataldo and Broadwell 1986), IAA (1 mM), a relatively specific irreversible inhibitor of glyceraldehyde 3-phosphate dehydrogenase (Sabri and Ochs 1971), was used to directly inhibit glycolytic metabolism. As with DG, four phases were observed (Fig. 1C): an early distinct hyperpolarizing response (phase 1, in 14 of 24 nerves) was followed by a rapid depolarization (phase 2) at a rate of 5 ± 2 mV/min (Table 4). Phase 2 began 20 ± 3 min after IAA application (Table 4), which was significantly faster than in zero-glucose/sucrose, 28 ± 6 min (P < 0.001), or in DG/zero-glucose, 38 ± 8 min (P < 0.01). As with DG, a second late hyperpolarizing response (phase 3) interrupted phase 2. Finally, a gradual depolarization after phase 3 (at a reduced rate in comparison to phase 2) reached 55 ± 6% of control potential after 60 min (Table 2). Addition of pyruvate (10 mM), a substrate used by the tricarboxylic acid cycle, completely prevented the IAA-induced changes (Fig. 1D, Table 2).

 
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TABLE 2. Effects of glycolytic block or chemical anoxia on optic nerve membrane potential


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FIG. 1. Effect of glycolytic inhibition on recorded (Vg) and calculated absolute (Vm) compound resting membrane potential in rat optic nerve. Vm was normalized to -80 mV for this and all subsequent figures. A: zero glucose/sucrose application evoked a gradual depolarization after a delay of approx 30 min. Ratio was calculated by dividing the potential at 60 min (right-arrow) by the time 0 potential. Deoxyglucose substituted zero-glucose (B) or iodoacetate (C) both elicited a characteristic response consisting of 4 distinct phases: an early hyperpolarizing phase, P-1, which was followed by a more rapid depolarization(P-2). P-2 was interrupted by a second late hyperpolarization, P-3, and the voltage trajectory then entered a final, more gradual depolarization at a lower rate (P-4). Final levels of depolarization were similar at the end of 2 h for iodoacetate and zero-glucose. Extent of depolarization was less for deoxyglucose than for iodoacetate or zero-glucose. D: pyruvate (10 mM) completely prevented iodoacetate-induced depolarization.

 
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TABLE 4. Time to onset and rate of rapid depolarization induced by glycolytic inhibition and chemical anoxia

 
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TABLE 7. Calculated absolute compound resting membrane potential

 
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TABLE 8. Calculated absolute compound resting membrane potential during metabolic inhibition and altered Na+ conductance

Chemical anoxia

Differences were observed between glycolytic block and chemical anoxia induced with 2 mM NaCN, an inhibitor of mitochondrial cytochrome oxidase aa3 (Albaum et al. 1946; Kauppinen and Nicholls 1986a; Tadic 1992). CN- resulted in a rapid depolarization (phase 2) at 4 ± 2 mV/min (Table 4) with minimal delay after application; in contrast to IAA and DG, the transient initial hyperpolarization (phase 1) was never seen (Fig. 2A). A small inflection (Fig. 2A, right-arrow), which was far less pronounced than with IAA, followed the rapid depolarizing response, leading to an abrupt slowing of the rate of depolarization (phase 4). Nerves depolarized to 34 ± 4% of control after 60 min (Table 2). To test the hypothesis that Na+,K+-ATPase was still partially active during CN- poisoning, ouabain (1 mM) was added to the perfusate after 90 min of CN-. An additional depolarization confirmed that residual pump activity was present during chemical anoxia (Fig. 2B). In contrast, no change in voltage trajectory was observed by adding ouabain to IAA-poisoned nerves(Fig. 2C).


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FIG. 2. Effect of chemical anoxia and ouabain on resting membrane potential in rat optic nerve. A: NaCN (2 mM) caused an immediate depolarizing response commonly interrupted by an inflection in the voltage trajectory (right-arrow), followed by a slower depolarization. Addition of ouabain (1 mM) produced an additional, although minor depolarization, in chemically anoxic nerves (B), but not in glycolytically inhibited nerves (C). This suggests that glycolysis is able to fuel the Na+,K+-ATPase to a minor extent during chemical anoxia. See Fig. 1 legend for definition of Vg and Vm.

Role of Ca2+ during glycolytic inhibition and chemical anoxia

The hyperpolarizing inflections observed with IAA and, to a far lesser extent, CN- were unexpected and may reflect interesting mechanisms of axonal response to energy failure. Ca2+-activated K+ conductances (KCa2+) have been implicated in other tissues (Leblond and Krnjevic 1989), therefore we investigated the role of Ca2+ on potential trajectories during IAA and CN- treatment. In Fig. 3A, simultaneous application of IAA and nominally zero Ca2+ caused an accelerated onset of depolarization, beginning within 5 ± 2 min, compared with 20 ± 3 min in IAA alone (Table 4). The depolarizing response, at a rate of 2 ± 0.3 mV/min (Table 4), was interrupted by a less prominent phase 3 (Fig. 3A, right-arrow). Resting potential depolarized to 65 ± 2% and44 ± 2% of control at 30 and 60 min (Table 3), respectively, significantly more than with IAA alone at these times(79 ± 7% and 55 ± 6%, P < 0.01 and P < 0.05, respectively). Pretreating the nerves with nominally zero Ca2+ for 60 min before IAA application caused phase 3 to become a brief response without a distinct hyperpolarization (Fig. 3B, arrow, Table 3) after the addition of IAA. A 60-min application of zero-Ca2+/EGTA (5 mM) elicited a small but rapid depolarization to 96 ± 1% of control (Fig. 3C, Table 6). As with nominally zero Ca2+ and IAA, the nerve depolarized quickly, beginning within 3 ± 1 min (Table 4), but distinct phases were not observed in the voltage trajectory. Exposure to zero-Ca2+/EGTA plus IAA resulted in a slower rate of depolarization (1 ± 0.1 mV/min) compared with IAA alone (5 ± 2 mV/min, P < 0.05, Table 4). Although zero-Ca2+/EGTA slowed the rate of depolarization because of a more rapid onset in Ca2+-depleted conditions, nerves depolarized to a greater degree at 30 and 60 min, 61 ± 3% and 43 ± 4%, respectively, versus IAA alone, 79 ± 7% and 55 ± 6%, (P < 0.0001 and P < 0.001, respectively) (Table 3). Similarly, the abrupt alteration in the depolarization rate after phase 3 was reduced progressively as the Ca2+ removal became more severe (compare Figs. 1C and 3, A-C), until the trajectory became monotonic under zero-Ca2+/EGTA conditions.


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FIG. 3. Effect of Ca2+-depleted perfusate on membrane potential during glycolytic inhibition with iodoacetate (IAA, 1 mM). A: exposure to iodoacetate and nominally zero Ca2+ simultaneously, accelerated the onset of depolarization, eliminated phase 1, and reduced the size of phase 3 (right-arrow; compare with Fig. 1C). B: pretreatment with zero Ca2+ caused phase 3 to become even less pronounced (right-arrow). C: zero-Ca2+/ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) solution alone elicited a small depolarization (right-arrow). Addition of the Ca2+ chelator eliminated all hyperpolarizing phases. D: tetraethylammonium chloride (TEA; 20 mM) alone depolarized nerves slightly (right-arrow) due to its broad-spectrum K+-channel-blocking properties (Hille 1992). TEA reduced the early hyperpolarizing response, P-1, and abolished the second late hyperpolarizing response, P-3. A Ca2+-dependent K+ conductance(s) is a possible mechanism for the early and late hyperpolarizing responses. See Fig. 1 legend for definition of Vg and Vm.

 
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TABLE 3. Role of Ca2+ during glycolytic inhibition and chemical anoxia

 
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TABLE 6. Effect of various experimental conditions on recorded compound resting membrane potential

TEA (20 mM), a broad-spectrum K+ channel antagonist (including some KCa2+) (Hille 1992) depolarized nerves to a surprisingly small extent when applied alone (Fig. 3D, Table 6). TEA exposure during IAA application reduced phase 1 and eliminated phase 3, with depolarization beginning within 17 ± 2 min (similar to IAA alone, 20 ± 3 min; Table 4) to 61 ± 4% of control after 60 min (similar to IAA alone, 55 ± 6%; Table 3). Although phase 3 was abolished, a distinct change in the rate of depolarization from phase 2 to phase 4 was still present.

During chemical anoxia and a nominally zero Ca2+ perfusate that was applied 60 min before CN-, nerves depolarized at a rate of 8 ± 2 mV/min (Table 4) to 30 ± 7% of control at 60 min (Table 3) with an inflection apparent (Fig. 4A). Similar to the voltage trajectory observed with IAA and zero-Ca2+/EGTA, addition of CN- after 60 min of zero-Ca2+/EGTA treatment evoked a depolarization at a rate of 9 ± 3 mV/min, without an inflection, to 27 ± 4% at 60 min (Fig. 4B, Table 3). For both nominally zero Ca2+ and zero-Ca2+/EGTA treatments, nerves depolarized at a significantly faster rate than with CN- alone (4 ± 2 mV/min, P < 0.05).


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FIG. 4. Effect of zero Ca2+ and zero Ca2+/EGTA on membrane potential during chemical anoxia. A: nominally zero Ca2+ significantly accelerated the rate of CN--induced depolarization (Table 4), but inflections, characteristic of induction of anoxia alone, were preserved. B: in contrast, addition of EGTA also caused accelerated depolarization with CN- but also abolished inflections in the voltage trajectory, suggesting the possible modulation of Ca2+-dependent conductances during chemical anoxia that were less pronounced than with glycolytic inhibition (compare with Figs. 1C and 3). See Fig. 1 legend for definition of Vg and Vm.

Role of Na+ during glycolytic inhibition and chemical anoxia

The role of transmembrane Na+ flux during metabolic inhibition was studied using a specific Na+ channel blocker and ion substitution experiments. In Fig. 5A, TTX (1 µM) alone elicited a hyperpolarizing response (Table 6), and with IAA addition, the extent of depolarization (to 95 ± 6% of control after 60 min, Table 5), was reduced greatly in comparison with IAA alone. This gradual depolarization continued for >= 2 h after IAA addition and did not attain a steady state level during the times examined. Notably, no hyperpolarizing inflections were ever seen (i.e., phases 1 or 3) in IAA-poisoned nerves pretreated with TTX. Further support for Na+-mediated depolarization was provided by experiments where Na+ was replaced with the impermeant cation choline. Zero-Na+/choline alone caused a prompt hyperpolarization followed by depolarization to a stable plateau (Fig. 5B; see also Fig. 3A in companion paper). Addition of IAA produced a small but prompt hyperpolarization (Fig. 5B, right-arrow) with little depolarization during 60 min of application (Table 5).


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FIG. 5. Role of Na+ during glycolytic inhibition and chemical anoxia. A: block of tetrodotoxin (TTX)-sensitive Na+ channels elicited a small hyperpolarizing effect. Under these conditions, glycolytic inhibition with iodoacetate (1 mM) resulted in a greatly reduced depolarization. B: replacement of Na+ with the impermeant cation choline caused a transient hyperpolarizing response. Addition of iodoacetate caused a prompt hyperpolarization (right-arrow) with complete preservation of membrane potential for >= 60 min. C: in contrast, TTX and NaCN application elicited significantly different responses between nerves A (recorded immediately) and nerves B (recorded after several additional hours of in vitro incubation, see text). For nerves A, NaCN application evoked a prompt depolarization that was much less extensive than without TTX. In contrast, nerves B failed to depolarize. These results suggest an additional Na+ influx pathway into RON axons, distinct from TTX-sensitive Na+ channels. This pathway appears to be downregulated after several hours of in vitro incubation (see text). D: similarly, NaCN and zero-Na+/choline solution caused a blunted but consistent depolarization for nerves A. Nerves B (Vm not normalized to -80 mV baseline) instead hyperpolarized slightly (right-arrow) after addition of CN- with little further depolarization. See Fig. 1 legend for definition of Vg and Vm.

 
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TABLE 5. Role of Na+ during glycolytic inhibition and chemical anoxia

For IAA and TTX, the trajectory of Vg was similar for nerves A (studied immediately after dissection) and nerves B (maintained in oxygenated aCSF at room temperature for several additional hours), which was not the case for CN-. For TTX-treated nerves A, addition of CN- resulted in a fast, albeit limited (compared with CN- without TTX, P < 0.0001), depolarization to 76 ± 13% of control after 60 min of treatment (Table 5). In contrast, nerves B failed to depolarize after CN- was added to the TTX perfusate (Vg = 99 ± 7% of control after 60 min of CN-), which was significantly different from nerve A (P < 0.05; Fig. 5C, Table 5). The nerve A and B difference also was observed for CN- and zero-Na+/choline (P < 0.001). Similar to the CN- and TTX nerves A, CN- exposure after 60 min of zero-Na+/choline produced an immediate, but slow depolarization (Fig. 5D) to 81 ± 5% of control after 60 min of CN- (Table 5). In contrast to nerves A, nerves B responded by hyperpolarizing (Fig. 5D, arrow) in CN-, then modestly depolarized to 98 ± 4% of control (Table 5). Results were similar regardless of whether CN- was applied as the Na+ or K+ salt (KCN solution, Table 1), indicating that the small amounts of Na+ from NaCN had little effect on the nominally zero-Na+/choline responses (data not shown).

We studied Vg during chemical anoxia with the use-dependent Na+ channel blockers, procaine and QX-314. In Fig. 6A, procaine (1 mM) applied for 60 min had a hyperpolarizing effect on Vg to 106 ± 2% of control (Table 6). Procaine greatly reduced the CN--induced depolarization (Vg = 84 ± 11% of control after 60 min, Table 5). QX-314 [300 µM, a concentration that does not block action potential generation (Stys et al. 1992a)], a quaternary analogue of lidocaine, hyperpolarized Vg to 105 ± 2%; this effect developed much more slowly than that of procaine (Fig. 6B, Table 6). The nerve A and B difference observed during CN-/TTX and CN-/zero-Na+/choline treatment also occurred for CN- and QX-314. Nerves A depolarized to 80 ± 9% of control whereas Vg for nerves B remained unchanged at 101 ± 6% (P < 0.05, Table 5).


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FIG. 6. Local anesthetics reduce anoxic RON depolarization. A: procaine (1 mM) alone had a rapid hyperpolarizing effect on membrane potential and blunted the depolarizing effect of chemical anoxia. B: QX-314 (300 µM) alone also elicited a small hyperpolarization, which developed much more slowly that with procaine, likely reflecting the limited rate at which this permanently charged compound could cross the axolemma to gain access to the cytosolic side of the Na+ channel. Differences were observed for nerves A and B (see Fig. 5 legend and text for details) after CN- was applied, causing limited depolarization in nerves A that was further reduced for nerves B. See Fig. 1 legend for definition of Vg and Vm.

    DISCUSSION
Abstract
Introduction
Methods
Results
Discussion
References

Glucose and oxygen are instrumental in sustaining adequate cellular energy metabolism because ATP is derived almost exclusively from a continuous supply of glucose in the brain (Erecinska and Silver 1989). Although a number of studies have been carried out on the effects of anoxia/glycolytic inhibition on peripheral myelinated axons (Brismar 1981; Lindstrom and Brismar 1991; Lundberg and Oscarsson 1954; Maruhashi and Wright 1967; Schoepfle and Bloom 1959), there is far less data on metabolically compromised central axons. Several studies have examined the effects of anoxia on action potential conduction in CNS fibers (Davis and Ransom 1987; Fern et al. 1995; Lee et al. 1993; Stys et al. 1990), to our knowledge there is no detailed account of the ionic determinants of resting membrane potential in central mammalian axons during anoxia or glycolytic block. We therefore investigated the effect of metabolic compromise on membrane potential in the RON, a representative model of mammalian CNS axons.

A large proportion of the ATP synthesized from glycolysis and oxidative phosphorylation is used by Na+,K+-ATPase to maintain membrane potential (Vm) in neural cells (Ritchie 1967). Unexpectedly, various methods of blocking glycolysis that, in theory, possessed similar modes of action, produced different depolarization trajectories. IAA likely blocked glycolysis profoundly, causing a more synchronous disruption of energy-dependent mechanisms, eliciting a response consisting of four distinct phases. Conversely, the energy depletion from zero glucose exposure occurred more gradually (probably due to slow removal of glucose from the extracellular space), resulting in a less severe insult and consequently a more gradual collapse of ion gradients and Vm (Tables 7 and 8). Axons also may have derived energy from the degradation of astrocytic glycogen (Cataldo and Broadwell 1986; Dringen and Hamprecht 1992; Swanson and Choi 1993), possibly via lactate transfer (Fern and Ransom 1996; Tsacopoulos and Magistretti 1996). Adding DG, an inhibitor of hexokinase (Devlin 1992), may have produced a somewhat harsher disruption of glycolysis, with emergence of the four characteristic phases typical of IAA (Fig. 1). Interference with glial glycogen breakdown by DG (Dringen and Hamprecht 1993) also may have contributed to the observed differences. Pyruvate completely prevented nerve depolarization induced by IAA, making it unlikely that nonspecific effects of this inhibitor were responsible for Vm changes in agreement with previous reports (Ochs and Smith 1971; Sabri and Ochs 1971). The results also indicated that RON axons are fueled preferentially by aerobic metabolism, which generates the majority of required ATP at rest, as in other parts of the CNS (Erecinska and Dagani 1990); glycolysis contributed insufficient and only minor amounts of ATP for the maintenance of Vm (Fig. 2).

A feature common to all three methods of glycolytic inhibition was a significant delay (20-30 min) before any effect on Vm was noticeable. Although it could be argued that slow washout of glucose was responsible for zero-glucose (± DG) treatments, a delayed penetration of IAA into the cells is unlikely as this inhibitor produced almost immediate effects under zero-Ca2+/EGTA conditions (Fig. 3C). It is possible that zero Ca2+ solutions altered the axon-myelin relationship as well as cell membranes (Schlaepfer and Bunge 1973), allowing more rapid entry of IAA. Alternatively, operation of the tricarboxylic acid cycle/oxidative phosphorylation may have continued temporarily in IAA poisoned nerves, using alternate substrates such as amino acids (Stryer 1988). Evidence for persistent oxidative phosphorylation activity during glycolytic inhibition was shown in cultured rat astrocytes and neurons (Pauwels et al. 1985) and guinea pig synaptosomes (Kauppinen and Nicholls 1986b). These studies suggested that the cells generated energy using substrates other than glycolytically derived pyruvate. Further support for maintenance of Vm mainly by aerobic metabolism (with minimal glycolytic contribution) was provided by the observation that nerves depolarized at a similar rate (phase 2) with IAA and CN-, despite a delayed onset with the former.

Role of Ca2+ during metabolic stress

The early distinct hyperpolarizing response (phase 1) observed in the majority of nerves during glycolytic inhibition with IAA and DG raised the possibility of a K+ conductance activation. These results are similar to guinea pig (Hansen et al. 1982) and rat (Belousov et al. 1995; Fujiwara et al. 1987; Leblond and Krnjevic 1989) hippocampal neurons in which hyperpolarization occurred during anoxia secondary to K+-conductance increase. It was suggested that a rise in [Ca2+]i was responsible for the hyperpolarizing response stimulating a KCa2+ channel (Leblond and Krnjevic 1989). Our data are also consistent with such a mechanism because phase 1 was Ca2+ dependent. Phase 3, which interrupted the rapid nerve depolarization with a hyperpolarizing inflection during glycolytic inhibition, was also dependent on Ca2+, raising the possibility that phase 3 is a continuation of phase 1 sharing similar mechanisms. Moreover, reduction or elimination of these responses by TEA further supports our hypothesis of a mechanism involving activation of a KCa2+ channel. These hyperpolarizing responses were reduced progressively with more severe Ca2+-depleting treatments (Fig. 3), suggesting that a relatively inaccessible source of Ca2+ may have been the trigger, perhaps originating from internal Ca2+ stores (Belousov et al. 1995) known to be present in myelinated axons (Takei et al. 1992). The absence of hyperpolarizing responses during CN- treatment, despite rapid and massive depolarization, may indicate preferential maintenance of internal Ca2+ stores by glycolytically derived ATP (Xu et al. 1995). Ca2+-depleted conditions accelerated the rate of depolarization during chemical anoxia, consistent with a more rapid loss of RON excitability (Stys, unpublished data). This may be due to removal of charge screening by the divalent cation, resulting in a leakier membrane (Hille et al. 1975; Woodhull 1973). However, zero-Ca2+ conditions markedly slowed the depolarization rates in glycolytically inhibited nerves, suggesting that Ca2+ depletion has more complex effects than simple charge screening removal at the membrane. We believe that the hyperpolarizing responses are not likely due to activation of the K+ conductance opened by low levels of ATP (KATP) (Jonas et al. 1991) because phases 1 and 3 (early and late hyperpolarizing responses, respectively) were abolished during zero Ca2+ conditions and, in addition, the KATP channel antagonist,glibenclamide, failed to blunt the hyperpolarizing phases (data not shown).

Phase 1 occurred without delay during zero-Na+/choline and IAA exposure as well as during zero-Na+/choline and CN- (nerve B) application. Moreover, it was absent during the concomitant application of IAA and the Na+ channel antagonist, TTX. The findings suggest a mechanism dependent on Na+ channels but not on Na+ influx (the response was still present with zero-Na+/choline application). It is possible that during IAA alone, the Na+/Ca2+ exchanger removed Ca2+ entering through Na+ channels (DiPolo et al. 1982; Stys and LoPachin 1997) delaying the rise in [Ca2+]i. During zero-Na+ perfusion, the exchanger was unable to buffer Ca2+, thereby permitting a faster increase in [Ca2+]i and activation of the Ca2+-dependent hyperpolarizing mechanism. We propose a dual role for the Na+/Ca2+ exchanger: at the beginning of metabolic stress, it operates to extrude Ca2+, but in the later stages of anoxia/ischemia, when the Na+ gradient has collapsed, it imports damaging quantities of Ca2+ (Stys et al. 1990, 1992b). The inflection recorded during CN- exposure alone may be analogous to the hyperpolarization in CN- and zero-Na+/choline (Fig. 5D, nerve B), truncated by the massive and rapid depolarization.

The abrupt reduction in depolarization rate coinciding with phase 3 (Fig. 1C), persisted in TEA despite the abolition of distinct hyperpolarizations by this blocker. This suggests that the mechanisms underlying the rate change are distinct from those mediating the hyperpolarizing phenomena, although both appear to be relatively Ca2+-dependent. Although the hyperpolarizing shifts were likely due to K+ conductance activations, given the rate limiting effect on depolarization of Na+ conductance (see below), it is likely that shifts in rate of decay of Vm were due to reductions in Na+ conductance, rather than increases in K+ conductance. Other studies have shown that anoxia decreases Na+ permeability and shifts the steady state inactivation curve to increasingly negative potentials (Brismar 1981, 1983; Cummins et al. 1991; Haddad and Jiang 1993). The mechanism for this presumed conductance change is unknown but may involve Ca2+-dependent phosphorylation of Na+ channels by protein kinase (Li et al. 1993). The reduction of Na+ permeability may represent a stereotyped response to metabolic stress, which would at the same time reduce the drain on already strained energy reserves and diminish the degree of depolarization and deleterious Na+/Ca2+ exchange-mediated Ca2+ influx.

Role of Na+ during metabolic stress

The immediate depolarizing response elicited by CN- demonstrated a major dependence of axonal polarization on aerobic metabolism, consistent with the rapid loss of excitability previously observed within minutes of anoxia (Stys 1996; Stys et al. 1990). Nerve depolarization by glycolytic inhibition or chemical anoxia was reduced by blocking TTX-sensitive Na+ channels. One explanation may be that Na+ influx is rate limiting and will in turn affect the rate of K+ loss for reasons of electroneutrality: reducing Na+ influx therefore will reduce loss of internal K+ and accumulation of extracellular K+ (Ransom et al. 1992), thereby maintaining membrane polarization. Relative preservation of membrane potential during metabolic inhibition where Na+ was replaced with the impermeant cation choline further supports this possibility.

Nerves A (studied immediately after dissection) and nerves B (stored in oxygenated aCSF at room temperature for several additional hours) exposed to Na+ channel blockers (TTX, QX-314) or Na+-depleted perfusate, responded very differently to the addition of CN-: nerves A depolarized (although not nearly to the same extent as with CN- alone), whereas nerves B did not (Fig. 5, C and D). Curiously, glycolytic inhibition did not distinguish between the two nerve populations. The persistent (albeit blunted) depolarization observed during anoxia in nerves studied promptly, despite Na+ channel block with TTX, may suggest an additional Na+ influx path that appeared to be downregulated with prolonged in vitro incubation, possibly due to washout of modulatory factors (for example see Zhang and Krnjevic 1993).

The local anesthetics procaine and QX-314 had similar effects to TTX. Both agents caused a small but reproducible hyperpolarization, indicating block of a persistent Na+ conductance at rest (Stys et al. 1993). These compounds act at the cytoplasmic side of the neuronal Na+ channel; the onset of the procaine effect was rapid likely due to its ability to permeate the membrane in its uncharged form. In contrast, the time of onset of the permanently charged QX-314 was much longer, reflecting the much slower movement across the axolemma by this molecule. As with TTX, both compounds greatly reduced nerve depolarization during chemical anoxia. Unlike TTX, however, QX-314 does not abolish electrogenesis at the concentration used (Stys et al. 1992a), and its membrane potential sparing effect is likely due to this drug's preferential action at open, noninactivating Na+ channels (Khodorov 1991; Wang et al. 1987). QX-314 appeared to reduce the extent of depolarization for nerves A even more effectively than TTX alone (compare Figs. 5C and 6B), similar to complete Na+ replacement with impermeant choline (Fig. 5D). Given the evidence suggesting additional Na+ influx pathways in RON axons (see above), perhaps QX-314 has activity at these as yet unidentified influx routes. One possibility is the inward rectifier, known to be present on RON axons (Eng et al. 1990) and to possess finite Na+ permeability (Karst et al. 1993; Solomon and Nerbonne 1993). This idea is supported further by observations that blocking this channel with Cs+ was highly protective against RON anoxia (Stys and Hubatsch 1996), likely due to a reduction of depolarization and Na+ influx.

In conclusion, we determined that the resting potential of the RON depends on energy generated primarily from aerobic metabolism with a minor contribution from glycolytic ATP. Inhibition of glycolysis or oxidative phosphorylation depolarized the recorded potential but with quite different trajectories. The effects of glycolytic inhibition alone were delayed, suggesting a limited ability of axons to generate aerobic ATP from alternate substrates. The inevitable massive depolarization was largely, although not exclusively, dependent on Na+ influx through TTX-sensitive Na+ channels [likely the noninactivating Na+ conductance previously demonstrated in the RON (Stys et al. 1993)], with evidence for additional TTX-insensitive Na+ influx pathways. The membrane potential-sparing effect of local anesthetics likely contributes to their neuroprotective actions. Finally, CNS axons may possess autoprotective mechanisms (Fern et al. 1996), including possible activation of a Ca2+-dependent K+ conductance(s) to delay depolarization onset, and perhaps a controlled reduction of Na+ permeability to slow the decay of Vm, which also appears to be Ca2+ dependent. We therefore suggest that during the initial stages of metabolic insult, the presumed rise in free [Ca2+]i may be transiently beneficial, reflecting the axon's intrinsic mechanisms designed to limit injury during anoxia/ischemia.

    ACKNOWLEDGEMENTS

  We thank Dr. Bin Hu for valuable comments on the manuscript.

  This work was supported in large part by the Heart and Stroke Foundation of Ontario.

    FOOTNOTES

  Address for reprint requests: P. K. Stys, Ottawa Civic Hospital, 1053 Carling Ave., Ottawa, Ontario K1Y 4E9, Canada.

  Received 6 January 1997; accepted in final form 17 June 1997.

    REFERENCES
Abstract
Introduction
Methods
Results
Discussion
References

0022-3077/97 $5.00 Copyright ©1997 The American Physiological Society