Department of Physiology, Kurume University School of Medicine, 67 Asahi-machi, Kurume 830-0011, Japan
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ABSTRACT |
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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.
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INTRODUCTION |
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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
).
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METHODS |
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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
M. 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.
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RESULTS |
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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 M
(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 M
(n = 8). This result suggests that the rapid
depolarization could trigger the membrane dysfunction.
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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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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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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, ). 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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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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DISCUSSION |
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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 -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
-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.
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ACKNOWLEDGMENTS |
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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.
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FOOTNOTES |
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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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REFERENCES |
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