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The Journal of Neuroscience, August 1, 2002, 22(15):6380-6387
Cholecystokinin Tunes Firing of an Electrically Distinct
Subset of Arcuate Nucleus Neurons by Activating A-Type
Potassium Channels
Denis
Burdakov and
Frances M.
Ashcroft
University Laboratory of Physiology, Oxford, OX1 3PT, United
Kingdom
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ABSTRACT |
The physiological activity of hypothalamic arcuate nucleus (ARC)
neurons is critical for dynamic maintenance of body energy homeostasis,
and its malfunction can result in common metabolic disorders, such as
obesity. It is therefore of interest to determine which set of ion
channels shapes electrical activity in the ARC. Whole-cell patch clamp
of ARC neurons in mouse brain slices identified three
electrophysiologically distinct types of neurons. These were
distinguished by their rebound "signatures" after hyperpolarizing current injection in current clamp and by the presence of transient inward (Type-B neurons) or outward (Type-A and Type-C neurons) subthreshold voltage-gated currents in voltage-clamp recordings. In
turn, the transient outward current (A-current) of Type-C neurons had a
lower activation threshold and different time and voltage dependence of
inactivation than that of Type-A neurons. The brain-gut peptide
cholecystokinin (CCK) has long been recognized to control food intake,
but how endogenous CCK modulates the activity of central
appetite-regulating networks remains unresolved. Here, we show that low
(picomolar) concentrations of CCK rapidly and reversibly slow the
firing of ARC Type-C neurons. This effect is mediated by postsynaptic
CCK-B receptors and is attributable to potentiation of the
A-current. Our study thus identifies several fundamental biophysical
mechanisms underlying the physiological activity of ARC neurons and
suggests a novel mechanism by which endogenous CCK may control appetite.
Key words:
hypothalamus; arcuate nucleus; appetite; energy
homeostasis; electrophysiology; CCK; A-currents; CCK-B receptor
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INTRODUCTION |
The hypothalamus maintains body
energy homeostasis by integrating neural, endocrine, and metabolic
information to elicit behavioral, autonomic, and endocrine responses
(Elmquist et al., 1999 ; Schwarz et al., 2000). Among the hypothalamic
regions involved, the arcuate nucleus (ARC) has recently gained
much prominence because of several findings. First, ARC neurons are the
source of several powerful neuropeptide modulators of food intake and
metabolic rate, such as  -melanocyte-stimulating hormone, cocaine-
and amphetamine-regulated transcript, neuropeptide Y, and agouti
protein (Elias et al., 1998; Hahn et al., 1998). Second, mRNA levels of
these peptides in the ARC are changed by perturbations of body energy
balance, such as fasting, and by changes in blood levels of insulin and leptin (Schwarz et al., 2000). Third, ARC neurons are strategically located near a "window" in the blood-brain barrier at the median eminence (Elmquist et al., 1999; Ganong, 2000), which allows them to
directly translate blood-borne signals of body energy status, such as
glucose, insulin, and leptin (Spanswick et al., 1997, 2000; Cowley et
al., 2001) into neuronal activity.
Together, these findings indicate that ARC neurons act as key
integrators of peripheral and central signals of body energy status and
are critical for central control of body energy balance (Schwarz et
al., 2000). Furthermore, defects in ARC function have been linked to
obesity (Barsch et al., 2000). It is therefore important to determine
the biophysical mechanisms that control the electrical activity of ARC
neurons and thereby the release of the neuropeptides they contain.
Within the hypothalamus, the region best characterized
electrophysiologically is the ventromedial nucleus. In neurons from
this region, spontaneous firing and responses to perturbations of
membrane potential are probably determined by currents with activation
thresholds below that of the voltage-gated Na+ channel (subthreshold currents)
(Minami et al., 1986; Miki et al., 2001). Some ARC neurons are also
spontaneously active (Rauch et al., 2000), suggesting that subthreshold
currents may shape their electrical activity. Of particular interest
are the currents that directly tune neuronal firing and are under
control of neuromodulators, such as hyperpolarization-activated
currents (Lüthi and McCormick, 1998) and A-type
K+ currents (Connor and Stevens, 1971;
Liss et al., 2001; Yang et al., 2001).
Cholesystokinin (CCK), which functions as both a hormone and a
neurotransmitter (Crawley and Corwin, 1994), has long been recognized
to play a key role in the short-term control of food intake by inducing
satiety (Gibbs et al., 1973; Moran, 2000). However, the mechanism by
which endogenous CCK controls appetite is still only poorly understood,
and both the site of CCK action (central or peripheral) and the type of
receptor involved (CCK-A or CCK-B) remain controversial (Baldwin et
al., 1998). The ARC is a target for both synaptic (Siegel et al., 1987;
Ciofi and Tramu, 1990; Baldwin et al., 1998) and probably also
blood-borne (Ganong, 2000) CCK. It is therefore possible that
endogenous CCK may modulate feeding by interacting with ARC neurons.
We used the whole-cell patch-clamp technique to investigate the
intrinsic biophysical properties and CCK responsiveness of ARC neurons
in mouse brain slices.
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MATERIALS AND METHODS |
Brain slice preparation. Procedures involving animals
were performed in accordance with the Animals (Scientific Procedures) Act, 1986 (United Kingdom). C57BL/6 mice (12-16 d postnatal) were killed by cervical dislocation. Brains were removed quickly, immersed in ice-cold solution, and then blocked for slicing. Coronal slices (250- to 300-µm-thick) containing the arcuate nucleus were cut with a
Vibroslice (Campden Instruments, London, UK), while bathed in ice-cold
artificial CSF (ACSF), containing (in mM)
the following: 118 NaCl, 25 NaHCO3, 3 KCl, 1.2 NaH2PO4, 1.5 CaCl2, 1 MgCl2, and 10 glucose (bubbled with a mixture of 95% O2 and
5% CO2). After sectioning, slices were allowed
to recover for >20 min in a chamber filled with gassed ACSF at room
temperature (22-24°C) before the experiment.
Electrophysiology and data analysis. For whole-cell
patch-clamp recordings, slices were continuously perfused at 2-4
ml/min with ACSF bubbled with a mixture of 95%
O2 and 5% CO2. Patch
pipettes were pulled from borosilicate glass (GC150TF/F; Clark,
Reading, UK) and had tip resistances of 3-5 M when filled with
internal solution, which contained (in mM) the
following: 130 K-gluconate, 1 NaCl, 0.1 EGTA, 1 MgCl2, 10 HEPES, and 5 K2-ATP, pH 7.3 adjusted with KOH. A low
concentration of EGTA was used to mimic physiological cytosolic
Ca2+ buffering and to preserve
Ca2+-dependent
K+ currents, which can affect the
spontaneous firing rates of neurons (Wolfart et al., 2001). Liquid
junction potentials were measured in separate experiments, in which the
bath (connected to the reference electrode by a 3 M KCl agar bridge) was perfused alternately with ACSF or the internal solution, and the zero-current pipette potentials in the two solutions were compared. Junction potentials were <8 mV and
were not corrected for. The arcuate nucleus was located using a mouse
brain stereotaxic atlas (Franklin and Paxinos, 1997). Neurons
were visualized by infrared interference contrast video microscopy with
a Newvicon camera (Hamamatsu, Hamamatsu City, Japan), mounted to an
upright microscope (Axioscop FS; Zeiss, Oberkochen, Germany) (Stuart et
al., 1993). Whole-cell recordings were performed in current- or
voltage-clamp mode using an EPC-9 patch-clamp amplifier (Heka
Elektronik, Lambrecht, Germany). Only cells with an input resistance of
>0.8 G were used. The series resistance was <15 M and was not
compensated. Data was sampled (at 3 kHz) and analyzed using the program
package Pulse+pulsefit (Heka Elektronik). The voltage dependence of
A-current inactivation (see Figs. 4B, 5B)
was fitted by the Boltzman equation,
 using Microcal Software (Northampton, MA) Origin software. Here, Vm is the membrane potential,
V0.5 is the membrane voltage at which
the normalized current amplitude
(I/Imax) is half-maximal, and k is the slope factor. Concentration-response data for
CCK were fitted according to the Hill equation,  I = Imax  Imax/{1 + ([CCK]/EC50)h}, where
I is the increase in current produced by CCK,
Imax is the maximum current
increase, [CCK] is the concentration of CCK, and h is the
Hill coefficient. Statistical significance was tested using a
two-tailed unpaired t test (Origin software; Microcal Software). Data are given as mean ± SEM unless stated otherwise.
Chemicals. CCK-8S (the main molecular form in both the CNS
and the periphery) (Dockray 1978; Baldwin et al., 1998) was obtained from Calbiochem (La Jolla, CA). CCK-8S, gastrin (Calbiochem), or
4-aminopyridine (4-AP) (Sigma, St. Louis, MO) were applied locally at a
rate of 35 µl/min (controlled by a syringe pump system) through a
quartz pipette (0.25 mm inner diameter) placed over the ARC using a
second manipulator. Tetrodotoxin (TTX) (Sigma) was included in the
extracellular solution in some experiments, as indicated.
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RESULTS |
Three distinct electrophysiological phenotypes of ARC neurons
Current-clamp analysis of ARC neurons revealed that they fall into
three electrically distinct types. "Type-A" neurons (membrane capacitance, 7-16 pF; n = 80) fired spontaneously at
5-10 Hz and immediately resumed normal firing after hyperpolarizing
current injections (Fig.
1A). "Type-B"
neurons (membrane capacitance, 10-25 pF; n = 16) fired
spontaneously at 2-8 Hz and showed a slow rebound depolarization spike
at the end of a hyperpolarizing current pulse (Fig.
1B). "Type-C" neurons (membrane capacitance,
15-35 pF; n = 34) exhibited spontaneous activity at
1-5 Hz (n = 27) or were electrically silent with
resting membrane potentials between 45 and 50 mV (n = 7); on recovery from hyperpolarizing pulses, these cells displayed a
rebound hyperpolarization, which caused them to resume firing with a
prominent delay (Fig. 1C). After a hyperpolarizing pulse to
80 mV, the latency to the first spike was 51 ± 6 msec in
Type-A cells (n = 80), 4.7 ± 3.5 msec in
Type-B cells (n = 16), and 648 ± 83 msec in
Type-C cells (n = 27). The latency was significantly
longer in Type-C cells than in either Type-A or Type-B cells
(p < 0.00001 in both cases).
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Figure 1.
Three distinct electrophysiological phenotypes of
ARC neurons. These phenotypes were distinguished by their recovery from
hyperpolarizing current injections (30-80 pA) in current clamp
(left) and by the currents elicited by a voltage step
from 90 to 40 mV in voltage clamp (right). Current-
and voltage-clamp protocols are displayed schematically below.
A, Type-A ARC cells show no rebound potential
(left) and a small, rapidly inactivating outward current
(right) in response to repolarization from
hyperpolarized potentials. B, Type-B ARC neurons exhibit
rebound depolarization (left) and a
prominent subthreshold inward current
(right). C, Type-C ARC neurons display
rebound hyperpolarization (left) and a large outward
current that inactivates slowly relative to that of Type-A cells
(right). Horizontal lines on the
current-clamp traces
(left) show the zero-current potential.
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To visualize the currents that underlie these three types of rebound
potential, we used a voltage-clamp protocol in which cells were held at
90 mV for 300 msec and then stepped to 40 mV. This protocol
revealed fundamental differences in subthreshold currents in the three
cell types. In Type-A cells, a voltage step from 90 to 40 mV evoked
a small (44 ± 3 pA), fast-inactivating outward current
(n = 80) (Fig. 1A), whereas in Type-C
cells, it induced a large (248 ± 14 pA), slowly activating and
inactivating outward current (n = 34) (Fig.
1C). The amplitudes of these currents were significantly
different (p < 0.00001). In Type-B cells, the same voltage step evoked a transient inward current (34 ± 3 pA; n = 16) (Fig. 1B).
The differences in the recovery from hyperpolarization and the
properties of voltage-gated currents were sufficient to identify unambiguously the type of each ARC neuron.
Absence of hyperpolarization-activated current in ARC neurons
ARC cells often exhibit regular spontaneous firing (Fig. 1). In
extracellular recordings, this activity is not abolished by blockade of
synaptic transmission (Rauch et al., 2000), which suggests that ARC
neurons may have intrinsic pacemaker activity. We therefore
investigated whether the hyperpolarization-activated current
(Ih), which can be critical for
neuronal pacemaking (Lüthi and McCormick, 1998), is present in
ARC neurons.
Hyperpolarizing current injections failed to elicit the typical
"sag" in membrane potential characteristic of
Ih activation in all three types of
ARC neurons (Fig.
2A-C), in contrast to neurons from the neighboring dorsomedial nucleus (n = 4) (Fig. 2D). A voltage-clamp step from 40 to 120
mV confirmed that Ih is elicited by
hyperpolarization in dorsomedial nucleus cells (n = 3)
(Fig. 2D) but not in ARC neurons (Fig.
2A-C). Thus, our results suggest that
Ih channels with fast activation
kinetics (i.e., those containing HCN1 subunits; Franz et al., 2000) are not present in all three types of ARC cells. We were unable to determine whether Ih currents with
slower activation kinetics (channels lacking HCN1; Franz et al., 2000)
are present in the ARC, because this requires holding cells at voltages
negative to 90 mV for >2 sec (Franz et al., 2000), which was not
well tolerated by ARC neurons in our experiments. However, because ARC
cells fire at 2-10 Hz (Fig. 1), it seems unlikely that a slowly activating Ih current would contribute
significantly to spike-to-spike pacemaking in the ARC.
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Figure 2.
Absence of fast Ih
currents in ARC neurons. Overlay of membrane potential responses to
hyperpolarizing current injections of increasing intensity
(left) and currents recorded in response to
hyperpolarization from 40 to 120 mV (right) in ARC
(A-C) or dorsomedial nucleus
(D) neurons. The prominent
Ih-mediated sag in potential and rapidly
activating inward current found for dorsomedial nucleus neurons
(D) are not observed for ARC neurons
(A-C). Current- and voltage-clamp protocols are
shown schematically below representative traces.
Horizontal lines on the current-clamp
traces (left) indicate the zero potential. Note
the difference in current scale in A-C
(right).
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Type-B ARC neurons express a low-threshold
Ca2+ current
The transient inward current and rebound depolarization in Type-B
neurons (Fig. 1B) were significantly reduced or
abolished by switching to ACSF containing low (0.3 mM) Ca2+ and high (9 mM) Mg2+
(n = 3), suggesting that they were attributable to
activation of a Ca2+ current. Voltage
steps to between 80 and +10 mV from a holding potential of 90 mV
revealed that this current activated between 60 and 50 mV
(n = 10) (Fig. 3). This
activation threshold is characteristic of neuronal T-type
Ca2+ channels (Tsien et al., 1988).
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Figure 3.
Voltage dependence of activation of
Ca2+ current in Type-B cells. TTX (700 nM) was added to block Na+ currents.
A, Representative example of membrane currents
(top) evoked by the voltage protocol below. The cells
were held at 90 mV for 300 msec and then stepped to +10
mV in 10 mV increments for 250-300 msec (interpulse interval of 2 sec). B, Peak amplitudes of resulting currents were
measured relative to the baseline at 90 mV and plotted against the
pulse potential. The data are representative of 10 cells. The
inset is an expansion of the threshold region and shows
that the inward current activates between 60 and 50 mV.
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A-Currents in Type-A and Type-C ARC neurons are different
The rapid activation and inactivation of the outward current
observed in Type-A and Type-C ARC neurons is typical of A-type potassium channels. Furthermore, the transient outward current was
completely eliminated by the A-current blocker 4-aminopyridine (10 mM; n = 4) (see Fig. 7B). A-Type
potassium currents are key determinants of neuronal firing frequency
(Rudy, 1988; Liss et al., 2001) and are modulated by neurotransmitters
(Aghajanian, 1985) and neurotrophic factors (Yang et al., 2001).
We therefore analyzed the properties of the A-current in Type-A and
Type-C ARC neurons in detail. Type-B neurons appeared to express a fast transient outward current similar to the A-current in Type-A cells (Fig. 3); we did not analyze this current further because of the low
numbers of Type-B cells in the ARC and because it was partially shunted
by inward currents activated in the same voltage range (Fig. 3).
Voltage steps to between 80 and +10 mV from a holding potential of
90 mV revealed that, in Type-A cells, the A-current activated between
40 and 30 mV (n = 12) (Fig.
4A), whereas in Type-C
cells, it activated at a much lower threshold, between 60 and 50 mV (n = 12) (Fig.
5A). The voltage dependence of
A-current inactivation was also significantly different in the two
cell types: half-maximal inactivation occurred at 68 ± 1 mV
in Type-A cells (Fig. 4B) and at 60 ± 1 mV in
Type-C cells (p < 0.0001) (Fig. 5B).
In addition, the time course of A-current inactivation in Type-C cells
was substantially slower than that in Type-A cells (Fig.
6), consistent with the differences in
rebound potentials in the two cell types (Fig.
1A,C). The rate of A-current
inactivation was essentially voltage independent in both cell types
(Fig. 6), as found for other neurons (Liss et al., 2001).
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Figure 4.
Voltage dependence of A-current activation and
inactivation in ARC Type-A neurons. TTX (700 nM) was added
to block Na+ currents. A , Current-voltage relationship. The protocol is the same as in Figure 3.
The data are representative of 12 cells. The inset is an
expansion of the threshold region and shows that the A-current diverges
from the linear leak current at potentials positive to 40 mV.
B, Voltage dependence of inactivation. The
middle and top parts show the
voltage-clamp protocol and an example of the data obtained,
respectively. The protocol consisted of a 300 msec prepulse to between
120 and 20 mV (in 10 mV increments), followed by a 250 msec test
pulse to 0 mV. The peak currents during the test pulse were expressed
as a fraction of the maximum and plotted against the prepulse potential
(bottom; n = 7). The line is the
best fit of the data to the Boltzman equation (see Materials and
Methods).
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Figure 5.
Voltage dependence of A-current activation and
inactivation in ARC Type-C neurons. TTX (700 nM) was added
to block Na+ currents. A, Whole-cell
currents (top), voltage protocol
(middle), and current-voltage relationship
(bottom). The data are representative of 12 cells. The
inset is an expansion of the threshold region and shows
that the A-current diverges from the linear leak current at potentials
positive to 60 mV. B, Voltage dependence of
inactivation (n = 6). The line is the best fit of
the data to the Boltzman equation. All protocols are the same as in
Figure 4.
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Figure 6.
A-Current inactivation is slower in Type-C than in
Type-A neurons. TTX (700 nM) was added to block
Na+ currents. A, Examples of
A-currents elicited by steps from 40 to 0 mV from a holding potential
of 90 mV. A monoexponential fit (I = I0 + A
exp{(t t0)/ }) to the A-current decay
was used to calculate the inactivation time constant, .
B, Voltage dependence of in Type-C cells
(open circles; n = 5) and Type-A
cells (filled circles; n = 5).
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CCK acutely slows firing and potentiates A-currents in ARC
Type-C neurons
We next examined whether CCK modulates the electrical
activity of ARC neurons. The firing frequency of Type-A
(n = 5) and Type-B (n = 4) neurons was
unaffected by 40 pM CCK. However, in Type-C ARC
neurons, CCK (40 pM) slowed firing within 0.5-5
min of application, without inducing detectable hyperpolarization (n = 4) (Fig.
7A). This type of behavior can
be caused by an increase in A-current activity (Liss et al., 2001). To
address this possibility, we monitored A-currents and firing frequency
before and after stimulation with CCK in the same Type-C neurons.
Because no detectable activation of delayed outward current was seen at
40 mV in Type-C cells (n = 10) (Fig. 7B),
we measured A-current amplitude at this potential. We found that the
magnitude of the A-current, but not of the steady-state current, was
greater after CCK application (n = 4) (Fig.
7B). Furthermore, the current activated by CCK was abolished
by 10 mM 4-AP. This suggests that CCK selectively
affects the A-current. Plotting the decrease in firing frequency
against the corresponding increase in A-current amplitude revealed a
strong linear correlation (r = 0.99) (Fig.
7C), as would be expected if CCK-induced slowing of firing
was attributable to an increase in the A-current.
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Figure 7.
CCK slows firing and activates A-currents in ARC
Type-C neurons. A, Membrane potential recordings of a
Type-C neuron before, during, and after CCK application. Action
potentials are truncated at 0 mV, and the dotted line
indicates 50 mV. B, Voltage-clamp recordings of the
corresponding A-currents taken from the same cell. The cell was held at
90 mV for 300 msec and then stepped to 40 mV for 800 msec to elicit
the A-current. Note that the peak A-current is greater in the presence
CCK, but the steady-state current amplitude is unchanged (see Results).
4-AP at 10 mM abolished the A-current. The
traces in A and B are
representative of four cells. C, CCK-induced decrease in
firing frequency plotted against the percentage of activation of the
A-current for four different neurons. There is a strong linear
correlation (r = 0.99; slope of 1.95).
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The mechanism of CCK modulation of A-currents in ARC
Type-C neurons
We next explored the mechanism underlying the activation of
A-currents in ARC Type-C cells by CCK (Fig.
8). Neurons were synaptically isolated by
including 1 µM TTX in the extracellular solution and voltage clamped at 60 mV to prevent membrane potential changes from
influencing A-current activity. A-Current amplitude was measured every
30 sec using the protocol shown in Figure 8A. Under
these conditions, CCK (2-40 pM) induced a
reversible (Fig. 8A) and dose-dependent (Fig.
8B) increase in A-current amplitude. Above 40 pM, CCK was less effective at increasing
A-currents (n = 4), probably because of the rapid
desensitization that is typical of high-affinity CCK receptors
(Burdakov and Galione, 2000). The best fit of the Hill equation to the
concentration-response data gave a maximum response of 31% and an
apparent EC50 of 18.7 pM
(n = 12).
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Figure 8.
Pharmacological and kinetic analysis of CCK
modulation of A-currents in ARC Type-C neurons. TTX (1 µM) was present to block Na+ currents.
A, Time course of A-current increase by CCK. The
inset shows the voltage-clamp protocol used to monitor
the A-current amplitude. The figure is representative of responses in
eight cells. B, Relationship between CCK concentration
and the increase in A-current ampli- tude. The line is drawn to the Hill equation with
EC50 of 18.7 and h = 1.32 (n = 12). For details, see Results.
C, The voltage dependence of A-current inactivation
(measured as in Fig. 5B) was not significantly shifted
by CCK (n = 5; control,
V0.5 = 58 ± 2 mV; CCK,
V0.5 = 60 ± 1 mV;
p > 0.1). D, Time dependence
of A-current recovery from inactivation. Cells were held at 0 mV for
500 msec to inactivate A-currents and then stepped to 90 mV for
varying durations to remove inactivation, after which the A-current
amplitude was measured at 40 mV. The inset shows the
protocol and an example of the data obtained. CCK did not significantly
change the time dependence of recovery from inactivation
(n = 3).
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The potentiation of the A-current by CCK was specific to ARC Type-C
cells: kinetically similar A-currents in neurons of the neighboring
premammillary nucleus were not modulated by the peptide (n = 4). The voltage and time dependence of A-current
inactivation were not significantly different before and after
application of CCK (Fig. 8C,D), suggesting that
CCK enhances the activation of the A-current without affecting its inactivation.
Two CCK receptor subtypes are known to exist. The affinity of the CCK-A
receptor for CCK-8S is 1000-fold higher than that for gastrin, whereas
the CCK-B receptor has similar affinities for gastrin and CCK-8S
(Crawley and Corwin, 1994). Application of 40 pM gastrin
mimicked the effects of 40 pM CCK on firing and A-currents
in ARC Type-C cells (n = 4) (Fig.
9). These results suggest that CCK slows
the firing and potentiates the A-current in ARC Type-C cells by
activation of postsynaptic CCK-B receptors.
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Figure 9.
CCK slows firing and activates A-currents in ARC
Type-C cells by acting on CCK-B receptors. Top trace,
Expansions of membrane potential recordings before, during, and after
application of 40 pM gastrin. Action potentials are
truncated at 0 mV. Bottom traces, Corresponding
A-currents, measured using the same protocol as in Figure
8A. The data are representative of four
cells.
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DISCUSSION |
In this study, we show that three biophysically distinct types of
neurons are found in the ARC, which can be readily identified on the
basis of their firing patterns and membrane currents. Each cell type
has a unique recovery behavior after a hyperpolarizing current
injection in current clamp (Fig. 1). Furthermore, voltage-clamp analysis shows that Type-B cells differ from Type-A and Type-C cells in
that they exhibit a pronounced low-threshold
Ca2+ current (Fig. 3), whereas Type-A and
Type-C cells can be readily distinguished by differences in their
A-currents (Figs. 4-6). The A-current of Type-C cells (Fig.
5B) is much less inactivated in the resting potential region
(40 to 50 mV) than that of Type-A cells (Fig.
4B), and it inactivates much more slowly (Fig. 6). These differences probably account for the finding that Type-C cells
fire at lower rates than Type-A cells.
Modulation of Ih by
neurotransmitter-generated intracellular second messengers such as cAMP
can influence cell firing (Brown et al., 1979; Lüthi and
McCormick, 1998). Our demonstration that ARC cells do not posses
Ih currents (Fig. 2) suggests that
such mechanisms probably do not play a role in modulating the
electrical activity of the ARC.
We found that the relatively small increases in A-current amplitude
produced by CCK in ARC Type-C cells led to large changes in neuronal
firing frequency (1.95% change in frequency per 1% A-current change)
(Fig. 7C). A similar steep linear dependence of firing
frequency on A-current amplitude was reported recently for dopaminergic
midbrain neurons (1.78% change in frequency per 1% A-current change;
Liss et al., 2001). Modulating A-current activity therefore seems to be
a powerful mechanism for controlling neuronal firing rates in several
brain regions.
Our finding that CCK acutely regulates firing frequency in ARC neurons
may help to explain how endogenous CCK controls appetite. There is
considerable evidence that CCK modulates food intake via both hormonal
(Lieverse et al., 1995) and synaptic (McLaughlin et al., 1985; Scallet
et al., 1985; Schick et al., 1986a, 1987; Sodersten and Linden, 1992)
modes of action. ARC neurons are likely to be exposed to CCK from both
of these sources. The absence of a blood-brain barrier at the median
eminence (Broadwell and Brightman, 1976; Ganong, 2000) implies that
they can directly sense circulating CCK. In this respect, it is
significant that, in our study, CCK modulated firing of ARC Type-C
cells in the same concentration range (low picomolar) (Fig.
8B) as that which is found physiologically in the
blood (Liddle et al., 1984) and within which the classical targets of
blood-borne CCK, such as the exocrine pancreatic cells, are activated
by the peptide (Petersen et al., 1991; Burdakov and Galione, 2000). It
is therefore possible that modulation of ARC neurons by CCK may be
involved in the negative feedback control of appetite by circulating
CCK, which is considered to be responsible for meal termination
(Crawley and Corwin, 1994). In turn, the presence of CCK-immunoreactive
nerve fibers and terminals in the ARC (Ciofi and Tramu, 1990) suggests
that ARC neurons are the target of synaptically released CCK. Release
of CCK within the ARC is triggered by conditions such as stress (Siegel
et al., 1987), and thus modulation of ARC neurons by synaptically
released CCK may be involved in stress-appetite interactions.
Both CCK-A and CCK-B receptors are found in the hypothalamus, but their
relative importance in appetite regulation by endogenous CCK is a
matter of considerable debate (for review, see Baldwin et al., 1998).
Our data suggest that CCK modulates the activity of the ARC appetite
center via CCK-B receptors (Fig. 9). The apparent EC50 of CCK-8S for increasing A-current amplitude
(Fig. 8B) may be an underestimate because of
desensitization. However, it is noteworthy that it is in reasonable
agreement with CCK-8S IC50 values determined for
the CCK-B receptor in radioligand displacement studies (Kopin et al.,
1994). The importance of the CCK-B receptor in appetite regulation is
supported by a number of other pharmacological studies. For example,
intracerebroventricularly administered CCK-8S and gastrin-17II were
equally potent in reducing food intake (Schick et al., 1986b), as would
be expected if CCK-B receptors are involved (Jensen et al., 1989).
Studies with the CCK-B-selective antagonist L365,260 (Dourish et al.,
1989; Schick et al., 1991; Corp et al., 1997) also suggest that CCK-B
receptors are involved in modulation of food intake.
CCK increased A-currents and slowed firing within 0.5-5 min of
application (Figs. 7, 8). The variability in the latency of this
response probably reflects differences in drug accessibility, because
different neurons were located at different depths within the slice.
However, no detectable response was observed within 0.5 min of
application, even in neurons lying on the slice surface. This suggests
that CCK action involves the generation of intracellular second
messenger(s). In some non-excitable cells, it has been shown that
activation of CCK-B receptors stimulates the phospholipase C/inositol
trisphosphate pathway and thereby mobilizes intracellular Ca2+ (Noble and Roques, 1999). Whether a
similar transduction pathway exists in ARC Type-C cells remains to be resolved.
In summary, our results provide a biophysical framework for
understanding the physiological and pathological activity of the ARC.
In particular, we show for the first time that circulating concentrations of CCK modulate the activity of a specific subset of ARC
neurons. This novel mechanism is likely to be involved in the
regulation of appetite by this important brain region.
| |
FOOTNOTES |
Received Feb. 19, 2002; revised May 9, 2002; accepted May 15, 2002.
We thank Dr. Stefan Trapp for instruction on working with brain slices
and the Wellcome Trust for support. D.B. holds a Wellcome Prize
Studentship, and F.M.A. holds the Royal Society GlaxoSmithKline Research Professorship.
Correspondence should be addressed to Prof. F. M. Ashcroft,
University Laboratory of Physiology, Parks Road, Oxford, OX1 3PT, UK.
E-mail: frances.ashcroft{at}physiol.ox.ac.uk.
| |
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ABSTRACT
INTRODUCTION
