Initiation and Propagation of Regenerative Ca2+-Dependent Potentials in Dendrites of Layer 5 Pyramidal Neurons

J. C. Oakley, P. C. Schwindt, and W. E. Crill

Department of Physiology and Biophysics, University of Washington School of Medicine, Seattle, Washington 98195-7290


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Oakley, J. C., P. C. Schwindt, and W. E. Crill. Initiation and Propagation of Regenerative Ca2+-Dependent Potentials in Dendrites of Layer 5 Pyramidal Neurons. J. Neurophysiol. 86: 503-513, 2001. The initiation and propagation of dendritic Ca2+-dependent regenerative potentials (CDRPs) were investigated by imaging the Ca2+-sensitive dye Fluo-4 during whole cell recording from the soma of layer 5 pyramidal neurons visualized in a slice preparation of rat neocortex by the use of infrared-differential interference contrast microscopy. CDRPs were evoked by focal iontophoresis of glutamate at visually identified sites 178-648 µm from the soma on the apical dendrite and at sites on the basal dendrites. Increases in [Ca2+]i were maximal near the site of iontophoresis and were graded with iontophoretic current that was subthreshold for evoking CDRPs. CDRP initiation was associated with a [Ca2+]i rise that differed from a just-subthreshold response in both magnitude and spatial extent but whose amplitude declined both proximal and distal to the iontophoretic site. These [Ca2+]i rises, whether associated with subthreshold or regenerative voltage responses, were minimally affected by blockade of N-methyl-D-aspartate receptors but were abolished by Cd2+, suggesting that Ca2+ influx through voltage-gated channels caused the rise of [Ca2+]i. On the assumption that the rise of [Ca2+]i during a CDRP marks the spatial extent of regenerative Ca2+ influx, we conclude that CDRPs can be evoked at any point on the main apical or basal trunk where membrane potential reaches CDRP threshold rather than at discrete "hot spots," the CDRP is initiated at a spatially restricted site, and it propagates decrementally both distal and proximal to its initiation site. These results raise the possibility that synaptic integration may occur first in the dendrites to evoke a CDRP. Because these responses propagate decrementally to the soma, they are able to sum with input from other regions of the cell so that the cell as a whole remains integrative.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In the classic view of synaptic integration, synaptic currents generated in the dendrites are summed at a region downstream from the soma to evoke the spike output of the cell. In this model, the output of the cell is determined by the sum of each synaptic input weighted by its distance from the downstream integration point. More recently it has been shown that depolarization of the apical dendrite of layer 5 neocortical neurons by focal iontophoresis of glutamate (Schwindt and Crill 1997, 1999), by current injection through a dendritic recording electrode (Kim and Connors 1993; Larkum et al. 1999; Schiller et al. 1997), or by synaptic stimulation (Kim and Connors 1993; Larkum et al. 1999; Schiller et al. 1997; Schwindt and Crill 1998) can evoke Ca2+-dependent action potentials. These Ca2+-dependent regenerative potentials (CDRPs) have been proposed as a mechanism both to amplify (Schiller et al. 1997; Schwindt and Crill 1997) and to limit (Oakley et al. 2001) the distal synaptic input that reaches the soma. If CDRPs are evoked locally by dendritic depolarization, there are at least two (and potentially many) sites of synaptic integration, one near the soma and the other(s) in the dendrites. According to this idea, synaptic integration would be determined in part by the number and the spatial extent of the site(s) of initiation of the regenerative dendritic potentials. If the dendritic regenerative responses remain localized, the downstream summing site may still integrate the whole cell current to evoke the cell's Na+ spike output, and the cell as a whole would remain integrative.

There is both direct (Larkum et al. 1999; Schiller et al. 1997) and indirect (Schwindt and Crill 1997) electrophysiological evidence, as well as evidence from Ca2+ imaging (Markram et al. 1995; Schiller et al. 1997), that the CDRPs are not propagated actively to the soma, but several questions remain. Are all portions of the dendrite capable of generating these regenerative potentials or only a few regions or only one region? Experiments employing Ca2+ imaging techniques (Markram and Sakmann 1994; Markram et al. 1995; Schiller et al. 1995; Yuste et al. 1994) suggest that Ca2+ channels exist over most or all of the apical dendrite, but other studies have suggested that CDRPs are evoked only at a few specific points ("hot spots") on the apical dendritic (Reuveni et al. 1993) or only in the distal apical tuft (Schiller et al. 1997). All available evidence is consistent with the decremental propagation of the CDRPs toward the soma, but the spatial extent of the spike-generating region(s) is not clear. It is not known, for example, if CDRPs are actively backpropagated. Because they are caused by Ca2+ influx, the backpropagation of these potentials might alter the efficacy of active distal synapses, which are known to be modulated by a rise of [Ca2+]i (Neveu and Zucker 1996; Yang et al. 1999). In the present study, we sought to answer these questions by the use of glutamate iontophoresis to depolarize sites on visualized dendrites of layer 5 neocortical neurons and the imaging of the Ca2+-sensitive dye, Fluo-4, which was loaded into the cell through the soma patch electrode. Since the CDRPs are known to depend on Ca2+ influx through voltage-gated Ca2+ channels, we measured changes in Fluo-4 fluorescence to determine the spatial-temporal increase in dendritic Ca2+ concentration ([Ca2+])i during CDRPs. We deduced the mode of initiation and propagation of CDRPs from the changes in [Ca2+].

The use of iontrophoresed glutamate to depolarize a visualized dendritic site is convenient for the several reasons discussed in Oakley et al. (2001). Foremost, it allows the amplitude and duration of long-lasting dendritic depolarization to be controlled conveniently by the experimenter at a precisely known distance from the soma. Importantly for the present study, it allows the response to the same depolarizing stimulus to be compared in the absence and in the presence of channel-blocking agents that would alter or block synaptic transmission.

In the study by Schwindt and Crill (1999), short iontophoretic pulses of glutamate evoked transient Ca2+ spikes that lasted ~100 ms, while longer iontophoretic pulses evoked a long-duration Ca2 spike (a "plateau") that persisted for the duration of the iontophoretic current. In this study, we focused on the rise in [Ca2+]i during the regenerative plateau response in part because these long-lasting plateaus have been less well studied than the transient Ca2+ spikes. A great experimental advantage of the plateau is that it lasts as long as the iontophoresis, which is under the experimenter's control. A long-lasting response allows the use of a sufficiently long CCD camera integration time to obtain a fluorescence signal with a satisfactory signal-to-noise ratio while also allowing a sufficient number of images to be obtained to define adequately the time course of the [Ca2+]i transient.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Tissue preparation and solutions

Sprague-Dawley rats of either sex aged 20-32 days postnatal were anesthetized with ketamine (150 mg/kg) and xylazine (10 mg/kg) and killed by carotid section. Following craniotomy, a section of dorsal frontoparietal (sensorimotor cortex) was removed, fixed with cyanoacrylite glue to the stage of a microslicer, and submerged in ice-cold physiological saline solution (PSS) containing (in mM) 130 NaCl, 3.0 KCl, 2.0 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, 2 CaCl2, and 20 glucose gassed with 95% O2-5% CO2 (carbogen). Coronal slices (300 µm) were cut and stored in a holding chamber filled with carbogenated PSS at 34°C. Individual slices were transferred to a recording chamber where they were maintained submerged in 29-32°C carbogenated PSS flowing at >2 ml/min. In some experiments TTX (1 µM, Sigma Chemical, St. Louis, MO) or D(-)-2-amino-5-phosphonopentanoic acid (AP-5) (100 µM, Precision Neurochemicals, Vancouver BC, Canada) were added to the PSS, or 200 µM CdCl2 was substituted for an equimolar amount of CaCl2 and NaH2PO4 was omitted to avoid precipitation.

The bottom of the 20-mm diam, 3-mm-deep recording chamber was formed by a No. 1 glass coverslip. The slice was held in place on the coverslip by thin glass bars straddling the two arms of a U-shaped platinum wire 300 µm thick. The gravity-fed inflow and suction outflow were positioned to promote laminar flow across the slice. The recording electrode and the iontophoretic electrodes opposed each other so that they could be placed independently anywhere in the chamber.

Recording methods

An upright microscope (Olympus BX50WI) fit with ×40 LWD water-immersion objectives and DIC optics was used to view cells in the top 100 µm of the slice. The slice was transilluminated using infrared (Omega 770 ± 40 nm band-pass filter) and viewed using a CCD camera (Hamamatsu) and a high-resolution video monitor (Sony). The microscope was fixed to an x-y translation stage that allowed the microscope to be positioned independently of the recording and iontophoretic electrodes so that different portions of the cell could be viewed without disturbing these pipettes.

Micropipettes pulled from VWR 75 ml capillary glass and filled with standard intracellular solution containing (in mM) 135 KMeSO4, 5 KCl, 2 MgCl2, 10 (3-[N-morpholino]propanesulfonic acid) (MOPS), 0.1 ethylene glycol-bis(beta -aminoethyl ether)N,N,N',N'-tetraacetic acid (EGTA), 2 Na-ATP, 0.2 Na-GTP, 0.15 Fluo-4 (Molecular Probes) and (in %wt/vol) 0.01 Lucifer yellow, K+ salt (Molecular Probes) were used to make whole cell recordings.

Extracellular DC resistance of the pipettes was 2-4 MOmega . The seal resistance formed with the soma membrane was >1 GOmega before break in to the whole cell configuration. Series resistance in the whole cell configuration was monitored by maintaining bridge balance during a hyperpolarizing current pulse. Recordings were discarded if series resistance became >50 MOmega .

An Axoclamp-2A amplifier (Axon Instruments, Foster City, CA) was used in bridge mode to record somatic membrane potential and inject current through the recording pipette. Stable recordings lasting 1-2 h were frequently obtained. Resting potential was taken as the difference between intracellular and extracellular potentials recorded on a strip chart recorder. Recorded potentials were corrected for a tip potential of 10 mV. Current and voltage recordings were filtered at 10 kHz and stored on videotape using pulse code modulation (Neurodata) and were later digitized (Axon Instruments TL-1 125 interface) and analyzed with the program WCP written and distributed by John Dempster and with the program IGOR (WaveMetrics, Lake Oswego, OR).

Glutamate iontophoresis

Fluorescence microscopy was used to simultaneously view the dendritic arbor and the iontophoretic electrode so that glutamate could be applied to visually identified segments of dendrite at a known distance from the soma. After obtaining a stable whole cell recording, short, hyperpolarizing current pulses (<500 pA) were applied to the recording pipette to iontophorese K+-Lucifer yellow and Fluo-4 dyes into the cell. The DIC analyzer was removed, and the light path was switched from transmitted light to epifluorescence illumination with excitation at 400 nm and emission at 535 nm. Illumination at 400 nm was chosen to excite K+-Lucifer yellow dye without bleaching Fluo-4 dye (peak excitation approx 485 nm).

Iontophoretic electrodes were pulled from 1 mm OD glass capillary tubes. Their tips were broken to 2 µm diam, and they were filled with (in mM) 150 glutamatic acid, 2.24 CaCl2, 4 KCl, 30 N-(2 hydroxyethyl)piperizine-N'-(2-ethanesulfonic acid) (HEPES), and (as % wt/vol) 0.05% K+-Lucifer yellow salt, pH 7.4. The second current amplifier and headstage of the Axoclamp-2A amplifier was used to provide positive DC holding current (+5 nA) and negative iontophoretic pulses (1-2 s duration, -5- to -100-nA amplitude). Once the dendrites contained sufficient dye, the iontophoretic electrode was visually guided to a site on the dendritic tree. Movement of the electrode tip over a range of 10-20 µm near the visualized dendrite caused the iontophoretically evoked response to vary from zero to maximal. The glutamate concentration 10 µm from the electrode tip caused by an iontophoretic current sufficient to evoke a plateau potential was estimated to be <= 10 mM (Oakley et al. 2001).

Imaging methods

Once the iontophoretic electrode was placed in the dendritic arbor and a stable iontophoretic response was obtained, the excitation wavelength was switched to 508 nm which was found to excite the Fluo-4 dye but not the K+-Lucifer yellow dye in the cell or in the iontophoretic electrode. Fluorescence signals were recorded at ×40 magnification. Imaging commenced >10 min after break-in to allow Fluo-4 to diffuse from the patch solution into the dendrites. Excitation wavelength was controlled by a TILL system (Photonics, Planeg, Germany). When not imaging, the excitation light was stepped to a long wavelength, and the scattered light from the TILL system was blocked by a high-pass filter. In most experiments, the iontophoretic electrode was centered within the imaged region. In some experiments, the imaged region proximal or distal to the iontophoretic electrode was increased by placing the iontophoretic electrode near one end of the imaged region. In those cells the spatial extent of the fluorescence signal was only measured in one direction from the iontophoretic electrode (the unmeasured direction is indicated by n.m. in Fig. 4).

Prior to the iontophoretic pulse, 10 images of resting fluorescence (FR) were acquired using a Pentamax cooled CCD camera (Princeton Instruments, EEV 1,024 × 512 chip) in frame transfer mode and Metafluor software (Universal Imaging). The first image contained no data and was thrown out. An additional 40 images were acquired (50 ms each) during and following the iontophoretic pulse. These images, taken at ×40, were acquired using a subchip defined around the dendrite with pixels binned by 3 to decrease the acquisition time required to give a satisfactory signal-to-noise ratio. Following each experiment images also were taken at ×10 using K+-Lucifer yellow excitation (425 nm) to define the straight-line distance from the center of the soma to the iontophoretic site during off-line analysis.

The ×40 image was used to define regions of interest (ROIs) that consisted of serial 8.5-µm-long sections of dendrite and that were analyzed off-line. The average fluorescence of each ROI was calculated with the use of the software package Metafluor. A shading image was acquired in a weak dye solution (which provided a bright, uniform fluorescence) for each subchip and binning combination employed. The raw fluorescence signals were corrected for shading by dividing each ROI by the value of the fluorescence at the same ROI in the shading image. The fluorescence signals were corrected for bleaching by fitting a line to the fluorescence of the nine frames taken at rest prior to the stimulus. The fitted line was then extrapolated and subtracted from the shading corrected fluorescence signals. After the application of these shading and bleaching corrections, the stimulus-linked change in fluorescence (Delta F) in each ROI were computed as Delta F = FS - FR, where FS is the average fluorescence of the ROI following stimulation and FR is the prestimulation resting fluorescence of the ROI. Data are presented as the relative change in fluorescence, Delta F/F = (FS - FR)/(FR - FB) where FB is the background fluorescence which was determined from an ROI away from the fluorescent dendrite.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell properties

Recordings were made from 27 neocortical layer 5 pyramidal neurons in slices from 20 rats 20-32 days old. Resting potentials ranged from -54 to -79 mV (mean: -72 mV). Input resistance, measured with long hyperpolarizing current pulses, varied from 24 to 100 MOmega (mean: 45 MOmega ). Cells were accepted for analysis if they had stable resting potentials and fired repetitive action potentials to depolarizing soma current. All recorded cells exhibited regular spiking (Connors and Gutnick 1990) in response to depolarization of the soma by injected current. No intrinsic bursters or fast spiking cells (Connors and Gutnick 1990) were recorded.

Plateau properties

In a previous study Schwindt and Crill (1999) found that a long-duration, all-or-none, Ca2+-dependent action potential (a "plateau"), which was usually preceded by an initial, transient, Ca2+ spike, could be evoked by focal iontophoresis of glutamate on the apical dendrite of layer 5 pyramidal cells. Similar responses were evoked by 1- to 2-s duration iontophoresis of glutamate in this study, which employed whole cell patch recording methods and direct visualization of the soma and dendritic tree (see METHODS). The properties of these responses, as recorded using whole cell, IR-CCD methods, are described in detail in Oakley et al. (2001). Figure 1 illustrates some key features of the Ca2+ spike and plateau. In this cell iontophoresis (-60 nA) at a site 460 µm from the soma on the apical dendrite evoked an initial, low-threshold, slow, action potential on which was superimposed a higher-threshold, fast spike (Fig. 1, black trace). The different time courses and thresholds of the fast and slow spikes are not obvious at the slow sweep speed used in this figure (cf. Schwindt and Crill 1997). Similar fast spikes were eliminated by TTX application, and similar slow spikes were eliminated by Cd2+ application (see following text). The membrane potential response during the remainder of this iontophoresis remained subthreshold for action potential initiation. A slightly larger iontophoretic current (-70 nA) again evoked the initial Ca2+ and Na+ spikes, which were then followed by a long-duration action potential (the plateau) that repolarized only when the iontophoretic current was terminated (Fig. 1, green trace). In all cells tested, a larger iontophoretic current (-80 nA in Fig. 1) decreased the latency to plateau initiation but did not increase plateau amplitude (Fig. 1, red trace). Similar plateaus were evoked by focal iontophoresis of glutamate at 22 identified sites on the apical dendrite in different experiments.



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Fig. 1. Plateau potential evoked all-or-none by focal iontophoresis of glutamate. Top: membrane potential responses color coded to iontophoretic currents (bottom) measured at the soma in response to focal glutamate iontophoresis 460 µm from the soma on the apical dendrite. Iontophoresis (-60 nA, 1-s duration) evoked an early Ca2+ spike which itself evoked a Na+ spike followed by a subthreshold response for the duration of the iontophoresis (black trace). Slightly larger iontophoretic current (-70 nA, green trace) evoked early Ca2+ and Na+ spikes followed by a plateau. Additional iontophoretic current (-80 nA, red trace) decreased the latency to plateau initiation but did not increase the amplitude of the plateau. Resting potential was -67 mV. Cell was held at -77 mV by DC current injection at soma to prevent Na+ action potentials during the plateau.

Location and properties of evoked [Ca2+]i increase in the apical dendrite

Since the plateau depends on Ca2+ influx through voltage-gated channels (Oakley et al. 2001; Schwindt and Crill 1999), its site of initiation and its propagation along the dendrite was expected to be indicated by the temporally correlated rise of intracellular Ca2+ concentration ([Ca2+]i). The rise of [Ca2+]i was estimated by measuring the change in relative fluorescence (Delta F/F, see METHODS) of the Ca2+-sensitive dye Fluo-4 during glutamate iontophoresis. An increase of Delta F/F signals an increase in [Ca2+]i. Fluorescence responses (Delta F/F) were measured and compared during iontophoreses that were subthreshold and suprathreshold for plateau initiation, both in physiological saline and in the presence of various blocking agents.

Typical results obtained in physiological saline are illustrated in Fig. 2, which is from the same cell as shown in Fig. 1. As indicated in the schematic drawing of Fig. 2A, the field of view was centered near the visually identified position of the iontophoretic electrode. In the cell of Fig. 2, this visualized region of the apical dendrite extended from 290 to 540 µm from the soma (indicated by red box in Fig. 2A). The visually identified location of the iontophoretic electrode is indicated by the dashed arrow in Fig. 2A. Images were taken every 50 ms, starting 500 ms before the iontophoretic pulse (to establish the resting fluorescence level) and ending 1 s after the iontophoretic pulse was terminated. The entire visualized length of the apical dendrite was divided into contiguous 8.5-µm-long segments in which average Delta F/F values were measured (see METHODS). Delta F/F values are represented according to the pseudocolor scale at the bottom left of Fig. 2B. In the pseudocolor plot of Fig. 2B, and in all other such plots in this paper, Delta F/F is plotted as a function of both distance along the apical dendrite (plot ordinate) and time after the start of the sweep (plot abscissa) to show the spatial-temporal increase in Delta F/F during an iontophoresis. Note that the ordinate in this (and all other such plots) starts at the most proximal part of the imaged region rather than at the soma. The membrane potential response (recorded at the soma) during the iontophoresis is shown at the same time scale below the pseudocolor plot (Fig. 2B, red trace). This is the same response to the -60 nA iontophoresis shown in Fig. 1. From the pseudocolor plot, it can be seen that the largest rise in [Ca2+]i occurred at the iontophoretic site (indicated by dashed line on pseudocolor plot). During the initial Ca2+ and Na+ spikes, [Ca2+]i increased above resting levels at locations both proximal and distal to the iontophoretic site. Following these initial spikes, [Ca2+]i decayed to the resting level at these proximal and distal locations, whereas [Ca2+]i remained above the resting level near the iontophoretic site for the remainder of the iontophoresis. After the iontophoresis was terminated, [Ca2+]i decayed to baseline. In this and all cells examined, an iontophoresis that was subthreshold for either the plateau or the initial transient Ca2+ spike caused an increase of [Ca2+]i, and the largest rise in [Ca2+]i occurred near the iontophoretic site.



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Fig. 2. Spatial-temporal pattern of rise in intracellular Ca2+ ([Ca2+]i) evoked by iontophoresis on apical dendrite. A: schematic drawing of same layer 5 pyramidal cell as shown in Fig. 1 showing extent of imaged area on apical dendrite (red box), distance from soma, and position of iontophoretic electrode (at arrow) during recording of membrane potential at soma (indicated by Vm) in this experiment. B, top: pseudocolor plot of the relative change in fluorescence (Delta F/F), an indicator of [Ca2+]i (see METHODS), in the imaged region as a function of distance along the apical dendrite 290-540 µm from the soma (ordinate) and time from the start of the sweep (abscissa). Shown below this pseudocolor plot are the corresponding membrane potential traces recorded at the soma (red trace) evoked by glutamate iontophoresis (black trace below) at the same time scale. Same record as -60 nA iontophoresis in Fig. 1 (with Na+ spike truncated.). C: similar to B but for same record as -70 nA iontophoresis in Fig. 1. In this and following figures the black dashed lines in the pseudocolor plots indicate the position of the iontophoretic electrode. The black contour line (in the plot of C) is drawn around a region with Delta F/F >=  50% of the peak Delta F/F evoked during the plateau (see RESULTS). Pseudocolor Delta F/F scale and 10-mV calibration bar in B also applies to C.

In Fig. 2C, the plateau was evoked (red trace at bottom) during a larger (-70 nA) iontophoresis at the same site. As indicated by the pseudocolor plot (Fig. 2C, top), during and immediately after the initial Ca2+ and Na+ spike the spatial-temporal increase of [Ca2+]i was similar to that of Fig. 2B. Subsequently, when the plateau was evoked, the increase of [Ca2+]i at the iontophoretic site was larger than during the subthreshold response of Fig. 2B or the subthreshold portion of the response that preceded the plateau in Fig. 2C. In addition, [Ca2+]i rose above its rest value both proximal and distal to the iontophoretic site following plateau initiation, and this rise persisted for the duration of the plateau. Thus the plateau was associated with a [Ca2+]i rise that differed from a subthreshold response in both magnitude and spatial extent.

Our primary finding is that the rise of [Ca2+]i associated with plateau generation declined with distance from the iontophoretic spike, both proximally and distally. This same pattern, highest around the iontophoretic site and lower at the proximal and distal boundaries of the imaged region, was evoked during plateau generation in all recorded cells. Assuming for the moment that the rise of [Ca2+]i is caused entirely by Ca2+ influx through voltage-gated channels, these results imply that dendritic membrane potential (i.e., plateau amplitude) also declined with distance. That is, the plateau propagates decrementally both proximal and distal to its site of initiation. In contrast, a regenerative response that propagated actively with constant amplitude, like a Na+ spike propagating down the axon, would be expected to result in a similar rise of [Ca2+]i along the whole region of active propagation. The assumptions on which this conclusion is based were tested in experiments described in the following text.

Because [Ca2+]i also rose during a subthreshold response and because [Ca2+]i did not decline abruptly with distance from the iontophoretic site during either the subthreshold response or the plateau, the precise spatial extent of active plateau generation (i.e., the region where net inward ionic current caused a regenerative response) was difficult to determine. Since subthreshold responses evoked a rise of [Ca2+]i, the amplitude of even a decrementally propagating plateau would be great enough to activate Ca2+ channels to cause a nonregenerative Ca2+ influx and a rise of [Ca2+]i. To provide some quantitative estimate of the spatial extent of the active region (in this and all recorded cells), a contour line was drawn around the region in which Delta F/F was >= 50% of the peak Delta F/F reached during the plateau (Fig. 2C, black line). This 50%-of-peak contour line was used to estimate the region of active plateau initiation.

Our rationale for adopting the 50%-of-peak contour line as an index of the spatial extent of the active plateau generating region is illustrated by the plots of Fig. 3. The plot of Fig. 3A is a more conventional representation of the florescence data of Fig. 2C. Florescence at three times during the response (at rest, 250 ms; during the subthreshold response after the initial Na+ and Ca2+ spikes, 1,000 ms, and during the plateau, 1,500 ms) is plotted versus distance from the soma over the field of view. Although we interpret the fluorescence data as resulting from variations of dendritic membrane potential, it should be realized that the relation between fluorescence signal (Delta F/F) and dendritic membrane potential is quite indirect and complex. Because of the sublinear nature of the binding relation between Ca2+ and this high-affinity dye, a small Delta F obtained when ambient [Ca2+]i is high may represent a larger increment of [Ca2+]i than does a larger Delta F obtained when ambient [Ca2+]i is low, and of course there is a highly nonlinear relation between membrane potential and Ca2+ current. Thus no simple or quantitative conclusions can be drawn from the plots of Fig. 3A about the variation of [Ca2+]i or dendritic membrane potential beyond the fact that both quantities must be larger where the fluorescence signal is larger (again assuming the signal arises solely from Ca2+ influx through voltage-gated channels). During the plateau in Fig. 3A, the fluorescence signal rose significantly above the subthreshold response over most of the 250 µm field of view, but the rise was particularly abrupt over the region bracketed by the vertical dotted lines, suggesting that Ca2+ influx was particularly large and relatively uniform over this region. We assume therefore that this region constitutes the extent of active plateau generation, and the vertical dotted lines represent the 50%-of-peak contour lines of Fig. 2C at this point in time.



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Fig. 3. Basis for estimating the region of regenerative plateau initiation. A: plot of relative fluorescence (Delta F/F) vs. distance from the soma at the 3 times indicated during the single sweep of Fig. 2C. Time of 250 ms is before the iontophoresis; 1,000 ms is during the subthreshold response after the initial Na+ and Ca2+ spikes, and 1,500 ms is during the plateau (cf. Fig. 2C). Vertical dotted lines mark the values of the 50%-of-peak black contour line of Fig. 2C obtained at 1,500 ms. Upward arrow marks site of iontophoresis. B: relation between iontophoretic current and the average fluorescence at the iontophoretic site in another cell. Each plotted Delta F/F value is the average of 4 sequential images taken during an iotophoresis. The imaged region of interest (ROI) extended 25.5 µm along the apical dendrite centered on the iontophoretic site 280 µm from the soma. The period of image acquisition, which was identical for each iontophoretic current, corresponded to the 1st 200 ms of the plateau, which was first evoked at -95 nA. Three smaller iontophoretic currents evoked only subthreshold responses whose data points are fitted with a solid line. The horizontal dashed line is drawn at 50% of the peak Delta F/F, which was used to estimate the region of regenerative plateau initiation. See text for further explanation.

The plot of Fig. 3B is from a different cell in which several subthreshold responses were obtained from iontophoresis 280 µm from the soma before evoking a plateau. The rightmost data point (labeled "plateau") is the average Delta F/F value measured over the first 200 ms (the 1st 4 images) of the plateau at the iontophoretic site. The data points to the left were obtained from the same region and over the same time interval but during smaller iontophoretic currents that evoked only the subthreshold responses. The subthreshold signals were graded with iontophoretic current as indicated by the line fitted to the three subthreshold data points. The average Delta F/F value measured during the plateau lies significantly above this line, suggesting there was a "jump" in [Ca2+]i after the plateau was triggered. Such a jump is expected because of the regenerative Ca2+ influx associated with the initiation of the plateau. The horizontal dashed line in Fig. 3 is drawn at 50% of the peak Delta F/F value. We assume Delta F/F values that lie above this arise from active (regenerative) plateau initiation, whereas values below the are ascribed to nonregenerative Ca2+ influx.

Based on the observations and rationale described in the preceding text, the proximal and distal extent of the active plateau-generating region was defined as the most proximal and distal extent of the 50%-of-peak contour line during a plateau. Using this criterion, the active plateau-generating region was remarkably small but of similar magnitude among the cells tested. Active plateau initiation extended 50 µm proximally and 38 µm distally from the iontophoretic site in the experiment of Fig. 2C. Plateaus were evoked on the apical dendrite of 12 cells bathed in physiological saline. Data from these 12 plateaus are plotted as squares in Fig. 4. The squares mark both the distance of the iontophoretic site from the soma in each cell and the distance where Delta F/F during the plateau was maximal, since both distances were identical. An estimate of the spatial extent of active plateau initiation in each cell (measured using the 50%-of-peak contour described in the preceding text) is indicated by the length of the vertical bars through each data point. In these experiments, the proximal extent of active initiation ranged from 20 to 113 µm (mean: 63.2 µm), and the distal extent ranged from 10 to 207 µm (mean: 51.4 µm). The proximal and distal extents were not significantly different (P = 0.47, paired, 2-tailed, Student's t-test).



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Fig. 4. The spatial extent of the regenerative plateau generating region was similar in all regions of the apical dendrites. Data are from all recorded cells. Plateaus were evoked by glutamate iontophoresis at 24 sites whose locations from the soma are given by the ordinate. Squares, sites on apical dendrites tested in physiological saline (12 cells). Triangles, sites on apical dendrite tested in 1 µM TTX plus 100 µM AP-5 (10 cells), and circles, sites on basal dendrites in physiological saline (2 cells). The estimated proximal and distal extent of regenerative plateaus generating regions are plotted as gray bars for each site. The proximal or the distal extent was not measured in some cells (labeled n.m.; see METHODS).

The plot of Fig. 4 also shows that plateaus were evoked over the whole extent of the apical dendrite examined (from 178 to 648 µm from the soma), and all plateaus were centered spatially about the iontophoretic site. The ability to evoke plateaus at sites over this length of the apical dendrite indicates that Ca2+ channel density is adequate for plateau generation over (at least) this length of the apical dendrite.

Increased [Ca2+]i results primarily from voltage-gated Ca2+ channel activation

It was assumed in the preceding text that the rise in [Ca2+]i is caused by influx of Ca2+ through voltage-gated Ca2+ channels. However, Ca2+ might also enter through N-methyl-D-aspartate (NMDA)-sensitive glutamate channels. The increase of [Ca2+]i observed during dendritic depolarizations that were subthreshold for plateau initiation (e.g., Figs. 2, A and B, and 3) particularly raised the possibility of Ca2+ influx through NMDA-sensitive channels.

To investigate the source of the Ca2+ responsible for the rise in [Ca2+]i, the spatial extent of plateaus evoked at the same site was investigated before and after the addition of 100 µM of the specific NMDA receptor antagonist AP-5 to physiological saline in two cells. In one cell, the extent of maximal [Ca2+]i rise (measured using the 50% contour criterion outlined in the preceding text) evoked by glutamate iontophoresis at a site 251 µm from the soma was not changed by the addition of AP-5. In the second cell, the proximal and distal extent of a plateau evoked by glutamate iontophoresis at a site 271 µm from the soma was decreased approx 10% by the addition of AP-5. In neither of these cells did the addition of AP-5 decrease the rise of [Ca2+]i evoked by a just-subthreshold iontophoretic current (data not shown).

To confirm that Ca2+ influx through NMDA channels was not a major influence on the measured rise in [Ca2+]i and to additionally test whether voltage-gated Na+ channel activity might influence the initiation site or spatial extent of the plateau, plateaus were evoked in an additional 10 cells bathed in 100 µM AP-5 plus 1 µM TTX. Figure 5 illustrates typical results when both Na+ channels and NMDA receptors were blocked. Plateaus were evoked in each cell tested under these conditions, and the spatial-temporal pattern of the rise of [Ca2+]i and the spatial extent of the plateaus were similar to those obtained in physiological saline. In these experiments, the estimated proximal extent of the plateaus ranged from 15 to 88 µm (mean: 53.1 µm) and the distal extent ranged from 22 to 146 µm (mean: 49.5 µm). These values were not significantly different from those obtained in physiological saline (for proximal extent P = 0.46; for distal extent P = 0.94; 2-tailed Student's t-test). As found for physiological saline, the proximal extent of plateaus evoked in AP-5 plus TTX was not significantly different from the distal extent (P = 0.81, 2-tailed Student's t-test). Data obtained from plateaus evoked in AP-5 plus TTX are plotted as triangles in Fig. 4 for comparison with plateaus evoked in physiological saline.



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Fig. 5. Increase in [Ca2+]i is caused by Ca2+ influx through voltage-gated Ca2+ channels. A: spatial-temporal rise of Delta F/F in the imaged region of the apical dendrite 225-450 µm from the soma during glutamate iontophoresis in a cell bathed in 1 µM TTX plus 100 µM AP-5. The corresponding depolarization (red trace) evoked by the iontophoresis (black trace) at 344 µm from the soma is shown at the same time scale for comparison. Iontophoresis evoked an early small Ca2+ spike followed by a plateau. B: after the addition of 200 µM Cd2+, the same iontophoresis evoked a passive response with no increase in Delta F/F. Pseudo-color scale, and 20 mV scale bar in A also applies to B. Resting potential was -74 mV.

To verify directly that increases in dendritic [Ca2+]i result from Ca2+ influx through voltage-gated Ca2+ channels, iontophoretically evoked changes in dendritic [Ca2+]i were compared before and after the addition of the Ca2+ channel blocker Cd2+ in three cells. Typical results are shown in Fig. 5. The iontophoretic current that evoked the plateau and rise of [Ca2+]i shown in Fig. 5A evoked only a passive membrane potential response and little or no increase in [Ca2+]i in the presence of 200 µM Cd2+ (Fig. 5B). In this and each cell tested, a subthreshold iontophoresis also evoked a significant rise of [Ca2+]i that also was blocked by Cd2+ (data not shown). Altogether, the experiments with AP-5 and particularly with Cd2+ suggest that the measured rise in [Ca2+]i, whether occurring during the plateau or during a subthreshold response, results predominantly from Ca2+ influx through voltage-gated Ca2+ channels. In addition, the results in TTX suggest that neither the site of initiation nor the spatial extent of the plateau depended significantly on Na+ channel activity.

Location and properties of evoked [Ca2+]i increase in basal dendrites

Plateaus could also be evoked by focal glutamate iontophoresis on basal dendrites (Oakley et al. 2001). In the present study, the spatial extent of these plateaus was examined in two cells. Figure 6 shows the results obtained in one of these cells. Focal glutamate iontophoresis at a site 128 µm from the soma on a basal dendrite evoked an all- (red trace, bottom)-or-none (gray trace, bottom) plateau. The corresponding pseudocolor plot (Fig. 6, top) shows that the plateau-evoked rise of [Ca2+]i was restricted to a small region surrounding the iontophoretic site (indicated by dashed line in pseudocolor plot). A 50% contour drawn around the region estimates the proximal extent of the plateau as 16 µm and the distal extent as 15 µm. In this cell, the spatial extent of the plateau was not significantly changed when 100 µM AP-5 was added to the bath and the experiment was repeated (data not shown). In the second cell tested, iontophoresis at a site 105 µm from the soma evoked a plateau, which was estimated to extend 20 µm proximally and 10 µm distally.



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Fig. 6. Plateaus evoked in basal dendrites also are restricted to the iontophoretic site. Glutamate iontophoresis on a basal dendrite 122 µm from the soma evoked a plateau (red trace). A just-subthreshold iontophoresis (gray trace) is also shown to illustrate that the plateau was evoked all-or-none. The pseudocolor plot above corresponds only to the sweep during which the plateau was evoked. The rise in [Ca2+]i temporally correlated with the plateau was localized around the iontophoretic site. Resting potential was -78 mV.


    DISCUSSION
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INTRODUCTION
METHODS
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DISCUSSION
REFERENCES

Focal iontophoresis of glutamate on dendrites of neocortical layer 5 pyramidal neurons evoked all-or-none CDRPs, the transient Ca2+ spike and the plateau, which were associated with a maximal rise of dendritic [Ca2+]i around the iontophoretic site. The rise of [Ca2+]i associated both with the CDRPs and with subthreshold iontophoretic responses was blocked by Cd2+ but was insensitive to AP-5, suggesting that Ca2+ influx through voltage-gated channels caused the rise of [Ca2+]i. Since the peak rise of [Ca2+]i always occurred around the iontophoretic site at all locations tested (from 178 to 648 µm on the apical dendrite; cf. Fig. 4), it is unlikely that the plateaus were initiated at discrete "hot spots" on the dendrite. These results strongly suggest that CDRPs can be initiated at any point on the main apical trunk where the membrane potential reaches plateau threshold.

The spatial-temporal pattern of [Ca2+]i rise was used to assess the site of active CDRP initiation, which appeared to extend only over a distance of some tens of micrometers. The decline of [Ca2+]i with distance from the ionotophoretic site strongly suggests that CDRP amplitude also declined with distance (thereby activating fewer Ca2+ channels to result in less Ca2+ influx), and the CDRP was thus propagated decrementally both distal and proximal to its site of initiation.

Most of our data were obtained from iontophoresis on the apical dendrite. In a previous study (Oakley et al. 2001) we found that plateaus with electrical properties similar to those evoked on the apical dendrite could be evoked on basal and apical oblique dendrites. In the present study, we measured the plateau-associated rise of [Ca2+]i in the basal dendrites of only two cells simply to examine whether its spatial-temporal properties were similar to those observed in the apical dendrite. The increase in [Ca2+]i in the basal dendrites did not extend as far proximally or distally as on the apical dendrite. This is probably explained by a greater electrotonic decay of current in basal dendrites than in apical dendrites. In support of this explanation, we found that the amplitude of basal dendrite plateaus (measured at the soma) was less than expected for a plateau evoked on the apical dendrite at a comparable distance from the soma (Oakley et al. 2001). Assuming equal plateau amplitudes at the dendritic site of initiation in both basal and apical dendrites, a greater decay of the potential with distance in basal dendrites would result in a smaller plateau amplitude at the soma. The greater decay of potential with distance in a basal dendrite would also result in less Ca2+ channel activation away from the initiation site and thus a more spatially restricted [Ca2+]i transient. A recent study that used a different method to apply exogenous glutamate to the dendritic membrane also found that CDRPs and an associated rise of [Ca2+]i could be evoked in basal dendrites (Schiller et al. 2000). The spatial extent of the [Ca2+]i rise measured in that study was very similar to our values. However, both the rise of [Ca2+]i and the dendritic spikes were attributed to current through NMDA receptor channels rather than voltage-gated Ca2+ channels in that study. We examined this question in only one of the two basal dendrites that we examined in this study, and we found both the plateau and the rise of [Ca2+]i to be AP-5 resistant.

In the present study, most tests of whether Ca2+ influx through NMDA-sensitive glutamate channels contributed to the rise of [Ca2+]i were performed on the apical dendrite. We used 100 µM AP-5 to block the NMDA-sensitive glutamate receptors. This concentration of AP-5 was chosen because its bath application was found to block the action of NMDA itself in layer 5 neurons using iontophoretic currents similar to those employed here (Flatman et al. 1986). There are two basic questions concerning Ca2+ influx through NMDA-sensitive glutamate channels. One question is whether NMDA currents are primarily responsible for the plateau itself. This possibility is raised by the fact that the cation current through NMDA channels can be voltage dependent and cause regenerative depolarizations in the presence of Mg2+ (Flatman et al. 1986). As discussed in Oakley et al. (2001), we found that Cd2+application blocked the plateau in every cell tested, and we concluded that Ca2+ currents through voltage-gated channels were essential for plateau initiation in most cases. Nevertheless, we also found that TTX or AP-5 application also blocked or altered the plateau in a minority of the cells in which these agents were tested, indicating that inward ionic currents other than Ca2+ can contribute to the plateau and that the contribution of these other currents can be essential for the regenerative response in some cases.

In the context of the present experiments, the important question is whether the rise of [Ca2+]i that we measured is caused by Ca2+ influx through NMDA channels. The findings of Oakley et al. (2001), that the addition of 100 µM AP-5 did abolish or alter plateaus in some cells, suggest that 100 µm AP-5 is sufficient to prevent iontophoresed glutamate from occupying NMDA receptors at those sites where the receptors play a significant role in the observed response. No blockade of the rise in [Ca2+]i by AP-5 was observed in the present study, but we did not examine the effect of AP-5 on every recorded cell. We examined the rise of [Ca2+]i before and after the addition of AP-5 in two cells and in the presence of AP-5 plus TTX in 10 others. The measured rise of [Ca2+]i and its spatial-temporal pattern in these cells was not significantly different from those cells not tested with AP-5. Furthermore, Cd2+ completely blocked the rise of [Ca2+]i (Fig. 5B). Thus we feel confident that the hypothesis that Ca2+ influx through NMDA channels significantly influenced our results was adequately tested and rejected. Similar results indicating a minimal role for Ca2+ influx through NMDA channels and a dominant role for Ca2+ influx through voltage-gated channels in the rise of [Ca2+]i that accompanied subthreshold, electrically evoked, dendritic excitatory postsynaptic potentials (EPSPs) were obtained in both hippocampal (Magee et al. 1995) and layer 5 pyramidal neurons (Markram and Sakmann 1994).

Recent experiments in hippocampal CA1 pyramidal neurons (Nakamura et al. 2000) found that the stimulation of metabotropic glutamate receptors by EPSPs coincident with influx of Ca2+ through voltage-gated channels (during action potentials) caused an IP3-dependent release of Ca2+ from internal stores in the apical dendrites of those neurons. The spatial extent of the rise of [Ca2+]i in a CA1 neuron dendrite was similar to the extent of the 50%-of-peak contour line that we used to define the active plateau region in our study. Could the rise of [Ca2+]i that we observed be caused by release from stores instead of from Ca2+ influx? Several of our observations suggest this is not likely to be the case. The application of specific metabotropic agonists to layer 5 neurons did not evoke plateaus (Greene et al. 1994; Linton et al. 1999). The metabotropic glutamate receptor (mGluR)-dependent release from stores occurred in an all-or-none manner in some CA1 cells during subthreshold EPSPs (without concomitant action potentials), whereas the rise of [Ca2+]i was graded with subthreshold depolarization in our experiments (Fig. 3B) and was abolished by Cd2+ (Fig. 5). If it is accepted that the rise in [Ca2+]i accompanying subthreshold, glutamate-evoked depolarizations up to plateau threshold is mainly due to Ca2+ influx through voltage-gated channels (because of its Cd2+sensitivity), then it is difficult to accept the idea that the [Ca2+]i rise associated with the very next data point, where the plateau is triggered (cf. Fig. 3B), is due instead to Ca2+ release from stores. The latter hypothesis would be plausible only if mGluR-dependent release from stores were the cause of the plateau, but as mentioned in the preceding text, specific metabotropic agonists do not evoke plateaus in the layer 5 neurons. Finally, the conditions that caused the greatest release from stores in the hippocampal neurons (glutamate dependent depolarization concomitant with Na+ spikes) resulted in a much smaller rise of [Ca2+]i and a different spatial-temporal pattern in our experiments than when the glutamate depolarization triggered a plateau in the absence of Na+ spikes (our unpublished observations). Although this question needs to be examined directly in the cortical neurons, the best explanation for the rise in [Ca2+]i in our experiments based on the available evidence is influx through voltage-gated channels, in which case the rise in [Ca2+]i reflects plateau amplitude.

We have interpreted the decline of [Ca2+]i proximal and distal to the iontophoretic site as reflecting decremental conduction of the plateau away from its site of initiation in both directions. We use the term "decremental" to express our uncertainty of whether conduction away from the initiation site is purely passive or, as seems more likely from the observed rise of [Ca2+]i, that the plateau is large enough to activate a nonregenerative Ca2+ current that may augment its conduction compared with the purely passive case. Both the spatially restricted site of plateau initiation and its decremental conduction toward the soma suggested by the imaging data are fully consistent with our previous observations that CDRP amplitude (as recorded in the soma) declines with the distance of the iontophoretic site from the soma (Oakley et al. 2001; Schwindt and Crill 1997). In contrast, an amplitude of the CDRP (measured in the soma) that was independent of iontophoretic position would be expected to result from active conduction to the soma. The decremental somatopedal conduction of a transient Ca2+ spike evoked in the distal dendrites has been observed directly during simultaneous intradendritic and intrasomatic recording (Larkum et al. 1999). Our present data suggest that conduction toward the distal dendritic tree is also decremental. Apparently, neither the initial Ca2+ spike nor the plateau backpropagates like a Na+ spike.

It is unlikely that this local initiation and decremental conduction of CDRPs is caused by a significant decline in the density of Ca2+ channels over the 200-300 µm of the dendrite that we visualized. Other imaging experiments have suggested that Ca2+ channels are present along the entire apical dendritic tree (Markram and Sakmann 1994; Markram et al. 1995; Schiller et al. 1995), and we found that Ca2+ channel density was sufficient to trigger a Ca2+ spike and plateau along the entire length of apical dendrite that we investigated (178-648 µm from the soma).

The absence of an abrupt spatial cutoff of the [Ca2+]i transient prevented a precise identification of the region of regenerative Ca2+ influx. A contour line was drawn around the region where Delta F/F was >= 50% of the peak Delta F/F measured during the plateau to estimate the extent of the region of active (regenerative) plateau initiation. Using this criterion, the extent of the active plateau generating region was remarkably small. Even if we were to assume the active region consisted of the entire distance over which fluorescence during the plateau was significantly higher than fluorescence during a subthreshold response, this would only amount to a couple hundred micrometers. Such a distance is still remarkably short considering the experimental evidence that layer 5 pyramidal neurons are electrically compact (Larkman et al. 1992; Stafstrom et al. 1984). However, the evidence for a long dendritic space constant was obtained under conditions that minimized the activation of voltage-gated conductances. The activation of K+ currents in particular would tend to decrease the effective space constant and thereby help localize a regenerative event (Wilson 1995). The conductance increase caused by the opening of dendritic glutamate channels would further shorten the effective space constant in the region of plateau initiation. The inability of the transient Ca2+ spikes to propagate actively was shown to depend on TEA-sensitive dendritic K+ channels (Schwindt and Crill 1997), and we suppose the same is true for the plateau. The importance of dendritic K+ channels in limiting dendritic excitability was indicated in the present study by the appearance of large-amplitude, repetitive, propagating, Ca2+ spikes in response to an iontophoresis that caused only a low-amplitude plateau before TEA application (our unpublished observations).

If Ca2+ spikes actively propagated to the soma in physiological saline as they do in the presence of TEA, they would dominate the cell's output because they are of such large amplitude. Our present imaging data, together with the electrophysiological results referenced in the preceding text, suggest that the regenerative Ca2+ spike or plateau is normally restricted to a rather small region of the dendrite and is propagated decrementally to the soma to cause a much smaller depolarization. In contrast to the actively propagated Ca2+ spike, the depolarizing current that reaches the soma from a spatially restricted Ca2+ spike can sum with depolarizing currents from other regions of the cell to depolarize the soma. Thus the cell as a whole remains integrative. According to this idea, the integration of synaptic input may occur in two stages. There could be a local integration of synaptic current at one or more dendritic sites to evoke a local transient Ca2+ spikes and/or a plateau. The depolarizing current from these local, dendritic, active responses could then sum with each other near the soma and with synaptic current from other regions of the cell to evoke the propagated Na+ spike that is the output of the cell The rules for integration of plateau currents evoked simultaneously at multiple dendritic sites were studied in a separate set of experiments (Oakley et al. 2001).


    ACKNOWLEDGMENTS

We thank G. Hinz for technical assistance.

This work was supported by National Institute of Neurological Disorders and Stroke Grant NS-16792 and by the Keck Foundation.


    FOOTNOTES

Address for reprint requests: W. E. Crill, Dept. of Physiology and Biophysics, Box 357390, University of Washington School of Medicine, Seattle, WA 98195-7290 (E-mail: wecrill{at}u.washington.edu).

Received 21 June 2000; accepted in final form 19 February 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

0022-3077/01 $5.00 Copyright © 2001 The American Physiological Society