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The Journal of Neuroscience, August 1, 2002, 22(15):6303-6308
BRIEF COMMUNICATION
Orexin-A Depolarizes Dissociated Rat Area Postrema Neurons
through Activation of a Nonselective Cationic Conductance
Bo
Yang and
Alastair V.
Ferguson
Department of Physiology, Queen's University, Kingston, Ontario,
Canada K7L 3N6
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ABSTRACT |
The area postrema (AP) is involved in the regulation of body fluid
balance, feeding behavior, and cardiovascular function. Orexin (ORX)-A
is a 33 aa peptide that regulates energy metabolism and sympathetic and
cardiovascular actions. ORX immunoreactive axons and their varicose
terminals have been found in AP. In this study, whole-cell, current- or
voltage-clamp recordings were obtained from 108 dissociated rat AP
neurons. The mean resting membrane potential of these neurons
(n = 48) was  59.24 ± 0.87 mV, the mean
input resistance was 3.57 ± 0.22 G , and the action potential amplitude of these cells was always >90 mV. Current-clamp studies showed bath application of ORX-A depolarized the majority of AP neurons
tested (68.8%; 33 of 48), whereas small proportions of cells were
either hyperpolarized (16.7%; 8 of 48) or unaffected (14.6%; 7 of
48). These depolarizing effects were found to be concentration
dependent from 10 8 to 1011
M. We then examined the contributions of specific ionic
conductances to the ORX-A-induced excitation of AP neurons through
whole-cell, voltage-clamp studies. Our results demonstrate that in
contrast to previous studies on other neuronal populations, ORX-A did
not affect net whole-cell potassium currents in AP neurons. Slow
depolarizing voltage ramps, however, revealed that ORX-A enhanced a
nonselective cationic conductance in AP neurons, effects which would
explain the depolarizing effects of the peptide. These data demonstrate that AP neurons are directly influenced by ORX-A and suggest that ORX-A
may exert its effects on the central control of feeding behavior and
cardiovascular function through direct actions in AP.
Key words:
area postrema; orexin-A; patch-clamp; nonselective
cationic conductance; electrophysiology; central control of feeding
behavior; cardiovascular function
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INTRODUCTION |
Orexins (ORX-A and -B)/hypocretins
(hypocretin-1 and -2) are novel neuropeptides discovered in 1998 (de
Lecea et al., 1998 ; Sakurai et al., 1998) that include two separate
peptides proteolytically derived from the same precursor protein
(Sakurai et al., 1998). Orexin-producing neurons are exclusively
distributed within and around the lateral hypothalamic area (LHA), the
dorsomedial hypothalamic nucleus (DMH), and the perifornical nucleus in
rats (de Lecea et al., 1998), areas which have been implicated
previously in the control of mammalian feeding behavior (Marshall and
Teitelbaum, 1974; Oltmans and Harvey, 1976). Central administration of
orexins stimulates feeding (Sakurai et al., 1998) and drinking (Kunii et al., 1999) in rats and mice and affects the behavioral satiety sequence in rats (Rodgers et al., 2000). Therefore, orexins are recognized as potent orexigenic peptides.
Although immunohistochemical studies using anti-ORX antiserum have
shown ORX-immunoreactive (IR) neurons specifically localized within the perifornical nucleus, LHA, DMH, and posterior hypothalamic area (de Lecea et al., 1998), ORX-IR axons and their varicose terminals
show a widespread distribution throughout the entire adult rat brain,
including the cerebral cortex, circumventricular organs (CVOs) [the
subfornical organ and the area postrema (AP)], limbic system, and
brain stem (Peyron et al., 1998). These results indicate that
the ORX system provides a link between the hypothalamus and other brain
regions and plays important roles in integrating the complex physiology
underlying feeding behavior and other autonomic functions. In addition
to effects on feeding, centrally administered orexins have also been
demonstrated to have sympathetic and cardiovascular actions (Shirasaka
et al., 1999), play a key role in the regulation of rapid eye movement
sleep and the pathophysiology of narcolepsy (Reilly, 1999; Bourgin et
al., 2000), and activate the hypothalamo-pituitary-adrenal axis (Kuru
et al., 2000). Therefore, orexin neurons project throughout the entire
CNS to nuclei known to be important in the control of feeding,
sleep-wakefulness, neuroendocrine homeostasis, and autonomic
regulation. In addition, genetic ablation of orexin neurons in mice
resulted in narcolepsy, hypophagia, and obesity (Hara et al.,
2001).
The area postrema is a midline structure located on the floor of the
caudal end of the fourth ventricle and is one of the most highly
vascularized regions in the brain (Wislocki and Putnam, 1920). In
addition, this circumventricular organ lacks the typical blood-brain
barrier that is normally impermeable to most blood-borne peptides
(Leslie, 1986). In addition to the description of AP as a
chemosensitive trigger zone (for review, see Borison, 1974), this
structure has also been shown to regulate central autonomic control
(Ferrario et al., 1979; Ferguson and Marcus, 1988), body fluid balance,
and feeding (Edwards and Ritter, 1981; Hyde and Miselis, 1983).
These studies combined with reports of immunoreactive ORX in the AP
suggest this CVO as a potential CNS site, where ORX-A acts to influence
central autonomic control. The present study uses whole-cell,
patch-clamp techniques to examinewhether ORX-A directly affects AP
neurons and to elucidate the membrane events underlying such actions.
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MATERIALS AND METHODS |
Cell dissociation. Cell dissociation and culture
methods were modified from those of Ferguson et al. (1997). In brief,
adult (100-150 gm) male Sprague Dawley (Charles River, Quebec, Canada) rats were decapitated, and the brains were rapidly removed and immersed
in ice-cold Hanks' buffer (nominally
Ca2+- and
Mg2+-free). Under a dissecting microscope,
the AP is visually identified (superficial location at the level of the
fourth ventricle, its v-shaped appearance, and its distinguishing
shallow orange color), cut away from the surrounding tissue, placed in
1.5 ml of the same solution containing 1 mg/ml trypsin (Sigma, St.
Louis, MO), and warmed to 37°C. After incubation (in 5%
CO2/95% O2 at 37°C) and
periodic gentle shaking at 5 min intervals for 15-20 min, the AP was
gently triturated through a tuberculin syringe fitted with a 20 gauge
needle. The cell suspension was transferred to a Hanks' solution
containing Ca2+ (1.3 mM), Mg2+ (0.9 mM), and 0.1% bovine serum albumin (BSA) (Sigma
type A-6003, essentially fatty acid-free) at room temperature
(22°C-24°C). After centrifugation, the pellet was resuspended in
this same solution and recentrifuged. The resultant pellet was again
resuspended in the BSA-containing solution, plated onto plastic Petri
dishes, and placed within a 5% CO2/95%
O2 environment at 37°C until the cells attached
to the dish (~1 hr). The Petri dishes were then further filled with
Neurobasal-A media (Invitrogen Canada, Inc., Ontario, Canada), which
contained antibiotics (100 U/ml penicillin/streptomycin; Invitrogen
Canada, Inc.) and was additionally supplemented with 0.5 mM L-glutamine (Invitrogen
Canada, Inc.). The cells were maintained in the
CO2 incubator at 37°C until use (12 hr to
3 d after dissociation). All procedures conformed to the standards
outlined by the Canadian Council on Animal Care and the Queen's
University Animal Care Committee.
Experimental solutions. For current-clamp recording and
measurement of the nonselective cation current as well as the
K+ conductances during voltage-clamp
recordings, the pipette solution contained (in
mM): 130 potassium-gluconate, 10 EGTA, 1 MgCl2, 10 HEPES, 4 Na2ATP,
and 0.1 GTP, pH adjusted to 7.2 with KOH. Tetrodotoxin (TTX) (0.1 µM) was added to the external solutions to
block voltage-gated sodium channels. The control bath solution consisted of artificial CSF (aCSF) of the following composition (in mM): 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES,
and 10 glucose, pH adjusted to 7.4 with NaOH. All chemicals, unless
otherwise stated, were obtained from Sigma.
Electrophysiological methods. Whole-cell, patch-clamp
recordings were obtained from cells defined as neurons by the presence of  90 mV action potentials in response to a depolarizing pulse (current-clamp recordings) or by the presence of large rapid
voltage-activated inward currents that were blocked by TTX
(voltage-clamp recordings). Electrodes/micropipettes of 3-5 M were
pulled from TW150 glass (World Precision Instruments, Sarasota,
FL) on a horizontal Flaming/Brown micropipette puller (model
P-87; Sutter Instrument Co., Novato, CA), fire-polished, and filled
with the appropriate filling solution. Signals were amplified,
collected, and processed using an Axopatch 200B (Axon Instruments,
Foster City, CA) amplifier and a 1401plus analog-to-digital
interface and Signal and Spike2 software packages from Cambridge
Electronic Design (Cambridge, UK). The recording chamber was the
35 mm plastic Petri dish in which the cells had been cultured, and
solution changes were made by a gravity-fed perfusion system. This flow
was adjusted to ~3 ml/min and was maintained constant throughout the
entire recording period.
Definition of response. The membrane potential and firing
frequency (20 sec bins) of cells were measured for 1 min before and for
 2 min after change in bath solutions, with ORX-A being applied to the
recording dishes via perfusion lines. Membrane potential changes were
assessed by evaluating changes expressed in relation to the average of
the 60 sec control period for at least three consecutive 20 sec time
periods after experimental treatments.
Statistical analysis. For statistical analysis of the
effects of ORX-A on AP neuronal properties, means were calculated from cells that were determined to have been affected using the above criteria. Changes in input resistance and peak and steady-state conductance in response to ORX-A were compared using the Student's t test. All values are plotted as means ± SEM. The
concentration-response curve was constructed from a sigmoidal function
of nonlinear regression (Prism; GraphPad Software Inc., San Diego, CA).
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RESULTS |
The distinct morphology of the AP neurons after 1-3 d in culture,
including phase-bright, 6- to 8-µm-diameter soma and sometimes 2-5
µm long thin processes, are readily distinguished from the flat,
large glial cells. In addition, AP neurons can also be distinguished from glial cells electrophysiologically (action potentials and sodium
currents) (Ferguson et al., 1997). A total of 108 AP neurons were
recorded with the whole-cell, patch-clamp technique. The mean resting
membrane potential of neurons recorded in current clamp
(n = 48) was 59.24 ± 0.87 mV, the mean input
resistance was 3.57 ± 0.22 G, and the action potential
amplitude of neurons included in this study was always >90 mV. The
frequency of spontaneous action potentials in these AP neurons ranged
from 0.2 to 9.9 Hz with a population average rate of discharge of
2.9 ± 1 Hz. Initial analysis of all current-clamp recordings
showed that responsiveness of cells could be split into three
distinctive subgroups according to the observed changes in membrane
potential (depolarization, 67%; hyperpolarization, 13%; no effect,
20%) in response to ORX-A (108
M) (Fig. 1). The
inset in Figure 1 shows that 108,
109, and
1010 M ORX-A
depolarized 68.3%, hyperpolarized 17.1%, and was without effect on
14.6% of AP neurons (n = 41). We could not identify any difference in the electrophysiological properties of these neurons,
which corresponded with their responsiveness to ORX-A. In accordance
with these observations, a neuron was thus deemed to be responsive if
membrane potential changed by >5 mV after any particular experimental
manipulation.
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Figure 1.
AP neurons respond to ORX-A application. AP
neurons can be divided into three groups in accordance with their
responses to 108 M ORX-A exposure
(depolarized, n = 10; hyperpolarized,
n = 2; with no effect, n = 3).
Inset, Pie chart showing that
108, 109, and
1010 M ORX-A depolarized 68.3%,
hyperpolarized 17.1%, and were without effect on 14.6% of AP neurons
(n = 41).
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ORX-A depolarizes AP neurons
Based on the criteria established (see Materials and Methods), 41 of 48 cells (85.4%) were classified as being responsive to changes in
extracellular ORX-A (Responders), whereas the remainder of the cells
(seven cells; 14.6%) did not respond in a sustained manner and were
therefore classified as Nonresponders. The vast majority of responsive
cells (80.5%; 33 of 41) demonstrated clear reversible depolarizations
in response to bath application of ORX-A, the magnitude of which was
concentration dependent, as illustrated in Figure
2B. During exposure of
a neuron to 108 M
ORX-A from a control bath aCSF (Fig. 2A, top
panel), cells exhibited rapid sustained depolarizations
accompanied in most cases by a rapid increase in firing frequency of
action potentials. After washout of ORX-A and replacement of the bath
solution with control aCSF, the membrane potential and action potential
frequency recovered to the control level. In 10 cells exposed to this
same concentration of ORX-A, the mean amplitude of depolarization was 9.96 ± 0.52 mV. Similar depolarizing responses of AP neurons to the exposure to 109 and
1010 M ORX-A are
shown in Figure 2A. These depolarizing effects were found to be concentration dependent at concentrations ranging from
108 to
1011 M, as
illustrated by the concentration response curve presented in Figure
2B (EC50 = 1.76 × 1010 M). Although
the depolarizing response of AP neurons to exposure to
1011 M ORX-A did
not reach the criteria established (see Materials and Methods) to be
defined as responders, all cells tested with this concentration were
included in the analysis to complete the concentration-response
curve.
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Figure 2.
ORX-A depolarizes AP neurons. A,
Bath application of various concentrations (108 to
1011 M) of ORX-A
depolarized 68.8% (33 of 48) of the AP neurons tested. During exposure
to ORX-A from a control bath aCSF, AP neurons exhibited rapid sustained
depolarizations accompanied in most cases by a rapid increase in firing
frequency of action potentials. After washout of ORX-A and replacement
of the bath solution with control aCSF, the membrane potential and
action potential frequency returned to the control level. ORX-A
application is represented by the horizontal bar above
each trace. Calibration: 60 sec, 10 mV. The dashed
line indicates the baseline of potentials. B,
Depolarization of AP neurons by ORX-A is concentration dependent.
Changes in membrane potential measured during responses to
1011 (n = 5),
1010 (n = 14),
109 (n = 4), and
108 (n = 10) M
extracellular ORX-A were plotted against bath ORX-A concentrations.
Data change presented as mean ± SEM data was fitted to a sigmoid
concentration-response function, and the corresponding curve was
overlaid; EC50 = 1.76 × 1010 M.
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ORX-A does not influence net whole-cell potassium currents
The modulation of voltage-gated potassium conductances has been
shown to be important in the regulation of neuronal excitability. Orexin-B has been reported to decrease potassium conductance (Ivanov and Aston-Jones, 2000) or reduce afterhyperpolarization (Horvath et
al., 1999) in locus ceruleus neurons. Therefore, we used voltage-clamp techniques to examine the effects of ORX-A on net whole-cell potassium currents evoked by depolarizing voltage steps. Figure
3A illustrates the whole-cell
currents recorded in response to depolarizing voltage steps applied
from a holding potential of 100 mV in 20 mV increments before and
after bath application of 108
M ORX-A, showing no clear effect of ORX-A on area
postrema on either the peak or sustained current. In all of the neurons
tested (n = 20 of 20), application of ORX-A
(108 M) did not
affect K+ conductances of area postrema
neurons at every voltage level, as illustrated in the group data
presented in Figure 3B.
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Figure 3.
ORX-A does not influence net whole-cell potassium
currents. A, Whole-cell net K+
currents isolated from area postrema neurons (with 0.1 µM
TTX in aCSF) evoked by 20 mV voltage steps (5 sec) from 100 mV
holding potential to +40 mV were unaffected by ORX-A exposure.
B, Summary data of K+ conductances
recorded before (solid) and after (open)
extracellular application of ORX-A (108
M) were not different at either the peak
(squares) or the sustained currents
(triangles).
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ORX-A activates nonselective cationic conductances of
AP neurons
In view of the considerable literature demonstrating peptidergic
effects on neuronal excitability mediated by modulation of nonselective
cationic conductances (NSCCs), we subsequently used slow voltage ramps
[100-0 mV (10 sec) after a prepulse of 100 mV (500 msec)] to
determine whether ORX-A influenced AP neurons as a consequence of
activation of such conductances. The data presented in the top
panel of Figure 4A
show average currents recorded from an AP neuron in response to such
ramps (each trace is the mean of five ramps) recorded before, during,
and after bath administration of ORX-A
(108 M). The
bottom panel of Figure 4A shows the
difference current (i.e., ORX-A-induced current) obtained by
subtracting control ramps from those obtained during ORX-A. Application
of ORX-A (108 M)
activated the NSCCs, and 5 min after washout of ORX-A and replacement
of the bath solution with aCSF, the NSCCs recovered toward control
levels. Interestingly, as illustrated by the difference current shown
in Figure 3A, we often saw increased noise in the depolarized range, which may be indicative of effects of ORX-A on
additional, as yet unidentified channels. Similar effects of ORX-A
(108 M) were
observed in 14 of 20 (70%) cells tested, a proportion that closely
matches the proportion of AP neurons identified as being depolarized by
ORX-A in our current-clamp experiments. Mean ORX-A-evoked currents for
this group of responsive neurons are presented in Figure
4B, with these observations suggesting a reversal potential of 43.57 ± 5.43 mV (n = 14).
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Figure 4.
ORX-A activates nonselective cationic conductance
in AP neurons. A, Top panel, Mean
whole-cell currents (each trace is the mean of 5 ramps) evoked from
slow depolarizing (10 mV/sec) voltage ramps before, during, and after
exposure to ORX-A (108 M).
Bottom panel, Difference current obtained by subtracting
the control current from the current recorded during ORX-A application.
This represents the ORX-A-evoked current. B, Mean
ORX-A-evoked current for responsive neurons (n = 14) and its 95% confidence interval.
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DISCUSSION |
Several lines of evidence have shown that ORX acts in the CNS to
modulate feeding, sleep-wakefulness, neuroendocrine homeostasis, and
autonomic regulation. The distribution of ORX-IR axons and their
varicose terminals in area postrema, combined with the established involvement of ORX-A in the central modulation of feeding (Sakurai et
al., 1998) and cardiovascular function (Shirasaka et al., 1999), suggest that the area postrema represents a significant site for potential neuroregulatory actions of this peptide. The data from this
study are the first to demonstrate direct effects of ORX-A on area
postrema neurons and, in addition, represent the first report of direct
effects of this peptide on nonselective cationic conductances.
A significant issue in all studies using either acutely dissociated or
cultured neurons is the degree to which the properties of these neurons
match those of the cell in vivo. We have developed a
methodology in these studies in which dissociated AP neurons are
prepared from adult as opposed to neonatal tissue (Hay and Lindsley,
1995; Hay et al., 1996), which alleviates the likelihood of
developmental changes influencing recorded properties. Our ability to
completely isolate AP from all surrounding tissue also ensures that the
cells we have recorded were derived exclusively from the AP. These
dissociated AP neurons also show properties similar to those reported
by other groups (Hay and Lindsley, 1995), as well as for AP neurons in
tissue slice preparations (Jahn et al., 1996), and demonstrate patterns
of spontaneous activity similar to those reported for AP neurons
in vivo (Papas et al., 1990). Finally, the demonstration of
effects of ORX on these dissociated cells argues strongly that the use
of trypsin in the dissociation procedure has not permanently stripped
these cells of their membrane-located peptide receptors.
Our initial current-clamp studies clearly illustrate the ability of
ORX-A to influence the membrane potential of what appears to be a
subpopulation of AP neurons. The fact that these effects were observed
on dissociated cells indicates that they are the result of direct
actions on the recorded neuron. The present lack of specific ORX
receptor antagonists precluded identification of the specific ORX
receptor mediating these effects. However, the clear reversibility and
concentration dependence of these effects argue strongly that they are
receptor mediated.
Previous studies have reported that ORX-B decreases potassium
conductances (Ivanov and Aston-Jones, 2000) and reduces
afterhyperpolarizations (Horvath et al., 1999) in locus ceruleus
neurons. In view of the accepted role of potassium conductances in
controlling neuronal excitability throughout the brain as well as in
the area postrema (Hay and Lindsley, 1995), we subsequently examined
whether the effects of ORX-A on these neurons might be mediated at
least in part by similar modulatory actions of this peptide on
voltage-gated potassium currents. Surprisingly, our data demonstrated
that ORX-A had no effects on whole-cell, voltage-dependent potassium
currents. In view of the clear lack of effect of the peptide on these
mixed currents, they were not dissected further for the current
analysis, although separate experiments have shown that
IA, IK, and
ID all likely contribute to the total current we
examined. However, it should be emphasized that we have not evaluated
potential effects of ORX on the iberiotoxin-sensitive
Ca2+-activated potassium channel, which
has also been reported in AP neurons (Li and Hay, 2000).
The nonselective cationic conductance is a membrane channel for cations
not specific for Na+,
K+, or Ca2+.
Its reversal potential is unaffected by changes in
Cl concentration and pH, and it is
voltage independent (Kramer and Zucker, 1985). Nonselective cationic
conductances have been shown to participate in controlling neuronal
excitability in many systems, including generation of the depolarizing
phase of bursting pacemaker activity in Aplysia burst-firing
neurons (Kramer and Zucker, 1985), and in the intrinsic activation of
rat supraoptic neurons by hyperosmotic stimuli (Bourque, 1989),
neurotensin (Kirkpatrick and Bourque, 1995), and P2 purinoceptor
agonists (Hiruma and Bourque, 1995). In addition, previous work from
our laboratory demonstrated that activation of the calcium receptor in
neurons of the subfornical organ, a forebrain circumventricular
structure, results in profound effects on neuronal excitability through
metabotropic actions on a nonselective cation channel (Washburn et al.,
1999). The results from the current study illustrate for the first time
the effect of ORX-A on an NSCC in area postrema neurons. Extracellular application of ORX-A activated NSCCs in reversible manner in a proportion of AP neurons similar to that which our current-clamp recordings demonstrated to be depolarized by this peptide. The conductance of this NSCC is 0.497 ± 0.031 nS. We usually held AP
neurons between 52 and 53 mV before ORX-A
(108 M)
administration. At this potential, ORX-A would be expected to activate
the NSCCs as a 2.5-3 pA inward current (Fig. 4B), which we calculate to evoke an 8-11 mV depolarization (average input
resistance of AP neurons is 3.6 G), which is close to the average
depolarization (9.96 ± 0.52 mV) caused by
108 M ORX-A application.
In conclusion, this study provides the first evidence that orexin-A
directly activates area postrema neurons by modulating nonselective
cationic conductance, whereas it does not affect the
K+ channel activities. These findings
suggest that orexin may have a functional role in the regulation of
feeding behavior and cardiovascular and autonomic nervous systems at
the AP level.
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FOOTNOTES |
Received Feb. 11, 2002; revised April 1, 2002; accepted April 11, 2002.
This work was supported by a grant to A.V.F. from the Heart and Stroke
Foundation of Ontario. We thank Dr. J. W. Anderson for helpful
comments on this manuscript.
Correspondence should be addressed to Dr. A. V. Ferguson,
Department of Physiology, Queen's University, Kingston, Ontario, Canada K7L 3N6. E-mail: FergusnA{at}post.queensu.ca.
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ABSTRACT
INTRODUCTION
