Ion Transport and Membrane Potential in CNS Myelinated Axons I. Normoxic Conditions

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. I. Normoxic conditions. J. Neurophysiol. 78: 2086-2094, 1997. Compound resting membrane potential was recorded by the grease gap technique during normoxic conditions (37°C) in rat optic nerve, a representative CNS myelinated tract. Mean potential was -47 ± 3 (SD) mV and remained stable for 2-3 h. Input impedance of a single optic nerve axon was calculated to be approx 5 GOmega . Contribution of the Na+ pump to resting axonal potential is estimated at -7 mV. Ouabain (10 µM to 10 mM) evoked a dose-dependent depolarization that was maximal at >= 1 mM, depolarizing the nerves to ~35-40% of control after 60 min. Inhibiting energy metabolism (CN- and iodoacetate) during high-dose ouabain (1-10 mM) exposure caused an additional depolarization, suggesting additional ATP-dependent, ouabain-insensitive ion transport systems. Perfusion with zero-Na+ (choline substituted) caused a transient hyperpolarization, that was greater than with tetrodotoxin (TTX; 1 µM) alone, indicating both TTX-sensitive and -insensitive Na+ influx pathways in resting rat optic nerve axons. Resting probability (P)K:PNa is calculated at 20:1. In contrast to choline-substituted solution, Li+-substituted zero-Na+ perfusate caused a rapid depolarization due to Na+ pump inhibition and the ability of Li+ to permeate the Na+ channel. TTX reduced, but did not prevent, ouabain- or zero-Na+/Li+-induced depolarization. We conclude that the primary Na+ influx path in resting rat optic nerve axons is the TTX-sensitive Na+ channel, with evidence for additional TTX-insensitive routes permeable to Na+ and Li+. In addition, maintenance of membrane potential is critically dependent on continuous Na+ pump activity due to the relatively high exchange of Na+ (via the above mentioned routes) and K+ across the membrane of resting optic axons.

    INTRODUCTION
Abstract
Introduction
Methods
Results
Discussion
References

Axons play the critical role of conducting electrical signals within the nervous system with high fidelity and efficiency. To sustain electrogenesis, transmembrane K+ and Na+ gradients maintain axons in a polarized state and provide energy for signaling, respectively. These electrochemical gradients are established by energy-dependent ion transport systems, the most important of which is the Na+,K+-ATPase (Gordon et al. 1990; Horisberger et al. 1991; Rossier et al. 1987; Sweadner 1995). Because membrane conductances responsible for initiation and termination of the action potential are voltage dependent, resting properties of an axon, such as membrane potential, ionic permeabilities, and the various transporters that move ions against their gradients, will influence its electrical behavior. Moreover, these same properties may affect the response of an axon to pathophysiological conditions, such as anoxia/ischemia, and may in turn modulate the degree of injury sustained by the fiber. For these reasons, it is important to define these properties as they will fundamentally influence the response of the axon to physiological and pathophysiological stimuli. We investigated the ionic determinants underlying the resting membrane potential in central myelinated axons of the rat optic nerve (RON) and explored some of the membrane transport systems involved. Our findings are extended in the companion paper, which describes the responses of axonal membrane potential to cellular energy depletion. Preliminary results have been presented in abstract form (Leppanen and Stys 1996).

    METHODS
Abstract
Introduction
Methods
Results
Discussion
References

Long-Evans male rats (150-175 g) were anesthetized with 80% CO2-20% O2 and decapitated, and the optic nerves dissected. The axonal compound resting membrane potential, Vg, was recorded in vitro with the grease gap technique (Fig. 1A and next section) (Stys et al. 1993). This technique provides a more stable recording in central mammalian axons at physiological temperatures than the conventional sucrose gap method (Eng et al. 1990; Stämpfli 1954). The small diameter of optic nerve axons [mean <1 µm (Foster et al. 1982)] and the long recording times required (up to 4 h) precluded the use of intracellular microelectrodes. The middle segment of one nerve was inserted into a silastic tube slit longitudinally (approx 2 mm long, 0.64 mm diam) filled with petroleum jelly for electrical isolation and to minimize intermixing of solutions from the two wells (Fig. 1A). One end was perfused at 2 ml/min with oxygenated artificial cerebrospinal fluid (aCSF) or test solutions at 37°C. The opposite end was depolarized using an isotonic K+ solution (NaCl replaced by KCl) containing 0.5 mM CaCl2 (to reduce potential Ca2+-mediated injury in strongly depolarized axon segments) flowing at 2 ml/min at room temperature. All solutions were gassed with 95% O2-5% CO2. To improve recording stability, the central nerve segment embedded in petroleum jelly was cooled by circulating a cold (approx 0°C) solution of 1:1 glycerol/water through tubing in contact with the silastic tube. Vg was recorded with 3 M KCl/agar bridge electrodes. Junction potentials were determined at the beginning and end of each experiment by shorting the two wells with a strand of paper. These potentials, which never exceeded 6 mV, drifted linearly over time (data not shown). Vg recorded during the course of an experiment therefore was corrected by subtracting estimated values of junction potentials obtained by linear interpolation between the readings obtained at the start and end of the experiment. The trans-gap resistance, Rg, was measured in some experiments by applying current steps (98 nA) across the two wells (ig, Fig. 1A), small enough to prevent activation of voltage-sensitive conductances. After dissection, one nerve was recorded immediately (nerve A), while the second (nerve B) was placed in an oxygenated chamber (95% O2-5% CO2) containing aCSF at room temperature for later study.


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FIG. 1. A: diagram of the grease gap recording chamber. A nerve is inserted into the slit silastic tube containing petroleum jelly, cooled for stability, with one end perfused with artificial cerebrospinal fluid or test solutions and the other end is depolarized with isotonic K+ solution. The potential, Vg, was recorded with 3 M KCl/agar bridge electrodes. Constant current steps (ig) were applied across the wells to measure Rg. B: electrical model of the grease gap technique. Vg is the recorded potential. Vm and V'm are the absolute transmembrane potentials at each end of the nerve. Re is the external resistance, composed of extracellular space, nerve sheath, residual solution and other parallel leakage pathways. Ri is the combined internal axoplasmic and membrane resistances. A current ig will flow when Vm and V'm are unequal, and the resultant voltage drop (Vg) is recorded across Re, representing a stable, constant fraction under control conditions of the absolute transmembrane potential, Vm (see text).

Composition of aCSF and zero-Na+ solutions [choline Cl (BDH), LiCl (Sigma), N-methyl-D-glucamine (NMDG; Sigma)] are listed in Table 1. Tetrodotoxin (TTX; Sigma) was diluted from stock solution in distilled water. Ouabain (Sigma, RBI), NaCN (BDH), and iodoacetate-Na+ salt (IAA; Sigma) were dissolved in aCSF.

 
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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

Vg recorded from an optic nerve by the grease gap technique during control conditions is illustrated in Fig. 2A. After nerve insertion, Vg usually stabilized within 90 min with potentials ranging from -40 to -55 mV [-47 ± 3 (mean ± SD) mV]. No differences were noted between nerves A and B (see METHODS; -47.3 ± 3.5 vs. -46.6 ± 4.2 mV, P = 0.18). Rg was typically ~60 kOmega . Both Vg and Rg remained relatively constant for several hours under control conditions (Fig. 2A, Table 2). Probable variation in the short-circuit factor (Stämpfli 1954) from nerve to nerve resulted in different absolute values of Vg. For this reason, we routinely 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 (Vm) over time in this and subsequent experiments (Table 5).


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FIG. 2. Recorded (Vg) and calculated absolute (Vm) compound resting membrane potential in rat optic nerve measured by the grease gap technique during control conditions and Na+,K+-ATPase inhibition with ouabain. A: representative trace of membrane potential under control conditions. Vg typically ranged from -40 to -55 mV and remained constant for 2-3 h (the preceding 90 min stabilization period that followed dissection and insertion is not shown). These and subsequent tracings were normalized to the estimated absolute resting membrane potential (Vm, -80 mV; see text). B: ouabain (10 µM-1 mM) evoked nerve depolarization in a dose-dependent manner, indicating a critical (although not exclusive, see C) dependence of RON axonal resting potential on continued pump operation. Ratios were calculated by dividing the potential at 60 min (right-arrow) or 120 min (right-arrowright-arrow) by the time 0 potential. C: metabolic inhibition with NaCN (2 mM) and iodoacetate-Na+ salt (IAA; 1 mM) caused further depolarization after ouabain application (1 mM), suggesting additional ATP-dependent ion transport mechanisms (see text).

 
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TABLE 2. Resting membrane potential recorded during control conditions in aCSF by the grease gap technique

 
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TABLE 5. Calculated absolute resting membrane potential (Vm) under various conditions

Na+,K+-ATPase inhibition

Na+,K+-ATPase plays a key role in maintaining transmembrane Na+ and K+ gradients. Ouabain (1 mM), an antagonist of the pump, caused a rapid depolarization that ended in a stable plateau at 39 ± 3% of control potential after 60 min (right-arrow) or more of exposure (Fig. 2B, Table 3). The initial rapid depolarization proceeded at a rate of6 ± 1 mV/min. Higher ouabain concentrations (2 and 10 mM) produced a slightly greater depolarization that was not statistically different from exposures to 1 mM ouabain (Table 3). Lower concentrations of ouabain (10 and 100 µM) produced proportionally less depolarization (Fig. 2B, Table 3). The substantial membrane potential remaining after presumably complete pump inhibition with 1-10 mM ouabain was surprising. We hypothesized that additional energy-dependent mechanisms, independent of Na+,K+-ATPase, may be in part responsible for maintenance of Vm. Nerves were depolarized by a 60-min exposure to ouabain (1 mM) as above. Mitochondrial and glycolytic ATP production then was blocked by adding CN- (2 mM) and IAA (1 mM), respectively, to the ouabain-containing perfusate. This treatment resulted in a further depolarization to 22 ± 4% of control (Fig. 2C, Table 3) exceeding that seen with ouabain alone (P < 0.0001). Similar results were obtained with 10 mM ouabain and metabolic inhibition and also with 1 mM ouabain and lowered temperature (room temperature) (data not shown).

 
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TABLE 3. Effect of Na+, K+-ATPase inhibition with ouabain on membrane potential

Na+ permeability

The influence of membrane Na+ permeability on Vg was studied by substitution of bath Na+ with other monovalent cations or pharmacological block of Na+ channels. Replacing Na+ with the impermeant cation choline elicited a transient hyperpolarizing response. The maximal hyperpolarization reached 109 ± 3% of control (Fig. 3A, right-arrow, Table 4) for nerve A (the nerve recorded immediately after dissection) and was greater for nerve B (113 ± 3%, P < 0.002). The hyperpolarization was followed by a slow depolarization to a stable potential (96 ± 8% of control, Table 4) within 60 min, with no difference between nerves A and B. NMDG-substituted zero-Na+ solution produced similar results (data not shown). To test whether Na+,K+-ATPase still was operating after perfusion with zero-Na+ solution, ouabain was added after 60 min, resulting in a prompt, albeit limited, depolarization (Fig. 3B) to 79 ± 5% of control potential in aCSF alone (Table 4).


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FIG. 3. Effect of Na+ substitution experiments on resting membrane potential. A: replacement of Na+ with the impermeant cation, choline, caused a hyperpolarizing response (right-arrow), followed by modest depolarization to a constant potential, which typically stabilized within 60 min. B: ouabain added to the zero-Na+/choline perfusate evoked a small, but limited depolarization, indicating continued pump operation under Na+-depleted conditions. C: application of the permeant cation, lithium, in the absence of Na+, elicited a small, brief hyperpolarization (right-arrow). The nerve then depolarized rapidly, followed by a hyperpolarizing sag (right-arrowright-arrow). D: zero-Na+/lithium application for 60 min produced a similar response described in C. Addition of ouabain to zero-Na+/lithium conditions elicited a substantial depolarization, indicating continued pump operation under these Na+-substituted conditions. See Fig. 2 legend for definition of Vg and Vm.

 
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TABLE 4. Effect of sodium substitution on Vm during various conditions

Substituting bath Na+ with Li+, a cation permeable at the Na+ channel (Hille 1992; Richelson 1977), resulted in a different voltage trajectory. In contrast to results with impermeant cations, zero-Na+/Li+ solution caused only a small, brief hyperpolarization (Fig. 3C, right-arrow) that persisted in zero external Ca2+ bath (data not shown). This was followed by a steep, rapid depolarization (rate: 4 ± 1 mV/min) to a plateau of65 ± 5% of control after 60 min (Table 4), significantly less depolarized than with 1 mM ouabain (39 ± 3%, P < 0.0001, Table 3). In several nerves, a small hyperpolarizing "sag" also was noted (Fig. 3C, right-arrowright-arrow). To examine whether the Na+,K+-ATPase was operating during Li+-substituted conditions, ouabain (1 mM) was added after 60 min of zero-Na+/Li+ exposure, resulting in a substantial additional depolarization to32 ± 3% of control (Fig. 3D, Table 4).

Choline substituted-zero-Na+ data provided evidence that ouabain-induced depolarization was dependent to a large extent on extracellular Na+. The role of voltage-gated Na+ channels in isolation was studied by applying TTX (1 µM), which evoked a small hyperpolarizing response (Fig. 4A) to 104 ± 1% of control membrane potential after 20 min (Table 4). The TTX-induced hyperpolarization was significantly smaller than that produced by zero-Na+/choline perfusion (P < 0.001), suggesting Na+ influx pathway(s) in addition to TTX-sensitive Na+ channels. Blocking TTX-sensitive Na+ conductance during zero-Na+/Li+ perfusion resulted in depolarization to 75 ± 10% of control, which proceeded at a significantly slower rate of 1 ± 0.3 mV/min compared with Li+ alone (P < 0.0001; Fig. 4B, Table 4). TTX blunted but did not abolish the depolarizing response, providing additional support for a TTX-insensitive route(s) of Na+ and Li+ entry. The hypothesis was supported by applying ouabain to nerves pretreated for 20 min withTTX (Fig. 4C). This still resulted in a marked depolarization to 58 ± 5% of control, less than with ouabain alone (39 ± 3% after 60 min, P < 0.001, Table 3) and also occurring at a considerably slower rate (1 ± 0.1 vs. 6 ± 1 mV/min, P < 0.005).


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FIG. 4. Effects of the Na+ channel blocker tetrodotoxin (TTX). A: TTX alone had a hyperpolarizing effect on Vm, reaching a stable potential within 20 min, indicating a finite, TTX-sensitive Na+ conductance in resting RON axons. B: switching to zero-Na+/lithium perfusate caused a depolarization that was more gradual and limited compared with TTX-free conditions (Fig. 3C). C: TTX also reduced the rate of nerve depolarization during ouabain treatment. Together these results suggest additional, TTX-insensitive Na+ influx pathways in resting RON axons. See Fig. 2 legend for definition of Vg and Vm.

    DISCUSSION
Abstract
Introduction
Methods
Results
Discussion
References

Grease gap electrical model

The potential recorded by the grease gap technique, Vg, is governed by the electrical model illustrated in Fig. 1B. If Vm and V'm are unequal, a current ig will flow, and the recorded potential will be
<IT>V</IT><SUB>g</SUB><IT>= i</IT><SUB>g</SUB><IT>R</IT><SUB>e</SUB> (1)
The magnitude of the current will be
<IT>i</IT><SUB>g</SUB>= (<IT>V</IT><SUB>m</SUB><IT>− V</IT><SUP></SUP>′<SUB>m</SUB>)/(<IT>R</IT><SUB>e</SUB><IT>+ R</IT><SUB>i</SUB>) (2)
As V'm approaches 0 in isotonic KCl solution, and eliminatingig from Eqs. 1 and 2, the expression reduces to (Stämpfli 1954)
<IT>V</IT><SUB>g</SUB><IT>= V</IT><SUB>m</SUB><IT>R</IT><SUB>e</SUB>/(<IT>R</IT><SUB>e</SUB><IT>+ R</IT><SUB>i</SUB>) (3)
where Vg is the recorded potential, Vm is the absolute transmembrane potential, Ri is the internal axoplasmic and combined membrane resistances, and Re is the external resistance, composed of extracellular space, nerve sheath, residual solution, and other parallel leakage pathways.

The term Re/(Re + Ri), defined as the "short circuit factor" (Jirounek and Straub 1971; Stämpfli 1954), will tend to reduce Vg to a constant fraction of the true transmembrane potential, Vm. When Re is assumed to be much greater than Ri [as might occur in sucrose gap recordings of single fibers (Julian et al. 1962)], Vg will approach Vm. In our preparation, this assumption may not be valid. Independent estimates of Vm in optic axons [approximately equal to -80 mV (Stys et al. 1997)] differ from our typical grease gap potentials (approximately equal to -50 mV) by a factor of approx 0.6, which represents an estimate of our short circuit factor (Eq. 3). Importantly, although absolute transmembrane potentials could not be recorded from optic axons using this technique, as long as the short circuit factor remains constant, Vg will represent a reliable fraction of Vm and will vary linearly with Vm. Our data showed that the potential recorded by the grease gap represented a reliable fraction of the true axonal compound resting membrane potential in a CNS myelinated axon bundle. Although the magnitude of Vg was less than Vm due to a short circuit factor less than unity, this disadvantage was offset by the long-term stability of our recordings (Fig. 2A), essential for the questions we wished to address in this preparation. The equivalent mean absolute membrane potential during control conditions varied from -80 to -81 mV during 150 min. Table 5 lists calculated absolute membrane potentials during various experimental conditions.

The measured potential, Vg, originates from a voltage drop across Re produced by a current ig (Fig. 1B). Current flow through each constituent fiber is inversely proportional to its axial resistance and therefore directly proportional to the square of axon diameter. Because the RON is composed of a parallel bundle of fibers with different diameters ranging from 0.3 to 3 µm (Hildebrand and Waxman 1984), it follows that the current contribution to the total current ig, and therefore to recorded potential Vg, will be biased in favor of larger fibers. Finally, because of the origin of the recorded potential and its dependence on the short circuit factor, we cannot exclude effects from some drug applications (see below and companion paper) that may affect the short circuit factor by virtue of altered membrane impedance or altered axo-glial relationships.

Input impedance of optic axons

Combining equations for the short circuit factor (f) and Rg (Fig. 1B) and solving for Ri we obtain
<IT>R</IT><SUB>i</SUB><IT> = R</IT><SUB>g</SUB>/<IT>f</IT>
where Ri represents the parallel combination of axial resistances of all constituent axons. Using a typical measured Rg value of 60 kOmega and f = 0.6, Ri is calculated to be 100 kOmega . Adult optic nerves contain ~100,000 axons (Fukuda et al. 1982), therefore the average Ri per fiber is 10 GOmega , which is an approximation of the axonal input impedance as a semi-infinite cable. The true input impedance of the infinite equivalent is one-half of this value or ~5 GOmega [calculated range of ~20 GOmega for the smallest (0.3 µm diam) to 600 MOmega in large (3 µm diam) optic nerve axons]. Peripheral rat motor axons with a mean diameter of 7.7 µm had a mean input impedance of 45 MOmega (David et al. 1995). As an approximation, scaling this input impedance to a smaller optic axon [mean diameter 0.75 µm (Foster et al. 1982)] as the inverse 3/2 power of diameter (Jack et al. 1983) yields a calculated value of 1.5 GOmega . Similarly, peripheral lizard axons with a mean diameter of 9.8 µm and impedance of 96 MOmega (David et al. 1995) would result in an estimated input impedance of 4.5 GOmega in an equivalent axon scaled to 0.75 µm diam. Although comparisons between different axon types might not always be applicable, these calculations illustrate that our estimate of 5 GOmega is within the same order of magnitude as more direct measures from larger myelinated fibers.

Na+,K+-ATPase inhibition

The importance of Na+,K+-ATPase in maintaining Vm was illustrated by the effect of the inhibitor, ouabain. The loss of ion gradients resulted in rapid nerve depolarization in a dose-dependent manner, which was exacerbated by either metabolic inhibition (CN- and IAA) or lowered temperature suggesting a ouabain-insensitive, ATP-dependent transport mechanism for partial maintenance of Vm. Similar observations were found in rat hypoglossal neurons (Jiang and Haddad 1991). Given that the Na+,K+-ATPase also may be located at the internodal axolemma (Mata et al. 1991), it is possible that some pump molecules were shielded from bath-applied ouabain by the myelin sheath and continued to operate: these pumps then failed when nerves were poisoned with CN- and IAA. Failure of additional energy-dependent ion transport mechanisms, or activation of PNa and/or reduction of PK by metabolic inhibition (which would cause additional depolarization), may have contributed as well. A finite potential remained after combined ouabain and metabolic block, which may be partly due to a passive Donnan equilibrium (Junge 1981), diffusion of K+ from the isotonic K+ well to the test well contributing to a higher [K+]i in the latter compartment, and/or a poorly explained hyperpolarizing shift caused by low-conductance media such as sucrose, observed in sucrose gap recordings (Jirounek et al. 1981; Julian et al. 1962).

Na+ permeability

TTX reduced resting Na+ permeability, producing a hyperpolarizing response that was shown previously (Stys et al. 1993). Replacing Na+ with an impermeant cation caused a greater hyperpolarization, indicating additional, TTX-insensitive resting Na+ permeability in RON axons. Our estimated absolute hyperpolarization of 7 mV in zero-Na+/choline [-87 mV (Table 5) vs. -80 mV (Stys et al. 1997)] was identical to that observed by Morita et al. (1993) in peripheral lizard axons recorded intra-axonally under the same bath conditions. Using Na+-replacement data (from nerve A only), the resting PK:PNa ratio was estimated at 20:1 using (Stys et al. 1993)
<FR><NU>PK</NU><DE>PNa</DE></FR> = <FR><NU><IT>E</IT><SUB>K</SUB><IT> − E</IT><SUB>Na</SUB><IT>R</IT><SUB>V</SUB><IT></IT></NU><DE>E<SUB>K</SUB><IT>R</IT><SUB>V</SUB><IT> − E</IT><SUB>K</SUB></DE></FR>
where Rv = Vzero-Na+/Vm, i.e., the ratio of membrane potentials after and before Na+ permeability blockade with zero-Na+ solutions. EK and ENa are ionic reversal potentials calculated from measurements of axoplasmic concentrations (LoPachin and Stys 1995; Stys et al. 1997).

This ratio was lower than a previous estimate of 35, which reflected only TTX-sensitive permeability (Stys et al. 1993). Interestingly, the PK:PNa ratio calculated using nerves B (subject to several additional hours of incubation at room temperature) was significantly lower at 15:1 (P < 0.01). In contrast, TTX-induced hyperpolarizations were not different between nerves A and B, indicating that the different PK:PNa ratios were either due to a higher TTX-insensitive Na+ permeability or a reduced K+ permeability in nerves B. Because PNa is less than PK at rest, Na+ permeability will be rate limiting for electroneutral K+ and Na+ exchange across the membrane during pump inhibition. Therefore, one would expect a more rapid depolarization during ouabain exposure or metabolic inhibition (see companion paper) in nerves B if its lower PK:PNa ratio was due to an absolute increase in PNa. Instead we found that neither the rate nor the extent of depolarization were different during ouabain or CN- treatment between the two sets of nerves (P > 0.2), suggesting that the lower PK:PNa ratio in nerves B was due to a reduction of PK rather than increase in PNa. The reasons for the difference are not known but may be related to transient exposure to lower temperature or washout of a soluble modulator (see below and companion paper).

Curiously, despite the observed difference in PK:PNa ratios between nerves A and B, resting membrane potentials were virtually identical. It is likely that hyperpolarizing influences increased with incubation time negating the depolarizing effect of a smaller PK:PNa ratio; for example axonal [K+]i has been shown to rise with in vitro incubation (LoPachin and Stys 1995), and washout of endogenous ouabain-like substance (Blaustein 1993) might increase Na+,K+-ATPase activity. Using a PK:PNa ratio of 20:1, we calculated the contribution of the pump to Vm at rest to be -7 mV (Martin and Levinson 1985), consistent with previous observations of electrogenic pumping in resting and stimulated myelinated fibers (Gordon et al. 1990; Morita et al. 1993).

All nerves depolarized to a variable extent after the zero-Na+/choline-induced hyperpolarization, which we interpret as partial pump inhibition due to depletion of axoplasmic Na+. A new steady state was reached with reduced pump activity and parallel reduction of membrane Na+ conductance. The small depolarization that occurred after the addition of ouabain (Fig. 3B) indicates that Na+,K+-ATPase was not totally inhibited by 60 min of zero-Na+ perfusion, consistent with reports that extensive depletion of intracellular Na+ is difficult to attain (Dunham and Senyk 1977; Stys et al. 1997). It is likely that a small proportion of extracellular Na+ ions were continuously recycled back into the axon to supply the pump at this new steady state rather than negotiate the sinuous extracellular space of the nerve to be removed by the perfusate.

In contrast to zero-Na+/choline, the nerve depolarized markedly during zero-Na+/Li+ because Li+ permeates the Na+ channel, partially inhibiting Na+,K+-ATPase and allowing electroneutral K+ efflux. This effect was strikingly different from that observed in peripheral myelinated axons of the frog, where Li+ is able to substitute for Na+ equally well with respect to maintenance of resting and action potentials (Huxley and Stämpfli 1951). Li+ is not equally permeable to Na+ at the Na+ channel [PNa:PLi is 0.9 (Hille 1992)], therefore, a decrease in [Na+]o may have evoked the brief hyperpolarizing response typically seen (Fig. 3C, arrow). The hyperpolarizing sag noted after depolarization may have resulted from a reduction of [K+]o, caused by a slow washout of the ion. The extent of depolarization with zero-Na+/Li+ was less than expected, given that Li+ inhibits Na+,K+-ATPase (Halm and Dawson 1983). The additional depolarization caused by ouabain after zero-Na+/Li+ application suggested the pump was maintaining Vm by using residual Na+. Electron microprobe data showed that [Na+]i remained at approx 6 mM after 60 min of zero-Na+/Li+ perfusion (Stys et al. 1997), and, at this concentration, the pump still is able to operate at ~20% capacity (Shyjan et al. 1990). It is also possible that the pump substituted Li+ for Na+ (Dunham and Senyk 1977; Ritchie and Straub 1980; Thomas et al. 1975). The application of TTX during zero-Na+/Li+ did not prevent nerve depolarization, which suggested the presence of a TTX-insensitive Li+/Na+ influx pathway in RON axons. Na+ channel blockade with TTX during ouabain exposure supported this notion because this blocker did not eliminate ouabain-induced depolarization, the extent of which exceeded the maximal steady state hyperpolarizing contribution of pump current [approximately equal to -10 mV for a coupling ratio of 3 Na+:2 K+ (Martin and Levinson 1985; Thomas 1972)], indicating transmembrane ion flux rather than mere elimination by ouabain of pump contribution to Vm. However, for both zero-Na+/Li+ or ouabain exposure, TTX reduced the rate of depolarization, due to partial reduction of resting TTX-sensitive Na+ permeability, with gradients dissipating more gradually because of the slowed exchange of Na+ and K+.

Taken together, Na+ influx into RON axons is a key step mediating membrane depolarization during Na+,K+ATPase inhibition. Although the primary Na+ influx path is the TTX-sensitive Na+ channel, our data support the existence of additional, TTX-insensitive Na+ influx pathways that may have important implications for axonal responses to physiological and pathological stimuli.

    ACKNOWLEDGEMENTS

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

  This work was supported in 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