Bidirectional Synaptic Plasticity Correlated With the Magnitude of Dendritic Calcium Transients Above a Threshold

R. J. Cormier, A. C. Greenwood, and J. A. Connor

Department of Neurosciences, University of New Mexico, School of Medicine, Albuquerque, New Mexico 87131


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cormier, R. J., A. C. Greenwood, and J. A. Connor. Bidirectional Synaptic Plasticity Correlated With the Magnitude of Dendritic Calcium Transients Above a Threshold. J. Neurophysiol. 85: 399-406, 2001. The magnitude of postsynaptic Ca2+ transients is thought to affect activity-dependent synaptic plasticity associated with learning and memory. Large Ca2+ transients have been implicated in the induction of long-term potentiation (LTP), while smaller Ca2+ transients have been associated with long-term depression (LTD). However, a direct relationship has not been demonstrated between Ca2+ measurements and direction of synaptic plasticity in the same cells, using one induction protocol. Here, we used glutamate iontophoresis to induce Ca2+ transients in hippocampal CA1 neurons injected with the Ca2+-indicator fura-2. Test stimulation of one or two synaptic pathways before and after iontophoresis showed that the direction of synaptic plasticity correlated with glutamate-induced Ca2+ levels above a threshold, below which no plasticity occurred (~180 nM). Relatively low Ca2+ levels (180-500 nM) typically led to LTD of synaptic transmission and higher levels (>500 nM) often led to LTP. Failure to show plasticity correlated with Ca2+ levels in two distinct ranges: <180 nM and ~450-600 nM, while only LTD occurred between these ranges. Our data support a class of models in which failure of Ca2+ transients to affect transmission may arise either from insufficient Ca2+ to affect Ca2+-sensitive proteins regulating synaptic strength through opposing activities or from higher Ca2+ levels that reset activities of such proteins without affecting the net balance of activities. Our estimates of the threshold Ca2+ level for LTD (~180 nM) and for the transition from LTD to LTP (~540 nM) may assist in constraining the molecular details of such models.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Learning and memory are widely associated with enduring forms of synaptic plasticity such as long-term potentiation (LTP) (Bliss and Gardner-Medwin 1973) and long-term depression (LTD) (Fujii et al. 1991), which respectively enhance and diminish synaptic strength. Despite their opposing effects, LTP and LTD share some common biochemical mechanisms, including intracellular Ca2+ transients. Here, we focus on the relation of transient Ca2+ levels to LTP and LTD in postsynaptic CA1 pyramidal neurons.

A wealth of data demonstrates the requirement for Ca2+ transients in synaptic plasticity. Direct measurements of Ca2+ using ratio-fluorescence microscopy showed micromolar levels in postsynaptic neurons in response to tetanic stimulation (Müller and Connor 1991; Petrozzino et al. 1995; Yuste and Denk 1995). In addition, raising postsynaptic Ca2+ directly by photolysis of caged-Ca2+ compounds loaded into postsynaptic neurons induced LTP as well as LTD (Malenka et al. 1988; Neveu and Zucker 1996). On the other hand, inhibiting Ca2+ transients by loading postsynaptic neurons with calcium chelators, such as EGTA (Lynch et al. 1983) or caged-chelators (Malenka et al. 1992), inhibited induction of LTP and LTD. Furthermore induction protocols associated with higher Ca2+ levels induced LTP, while protocols associated with more moderate Ca2+ levels induced LTD (Hansel et al. 1996, 1997). Despite these compelling studies relating Ca2+ to synaptic plasticity, a quantitative relationship between measured Ca2+ levels and synaptic plasticity in the same neurons has not been directly demonstrated, prompting the present study.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

We prepared coronal hemispheric brain slices (350- to 400-µm thick) from adult male Sprague-Dawley rats (Harlan; 5-10 wk old) by standard humane methods involving quick decapitation of anesthetized animals (Connor and Cormier 2000). The slices were then maintained at 28°C in artificial cerebrospinal fluid (ACSF; containing, in mM: 126 NaCl, 3 KCl, 1.25 NaH2PO4, 26 NaHCO3, 1 MgCl2, 2 CaCl2, and 10 dextrose; gassed with 95% O2-5% CO2) until use. All chemicals were from Sigma.

For experiments, individual slices were placed in a submersion chamber and perfused at 1-2 ml/min with ACSF at 31 ± 0.5°C. Intracellular recordings were made from CA1 neurons ~75 µm below the slice surface, using glass micropipettes with tips filled with 12 mM fura-2 and shanks filled with a solution of 3 M KCl and 1 M K-acetate. The electrode resistance was initially ~200 MOmega with fura-2 present but dropped to ~120 MOmega in ~20 min before data collection. An Axoclamp 2A amplifier in bridge mode was used to record membrane potential. To evoke excitatory postsynaptic potentials (EPSPs), one or two monopolar stimulating electrodes were placed in stratum radiatum with the tip (20 µm diameter) ~100 µm below the slice surface. Stimulating electrodes were placed in the proximal third of s. radiatum to activate afferents on proximal dendrites or in the distal third to activate afferents on distal dendrites. Pairs of EPSPs separated by 50 ms were evoked by constant-current pulses (50-100 µA) every 15 s.

After stopping the afferent stimulation used to evoke baseline EPSPs, an iontophoresis pipette (1 M glutamate, pH 7.0, ~10 MOmega ) was positioned 20-50 µm from the primary apical dendrite of the fura-filled neuron. Glutamate was ejected by five iontophoretic pulses (duration: 10 s, amplitude: 4 µA, interval: 60 s), and the pipette was then withdrawn from the slice >= 5 min before afferent stimulation was resumed. In control experiments, 1 M NaCl was used in place of glutamate.

Ca2+ levels were measured by successively illuminating fura-filled neurons with 350 and 380 nm wavelength light for 200 or 400 ms and imaging the resulting fluorescence with a cooled-CCD camera on an upright microscope (Zeiss Axioskop) with a water-immersion, ×40 objective lens. Ca2+ levels were calculated by standard ratiometric methods after fluorescent-background correction, assuming a Ca2+-affinity KD of 225 nM for fura-2 (Grynkiewicz et al. 1985). Fura levels were estimated as follows. Cells were injected with 0.7 nA for 20 min. Conservatively ignoring dilution by the backfilling solution and assuming a cell volume of 3 nl, the final intracellular [fura] would be ~60 µM, by the following equation: [Fura-2] = (n · i · t)/(z · F · v). Here n is the transport number (0.1), i is the iontophoresis current, t is the time of iontophoresis, z is the net charge, F is Faraday's constant, and v is the cell volume. A published comparison of similar sharp-electrode methods to whole cell fura-injection suggests that our methods achieved intracellular fura levels of 20-30 µM (Petrozzino et al. 1995), unlikely to slow the already slow Ca2+ kinetics associated with glutamate iontophoresis.

Statistical comparisons were made using Tukey's two-tailed t-test, corrected for multiple comparisons by Bonferroni's method (Snedecor and Cochran 1980). All error bars show the SE. Error propagation to determine uncertainty in plasticity factor followed standard methods (Bevington and Robinson 1992).


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

We measured intracellular Ca2+ levels during iontophoresis of glutamate onto the apical dendrites of CA1 pyramidal cells in coronal brain slices from rats (n = 67). The bulk of these experiments (n = 53) served to characterize the ensuing Ca2+ transients, guiding a study of the relation between these transients and effects on transmission in 20 synaptic inputs to 14 cells with one (n = 8) or two (n = 6) stimulated synaptic pathways. Cells that were included in this study had stable resting potentials more negative than -65 mV and stable resting Ca2+ levels averaging 71 ± 11 (SE) nM.

Glutamate iontophoresis (10 s, 4 µA) typically depolarized the membrane to values between -50 and -10 mV (data described in this paragraph not shown) (see Cormier and Kelly 1996; Cormier et al. 1993). After a glutamate pulse, cells repolarized and sometimes also hyperpolarized, perhaps from activation of Ca2+-gated potassium channels (Hotson and Prince 1980). Action potentials were often observed during the first two glutamate pulses in a train (at 1 pulse/min) but rarely during later pulses, though these later pulses were associated with more rapid depolarization than were the first two pulses. Control experiments with equimolar substitution of sodium chloride for glutamate demonstrated that iontophoresis current (up to 10 times that used here) had no detectable effect on membrane potential, intracellular Ca2+, or synaptic transmission (n = 3) (see Cormier and Kelly 1996; Cormier et al. 1993).

Focusing first on the temporal information in our imaging data, we measured Ca2+ levels in the region of dendrite to which glutamate was applied in most experiments, ~70 µm from the soma (marked in Fig. 1D, ---). In this region, Ca2+ levels typically rose during 6-10 s of the standard 10-s pulse, reached a peak level between 100 and 600 nM, and returned to basal levels by 5-120 s after the pulse (Fig. 1A: data from typical cell). Increasing the iontophoretic current produced higher Ca2+ levels (Fig. 1B: data from same cell) and very prolonged Ca2+ transients in some cases, as reported elsewhere (Connor and Cormier 2000). In our study relating Ca2+ transients to plasticity, we set the iontophoretic current at an intermediate level (4 µA) that produced Ca2+ levels <1 µM to reduce the risk of excitotoxic effects.



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Fig. 1. Glutamate iontophoresis elicited reproducible Ca2+ transients. A: time course of a representative transient elicited by a single 10-s pulse of glutamate (: 4.2 µA). Glutamate was applied and Ca2+ was measured ~70 µm from the apical base. B: increasing the glutamate iontophoresis current increased the peak Ca2+ level in the same cell (: 3.3, 4.2, and 5 µA). Only 4-µA pulses were used to generate the Ca2+ transients that were related to plasticity in this study. C: a series of 5 glutamate pulses () had somewhat different effects in different cells. In 2 typical cells, successive Ca2+ transients were either larger (C1) or similar (C2). D: spatial profiles of a Ca2+ transient induced by a single glutamate pulse and a fluorescent image of the fura-2 filled apical dendrite (excited at 380 nm) from which the data came. The numbers to the left of the profiles indicate the measurement times relative to the start of the 10-s glutamate pulse. Vertical scale bar: 100 nM Ca2+. The location of the horizontal scale bar (37 µm) indicates the region closest to the glutamate pipet (~70 µm from the apical base), coincident with the location of the peak Ca2+ levels (---). E: spatial profile of a Ca2+ transient peaking at the 8-s point during glutamate application to the distal dendrite (~250 µm from the apical base), as indicated by location of the horizontal scale bar (37 µm) in the accompanying fura-2 picture. Not surprisingly, Ca2+ levels peaked more distally (in the vicinity of the glutamate pipette), when glutamate was applied distally instead of to the proximal dendrite. The flat profile shows baseline Ca2+ levels. Vertical scale bar: 100 nM Ca2+.

A series of five successive pulses of glutamate at one-minute intervals led to multiphasic Ca2+ transients (Fig. 1C). In some experiments, the basal Ca2+ level was higher for successive pulses (n = 20 of 67; typical data shown in Fig. 1C1). Also the peak Ca2+ level often increased with successive pulses (n = 38 of 67; Fig. 1C1). Saturation of glutamate uptake (Asztely et al. 1997; Mennerick and Zorumski 1995) or of Ca2+ sequestration or extrusion, among other mechanisms, may have contributed to these phenomena. On the other hand, in many experiments, the peak Ca2+ level was very similar from pulse to pulse (n = 29 of 67; Fig. 1C2 shows typical data).

The spatial distribution of glutamate-induced Ca2+ transients within cells was inhomogeneous, suggesting that synapses at widely separated locations might show synaptic plasticity of differing magnitude and direction. In our initial experiments, we placed the iontophoresis pipet at ~70 µm along the proximal apical dendrites, producing Ca2+ transients centered near the point of application. Typically, the initial peak was closer to the soma than the pipette was (Fig. 1D, 4 s), possibly the result of activating dendritic voltage-gated Ca2+ channels (Miyakawa et al. 1992; Regehr and Tank 1992). However, by 8 s, the Ca2+ levels most proximal to the soma had declined, leaving an increasingly well-defined peak centered near the glutamate source. In some later experiments (n = 6), the iontophoresis pipette was placed in distal stratum radiatum, ~250 µm from the base of the apical shaft. Distal glutamate led to Ca2+ transients resembling those that followed proximal glutamate application, except that the location of maximal change tended to be shifted distally. An example is shown in Fig. 1E.

Having examined the effects of glutamate application on dendritic Ca2+, and as a prelude to correlating glutamate-induced Ca2+ levels with synaptic plasticity, we next asked what does glutamate iontophoresis do to synaptic transmission near and away from the application site in fura-filled cells? To answer this question, we used the fura-filled micropipette to record evoked intracellular EPSPs before and 15 min after glutamate application. In eight experiments, a single stimulating electrode was placed close to the cell-body layer to activate synapses on the proximal dendrites, the site of glutamate application (Fig. 2A1). These experiments were classified into three groups based on their plasticity factor, defined as mean slope of 20 consecutive EPSPs arbitrarily selected from 40 to 60 min after the end of iontophoresis, normalized with respect to a similarly calculated mean baseline slope: LTP >=  115% (A); LTD <=  85% (B); 85% <=  no plasticity <=  115% (C). These specific group boundaries were chosen because the 95% confidence interval for the baseline mean was 100 ± ~15% in every experiment. However, varying the group boundaries to 100 ± 10 or 100 ± 30% had no effect on group membership. Furthermore due to the stability of the recorded EPSPs, the 95% confidence interval for a plasticity factor was never more than 7.1% of the plasticity factor for any pathway in this study, and the mean normalized confidence interval for all plasticity factors was only 2.9%. Thus no 95% confidence interval for a pathway's plasticity factor ever crossed a group boundary as initially defined. Applying this classification scheme to EPSPs that were evoked in eight single-pathway experiments, we observed LTD of synaptic transmission in five cells (to 56 ± 10% of baseline) and LTP in three cells (to 166 ± 11%), as summarized in Fig. 2B (single pathway). These results showed that glutamate applied to the proximal dendrites affected synaptic transmission in that dendritic region, prompting a series of two-pathway experiments with distal glutamate application.



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Fig. 2. Glutamate-induced synaptic plasticity. A: schematics show electrode placement for 1 (A1) and 2 (A2) pathway experiments. Stimulating electrodes on Shaffer fibers are indicated by "Stim." The intracellular recording electrode ("Fura") was present throughout the experiment; the glutamate iontophoresis pipette ("Glu") was present only at the time of glutamate delivery. B: summary of plasticity in experiments sorted by stimulation pathway. Each point represents 1 pathway's plasticity factor: the mean of 20 consecutive slopes 40-60 min after iontophoresis, normalized with respect to a similar baseline mean. , indicate group means. Values from the proximal and distal dendrites in each 2-pathway experiment are connected (---). C: data from a 2-pathway experiment in which the distal pathway underwent LTP () and the proximal pathway underwent no plasticity (). Each point shows the average slope of 4 consecutive EPSPs, normalized with respect to 20 consecutive baseline EPSPs. Some of the distal EPSPs triggered spikes after LTP (e.g., truncated response in inset). C-F, insets: waveforms from the times indicated by a and b (scale bars: 10 mV, 10 ms). Left waveforms show the voltage response to a hyperpolarizing current pulse (0.1 nA), monitored to rule out confounding changes in recording quality or cell viability. Right waveforms show EPSPs. D-F: the time course of the group mean ± SE of the normalized EPSP slope was plotted for each of 3 plasticity groups, defined by plasticity factor. The 3 groups were LTP >=  115%, LTD <=  85%, and 85% < no plasticity <115% (n = 7, 7, and 6). The 5 vertical bars on each x-axis represent pulses of glutamate. Afferent stimulation was suspended from 5-10 min before iontophoresis until 10-20 min after the last pulse.

In these two-pathway experiments (n = 6), the first stimulating electrode in proximal s. radiatum was complemented with a second one in distal s. radiatum, as diagrammed in Fig. 2A2. Glutamate was applied to the dendrites in the distal third of s. radiatum (as in Fig. 1E) to increase the likelihood that EPSPs from the distal synapses would undergo LTP (as in the representative experimental time course in Fig. 2C). We expected that we might also observe plasticity in the proximal pathway due to the activation of voltage-gated Ca2+ channels on the primary apical dendrites (Miyakawa et al. 1992; Regehr and Tank 1992). In fact, when we applied our plasticity-factor classification scheme to the proximal pathway, we observed no plasticity (98 ± 3%) in four cells, LTD (72%) in one cell, and LTP (302%) in one cell1 at 40-60 min after the glutamate treatment. Meanwhile, the distal pathway showed no plasticity (106 ± 1%) in two cells, LTD (32%) in one cell, and LTP (231 ± 51%) in three cells. These data are plotted on the right in Fig. 2B, with proximal and distal plasticity factors from individual neurons connected by lines. Thus both sides of Fig. 2B show that the range of observed plasticity factors was well suited to answer the question: do the Ca2+ levels reached during glutamate iontophoresis and subsequent effects on synaptic transmission correlate as they would in a simple model in which Ca2+ caused these effects?

To answer this question, we first pooled the electrophysiological data from the 8 single-pathway experiments and the six two-pathway experiments to obtain a total of 20 experimental time courses of EPSP slope, divided as before into three groups according to plasticity factor. The average time courses for the LTP, LTD, and no-plasticity groups of pooled data are plotted in Fig. 2, D-F, respectively (n = 7, 7, and 6). Next, we sorted the glutamate-induced Ca2+ transients into the same three groups. In these data, the distal or proximal location of stimulus-activated synapses was inferred from the position of the stimulating electrode. To quantify the Ca2+ levels that may have affected synaptic transmission, we analyzed the iontophoresis imaging data with measurement boxes placed on the dendrite near these putative synaptic locations. Figure 3A shows such box placement on a typical cell in a two-pathway experiment, while Fig. 3B shows the average multiple-pulse Ca2+-level time course obtained from all such boxes for synaptic pathways that exhibited LTP. For comparison between pathways or groups of pathways, multiple-pulse time courses were collapsed to a single-pulse time course by averaging across the five glutamate pulses, as illustrated for the LTP group in Fig. 3C. Then, the three points at 6, 8 and 10 s during the single-pulse time courses were averaged to obtain a single measure of peak-Ca2+ level for each pathway or for each group in the case of the averaged data. Fura saturation had minimal effect on the average peak-Ca2+ level, as at most one of five peaks in the average approached 1 µM (e.g., 850 nM in Fig. 1C1).



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Fig. 3. Plasticity correlated with Ca2+ levels above a threshold. A: fluorescent image of a fura-filled dendrite excited by 380 nm light (scale bar: 37 µm). Boxes show locations of Ca2+ measurements for the proximal and distal synapses (left box: ~70 µm from apical base, right box: ~250 µm). B: time course of the mean Ca2+ level in the LTP group during the 5 glutamate pulses (horizontal bars; n = 7). Recovery to baseline Ca2+ levels was verified with a delay (R = 2.3 ± 0.7 min after last pulse). C: the Ca2+ transient obtained by averaging across pulses and within the LTP group. The horizontal bar represents the canonical pulse. The filled circles indicate the times (6, 8 and 10 s) at which transients were sampled for averaging to calculate "peak" Ca2+ levels for all pathways and groups. D: plasticity factor was plotted against peak Ca2+ level for each pathway (+, single pathway; , proximal pathway of 2; x, distal pathway). The data were fit with a line and a 4th-order polynomial to elucidate nonlinear trends. The vertical dashed lines show estimates of the Ca2+ threshold for LTD (180 nM) and the half-way point between LTD and LTP (541 nM), discussed in the text. The diamond represents mean basal [Ca2+]. E: group-mean peak Ca2+ levels are plotted for the LTP, LTD, and no-plasticity groups (solid bars) and the high-Ca2+ and low-Ca2+ no-plasticity subgroups (striped bars). Pair-wise comparisons were made among the 3 groups. LTP was associated with higher Ca2+ levels than LTD (P < 0.05; n = 7 and 7). The Ca2+-dependent definition of the no-plasticity subgroups precluded conventional comparisons involving them. However, the difference between their mean peak Ca2+ levels was large relative to the SEs, and dividing the no-plasticity group into subgroups accounted for most of the group SE (see error bars; n = 3 and 3). F: plasticity factor is shown for the same groups.

Looking first at the results for each pathway separately, Fig. 3D shows a scatter plot of percentage plasticity factor versus the calculated peak Ca2+ level. Different symbols indicate data collected from single (proximal) pathways and distal and proximal pathways in two-pathway experiments (see legend). Despite a statistically adequate fit (r = 0.60, P < 0.005), a linear model was rejected on physiological grounds because it predicted the ongoing induction of profound LTD at resting Ca2+ levels, which of course was not observed. A linear model constrained to start from no plasticity at Ca2+ baseline was rejected (r = 0.12, P < 0.59, not plotted). A fourth-order polynomial fit is also plotted in Fig. 3D to help visualize the multiphasic dependence of plasticity on Ca2+ that a satisfactory model would need to explain. One satisfactory class of models would involve a balance of effectors with opposing effects on synaptic strength, with the activity of at least one effector being reset by Ca2+ levels above a threshold (Grzywacz and Burgi 1998; Lisman 1989). In such a model, the no-plasticity group would be predicted to consist of two subgroups, one that was exposed to Ca2+ levels below that required to reset the enzyme activities and a second that was exposed to Ca2+ levels at which the activities of individual enzyme molecules were reset without affecting the balance of activities.

Turning next to the summary data shown in Fig. 3E, pair-wise comparisons among all three plasticity groups showed that the peak Ca2+ levels in the LTP group were significantly larger than in the LTD group (P < 0.05, n = 7 and 7). Thus our data support the conventional view that low Ca2+ levels lead to LTD, while higher levels lead to LTP. In contrast, the mean peak Ca2+ level of the no-plasticity group barely differed from that of the LTD group (not significant, P > 0.9, n = 6 and 7). Meanwhile, the standard deviation (SD = 220) of the peak Ca2+ levels in the no-plasticity group was so large that its mean was not significantly different from that of the LTP group (P > 0.2, n = 6 and 7). However, as suggested by Fig. 3D, this extreme variability within the no-plasticity group could be accounted for by dividing it into two subgroups ("low" and "high" Ca2+) with markedly different average peak Ca2+ levels with small standard deviations [147 ± 20 and 541 ± 63 (SD) nM, n = 3 and 3].

Of course, the significance of this difference in Ca2+ levels could not be tested because membership in the subgroups depended on Ca2+, violating the standard assumption of independence. However, the apparent bimodality of the distribution of peak Ca2+ levels within the no-plasticity group was supported by the failure of a normal model to fit this distribution (P 0.001, chi 2 = 35). Also, despite the marked difference between the Ca2+ levels associated with the high-Ca2+ and low-Ca2+ no-plasticity subgroups, it followed from the no-plasticity group's definition that the mean plasticity factors associated with these subgroups were statistically indistinguishable (Fig. 3F, P = 0.27, n = 3 and 3). Finally, although the mean peak Ca2+ level associated with LTP was not much higher than that of the high-Ca2+ no-plasticity subgroup (Fig. 3E, no statement of significance possible), this result is not surprising because the iontophoretic current (4 nA) was selected to achieve Ca2+ levels below 1 µM, typically ~500 nM. Previous work showed that larger iontophoretic currents led to LTP (Cormier et al. 1993) and larger Ca2+ transients (Connor and Cormier 2000).

Returning to Fig. 3D, the fourth-order polynomial fit provided initial estimates for the Ca2+ levels associated with the low-Ca2+ transition between no plasticity and LTD (165 nM) and the 100% cross-over point between the LTD and LTP groups (485 nM). To address the arbitrariness of the fourth-order polynomial, we also estimated each of these transition points by a second method. For a second estimate of the low-Ca2+ transition, linear interpolation between the data points straddling the 85% plasticity line yielded an 85% intercept of 180 nM. For a second estimate of the 100% cross-over point, we took the mean peak Ca2+ level in the high-Ca2+ no-plasticity subgroup (541 nM). As they include no arbitrary assumptions, we accepted these second estimates in place of the estimates from the fourth-order polynomial, with which they are in reasonable agreement.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

This paper reports that glutamate iontophoresis elevated intracellular Ca2+ in the apical dendrites of CA1 pyramidal cells and induced synaptic plasticity in the same cells. Although glutamate iontophoresis was previously shown to induce synaptic plasticity in hippocampal cultures (Malgaroli and Tsien 1992) and slices (Cormier and Kelly 1996; Cormier et al. 1993), the present study included Ca2+ imaging, allowing us to relate glutamate-induced Ca2+ levels to synaptic plasticity quantitatively.

Glutamate iontophoresis, Ca2+, and cell viability

The iontophoretic protocol produced Ca2+ transients that returned to baseline within 120 s after a series of five 10-s pulses at 1-min intervals. Earlier work suggested that exposure to appropriate Ca2+ levels for 5 min was sufficient to induce LTD (Mulkey and Malenka 1992), while only 2.5 s of appropriate Ca2+ levels was reported to be required for LTP induction (Malenka et al. 1992). With these putative temporal requirements in mind, our protocol was designed to reduce the importance of Ca2+-transient kinetics as a factor affecting synaptic plasticity and to result in LTD or LTP, depending on the peak Ca2+ levels achieved. The between-cell variability that we observed in peak Ca2+ levels is consistent with the many sources of variability inherent in the iontophoretic application of glutamate in the slice, including cell depth, variations in local perfusate flow, and distance from the dendrite. This variability is also consistent with an earlier study of glutamate-induced Ca2+ transients in dentate granule cells, not involving synaptic plasticity (Kudo et al. 1987).

In contrast to our protocol, which required five sustaining 10-s pulses of glutamate to achieve ~5 min of elevated Ca2+, a neurotoxic exposure of acutely dissociated hippocampal cells to glutamate or N-methyl-D-aspartate (NMDA) can induce Ca2+ transients that last for many minutes after iontophoresis (Connor et al. 1988; Wadman and Connor 1992). Although we found that a similarly prolonged recovery could be achieved using glutamate iontophoresis in hippocampal slices, in our hands it required twice as much iontophoresis current as was used in the present study (Connor and Cormier 2000). Thus our five-pulse protocol was designed to achieve relatively prolonged Ca2+ transients, while avoiding the prolonged single-pulse recovery that we feared would be associated with neurotoxic effects. As the higher Ca2+ levels were associated with LTP, whereas sick cells might be expected to show LTD, we infer that we successfully avoided complicating neurotoxic effects. This inference is further supported by our observation of stable resting potential and input resistance throughout the experiments.

Glutamate iontophoresis and synaptic plasticity

The clear correlation between Ca2+ levels and plasticity that we observed suggests that Ca2+ is a key mediator of plasticity induced by glutamate iontophoresis. However, our experiments do not rule out effects of glutamate on synaptic plasticity that are not entirely mediated by postsynaptic Ca2+. For example, in addition to their role in mobilizing intracellular Ca2+ (Jaffe and Brown 1994; Linden et al. 1994; Llano et al. 1991; Murphy and Miller 1988), metabotropic glutamate receptors can affect synaptic transmission in a variety of other ways (Conn and Pin 1997). Also, glutamate iontophoresis may release neuromodulators, such as nitric oxide or arachidonic acid (Medina and Izquierdo 1995). Addressing the possibility that these substances or glutamate contributed to presynaptic plasticity in our experiments, we note that we did not stimulate afferent fibers during glutamate exposure and that previous work established that presynaptic action potentials and transmitter release are not required for the induction of plasticity by glutamate iontophoresis (Cormier et al. 1993).

In contrast to the absence of synaptic stimulation in our induction protocol, an earlier study found that sustained dendritic Ca2+ transients elicited by action potentials evoked with current pulses at 3 Hz for 5 min were insufficient to elicit LTD without paired synaptic stimulation (Christie et al. 1996). This study also differed from the present study in that Delta F/F measurements of fura-2 fluorescence were reported instead of ratiometrically determined Ca2+ levels, preventing a direct comparison of results. However, in harmony with the present study, these authors interpreted their results to suggest that Ca2+ played a critical role in the efficacy of their LTD-induction protocol, noting that LTD induction was blocked by nimodipine, Ni2+, or APV (independently) and suggesting that the essential role of paired synaptic stimulation may have been to enhance Ca2+ influx specifically into spines (see also Yuste and Denk 1995). These localized synaptic Ca2+ transients apparently did not add significantly to the dendritic Delta F/F observed with action potentials alone. In contrast, our admittedly less "physiological" induction protocol may have permitted a more direct assessment of the Ca2+ levels affecting transmission, as glutamate-induced Ca2+ transients are relatively homogeneous spatially, especially in fura-filled cells (see following section).

Ca2+ measurements with fura-2

Two concerns naturally arise regarding the use of fura-2. First, its high affinity for Ca2+ (~225 nM) could lead to saturation. However, saturation occurs at ~1 µM Ca2+, above the levels that we obtained during plasticity induction. Second, it has been reported that exogenous Ca2+ buffers, like fura-2, can interfere with the induction of LTP by diminishing the peak amplitude of brief Ca2+ transients while incidentally prolonging the recovery (Hansel et al. 1996, 1997; Kimura et al. 1990; Lynch et al. 1983; Malenka et al. 1988). However, the use of fura-2 in our experiments is unlikely to have interfered with the relation between our Ca2+ measurements and plasticity for the following reasons. Similar methods were reported to lead to intracellular fura levels of 20-30 µM (Petrozzino et al. 1995), while our conservatively high calculated estimate was 60 µM. Also, as the postsynaptic Ca2+ transients were induced by iontophoresis of glutamate at some distance from the putative synaptic location, the transients were expected to be inherently slow, reducing concerns that fura-2 would further retard their kinetics or diminish their magnitude. Granted, it is likely that mobile fura-2 molecules tended to keep the Ca2+ level at the synapse and in the dendrite more homogeneous than if local Ca2+ level depended only on local Ca2+ channels and other physiological sources (Carnevale and Rosenthal 1992). However, this enhanced homogeneity coupled with our slow Ca2+-transient kinetics would have the advantage of reducing extra variance in the relationship between measurements of plasticity and dendritic Ca2+. In summary, the main goal of these experiments was to study effects of Ca2+-transient magnitude in relative isolation from duration effects, irrespective of "normal" Ca2+-level kinetics and spatial inhomogeneity.

Synapse localization

Nonetheless, imprecision in synapse localization must have contributed to variance in the relationship between the Ca2+ measurements and synaptic plasticity, tending to obscure our results. To limit this source of variance, we placed a fine stimulating electrode (20 µm diameter tip) as close as 0.5 mm from the dendrite and measured "plasticity-related" Ca2+ along a ~40 µm length of dendrite. Also relevant to the probable magnitude of this source of variance, the Ca2+ gradient along the apical dendrite was typically ~1 nM/µm within 50 µm of the presumed synaptic locations at the time of peak Ca2+. In spite of this variance, however, some significant and intriguing results emerged from the data.

Distinct Ca2+ thresholds for LTD and LTP

These data provided indirect support for a class of models in which Ca2+ levels above a threshold reset the activity-level balance between opposing molecular effectors of synaptic strength (e.g., Lisman 1989). Specifically, we observed LTD at lower peak Ca2+ levels and LTP at higher levels (means: 335 ± 46 and 574 ± 27 nM, P < 0.05). Our data also provide estimates of the transition Ca2+ levels that separate the LTD plasticity group from the no-plasticity subgroup on the low-Ca2+ side (180 nM) and from the LTP group on the high-Ca2+ side (541 nM). The latter transition level is the mean peak Ca2+ level of the no-plasticity subgroup intermediate between LTD and LTP, with peak Ca2+ levels ranging from 450 to 600 nM. Ours is the first study to document the existence of this no-plasticity subgroup, an essential prediction of a Ca2+-based version of the BCM learning rule (Bienenstock et al. 1982).

In this study, we focused on peak Ca2+ levels because prior evidence suggested that peak Ca2+ levels were related to synaptic plasticity (Hansel et al. 1996, 1997; Malenka et al. 1988; Müller and Connor 1991; Neveu and Zucker 1996; Petrozzino et al. 1995; Yuste and Denk 1995). Of course, it is likely that Ca2+ levels during the decay phase also affect synaptic plasticity. However, because decay is fast when Ca2+ is high, most of the recovery phase (the long tail) is similar for a transient peaking at 600 nM and one peaking at 300 nM, for example. To a rough approximation, the main difference between such transients is the peak at 600 nM and swift decay to 300 nM. This difference in peak Ca2+ may contribute a difference in net plasticity, while the remaining decay phase in common may make similar contributions to net plasticity. Our use of five relatively closely spaced transients, interrupting the first four decay phases, may accentuate the effects on plasticity of differences in peak Ca2+.

Relating our findings to earlier work, this laboratory previously showed that increasing Ca2+ levels to 20 µM by tetanic stimulation (Petrozzino et al. 1995) or bath application of tetraethylammonium (Petrozzino and Connor 1994) reliably induced robust LTP. These early studies differ from the present study in that a low-affinity indicator (mag-fura-5) was used to focus exclusively on LTP induced by large Ca2+ transients in spines and fine dendrites. Also an early study where Ca2+ transients were produced by flash-photolysis of the caged-Ca2+ compound nitr-5 that was injected into the postsynaptic cell showed that 2-4 µM Ca2+ induced only LTP (Malenka et al. 1988). A more recent nitr-5 study suggested that Ca2+ levels of 300-500 nM induced LTD and LTP in separate cells (Neveu and Zucker 1996). Unlike the present study, this study did not measure the Ca2+ levels resulting from the plasticity-induction protocol (flash photolysis) and, instead, estimated Ca2+ from the duration of the flash, cell depth, and model assumptions. Further support for distinct Ca2+ thresholds came from imaging studies where different stimulation protocols induced LTD or LTP and caused relatively small and large Ca2+ transients, respectively (Hansel et al. 1996, 1997). However, the synaptic plasticity and imaging measurements in these studies were conducted in separate experiments and calibrated estimates of Ca2+ levels were not provided (Hansel et al. 1996, 1997).

Another study used two tetanic protocols in the presence or absence of picrotoxin to evoke Ca2+ transients that were related to LTP and LTD (Otani and Connor 1998). Several factors motivated the execution of the present study with a single protocol of glutamate iontophoresis, achieving results that extend the results of these earlier experiments without contradiction. Compared to glutamate iontophoresis, tetani are likely to activate different mechanisms of Ca2+ entry and additional plasticity factors. As a result of recruitment of additional pathways, the spatial domain of the Ca2+ transients associated with tetani may not accurately reflect the synaptic locations under test conditions. Also, Ca2+ flows directly into the postsynaptic structure during tetani, reaching transient local peaks that are likely to affect plasticity but are difficult to measure accurately. In contrast, glutamate iontophoresis generated slow and relatively homogeneous Ca2+ increases that could be measured accurately. Despite these methodological differences, however, no contradiction exists between the results of these two studies. The mean peak Ca2+ associated with LTD in the earlier tetanic study was ~460 nM, consistent with the range of peak levels determined for LTD in the present study (180-500 nM). The tetanic value could be a little high within the present range because Ca2+ levels were near their peak for only ~10 s with the tetanic protocol or because baseline Ca2+ was relatively high in the earlier study. Conditions of insufficient Ca2+ increase for plasticity were also consistent between studies. In the earlier study, no plasticity was observed under tetanic conditions leading to a peak Ca2+ increase of ~30 nM from baseline, safely below our new threshold for the induction of LTD (an increase of 120 nM from baseline). Finally, the mean Ca2+ increase associated with LTP in the earlier tetanic study was >1 µM, well above our minimum threshold for LTP induction.

In summary, our new results are consistent with earlier data while going beyond them to show, with one induction protocol and one Ca2+ indicator, that low Ca2+ levels induce LTD, high Ca2+ levels induce LTP, and intermediate Ca2+ levels are associated with a no-plasticity domain. In general, of course, Ca2+ thresholds for the induction of synaptic plasticity are likely to be affected by Ca2+-signal duration and various pre- and postsynaptic factors, perhaps including prior neuronal activity (Abraham and Tate 1997; Bienenstock et al. 1982). During our iontophoretic protocol, however, prolonged postsynaptic Ca2+ transients are uncoupled from presynaptic activity. With this protocol, it may be that a steady-state balance of Ca2+-dependent enzymes plays a relatively large role in determining the ensuing synaptic plasticity. Thus our estimates of the threshold Ca2+ level for LTD (~180 nM) and for the transition from LTD to LTP (~540 nM) in this simplified context may contribute to the ongoing effort to construct a molecular model of Ca2+-dependent synaptic plasticity (Grzywacz and Burgi 1998; Lisman 1989; Malenka and Nicoll 1999; Soderling and Derkach 2000).


    FOOTNOTES

Present address and address for reprint requests: R. J. Cormier, Dept. of Psychiatry, Washington University School of Medicine, St. Louis, MO 63110 (E-mail: cormierb{at}psychiatry.wustl.edu).

1 In one cell, both pathways were unusually responsive to distal glutamate application. However, no other basis existed for excluding these data and their relation to corresponding Ca2+ levels provided nonessential support for this study's conclusions.

Received 1 March 2000; accepted in final form 21 September 2000.


    REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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0022-3077/01 $5.00 Copyright © 2001 The American Physiological Society