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1 Department of Anatomy and Cell Biology, University of Illinois, Chicago, Illinois 60612-7308; 2 Department of Pharmacology, University of Illinois, Chicago, Illinois 60612-7308; 3 Howard Hughes Medical Institute, Department of Molecular Genetics, University of Texas Southwestern Medical Center, Dallas, Texas 75235-9050
Submitted 2 January 2003; accepted in final form 10 April 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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S-loaded cells, orexin A (OXA, 3 µM)
inhibited GIRK currents that had previously been activated by somatostatin (in
LC cells), nociceptin (TM cells), or the mu opioid agonist DAMGO (HEK cells).
In guanosine triphosphate (GTP)–loaded HEK cells, in which GIRK currents
were not preactivated, OXA induced a biphasic response through both types of
orexin receptors: an initial current increase and a subsequent decrease to
below resting levels. Current–voltage (I–V) relationships revealed
that both the OXA-induced and suppressed currents are inwardly rectifying with
reversal potentials around EK. The OXA-induced
initial current was partially pertussis toxin (PTX) sensitive and partially
PTX insensitive, whereas the OXA-suppressed current was PTX insensitive. These
data suggest that orexin receptors couple with more than one type of
G-protein, including PTX-sensitive (such as Gi/o) and
PTX-insensitive (such as Gq/11) G-proteins. The modulation of GIRK
channels by orexins may be one of the cellular mechanisms for the regulation
of brain nuclei (e.g., LC and TM) that are crucial for arousal, sleep, and
appetite. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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;
Sakurai et al. 1998). Their
receptors, orexin receptor type 1 (OX1R) and type 2
(OX2R), are cloned and found to be G-protein coupled
(Sakurai et al. 1998). Further
studies implicate orexins and OX2R in the sleep disorder narcolepsy
(Chemelli et al. 1999;
Lin et al. 1999; Nishino et
al. 2000,
2001;
Thannickal et al. 2000).
Orexin receptors are present in brain nuclei constituting the ascending
arousal system, such as the locus coeruleus (LC) and the nucleus
tuberomammillaris (TM). The LC, the major noradrenergic nucleus of the brain,
contains predominantly OX1R, whereas the TM, the histaminergic
nucleus, contains predominantly OX2R
(Trivedi et al. 1998;
Eriksson et al. 2001;
Greco and Shiromani 2001;
Hervieu et al. 2001;
Marcus et al. 2001;
Yamanaka et al. 2002). Orexins
have been shown to have a direct excitatory effect on LC and TM neurons
(Hagan et al. 1999;
Horvath et al. 1999;
Bourgin et al. 2000;
Ivanov and Aston-Jones 2000;
Bayer et al. 2001;
Eriksson et al. 2001;
Soffin et al. 2002;
van den Pol et al. 2002;
Yamanaka et al. 2002). The
mechanism of orexin-induced excitation has been suggested to involve: a
decrease in K+ conductance
(Ivanov and Aston-Jones 2000),
an induction of TTX-insensitive Na+ inward currents
(van den Pol et al. 2002), and
an activation of the electrogenic Na+/Ca2+
exchanger and a Ca2+ current
(Eriksson et al. 2001).
However, precise ionic and signal transduction mechanisms of orexin effects
have not been fully clarified.
Here, using cultured LC and TM neurons and a reconstituted system (HEK293A cells), we show that when G-protein–coupled inward rectifier (GIRK, Kir3) channels are previously activated by inhibitory transmitters, OXA suppresses GIRK channel activity, which likely leads to neuronal excitation. We also show that when applied to a reconstituted system in which GIRK currents are not activated, OXA induces a biphasic response through both receptor types: an initial, partially pertussis toxin (PTX) sensitive increase and a subsequent, PTX-insensitive decrease in GIRK channel activity to below resting levels.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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Locus coeruleus neurons were cultured from 3- to 4-day-old and 10-day-old
postnatal Long-Evans rats (Charles River Laboratories, Wilmington, MA) by
using the procedures reported previously
(Masuko et al. 1986;
Nakajima and Masuko 1996).
Tuberomammillary neurons were similarly cultured
(Bajic et al. 2003) from 3- to
4-day-old rats. The rats were anesthetized with ether. After the rats became
completely unconscious (coma), the scalp and skull were removed exposing the
brains. Brain stems and hypothalamic regions were removed. Immediately
afterward, the animals were decapitated to ensure euthanasia. The removed
brain regions were embedded in agar and sectioned into 400-µm-thick slices
with a Vibratome. The LC and TM were isolated from the slices under a
dissecting microscope. The nuclei were then dissociated with papain (12 U/ml),
plated on a glial feeder layer, and incubated at 37°C with 10%
CO2 in medium consisting of minimum essential medium with Earle's
salt (88%; Gibco BRL, Gaithersburg, MD) modified by adding
L-glutamine (0.292 mg/ml, final concentration), NaHCO3
(3.7 mg/ml), D-glucose (5 mg/ml), L-ascorbic acid (10
µg/ml), penicillin (50 U/ml), streptomycin (50 µg/ml), and
heat-inactivated rat serum (2%, prepared in our laboratory). Conditioned
medium (Baughman et al. 1991)
was used throughout the culture. Cultures from 3- to 4-day-old rats and those
from 10-day-old rats were maintained for 50–92 days and for 28 days,
respectively. Experiments were performed on large neurons, likely to be
noradrenergic LC neurons (Masuko et al.
1986) or histaminergic TM neurons
(Bajic et al. 2003). The
average diameter was 34.5 ± 6.9 µm (mean ± SD) among chosen
LC neurons, and 24.7 ± 2.0 µm among TM neurons. All cells used had
resting potentials more negative than –59 mV in 10 mM K+
Krebs solution.
Cell Line Culture and Transfection
Human embryonic kidney 293A cells (HEK293A, Qbiogene, Carlsbad, CA) were maintained in DMEM supplemented with 10% FBS, penicillin (100 U/ml), and streptomycin (100 µg/ml). One day before transfection, 6-cm culture dishes were plated with 2.5 x 105 HEK293A cells/dish. Using Effectene Transfection Reagent (Qiagen, Chatsworth, CA), cells were transfected, for most experiments, with: GIRK1 and GIRK2 (0.15 µg cDNA each), human mu opioid receptor (MOR, 0.5 µg), either OX1R or OX2R (0.5 µg, human receptors), and green fluorescent protein (GFP, 0.05 µg). One day after transfection, cells were replated onto 3.5-cm dishes coated with rat tail collagen (Boehringer Mannheim Biochemicals, Indianapolis, IN). Electrophysiological experiments were performed 72–84 h after transfection on isolated cells displaying strong GFP fluorescence and resting potentials more negative than –59 mV.
For the dose–response experiments, either OX1R or
OX2R (0.5 µg) was transfected along with G
and G
subunits (0.4 µg each), GIRK1 and GIRK2 (0.15 µg each), and GFP (0.05
µg).
For the pertussis toxin experiments, either OX1R, OX2R, or MOR (0.5 µg) was transfected along with GIRK1 and GIRK2 (0.15 µg each) and GFP (0.05 µg). The total amount of cDNA was kept constant by adding empty expression vectors. Ninety-one to 93 h after transfection, cultures were treated with 250 ng/ml of PTX (List Biological Laboratories, Campbell, CA) or heat-inactivated PTX (30 min at 100°C) and incubated for 9 to 20 h. PTX was freshly dissolved in 0.04% BSA (Calbiochem Biosciences, La Jolla, CA) for each experiment.
Electrophysiology
The main experiments were done using the whole cell version of patch clamp
(voltage clamp). The external solution contained (in mM) 141 NaCl, 10 KCl, 2.4
CaCl2, 1.3 MgCl2, 11 D-glucose, 0.0005
tetrodotoxin (TTX), and 5 HEPES-NaOH (pH 7.4). The patch pipette solution
contained (in mM): 141 K D-gluconate, 10 NaCl, 5 HEPES-KOH, 0.5
EGTA-KOH, 0.1 CaCl2, 4 MgCl2, 3 Na2ATP, and
0.2 GTP (pH 7.2). In guanosine 5'-[-thio]triphosphate
(GTPS) experiments (Figs.
2 and
3), GTP was replaced with 0.2
or 0.3 mM GTPS. The holding potential was –84 mV.
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For the action potential recordings (Fig. 1), the external solution contained 5 mM K+ with no TTX (the osmolarity was adjusted by increasing Na+).
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Data were analyzed with pCLAMP programs (version 6, Axon Instruments, Burlingame, CA). Membrane potential values were corrected for the liquid junctional potential between the bath and the patch pipette solutions. Drugs were applied either through a thoroughly washed glass capillary by pressure ejection or through a sewer pipe system (ALA Scientific Instruments, Westbury, NY). Unless otherwise specified, the following concentration of drugs were used: OXA (3 µM, Peptides International), somatostatin (0.3 µM, Peptides International, Louisville, KY), nociceptin (1 µM, Peptides International), and DAMGO (3 µM, Bachem, Torrance, CA). Experiments were performed at room temperature.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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To examine whether OXA actually depolarizes and excites our cultured brain
neurons, current clamp recordings were performed on LC neurons. With the
membrane potential near the resting potential, application of a low
concentration of somatostatin (SOM, 0.03 µM) elicited a hyperpolarization
(Fig. 1, A1 and
B1). When the potential reached about –77 mV, the
application of OXA (3 µM) (together with SOM) was started. This produced a
depolarization (Fig. 1, A1 and
B1). As reported by
Hagan et al. in 1999, at or
near the resting potential of LC cells in slice preparations, there existed a
spontaneous action potential firing activity. The membrane potential was
therefore depolarized to about the level of threshold where low frequency
action potential firing occurred (Fig. 1,
A2 and B2). Application of SOM elicited a
hyperpolarization, which prevented the firing of action potentials
(Fig. 1, A2 and
B2). When the potential reached about –59 mV, the
application of OXA (together with SOM) was started. This produced a
depolarization, eventually eliciting the firing of action potentials once
again. The firing frequency was now higher than that before the SOM
application. This experiment demonstrates that OXA indeed produces
depolarization and excitation of our primary cultured brain neurons. Therefore
the main theme of the present experiments is to elucidate the ionic mechanisms
of this OXA-induced change in excitability by focusing on the roles of GIRK
channel activity.
In primary cultured neurons, OXA suppressed a GIRK current
LC neurons are known to be rich in OX1R and TM neurons rich in
OX2R (Eriksson et al.
2001; Greco and Shiromani
2001; Hervieu et al.
2001; Marcus et al.
2001; Trivedi et al.
1998; Yamanaka et al.
2002). When OXA (3 µM) was applied to cultured LC neurons under
voltage clamp, OXA had little or no effect on membrane conductance (n
= 8). When applied to TM neurons, a decrease in whole cell conductance was
seen in only one out of the 3 recorded cells. We suspected that this was
because the basal activity of GIRK channels is very low under these
conditions. We therefore first activated GIRK currents with inhibitory
transmitters, SOM for LC neurons and nociceptin (NOCI) for TM neurons,
followed by the application of OXA. It is known that GIRK currents are
activated by SOM in LC neurons (Inoue et
al. 1988; Velimirovic et al.
1995) and by NOCI in TM neurons
(Eriksson et al. 2000). In
these experiments neurons were loaded with GTPS, a nonhydrolyzable GTP
analogue, to maintain G-protein–mediated inhibitory and excitatory
transmitter effects, avoiding complications from receptor desensitization. SOM
or NOCI was applied 2 to 4 min after rupturing the patch.
As shown in Fig. 2A1, application of SOM (0.3 µM) to LC neurons increased the membrane conductance, which was suppressed by the subsequent application of OXA (3 µM, n = 10). Similarly, in Fig. 2B1, application of NOCI (1 µM) to TM neurons increased the membrane conductance, which was suppressed by the subsequent application of OXA (n = 5). We also observed that a second application of SOM in LC neurons (n = 6) or NOCI in TM neurons (n = 4) resulted in no conductance increase.
In Fig. 2A2 we measured the current–voltage (I–V) relationship of an LC neuron while the conductance was enhanced by SOM (solid circles, solid line). This was followed by the measurement of the I–V relation after the conductance was declined by the application of OXA (open circles, dotted line). The comparison of the two curves indicates that the resting potential (potential at zero current) shifted from –62 mV (solid arrow) to –57 mV (open arrow) with a net depolarization of about 5 mV induced by OXA. Differences of the two curves in Fig. 2A2 represent the portion of the conductance that was affected by the OXA application (i.e., the "OXA-suppressed current") and are presented in Fig. 2A3. The I–V relations of the OXA-suppressed currents (Fig. 2, A3 and B2) exhibited an inward rectification with a reversal potential near the K+ equilibrium potential (EK, –71 mV). The mean reversal potential for LC neurons and that for TM neurons were –69 ± 2 mV (mean ± SEM, n = 4) and 70 ± 2 mV (n = 4), respectively.
In conclusion, the results in Fig. 2 indicate that OXA suppresses GIRK currents that were previously activated, and that the GIRK current inhibition by OXA was, under the present conditions, stronger than the GIRK current activation by SOM or NOCI.
In a heterologous system, OXA suppressed a GIRK current that had been enhanced by the mu opioid receptor
Neurons generally express two types of orexin receptors, although some
neurons such as LC and TM neurons predominantly, but not exclusively, express
one receptor type. To investigate the OX1R and OX2R in
isolation from each other, we employed a heterologous system and reconstituted
orexin effects by transfecting HEK293A cells with either
OX1RorOX2R cDNA along with mu opioid receptor (MOR),
GIRK1, GIRK2, and GFP cDNAs. We chose these specific GIRK channel subunits
because our single-cell RT-PCR study revealed that LC neurons contain
predominantly GIRK1 and GIRK2 mRNAs
(Kawano et al. 2002). In
transfected HEK cells, GIRK currents were induced by DAMGO
([D-Ala2, N-Me-Phe4,
Gly-ol5]-enkephalin), a specific MOR agonist
(Handa et al. 1981).
Using this heterologous system, we investigated the interaction of DAMGO (3
µM) and OXA (3 µM) effects in cells loaded with GTPS (0.2 or 0.3
mM). DAMGO was applied 2 to 4.5 min after rupturing the patch. In both
OX1R-expressing (Fig.
3A) and OX2R-expressing cells
(Fig. 3B), DAMGO
application induced an inward current accompanied by a conductance increase,
which was suppressed by a subsequent application of OXA. The time course of
the OXA-induced inhibition was slow in both OX1R-expressing cells
[with a half time (t0.5) of 42.9 ± 3.0 s; n = 10]
and OX2R-expressing cells (t0.5 = 31.0 ± 7.5 s;
n = 5). After the conductance had been suppressed by OXA, a second
DAMGO application could not increase the K+ conductance, indicating
that suppression of GIRK channels by OXA is stronger than their activation by
DAMGO. These results obtained in the reconstituted system are essentially the
same as those obtained in LC and TM neurons.
In Fig. 3C, the conductance enhanced by DAMGO (GDAMGO) was compared with the conductance suppressed by the subsequent OXA application (GOXA). GDAMGO and GOXA are expressed relative to the control conductance measured before the drug application (G1; diagrammed in Fig. 3D). The figure shows that the larger the DAMGO-induced conductance, the larger the OXA-induced conductance suppression, strongly suggesting that OXA suppressed the same channels (GIRK channels) as those activated by DAMGO.
OXA effects in HEK293A cells loaded with GTP
In the preceding experiments, GIRK currents were maintained by loading
cells with GTPS. Because the cytoplasm of living cells contains GTP, we
investigated the interaction between DAMGO and OXA effects in cells loaded
with GTP. In OX1R-expressing cells, application of DAMGO induced a
conductance increase (Fig.
4A1). About 7 min later, a second DAMGO application
(A2) was immediately followed by the application of OXA and DAMGO
together to test the OXA effect during the DAMGO-induced current. This
procedure resulted in the OXA-induced suppression of the DAMGO-activated
current. About 10 min later, a third DAMGO application induced a conductance
increase once again (A3), this time with a smaller amplitude than
that in the first application, partly reflecting receptor desensitization. The
I–V relations in Fig. 4, A4
and A5, show that the DAMGO-induced current and the
OXA-suppressed current were inwardly rectifying with reversal potentials
approximately coinciding with EK, indicating that
both are GIRK currents. Similar OXA effects were also observed in cells
expressing OX2R (Fig.
4B). These results demonstrate that in GTP-loaded cells,
OXA also suppresses GIRK currents that were previously enhanced by MOR
activation in both OX1R-expressing (n = 5) and
OX2R-expressing cells (n = 4).
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Concentration–response relationships for GIRK inhibition induced by OXA in HEK293A cells expressing OX1R or OX2R
By using the HEK293A cell reconstituted system, we determined
concentration–response relationships for GIRK inhibition by OXA. HEK293A
cells were transfected with either OX1R or OX2R cDNA
along with G, G, GIRK1, GIRK2, and GFP cDNAs. In these cells,
GIRK currents were robustly activated by overexpressing G and G.
In Fig. 5 the magnitude of the
conductance decrease was represented as a percent, relative to the resting
conductance before OXA application. Data were fitted with the equation
Axp/(K'p + x
p), where x is the OXA concentration, A is
the maximal value of OXA-induced inhibition, K' is the
half-effective concentration (EC50), and p is Hill's
coefficient. The EC50 was 0.41 µM and 0.27 µM in
OX1R- and OX2R-expressing cells, respectively. The
experiments also indicated that 1 µM OXA is a nearly saturating
concentration for both receptor types.
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Sakurai et al. (1998)
reported that OXA has a slightly higher affinity to OX1R
(IC50 = 20 nM) than to OX2R (IC50 = 38 nM) in
a ligand binding assay, whereas OXA has almost the same efficacy to both types
of OX receptors in a [Ca2+] transient assay
(EC50 = 30 nM in OX1R and EC50 = 34 nM in
OX2R). Thus the OXA effect on GIRK channels appears to have a lower
affinity than that of the binding assay or the [Ca2+]
transient assay.
PTX sensitivity of OXA effects on GIRK currents that were not previously enhanced
When OXA alone was applied to GTP-loaded HEK293A cells, in which G
and G cDNA were not transfected, it unexpectedly induced a
biphasic response consisting of an initial increase and a
subsequent decrease in the membrane conductance to below resting
levels (Fig. 6A1). PTX
pretreatment was used to determine the type(s) of G-protein responsible for
these conductance changes. HEK293A cells were transfected with cDNA of either
OX1R or OX2R together with cDNAs of GIRK1, GIRK2, and
GFP. Ninety-one to 93 h after transfection, cells were treated with either PTX
or heat-inactivated PTX (control) for 11–20 h. Whole cell recordings
were then performed.
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Figure 6A1 shows that application of OXA to control cells expressing OX1R induced a conductance increase, followed by a long-lasting decrease. Both the initial OXA-induced conductance increase and the subsequent OXA-induced conductance decrease were inwardly rectifying with reversal potentials around EK, suggesting that both the OXA-induced and -suppressed currents are GIRK currents (Fig. 6, A2 and A3). In PTX-treated cells expressing OX1R, the application of OXA resulted in a smaller increase in conductance compared with that observed in the control experiments (Fig. 6B1). This was followed by a decrease in conductance, similar in magnitude to control values (Fig. 6B1). I–V relations revealed that the PTX-insensitive, OXA-induced current (Fig. 6B2) and the OXA-suppressed current (Fig. 6B3) are both inwardly rectifying with reversal potentials around EK, indicating that they are GIRK currents. Similar effects of OXA were observed in OX2R-expressing cells (figures not shown).
The summary figure (Fig. 6C) shows that in both receptor types, there was a significant difference between the OXA-induced conductance increase in control cells (treated with heat-inactivated PTX) and that found in PTX-treated cells, suggesting that OXA induces conductance increase through a mechanism that is partially PTX insensitive and partially PTX sensitive. In contrast, there was no significant difference in the OXA-induced conductance decrease between control cells and PTX-treated cells in either receptor type (Fig. 6D), suggesting that OXA suppresses conductance in both OX1R- and OX2R-expressing cells through a PTX-insensitive mechanism.
As a positive control for PTX activity, we used HEK293A cells expressing MOR and GIRK channels. As shown in Fig. 7, the same PTX treatment completely prevented DAMGO from activating the GIRK current, indicating that the PTX used was potent.
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In summary, in both OX1R- and OX2R-expressing cells, the OXA-induced conductance increase was partially inhibited by PTX treatment, whereas the OXA-suppressed conductance was unaffected. These results suggest that both OX1R and OX2R couple with more than one type of G-protein, involving both PTX-sensitive (such as Gi/o) and PTX-insensitive (such as Gq/11) G-proteins. Interestingly, the PTX sensitivity is only partial for the OXA-induced enhancement of GIRK currents.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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Our main observation was that orexin inhibited the activity of GIRK channels. This was observed in both a heterologous system (HEK cells) and primary cultured neurons from the LC and TM. We used brain neurons from the LC and TM that had been cultured for an extended period of time (50–92 days). The density of neurons in these cultures was low, resulting in minimal synaptic interactions among neurons. Is our observation on these neurons relevant to the regulation of neuronal excitability in the in situ brain? In the following discussion, we suggest possible physiological roles played by the orexin-induced GIRK inhibition.
Unlike neurons cultured for a extended period of time, which are rather
isolated from each other, brain neurons in situ are usually packed densely,
and therefore would be influenced almost constantly by slow excitatory or
inhibitory transmitters (such as substance P, or enkephalins) arriving from
neighboring neurons. The balance between these two influences would be an
important determinant for neuronal excitability. Many of the slow inhibitory
transmitters are known to produce their effect by activating GIRK channels in
the brain (Stanfield et al.
2002) and, consequently, the neutralization of GIRK channels (by
orexins) would result in an increase in excitability. Aghajanian and
colleagues (1977) observed
excellent examples of this balancing mechanism in LC and mesenkephalic
dopaminergic neurons. Here, neurons are constantly producing trains of action
potentials. The spike frequency is substantially determined by
"auto-inhibition," derived from the inhibitory transmitters
released from the dendrites of the same or neighboring neurons. The released
transmitter constantly activates GIRK channels through
2-adrenoceptors (LC) or D2-receptors (substantia
nigra). Therefore excitability would be determined by the balance between GIRK
activity (auto-inhibition) and the spontaneous excitatory influences on the
neuron (Aghajanian et al. 1977;
Bunney et al. 1973;
Cheramy et al. 1981;
Kim et al. 1995). Thus the
application of an -adrenergic antagonist (2-blocker)
to LC neurons (Aghajanian et al.
1977) or the D2-antagonist chlorpromazine to
dopaminergic neurons (Bunney et al.
1973) is sufficient to excite these neurons, strongly suggesting
that the elimination of GIRK activity induces neuronal excitation.
Evidence supporting the role of GIRK channels in modulating neuronal excitability can be seen in Fig. 2A2. The plots of the I–V relations before (solid line) and after OXA application (dotted line) reveal that by closing GIRK channels in an LC neuron, OXA depolarized the resting potential by about 5 mV. Additionally, the experiment in Fig. 1 suggests that OXA might have produced depolarization partly by suppressing SOM-induced GIRK current, although other mechanisms would also play various roles over the voltage range in Fig. 1.
Further support of the role of GIRK channels in setting excitability comes
from Torrecilla et al. (2002).
Using knockout mice, the authors showed that in the mouse LC, GIRK channels
may contribute 
20 mV of the resting potential.
Our focus in this study was on the GIRK channel. It should be stressed,
however, that we do not intend to conclude that GIRK inhibition is the only
mechanism underlying orexin's excitatory effects. Previously Eriksson et al.
(2001) showed that orexins
activate the electrogenic Na+/Ca2+ exchanger
and a Ca2+ current. Van den Pol et al.
(2002) observed that orexins
induce TTX-insensitive Na+ inward currents. Additionally, Ivanov
and Aston-Jones (2002) observed a decrease in K+ conductance by
orexins; this K+ conductance may involve K+ channels
other than GIRK. We conclude that orexin-induced excitation is a complex
process, involving more mechanisms than solely the closing of GIRK
channels.
Signal transduction mechanism of orexin suppression of GIRK channels
At present not much is known about the signal transduction mechanism by which orexins suppress GIRK channels that were previously activated by inhibitory transmitters. This OXA effect can be mediated by either OX1R or OX2R receptors. We have also shown that this signaling involves a PTX-insensitive G-protein such as Gq/11.
The concentration–response relation for the GIRK inhibition showed
EC50 was 410 nM for OX1R and 270 nM for OX2R.
These values are higher than that obtained from the orexin binding to the
receptors (20–38 nM) (Sakurai et al.
1998). In the present experiment, because of the relatively short
application duration, OXA could have been washed out before its inhibitory
effect had reached maximum. With a longer exposure time, the inhibition at
each dose could be stronger, and this would be particularly important at low
doses. Therefore it is safe to conclude that 1 µM is a nearly saturating
concentration for both receptor types, although our EC50 values
could have been overestimates. Still, it is possible that the overall
transduction of the GIRK channel inhibition by orexins may take place with a
lower affinity than the binding of the agonist to the receptors.
Interestingly, light microscopic immunocytochemical studies on LC and TM
neurons reveal that these neurons receive abundant innervations of
orexin-containing nerve fibers (Chemelli et
al. 1999; Horvath et al.
1999). Additionally, an electron microscopic study reveals the
abundant presence of excitatory (asymmetric) synaptic contacts between
orexin-containing axon terminals and locus coeruleus neurons
(Horvath et al. 1999). These
morphological characteristics suggest synaptic transmission at high
concentrations of the transmitter.
GIRK channels are activated by somatostatin (LC neurons) and nociceptin
(TM) through signal transduction by PTX-sensitive G-proteins. The final agent
that activates GIRK channels is the G-protein subunits
(Logothetis et al. 1987). We
have now shown that GIRK channels are suppressed by OXA through a
PTX-insensitive G-protein. The GIRK activity is thus controlled by two
opposing signals. This dual regulation was previously reported in LC neurons
and in dopaminergic neurons in relation to the effect of somatostatin (or
metenkephalin) versus substance P (Koyano
et al. 1993; Velimirovic et
al. 1995), or D2-dopaminergic agonist versus
neurotensin (Farkas et al.
1997). These opposing regulations are unique for GIRK in the sense
that the effects of the two transmitters converge on the same channel. In the
hyperpolarization-activated cation channels (Ih),
apparently opposing transmitter regulations are also documented. In the case
of Ih, however, the antagonistic influences
converge on adenylate cyclase, not on the channel itself
(DiFrancesco 1993). It is
important to note that OXA-induced GIRK channel suppression occurred as long
as there was some level of GIRK channel activity present, regardless of
whether this activity was induced by SOM, NOCI, DAMGO, or G
overexpression. Therefore the observed OXA effects are unlikely to be
attributable to the modulation of the inhibitory transmitter effects, but
rather to direct effects on the GIRK activity.
The signaling pathways for GIRK inhibition are the focus of a controversial
debate. Several investigators concluded that transmitter-induced GIRK
inhibitions are caused by depletion of the phosphatidylinositol
4,5-bisphosphate (PIP2) level in the membrane
(Cho et al. 2001;
Kobrinsky et al. 2000;
Lei et al. 2001), whereas
others emphasized the role of PKC-induced phosphorylation
(Hill and Peralta 2001;
Leaney et al. 2001;
Sharon et al. 1997). Our
recent data suggest that substance P (not orexin)–induced GIRK
inhibition could originate from direct interaction between
Gq and GIRK (Koike et
al. 2003). It is important to note that the time course of the
orexin-induced GIRK inhibition is slower than that of the substance
P–induced GIRK inhibition; hence, the conclusion based on the effect of
substance P might not be applicable to the orexin-induced event.
Orexin causes biphasic effects in HEK293A cells: activation and subsequent suppression of GIRK channels
In HEK293A cells expressing OX1R or OX2R, we observed
OXA induces a biphasic response: an initial phase of GIRK activation followed
by a second phase of long-lasting GIRK inhibition. This biphasic effect became
evident when OXA was applied to HEK cells lacking a precedent GIRK activation
through an inhibitory transmitter or through the overexpression of
G. Such a precedent GIRK activation would have fully activated
the GIRK channel, leaving no room for further conductance increase.
The OXA-induced initial phase of GIRK activation was partially PTX
sensitive and partially PTX insensitive, whereas the subsequent phase of
conductance decrease was entirely PTX insensitive. This result suggests that
both OX1R and OX2R couple to more than one type of
G-protein, including PTX-sensitive (such as Gi/o) and
PTX-insensitive (such as Gq/11) G-proteins. The initial GIRK
activation could be mediated partly by the G released from a
PTX-sensitive G-protein (such as Gi/o) and partly by the
G released from a PTX-insensitive G-protein (such as
Gq/11 and/or Gz), whereas the subsequent inhibitory
action on GIRK activity could be caused by Gq/11.
A biphasic effect through Gq/11-activating receptors was first
reported by Sharon et al.
(1997) in oocytes expressing
metabotropic glutamate receptors. They proposed that its inhibitory effect on
GIRK activity is mediated by a Ca-independent PKC. More recently, Leaney et
al. (2001) reported a biphasic
response caused by the M1 and M3 muscarinic receptors in
the heterologous system of HEK293 cells. In this case, however, the initial
GIRK activation was PTX sensitive in M1 receptors, but not in
M3 receptors.
In cultured cells where GIRK channels were not preactivated by inhibitory transmitters, GIRK activation on OXA application was not observed in either LC neurons (n = 8) or TM neurons (n = 3).
A possible explanation is that when receptors are overexpressed in a
heterologous system, the "wrong" G-proteins could couple to the
receptors (Gudermann et al.
1997), suggesting that the biphasic response in our HEK293 cell
experiments may not be readily observed in native neurons. Another possibility
is that the type(s) of G-protein that couple to orexin receptors may vary in
different kinds of cells. Laburthe et al.
(2002) reported that the types
of G-proteins that couple to VIP receptors (VPAC1 and VPAC2) vary among cell
types and animal species. These are merely speculative explanations, which
need to be verified.
In summary, OXA suppresses GIRK channels that were previously activated by inhibitory transmitters in primary cultured neurons from the LC and TM, as well as a heterologous system. When applied to a reconstituted system in which GIRK channels are not preactivated, OXA induces a biphasic response through both receptor types: an initial, partially PTX sensitive increase, and a subsequent PTX-insensitive decrease in GIRK channel activity. The modulation of GIRK channels by orexins may be one of the cellular mechanisms for the regulation of brain nuclei that are crucial for arousal, sleep, and appetite.
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DISCLOSURES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESACKNOWLEDGMENTSREFERENCES |
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FOOTNOTES |
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Address for reprint requests: Y. Nakajima, Department of Anatomy and Cell Biology, University of Illinois at Chicago, 808 S. Wood Street (M/C 512), Chicago, Illinois 60612-7308 (E-mail: yasukon{at}uic.edu).
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