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Wake Forest University Health Sciences, Department of Physiology and Pharmacology, Winston-Salem, North Carolina 27157
Submitted 23 December 2002; accepted in final form 8 March 2003
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ABSTRACT |
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-aminobutyric acid-A (GABAA) receptor-mediated inhibitory
postsynaptic currents (sIPSCs and eIPSCs, respectively) were recorded prior to
and following depolarization of CA1 hippocampal pyramidal cells. Depolarizing
voltage pulses were shaped to evoke currents in QX-314-treated cells similar
to those accompanying single spontaneous voltage-clamped action potentials
recorded from the soma. Attempts were made to elicit DSI with trains of these
pulses that mimicked hippocampal cell firing patterns in vivo, for instance,
when animals traverse place fields or are performing a short-term memory task.
DSI could not be elicited by such pulse trains or by a number of other
combinations of behaviorally specific firing parameters. The minimum duration
of depolarization necessary to elicit DSI in hippocampal neurons determined by
paired-pulse manipulation was 50 –75 ms at a critical interval of 20
–30 ms between pulse pairs. Under the conditions tested, the normal
firing patterns of hippocampal neurons that occur in vivo do not appear to
elicit DSI. |
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INTRODUCTION |
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,
1994a,b)
to the release of endogenous cannabinoids (endocannabinoids) in hippocampus
(Wilson and Nicoll 2001,
2002), has important
implications. The reduction in GABAergic transmission, as signaled by the
decrease in amplitude of inhibitory postsynaptic currents (IPSCs) that occurs
following 1.0-s depolarizations of hippocampal pyramidal cells in slices
(Wilson and Nicoll 2001;
Wilson et al. 2001) and
cultured hippocampal neurons (Ohno-Shosaku
et al. 2002), is prevented by the CB1 cannabinoid
receptor-antagonist SR141716A. A similar depolarization-dependent cannabinoid
suppression of presynaptic release of excitatory transmitter has been
demonstrated in the rat cerebellum
(Kreitzer and Regehr 2001). In
hippocampus, DSI involves activation of CB1 receptors on GABAergic terminals
of hippocampal interneurons (Katona et al.
1999; Wilson and Nicoll
2001), which presumably lead to a reduction in voltage-sensitive
Ca2+ conductances
(Mackie and Hille 1992) and a
corresponding decrease in GABA release, signaled by a marked reduction in both
evoked (eIPSCs) and spontaneous inhibitory postsynaptic currents (sIPSCs) in
hippocampal pyramidal cells (Wilson et al.
2001).
We investigated DSI from two different perspectives: first, as to whether
this process could be initiated in vitro by normal patterns of action
potentials recorded from animals performing hippocampal-dependent behavioral
tasks. This is important because it addresses the functional significance of
DSI and associated endocannabinoids that are involved, since it appears that
hippocampal neurons need to be significantly depolarized to release
endocannabinoids (Lenz and Alger
1999; Wilson and Nicoll
2002). The second purpose was to define precisely the range of
frequencies and minimum duration of depolarizing pulses required to elicit DSI
in hippocampal pyramidal neurons in vitro. Both objectives were directed at
determining whether release of endocannabinoids is possible under what can be
considered normal firing conditions for hippocampal neurons in vivo. To
address this issue, we utilized trains of depolarizing pulses similar to those
that occur during spontaneously increased firing, in various behavioral
contexts (Deadwyler et al.
1996; Hampson et al.
1996; Hampson and Deadwyler
2000). This was performed in hippocampal pyramidal cells recorded
in vitro in which DSI could in fact be demonstrated. Hence, the following
findings: 1) describe the minimal activation conditions required to
provoke DSI in hippocampal pyramidal neurons, and therefore, and 2)
indicate the circumstances in which endocannabinoid release could contribute
to hippocampal operation in vivo.
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METHODS |
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The preparation of hippocampal brain slices was similar to that described
in several previous reports (Alger
1999; Kim et al.
2002; Lenz and Alger
1999; Pitler and Alger 1994;
Wilson and Nicoll 2001).
One-week-old male Sprague-Dawley rats (Harlan) were CO2
anesthetized and rapidly decapitated. The brains were removed and the
hippocampi dissected in cold (4°C) buffered saline medium (see bathing
medium, below) and then sliced at 300 µm thickness along the transverse
axis using a Leica vibratome. Slices were incubated in oxygenated buffered
saline for 1 h at room temperature before recording. Prior to use, slices were
transferred to a custom perfusion chamber
(Staff et al. 2000) and viewed
through a Zeiss Axioskop2 near-infrared differential interference contrast
(DIC) microscope. Slices were constantly perfused with warmed (35°C)
bathing medium [140 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 2 mM
MgCl2, 10 mM glucose, 10 mM HEPES, 20 µM
6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), and 50 µM
2-amino-5-phosphovaleric acid (APV)]. Carbachol (3.0 µM) was added to the
bathing medium after formation of the whole cell patch to increase the
frequency of sIPSCs. All experiments were performed on slices within 2 h
following transfer to the recording chamber.
Recording methods
Whole cell patch-clamp procedures, similar to that reported previously
(Deadwyler et al. 1993;
Mu et al. 1999), were
performed using the DIC microscope to visually localize individual CA1
pyramidal neurons in slices of hippocampus. Briefly, patch electrodes were
prepared from 1.5 mm outer diameter, 1.1 mm inner diameter borosilicate glass
capillaries to produce 1–2 µm (2–5 M
) tip openings.
Electrodes were filled by suction and backfilling with a standard
intracellular solution [140 mM KCH3SO3 (or 100 mM
CsCH3SO3, 40 mM CsCl, 3 mM KCl), 0.2 mM EGTA, 0.02 mM
CaCl2, 1 mM MgCl2, 2 mM ATP, 300 µM GTP, 10 mM HEPES
buffer (Sigma), and 5 mM QX314]. A critical aspect of pipette solutions that
affects DSI is the buffered calcium level in the cell, since increased
internal calcium increases the depolarization threshold for DSI. Internal
calcium concentrations were calculated to be 20 nM, with low calcium
buffering, a condition that provides the lowest threshold for provoking DSI
(Lenz and Alger 1999).
Sealing the pipette to the neuron and obtaining access to the whole cell
followed previously described patch-clamp procedures
(Hamill et al. 1981;
Mu et al. 1999). Voltage-clamp
recordings and command voltage steps were performed with an AxoClamp 2A
amplifier and Digidata 1322A controller (Axon Instruments). Whole cell current
and voltage records were acquired and stored on magnetic disk using pClamp
CLAMPEX software (Axon Instruments). Pipette tip junction potentials were
continuously monitored and compensated as necessary at the amplifier prior to
breakage of the seal. Access (series) resistance was typically 5–10
M. Series resistance compensation was usually not necessary because of
low pipette resistance. Leakage correction and capacitance compensation
(typically 10 –30 pF) utilized dialed-in compensation adjustment at the
amplifier, as well as a P/–4 subtraction procedure within the
acquisition program. Cells were voltage-clamped and held near resting membrane
potential (–70 mV) during testing.
Evoked inhibitory postsynaptic currents (eIPSCs) were produced by
stimulation through a bipolar concentric electrode (Frederic Haer) placed
within 50 –100 µm of the pipette in the CA1 cell layer. CNQX (20
µM) and APV (50 µM) were present in the bathing medium to block
glutamatergic synaptic responses. The frequency of sIPSCs was enhanced by
addition of carbachol (3 µM) to the medium. DSI was elicited by
depolarizing pulse steps (from –70 to 0 mV) delivered through the
patch-clamp electrode. Measurement of current amplitudes and time constants
utilized pClamp software. Measurement of DSI consisted of calculating the area
of individual sIPSCs and summing all sIPSCs in the 3-s period prior to the
depolarization step, with a similar measure taken over 4 –7 s following
depolarization at the stage of maximum suppression of IPSCs
(Alger 1999). In most cases,
DSI was computed from pre- and post-sIPSC area [DSI = (pre ÷ post)
x 100%] and reported as mean ± SE
(Alger 1999;
Lenz and Alger 1999). Bar
graphs were plotted using mean (±SE) percentage of prepulse sIPSC area
[(1 – (pre ÷ post)) x 100%] for comparison with current
traces. Statistics were calculated from analysis of variance (ANOVA) of
within-cell comparisons of the effects of depolarization patterns and drug
effects, with individual comparisons calculated as orthogonal linear contrasts
either between conditions or against no change in sIPSC area (mean percentage
of prepulse sIPSC area = 100%, DSI = 0%).
Drug preparation
WIN 55,212-2 (Research Biochemicals, Natick, MA) was prepared daily from a
10 mM stock solution in ethanol, diluted with extracellular bathing medium,
and the ethanol evaporated under a constant stream of nitrogen
(Mu et al. 1999). The drug was
added to the bathing solution in either 100 or 500 nM concentrations.
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RESULTS |
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DSI was produced in the same manner as described previously
(Alger 1999;
Lenz and Alger 1999;
Pitler and Alger 1994b;
Wilson and Nicoll 2001). Using
two measures of -aminobutyric acid (GABA) release, eIPSCs, or sIPSCs,
DSI could be elicited in the majority of neurons tested (n = 150).
Figure 1A shows the
decrease in sIPSCs resulting from a 1.0-s depolarizing pulse. The duration and
magnitude of DSI suppression of sIPSCs by this standard depolarizing parameter
was appreciable (mean decrease in amplitude from the predepolarization
baseline = 52.2 ± 0.6%). Figure
1A also illustrates that sIPSCs were bicuculline
sensitive and suppressed by application of the cannabinoid CB1 receptor
agonist WIN55,212-2 (500 nM) but recovered in the presence of the CB1 receptor
antagonist SR141716A (100 nM). Also, DSI was completely blocked by pretreating
slices with SR141716A as evidenced by absence of decrease in sIPSC area from
prebaseline (1.1 ± 2.3%; Fig.
1B), indicating reliance on release of endogenous
cannabinoids (Kim et al. 2002;
Ohno-Shosaku et al. 2002;
Wilson and Nicoll 2001;
Wilson et al. 2001).
Depolarizing pulses of 100-ms duration elicited significant DSI but at an
increased latency to onset (2.8 ± 0.06 s) compared with 1.0-s pulses
(Wilson et al. 2001). The
traces and graph in Fig.
1B show the effects of several agents used to test DSI
produced by 1-s depolarizations (DSI = 52.3 ± 2.8%,
F(1,283) = 257.3, P < 0.001). DSI was not
altered by pretreating slices with the phosphatase inhibitor calyculin A (DSI
= 33.7 ± 3.1%, F(1,283) = 108.6, P <
0.001) nor the adenylate cyclase activator, forskolin (DSI = 31.3 ±
2.4%, F(1,283) = 96.2, P < 0.001), but was
blocked by the N-type Ca2+ channel blocker
-conotoxin [DSI = 2.8 ± 1.6%, F(1,283) =
0.87, not significant (NS)]. Thus, by a variety of pharmacological criteria,
DSI observed under our recording conditions was similar to that described in
several other studies (cf. Alger
1999; Wilson and Nicoll
2002).
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Involvement of K-currents in DSI
We next examined the whole cell currents associated with depolarizing
pulses that elicited DSI, in comparison to simulated action potential and
other depolarizing currents that occur in vivo. The outward potassium current
associated with spontaneous action potentials evoked under voltage clamp was
found to be 800 pA with a duration of 3–5 ms. Voltage parameters that
produced action potential-like currents were then systematically examined when
voltage-sensitive sodium channels were blocked as in most demonstrations of
DSI (Lenz and Alger 1999;
Wilson et al. 2001). Injected
depolarizing command pulses of <1.0-ms duration were not sufficient to
elicit outward potassium current similar to that exhibited by spontaneous
action potentials. Depolarizing pulses between 1 and 10 ms produced outward
currents identified as a fast inactivating potassium A current of the Kv1.4
type (Dolly and Parcej 1996;
Grosse et al. 2000;
Sheng et al. 1992). If the
depolarizing pulse was ≥10 ms, a subsequent late onset K-current
(Dolly and Parcej 1996;
Storm 1990) was apparent,
which lasted for the duration of the injected voltage with no sign of
inactivation (Fig. 2, Control).
The outward potassium currents evoked by these depolarizing pulses were found
to be altered by either 4-aminopyridine (4-AP) (A-current) or
tetraethylammonium (TEA; K-current, Fig.
2, TEA) as documented in prior studies
(Hille 1998;
Mu et al. 1999;
Rudy 1988;
Serodio and Rudy 1998;
Storm 1990).
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The other variable relevant to producing DSI is the level of
depolarization. Command voltage steps that produce less depolarization (i.e.,
steps to 0 mV) typically did not elicit DSI
(Lenz and Alger 1999). As
such, there are few membrane transients that meet such a requirement in
vivo—with the exception of the action potential itself. Injected
voltages were therefore tailored to evoke currents that were the following:
1) similar in duration and amplitude to those produced by spontaneous
action potentials recorded under voltage clamp in similar bath and pipette
conditions with the exception of QX-314; 2) similar in voltage
dependence and pharmacological sensitivity with respect to outward potassium
currents that occur during spontaneous or evoked action potentials in vitro.
Using these guidelines, we subsequently showed that if the injected pulses
were prolonged, they not only produced the expected slow onset and
noninactivating K-current, but DSI as well
(Fig. 2, Control). Blockade of
the late onset K-current by TEA eliminated DSI even though A-current remained
(Fig. 2, TEA). This indicates
that the time course, magnitude, and threshold for producing DSI were related
to the duration of depolarizing K-current and hence the charge transfer
necessary for activation of voltage-dependent Ca2+
channels (Lenz and Alger
1999). A minimum –70 to 0 mV depolarizing pulse of 50-ms
duration produced only 5% DSI. Figure
3 shows a comparison of DSI as a percentage of sIPSP area elicited
by injected depolarizing pulses ranging from 25 ms (DSI = 1.1 ± 1.4%,
F(1,283) = 0.11, NS) to 1.0 s (DSI = 52.3 ± 2.8%,
F(1,283) = 257.3, P < 0.001). It is clear that
as the duration of the depolarizing pulse decreased from 1.0 s, the percentage
of maximum DSI decreased as well; however, it is also apparent from
Fig. 3 that this relationship
was not entirely linear across the range of pulses tested. For depolarizing
pulses of 25–200 ms (DSI = 37.4 ± 3.6%,
F(1,283) = 132.4, P < 0.001) decrease in sIPSC
frequency (increase in DSI) was close to linear and steep (slope = 0.23% per
ms), but as depolarizing pulses exceeded 250 ms
(Fig. 3), the slope of this
function became much more shallow (slope = 0.02% per ms).
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Simulated in vivo neuron firing and DSI
Under most conditions in vivo, pyramidal cell action potentials are not
accompanied by large calcium-mediated depolarizing shifts and are of a
duration (1–10 ms), even in dendrites, that would be subthreshold for
inducing DSI (Staff et al.
2000). It is possible, however, that frequency-summation of action
potentials in vivo might produce sufficient depolarization per unit time to
elicit DSI. Therefore we tested pulse frequencies and patterns that mimicked
increases in hippocampal pyramidal cell firing in vivo for their ability to
elicit DSI. Two major conditions that produce increased firing in vivo are as
follows: animals traversing place fields in an open arena
(Frank et al. 2000;
Hampson et al. 1996;
Muller et al. 1987) and
performance of hippocampal-dependent learning and memory tasks
(Deadwyler et al. 1996;
Hampson et al. 2002;
Eichenbaum et al. 1989;
Wood et al. 1999).
Figure 4 shows two examples of
spike trains recorded while animals either 1) traversed a place field
specific for that cell, or 2) responded to behaviorally significant
stimuli in a delayed-nonmatch-to-sample (DNMS) memory task. The frequencies
ranged from 0.5 to 10.1 Hz for place field firing, and 0.5 to 17.8 Hz for the
DNMS task. When injected into cells in hippocampal slices, neither firing
pattern was sufficient to produce DSI, even if several trains were delivered
within short intervals (1.0-min separations). The bottom of
Fig. 4 shows superimposed
traces of sIPSCs during five repeated presentations of the DNMS firing pattern
(Fig. 4, middle
right), with no indication of DSI (2.7 ± 0.9% DSI,
F(1,283) = 0.84, NS).
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To examine further the frequency dependence of the depolarizing pulses
required to elicit DSI, several additional types of pulse trains were also
tested (Fig. 5, A and
B). These trains employed depolarizing pulses of
differing durations and frequencies, some of which were not typical of normal
pyramidal cell activation, but might occur in vivo under specific
circumstances (LeBeau and Alger
1998). Frequencies between 5 and 400 Hz in patterns consisting of
1– 400 pulse trains with pulse durations ranging between 5 and 20 ms
were all ineffective in eliciting significant DSI
(Fig. 5B). No
significant DSI was detected when irregular bursts of five 10-ms pulses were
delivered at an average interval of 50 ms (DSI = 0.2 ± 1.4%,
F(1,283) = 0.03, NS). The maximum effect produced was by
trains with pulse durations of 2 ms at frequencies ≤400 Hz (DSI = 3.5
± 2.1%, F(1,283) = 1.15, NS,
Fig. 5B). Even trains
of 6 –10 pulses at 100 Hz, repeated in 6-Hz bursts, which mimicked
-bursts and effectively produced long-term potentiation (LTP) in
hippocampal neurons (Rose and Dunwiddie
1986), proved ineffective for inducing DSI (DSI = 6.3 ±
2.2%, F(1,283) = 3.38, NS). Since depolarizations to 0 mV
for 70 –100 ms (Fig. 3),
the threshold for DSI in vitro, may not be normal occurrences in healthy adult
pyramidal neurons (Jefferys
1994; LeBeau and Alger
1998; Leinekugel et al.
2002), it is possible that frequency and duration of cellular
depolarization must interact to produce DSI.
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To examine the temporal specificity of DSI, a paired-pulse paradigm was used in which two pulses of 35-ms duration were initially separated by >100 ms, a protocol insufficient to elicit DSI. The inter-pulse interval between these pulses was then systematically shortened until DSI was produced, without changing level of depolarization. Figure 6, A and B, shows that the threshold for significant DSI was achieved at an interval of 30 ms and that DSI systematically increased in magnitude when the two pulses were moved closer together until at 10-ms separation, DSI was similar to that effected by a single 70-ms duration pulse (DSI = 18.8 ± 2.4%, F(1,283) = 22.4, P < 0.01, Fig. 6B). This indicated that at least one critical parameter for producing DSI was the interval (30 –35 ms) between depolarizing pulses. DSI could not be elicited, however, by simply delivering the same total amount of depolarization per unit time over the interval. Higher frequency shorter duration pulses (i.e., 7 pulses of 10-ms duration), delivered in the same 100 ms interval of time (e.g., 70 Hz) as the two 35-ms duration pulses (Fig. 6A), did not produce DSI (DSI < 3.0%, F(1,283) < 2.3, NS). This was further verified by systematically decreasing the duration of the first 35-ms pulse while the interval between the two pulses was held constant (20 ms). If the initial pulse duration was decreased to <25 ms, DSI was effectively eliminated, even though the interpulse interval (20 ms) that was effective if both pulses were ≥35 ms was maintained (DSI < 2.5%, F(1,283) < 1.8, NS, see Fig. 6B).
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Simulated in vivo firing and membrane depolarization
The finding that DSI was temporally dependent in this manner indicates that
the pulse trains shown in Figs.
4 and
5 did not meet minimal criteria
for eliciting DSI. To determine whether the depolarizing pulse trains that did
not produce DSI in voltage clamp might sum and produce sufficient membrane
depolarization if the membrane were not clamped
(Pitler and Alger 1994b), the
same trains were delivered to cells in current-clamp mode. Cells were held at
–74 mV by application of slight hyperpolarizing current and recorded
without QX 314 in the pipette solution to allow spontaneous action potentials
to occur. Figure 7A
(left) shows an action potential elicited by 25 pA (1.6 ms)
depolarizing current, and a train of 20 such action potentials
(Fig. 7A,
right), delivered in a pattern similar to that shown in
Fig. 4. The accompanying
depolarizing shift in the unclamped membrane was far from sufficient to
produce DSI. The bar graph in Fig.
7B shows that neither the simulated baseline firing rate
(5 pulses, mean rate 0.5 Hz) nor the increases produced by several different
patterned trains of 20 pulses (mean rate 20 Hz) produced significant DSI (DSI
< 1.8%, F(1,283) < 0.7, NS). However, a depolarizing
current step of 100-ms duration of the same amplitude did induce significant
DSI (DSI = 38.7 ± 4.9%, F(1,283) = 71.2, P
< 0.001). The graph also shows that addition of WIN 55,212-2 (500 nM)
reduced sIPSP amplitudes (DSI = 24.8 ± 4.2%,
F(1,283) = 44.6, P < 0.001) and that the
reduction was blocked by co-administration of (100 nM) SR141716A (DSI = 2.7
± 3.1%, F(1,283) = 3.1, NS). Thus in current-clamp
mode, DSI could be generated using conventional long-duration depolarizing
protocols, but patterns of action potentials that occur in hippocampus in vivo
were not sufficient to elicit DSI.
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DISCUSSION |
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). The
investigation revealed quite precisely the limits of both frequency and pulse
duration required to elicit DSI in hippocampal neurons
(Fig. 3). We confirmed that
blockade of voltage-dependent N-type Ca2+ channels was
sufficient to eliminate DSI (Fig.
1B) and showed that DSI was systematically increased as
the duration of the depolarizing pulse delivered to the cell
(Fig. 3) was also increased as
previously reported (Lenz and Alger
1999). In addition the threshold values for eliciting DSI
described here are in the same range (75- to 100-ms duration) reported by
Wilson et al. (2001) with
similar calcium buffering to that reported by Lenz and Alger
(1999). At the same time the
results question whether normal in vivo firing patterns of hippocampal neurons
are sufficient to induce DSI (Fig.
4). Attempts were made to elicit DSI with an extensive array of
depolarizing pulse parameters and protocols (Figs.
4,5,6),
patterns of stimulation that mimicked the range of firing of hippocampal
neurons in behaviorally relevant circumstances
(Deadwyler et al. 1996; Hampson
et al. 1996,
1999,
2002). However, none of the
parameters resulted in significant DSI.
In the preceding testing it was assumed that a 10-ms duration depolarizing
pulse under voltage-clamp conditions was within the range of membrane changes
that would occur in the soma of active hippocampal pyramidal neurons in vivo
(Hille 1998;
Staff et al. 2000). The
standard pulse used for testing DSI production was patterned to replicate the
duration of outward potassium currents that occurred under voltage clamp
during spontaneous action potentials elicited without QX314 blockade. While
logical, this assumption is not necessarily correct as action potential shapes
and durations can vary considerably depending on where generated and whether
they traverse into dendrites or are localized to somal regions of the cell
(Golding et al. 2001;
Magee 2001;
Watanabe et al. 2002).
However, depolarizations resulting from inputs to dendrites are typically
associated with fewer action potentials than those elicited by the same
depolarizing inputs to the soma (Harris et
al. 2001; Leinekugel et al.
2002; Magee 2001)
and bursts of action potentials in hippocampal pyramidal cells are more likely
to be generated nearer to than further from the soma
(Harris et al. 2001). Hence,
pulses injected into the soma would appear to be the most potent as well as
the most capable of producing DSI (Fig.
3). From this perspective, burst-type firing in pyramidal cells
would provide the best possible means of producing DSI
(LeBeau and Alger 1998).
However, as shown here, the timing and duration of such bursts are extremely
critical (Fig. 6). Alger and
colleagues have shown definitively that DSI is directly dependent on calcium
charge differences in depolarized pyramidal cells
(Lenz and Alger 1999). Hence
it is possible, as suggested, that DSI and this form of endocannabinoid
release is related to processes not routinely encountered in behaviorally or
cognitively activated hippocampal neurons
(Christie and Vaughan 2001;
Seward et al. 1995; Seward and
Nowicky 1996).
Because the duration of depolarizing pulses required (Figs.
3 and
6) to produce DSI (Pitler and
Alger 1994), certain outward currents can be eliminated as factors responsible
for its occurrence. These include potassium A and D currents whose time course
of activation is shorter, resulting in nearly complete inactivation
(Fig. 2) during sustained
depolarizations ≥25 ms (Hille
1998; Jan and Jan
1994; Mu et al.
1999; Storm 1990;
Watanabe et al. 2002). This
minimum duration necessary to provoke DSI leaves only "K" and
"leak" potassium currents as possible candidates for providing the
outward current during depolarizations necessary for DSI. We confirmed this
relationship by showing that DSI was attenuated in the presence of TEA which
blocks K but not A and D voltage-dependent potassium current
(Hille 1998;
Storm 1990). Since the
reduction in K-current by TEA also blocked DSI
(Fig. 4), it is likely that the
depolarizing pulse parameters sufficient to elicit DSI involve TEA-sensitive
potassium conductances. This dependence between DSI and K-current may be
circumstantial due to the fact that both are a direct function of injected
depolarizing pulses, but since the specific nature of this interaction with
respect to presynaptic "expression" of DSI is undetermined at this
point, it remains a possibility.
The fact that DSI could not be evoked within the range of characteristic in
vivo firing patterns of hippocampal pyramidal cells (Figs.
4,5)
questions the relevance of this type of endocannabinoid release process in the
control or modulation of behaviorally relevant hippocampal pyramidal cell
activity. This view is reinforced when it is noted that the above experiments
were carried out under conditions that have been shown to be optimal for
inducing DSI with the lowest possible duration of depolarizing pulses
(Pitler and Alger 1994a),
including high concentrations of carbachol (3 µM) and calculated
concentrations (20 nM) of intracellular free Ca2+
(Lenz and Alger 1999). At the
same time, it was demonstrated that the DSI evoked with longer duration pulses
was in fact cannabinoid receptor (CB1) sensitive
(Fig. 1A) and that
reductions in sIPSCs were readily produced with WIN 55,212-2 and reversed by
SR141716A applied to these same cells. Because there was no effect of the CB1
receptor antagonist (SR) alone on sIPSCs, constitutive "release"
of endocannabinoids was apparently nonexistent in the absence of
suprathreshold depolarizing pulses necessary for DSI
(Wilson and Nicoll 2002).
These results are consistent with our prior reports that the antagonist alone
had no effects on cannabinoid-altered memory processes
(Hampson and Deadwyler 2000).
However since the precise mechanism of endocannabinoid release under these
conditions is not known (Davies et al.
2002; Kim et al.
2002; Kreitzer and Regehr
2002; Piomelli et al.
2000; Wilson and Nicoll
2002), it is not possible at this time to rule out other factors
that might induce endocannabinoid action in vivo.
The threshold duration for evoking DSI with a single pulse under recording
conditions employed here was 50 –75 ms; single pulses of less duration
(Fig. 5), no matter the
frequency, failed to elicit significant DSI
(Fig. 3). At the same time, the
process appears to have a dynamic that has not been described in prior
reports, namely that mere depolarization per unit time is not adequate to
induce endocannabinoid release. This was demonstrated by injecting pulse pairs
of the same amplitude in which DSI was dependent on pulse separation
(25–30 ms, Fig.
6B). This temporal dependence of DSI could not be
mimicked by pulses of shorter duration presented at higher frequencies, nor by
decreasing the duration of the first pulse and maintaining the temporal
separation between pulses. These results provide clues as to the potential
kinetics of intracellular events underlying endocannabinoid release and
support the proposal by Lenz and Alger
(Lenz and Alger 1999;
Lenz et al. 1998) for a
two-step process, possibly involving diffusion of intracellular calcium to a
prescribed release site. The recent demonstration of enhancement of
endocannabinoid-mediated DSI by co-activation of group I metabotropic
receptors (Varma et al. 2001)
suggests facilitation in this process. However, since
Ca2+ appears to be required for release of
endocannabinoids in most contexts
(Piomelli et al. 2000), it is
not clear how reduced sIPSCs produced by activation of group I metabotropic
receptors alone (as recently reported by
Kim et al. 2002) could explain
the relationship between depolarization-induced endocannabinoid release (i.e.,
DSI), and how this might be facilitated during normal firing of hippocampal
neurons in vivo.
The failure to demonstrate DSI with trains of depolarizing pulses that
mimicked those generated in animals engaged in hippocampal-dependent tasks
does not rule out cannabinoid participation in these processes. However,
previous research showing effects of cannabinoids on hippocampal cells
indicates that systemically administered cannabinoids suppress firing of
hippocampal neurons in behaviorally relevant circumstances (Hampson and
Deadwyler 2000, 2003;
Heyser et al. 1993). Similar
suppressive effects have been reported on hippocampal
GABAA-mediated potentials in vitro
(Hoffman and Lupica 2000) and
in vivo (Hajos et al. 2000).
While there may be circumstances in which the threshold for producing
endocannabinoid-dependent DSI is exceeded in hippocampal neurons
(LeBeau and Alger 1998), such
as depolarization shifts as a result of asphyxia, excitotoxicity, or
epileptogenesis, most of these conditions are considered to be pathologic
(Arabadzisz et al. 2002;
Gorter et al. 2002;
Perez-Velazquez et al. 1997;
Shuttleworth and Connor 2001;
Tanaka et al. 2002;
Yin et al. 2002). The one
normal circumstance in which prolonged action potential bursts occur
frequently in hippocampal neurons in the rat is during sleep
(Nadasdy et al. 1999);
however, even these episodes are transient and, outside of providing a
synchronizing influence, tend to mimic firing patterns that occur in waking
(Louie and Wilson 2001).
Whatever the process responsible for release of endogenous cannabinoids in
vivo and its relation to DSI, its significance will not be completely
understood until cannabinoid-mediated DSI is demonstrated in vivo
(Piomelli et al. 2000). The
above paired-pulse analyses revealed the required frequency range and duration
of such depolarizing events (Figs.
3 and
6B) and showed that
production of the same depolarization per unit time via trains of shorter
action potential-like pulses was not sufficient to induce release of
endocannabinoids (Fig.
7B). This temporal dependence appears to have a direct
relationship to events that provoke calcium entry (see
Wilson and Nicoll 2002), but
how this form of depolarization is induced, either synaptically or by other
cellular events in vivo, has not yet been identified. That endogenous
cannabinoids may in fact have an important role in hippocampal function and in
the sculpting of critical firing patterns, remains a distinct possibility,
especially with respect to the profound and selective effects of systemically
administered CB1 receptor agonists on hippocampal-mediated short-term memory
processes (Hampson and Deadwyler
2000, 2003). However, since
administration of the CB1 receptor antagonist (SR) alone is ineffective in the
latter circumstance and does not alter GABA sIPSCs in the absence of DSI
(Wilson and Nicoll 2002), the
role of endocannabinoid signaling in hippocampal-dependent behavior remains
unresolved.
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ACKNOWLEDGMENTS |
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This work was supported by National Institute on Drug Abuse Grants DA-07625, DA-03502, and DA-00119 to S. A. Deadwyler and DA-08549 to R. E. Hampson.
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FOOTNOTES |
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Address for reprint requests: S. A. Deadwyler, Department of Physiology and Pharmacology, Wake Forest University Health Sciences, Winston-Salem, NC 27157 (E-mail: sdeadwyl{at}wfubmc.edu).
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INTRODUCTION
