Membrane Dysfunction Induced by In Vitro Ischemia in Rat Hippocampal CA1 Pyramidal Neurons

E. Tanaka, S. Yamamoto, H. Inokuchi, T. Isagai, and H. Higashi

Department of Physiology, Kurume University School of Medicine, 67 Asahi-machi, Kurume 830-0011, Japan


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Tanaka, E., S. Yamamoto, H. Inokuchi, T. Isagai, and H. Higashi. Membrane dysfunction induced by in vitro ischemia in rat hippocampal CA1 pyramidal neurons. Intracellular and single-electrode voltage-clamp recordings were made to investigate the process of membrane dysfunction induced by superfusion with oxygen and glucose-deprived (ischemia-simulating) medium in hippocampal CA1 pyramidal neurons of rat tissue slices. To assess correlation between potential change and membrane dysfunction, the recorded neurons were stained intracellularly with biocytin. A rapid depolarization was produced ~6 min after starting superfusion with ischemia-simulating medium. When oxygen and glucose were reintroduced to the bathing medium immediately after generating the rapid depolarization, the membrane did not repolarize but depolarized further, the potential reaching 0 mV ~5 min after the reintroduction. In single-electrode voltage-clamp recording, a corresponding rapid inward current was observed when the membrane potential was held at -70 mV. After the reintroduction of oxygen and glucose, the current induced by ischemia-simulating medium partially returned to preexposure levels. These results suggest that the membrane depolarization is involved with the membrane dysfunction. The morphological aspects of biocytin-stained neurons during ischemic exposure were not significantly different from control neurons before the rapid depolarization. On the other hand, small blebs were observed on the surface of the neuron within 0.5 min of generating the rapid depolarization, and blebs increased in size after 1 min. After 3 min, neurons became larger and swollen. The long and transverse axes and area of the cross-sectional cell body were increased significantly 1 and 3 min after the rapid depolarization. When Ca2+-free (0 mM) with Co2+ (2.5 mM)-containing medium including oxygen and glucose was applied within 1 min after the rapid depolarization, the membrane potential was restored completely to the preexposure level in the majority of neurons. In these neurons, the long axis was lengthened without any blebs being apparent on the membrane surface. These results suggest that the membrane dysfunction induced by in vitro ischemia may be due to a Ca2+-dependent process that commences ~1.5 min after and is completed 3 min after the onset of the rapid depolarization. Because small blebs occurred immediately after the rapid depolarization and large blebs appeared 1.5-3 min after, it is likely that the transformation from small to large blebs may result in the observed irreversible membrane dysfunction.


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Neurons of the CA1 area of the hippocampus are known to be among the most vulnerable in the CNS to ischemia or anoxia (Brierly and Graham 1984). After 5-30 min of ischemia after occlusion of the carotid artery, hippocampal CA1 neurons show cytoplasmic vacuoles, transient mitochondrial swelling associated with disintegration of internal cristae and of microtubles, and dilatation of rough endoplasmic reticulum and of Golgi cisternae (Petito and Pulsinelli 1984; Yamamoto et al. 1986, 1990; also see Schmidt-Kastner and Freund 1991). In response to oxygen and glucose deprivation, hippocampal CA1 neurons in vitro show a stereotyped response characterized by an initial hyperpolarization followed by a slow depolarization, which leads to a rapid depolarization after ~6 min of deprivation. When oxygen and glucose are reintroduced immediately after generating the rapid depolarization, the membrane potential depolarizes further and approaches 0 mV (the persistent depolarization) (Rader and Lanthorn 1989). Thus the neuron shows no functional recovery (Higashi 1990; Higashi et al. 1990; Kudo et al. 1989; Rader and Lanthorn 1989; Tanaka et al. 1997; also see Martin et al. 1994). Simultaneous recordings of changes in intracellular Ca2+ concentration ([Ca2+]i) and membrane potential recorded in Fura-2/AM-loaded slices revealed a rapid increase in [Ca2+]i corresponding to the rapid depolarization in all CA1 layers (Tanaka et al. 1997; also see Hansen and Zeuthen 1981; Silver and Erecinska 1990; Uematsu et al. 1988). Moreover, pretreatment with N-methyl-D-aspartic acid (NMDA) receptor antagonists or a non-NMDA receptor antagonist inhibits the persistent depolarization and restores the membrane potential to the preexposure level when oxygen and glucose have been reintroduced immediately after generating the rapid depolarization (Rader and Lanthorn 1989; Tanaka et al. 1997; Yamamoto et al. 1997). An inorganic Ca2+ channel blocker, Co2+ (2 mM) or Ni2+ (2 mM), low Ca2+ (0.25 mM) medium, an inhibitor of Ca2+-induced Ca2+ release from intracellular store sites, 8-(diethylamino)octyl-3,4,5-trimethoxybenzoate hydro-chloride (TMB-8) or a Ca2+ chelator, 1,2-bis(2-aminophenyoxy)-ethane-N,N,N',N'-tetraacetic acid tetraacetoxymethyl ester (BAPTA-AM) have similar effects (Yamamoto et al. 1997). These results suggest that the activation of non-NMDA and NMDA receptors and the accumulation of [Ca2+]i have important roles in the induction of membrane dysfunction induced by in vitro ischemia. Nevertheless, the critical period for generating the irreversible change in membrane function is still unclear.

The rapid depolarization induced by in vitro ischemia corresponds to the terminal depolarization (or phase II depolarization) (Hansen 1985) produced by in situ ischemia or asphyxia. The changes occurring in the neuron during and after the rapid depolarization are of interest because they may represent the trigger for the irreversible change that leads to neuronal death (Tanaka et al. 1997; Yamamoto et al. 1997). Thus the present study is concerned with processes involved in the resultant membrane dysfunction induced by in vitro ischemia in hippocampal CA1 neurons in the slice preparation of adult rat. We have examined whether or not the rapid depolarization triggers membrane dysfunction and whether membrane dysfunction occurs during or after the rapid depolarization. In addition, to clarify the structural disorder that leads to the irreversible change, we have compared the morphological structure of the recorded neurons stained by intracellular injection of biocytin before, during, and after the rapid depolarization. Preliminary accounts of some of the data have been presented previously (Higashi et al. 1997; Tanaka et al. 1998).


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The preparation and recording techniques employed were similar to those described in the preceding paper (Isagai et al. 1999; see also Tanaka et al. 1997). Briefly, the forebrain of adult Wistar rats (male 200-250 g weight) was removed quickly under ether anesthesia and placed in chilled (4-6°C) Krebs solution aerated with 95% O2-5% CO2. The composition of the solution was (in mM) 117 NaCl, 3.6 KCl, 2.5 CaCl2, 1.2 MgCl2, 1.2 NaH2PO4, 25 NaHCO3, and 11 glucose. The hippocampus was dissected and then sliced with a Vibratome (Oxford) at a thickness of ~400 µm. Slice preparations were submerged completely in the superfusing solution and preheated to and maintained at 36.5 ± 0.5°C.

Intracellular recordings from CA1 pyramidal neurons were made with glass micropipettes filled with K acetate (2 M), KCl (2 M), or K acetate (2 M) with biocytin (2%). The electrode resistance was 40-80 MOmega . The recording electrodes filled with 2 M KCl were used for single-electrode voltage-clamp experiments. Voltage-clamp recordings were obtained with a single-electrode voltage-clamp amplifier (Axon Instruments, Axoclamp 2B), employing a switching frequency of 5 kHz and a 30% duty cycle. The headstage voltage was continuously monitored to ensure complete settling of the voltage at the end of each switching cycle, and capacitance compensation adjusted while maximizing the sampling rate, according to methods described by others (Finkel and Redman 1984).

Slices were made "ischemic" by superfusion with medium equilibrated with 95% N2-5% CO2 and deprived of glucose, which was replaced with NaCl isoosmotically (ischemia-simulating medium). Ca2+-free with Co2+ (2.5 mM) medium was made by replacement of CaCl2 with CoCl2. When switching the superfusing media, there was a delay of 15-20 s before the new medium reached the chamber due to the volume of the connecting tubing. Thus the chamber was filled with the test solution ~30 s after switching of solution.

For biocytin staining, slices were transferred to 0.1 M phosphate buffer solution with 4% paraformaldehyde buffered to pH 7.4 within 20 s of withdrawal of the recording electrodes filled with K acetate (2 M) and biocytin (2%). After overnight fixation, slices were washed with alcohol (80%) and subsequently dimethylsulfoxide (DMSO). Slices then were transferred to 0.1 M phosphate buffered saline (NaCl, 150 mM, pH 7.0) and rinsed. The slices were pretreated with triton-X (0.05%) containing Tris buffer (pH 7.0), followed by addition of extravidin-horseradish peroxidase conjugates (buffer: extravidin = 1,000:1). After overnight incubation with extravidin-horseradish peroxidase conjugate, the slices were reacted with diaminobenzidine (0.05%) and hydroxiperoxide (0.03%). The slices were rinsed in Tris buffer and then mounted in glycerol and examined by light microscopy. The drugs used were biocytin, extravidin-horseradish peroxidase conjugate, and diaminobenzidine (all from Sigma Chemical); DMSO (Wako Chemicals); hydroxiperoxide (Mitsubishi Kasei).

The onset potential of the rapid depolarization was measured as the membrane potential at which the extrapolated slopes of the slow depolarization and the rapid depolarization intersect. The peak potential was measured as the membrane potential deflection from the rapid depolarization to the persistent depolarization. The amplitude of the rapid depolarization was measured between the peak potential and the onset potential as described before (Tanaka et al. 1997). When testing the effects of Ca2+-free with Co2+-containing medium on the peak potential of the persistent depolarization, we arbitrarily have chosen to measure the potential 1 min after generating the rapid depolarization as the peak because the persistent depolarization was maintained for 1-2 min when the membrane potential began to recover after reintroduction of oxygen and glucose in test media (Yamamoto et al. 1997). Recovery after reintroduction of oxygen and glucose is defined as follows: no recovery, 30-60 min after reintroduction the membrane potential lay between 0 and -19 mV; complete recovery, the membrane potential was more negative than -60 mV; partial recovery, membrane potential repolarized to a value between -20 and -59 mV (Yamamoto et al. 1997). In most neurons with complete recovery, action potentials and fast excitatory postsynaptic potentials elicited by direct and focal stimulation, respectively, were similar to those observed during the preexposure period, i.e., in normal medium. All quantitative results were expressed as means ± SD. The number of neurons examined was given in parentheses. The one-way ANOVA with Scheffé post hoc comparisons was used to compare data, with P < 0.05 considered significant unless specified otherwise.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

This study was based on intracellular recordings obtained with K acetate-, KCl- or K acetate with biocytin-filled electrodes from 127 CA1 pyramidal neurons of adult rats with stable membrane potentials more negative than -60 mV. The mean resting membrane potential and the apparent input resistance of these neurons were -70 ± 6 mV and 45 ± 14 MOmega (n = 127), respectively.

Responses to superfusion with ischemia-simulating medium

As described previously, deprivation of oxygen and glucose produced a sequence of potential changes consisting of an initial hyperpolarization, a slow depolarization, a rapid depolarization, and a persistent depolarization (Higashi et al. 1990; Tanaka et al. 1997; Yamamoto et al. 1997). All responses were accompanied by decreases in apparent input resistance (Fig. 1A). The peak amplitude and duration of the initial hyperpolarization was 5 ± 2 mV and 2.8 ± 1.0 min (n = 49), respectively. The peak amplitude and duration of the slow depolarization was 7 ± 6 mV and 1.9 ± 0.9 min (n = 49), respectively. At a temperature of 36.5 ± 0.5°C, the latency, amplitude, and peak potential of the rapid depolarization was 6.1 ± 1.5 min, 49 ± 6 mV, and -15 ± 4 mV (n = 49), respectively. When oxygen and glucose were reintroduced to the slice immediately after generating the rapid depolarization, the neuron did not repolarize and the membrane potential reached 0 mV after ~5 min (persistent depolarization). The peak potential of the persistent depolarization was -5 ± 3 mV (n = 49). The tissue slices exposed to the medium deprived of oxygen and glucose for 6-7 min did not recover after returning to control solution. Thirty to 90 min after returning to the normal medium, neurons were difficult or impossible to impale. Even when cells could be impaled successfully, the resting potentials were between -10 and -20 mV, and the input resistances were <3 MOmega (n = 8). This result suggests that the rapid depolarization could trigger the membrane dysfunction.



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Fig. 1. Responses produced by oxygen and glucose deprivation in hippocampal CA1 neurons. In this and subsequent figures, ischemia-simulating medium was applied as indicated between down-arrow  and up-arrow , and in each trace, dotted line indicates the preexposure level of membrane potential, unless specified otherwise. A: changes in membrane potential and apparent input resistance induced by oxygen and glucose deprivation. Preexposure level was -70 mV. Downward deflections are hyperpolarizing potential elicited by anodal current pulses (0.29 nA for 200 ms every 3 s). B: change in membrane current and conductance induced by oxygen and glucose deprivation. In top trace, dotted line indicates the preexposure level of holding current. Preexposure level of holding potential and current were -70 mV and 0.07 nA, respectively. Downward deflections are inward currents elicited by hyperpolarizing voltage steps (16 mV for 200 ms every 3 s). Single-electrode voltage-clamp recordings were terminated and switched to the current-clamp mode after 20 min of the reintroduction of oxygen and glucose. Resting membrane potential subsequently was estimated from the 0 potential level after withdrawal of the electrode from the neuron.

To investigate this further, membrane currents induced by oxygen and glucose deprivation were recorded using single-electrode voltage-clamp techniques with KCl (2 M)-filled electrodes. The membrane potential was held at -70 mV, which was close to the resting membrane potential. Deprivation of oxygen and glucose induced a sequence of currents corresponding to the membrane potential changes observed in current-clamp mode. This consisted of an initial outward current, followed by a slow inward current and a rapid inward current (Fig. 1B). Under relatively adequate voltage-clamp condition, the amplitudes of the initial outward current, the slow inward current, and the rapid inward current were 0.14 ± 0.08 nA, 0.35 ± 0.23 nA, and 1.71 ± 0.39 nA, respectively (n = 7). In addition, the membrane depolarization of 3 ± 1 mV (n = 7) was produced during the generation of the rapid inward current. Reintroduction of oxygen and glucose immediately after generating the rapid inward current progressively reduced and restored the inward current nearly to the preexposure level 20 min after the reintroduction. When voltage-clamp recording was terminated at this time, the membrane potential measured in current-clamp mode was -20 ± 20 mV (n = 7) (Fig. 1B, bottom). Four of seven neurons showed a partial recovery, and the remaining neurons showed no recovery after terminating relatively adequate voltage clamp. The holding currents at -70 mV were 0.00 ± 0.04 nA (n = 7) and 0.48 ± 0.21 nA (n = 7) before ischemic exposure and 20 min after the reintroduction, respectively. Together with the previous finding that the rapid depolarization and the persistent depolarization occur in all layers of CA1 (Tanaka et al. 1997), the results indicate that the space clamp was incomplete even in adequate voltage-clamp condition.

Effects of Ca2+-free with Co2+-containing medium

Tanaka et al. (1997) reported that [Ca2+]i elevates markedly during the rapid depolarization and is sustained with the persistent depolarization. Furthermore, pretreatment with a low Ca2+ (0.25 mM) medium or an inorganic Ca2+ channel blocker (Co2+ or Ni2+)-containing medium inhibits the peak of the persistent depolarization, and the membrane potential completely recovers to the preexposure membrane level after reintroduction of oxygen and glucose (Tanaka et al. 1997; Yamamoto et al. 1997). To identify a period when the membrane function turns into the irreversible change, Ca2+-free (0 mM) with Co2+ (2.5 mM)-containing medium including oxygen and glucose was superfused at various times after the onset of the rapid depolarization. When the Ca2+-free with Co2+-containing medium was superfused within 1 min after the rapid depolarization, neurons showed a complete recovery in 16 of 18 tested, the remaining 2 neurons showing a partial recovery. When the Ca2+-free with Co2+-containing medium was superfused 1 min after the rapid depolarization, the membrane potential was completely recovered in three neurons, partially recovered in four neurons, and one neuron showed no recovery. In contrast, neurons showed either partial or no recovery when the Ca2+-free with Co2+-containing medium was superfused >1.5 min after the rapid depolarization (Fig. 2). These results suggest that the membrane dysfunction may occur or be triggered between 1 and 2 min after the reintroduction of oxygen and glucose, taking into account a time lag for the chamber to fill with the test solution (Ca2+-free with Co2+-containing medium) of ~0.5 min after switching to the test solution.



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Fig. 2. Effects of Ca2+-free with Co2+-containing medium on the persistent depolarization. Downward deflections are hyperpolarizing potentials elicited by anodal current pulses (range 0.2-0.3 nA for 200 ms every 3 s). A: Ca2+-free with Co2+-containing medium was applied with oxygen and glucose immediately after (top), 1 min after (2nd trace), 1.5 min after (3rd trace), or 2 min after (bottom) the rapid depolarization. Note that Ca2+-free with Co2+-containing medium including oxygen and glucose restored the persistent depolarization to the preexposure level when the medium was applied within 1 min after the rapid depolarization. B: effects of Ca2+-free with Co2+-containing medium including oxygen and glucose on percentage of neurons exhibiting recovery. , , and , complete, partial, and no recovery, respectively. Note that majority of neurons tested showed complete recovery when the Ca2+-free with Co2+-containing medium was superfused within 1 min after the rapid depolarization.

Table 1 shows the peak potential of the persistent depolarization, the membrane potential level after recovery, and the apparent input resistances before and after ischemic exposure in the neurons tested. Comparing membrane potentials after recovery in the Ca2+-free with Co2+-containing medium, the recovery membrane potentials after a delay of >1.5 min of introduction of Ca2+-free with Co2+ were significantly more positive than in cells exposed to the same medium immediately after the onset of the rapid depolarization (nondelay). These results suggest that the membrane dysfunction may occur or be triggered ~2 min after the generation of the rapid depolarization.


                              
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Table 1. Effects of Ca2+-free and Co2+ (2.5 mM)-containing medium on the persistent depolarization

Light microscopic aspects for CA1 neurons during and after ischemic exposure

The morphology of CA1 pyramidal neurons, revealed by biocytin, was compared at the various phases of membrane potential changes (before exposure, during the initial hyperpolarization, during the slow depolarization, and immediately after, 1 min after, and 3 min after the rapid depolarization). In the normal medium, biocytin was injected for 1 h by applying hyperpolarizing current pulses (0.1-0.3 nA for 200 ms every 3 s) through the recording electrode. The resting membrane potential and apparent input resistance after injection were not significantly different from those before injection. At the various periods of the potential change produced by oxygen and glucose deprivation, the recording electrode was withdrawn and the slice transferred into fixatives, maneuvers which took 15-20 s. After overnight fixation, the recorded neurons were stained, and the morphological aspects examined under light microscopy.

Figure 3 shows the typical appearance of the cell body and the proximal site of the apical dendrite (top) and the distal dendrites (bottom) of CA1 pyramidal neurons recorded before and during ischemic exposure for 6-9 min. Before application of ischemia-simulating medium (control condition), the cell body and proximal site of the apical dendrite were stained densely, and the surface of these structures appeared smooth (Fig. 3A1). The distal dendrites also were stained densely, and many spines were seen on the surface membrane (Fig. 3A2). The appearance of the cell body and dendrites was not significantly changed in neurons stained during the initial hyperpolarization and the slow depolarization (~2 and 4 min of ischemic exposure, respectively; not shown). In neurons stained immediately after the rapid depolarization (~6 min of exposure), the cell body and proximal site of the apical dendrite were stained poorly as compared with the control condition (Fig. 3B1). Small blebs were observed on the surface membrane of the cell body and proximal site of the apical dendrite; a small, round core stained poorly was surrounded by the relatively dense shell (Fig. 4, black-triangle). The distal dendrites were stained densely but spines were not distinct (Fig. 3B2). One minute after the rapid depolarization (~7 min of exposure), large blebs appeared on the surface membrane of the cell body (Fig. 3C1). The distal dendrites were stained densely, but the appearance was bead-like (Fig. 3C2). Three minutes after (~9 min of exposure), the cell body and proximal site of the apical dendrite were swollen (Fig. 3D1), the distal dendrites were stained poorly, and the stained dendrites were fragmented into pieces (Fig. 3D2). Similar results were observed in another five neurons. In addition, reintroduction of oxygen and glucose immediately after the rapid depolarization showed similar effects in six neurons tested; large blebs appeared 1 min after starting reintroduction and the cell body was swollen 3 min after (Table 2).



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Fig. 3. Effects of ischemia-simulating medium on the appearance of CA1 neurons. Light photomicrograph of CA1 pyramidal neurons stained intracellularly with biocytin after recording. Each photograph was taken from a different neuron. Top: pyramidal cell body and proximal site of the apical dendrite. Bottom: distal site of the apical dendritic shaft and secondary dendrites. A: appearance of the neuron before exposure to ischemia-simulating medium (control condition). B: appearance of the neuron immediately after the rapid depolarization. C: appearance of the neuron 1 min after the rapid depolarization. D: appearance of the neuron 3 min after the rapid depolarization. Scale bar represents 10 µm.



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Fig. 4. High-magnification photograph of the same CA1 pyramidal neuron shown in Fig. 3B. black-triangle, small blebs. Scale bar represents 10 µm.


                              
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Table 2. Parameters of CA1 pyramidal cell appearance before, during, and after in vitro ischemia

Table 2 shows a quantitative analysis of the long and transverse axes, and the cross-sectional cell body area of stained CA1 pyramidal cells before, during, and after ischemic exposure. In neurons with continuous ischemic exposure, the long axis and transverse axis 1 and 3 min after the rapid depolarization were lengthened significantly compared with the control values (P < 0.05 for 1 min and P < 0.005 for 3 min). The cell body area 1 and 3 min after also was increased significantly (P < 0.05 for 1 min and P < 0.005 for 3 min). In contrast, the long axis, transverse axis, and cell body area immediately after the rapid depolarization were not significantly changed. The number of blebs was reduced significantly 1 and 3 min after the rapid depolarization as compared with the appearance of blebs immediately after (P < 0.01) (Table 2). The distribution histograms for the diameter of small and large blebs in the neurons immediately, 1 and 3 min after were different from normal distribution; the histograms were skewed toward large diameters. Therefore the Kruskal-Wallis test and Mann-Whitney test with Bonferroni method were used to analyze and compare the diameter of blebs in neurons recorded up to, immediately after, and 1-3 min after the rapid depolarization. The blebs in neurons observed immediately after the rapid depolarization were significantly smaller than those of 1 and 3 min after (P < 0.001) (Table 2). In neurons with reintroduction of oxygen and glucose, there were also significant differences in long axis, transverse axis, number of blebs, diameter of blebs, and cross-sectional area 3 min after the rapid depolarization but not 1 min after compared with the control values (Table 2).

There is a possibility that cell swelling produced by ischemic exposure is a property common to only impaled neurons. To elucidate effects of the impalement on the morphological change, we compared the neuronal structure between the biocytin-stained neurons in which the recording electrode was withdrawn before ischemic exposure (unimpaled neurons) and the neurons impaled during and after ischemic exposure (impaled neurons). The distance between the neurons was ~300 µm in the same tissue slice. The impaled neuron showed swelling of the cell body and the proximal site of the apical dendrite 3 min after reintroduction of oxygen and glucose, whereas the unimpaled neuron showed the small blebs on the surface of the cell body and the proximal site of the apical dendrite without cell swelling. Thus the long axis and the transverse axis in unimpaled neurons were 40.1 ± 5.4 µm and 17.8 ± 1.7 µm (n = 6), respectively, which were not significantly different from the values of the control neurons before ischemic exposure. The cross-sectional cell body area in unimpaled neurons was 432.7 ± 27.9 µm2 (n = 6), however, not significantly different from the control values. From these results, it is likely that the essential change in membrane structure is similar in impaled and unimpaled neurons, but the cell swelling of unimpaled neurons occurs much later than that of impaled neurons.

Morphological aspects after reintroduction of oxygen and glucose in Ca2+-free with Co2+-containing medium

When oxygen and glucose were reintroduced with the Ca2+free with Co2+-containing medium immediately after the rapid depolarization, a complete recovery of the membrane potential was observed in the majority of neurons (Fig. 2). Morphological aspects of the biocytin-stained neurons were compared before and after ischemic exposure. Figure 5 shows the cell body and proximal apical dendrite (Fig. 5A1), and distal dendrites (Fig. 5A2) of a CA1 pyramidal neuron recorded in control medium and those (Fig. 5B, 1 and 2) in a neuron that showed a complete recovery by superfusion with the Ca2+-free with Co2+-containing medium, also containing oxygen and glucose, immediately after the rapid depolarization. In control conditions, cell bodies and proximal apical dendrites were stained densely, and the surface of these structures was smooth (n = 6, Fig. 5A1). The distal dendrites also were stained densely (n = 6, Fig. 5A2). In contrast, the cell bodies and proximal apical dendrites of neurons after complete recovery in Ca2+-free with Co2+-containing medium were stained poorly compared with those of the control neurons (n = 6, Fig. 5B1). Blebs were not observed on the membrane surface, but the long axis and cell body area were increased significantly (n = 6, P < 0.005, Fig. 5B and Table 2).



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Fig. 5. Morphological appearances of a control neuron and a neuron that recovered after reintroduction of oxygen and glucose in Ca2+-free with Co2+-containing medium. Each photograph was taken from a different neuron. Top: pyramidal cell body and proximal site of the apical dendrite. Bottom: distal site of the apical dendritic shaft and secondary dendrites: A: control condition. B: a typical morphological appearance of neuron 20 min after starting reintroduction of oxygen and glucose in Ca2+-free with Co2+-containing medium (see text). Scale bar represents 10 µm.


    DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Membrane dysfunction produced by ischemia-simulating medium

The present study demonstrates that in the majority of neurons, the persistent depolarization produced by ischemia-simulating medium was decreased significantly and the membrane potential restored to almost the preexposure potential level when a Ca2+-free with Co2+-containing medium was superfused within 1 min after the rapid depolarization. When switching the superfusing media, there was a delay of ~0.5 min before the chamber was filled with the test solution. These results indicate that the irreversible membrane dysfunction may occur or be triggered later than 1.5 min after the rapid depolarization. In contrast, the neurons showed only a partial recovery when the Ca2+-free with Co2+-containing medium was superfused 1.5-2 min after the rapid depolarization, suggesting that the irreversible membrane damage may be complete later than 2 min after the onset of the rapid depolarization.

We have reported previously that a sustained increase in [Ca2+]i has a dominant role in causing irreversible changes in membrane function (Yamamoto et al. 1997). It is therefore likely that reduction in Ca2+ influx into the neuron in the Ca2+-free with Co2+-containing medium is one of the factors that allows the membrane potential to be restored to the preexposure level. Under current-clamp condition, the membrane potential does not recover when oxygen and glucose were reintroduced immediately after the rapid depolarization (Tanaka et al. 1997; Yamamoto et al. 1997). The present voltage-clamp study revealed that the membrane clamped adequately was restored partially after the reintroduction of oxygen and glucose in four of seven neurons and not recovered in the remaining three neurons, suggesting that the membrane depolarization (the persistent depolarization) has an important role in the induction of membrane dysfunction. The rapid inward current, which was sustained <1 min, was followed by the persistent inward current for 5 min after reintroduction of oxygen and glucose, suggesting that a sustained increase in [Ca2+]i during the persistent depolarization triggers the irreversible membrane dysfunction.

Pretreatment with NMDA receptor antagonists or a non-NMDA receptor antagonist inhibits the persistent depolarization and allows the membrane potential to recover when oxygen and glucose have been reintroduced immediately after generating the rapid depolarization (Rader and Lanthorn 1989; Tanaka et al. 1997; Yamamoto et al. 1997). Inorganic Ca2+ channel blockers, reduction in external Ca2+, an inhibitor of Ca2+-induced Ca2+ release from store sites, or an acetoxymethyl ester compound of Ca2+ chelator have similar effects, but organic Ca2+ channel blockers do not (Yamamoto et al. 1997). These results suggested that the Ca2+ influx into the neuron during the persistent depolarization is due to the leaky membrane, activation of ionotropic-glutamate receptor channels as well as Ca2+ release from internal stores but is not due to the influx of Ca2+ via L- and T-type voltage-gated Ca2+ channels, as reported before (Yamamoto et al. 1997). It is therefore likely that Ca2+ influx during the persistent depolarization, which removes the voltage-dependent Mg2+ block in NMDA receptor channels and activates voltage-independent R-type Ca2+ channels, may cause the irreversible membrane dysfunction. Nevertheless, Ebine et al. (1994) have reported that even in Ca2+-free with Ca2+ chelator-containing solution, deprivation of oxygen and glucose causes an excessive increase in [Ca2+]i. The peak of the persistent depolarization was not affected by superfusion with the Ca2+-free with Co2+-containing medium, suggesting the possibility that Ca2+ release from internal stores may play a major role in the increase in [Ca2+]i during the early phase of the persistent depolarization.

The present study demonstrates that the morphological appearance of biocytin-stained neurons before the rapid depolarization were not significantly different from the control neurons, whereas biocytin-stained neurons immediately after the rapid depolarization showed small blebs on the cell soma and the proximal site of the apical dendrite. These blebs increased in diameter while decreasing in number and transformed to larger blebs 1 min after the end of the rapid depolarization. Taking into account that it took ~20 s for the slice to be transferred into fixatives, it is likely that the transformation from small blebs to large blebs occurred later than 1.5 min after the end of the rapid depolarization. Finally, neurons became swollen and membrane dysfunction occurred 3 min after the rapid depolarization.

The generation of the rapid depolarization was correlated to the formation of the small blebs because small blebs were found only immediately after the rapid depolarization. The neurons immediately after the rapid depolarization always showed a weak staining in comparison with the control neurons. This weak staining of the neurons suggests that the formation of small blebs may induce leakage of biocytin (M. W. 372) from the neurons. It is, however, possible that the weak staining may have resulted from the cell swelling. The long axis, transverse axis, and cell body area of the neurons were not significantly changed immediately after the rapid depolarization as compared with the control neurons, suggesting that the contribution made by expansion of the cell volume to the weak staining is, at least, minimal. These results support the idea that the formation of micro-pores in the small blebs may generate the rapid depolarization because the rapid depolarization is voltage independent and is due to a nonselective increase in permeability to all participating ions, which probably occurs only in pathological conditions (Tanaka et al. 1997). In mouse hippocampal CA1 neurons, the pH-sensitive fluorescent dye, 2',7'-bis(carboxyethyl)-carboxyfluorescein (BCECF, M. W. 520), is leaked rapidly after 5-7 min application of ischemia-simulating medium (Fujiwara et al. 1992). On the other hand, fluorescence of Fura-2 (M. W. 641) induced by excitation at both 340 and 380 nm during ischemic exposure does not diminish after generating the rapid depolarization (Tanaka et al. 1997). Taken together, these results suggest that the formation of micro-pores may underlie the generation of the rapid depolarization, and the newly formed pores may provide a path for molecules with molecular weight <520.

Plasma membrane blebbing is a phenomenon associated with toxic and ischemic cell injury in hepatocytes and mouse embryo cells. An increase in [Ca2+]i concentration causes the dissociation of actin microfilaments from alpha -actinin, which associates microfilaments with actin-binding proteins in the plasma membrane. In addition, increased [Ca2+]i activates proteases that cleave actin-binding proteins, eliminating the plasma membrane anchor for the cytoskeleton. The formation of the weakened membrane, where the cytoskeleton has dissociated from the plasma membrane, may lead to the production of surface blebs (Gores et al. 1990; Orrenius et al. 1989). In hippocampal CA1 neurons, the [Ca2+]i begins to increase slowly 1 min after starting superfusion of ischemia-simulating medium (Tanaka et al. 1997). In dissociated rat hippocampal CA1 neurons, concurrent with the increase in [Ca2+]i with anoxia, small blebs appear on the dendrites, which then increase in diameter (Friedman and Haddad 1993). Furthermore in gerbil hippocampal CA1 region, transient forebrain ischemia is followed within 15 min by accelerated proteolysis of the cytoskeletal protein spectrin (Seubert et al. 1989). It is therefore, likely that the formation of small blebs may be the result of the proteolysis of actin-binding proteins or the dissociation of actin microfilaments from alpha -actinin, which is induced by the increase in [Ca2+]i.

The temperature coefficient of latency and maximal slope of the rapid depolarization is 2.5 and 2.9, respectively, and these values are similar to that of many enzyme mechanisms (Onitsuka et al. 1998). This may support the idea that the formation of small blebs may underlie the generation of micro-pores and the rapid depolarization. However, it is possible that the increase in voltage-independent and nonselective ion permeability during the rapid depolarization may induce the formation of the small blebs. It is difficult to elucidate whether the formation of small blebs induces the rapid depolarization or vice versa because there is approximately a 20-s delay to fix the slice that contained the recorded neuron.

The biocytin-stained neurons, in which the recording electrode was withdrawn before ischemic exposure, showed small blebs on the surface of cell body and proximal sites of the apical dendrite 3 min after the end of the rapid depolarization, suggesting that the intracellular recordings may accelerate the degeneration of the neuron membrane. The time course of degeneration in the neuron membrane that was not impaled by an electrode may be slower than that of the impaled neurons. It was, however, difficult to impale CA1 pyramidal neurons after reintroduction of oxygen and glucose, suggesting that there is no functional recovery even in the unimpaled CA1 neurons.

In rodent hippocampal CA1 neurons, microvacuoles and/or swollen mitochondria with disintegration of internal cristae are observed after ischemia (occlusion of carotid artery) for 5-30 min (Petito and Pulsinelli 1984; Yamamoto et al. 1986, 1990; also see Schmidt-Kastner and Freund 1991). These microvacuoles and/or swollen mitochondria are found in the cell soma or dendrites. Similarly in hepatocytes, viewed by phase contrast microscopy, blebs appear as bubble-like projections extending from the cell surface, whereas mitochondria, lysosomes, Golgi apparatus, and peroxisomes are excluded from the blebs, which accounts for their phase-lucent appearance (Herman et al. 1988; Lemasters et al. 1987; also see Gores et al. 1990). On the other hand, the present study showed that the small blebs were found on the membrane surface, suggesting that the small blebs may not consist of vacuoles or swollen mitochondria.

Functional membrane recovery from the reversible damage

After the recovery of the membrane potential in the Ca2+-free with Co2+-containing medium, blebs were not observed on the cell surface, and the transverse axis was not different from the control neurons, whereas the long axis was lengthened and the cross-sectional cell body area was increased in the recovered neurons. Thus the apex of the pyramidal cell body and the proximal site of the apical dendrite appear susceptible to ischemic exposure.

In conclusion, in vitro ischemia causes irreversible membrane dysfunction during the persistent depolarization. The membrane dysfunction may be a Ca2+-dependent process that starts 1.5 min after and may be complete 3 min after the rapid depolarization. Morphologically, small blebs appear immediately after, and the transformation from small blebs to large blebs occurs 1.5-2 min after the onset of rapid depolarization. Finally, cell swelling occurs 3 min after the onset of rapid depolarization. These results suggest that the irreversible membrane dysfunction may involve the transformation of small blebs to large blebs.


    ACKNOWLEDGMENTS

We thank Dr. D. C. Spanswick for comments and suggestions on the manuscript.

This work was supported in part by a grant-in-aid for Scientific Research of Japan, an Ishibashi Foundation grant, and an Ichiro Kanehara Foundation grant.


    FOOTNOTES

Address reprint requests to E. Tanaka.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 8 September 1998; accepted in final form 1 December 1998.


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