Volen Center for Complex Systems, Biology Department, Brandeis University, Waltham, Massachusetts 02454
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ABSTRACT |
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Chen, Huan-Xin, Nikolai Otmakhov, and John Lisman. Requirements for LTP Induction by Pairing in Hippocampal CA1 Pyramidal Cells. J. Neurophysiol. 82: 526-532, 1999. The induction of long-term potentiation (LTP) in the hippocampal CA1 region requires both presynaptic activity and large postsynaptic depolarization. A standard protocol for inducing LTP using whole-cell recording is to pair low-frequency synaptic stimulation (100-200 pulses, 1-2 Hz) with a depolarizing voltage-clamp pulse (1-3 min duration). In this standard protocol, a Cs+-based internal solution is used to improve the fidelity of the depolarization produced by voltage-clamp. In an attempt to induce LTP more rapidly, we tried to induce LTP by pairing high-frequency stimulation (200 pulses, 20-100 Hz) with a short depolarization (~15 s). Surprisingly, we found that this protocol failed to induce LTP, even though large LTP (~300% of baseline) could be induced by a subsequent standard protocol in the same cell. Pairing brief high-frequency stimulation at the beginning of a long depolarization (3 min) also did not induce LTP. However, the same high-frequency stimulation at the end of the long depolarization did induce LTP. When similar experiments were done with a K+-based internal solution, pairing high-frequency stimulation with a short depolarization did induce LTP. This indicates that the requirement for long depolarization is related to the use of Cs+. We speculate that, when recording is made with Cs+, a tetanus given at the beginning of depolarization initiates a process that inhibits N-methyl-D-aspartate (NMDA)-dependent LTP. This inhibitory process itself decays away during prolonged depolarization.
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INTRODUCTION |
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Long-term potentiation (LTP) of the Schaffer
collateral synapses in the CA1 region of the hippocampus is the primary
model system for the study of the associative synaptic modification thought to underlie learning and memory (Bliss and Collingridge 1993). The form of LTP at these synapses has a Hebbian
property: synapses are strengthened if there is both presynaptic
activity and substantial postsynaptic depolarization (Brown et
al. 1990
). The requirement for postsynaptic depolarization is
due to the properties of the
N-methyl-D-aspartate (NMDA) channel. To open, the Mg2+ block of these channels must be relieved
by depolarization (Mayer et al. 1984
; Nowak et
al. 1984
). Once NMDA channels open, they allow the influx of
Ca2+ that triggers synaptic strengthening
(Bliss and Collingridge 1993
). A commonly used protocol
for inducing LTP is to give "tetanic" stimulation in which a large
number of axonal inputs is stimulated at high frequency (100 Hz) for
1 s. This synaptic input produces the postsynaptic depolarization
required to open the NMDA conductance, but the quantitative properties
and mechanism of this depolarization are not yet established. What is
clear is that many factors are likely to be involved, including
temporal and spatial summation of excitatory and inhibitory
postsynaptic potentials (EPSPs and IPSPs, respectively), the triggering
of back-propagating Na+ spikes (Magee and
Johnston 1997
), the triggering of bursts (Thomas et al.
1998
), and the function of a variety of other voltage-dependent conductances (Magee 1998
; Magee et al.
1998
).
Because of the complexity of the postsynaptic processes that affect
depolarization during tetanus-induced LTP, many investigators have
sought to evoke LTP in a simpler and more defined way by using what is
termed a "pairing protocol." This protocol circumvents the
complexities of synaptically induced postsynaptic depolarization by
simply imposing depolarization by current injection through the
microelectrode (Gustafsson et al. 1987) or by voltage
clamp, using the somatic patch electrode (Malinow 1991
).
Typically Cs+ is used as the major internal
cation because it blocks K+ channels and makes it
possible to achieve a larger and more uniform dendritic depolarization.
Because postsynaptic depolarization is produced by the clamp rather
than by synaptic stimulation, only a few input fibers need to be
stimulated (Kullmann and Nicoll 1992
; Malinow
1991
). This improves accuracy with which synaptic currents can
be quantified, because small currents reduce voltage-clamp errors. A
further aspect of standard pairing protocols is that a much lower
frequency of synaptic stimulation (0.1-2 Hz) is used (Colino et
al. 1992
; Malinow and Tsien 1990
; Manabe et al.
1992
; Perkel and Nicoll 1993
) than during tetanic
stimulation (100 Hz). This produces an additional simplification by
avoiding the transient forms of presynaptic plasticity that occur
during high-frequency stimulation. A consequence of using low-frequency
stimulation is that the depolarization during pairing must be long.
Typically, this depolarization is applied for more than one minute
(Colino et al. 1992
; Manabe et al. 1992
;
Otmakhov et al. 1997
).
The experiments reported began with the attempt to devise a much briefer pairing protocol that more closely resembles the duration of depolarization during tetantically induced LTP. Based on what is known about LTP, we reasoned that it should be possible to induce LTP by tetanic stimulation during a brief depolarizing voltage-clamp pulse. We found, however, that it is not possible to do so. Our results further show that this surprising inability to induce LTP is related to the use of Cs+ as the major internal cation.
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METHODS |
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Transverse hippocampal slices were prepared from male Long-Evans
rats (14-19 days old) as described previously (Otmakhov et al.
1997). The CA3 region of each slice was removed from the slice by a surgical cut. Slices were incubated on cell culture inserts (Falcon, 8 µm pore diameter) covered by a thin layer of artificial cerebrospinal fluid (ACSF containing 2 mM Ca2+
and 6 mM Mg2+) and surrounded by a humidified
95% O2-5% CO2 atmosphere
at room temperature (~22°C). For recording, a single slice, after
at least 2 h incubation, was transferred to a submerged recording
chamber with continuous flow (1.5-2 ml/min) of ACSF. The ACSF
contained (in mM) 124 NaCl, 26 NaHCO3, 1.25 NaH2PO4, 2.5 KCl, 4 CaCl
2, 4 MgSO4, 10 D-glucose, and 0.05 picrotoxin, gassed with 95%
O2-5% CO2 giving pH 7.4. All experiments were carried out at room temperature.
Whole-cell voltage-clamp recordings were performed from CA1 pyramidal
cells located 30-90 µm beneath the slice surface under visual
control using infrared dark-field illumination and a charge-coupled device (CCD) TV camera. The patch electrodes were made from hard borosilicate glass and filled with (in mM) 120 Cs-gluconate, 10 CsCl,
10 HEPES, 8 NaCl, 0.2 EGTA, 2 MgATP, 0.3 Na3GTP,
and 10 phosphocreatine (pH 7.3 with CsOH, osmolarity 290-300
mosM), or 125 K-gluconate, 10 HEPES, 8 NaCl, 0.2 EGTA, 2 MgATP,
0.3 Na3 GTP, and 10 phosphocreatine (pH 7.3 with
KOH, osmolarity 290-296 mosM). The electrodes had resistance 3-5 M
when filled with internal solution. Whole-cell recording was made in
voltage-clamp mode using an Axopatch-1D (Axon Instruments, Foster
City, CA) and cells were voltage-clamped at
68 mV. To evoke synaptic
current, two glass electrodes filled with ACSF (300 K
) were
placed in the dendrite region 70 and 150 µm away from the cell body
layer to stimulate two separate groups of Schaffer collaterals. Stimuli (100 µs) were delivered alternatively to each input pathway through current output isolation units. The interval between stimuli in each
pathway was 6 s with a 3 s interval between pathways. Stimulation intensity was adjusted to produce an excitatory postsynaptic
current (EPSC) with an amplitude of ~100 pA at the beginning of
recording. Series and input resistances during the recording were
monitored every 3 s by measuring the peak and steady-state
currents in response to 2 mV, 38 ms depolarizing pulses. The series
resistance ranged from 6 to 12 M
. The input resistance ranged from
300 to 100 M
.
In all pairing protocols used for inducing LTP, a total of 200 stimuli were delivered to the test pathway during the depolarization. During all induction protocols, stimulation of the control pathway was stopped altogether. The specific LTP protocols were 1) four brief high-frequency tetani (50 pulses at 50 or 20 Hz per each; 4 s intervals) paired with a short depolarization (~15 s to 0 mV); 2) four brief high-frequency tetani (50 pulse of 50 or 20 Hz per each; 4 s intervals) paired with a long depolarization (~3 min to 0 mV) given either at the beginning or at the end of the long depolarization; 3) the standard pairing protocol: low-frequency stimulation (200 pulses, 1.4 Hz) paired with a long depolarization (~3 min to 0 mV).
Data were acquired using a 486 PC computer, Labmaster DMA ADC and program written in Axobasic. The amplitude of a synaptic response was calculated as the difference between the average of data points in a window before the stimulus and in a window around the peak of the synaptic response. The average of responses during a 5 min period before LTP induction was taken as the baseline, and all values were normalized to this baseline. The level of LTP was calculated from this normalized data as the average (over a 3 min period) at the times after LTP induction indicated in each case. Values were expressed as means ± SE. Two-tail paired and unpaired t-test were used for calculation of the statistical significance of differences. Drugs used included D-2-amino-5-phosphonovaleric acid (D-AP5, Research Biochemicals International) and picrotoxin (Sigma).
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RESULTS |
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Whole-cell recordings were made under visual control from CA1
pyramidal cells using a Cs+-based internal
solution (see METHODS), and test stimuli were given every
6 s to two independent pathways. In the form of the "standard"
pairing procedure for LTP induction used in our laboratory, 200 synaptic stimuli are given at 1.4 Hz during a ~3 min depolarization to 0 mV. We sought to test whether the same number of stimuli given at
high-frequency during a brief depolarization could also induce LTP. In
these experiments we first established a stable baseline for synaptic
responses. The pairing protocol was then given as follows: the holding
voltage was increased from 68 to 0 mV over a 3 s period. Immediately
after that a series of four high-frequency tetani (50 stimuli at 50 Hz)
were given at 4 s intervals to one of the pathways. The holding voltage
was then rapidly restored to
68 mV. The total period of
depolarization was thus around 15 s. Figure
1, A-C, shows that this brief
pairing with high-frequency stimulation induced only a small, transient potentiation. In most cases, this transient potentiation was seen in
both the stimulated (test) and unstimulated (control) pathways (123 ± 9.3% of the baseline in the test and 118 ± 6% in
the control input, control 10 min after the pairing, n = 6; Fig. 1C). Ten minutes later, a standard pairing
protocol was used (200 stimuli at 1.4 Hz, depolarization to 0 mV
for ~3 min). This standard pairing produced a very large potentiation
(307 ± 36%, 30 min after induction, n = 6), and
the potentiation was specific to the stimulated pathway. The efficacy
of the standard pairing stimulus cannot be attributed to the greater
delay after the onset of whole-cell recording because the standard
protocol also induces large LTP if given after shorter delays (data not
shown). The failure of the high-frequency stimulation to induce LTP was
surprising. We explored several minor variants that used higher or
lower frequencies (100 Hz, n = 2; 20 Hz,
n = 8), and they also failed to induce strong LTP.
Rather, they induced only weak potentiation that was evident in both
test and control pathways. Ten minutes after 100 Hz tetani given during short depolarization, the level of potentiation was 116 ± 11% in
the test and 122 ± 5% in control input. Ten minutes after 20 Hz
tetani given during short depolarization, the potentiation was 138 ± 10% in the test and 119 ± 8% in the control input. The standard pairing protocol given later in the same experiments produced
robust synapse specific LTP (300 ± 7% in the test and 116 ± 8% in the control input, n = 8, 30 min after
induction).
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There are two major differences between the brief protocol and the standard protocol: the length of the depolarization and the frequency of synaptic stimulation. We next sought to determine which of these differences is important. In the experiments illustrated in Fig. 2, pairing was done with the long depolarization (~3 min) used in the standard pairing protocol, but synaptic stimulation was done using brief periods of high-frequency (50 Hz) stimulation. Two variants of this experiment were done, one in which the high-frequency stimulation was given at the beginning of the long depolarization, the other in which the same high-frequency stimulation was given at the end of the long depolarization. When stimulation was given at the beginning of the depolarization (Fig. 2A), there was virtually no potentiation (101.5 ± 15.7%, at 20 min after pairing, n = 5). However, when stimulation was given at the end of the depolarization (Fig. 2B), large input-specific LTP was induced (243 ± 22% in the test and 101 ± 11% in the control at 20 min after induction of LTP, n = 7). In a separate set of experiments when 20 Hz tetani were given at the end of the long depolarization, strong and synapse-specific LTP was also induced (197 ± 6% in the test and 102 ± 5% in the control at 20 min after induction of LTP, n = 7). These results indicate that LTP can be induced by a high-frequency stimulation during pairing, but that stimulation must be preceded by a long depolarization.
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The LTP that can be induced by the standard low-frequency pairing protocol can be blocked by 2-amino-5-phosphonovaleric acid (APV), a blocker of NMDA channels (n = 4, data not shown). To check whether the LTP produced by high frequency given at the end of a long depolarization was also dependent on the NMDA channels, we repeated these experiments in the presence of APV (50 µM). Figure 3A shows that this type of LTP was dependent on NMDA channels. We also tested whether the small, transient potentiation produced by pairing high-frequency stimulation with a brief depolarization was NMDA dependent. Figure 3B shows that it was not sensitive to APV (50 µM; 126 ± 6%, 10 min after tetanus). This level of potentiation was not significantly different from the level of potentiation produced in control ACSF (P > 0.05). It should also be noted that the NMDA-independent potentiation was not input specific (123 ± 9% potentiation on the control input 10 min after tetanus).
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It is of interest to know whether the properties of the synaptically evoked current during pairing give any hint about why LTP occurs under some conditions and not others. Figure 4A shows the currents during each of four tetani that were given early (left column) or late (right column) during a long depolarization. The records were from the same cell and the same input. Figure 4B shows an example from another cell, but where different inputs were used for early and late responses. These currents have the slow kinetics characteristic of the NMDA current and are almost completely blocked by APV (Fig. 4C). It can be seen that the current is larger when the tetani are given early than when they are given late during the depolarization. Similar results were obtained in all cells examined. One curious feature of these records that might be significant is that the tetanus given early sometimes evoked currents with seemingly supralinear summation on the rising edge, followed by a rapid collapse of the current. These features were not observed when the tetanus was given at the end of the depolarization. Possible interpretations of these complex kinetics are given in the DISCUSSION. An additional feature of these experiments is that the total outward membrane current falls gradually during a long depolarization (Fig. 4D).
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In a final series of experiments, we sought to determine whether the failure of the short depolarization protocol might be related to the use of Cs+ as the major internal cation. To examine this issue, we changed the internal solution to one containing K+ instead of Cs+. Figure 5 shows that, with the use of this solution, it became possible to induce LTP by pairing high-frequency stimulation (50 Hz) with a brief (15 s) depolarization to 0 mV. Although the amount of LTP is large (217 ± 21.6%, n = 6), it is significantly smaller than that induced by the standard pairing protocol with Cs+ as the internal cation (307 ± 36%, P < 0.05).
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DISCUSSION |
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We have investigated the requirements for inducing LTP using
variants of the standard pairing protocol. In the standard protocol, depolarization is imposed on the postsynaptic neuron by voltage clamping the membrane to 0 mV while stimulating the input axons at
fairly low frequency (0.1-2 Hz) for several minutes (Colino et
al. 1992; Manabe et al. 1992
; Otmakhov et
al. 1997
). Generally, Cs+ has been used
as the major internal cation to block several K+
channels and thereby make it easier to impose depolarization. We have
found that if we modified the standard protocol and used high-frequency
(20-100 Hz) stimulation given during a brief (15 s) depolarization to
0 mV, LTP cannot be induced. This failure is not simply due to the use
of high-frequency stimulation because LTP can be induced by
high-frequency stimulation given at the end of a long depolarization.
Furthermore, we have found that the failure to induce LTP during brief
depolarization is related to the use of Cs+-based
internal solution. When K+ is the major cation,
high-frequency stimulation given during a brief depolarization does
induce LTP. Thus it would appear that Cs+ somehow
prevents LTP induction when short high-frequency stimulation is paired
with short depolarization. Importantly, this blocking action of
Cs+ must then dissipate, because if
high-frequency stimulation is given at the end of a long
depolarization, LTP occurs normally.
These findings are quite surprising and cannot be simply
explained in terms of what is known about LTP.
Cs+ is used to block K+
channels and should therefore enhance depolarization. It has been shown
that depolarization up to the reversal potential for EPSC should
enhance LTP induction, not inhibit it (Perkel et al. 1993). Because we do pairing at a potential somewhat negative of reversal potential, any dendritic depolarization may occur during
synaptic stimulation (due to clamp failure) should cause the voltage to
become closer to the reversal potential and therefore lead to larger
potentiation rather than the very small potentiation we observed. The
use of brief high-frequency stimulation should bring about facilitation
of transmitter release and enhance LTP induction, not inhibit it.
Unfortunately, we are unable to provide any clear explanation of our
findings and can only point out several possibilities. A key unresolved
question is how the NMDA current itself is affected by various
induction conditions. The total synaptically evoked current during
pairing can be measured and gives hints about what might be going on.
This current is inward and appears larger when high-frequency
stimulation is given during a brief depolarization (or early during a
long depolarization) than when the same stimulation is given toward the
end of a long depolarization (Fig. 4). A curious and unexplained
feature of this current is that it often has an accelerating
rising edge and a transient character. The inward current is
strongly reduced by APV, suggesting that it is either largely NMDA
current or consists of both NMDA current and components triggered as a
consequence of current through the NMDA channel. On the assumption that
the current is primarily NMDA current, one might ask why the current
has such complex kinetics (Fig. 4). Recent work showed that the
NMDA conductance is enhanced by intracellular
Na+ (Yu and Salter 1998). So it is
possible that the odd kinetics of the current during the tetanus are
the consequence of Na+-dependent regulation. If
the total current is indeed largely NMDA current, then the failure to
induce LTP during brief depolarization can certainly not be
attributed to a reduced NMDA conductance, because the total synaptic
current is larger at early times (Fig. 4). If anything, the results
raise the possibility that there could be an optimal NMDA-mediated
Ca2+ entry and that if this optimum is exceeded,
smaller LTP will be induced (Lisman 1985
).
The transient nature of the synaptically evoked current during
pairing suggests a second interpretation in which the initial NMDA-mediated synaptic current triggers a dendritic
Ca2+ spike (assuming that clamping is not
adequate in the dendrites). There are indications that
Ca2+ elevations can inhibit the NMDA conductance
itself (Legendre et al. 1993), and this might explain
why LTP induction was blocked. Ca2+ spikes might
be much more difficult to induce when K+ channels
are functional, and this might explain why LTP can be induced when
K+ is the major internal cation. In a related
way, it might be supposed that inactivation of the
Ca2+ conductance by a long depolarization
prevents initiation of a Ca2+ spike late during
the depolarization, and this could explain why stimulation given late
during a long depolarization can induce LTP.
It is possible that the failure to induce LTP by high-frequency
stimulation given at the beginning of long depolarization (or during
short depolarization) is related to both the high frequency and the timing when the tetanus is given. In this case, short low-frequency stimulation would induce LTP even if it is given at the
beginning of long depolarization (or during short depolarization). Indeed the work has indicated that weak NMDA-dependent potentiation can
be produced when a small number of stimuli are given at low frequency
during brief depolarizations (Petersen et al. 1998). One
possible explanation in terms of the Ca2+ spike
model given above would be that only high-frequency stimulation can
produce sufficient depolarization to trigger a
Ca2+ spike.
Our work is not the first attempt to induce LTP by combining
high-frequency stimulation with imposed depolarization. A number of
works reported that LTP could be induced by pairing procedures of this
kind (Hjelmstad et al. 1997; Kato et al.
1993
; Manabe et al. 1993
; Perkel and
Nicoll 1993
). These experiments were done using
Cs+. However, the detailed timing of synaptic
stimulation during the depolarization is not clear. These previous
results may therefore not be incompatible with our findings. In a study
done under experimental conditions (5-wk-old guinea pigs, at 32°C)
(Chen et al. 1998
) somewhat different from those used
here (<3-wk-old rats, room temperature, Cs+ as
the major intracellular cation was used in both cases), it was possible
to induce LTP by delivering two 20 Hz trains given 10 s apart
during continuous depolarization lasting overall ~15 s. It is unclear
whether these technical differences are important. One previous paper
reported that intracellular Cs+ can block
tetanus-induced LTP (Haas and Rose 1984
). Their results are consistent with ours, although they used sharp microelectrode recording, and depolarizing current was not injected during the tetanus.
Our results indicate that there are aspects of the standard
pairing procedure that are not understood, specifically, the
requirement for long depolarization. This raises the question of
whether pairing protocols are a good model for the LTP induced by
tetanic stimulation or whether there are fundamental differences. One
difference is the size of the LTP induced by these different protocols.
We and others find that the standard pairing protocol produces a very large LTP. The synaptic response after LTP induction is often 400% and
can sometimes be 1,000% (Malinow 1991; Otmakhov
et al. 1997
; Otmakhov, unpublished results). In contrast, the
LTP induced by tetanic stimulation and field recording methods
typically show an LTP that is <200% and is often only 150%. We find
that the brief pairing protocol with K+ as the
internal cation evokes an LTP that is intermediate in size (220%).
Similarly, Lu et al. (1998)
using whole-cell recording with K+ found an LTP of 200%. The factors that
determine the size of LTP remain generally unclear. In particular,
there is no good explanation for why multiple tetani evoke LTP of the
field EPSP, which is at most 200% (saturation occurs with ~3
tetani), whereas pairing in Cs+ can evoke a much
larger LTP of up to 400%. Despite this difference in magnitude, there
are no other reasons to think that fundamentally different mechanisms
are involved in the LTP produced by pairing and tetanus protocols. Both
forms of LTP are dependent on NMDA channels and can be blocked by
kinase inhibitors (Otmakhov et al. 1997
).
In summary, we have studied the requirements for inducing LTP by pairing protocols. We unexpectedly found that when a tetanus is given at the beginning, but not at the end of depolarization, LTP induction is inhibited. The generation of this transient inhibition is related to the use of Cs+ as a major intracellular cation. These results are not expected from the known properties of LTP and suggest that there is a process that can powerfully regulate LTP induction that has not yet been identified.
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ACKNOWLEDGMENTS |
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The authors thank Dr. Nonna Otmakhova for useful comments.
This work was supported by National Institute of Neurological Disorders and Stroke Grants 5 R01 NS-35083 and 5 R01 NS-27337. The authors gratefully acknowledge the support of the W. M. Keck Foundation.
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FOOTNOTES |
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Address reprint requests to J. Lisman.
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 21 December 1998; accepted in final form 21 April 1999.
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REFERENCES |
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