Department of Cell Biology, Duke University Medical Center, Durham, North Carolina 27710
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ABSTRACT |
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Müller, Michael and
George G. Somjen.
Na+ and K+ Concentrations, Extra- and
Intracellular Voltages, and the Effect of TTX in Hypoxic Rat
Hippocampal Slices.
J. Neurophysiol. 83: 735-745, 2000.
Severe hypoxia causes rapid depolarization
of CA1 neurons and glial cells that resembles spreading depression
(SD). In brain slices in vitro, the SD-like depolarization and the
associated irreversible loss of function can be postponed, but not
prevented, by blockade of Na+ currents by tetrodotoxin
(TTX). To investigate the role of Na+ flux, we made
recordings from the CA1 region in hippocampal slices in the presence
and absence of TTX. We measured membrane changes in single CA1
pyramidal neurons simultaneously with extracellular DC potential
(Vo) and either extracellular
[K+] or [Na+]; alternatively, we
simultaneously recorded [Na+]o,
[K+]o, and Vo.
Confirming previous reports, early during hypoxia, before SD onset,
[K+]o began to rise, whereas
[Na+]o still remained normal and
Vo showed a slight, gradual, negative shift;
neurons first hyperpolarized and then began to gradually depolarize.
The SD-like abrupt negative Vo
corresponded to a near complete depolarization of pyramidal neurons and
an 89% decrease in input resistance. [K+]o
increased by 47 mM and [Na+]o dropped by 91 mM. Changes in intracellular Na+ and K+
concentrations, estimated on the basis of the measured extracellular ion levels and the relative volume fractions of the neuronal, glial,
and extracellular compartment, were much more moderate. Because
[Na+]o dropped more than
[K+]o increased, simple exchange of
Na+ for K+ cannot account for these ionic
changes. The apparent imbalance of charge could be made up by
Cl
influx into neurons paralleling Na+ flux
and release of Mg2+ from cells. The hypoxia-induced changes
in interneurons resembled those observed in pyramidal neurons.
Astrocytes responded with an initial slow depolarization as
[K+]o rose. It was followed by a rapid but
incomplete depolarization as soon as SD occurred, which could be
accounted for by the reduced ratio,
[K+]i/[K+]o. TTX (1 µM) markedly postponed SD, but the SD-related changes in
[K+]o and [Na+]o
were only reduced by 23 and 12%, respectively. In TTX-treated pyramidal neurons, the delayed SD-like depolarization took off from a
more positive level, but the final depolarized intracellular potential
and input resistance were not different from control. We conclude that
TTX-sensitive channels mediate only a fraction of the Na+
influx, and that some of the K+ is released in exchange for
Na+. Even though TTX-sensitive Na+ currents are
not essential for the self-regenerative membrane changes during hypoxic
SD, in control solutions their activation may trigger the transition
from gradual to rapid depolarization of neurons, thereby synchronizing
the SD-like event.
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INTRODUCTION |
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Acute severe hypoxia of forebrain gray matter induces
self-regenerating rapid depolarization of neurons that closely
resembles spreading depression (Leão 1947).
Spreading depression (SD) is characterized by loss of neuronal
activity, nearly complete depolarization of neurons and glial cells,
and substantial disturbance of the ionic distribution (Grafstein
1956
; Marshall 1959
). During both normoxic SD
and hypoxic SD-like depolarization, the extracellular concentrations of
the major inorganic cations and anions are shifted far beyond the
physiological range, greatly in excess of the ionic changes associated
with tetanic stimulation or seizures (Hansen 1985
;
Nicholson 1984
; Nicholson and Kraig
1981
). From the outset, the question has been raised whether SD
represents a breakdown of the normal selective permeability of neuron
membranes or whether it is produced by the abnormal operation of
physiological ion channels. This basic question has not been
satisfactorily answered.
SD can be provoked even when action potentials and synapses are blocked
by tetrodotoxin (TTX) (Sugaya et al. 1978;
Tobiasz and Nicholson 1982
). Similarly, hypoxic SD-like
depolarization is not usually blocked by TTX, but its onset is delayed
considerably and, in a minority of cases, prevented by the drug
(Aitken et al. 1991
; Xie et al. 1994
).
Yet several observations suggest that SD is not the consequence of
membrane breakdown or the complete loss of membrane resistance
(Czéh et al. 1993
; Müller and Somjen 1998
; Phillips and Nicholson 1979
). Moreover, in
a previous study (Müller and Somjen 1998
), we
demonstrated that hypoxic SD can be prevented completely if
voltage-sensitive Na+ and
Ca2+ channels as well as AMPA/kainate and
N-methyl-D-aspartate (NMDA) glutamate receptors
are all blocked. A role for physiological channels also is suggested by
the fact that, in current-clamp recordings from CA1 pyramidal neurons,
the hypoxic SD-like depolarization is triggered at an apparent
threshold potential of approximately
52 mV, which is almost identical
to the Na+ spike threshold of
53 mV reported by
Dingledine (1983)
.
These observations reopened the question, what role, if any, is played by voltage-gated Na+ channels in the evolution of SD? The main purpose of the present study was to answer this question. To this end, we obtained current-clamp recordings from CA1 pyramidal neurons, interneurons, and glial cells during severe hypoxia while extracellular potential and either [K+]o or [Na+]o also were monitored. Unlike in previous studies, these parameters were not measured one by one but were monitored simultaneously in the same spot of a given slice. In other trials, we used triple-barreled microelectrodes, simultaneously measuring extracellular potential, [K+]o and [Na+]o in a single point to resolve the time course and interrelationship of these parameters. These experiments first were performed in control slices and then repeated after application of TTX, revealing the TTX sensitivity of each of the measured variables.
Parts of this study have been published in abstract form
(Müller and Somjen 1999a,b
).
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METHODS |
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Preparation
Hippocampal tissue slices were prepared from ether-anesthetized,
male Sprague-Dawley rats of 119-265 g body wt (4-7 wk old). After
decapitation, the brain was removed rapidly from the skull and placed
in chilled artificial cerebrospinal fluid (ACSF) for 1-2 min. The two
hemispheres were separated, one hippocampus was isolated, and
transverse slices of 400-µm thickness were cut using a tissue
chopper. Slices were transferred to an interface recording chamber of
the Oslo style and were left undisturbed for 90 min. The recording
chamber was kept at a temperature of 34.5-35.5°C. It was aerated
continuously with 95% O2-5%
CO2 (400 ml/min), and perfused with oxygenated
ACSF (1.5 ml/min). Hypoxia was induced by switching the chamber's gas
supply to 95% N2-5% CO2.
To protect the slices from drying out and to prevent oxygenation from
the air during hypoxic episodes, the slice chamber was covered by a lid
with a small (2 cm2) opening for the positioning
of the electrodes. Exchange of the bathing solution and diffusion of
applied drugs into the slice took ~15 min.
Solutions
The ACSF had the following composition (in mM): 130 NaCl, 3.5 KCl, 1.25 NaH2PO4, 24 NaHCO3, 1.2 CaCl2, 1.2 MgSO4, and 10 dextrose; aerated with 95% O2-5% CO2 to adjust pH to 7.4. TTX (tetrodotoxin, citrate buffered, Calbiochem and Sigma) was prepared as 1 mM stock solution in distilled water and was kept frozen.
Microelectrodes
Single-barreled glass microelectrodes for extracellular
recordings were pulled from thin-walled borosilicate glass (TW150F-4, WPI) using a horizontal puller (P-80/PC, Flaming Brown). They were
filled with ACSF, and their tips were broken to a final resistance of
5-10 M. Sharp microelectrodes for current-clamp recordings were
made from thick-walled borosilicate glass (1B150F-4, WPI) and filled
with 2 M K-Acetate +5 mM KCl +10 mM
N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES; Sigma); pH 7.4. Their resistances were 60-80 M
.
Extracellular Na+ and K+ concentrations were measured using double-barreled Na+-or K+-sensitive microelectrodes. They were pulled from theta type capillaries (GCT 200-10, Clark Electromedical Instruments) on the horizontal puller. The designated ion-sensitive barrel was silanized by 60-min exposure to HMDS vapors (hexamethyldisilazane, 98%, Fluka; vaporized at 40°C) and subsequent baking in the oven (200°C, 2 h). Silanization of the reference barrel was prevented by perfusing it with compressed air (1.5 bar).
The tip of the K+-sensitive barrel was filled
with the valinomycin-based K+ ion neutral carrier
(Potassium Ionophore I -Cocktail A, Fluka 60031) and backfilled with
150 mM KCl +10 mM HEPES, pH 7.4. The reference barrel contained 150 mM
NaCl +10 mM HEPES, pH 7.4. Mean electrode resistances of the reference
and ion-sensitive barrel were 20-40 and 80-110 M, respectively.
The widely used ETH 157 Na+ neutral carrier
(Sodium Ionophore II Cocktail A, Fluka 71178) was found to be unsuited
for extracellular Na+ measurements during SD, due
to its high K+ sensitivity. Because this
Na+ carrier is only 2.5 times more sensitive to
Na+ than to K+ (log
KNaKpot = 0.4), the electrode response strongly
deviated from linearity in the range where
[Na+]o was expected to be
low and [K+]o high, and
the constructed electrodes exhibited an average detection limit of 23 mM Na+. We therefore prepared a
Na+-sensitive cocktail based on the more
selective Na+ Ionophore VI (Fluka 71739), which
is 100 times more sensitive to Na+ than to
K+ (log KNaKpot =
2.0).
According to Deitmer and Munsch (1995)
the
Na+ ionophore VI, the organic solvent
2-nitrophenyl octyl ether (Fluka 73732) and the lipophilic salt
potassium tetraphenylborate (Fluka 72018) were mixed at the weight
percent ratio 10.0:89.5:0.5, respectively.
Ion-sensitive electrodes were calibrated before and after each experiment by detecting their response generated in standard solutions (0, 1, 2, 5, 10, 20, 50, and 100 mM K+ for K+-sensitive electrodes and 150, 100, 50, 20, 10, 5, 1, and 0 mM Na+ for Na+-sensitive electrodes). To maintain constant ionic strength similar to that in interstitial fluid, Na+ in the calibration solutions was replaced by K+ and vice versa (reciprocal calibration method). Average slopes of the K+- and Na+-sensitive electrodes were 54.6 ± 2.9 mV/decade K+ and 50.8 ± 3.6 mV/decade Na+; their detection limits were 0.25 ± 0.23 mM K+ (n = 11) and 3.7 ± 0.3 mM Na+ (n = 20).
Occasionally we used triple-barreled microelectrodes of the twisted
type for simultaneous Na+ and
K+ recordings. A single-barreled capillary
(1B150F-4, WPI) was glued to the double-barreled theta-type capillary
(GCT 200-10, Clark Electromedical Instruments) using a slow-setting
two component epoxy glue (Poxy Pro, Power Poxy Adhesives). To achieve
maximum adhesion strength, the glue was hardened by placing the
capillary tubes in the oven (60°C, 1 h). The capillaries then
were pulled on a vertical puller (Narishige PE-2). In a first step, the
glass was melted and the capillaries were pulled by only 2-4 mm,
simultaneously twisting the attached reference barrel by 180° around
the centered theta capillary. After cooling, capillaries were pulled
apart in a second pulling step. Both barrels of the theta capillary were silanized by exposure to HMDS vapors and subsequent baking in the
oven. Silanization of the attached reference barrel was prevented by
filling it with distilled water. Fillings of the Na+- and K+-sensitive
barrels were the same as for the double-barreled electrodes and the
reference barrel contained 150 mM NaCl +10 mM HEPES, pH 7.4. Electrode
resistances of the Na+-sensitive,
K+-sensitive, and reference barrels were 200, 130, and 40 M, respectively. Electrode slopes and detection limits
were comparable with those of the double barreled electrodes. There was
no noticeable interference between adjacent barrels.
Electrical recordings
Ion-sensitive electrode signals were referred to an Ag/AgCl bridge electrode embedded in 2% agar in 3 M KCl. They were recorded by a DC amplifier (constructed locally) and digitized by a TL-1/Lab Master acquisition system at sampling rates of 25 Hz ([Na+]o, [K+]o measurements only) or 1 kHz (combined current-clamp and [Na+]o, [K+]o measurements). Because electrodes were calibrated to Na+ and/or K+ concentrations and the activity coefficient of the measured ion was held constant, changes in [Na+]o and [K+]o could be calculated directly from the electrode responses using the electrodes' averaged slope of pre- and postexperiment calibration.
All signal amplitudes were measured between the prehypoxia baseline and
the maximal change. Only rapid negative extracellular DC potential
changes (Vo) of
10 mV amplitude
were considered as SD. SD onset was defined as occurrence of the sudden
Vo.
Current-clamp recordings from CA1 pyramidal neurons were performed with
an intracellular recording amplifier (Neuro Data, IR-283). Bridge
balance and electrode-capacitance compensation were adjusted before
insertion of the electrode and continuously controlled during the
entire recording. CA1 pyramidal neurons were identified by their
location in stratum pyramidale, membrane potential, spontaneous
activity, action-potential shape, and input resistance (Morin et
al. 1996). Interneurons were identified by their location in
st. radiatum close to the pyramidal cell layer, their nonadaptive spike
firing in response to depolarizing stimuli, prominent
spike-afterhyperpolarizations as well as a pronounced inward
rectification during hyperpolarizing current pulses (Morin et
al. 1996
). Astrocytes were identified by their location in st.
radiatum lining the pyramidal cell layer (D'Ambrosio et al. 1998
), very negative membrane potential, low input resistance, and absence of spike discharges in response to depolarizing stimuli. Successful cell impalement was achieved by slowly advancing the electrode into the slice and applying a brief high-frequency
oscillating AC signal to the electrode. Cell recovery was facilitated
by injecting a hyperpolarizing current for the first few minutes after
impalement. Only neurons with a stable membrane potential of at least
55 mV and glial cells with a membrane potential of at least
75 mV were accepted. Neuronal input resistance was determined every 10 s
by injecting a hyperpolarizing current of 400-pA amplitude and 200-ms
duration. Data were sampled at 1 kHz using the TL-1/Labmaster acquisition system and the Axotape V2 software (Axon Instruments). Input resistances were measured at the steady state level of the voltage deflections and averaged over 10 successive current injections. Changes in input resistance were expressed as a percent of pretreatment value.
Estimates of intracellular Na+ and K+ changes
On the basis of the measured
[Na+]o and
[K+]o changes during SD,
we calculated the intracellular ion changes. These estimates were based
on an averaged interstitial volume fraction (ISVF) of 0.15 (McBain et al. 1990; Pérez Pinzón et
al. 1995
) and intracellular volume fractions (ICVF) of 0.45 for
neurons and 0.40 for glial cells, based on Kuffler and Nicholls
(1966)
much-quoted figure of glial cells occupying
50% of
the cellular space. The fraction of glial volume may in hippocampus be
<50% (see e.g., Wolff 1966
; reviewed by Somjen
1975
). Also the ISVF at rest may actually be >15%
(Mazel et al. 1998
), whereas during SD it shrinks to a
much smaller size (Jing et al. 1994
;
Pérez-Pinzón et al. 1995
). For now, the
calculations are only intended to show limits; the true intracellular
changes are probably within the calculated range. The intracellular ion
changes (
[X]i) were calculated
according to the formula
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Statistics
The data were obtained from 32 rats, and because most experiments did not last longer than 2 h, up to four slices could be used from each brain. All numerical values are represented as means ± standard deviations. Significance of the observed changes was tested using a two-tailed, unpaired Student's t-test and a significance level of 5%. In the diagrams, significant changes are marked by asterisks (*P < 0.05; **P < 0.01). Statistical calculations and linear regressions were done with the Excel 7.0 or QuattroPro 3.0 software.
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RESULTS |
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Interrelationship of pyramidal neuron membrane properties, Vo, [Na+]o, and [K+]o
To investigate the interrelationship of intra- and extracellular voltage changes and extracellular ion concentrations during hypoxic SD, we recorded the intracellular potential of CA1 pyramidal neurons and measured Vo combined with either [Na+]o or [K+]o in the extracellular space of st. pyramidale close to the impaled neuron. The extracellular ion changes are summarized in Table 1.
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The mean resting membrane potential of pyramidal neurons was
62.6 ± 5.0 mV and their input resistance averaged 39.0 ± 7.3 M
(n = 22). Oxygen withdrawal almost immediately
caused an initial hyperpolarization of 4.3 ± 2.5 mV and a
decrease of the input resistance by 37.8 ± 14.8% (Figs.
1 and 2).
The initial hyperpolarization abolished spontaneous activity within the
first 30 s of hypoxia. At the same time
[K+]o increased slowly,
which suggests release of K+ from neurons via the
activated K+ channels that generated the
hyperpolarization (Hansen et al. 1982
) (Fig. 1). After
~1 min, the initial hyperpolarization turned into a gradual
depolarization, whereas input resistance remained low and
[K+]o continued to
increase. During this initial phase of hypoxia, [Na+]o remained unchanged
(Fig. 2).
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Within 1.8 ± 0.5 min of hypoxia the intracellular potential
(Vi) depolarized to 51.6 ± 4.3 mV (n = 22), the apparent threshold potential at which
SD started. This apparent SD threshold is almost identical to the
Na+ spike threshold of
53 mV (Dingledine
1983
). At the onset of SD
[K+]o averaged 9.0 ± 3.6 mM (n = 10), which is below the so called "K+ ceiling level," a limit that is not
exceeded during tetanic stimulation or even seizures (Heinemann
and Lux 1977
; Nicholson 1984
).
Immediately before SD onset, indicated by the sudden
Vo, pyramidal neurons discharged
multiple spikes and then their Vi
rapidly rose to
23.2 ± 4.9 mV followed by a more gradual
depolarization to a final level of
7.8 ± 5.0 mV. The input
resistance decreased by 88.5 ± 11.9% (n = 22).
The SD-related sudden negative
Vo
averaged
17.8 ± 4.5 mV at the same time
[Na+]o dropped from its
155 mM baseline to 64.3 ± 13.3 mM (n = 15, Fig.
2) and the already elevated
[K+]o rapidly increased
further to 50.9 ± 28.9 mM (n = 10, Fig. 1). In
all slices, the negative
Vo reached
its maximum amplitude somewhat earlier than the extracellular ion
concentrations; this may be due to the slower response time of
ion-sensitive electrodes. Both
[Na+]o and
[K+]o began to recover
already during hypoxia but at different rates, [Na+]o moving faster
toward its baseline than
[K+]o and reaching a
plateau level of 92.1 ± 8.7 mM (n = 15). By contrast, neither Vi nor the input
resistance recovered before oxygen was readmitted (Figs. 1 and 2).
When reoxygenation was started ~100 s after SD onset, the extracellular ion concentrations immediately started to recover, whereas the recovery of Vi and input resistance was somewhat delayed. In fact, in most pyramidal neurons the depolarization continued briefly before repolarization began (Figs. 1 and 2). During reoxygenation the membrane potential usually became more negative and the input resistance higher than before hypoxia. Spontaneous action potentials returned within 3-5 min. [Na+]o returned to its prehypoxic baseline, whereas [K+]o consistently undershot its 3.5 mM baseline, reaching 1.8 ± 0.6 mM (n = 10; Fig. 1).
Hypoxia-induced membrane changes in interneurons and glial cells
Occasionally we also recorded from current-clamped interneurons
(n = 11) and astrocytes (n = 6), which
were identified according to their location in st. radiatum and their
electrophysiological properties (see METHODS). The membrane
changes induced by hypoxia in different cell types are compared in
Table 2. St. radiatum interneurons had a
somewhat more negative membrane potential and a lower input resistance
than pyramidal neurons. The hypoxia-induced membrane changes were,
however, not noticeably different from those observed in pyramidal
neurons. Interneurons also showed an initial hyperpolarization parallel
to the increasing [K+]o,
and they responded with a near complete depolarization and drastically
reduced input resistance as soon as the hypoxic
Vo occurred (Fig.
3A).
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Glial cells (astrocytes) had the most negative membrane potential
(Table 2) and their input resistance was less than a third of that of
neurons (see also Leblond and Krnjevíc 1989).
After oxygen withdrawal, they immediately started to depolarize without the initial hyperpolarization seen in neurons (Fig. 3B). The
initial depolarization coincided with the increase in
[K+]o. As soon as the
hypoxic
Vo occurred, glial cells
also responded with a sudden depolarization. The amplitude of the
SD-like depolarization in glial cells was not different from that in
neurons (Table 2), but because glial cells started from more negative
resting potentials, they retained a more negative intracellular
potential, averaging
28.2 ± 2.8 mV (n = 6) at
the height of SD (Fig. 3B).
Because of the low glial input resistance, we were not able to
measure its changes during hypoxia. Successful glial cell impalement required electrode resistances in excess of 70 M, and current pulses
of 1.6-2.0 µA were required to evoke 10- to 15-mV membrane potential
deflections. Because electrode resistances were much higher than cell
input resistances and tended to change after cell impalement, a stable
bridge adjustment could not be maintained. From the few trials it
appeared, however, that during hypoxic SD the resistance changes in
glial cells are more moderate than those in neurons.
Estimate of intracellular ion changes and electromotive forces
Because the intracellular potential,
Vi, was recorded in reference to bath
("ground") potential, the measured change in voltage was the sum of
the shifts of the true membrane potential,
Vm, plus the extracellular potential
shift, Vo. The difference between Vi and
Vm becomes maximal at the height of
SD. Because the extracellular DC potential does not markedly change
during the initial phase of hypoxia, before SD onset, the deviation,
however, should be negligible until the rapid depolarization and the
negative DC shift start to build up. To estimate
Vm at the height of hypoxic SD,
Vi must be corrected for
Vo. Figure
4 shows the effect of this correction.
During SD Vm approached but did not
quite reach 0 mV. Also the corrected SD-related depolarization has a
more "flat top" than in the raw recordings of
Vi, indicating that the slow
continuing positive shift of Vi (Figs.
1 and 2) is, in part at least, an artifact caused by contamination by
Vo.
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We estimated the changes in intracellular ion concentrations
based on the measured extracellular ion changes and the relative volume
fractions (see METHODS). We assumed "resting"
intracellular concentrations of 10 mM Na+
(Rose and Ransom 1997a,b
) and 140 mM
K+. If during SD Na+ influx
and K+ release occurs only in neurons, then the
calculated [Na+]i
increased to 41.0 ± 5.0 mM (n = 9) and
[K+]i dropped to
125.6 ± 5.5 mM (n = 9). These changes would be
more moderate if both neurons and glial cells were involved equally (Fig. 4). In this case,
[Na+]i would increase to
26.4 ± 2.6 mM (n = 9), whereas
[K+]i would drop to
132.4 ± 2.9 mM (n = 9). If cell swelling is taken into account, then the computed changes of intracellular ion
concentrations are even smaller by ~5%.
From the measured change in
[K+]o and the estimated
change in [K+]i, the
K+ equilibrium potential
(EK) during SD is calculated to shift
from 97.7 to
27.8 ± 10.8 mV, whereas
ENa shifts from 72.7 to 9.8 ± 10.2 mV (n = 9), if we assume that ions flow only into
and out of neurons. The assumption that neurons and glial cells
participate equally in ion fluxes yields an
EK of
29.2 ± 10.3 mV and an
ENa of 21.4 ± 9.7 mV.
Simultaneous recordings of Vo, [Na+]o, and [K+]o in control and in TTX-treated slices
In previous studies, extracellular Na+ and
K+ usually were recorded separately in different
slices. We now used triple-barreled microelectrodes to simultaneously
measure Vo,
[Na+]o, and
[K+]o from one point in
st. radiatum in a slice to resolve the magnitude, time course,
interrelationship, and TTX sensitivity of these variables. As in st.
pyramidale, the onsets of the sharp
[K+]o increase, the
sudden [Na+]o drop, and
the Vo coincided (Fig.
5). The magnitude of the ionic changes in
st. radiatum did not differ from those measured in st. pyramidale
(Table 1). With the restricted time resolution, inevitable for
ion-sensitive microelectrodes, a noticeable time delay between the
onsets of the two ion fluxes was not observed. Both ionic changes
reached their maximum shortly after SD onset and then began to return
toward their normal level,
[Na+]o more rapidly than
[K+]o. The changes in
[Na+]o were, however,
consistently greater than those in
[K+]o, resulting in an
apparent imbalance in cation concentrations averaging 44.3 ± 14.0 mM (n = 9).
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After the slices recovered from the first (control) hypoxic SD, we
applied 1 µM TTX for 30 min to ensure complete inhibition of
TTX-sensitive Na+ channels, before the second
hypoxic SD was induced. In a previous study (Müller and
Somjen 1998
), we already demonstrated that repetitive induction
of hypoxic SD in a single slice kept under control conditions did not
induce any significant changes in duration, onset and amplitude of the
extracellular DC potential shift
(
Vo) for the first four hypoxic SDs.
TTX markedly delayed the onset of hypoxic SD but only moderately damped
the Vo amplitude and the changes in
[Na+]o and
[K+]o (Fig. 5). After
30-min application of 1 µM TTX, the amplitude of the hypoxic
Vo decreased to 78.0 ± 7.9%
of control and its onset was delayed by 143.9 ± 57.0%
(n = 8).
TTX reduced the drop in
[Na+]o during SD only
slightly (Fig. 5). In contrast to control slices,
[Na+]o began to decrease
slightly already before SD onset. In the presence of TTX SD was
delayed, whereas [K+]o
continued its slow, pre-SD rise. Therefore at the time of SD onset,
[K+]o now averaged
11.6 ± 2.4 mM, which is higher than normally but still below the
K+ ceiling level (Heinemann and Lux
1977; Nicholson 1984
). The sharp [K+]o increase coinciding
with SD, however, was damped by 23.1 ± 20.2%, whereas the
undershoot of the K+ baseline after reoxygenation
was not significantly affected. The sudden drop in
[Na+]o coinciding with SD
onset decreased by only 12.2 ± 8.9%, and the plateau level
decreased by 18.3 ± 17.0% (n = 8).
Effects of TTX on the SD-like depolarization in single pyramidal neurons
As TTX partially reduced the amplitude of the hypoxic
Vo, we asked whether it also
reduced the amplitude of the SD-like depolarization in single cells.
Because stable impalement could usually not be maintained for >45 min
and recordings often were disrupted during reoxygenation, hypoxia under
control conditions and in the presence of TTX could not be investigated
in the same cells. We therefore compared hypoxia-induced membrane
changes during TTX administration to the changes observed in untreated
control slices ("control group"). TTX was administered for
30 min
before withdrawing oxygen.
As expected, application of 1 µM TTX blocked spontaneous impulse
firing. The membrane potentials of TTX-treated pyramidal neurons was 6 mV more negative than that of control cells and their input resistance
was 22% lower (see also Fung and Haddad 1997). When
oxygen was withdrawn in the presence of TTX, the onset of the SD-like
depolarization occurred after 504 ± 131 s (n = 12) of hypoxia, which is >6 min later than in untreated cells (Fig.
6). The initial hyperpolarization was
reduced to 51% of the control amplitude, which is probably due to the
more negative prehypoxic membrane potential, and the gradual
depolarization before SD onset showed a markedly slower time course
(Fig. 6, A and B). The final level of the SD-like
depolarization and the magnitude of the resistance decrease were not
significantly affected by TTX, but the apparent threshold potential of
the SD-like depolarization shifted to more positive membrane
potentials, averaging now
47.3 ± 5.8 mV (n = 12; Fig. 6C).
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DISCUSSION |
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Changes in [Na+]o and [K+]o
Simultaneous measurements of extracellular and intracellular changes demonstrated that the onsets of sudden SD-like depolarization of single cells, the drop in [Na+]o, and the increase in [K+]o exactly coincide with the negative deflection of the extracellular DC potential that signals the onset of hypoxic SD. The extracellular ion changes measured during severe hypoxia in the cell body layer (st. pyramidale) and the dendritic layer (st. radiatum) did not show significant differences (Table 1).
Whereas [K+]o already
increased during the initial, pre-SD phase of hypoxia,
[Na+]o remained virtually
unchanged until the onset of SD. Moderate cell swelling begins during
the initial phase of hypoxia restricting extracellular space by ~10%
(Jing et al. 1994; Müller and Somjen 1999d
), indicating the uptake of solute, most probably NaCl,
into neurons and KCl into glial cells. The stability of
[Na+]o in the face of
pre-SD swelling suggests nearly isotonic influx of salt and water with
moderate cell swelling and restriction in extracellular space exactly
balancing the amount of extracellular Na+ already
entering neurons and possibly also glial cells during the initial phase
of hypoxia.
The recorded ion concentrations,
[K+]o, which increased to
51-54 mM, and [Na+]o,
which dropped to 61-64 mM (Table 1), are in the ranges previously reported for hypoxic and normoxic SD or ischemia. During anoxia of rat
and cat cortex, [K+]o
increased to levels between 25 and 100 mM and
[Na+]o decreased to
48-53 mM, whereas ischemia raised
[K+]o to 75 mM and
depressed [Na+]o to 48 mM
(reviewed by Hansen 1981, 1985
). Normoxic spreading depression in rat cerebellum shifted
[K+]o to 40 mM and
[Na+]o to 60 mM
(Nicholson 1984
; Nicholson and Kraig
1981
).
After reaching its peak but before reoxygenation,
[Na+]o already noticeably
recovered, whereas [K+]o
recovered only slightly during hypoxia (Figs. 1, 2, and 5). The partial
recovery of extracellular ion concentrations in the absence of oxygen
may reflect exchange by diffusion between tissue and bath, but this
cannot be the whole explanation because the Na+
diffusion coefficient is smaller than that for K+
(DNa = 1.33 *
105 cm2/s,
DK = 1.96 *
10
5 cm2/s) (Hille
1992
) yet [Na+]o
recovered faster than
[K+]o. Instead, these
differences might reflect substantially differing kinetics of uptake
and/or release of Na+ and
K+. Because the difference in the time courses of
the two ion concentration changes also was observed in the presence of
TTX (Fig. 5), inactivation of voltage-gated Na+
channels is probably not the explanation for this early
Na+ recovery.
Another interesting aspect is the imbalance of extracellular
Na+ and K+ changes observed
at the height of hypoxic SD. In falling from 155 to 64. 3 mM,
[Na+]o dropped by 90.7 mM, whereas in rising from 3.5 to 50.9 mM, [K+]o increased by only
47.4 mM, leaving an apparent arithmetic cation deficit of 43.3 mM.
Therefore simple exchange of Na+ for
K+ cannot fully explain the ionic changes during
hypoxic SD. Probably most of the difference represents the coupled
influx of Na+ and anions like
Cl or HCO3
, causing neuronal
swelling, with a possible moderate contribution by the release of other
cations, especially Mg2+ from cells. Because a
large fraction of the total intracellular Mg2+ is
bound to ATP, depletion of ATP caused by hypoxia could elevate intracellular Mg2+ activity and thus facilitate
Mg2+ release into the extracellular space. In
fact, Taylor and coworkers (1999)
observed a gradual
loss of Mg2+ from cytoplasm, mitochondria, and
nucleus during oxygen/glucose deprivation in hippocampal slices.
Estimates of intracellular Na+ and K+ changes during hypoxic SD
While hypoxic SD dramatically changed the ionic composition of the
extracellular compartment, our estimates indicate much more moderate
effects on [Na+]i and
[K+]i (Fig. 4). The
difference is due to the smallness of the interstitial volume fraction,
which amounts to only 12-20% of the total tissue volume (Mazel
et al. 1998; McBain et al. 1990
;
Pérez-Pinzón et al. 1995
). The changes of
[Na+]i and
[K+]i were calculated for
pyramidal neurons on the basis of the measured extracellular ion
concentrations during SD, and assuming two theoretical conditions: that
glial cells do not participate in Na+ uptake and
K+ release during SD and that glial cells and
neurons participate equally. The physiological conditions are likely to
lie somewhere between these two extremes and will have to be
established experimentally by microfluorimetric recordings or the use
of ion-sensitive microelectrodes. For now, these calculations are
intended only to define the expected limits of intracellular ion
changes. According to Friedman and Haddad (1994)
,
[Na+]i increased in
cultured cortical neurons during anoxia by 27 mM; this fits well into
the predicted range.
Membrane responses in single CA1 neurons and glial cells
Neurons as well as glial cells both underwent substantial
depolarizations as soon as SD occurred. Yet their membrane responses during the initial phase of hypoxia, before SD onset, clearly differed.
Pyramidal neurons and interneurons hyperpolarized by releasing
K+ into the extracellular space. This neuronal
K+ loss apparently reflects activation of
KCa channels and/or
KATP channels (Erdemli et al.
1998; Fujimura et al. 1997
; Hansen et al.
1982
; Leblond and Krnjevíc 1989
), and at
least a partial contribution also could be the failure of the
Na+/K+ ATPase due to energy
shortage. By contrast, glial cells directly responded with a
depolarization during the initial phase of hypoxia, as expected from
the K+ sensitivity of glial membrane potential
(Kuffler and Nicholls 1966
; Somjen 1987
).
Although we did not observe noticeable differences in the hypoxic
changes induced in pyramidal neurons and interneurons, Congar et
al. (1995) reported anoxia to induce a more pronounced outward current in interneurons than in pyramidal neurons. Because of the
differing experimental conditions (fully submerged slices, kept at
30-32°C) in the studies of Congar et al. (1995)
hypoxic SD did not occur. Instead the anoxic outward currents reported by these authors correspond to the hyperpolarization and the decrease in input resistance we observed during the initial phase of hypoxia before SD onset.
The intracellular potential changes in both, neurons and glial cells
exhibited a clearly defined apparent threshold potential at which the
slow depolarization turned into the rapid, near complete depolarization
(Figs. 1-3). In neurons, this threshold potential is almost identical
to the Na+ spike threshold reported by
Dingledine (1983), and it shifted toward more positive
potentials after TTX treatment (Fig. 6). This suggests that, under
control conditions, this threshold activation of voltage-gated
Na+ channels triggered SD or at least contributed
to the process.
Unclear is, whether activation of glial Na+
channels contributes to the rapid glial depolarization. Although
hippocampal astrocytes do posses voltage-gated
Na+ channels, the K+
conductance of their membranes is at least fourfold higher than the
Na+ conductance, and their membrane potential is
determined mostly by the distribution of K+
across their membrane (Bordey and Sontheimer 1997;
Dennis and Gerschenfeld 1969
; Kuffler and
Nicholls 1966
). The rapid depolarization observed in astrocytes
at the onset of SD is therefore probably the result of the elevated
[K+]o rather than
Na+ influx. The measured increase in
[K+]o from 3.5 to 50.9 mM
is expected to cause a "Nernstian response" of 70.9 mV, at least if
[K+]i is assumed to
remain constant. This calculated potential shift is larger than the
57-mV depolarization observed in glial cells. The difference could be
explained by Cl
influx into glial cells
(Coles et al. 1989
). If, however, glial cells would lose
K+ during SD, then their depolarized
intracellular potential,
28.2 mV, would be near the calculated
EK. In the few instances that it could
be measured, glial input resistance was less dramatically reduced than
that of neurons. This agrees with whole cell patch-clamp recordings
made by Czéh et al. (1992)
, whereas in cat cortex, Sugaya and coworkers (1978)
found no change in glial
input resistance during normoxic SD. These facts point to neurons as
the primary generators of hypoxic SD, whereas glial cells appear to
play a more passive role.
Glial cells probably protect neurons before SD onset by buffering the
excess of [K+]o and
controlling the ionic composition of the extracellular space
(Orkand et al. 1966; Somjen 1987
). The
importance of glial K+ buffering was demonstrated
by Janigro and coworkers (1997)
, who observed
interictal-like events and epileptiform afterdischarges when glial
K+ uptake was blocked by
Cs+. Eventually, however, the glial buffering
capacity is overwhelmed because
[K+]o continues to rise
during hypoxia and glial gap junctions might be closed due to
intracellular acidification.
A more than purely passive role of the glial syncytium in the
generation and propagation of SD may be indicated by the inhibition of
normoxic SD in the presence of the gap-junction uncoupling agent
heptanol (Largo et al. 1997b) as well as the observation that glial cells release glutamate during SD (Basarsky et al. 1999
). Heptanol treatment did, however, not affect the
generation and propagation of hypoxic SD (Aitken et al.
1998
). Also the amount of glutamate released from glial cells
is quite small (only 20% of total release) compared with the amount
released via Ca2+-dependent exocytosis
(Basarsky et al. 1999
). Therefore one might wonder
whether glial glutamate release really could indicate a pivotal glial
contribution to the generation of SD. Furthermore the efficacy of
glutamate antagonists varies among normoxic and hypoxic SD. Although
the generation and propagation of normoxic SD can be blocked by
glutamate antagonists (Lauritzen and Hansen 1992
;
Marranes et al. 1988
), in the case of hypoxic SD
these treatments only caused a postponement of SD onset and a reduction
in
Vo amplitude. Neither could a complete
inhibition of hypoxic SD be achieved by combined application of NMDA
and non-NMDA inhibitors (Jing et al. 1993
; unpublished
observations). From these findings, it appears that glial cells may
contribute differently to the generation and propagation of normoxic
and hypoxic SD, being apparently more important under normoxic than
hypoxic conditions.
A predominantly active contribution of glial cells to the generation of
SD also is questioned by the ineffectiveness of fluoroacetate and
fluorocitrate. Both hypoxic as well as normoxic SD still can be induced
after metabolic poisoning of glial cells (Largo et al.
1997a,b
; Müller and Somjen 1999c
,d
). Of
course the poisoning actions of these agents are not restricted to
glial cells only, but earlier studies showed clearly different time
scales in the decline of glial and neuronal function. Morphological
damage occurred first in glial cells, and also the gradual glial
depolarization, tissue acidosis, and partial loss of
[K+]o regulation do suggest an early loss in
glial function, whereas neurons are at first mostly unaffected
(Largo et al. 1996
, 1997a
).
Effects of TTX
Similarly to earlier studies of both normoxic and hypoxic SD
(Aitken et al. 1991; Sugaya et al. 1978
;
Tobiasz and Nicholson 1982
; Xie et al.
1994
), application of TTX postponed the onset and depressed the
intensity of hypoxic SD but failed to prevent it. The protective effect
of TTX (Fung and Haddad 1997
; Yamasaki et al.
1991
) appears to depend mainly on the delayed onset of SD and
on the slower depolarization and decrease in input resistance during
the initial, pre-SD phase of hypoxia (Figs. 5 and 6, A and
B). Once SD occurred, the extracellular ion changes were
only slightly depressed and the final amplitude of the depolarization during SD of pyramidal neurons was not changed at all by TTX (Fig. 6C). As in this study, TTX also failed to prevent
[Na+]o changes during
normoxic SD in rat cerebellum (Tobiasz and Nicholson 1982
). It appears therefore that during normoxic as well as
hypoxic SD only a small amount (12-18%) of the total
Na+ flux involves TTX-sensitive
Na+ channels.
In single pyramidal neurons, TTX caused a small but consistent positive
shift of the SD threshold potential (Fig. 6), whichin combination
with the slowed gradual, pre-SD depolarization
is apparently
responsible for the marked delay in SD onset. The rapidly depolarizing
segment of the Vi trajectory (the b-c
segment in Fig. 6), the final level of
Vi reached during SD, and the massive reduction in input resistance were, however, not changed. Therefore the
observed 22% decrease in
Vo
amplitude (Fig. 5) cannot be the result of reduced changes on the
single cell level but probably reflects less extracellular current flow
due to desynchronized activity in single neurons.
SD appeared more delayed by TTX in the recordings of Vi from neurons than in the extracellular recordings made with triple-barreled ion-sensitive microelectrodes. This may be a matter of sampling individual units out of a desynchronized population, although localized tissue damage inflicted by the triple-barreled microelectrode, facilitating SD onset, also may have played a role.
Role of TTX-sensitive Na+ channels in spreading depression
Without doubt, the rapid SD-like depolarization of hippocampal
neurons implies massive Na+ influx. This is shown
not only by the drop in
[Na+]o but also in the
reported shift of the reversal potential of the ischemic depolarization
when the Na+ concentration in the bath of brain
tissue slices was reduced (Tanaka et al. 1997). In
dissociated cells and cell cultures, anoxia raised
[Na+]i (Friedman
and Haddad 1994
). The mechanism of Na+
entry during SD is less clear. Voltage-activated "fast"
Na+ channels inactivate within a few
milliseconds. They may play a part in triggering SD but are unlikely to
contribute much to the SD-related inward current itself. A
TTX-sensitive persistent Na+ current is, however,
also present in hippocampal neurons (French et al.
1990
). Although its maximum amplitude in isolated cells originally was reported to be quite small, it appears to grow considerably as a consequence of cyanide poisoning and hypoxia (Hammarström and Gage 1998
) as well as in elevated
[K+]o (Somjen,
unpublished observations). It therefore may be responsible for a
substantial fraction of the SD-like depolarization, but much of the
Na+ appears to enter cells through a
TTX-insensitive mechanism. A TTX-insensitive slow
Na+ current has been reported (Hoehn et
al. 1993
), but this finding is disputed by (Chao and
Alzheimer 1995
).
Inward flow of K+ through voltage-gated K+ channels has to be ruled out as a mechanism driving the self-regenerative depolarization because even at the height of hypoxic SD EK was more negative than Vm and therefore the K+ driving force was directed outwardly at all times.
The events that cause the depolarization in the presence of TTX
therefore remain uncertain for now. In a previous study
(Müller and Somjen 1998), we demonstrated that
hypoxic SD can be prevented by the combined application of
Ni2+, TTX,
6,7-dinitroquinoxaline-2,3-dione, and
(±)-3-(2-carboxypiperazin-4-yl)-propyl-1-phosphonic acid. These
observations suggest that activation of glutamate receptors might
contribute to the ion fluxes during SD. Even though synaptic function
and axonal conduction are blocked during hypoxia in the presence of
TTX, release of glutamate and/or aspartate still does occur.
Depolarization of synaptic terminals because of increasing
[K+]o may be sufficient to trigger vesicle
fusion and transmitter release. In addition, excitatory amino acids
also may be released during SD by two extrasynaptic mechanisms:
Na+-driven glutamate uptake may reverse due to decreased
Na+ driving force (Attwell et al. 1993
) and
swelling-activated nonselective anion channels in the membrane of glial
cells may allow glutamate release into the extracellular space
(Basarsky et al. 1999
; Kimelberg et al.
1990
). The relative contribution of these and other possible Na+ pathways has yet to be investigated in more detail.
Concluding remarks
Our present study demonstrates that the massive depolarization of
pyramidal neurons, interneurons, and glial cells coincided with the
sudden drop in [Na+]o,
the increase in [K+]o,
and the negative Vo. Although
neuronal depolarization is characterized by a marked decrease of input
resistance and Na+ influx, the glial
depolarization seems to be largely "passive," caused by increased
[K+]o. Administration of
TTX delayed the onset of hypoxic SD but did not prevent it. The final
amplitude of the hypoxic SD-like depolarization in single TTX-treated
pyramidal neurons did not differ from untreated control slices, and
once SD set in, ionic maintenance was not much improved by TTX.
We conclude that TTX-sensitive Na+ channels mediate only a small portion of the neuronal Na+ influx during SD. In the absence of TTX, activation of voltage-gated Na+ channels appears to be responsible for the initiation but not the shaping of the hypoxia-induced SD.
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ACKNOWLEDGMENTS |
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This study was supported by National Institute of Neurological Disorders and Stroke Grant NS-18670.
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FOOTNOTES |
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Address for reprint requests: G. G. Somjen, Dept. of Cell Biology, Box 3709, Duke University Medical Center, Durham, NC 27710.
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 26 June 1999; accepted in final form 5 October 1999.
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REFERENCES |
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