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Department of Molecular and Integrative Physiology, University of Illinois; Department of Pharmacology, College of Medicine, University of Illinois; and Beckman Institute, University of Illinois, Urbana-Champaign, Urbana, Illinois
Submitted 21 October 2005; accepted in final form 17 April 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; von Krosigk et al. 1993; Steriade et al. 1993; Huguenard and Prince 1994; Warren and Jones 1994). The rhythmic activities arise from the reciprocal synaptic connectivity between thalamic relay nuclei and the adjacent thalamic reticular nucleus (TRN) in conjunction with the intrinsic properties of thalamic neurons (Deschênes et al. 1982;Jahnsen and Llinás 1984a; von Krosigk et al. 1993). Although the cellular mechanisms required to maintain such rhythmic activities are well understood, those involved in the initiation and termination of these rhythmic activities are unclear. Certain neuromodulators, which arise from brain stem nuclei, such as acetylcholine, norepinephrine, and serotonin, have been found to attenuate the rhythmic activities (Lee and McCormick 1997; Lee and McCormick 1996; McCormick 1992). This termination of the rhythmic activity is closely related to changes in behavioral states.
There is increasing evidence that another group of neuromodulators namely, neuropeptides can also alter thalamic activity. A variety of neuropeptides (e.g., cholecystokinin, neuropeptide Y, nociceptin/orphanin FQ, somatostatin, substance P, and vasoactive intestinal peptide [VIP]), along with their respective receptors, have been localized within the thalamus and have been shown to alter the excitability of thalamic neurons by changing intrinsic properties of these cells such as resting membrane potential, input resistance, membrane conductance, and action potential firing mode or affecting synaptic transmission (Cox et al. 1995; Leresche et al. 2000; Sun et al. 2001; Sun et al. 2002; Lee and Cox 2003; Sun et al. 2003). Because neuropeptides have been found to be colocalized with classical neurotransmitters (i.e., 
-aminobutyric acid [GABA] and glutamate) in other brain regions, released in an activity-dependent manner, and produce long-lasting changes in neuronal excitability, they may play an important role in regulating thalamic activity (Lundberg and Hökfelt 1983).
VIP and pituitary adenylate cyclase–activating polypeptide (PACAP) belong to the glucagons-secretin-VIP family of peptides. VIP was originally isolated from porcine intestine as a 28–amino acid peptide and is involved in many regulatory functions, including vasodilation, gastrointestinal secretion and motility, and glycogenolysis (Gozes and Brenneman 1989). VIP is also widely distributed in the central and peripheral nervous system, where it has been found to produce a variety of actions (Gozes and Brenneman 1989). PACAP is a 38–amino acid peptide that shares 68% identity with VIP and was originally isolated from ovine hypothalamus and found to increase cyclic AMP (cAMP) accumulation in anterior pituitary cells (Miyata et al. 1989). PACAP is also localized within a variety of brain areas and peripheral organs and exerts a broad range of physiological actions, including the regulation of anterior pituitary cell function, adrenal gland activity, endocrine and exocrine pancreas secretions, and testicular spermatogenesis (Arimura and Shioda 1995).
VIP and PACAP can activate three different subtypes of receptors in the CNS: VPAC1, VPAC2, and PAC1 (Gozes and Brenneman 1989; Harmar et al. 1998). VIP and PACAP have similar affinities for VPAC1 and VPAC2 receptors; however, PACAP has a relatively higher affinity than VIP for PAC1 receptors. All three receptor subtypes are GTP-dependent protein-coupled receptors and are coupled to cAMP and/or inositol phosphate turnover (Ishihara et al. 1992; Deutsch and Sun 1992; Spengler et al. 1993; Van Rampelbergh et al. 1997; MacKenzie et al. 2001). These receptor subtypes are differentially distributed throughout the brain: VPAC1 receptors are most abundant in the cerebral cortex and hippocampus (Ishihara et al. 1992), VPAC2 receptors are primarily found in the thalamus and suprachiasmatic nucleus (Usdin et al. 1994; Sheward et al. 1995), and PAC1 receptors are more evenly distributed in various brain regions with relatively higher concentrations within the olfactory bulb, thalamus, hypothalamus, hippocampus, and cerebellum.
VIP is localized within TRN neurons, and these neurons in turn innervate almost all thalamic nuclei (Jones 1985). Both VPAC2 and PAC1 receptors are localized within primary sensory thalamic nuclei, including the ventrobasal nucleus (VB) and the dorsal lateral geniculate nucleus (Usdin et al. 1994; Sheward et al. 1995; Vertongen et al. 1997; Burgunder et al. 1999). VPAC1 receptors, however, have not been localized within the thalamus (Usdin et al. 1994). We have shown elsewhere that VIP, via an unidentified receptor subtype, selectively depolarizes VB relay neurons and attenuates slow intrathalamic oscillations (Lee and Cox 2003; Sun et al. 2003). In a previous study, the VIP and PACAP depolarization of thalamic neurons was claimed to be via activation of PAC1 receptors based on agonist concentrations used (Sun et al. 2003). These pharmacological agents, however, are not particularly selective. In light of the availability of transgenic mice lacking the VPAC2 receptor, we can determine the role of VPAC2 receptor-mediated actions as well as PAC1 receptor-mediated actions on relay neurons. Our results indicate that VIP depolarizes relay neurons and attenuates intrathalamic oscillation via activation of VPAC2 receptors in mouse thalamus. PACAP, presumably through the activation of PAC1 receptors, also depolarizes relay neurons via a common mechanism, the enhancement of the hyperpolarization-activated mixed cation current, Ih. Finally, the VPAC2 receptor-mediated actions require the activation of the cAMP-dependent second messenger pathways. Some of these findings have been presented in abstract form (Lee et al. 2003).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). Mice deficient in VPAC2 receptor (VPAC2–/–) were originally generated by Dr. A. Harmer (Harmar et al. 2002). The University of Illinois colony of VPAC2–/– and VPAC2+/+ mice was derived from five breeding pairs of heterozygous mice (C57BL/6J) kindly provided by Dr. Harmar. Procedures used for genotyping the mice in these experiments were similar to those previously described (Harmar et al. 2002). Young VPAC2–/– and VPAC2+/+ mice (postnatal age, 10–21 days) were deeply anesthetized with sodium pentobarbital (50 mg/kg) and decapitated. The brain was quickly removed and placed into cold, oxygenated slicing medium containing (in mM): 2.5 KCl, 10.0 MgCl2, 0.5 CaCl2, 1.25 NaH2PO4, 26.0 NaHCO3, 11.0 glucose, and 234.0 sucrose. Tissue slices (300–400 µm thickness) were cut in the horizontal plane using a vibrating tissue slicer, transferred to a holding chamber, and incubated 
1 h prior to recording. Individual slices were then transferred to a recording chamber and continuously superfused with warm (30°C), oxygenated physiological saline containing (in mM): 126.0 NaCl, 2.5 KCl, 1.25 MgCl2, 2.0 CaCl2, 1.25 NaH2PO4, 26.0 NaHCO3, and 10.0 glucose. This solution was gassed with 95% O2-5% CO2 to a final pH of 7.4. Intracellular recording procedures
Intracellular recordings using the whole-cell configuration were obtained with the visual aid of an Axioskop 2FS equipped with differential interference contrast optics (Zeiss Instruments, Thornwood NY). Individual slices were transferred to a submersion-type recording chamber. A low-power objective (5x) was used to identify specific thalamic nuclei, and a high-power water immersion objective (63x) was used to visualize individual neurons. Recording pipettes were pulled from 1.5 mm outer diameter capillary tubing and had tip resistances of 3–6 M
when filled with the following intracellular solution (in mM): 117 K-gluconate, 13 KCl, 1.0 MgCl2, 0.07 CaCl2, 0.1 EGTA, 10.0 HEPES, 2.0 Na2-ATP, and 0.4 Na-GTP. The pH was adjusted to 7.3 and osmolarity was adjusted to 290–300 mosm. Initial access resistances following break-in typically ranged from 10 to 25 M and remained stable during most recordings included for analyses in this study. An Axoclamp2B amplifier (Axon Instrument, Foster City, CA) was used in bridge mode for voltage recordings (current clamp mode) or switching single-electrode voltage clamp mode for current recordings. Current and voltage protocols were generated by use of pClamp software (Axon Instruments), and data were stored on a computer. For current clamp recordings, an active bridge circuit was continuously adjusted to balance the drop in potential produced by passing current through the recording electrode. The apparent input resistances of individual neurons were calculated from the linear slope of the voltage-current relationship obtained by applying constant current pulses ranging from –100 to +40 pA (800 ms duration). During VIP and PACAP38 application, changes in input resistance were determined by membrane responses to single-intensity constant current hyperpolarizing pulses (5–40 pA, 500 ms, 0.2 Hz). For voltage clamp recordings, the amplifier was used in discontinuous mode. In these recordings, the switching frequency ranged from 2.5 to 3.5 kHz, with a gain of 150 to 800 pA/mV, and the headstage was continually monitored to ensure that the current transients had completely decayed before voltage measurements. Voltage-clamp recordings were limited to neurons that had a stable access resistance <30 M.
Extracellular recording procedures
Extracellular multiple unit recordings were obtained using sharpened tungsten microelectrodes (Frederick Haer, Inc., Bowdoinham, ME). All data were digitized (1–2 kHz) and stored using pClamp software (Axon Instruments). Monopolar electrical stimulation was applied to either TRN or internal capsule by use of sharpened tungsten electrodes (200–600 k). Analyses of intrathalamic rhythmic activities were similar to those described previously (Lee and Cox 2003). In brief, autocorrelograms were constructed from the extracellular multiple-unit data during a 3–9 s period with a bin size of 30 ms to quantify degree of synchrony and duration of intrathalamic oscillations (Minianalysis, Synaptosoft Inc., Fort Lee, NJ). Three measures were used to quantify oscillatory activity in autocorrelograms: number of peaks, oscillation amplitude (Amposc), and frequency of oscillation. Alterations in Amposc indicate a change in the number of unit discharges in the rhythmic activity. The number of peaks indicates the number of cycles in the rhythm, and the oscillation frequency reflects the principle frequency of the rhythmic activity.
Pharmacological agents
Concentrated stock solutions of VIP and PACAP38 were prepared in distilled water and diluted in physiological saline to a concentration of 0.4–4.0 µM. VIP and PACAP38 were applied by injecting a bolus into the input line of the chamber using a motorized syringe pump. On the basis of the rate of agonist injection and the rate of chamber perfusion, the final bath concentration of drugs was estimated to be one eighth of the concentration introduced in the flow line and this dilution is reflected within the results (Cox et al. 1995; Lee and Cox 2003). Control injections of physiological saline did not alter the intrathalamic activity during extracellular recording or intrinsic properties (e.g., membrane potential, input resistance) of neurons during current-clamp recordings, which suggests that the temporary increase in flow rate during the bolus injections had no effect on the recordings. VIP and PACAP38 were purchased from Calbiochem (San Diego, CA) and ZD7288 from Tocris (Ellisville, MO). All remaining compounds were purchased from Sigma (St. Louis, MO).
Statistics
Data are presented as mean ± SD. Most statistical analyses consist of Mann-Whitney U test and, when appropriate, the Wilcoxon test for paired samples. In some noted instances, a students’ t-test or paired t-test was used for testing statistical significance. Results were considered statistically significant when P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Extracellular recording electrodes were placed in the VB to monitor intrathalamic rhythmic activity that arises from the reciprocal synaptic connectivity between VB and the adjacent TRN. Single electrical stimuli in or near TRN typically evoked a spindle-like rhythmic activity (5–8 Hz) in slices from VPAC2+/+ and VPAC2–/– mice that could last for many seconds (Fig. 1Ai). These spindle-like rhythms were produced in 76% of slices from VPAC2+/+ mice (13/17 slices) and 78% of slices from VPAC2–/– mice (14/18 slices). This incidence rate of obtaining the rhythmic activity was not significantly different between the two groups (P > 0.5, Fisher’s Exact test). The autocorrelogram clearly indicates a highly synchronized response with duration of approximately 2 s (approximately 10 cycles) in a slice from a VPAC2+/+ mouse (Fig. 1B, left panel, black line) and a similar duration (2 s, approximately 12 cycles) in a slice from a VPAC2–/– mouse (Fig. 1B, right panel, black line). Under these conditions, the frequency and duration of the rhythmic activity were stable from trial to trial. From the autocorrelograms, we quantified the number of peaks and interburst frequency. The interburst periods of the spindle-like activity produced in slices from VPAC2–/– mice (170 ± 35 ms, n = 12) were significantly shorter than those evoked in slices from VPAC2+/+ mice (213 ± 45 ms, n = 10; P < 0.05, Mann-Whitney U test), but there was no significant difference in the number of peaks (Fig. 1C and D).
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; Bal et al. 1995; McCormick et al. 1995), addition of the GABAA receptor antagonist, bicuculline methiodide, (BMI, 10 µM), slowed the 5–8 Hz activity to a 2–4 Hz rhythmic activity in all slices tested (Figs. 1A and 2A; n = 20). Under these conditions, the frequency and duration of the rhythmic activity was very stable from trial to trial (Figs. 1Aii and 2Aii). The autocorrelograms clearly indicate a highly synchronized response that lasts four cycles (approximately 1.5 s) in a slice from a VPAC2+/+ mouse (Fig. 1B, left panel, gray line) and three cycles (approximately 1.5 s) in a slice from a VPAC 2–/– mouse (Fig. 1B, right panel, gray line). The interburst periods and the number of peaks of the rhythmic activity did not significantly differ in slices from VPAC2+/+ and VPAC2–/– mice (Fig. 1, C and D; P > 0.05; Mann-Whitney U test).
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). The contour plot illustrates that the maximum effect occurred 120 s after VIP treatment and recovered near predrug levels within 5 min following VIP application (Fig. 2Aii, left column). VIP significantly reduced the number of peaks and Amposc in all VPAC2+/+ slices (Fig. 2B, P < 0.05; Mann-Whitney U test). In addition to the suppression of the rhythmic activity, the interburst period was shortened from 220 ms to 150 ms after VIP application in two slices in which VIP severely attenuated but did not eliminate the rhythmic activity. Following the addition of BMI (10 µM) to transform the spindle-like rhythm to a slow-wave rhythm, VIP (0.5 µM, 60 s) also dramatically suppressed the slow rhythmic activity in a reversible manner in all slices tested from VPAC2+/+ mice (Fig. 2Ai, ii, left panel; n = 7). In slices from VPAC2–/– mice, this relatively high concentration of VIP (0.5 µM) did not alter the spindle-like or slow-wave rhythmic activity (Fig. 2Ai, ii, iii). Our population data indicate that VIP did not significantly alter the number of peaks, Amposc, or interburst period in slices from VPAC2–/– mice (Fig. 2B, P > 0.1, Mann-Whitney U test). Our data indicate that the antioscillatory action of VIP is mediated through the activation of VPAC2 receptors. VIP depolarizes relay neurons in VPAC2+/+ slices, not in VPAC2–/– slices
We next examined the effects of VIP on thalamic relay neurons in slices from VPAC2+/+ and VPAC2–/– mice. Intracellular recordings were obtained from 68 relay neurons from VPAC2+/+ mice and 61 relay neurons from VPAC2–/– mice. The average resting membrane potential (VPAC2+/+: –67.7 ± 3.1 mV; VPAC2–/–: –68.7 ± 2.5 mV) and apparent input resistance (VPAC2+/+: 210.0 ± 82.5 M; VPAC2–/–: 218.2 ± 87.0 M) of relay neurons from VPAC2+/+ and VPAC2–/– mice did not differ significantly (P > 0.1, Mann-Whitney U test).
Similar to what we had previously observed in rat thalamic relay neurons, VIP (0.5 µM) consistently depolarizes thalamic relay neurons from wild-type mice (Fig. 3A). This VIP concentration (0.5 µM) depolarized all relay neurons tested from VPAC2+/+ mice with an average amplitude of 3.9 ± 1.5 mV (n = 23). The latency to peak of the VIP-mediated depolarization occurred 115 ± 26 s (n = 23) after VIP application, and the depolarization lasted an average of 333 ± 71 s in 12 of 17 neurons that completely recovered to the pre-VIP membrane potential. A smaller depolarization was produced by lower VIP concentration (0.1 µM, 3.4 ± 0.8 mV, n = 3, data not shown). A reduction in the apparent input resistance was commonly associated with the VIP-mediated membrane depolarization (Fig. 3A). In a subset of neurons in which the membrane potential was manually repolarized to predrug levels during the peak of the VIP-mediated depolarization, the input resistance was reduced by 36 ± 4% (n = 4).
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) and VPAC2–/– (
) animals. It is clear that the excitatory action of VIP on thalamic relay neurons is predominantly mediated via activation of VPAC2 receptors.
In our previous study in rat thalamus, we found that VIP enhances Ih in relay neurons (Lee and Cox 2003). We sought to extend this finding to the mouse thalamus but, more important, determine whether the alteration of Ih is abolished in the VPAC2–/– animals. Voltage-clamp recordings were used to characterize the VIP-mediated conductance changes in relay neurons from VPAC2+/+ and VPAC2–/– animals. Long-duration (2.4 s) voltage step commands (–50 to –120 mV; 10 mV increments; 0.125 Hz) were used to activate Ih in relay neurons. The current responses to the step commands consisted of an initial instantaneous response followed by a slow inward current that typically reached a steady state after a couple of seconds (Fig. 4A). We calculated Ih as the difference between the initial instantaneous response (Iins) and the steady-state level (Iss). Little Ih was evoked in response to the initial small voltage commands, but with stronger hyperpolarizing voltage commands there was an increase in the slow inward current (Fig. 4A, black line). Following VIP application (0.5 µM; 60 s duration), Ih was enhanced similar to that observed in rat thalamic relay neurons (Fig. 4A, VPAC2+/+, gray lines). VIP produced a statistically significant shift in the depolarizing direction of the membrane potential at half-maximal activation in a reversible manner (6.6 ± 3.5 mV, n = 5, P < 0.02; paired t-test, Fig. 4B, VPAC2+/+). In contrast, VIP did not alter Ih in relay neurons from VPAC2–/– mice (Fig. 4A, B, VPAC2–/–, n = 6). It is important to note that, despite our negative findings with VIP, Ih was present in the VPAC2–/– animals and was actually slightly greater in amplitude than that recorded from neurons of VPAC2+/+ animals (Fig. 4C). Furthermore, the voltage dependence of Ih did not differ in the two different types of animals (Fig. 4D). The membrane potential at half-maximal activation was –96.4 ± 2.0 mV (n = 10) and –97.8 ± 3.0 mV (n = 13) in relay neurons from VPAC2+/+ and VPAC2–/– mice, respectively (P > 0.2, Mann-Whitney U test; Fig. 4D). These data support the role of VPAC2 receptors underlying the VIP-mediated enhancement of Ih and membrane depolarization in thalamic relay neurons.
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The fact that VIP produced a small depolarization in 2 of 10 neurons in slices from VPAC2–/– mice suggests VIP may also acton other receptor subtypes (i.e., VPAC1, PAC1). Both VPAC2 and PAC1 receptors are localized in the thalamus (Masuo et al. 1992; Hashimoto et al. 1996; Shioda et al. 1997). Because VIP has high affinity for both VPAC1 and VPAC2 receptors, but a much lower affinity (approximately 1,000-fold less) for the PAC1 receptor, it is possible that the VIP concentrations used in our current study (0.5 µM) could also activate PAC1 receptors. It is important to note, however, that this VIP-mediated action was only in a small minority of neurons in the VPAC2–/– slices. To probe for PAC1 receptor–mediated actions, we next used the agonist, PACAP38, which has a relatively higher affinity for PAC1 receptors.
In neurons from VPAC2–/– mice, PACAP38 (0.5 µM) depolarized 10 of 11 neurons tested (Fig. 5, A and C). The average response to PACAP38 (0.5 µM) was 2.6 ± 0.9 mV (n = 10). Of the 10 cells tested, 5 were in control physiological solution, and the remaining 5 in TTX (1.0 µM). The amplitudes of the PACAP38-mediated responses did not significantly differ between these two conditions (P > 0.1; Mann-Whitney U test), and thus these data have been combined. The latency to peak of the PACAP38-mediated depolarization occurred 176 ± 48 s (n = 10) after application, and the membrane potential partially recovered to predrug levels. Similar to VIP, the PACAP38-mediated membrane depolarization was associated with a decrease in apparent input resistance (17 ± 16%, n = 5).
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; Lam and Srivastava 1990). In slices from VPAC2+/+ mice, PACAP38 (0.5 µM) produced a depolarization in 9 of 11 neurons, which averaged 3.4 ± 0.9 mV (n = 9; Fig. 5, B and C). Of the 9 cells tested, 4 were in control ACSF, and the remaining 5 in 1.0 µM TTX. The amplitude of the PACAP38-mediated depolarization did not differ between these two conditions, and thus these data have been combined (P > 0.1, t-test). The latency to peak of the PACAP38-mediated depolarization occurred 169 ± 42 s (n = 9) after application and the membrane potential partially recovered to predrug levels. The PACAP38-mediated membrane depolarization was associated with a reduction in apparent input resistance (32 ± 18%, n = 5; P < 0.05, paired t-test). Because relay neurons from VPAC2+/+ mice may have more receptors (i.e., VPAC2, PAC1), it is interesting that the PACAP38-mediated depolarization in relay neurons from VPAC2+/+ mice did not significantly differ from that in relay neurons from VPAC2–/– mice (P > 0.1; Mann-Whitney U test). Thus, we next tested whether VIP and PACAP38 share identical pathways involved in membrane depolarization. We attempted whether VIP could occlude the membrane depolarization produced by PACAP38. As illustrated in Fig. 5D, VIP (0.5 µM) depolarized all relay neurons tested from VPAC2+/+ slices (n = 6, 4.2 ± 1.1 mV). At the peak of VIP effect, the membrane potential was manually adjusted to resting levels and PACAP38 (0.5 µM) was applied. Under these conditions, PACAP38 produced no obvious depolarization in all cells tested (n = 6; Fig. 5D). Following washout of PACAP38 and VIP, a subsequent PACAP38 application (16 min interval) produced membrane depolarizations in four of six neurons tested (n = 4, 1.9 ± 0.1 mV).
PACAP38 enhances Ih in relay neurons from VPAC2 –/– slices
Our initial focus in this study was to identify the specific receptor subtype underlying the VIP-mediated depolarization in thalamic relay cells; however, in light of the small population of neurons in which VIP still depolarizes neurons in the VPAC2–/– animal, this animal model also provides a unique tool to study the role of PAC1 receptors in the thalamus. Voltage-clamp recordings were used to characterize the conductance changes produced by PACAP38 in relay neurons from VPAC2–/– mice. Analogous to our current clamp recordings above, PACAP38 (0.05–0.5 µM) produced an inward current that partially recovered to predrug holding current levels (Fig. 6A). In TTX (1.0 µM), PACAP38 (0.05–0.5 µM) produced an inward current that averaged 18 ± 15 pA (n = 8), and the time to peak of the inward current averaged 198 ± 56 s, which is similar to our current clamp recordings. Slow voltage command ramps (–60 mV to –110 mV, 2 s duration, 0.1 Hz) were used to determine the voltage dependence of the PACAP38-mediated alteration in conductance (Fig. 6, A and B). In the predrug condition, the current response to the ramped voltage command is nonlinear; during the ramped voltage command there is an increase in conductance at more hyperpolarized potentials, which is likely a result of Ih activation. To quantify changes in the "resting" conductance of the neuron, we analyzed the initial portion of the current response near resting membrane potential (–60 to –80 mV). The resting conductance of the neurons prior to PACAP38 averaged 4.7 ± 2.1 nS. Following PACAP38 application, the conductance was significantly increased to 5.6 ± 2.6 nS (n = 9; P < 0.02, Wilcoxon test). Of the nine cells tested, 0.1 µM PACAP38 was applied to four cells, 0.5 µM to four cells, and 0.05 µM to the remaining cell, and these data have been combined for this analysis.
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). In light of our previous work demonstrating that the VIP-mediated response was completely attenuated by the Ih blocker, ZD7288, we next tested whether the PACAP38-mediated response was also sensitive to this agent. In ZD7288 (100 µM), PACAP38 (0.5 µM) did not produce inward current, nor was there any change in the current response to the ramped voltage commands (Fig. 6D-E, n = 5). Our data suggest that PACAP38 enhances Ih in relay neurons from VPAC2–/– animals. Role of cAMP in VIP-mediated actions
Previous studies have demonstrated that increases in cAMP activity can enhance Ih in thalamic relay neurons (Budde et al. 1997; Lüthi and McCormick 1999; Sun et al. 2003). In light of the absence of the VIP-mediated enhancement of Ih in neurons from VPAC2–/– animals, we tested whether this pathway may have been disrupted by the genetic manipulation. Voltage steps commands were used to activate Ih in neurons of slices from VPAC2+/+ and VPAC2–/– animals (Fig. 7). Similar to VIP, the membrane permeable, cAMP analogue, 8-cpt-cAMP (1 mM; 60 s application) produced a statistically significant depolarizing shift in the Ih activation curve (4.8 ± 0.4 mV; n = 5) in neurons from VPAC2+/+ mice (Fig. 7B, P < 0.01; paired t-test). In relay neurons from VPAC2–/– mice, 8-cpt-cAMP (1 mM) also produced a significant depolarizing shift in the Ih activation function (4.2 ± 1.1 mV; n = 5; P < 0.02; paired t-test, Fig. 7B). These results indicate that the deletion of VPAC2 receptor gene did not disrupt the modulation of Ih by cAMP, which thereby suggests that the lack of VIP mediated responses in the VPAC2–/– mice is not due to a disruption of the second messenger pathway.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Intrathalamic rhythmic activities result from the intrinsic properties of thalamic neurons and the reciprocal synaptic connections between thalamic relay nuclei and the TRN (Steriade and Llinás 1988; von Krosigk et al. 1993; Steriade et al. 1993; Huguenard and Prince 1994; Warren and Jones 1994). Spindle-like and slow-wave rhythms have been observed in vivo in many animal species, but in vitro such rhythms have clearly been observed in ferrets, guinea pigs, and, to a lesser extent, in rats. Despite the presence of slow-wave rhythms in mice, spindle-like rhythms have not been obvious (Warren et al. 1994; Huntsman et al. 1999). Our study provides clear evidence that the mouse thalamus is clearly capable of both spindle-like rhythmic activities, and attenuating GABAA receptor-mediated activity transformed the spindle-like rhythmic activity to a slow-wave rhythmic activity similar to that observe in other animal models (Huguenard and Prince 1994; Bal et al. 1995; McCormick et al. 1995). The presence of these intrathalamic rhythms in wild-type mice allows the power of transgenic approaches to be used for the studies of these activities, in a manner similar to our use with the VPAC2–/– mouse.
In our earlier study, we found that VIP attenuates slow-wave rhythmic activity via an unidentified receptor subtype in rat thalamic slices (Lee and Cox 2003). In the present study, we found that VIP attenuates both spindle-like and slow 2–4 Hz intrathalamic rhythms in slices from wild-type (VPAC2+/+) mice. In slices from VPAC2 receptor knock-out animals (VPAC2–/–), VIP did not alter the rhythmic activity. These data clearly indicate that the antioscillatory actions of VIP are mediated via activation of VPAC2 receptors.
The antioscillatory actions of VIP are likely a consequence of the depolarizing actions of VIP on thalamic relay neurons. Within mouse thalamic relay neurons, we found that VIP produced a long-lasting robust membrane depolarization. Despite numerous reports of VIP-mediated membrane depolarizations in different regions of the CNS, the identification of the specific receptors underlying these actions has remained elusive because of poorly selective pharmacological agents for VIP/PACAP receptors (Phillis et al. 1978; Jeftinija et al. 1982; Kohlmeier and Reiner 1999; Liu and Morris 1999; Lee and Cox 2003; Sun et al. 2003). The development of this VPAC2–/– mouse may serve as a valuable tool to investigate the functional role of VPAC2 receptors in various brain regions (Harmar et al. 2002; Aton et al. 2005). Immunocytochemical studies indicate that VPAC2 and PAC1 recepters are present within thalamic nuclei, though distinguishing the functional consequences of these specific receptor subtypes has been difficult with current pharmacological agents (Usdin et al. 1994; Sheward et al. 1995; Vertongen et al. 1997; Burgunder et al. 1999).
The integral role of VPAC2 receptors
The VPAC2–/– mice have served as a very important tool in further understanding of VIP-mediated actions in the thalamus. In slices from VPAC2+/+ mice, we have found that in relay neurons VIP produces a membrane depolarization that is associated with a decreased input resistance. Voltage clamp recordings clearly indicate that VIP enhances Ih, which is similar to what has been observed in rat thalamic neurons (Lee and Cox 2003; Sun et al. 2003). In slices from VPAC2–/– mice, however, the VIP-mediated depolarization in response to a relatively high VIP concentration (0.5 µM) is absent in 80% of the neurons and is severely reduced in amplitude in the remaining cells. In addition, VIP does not alter Ih in relay neurons in slices from VPAC2–/– mice. Thus, our data strongly support the notion that the excitatory actions of VIP are predominantly via VPAC2 receptors. Our results differ from the conclusions of Sun et al. (2003), who concluded that the predominant action of VIP/PACAP was due to PAC1 receptors based on (1) the concentration dependence of their agonists responses (PACAP more potent than VIP) and (2) the VPAC1 receptor agonist, (Ala) VIP, did not depolarize thalamic relay neurons, which suggests the lack of VPAC1 receptor–mediated actions on thalamic relay neurons (Nicole et al. 2000; Sun et al. 2003). Our results strongly indicate the role of the VPAC2 receptor, unless there is a complete remodeling of the VIP/PACAP receptor distribution as a result of the VPAC2 knock-out, but such a remodeling has not been observed in other brain regions of these VPAC2 mice and thus would be specific only to the thalamic nuclei (Aton et al. 2005).
As mentioned above, although the majority of relay neurons from VPAC2–/– mice did not respond to VIP, in the remaining 20% of neurons, VIP produced a smaller depolarization. Because VIP has a lower affinity for PAC1 receptors relative to VPAC2 receptors, the VIP concentration (0.5 µM) used in this study could possibly activate PAC1 receptors. Our finding supports such a conclusion because the agonist, PACAP38, which has a higher affinity for PAC1 receptors than VIP, depolarized the majority of neurons from VPAC2–/– mice. In light of the anatomical data indicating the presence of VPAC2 and PAC1 receptors in thalamic relay nuclei, the VPAC2–/– mice serve as a potentially useful model to understand PAC1-mediated actions within thalamic neurons. We found that PACAP38 not only depolarized thalamic neurons from VPAC2–/– mice, but our voltage clamp recordings also reveal that PACAP38 also increases the conductance of relay neurons and appears to also enhance Ih, similar to that observed by VIP via VPAC2 receptors. Thus, our data clearly indicate that activation of PAC1 receptors can enhance Ih, but, perhaps more important, also suggest that these two different receptors may converge onto a common effector mechanism to alter the excitability of thalamic neurons. Consistent with this idea, our data strongly suggest that VIP occludes PACAP38-mediated membrane depolarization in thalamic neurons from VPAC2+/+ mice.
The dependence of VIP-mediated response on cAMP
VIP-sensitive receptors are G protein–dependent receptors that have been found to engage cAMP synthesis as well as inositol phosphate turnover pathways (Ishihara et al. 1992; Deutsch and Sun 1992; Spengler et al. 1993; Van Rampelbergh et al. 1997; MacKenzie et al. 2001). In addition, hyperpolarization-activated cyclic nucleotide-gated (HCN) channels that underlie Ih, are modulated by alteration in cAMP concentrations (Beaumont and Zucker 2000; Wainger et al. 2001). Several neuromodulators within the thalamus (e.g., norepinephrine, serotonin, VIP, PACAP), which activate receptors coupled to cAMP dependent pathways, enhance Ih and thereby depolarize thalamic neurons (McCormick and Pape 1990; Pape 1996; Sun et al. 2003; Frere and Luthi 2004; present study). Our data demonstrate that cAMP analogues enhance Ih, and produce membrane depolarizations similar to VIP. Furthermore, we have found that activation of cAMP can occlude the VIP-mediated membrane depolarization, which indicates the dependence of the depolarizing action on cAMP dependent processes. Our findings are consistent with previous reports that VPAC2 receptors are primarily coupled to cAMP-dependent pathways (Lutz et al. 1993; Lutz et al. 1999; Itri and Colwell 2003).
Functional role of VIP/PACAP in the thalamus
Thalamic neurons discharge action potentials in two basic firing modes: tonic and burst. The firing mode of thalamic neurons is dependant on the low threshold, transient calcium current, IT (Jahnsen and Llinás 1984a; Steriade and Llinás 1988; Crunelli et al. 1989). At relatively hyperpolarized membrane potentials, channels underlying IT are deactivated, and with membrane depolarization, IT can be activated giving rise to a transient calcium-dependent depolarization, a low-threshold calcium spike, on which a short-duration (100–200 ms), high-frequency burst (>200 Hz) of sodium-dependent action potentials can occur (Jahnsen and Llinás 1984a; Steriade and Llinás 1988; Crunelli et al. 1989). At relatively depolarized membrane potentials IT is inactivated, and subsequent depolarization of the membrane potential will result in tonic action potential discharge at a frequency that is relatively linear with the degree of membrane depolarization (Jahnsen and Llinás 1984a;Jahnsen and Llinás 1984b; Huguenard and McCormick 1992; Zhan et al. 1999).
Several neuromodulators depolarize thalamic neurons, predisposing the neurons to a tonic mode and thereby terminating the intrathalamic rhythmic activity. It is interesting that many of these modulators alter the excitability of thalamic neurons via common effector mechanisms. Two obvious candidates are the resting K current, Kleak, and Ih. Within thalamic neurons, acetylcholine, glutamate (via metabotropic receptors), norepinephrine, serotonin, cholecystokinin, as well as histamine depolarize thalamic neurons by decreasing a resting leak potassium current and thereby alter the action potential discharge mode of these neurons and ultimately terminating the rhythmic activity (McCormick and Prince 1986; McCormick and Prince 1987; McCormick and Prince 1988; McCormick 1989; McCormick and Wang 1991; McCormick and Williamson 1991; McCormick and von Krosigk 1992; McCormick 1992; Cox et al. 1995; Cox et al. 1997; Cox and Sherman 1999). Furthermore, norepinephrine, serotonin, and histamine have been found to enhance Ih, thus leading to membrane depolarization and ultimately terminating the intrathalamic oscillation (Pape and McCormick 1989; McCormick and Pape 1990; McCormick and Williamson 1991). Our data strongly indicate VIP and PACAP38 depolarize relay neurons via an enhancement of Ih (Lee and Cox 2003; Sun et al. 2003). A common aspect of these modulators is their ability to engage particular intracellular second-messenger pathways namely, cAMP which ultimately alters Ih.
Despite sharing a common effector mechanism, a key to understanding the functional aspects of VIP and PACAP likely depends on the source of these peptides. VIP-containing neurons are in the TRN and therefore VIP is endogenous to the intrathalamic circuit, and its release is likely correlated to intrathalamic circuit activity (Burgunder et al. 1999). Other neuropeptides (e.g., cholecystokinin, somatostatin, neuropeptide Y, and VIP) can also alter the firing mode of thalamic neurons and attenuate intrathalamic rhythmic activity (Cox et al. 1997; Leresche et al. 2000; Sun et al. 2002; Lee and Cox 2003; Sun et al. 2003). Similar to VIP, these other neuropeptides are localized within the thalamic neurons and may play a potentially important role in regulating thalamic neuron excitability, but further studies are needed. In contrast to the above-mentioned neuropeptides, PACAP-containing neurons that innervate the thalamus originate from brain stem regions in a somewhat similar manner as other known modulators such as acetylcholine, norepinephrine, serotonin, and histamine (Hashimoto et al. 1996; Shioda et al. 1997; Hannibal 2002). The activity of neurons in these areas is closely associated with levels of arousal and sleep-wake states (Steriade et al. 1993; McCormick and Bal 1997). Thus, the functional distinction between peptides such as VIP and PACAP, which appear to share common cellular mechanisms, likely involves their sites of origin. In the case of these two peptides, VIP appears to be endogenous to the thalamic circuitry, whereas PACAP would presumably be released by increase activity of brain stem neurons.
Our working hypothesis is that high-frequency burst activity of TRN required for intrathalamic rhythmic activity may be optimal for peptide release and therefore produce the release of VIP. This synaptically released VIP activates VPAC2 receptors on relay neurons, increases cAMP activity, enhancing Ih, and thereby depolarizes the relay neurons. This depolarization and decrease of membrane resistance biases the cells to tonic-discharge mode and may contribute to the termination of rhythmic activity. We speculate that VIP acts as an intrinsic modulator of intrathalamic oscillation that could regulate the duration of rhythmic activity. We predict that overall decreases in VIP release could lead to prolonged durations of intrathalamic rhythmic activities similar to that during absence seizure activity. The role of neuropeptides in the thalamus may not only influence rhythmic activities, but considering their long-lasting actions, these compounds may also play an important role in regulating the overall information transfer through thalamocortical circuits.
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Address for reprint requests and other correspondence: Charles L. Cox, Department of Molecular and Integrative Physiology, University of Illinois, 2357 Beckman Institute, 405 North Mathews, Urbana, IL 61801 (E-mail: clcox{at}life.uiuc.edu)
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Aton SJ, Colwell CS, Harmar AJ, Waschek J, and Herzog ED. Vasoactive intestinal polypeptide mediates circadian rhythmicity and synchrony in mammalian clock neurons. Nat Neurosci 8: 476–483, 2005.[Web of Science][Medline]
Bal T, von Krosigk M, and McCormick DA. Synaptic and membrane mechanisms underlying synchronized oscillations in the ferret lateral geniculate nucleus in vitro. J Physiol 483: 641–663, 1995.
Beaumont V and Zucker RS. Enhancement of synaptic transmission by cyclic AMP modulation of presynaptic Ih channels. Nat Neurosci 3: 133–141, 2000.[CrossRef][Web of Science][Medline]
Budde T, Biella G, Munsch T, and Pape HC. Lack of regulation by intracellular Ca of the hyperpolarization-activated cation current in rat thalamic neurones. J Physiol 503 (Pt 1): 79–85, 1997.
Burgunder JM, Heyberger B, and Lauterburg T. Thalamic reticular nucleus parcellation delineated by VIP and TRH gene expression in the rat. J Chem Neuroanat 17: 147–152, 1999.[CrossRef][Web of Science][Medline]
Cox CL, Huguenard JR, and Prince DA. Cholecystokinin depolarizes rat thalamic reticular neurons by suppressing a K conductance. J Neurophysiol 74: 990–1000, 1995.
Cox CL, Huguenard JR, and Prince DA. Peptidergic modulation of intrathalamic circuit activity in vitro: Actions of cholecystokinin. J Neurosci 17: 70–82, 1997.
Cox CL and Sherman SM. Glutamate inhibits thalamic reticular neurons. J Neurosci 19: 6694–6699, 1999.
Crunelli V, Lightowler S, and Pollard CE. A t-type current underlies low-threshold Ca potentials in cell of the cat and rat lateral geniculate nucleus. J Physiol 413: 543–561, 1989.
Deschênes M, Roy JP, and Steriade M. Thalamic bursting mechanism: an inward slow current revealed by membrane hyperpolarization. Brain Res 239: 289–293, 1982.[CrossRef][Web of Science][Medline]
Deutsch PJ and Sun Y. The 38-amino acid form of pituitary adenylate cyclase–activating polypeptide stimulates dual signaling cascades in PC12 cells and promotes neurite outgrowth. J Biol Chem 267: 5108–5113, 1992.
Frere SG and Luthi. A Pacemaker channels in mouse thalamocortical neurones are regulated by distinct pathways of cAMP synthesis. J Physiol 554: 111–125, 2004.
Gottschall PE, Tatsuno I, Miyata A, and Arimura A. Characterization and distribution of binding sites for the hypothalamic peptide, pituitary adenylate cyclase–activating polypeptide. Endocrinology 127: 272–277, 1990.
Gozes I and Brenneman DE. VIP: Molecular biology and neurobiological function. Mol Neurobiol 3: 201–236, 1989.[Web of Science][Medline]
Hannibal J. Pituitary adenylate cyclase–activating peptide in the rat central nervous system: an immunohistochemical and in situ hybridization study. J Comp Neurol 453: 389–417, 2002.[CrossRef][Web of Science][Medline]
Harmar AJ, Arimura A, Gozes I, Journot L, Laburthe M, Pisegna JR, Rawlings SR, Robberecht P, Said SI, Sreedharan SP, Wank SA, and Waschek JA. International Union of Pharmacology. XVIII. Nomenclature of receptors for vasoactive intestinal peptide and pituitary adenylate cyclase–activating polypeptide. Pharmacol Rev 50: 265–270, 1998.
Harmar AJ, Marston HM, Shen S, Spratt C, West KM, Sheward WJ, Morrison CF, Dorin JR, Piggins HD, Reubi JC, Kelly JS, Maywood ES, and Hastings MH. The VPAC(2) receptor is essential for circadian function in the mouse suprachiasmatic nuclei. Cell 109: 497–508, 2002.[CrossRef][Web of Science][Medline]
Hashimoto H, Nogi H, Mori K, Ohishi H, Shigemoto R, Yamamoto K, Matsuda T, Mizuno N, Nagata S, and Baba A. Distribution of the mRNA for a pituitary adenylate cyclase–activating polypeptide receptor in the rat brain: an in situ hybridization study. J Comp Neurol 371: 567–577, 1996.[CrossRef][Web of Science][Medline]
Huguenard JR and McCormick DA. Simulation of the currents involved in rhythmic oscillations in thalamic relay neurons. J Neurophysiol 68: 1373–1383, 1992.
Huguenard JR and Prince DA. Intrathalamic rhythmicity studies in vitro: Nominal t-current modulation causes robust antioscillatory effects. J Neurosci 14: 5485–5502, 1994.[Abstract]
Huntsman MM, Porcello DM, Homanics GE, DeLorey TM, and Huguenard JR. Reciprocal inhibitory connections and network synchrony in the mammalian thalamus. Science 283: 541–543, 1999.
Ishihara T, Shigemoto R, Mori K, Takahashi K, and Nagata S. Functional expression and tissue distribution of a novel receptor for vasoactive intestinal polypeptide. Neuron 8: 811–819, 1992.[CrossRef][Web of Science][Medline]
Itri J and Colwell CS. Regulation of inhibitory synaptic transmission by vasoactive intestinal peptide (VIP) in the mouse suprachiasmatic nucleus. J Neurophysiol 90: 1589–1597, 2003.
Jahnsen H and Llinás R. Electrophysiological properties of guinea-pig thalamic neurones: an in vitro study. J Physiol 349: 205–226, 1984a.
Jahnsen H and Llinás R. Ionic basis for the electroresponsiveness and oscillatory properties of guinea-pig thalamic neurones in vitro. J Physiol 349: 227–247, 1984b.
Jeftinija S, Murase K, Nedeljkov V, and Randic M. Vasoactive intestinal polypeptide excites mammalian dorsal horn neurons both in vivo and in vitro. Brain Res 243: 158–164, 1982.[CrossRef][Web of Science][Medline]
Jones EG. The Thalamus. New York: Plenum Press, 1985.
Kohlmeier KA and Reiner PB. Vasoactive intestinal polypeptide excites medial pontine reticular formation neurons in the brainstem rapid eye movement sleep-induction zone. J Neurosci 19: 4073–4081, 1999.
Lam KS and Srivastava G. Sex-related differences and thyroid hormone regulation of vasoactive intestinal peptide gene expression in the rat brain and pituitary. Brain Res 526: 135–137, 1990.[CrossRef][Web of Science][Medline]
Lee KH and McCormick DA. Modulation of spindle oscillations by acetylcholine, cholecystokinin and 1S,3R-ACPD in the ferret lateral geniculate and perigeniculate nuclei in vitro. Neurosci 77: 335–350, 1997.[CrossRef][Web of Science][Medline]
Lee KH and McCormick DA. Abolition of spindle oscillations by serotonin and norepinephrine in the ferret lateral geniculate and perigeniculate nuclei in vitro. Neuron 17: 309–320, 1996.[CrossRef][Web of Science][Medline]
Lee SH and Cox CL. Vasoactive intestinal peptide selectively depolarizes thalamic relay neurons and attenuates intrathalamic rhythmic activity. J Neurophysiol 90: 1224–1234, 2003.
Lee SH, Harmar AJ, and Cox CL. Excitatory and anti-oscillatory actions of vasoactive intestinal peptide in mouse thalamus mediated by VPAC2 receptors. Soc Neurosci Abst 233, 2003.
Leresche N, Asprodini E, Emri Z, Cope DW, and Crunelli V. Somatostatin inhibits GABAergic transmission in the sensory thalamus via presynaptic receptors. Neurosci 98: 513–522, 2000.[CrossRef][Web of Science][Medline]
Liu XH and Morris R. Vasoactive intestinal polypeptide produces depolarization and facilitation of C-fiber evoked synaptic responses in superficial dorsal horn neurones (laminae I-IV) of the rat lumbar spinal cord in vitro. Neurosci Lett 276: 1–4, 1999.[CrossRef][Web of Science][Medline]
Lundberg JM and Hökfelt T. Coexistence of peptides and classical neurotransmitters. Trends Neurosci 6: 325–333, 1983.[CrossRef][Web of Science]
Lüthi A, and McCormick DA. Modulation of a pacemaker current through Ca-induced stimulation of cAMP production. Nat Neurosci 2: 634–641, 1999.[CrossRef][Web of Science][Medline]
Lutz EM, MacKenzie CJ, Johnson M, West K, Morrow JA, Harmar AJ, and Mitchell R. Domains determining agonist selectivity in chimaeric VIP2 (VPAC2)/PACAP (PAC1) receptors. Br J Pharmacol 128: 934–940, 1999.[CrossRef][Web of Science][Medline]
Lutz EM, Sheward WJ, West KM, Morrow JA, Fink G, and Harmar AJ. The VIP2 receptor: molecular characterisation of a cDNA encoding a novel receptor for vasoactive intestinal peptide. FEBS Lett 334: 3–8, 1993.[CrossRef][Web of Science][Medline]
MacKenzie CJ, Lutz EM, Johnson MS, Robertson DN, Holland PJ, and Mitchell R. Mechanisms of phospholipase C activation by the vasoactive intestinal polypeptide/pituitary adenylate cyclase–activating polypeptide type 2 receptor. Endocrinology 142: 1209–1217, 2001.
Masuo Y, Ohtaki T, Masuda Y, Tsuda M, and Fujino M. Binding sites for pituitary adenylate cyclase–activating polypeptide (PACAP): comparison with vasoactive intestinal polypeptide (VIP) binding site localization in rat brain sections. Brain Res 575: 113–123, 1992.[CrossRef][Web of Science][Medline]
McCormick DA. Cholinergic and noradrenergic modulation of thalamocortical processing. Trends Neurosci 12: 215–221, 1989.[CrossRef][Web of Science][Medline]
McCormick DA. Neurotransmitter actions in the thalamus and cerebral cortex and their role in neuromodulation of thalamocortical activity. Prog Neurobiol 39: 337–388, 1992.[CrossRef][Web of Science][Medline]
McCormick DA and Bal T. Sleep and arousal: Thalamocortical mechanisms. Ann Rev Neurosci 20: 185–215, 1997.
McCormick DA and Pape H-C. Noradrenergic and serotonergic modulation of a hyperpolarization-activated cation current in thalamic relay neurons. J Physiol 431: 319–342, 1990.
McCormick DA, and Prince DA. Acetylcholine induces burst firing in thalamic reticular neurones by activating a potassium conductance. Nature 319: 402–405, 1986.[CrossRef][Medline]
McCormick DA and Prince DA. Actions of acetylcholine in the guinea-pig and cat medial and lateral geniculate nuclei, in vitro. J Physiol 392: 147–165, 1987.
McCormick DA, and Prince DA. Noradrenergic modulation of firing pattern in guinea pig and cat thalamic neurons, in vitro. J Neurophysiol 59: 978–996, 1988.
McCormick DA Trent F, and Ramoa AS. Postnatal development of synchronized network oscillations in the ferret dorsal lateral geniculate and perigeniculate nuclei. J Neurosci 15: 5739–5752, 1995.[Abstract]
McCormick DA and von Krosigk M. Corticothalamic activation modulates thalamic firing through glutamate "metabotropic" receptors. Proc Natl Acad Sci USA 89: 2774–2778, 1992.
McCormick DA and Wang Z. Serotonin and noradrenaline excite GABAergic neurones of the guinea pig and cat thalamic reticular nucleus. J Physiol 442: 235–255, 1991.
McCormick DA and Williamson. A Modulation of neuronal firing mode in cat and guinea pig LGNd by histamine: possible cellular mechanisms of histaminergic control of arousal. J Neurosci 11: 3188–3199, 1991.[Abstract]
Miyata A, Arimura A, Dahl RR, Minamino N, Uehara A, Jiang L, Culler MD, and Coy DH. Isolation of a novel 38 residue-hypothalamic polypeptide which stimulates adenylate cyclase in pituitary cells. Biochem Biophys Res Commun 164: 567–574, 1989.[CrossRef][Web of Science][Medline]
Nicole P, Lins L, Rouyer-Fessard C, Drouot C, Fulcrand P, Thomas A, Couvineau A, Martinez J, Brasseur R, and Laburthe M. Identification of key residues for interaction of vasoactive intestinal peptide with human VPAC1 and VPAC2 receptors and development of a highly selective VPAC1 receptor agonist: Alanine scanning and molecular modeling of the peptide. J Biol Chem 275: 24003–24012, 2000.
Pape H-C and McCormick DA. Noradrenaline and serotonin selectively modulate thalamic burst firing by enhancing a hyperpolarization-activated cation current. Nature 340: 715–718, 1989.[CrossRef][Medline]
Pape HC. Queer current and pacemaker: the hyperpolarization-activated cation current in neurons. Annu Rev Physiol 58: 299–327, 1996.[CrossRef][Web of Science][Medline]
Phillis JW, Kirkpatrick JR, and Said SI. Vasoactive intestinal polypeptide excitation of central neurons. Can J Physiol Pharmacol 56: 337–340, 1978.[Web of Science][Medline]
Sheward WJ, Lutz EM, and Harmar AJ. The distribution of vasoactive intestinal peptide2 receptor messenger RNA in the rat brain and pituitary gland as assessed by in situ hybridization. Neurosci 67: 409–418, 1995.[CrossRef][Web of Science][Medline]
Shioda S, Shuto Y, Somogyvari-Vigh A, Legradi G, Onda H, Coy DH, Nakajo S, and Arimura A. Localization and gene expression of the receptor for pituitary adenylate cyclase–activating polypeptide in the rat brain. Neurosci Res 28: 345–354, 1997.[CrossRef][Web of Science][Medline]
Spengler D, Waeber C, Pantaloni C, Holsboer F, Bockaert J, Seeburg PH, and Journot L. Differential signal transduction by five splice variants of the PACAP receptor. Nature 365: 170–175, 1993.[CrossRef][Medline]
Steriade M and Llinás R. The functional states of the thalamus and the associated neuronal interplay. Physiol Rev 68: 649–742, 1988.
Steriade M, McCormick DA, and Sejnowski TJ. Thalamocortical oscillations in the sleeping and aroused brain. Science 262: 679–685, 1993.
Sun QQ, Huguenard JR, and Prince DA. Neuropeptide Y receptors differentially modulate G-protein-activated inwardly rectifying K channels and high-voltage-activated Ca channels in rat thalamic neurons. J Physiol 531: 67–79, 2001.
Sun QQ, Huguenard JR, and Prince DA. Somatostatin inhibits thalamic network oscillations in vitro: Actions on the GABAergic neurons of the reticular nucleus. J Neurosci 22: 5374–5386, 2002.
Sun QQ, Prince DA, and Huguenard JR. Vasoactive intestinal polypeptide and pituitary adenylate cyclase–activating polypeptide activate hyperpolarization-activated cationic current and depolarize thalamocortical neurons in vitro. J Neurosci 23: 2751–2758, 2003.
Usdin TB, Bonner TI, and Mezey E. Two receptors for vasoactive intestinal polypeptide with similar specificity and complementary distributions. Endocrinology 135: 2662–2680, 1994.[Abstract]
Van Rampelbergh J, Poloczek P, Francoys I, Delporte C, Winand J, Robberecht P, and Waelbroeck M. The pituitary adenylate cyclase–activating polypeptide (PACAP I) and VIP (PACAP II VIP1) receptors stimulate inositol phosphate synthesis in transfected CHO cells through interaction with different G proteins. Biochim Biophys Acta 1357: 249–255, 1997.[Medline]
Vertongen P, Schiffmann SN, Gourlet P, and Robberecht P. Autoradiographic visualization of the receptor subclasses for vasoactive intestinal polypeptide (VIP) in rat brain. Peptides 18: 1547–1554, 1997.[CrossRef][Web of Science][Medline]
von Krosigk M, Bal T, and McCormick DA. Cellular mechanisms of a synchronized oscillation in the thalamus. Science 261: 361–364, 1993.
Wainger BJ, DeGennaro M, Santoro B, Siegelbaum SA, and Tibbs GR. Molecular mechanism of cAMP modulation of HCN pacemaker channels. Nature 411: 805–810, 2001.[CrossRef][Medline]
Warren RA, Agmon A, and Jones EG. Oscillatory synaptic interactions between ventroposterior and reticular neurons in mouse thalamus in vitro. J Neurophysiol 72: 1993–2003, 1994.
Warren RA and Jones EG. Glutamate activation of cat thalamic reticular nucleus: Effects on response properties of ventroposterior neurons. Exp Brain Res 100: 215–226, 1994.[Web of Science][Medline]
Zhan XJ, Cox CL, Rinzel J, and Sherman SM. Current clamp and modeling studies of low-threshold calcium spikes in cells of the cat’s lateral geniculate nucleus. J Neurophysiol 81: 2360–2373, 1999.
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