Department of Anatomy and Neurobiology, University of Tennessee, Memphis, Tennessee 38163
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ABSTRACT |
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Stewart, Ansalan and Robert C. Foehring. Calcium Currents in Retrogradely Labeled Pyramidal Cells From Rat Sensorimotor Cortex. J. Neurophysiol. 83: 2349-2354, 2000. Our previous studies of calcium (Ca2+) currents in cortical pyramidal cells revealed that the percentage contribution of each Ca2+ current type to the whole cell Ca2+ current varies from cell to cell. The extent to which these currents are modulated by neurotransmitters is also variable. This study was directed at testing the hypothesis that a major source of this variability is recording from multiple populations of pyramidal cells. We used the whole cell patch-clamp technique to record from dissociated corticocortical, corticostriatal, and corticotectal projecting pyramidal cells. There were significant differences between the three pyramidal cell types in the mean percentage of L-, P-, and N-type Ca2+ currents. For both N- and P-type currents, the range of percentages expressed was small for corticostriatal and corticotectal cells as compared with cells which project to the corpus callosum or to the general population. The variance was significantly different between cell types for N- and P-type currents. These results suggest that an important source of the variability in the proportions of Ca2+ current types present in neocortical pyramidal neurons is recording from multiple populations of pyramidal cells.
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INTRODUCTION |
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Neocortical pyramidal cells express at least five
high-voltage activated (HVA) Ca2+ currents: N-,
L-, P-, Q-, and R-type (Brown et al. 1994;
Lorenzon and Foehring 1995
; Mermelstein et al.
1999
; Ye and Akaike 1993
). Individual pyramidal
cells from the primary motor and primary somatosensory cortices vary
extensively in both the percentage contribution of each
Ca2+ channel type to the whole cell
Ca2+ current (Lorenzon and Foehring
1995
) and in the extent that they are modulated by transmitters
(Foehring 1996
; Stewart et al. 1999
). One
possible source of this variability when recording from dissociated cells is that different amounts of dendrite are associated with the
soma and that there is subcellular compartmentalization of channel
types. Another possible source of variability is recording from
different types of pyramidal cells.
Neocortical pyramidal cells differ in their projections, size, laminar
distribution, dendritic extent, and firing pattern. Pyramidal cells are
classified based on their firing pattern as either regular spiking or
intrinsic bursting (Connors and Gutnick 1990;
McCormick et al. 1985
). These classes roughly correlate with the anatomic connections of the cells (Chagnac-Amitai et al. 1990
; Mason and Larkman 1990
; Wang
and McCormick 1993
). For instance, the majority of
corticotectal neurons are intrinsic bursters (Chagnac-Amitai et
al. 1990
; Mason and Larkman 1990
; Wang
and McCormick 1993
), whereas corticocortical cells tend to be
regular spiking (Gutnick and Crill 1995
). Pyramidal
cells from the neonatal visual cortex with distinct projection targets
(corticocortical vs. corticotectal) display different proportions of
voltage-gated currents such as Ih
(Solomon et al. 1993
) and calcium currents (HVA and
T-type) (Giffin et al. 1991
).
In the study by Giffin et al. (1991), N- and L-type HVA
Ca2+ currents were distinguished based on their
kinetics. The inactivating component of the whole cell
Ba2+ current was assumed to represent the N-type
current whereas the noninactivating component was designated L-type.
Recent studies of Ba2+ currents in pyramidal
cells of the sensorimotor cortex identified with specific organic
calcium channel antagonists indicate that N- and L-type currents have
similar inactivation kinetics in these cells (Brown et al.
1993
; Lorenzon and Foehring 1995
;
Mermelstein et al. 1999
). Furthermore, neocortical
pyramidal cells also express P-, Q-, and R-type
Ca2+ currents (Brown et al.
1994
; Lorenzon and Foehring 1995
;
Mermelstein et al. 1999
; Ye and Akaike
1993
). Thus inactivation kinetics are not a reliable indicator
of channel subtypes in these cells (see also Brown et al.
1993
). Distinguishing Ca2+ current types
using pharmacology is presently considered more reliable and generally
correlates with molecular classification schemes (e.g.,
Birnbaumer et al. 1994
).
There are biophysical differences between high-voltage activated (HVA)
Ca2+ currents in neocortical pyramidal cells
(Brown et al. 1994; Lorenzon and Foehring
1995
; Mermelstein et al. 1999
). Q- and R-type
currents inactivate faster and more completely than L-, P-, and N-type currents (Mermelstein et al. 1999
), and R-type current
activates at more negative voltages than the other four HVA currents
(Lorenzon and Foehring 1995
; Mermelstein et al.
1999
). HVA subtypes also differ in their contribution to
specific cellular events such as neurotransmitter release
(Wheeler et al. 1994
) and afterhyperpolarizations (AHPs)
(Pineda et al. 1998
).
We tested the hypothesis that a major source of the variability in the
percent of Ca2+ channel types expressed in
pyramidal cells is recording from different types of pyramidal cells.
Dissociated cells do not retain detailed morphology and the firing
pattern may depend on intact dendrites (Amitai et al.
1993; Kim and Connors 1993
; Schwindt and
Crill 1999
). Therefore we identified the cells by their
projection pattern. We injected fluorescently labeled beads into
several brain regions to which cortical pyramidal cells send efferents. Pyramidal cells projecting to the injected area were retrogradely labeled (Katz et al. 1984
). Three populations of
retrogradely labeled cells were compared as to the percentage of each
calcium channel type expressed. Our hypothesis predicts that the amount of variability observed within each identified cell type will be less
than the variability in the overall recorded sample. A second
prediction is that the average percentage of each channel type
expressed varies between pyramidal cell types.
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METHODS |
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Retrograde labeling
Neocortical pyramidal cells projecting to either the
contralateral sensorimotor cortex, [callosal projecting (CP)],
striatum (CS), or tectum (CT), were studied. The projection targets of 2-6 wk old Sprague-Dawley rats were pressure injected with
fluorescent red (rhodamine) or green beads (fluorescein) (Katz
and Iarovici 1990; Katz et al. 1984
) using
stereotaxic coordinates from Paxinos and Watson (1986)
scaled to accommodate immature animals. These coordinates were as
follows (distances measured from Bregma). Striatum: anterior posterior
(AP), ~0.7 to
0.2 mm and medial lateral (ML), ~2.0-3.0 mm.
Distance from pial surface: dorsal ventral (DV), ~4-4.5 mm.
Contralateral cortex: AP, ~4-4.5 mm; ML, 1.5-2 mm; and DV, ~2.5
mm. Tectum: AP, approximately
4.2 to
5.5 mm; ML, ~1-1.5 mm, and
DV, ~3.5 mm. Fluorescent latex beads do not alter the
electrophysiological properties of the cell, show minimal diffusion,
and resist fading under fluorescent illumination (Katz and
Iarovici 1990
; Katz et al. 1984
).
Acute isolation of pyramidal cells
One to two weeks after bead injection, the pyramidal cells were
acutely isolated. The rats were anesthetized with methoxyfluorane. Under anesthesia, the rats were decapitated and the brains were extracted. The brains were sectioned into 400 µM slices using a
vibrating tissue slicer (Cambden Instruments) in an oxygenated high-sucrose solution which contained (in mM) 250 sucrose, 2.5 KCl, 1 NaH2PO4, 11 glucose, 4 MgSO4, 0.1 CaCl2, and 15 HEPES (pH = 7.3 adjusted with 1N NaOH; 300 mOsm/l). The slices
were held for a minimum of 1 h in a carboxygen (95%
O2-5% CO2) bubbled
artificial spinal fluid (ACSF). The ACSF contained (in mM) 125 NaCl, 3 KCl, 2 CaCl2, 2 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, 20 glucose, 1 kynurenic acid, 1 pyruvic
acid, 0.1 nitroarginine, and 0.05 glutathione (pH = 7.4 adjusted
with 1 N NaOH; 310 mOsm/l). The sensorimotor cortex (the combined
primary motor and primary somatosensory cortices) was dissected from
these slices with the aid of a stereomicroscope. The dissected cortex
was incubated 20-30 min in oxygenated ACSF containing Pronase E (Sigma
protease type XIV, 1.2 mg/ml at 32°C; modified from Lorenzon
and Foehring 1995). After the incubation period, the tissue was
rinsed in a sodium isethionate solution which contained (in mM) 140 Na
isethionate, 2 KCl, 1 MgCl2, 23 glucose, 15 HEPES, 1 kynurenic acid, 1 pyruvic acid, 0.1 nitroarginine, and 0.05 glutathione (pH = 7.3 adjusted with 1 N NaOH; 310 mOsm/l). The
tissue was triturated in the same solution using fire-polished Pasteur
pipettes. The supernatant was collected and poured into a plastic petri
dish (Lux) positioned on the stage of an inverted microscope (Nikon
Diaphot 300). The cells were allowed several minutes to adhere to the
petri dish and the background flow of HEPES-buffered saline solution
(HBSS) was initiated (~1 ml/min). HBSS contained (in mM) 10 HEPES,
138 NaCl, 3 KCl, 1 MgCl2, and 2 CaCl2 (pH = 7.3 adjusted with 1N NaOH, 300 mOsm/l).
Pyramidal cells innervating the injected brain regions were identified with epifluorecence and excitation filters of either 530-560 nm (to visualize rhodamine-labeled cells) or 450-490 nm (to visualize fluorescein-labeled cells; Fig. 1). Cells were identified as pyramidal by soma shape, the presence of an apical dendrite, and retrograde labeling. We selected cells with apical dendrites of <25 µm length to facilitate voltage control. We recorded from 21 CP, 21 CS, and 18 CT fluorescently labeled pyramidal cells using the whole cell variation of the patch-clamp technique.
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Recording solutions and pharmacological agents
The external recording solution used to isolate the Ca2+ channel currents consisted of (in mM) 125 NaCl, 20 CsCl, 1 MgCl2, 10 HEPES, 5 BaCl2, 0.001 tetrodotoxin (TTX, to block Na+ currents), and 10 glucose (pH = 7.3 adjusted with TEA-OH; 300-305 mOsm/l). The internal recording solution included the following (in mM): 180 N-methyl-D-glucamine (NMG), 4 MgCl2, 40 HEPES, 10 EGTA, 0.1 leupeptin, 0.4 GTP, 2 ATP, and 0.007-0.015 phosphocreatine (pH = 7.2 adjusted with 0.1 N H2SO4; 265-275 mOsm/l).
The stock solutions of the calcium channel antagonists
-conotoxin-GVIA (GVIA; 500 µM),
-conotoxin-MVIIC (MVIIC; 500 µM), and
-agatoxin-IVA (AgTX; 100 µM) were dissolved in water,
measured into aliquots, and frozen. Each of the stocks were diluted to the appropriate concentration in the external recording solution immediately before the experiment. Nifedipine was dissolved in 95%
ethanol before being added to the external solution resulting in a
final concentration of ethanol of <0.05%. This concentration of
ethanol has no effect on Ca2+ currents in these
cells (Lorenzon and Foehring 1995
). Nifedipine was
protected from ambient light. Cytochrome C, at a final dilution of
0.01%, was combined with solutions containing AgTX to prevent nonspecific binding of AgTX to glass and plastic (Bargas et al. 1994
; Lorenzon and Foehring 1995
).
Most chemicals were obtained from Sigma (St. Louis, MO). GTP, ATP and EGTA were obtained from Calbiochem (La Jolla, CA), GVIA and MVIIC were obtained from Bachem (Torrance, CA), and AgTX was a gift from Dr. N. Saccamano (Pfizer).
Whole cell recording
Whole cell recordings were acquired using an Axopatch 200A
electrometer. The recordings were monitored and controlled by pCLAMP6 (Axon Instruments) installed on a 486 computer. The electrodes were
pulled from 7052 glass (Garner) and fire polished. Typically, series
resistance compensations of 70-80% were employed. Cells were not
included in the comparisons of biophysical properties if the estimated
series resistance error was >5 mV (calculated using Ohms Law,
V = I × R, as peak current
multiplied by uncompensated series resistance). Voltage control was
also assessed by observing tail currents after brief voltage steps (see
Lorenzon and Foehring 1995). A gravity-fed parallel
array of glass tubes was used to apply the solutions to the cell being studied.
SYSTAT (SYSTAT, Evanston, IL) was used to carry out all statistical
calculations. Unless otherwise stated, the population data are
represented as median and/or mean ± SE. Data are also presented graphically as box plots. In the box plots, the internal line
represents the median whereas the outer edges of the box represent the
inner quartiles of the data. The bars extending from the box depict the
two outer quartiles of the data. Data points more than two times the
difference between the box edges were considered outliers and are
indicated by an asterisk in the plots (Tukey 1977).
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RESULTS |
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The percentage contributions of each of five HVA
Ca2+ current types described in the cortex
(Lorenzon and Foehring 1995; Mermelstein et al.
1999
; Sayer et al. 1990
) were determined using
selective pharmacological antagonists. The whole cell current was
elicited by a 30-ms voltage step from
90 to
10 mV, which was
repeated every 5 s (Fig. 2). Our
previous studies indicate little run-down of currents with this
protocol (Foehring 1996
; Lorenzon and Foehring 1995
; Mermelstein et al. 1999
)(see also Fig. 2).
L-, P-, and N-type currents were blocked using 5 µM nifedipine, 25 AgTX, and 1 µM GVIA, respectively (Birnbaumer et al.
1994
; Lorenzon and Foehring 1995
). MVIIC blocks
N-, P-, and Q-type currents (Birnbaumer et al. 1994
).
After N- and P-type currents were blocked with nifedipine, AgTX, and
GVIA, Q-type currents could be identified as the current blocked by
MVIIC (Mermelstein et al. 1999
; Randall and Tsien
1995
, 1997
). Because the time constant for the block of Q-type
current is ~140 s (Mermelstein et al. 1999
), we
allowed at least 8-10 min for MVIIC to completely block the Q-type
current. The remaining current that was insensitive to organic
blockers, but was blocked by 400 µM Cd2+, was
designated R-type (Fig. 2) (Randall and Tsien 1995
,
1997
).
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Figure 3 shows the percentage of calcium current contributed by each of the current types in CP cells, CS cells, CT cells, and the overall sample. Each individual box plot indicates the median percentage and the variability (inner and outer quartile ranges) of percent currents within a given cell type. Comparison of the range of values in box plots from different cell types indicates the variability among cell types. Figure 3 shows that the range of percentage of L-type current is similar for CP cells and the general population and is wider in CP cells than for CS or CT cells. The median percentage of N-type current varies among the cell types, with the range of values greater for CP cells and for the general population than for CS and CT cells. The median percentage of P-type current also varies among cell types, with the range of values again greater for CP cells and for the general population than for CS and CT cells. The range of percentage for Q-type currents is wide in all cell types and the medians and ranges are similar among cell groups. For R-type current, the range is narrow for all four cell types, with similar variability in each type.
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We hypothesized that if the variability in the proportions of the
various current types in the overall sample was caused by recording
from multiple pyramidal cell types, each of which is relatively
homogeneous, then the variability observed within each cell type would
be less than the variability in the whole population (including labeled
and unlabeled cells; n = 41). We tested our hypothesis
using both Levene's test for homogeneity of variance and a Bartlett
test (Neter et al. 1996). With both tests we found significant differences in the variance among cell types for N- and
P-type currents (P < 0.05) but not the other channel
types (Fig. 3). The variance is less in CS and CT cells than in CP
cells or the overall sample.
We also hypothesized that the average percent contribution of each
current type would vary significantly among cell types. We tested this
hypothesis with a one-way analysis of variance (ANOVA) with planned
contrasts (Neter et al. 1996). We found significant differences among cell types for the means for L-, N-, and P-type currents (P
0.02), but not Q- or R-type currents
(Table 1). CP cells were similar to the
overall sample in mean percentages. CS cells had a lower percentage of
P-type and CT cells had a lower percentage of L- and N-type than the
other cell types.
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DISCUSSION |
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Pyramidal cells are the output cells of the neocortex, and
the projection target of these cells generally depends on the layer in
which their soma resides. The cell bodies of pyramidal cells sending
efferents to the contralateral cortex can be found mainly in layers III
and VI. CP cells are a heterogeneous group in that some cells also send
collaterals to the striatum (Wilson 1987) or the spinal
cord (White 1989
). Corticostriatal cells originate primarily from layer III and superficial layer V. Some pyramidal cells
in layer V also project to the tectum (White 1989
).
As a whole, pyramidal cells vary extensively in terms of the percentage
contribution of each Ca2+ current type to the
whole cell Ca2+ current (Lorenzon and
Foehring 1995) and the extent to which they are modulated
(Foehring 1996
; Stewart et al. 1999
).
This variation is potentially functionally significant; therefore we used organic and inorganic Ca2+ channel
antagonists to test the hypothesis that this variability resulted in
part from sampling from a heterogeneous population of pyramidal cells.
We identified cells with retrograde labeling and determined the
percentage contribution of L-, N-, P-, Q-, and R-type currents to the
whole cell Ba2+ current in three populations of
pyramidal cells (CP, CS, and CT).
If the variability in the overall sample was a result of recording from multiple populations of pyramidal cells, then we predict that the mean contribution of Ca2+ channel types would differ between defined pyramidal cell populations. We found significant differences among the three pyramidal cell types in the mean percent of L-, P-, and N-type currents but no significant differences for Q- or R-type currents.
A second prediction is that the variability of percentage contribution of channel types should be less within defined pyramidal cell populations than in the overall sample. The medians and ranges of values can be seen in the box plots (Fig. 3). For L-type current, the range is small for CS and CT cells but large for CP and the combined population. No statistical differences were found in the variance of L-type contribution among cell types. For both N- and P-type currents, the range of values is small for CS and CT cells as compared with CP cells and the general population ("All" in Fig. 3) and the variance significantly differs across cell types. The percentage of Q-type current varies greatly within all cell types and as a result each cell group has a similar range. The range of values for R-type currents are small and similar in all cell types. There were no differences in the variance between cell types for Q- or R-type currents.
Our combined findings suggest that much of the variability across
pyramidal cells in the percentage each current type contributes to the
whole cell current is a result of differences among multiple pyramidal
cell populations. These differences may have functional consequences,
as somatodendritic N-, P-, and Q-type channels (but not L-type) elicit
Ca2+ -dependent K+ currents
underlying spike frequency adaptation in these cells (Pineda et
al. 1998). L-type currents contribute to inward
currents underlying repetitive firing in neocortical pyramidal neurons (Pineda et al. 1998
) and are preferentially
involved in regulating gene expression in many cell types (Bito
et al. 1997
). Both CS and CT cells were more homogeneous in
percentage contribution of channel types than either CP cells or the
overall sample. Interestingly, CP cells were similar to the total
population in both mean percentages and in the degree of variability.
This may reflect the fact that CP cells, which send collaterals to
striatum and other targets, are a more heterogeneous group than CS or
CT cells. Collectively, the findings suggest that recording from a
diverse population of neurons is a significant source of the
variability in the proportions of HVA currents noted in earlier studies
of pyramidal cells. Further tests are required to determine the
functional correlates of this variability.
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ACKNOWLEDGMENTS |
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The authors thank Drs. D. Mendelowitz, R. Scroggs, D. J. Surmeier, S. Timmons, R. Waters, and C. J. Wilson for comments on earlier versions of this manuscript. Statistical help was provided by K. L. Arheart. Excellent technical support was provided by C. Windham.
This work was supported by National Institutes of Health Grants NS-33579 to R. C. Foehring and 5T32MH-19547 to A. E. Stewart.
Present address of A. E. Stewart: Dept. of Pharmacology, The George Washington University, 634 Ross Hall, 2300 Gye St., N.W., Washington, DC 20006.
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FOOTNOTES |
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Address for reprint requests: R. C. Foehring, Dept. of Anatomy and Neurobiology, University of Tennessee, 855 Monroe Ave., Memphis, TN 38163.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 23 August 1999; accepted in final form 21 December 1999.
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REFERENCES |
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