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. 1999 Dec;128(8):1779–1787. doi: 10.1038/sj.bjp.0702971

Nociceptin, Phe1ψ-nociceptin1–13, nocistatin and prepronociceptin154–181 effects on calcium channel currents and a potassium current in rat locus coeruleus in vitro

Mark Connor 1,*, Christopher W Vaughan 1, Ernest A Jennings 1, Richard G Allen 3, Macdonald J Christie 2
PMCID: PMC1571808  PMID: 10588934

Abstract

  1. The actions of the neuropeptide nociceptin, the putative nociceptin receptor antagonist [Phe1ψ(CH2-NH)Gly2]-nociceptin-(1–13)NH2 (Phe1ψ-nociceptin1–13) and the putative nociceptin precursor products nocistatin (rat prepronociceptin125–132) and rat prepronociceptin154–181 were examined on membrane properties of rat locus coeruleus (LC) neurons using whole cell patch clamp techniques.

  2. Nociceptin inhibited IBa in all LC neurons, (pD2 of 8.9, maximum inhibition 50%). The inhibition of IBa by nociceptin was associated with slowing of the activation of IBa and could be significantly reversed by a strong depolarizing prepulse. Phe1ψ-nociceptin1–13 also inhibited IBa in LC neurons (notional pD2 of 7.6, maximum inhibition 18%). Application of Phe1ψ-nociceptin1–13 (1 μM) significantly occluded the subsequent effects of a co-application of nociceptin (3 nM) on IBa.

  3. As previously reported for nociceptin, Phe1ψ-nociceptin1–13 caused an outward current in LC neurons voltage clamped at −60 mV (pD2 of 7.1, maximum current 50% of that of methionine enkephalin, 10 μM). The Phe1ψ-nociceptin1–13 induced current reversed polarity at −112 mV and exhibited pronounced inward rectification. Phe1ψ-nociceptin1–13 (1 μM) reversibly inhibited the current caused by nociceptin (300 nM) by 30%.

  4. Neither nocistatin nor rat prepronociceptin154–181 inhibited IBa in LC neurons, or prevented the subsequent inhibition by nociceptin. Neither nocistatin or prepronociceptin154–181 affected the membrane properties of LC neurons.

  5. This study demonstrates that nociceptin modulates somatic IBa in rat LC neurons. The putative ORL1 antagonist Phe1ψ-nociceptin1–13 exhibited partial agonist activity at inhibiting IBa and opening K+ channels in LC. Other putative nociceptin precursor products were without effect on LC cells.

Keywords: Nociceptin, orphanin FQ, nocistatin, ORL1, locus coeruleus, calcium channels, partial agonist, potassium channels

Introduction

Nociceptin (Meunier et al., 1995), also called orphanin FQ (Reinscheid et al., 1995), is an endogenous ligand for the opioid-like receptor, ORL1 (Mollereau et al., 1994; reviewed in Henderson & McKnight, 1997). To date, nociceptin has been shown to modulate a similar range of ion channels and second messenger cascades as opioids (Henderson & McKnight, 1997) and the nociceptin/ORL1 system has been implicated in a wide variety of physiological processes (Darland et al., 1998). Nociceptin is one of three putative peptide products of the prepronociceptin gene (Houtani et al., 1996; Okuda-Ashitaka et al., 1998) that have been shown to produce behavioural effects in mice (Florin et al., 1997; Okuda-Ashitaka et al., 1998; Rossi et al., 1998). Intriguingly, one of these other prepronociceptin peptides, nocistatin (rat prepronociceptin116–132), was reported to reverse the effects of nociceptin both in vivo and in vitro (Nicol et al., 1998; Okuda-Ashitaka et al., 1998). The cellular basis for these effects is not established, but nocistatin does not appear to directly interact with ORL1 (Okuda-Ashitaka et al., 1998).

Detailed investigations of the role of nociceptin have been hampered by the lack of effective antagonists for the ORL1 receptor. A synthetic analogue of nociceptin, [Phe1ψ(CH2-NH)Gly2]-nociceptin-(1–13)NH2 (Phe1ψ-nociceptin1–13), was reported to antagonize the inhibitory effects of nociceptin on contractions of the guinea-pig ileum and mouse vas deferens (Guerrini et al., 1998), and to be devoid of significant agonist activity (Calo et al., 1998a; Guerrini et al., 1998; Meis & Pape, 1998). However, subsequent studies have shown that Phe1ψ-nociceptin1–13 is a potent agonist at recombinant ORL1 receptors (Butour et al., 1998; Okawa et al., 1999) and that it mimics the inhibitory effects of nociceptin on nociceptive responses in rats (Carpenter & Dickenson, 1998; Xu et al., 1998), as well as the nociceptin reversal of morphine-induced supra-spinal analgesia in mice (Calo et al., 1998b; Grisel et al., 1998). Phe1ψ-nociceptin1–13 acts as a partial agonist with respect to nociceptin in assays of noradrenaline release in mouse brain (Schlicker et al., 1998), substance P release in guinea-pig airway (Shah et al., 1998), and activation of K+ channels (IK) in suprachiasmatic nucleus neurons (Allen et al., 1999).

We have previously shown that nociceptin activates an inwardly rectifying potassium conductance in rat locus coeruleus (LC) neurons (Connor et al., 1996). In this study we have compared the effects of nociceptin and Phe1ψ-nociceptin1–13 on calcium and potassium channels in locus coeruleus neurons, as well as examining the effects of several other putative prepronociceptin products on LC neurons.

Methods

Sprague-Dawley rats of either sex (post natal days 19–20, for slice experiments; post natal days 28–35 for dissociated cell experiments) were used for this study. The rats were anaesthetized with halothane and then killed by cervical dislocation. Horizontal slices (250 μm thick for slice experiments; 290–310 μM thick for preparation of dissociated cells) containing the LC were cut with a vibratome in ice cold physiological saline (ACSF) of composition (mM) NaCl 126, KCl 2.5, MgCl2 1.2, CaCl2 2.4, NaH2PO4 1.2, NaHCO3 24 and glucose 11; gassed with 95% O2/5% CO2. For recordings of IK, brain slices were placed in a chamber (1.5 ml volume) mounted on the stage of an upright microscope (Olympus BH-2 with a fixed-stage modification) and viewed using a water immersion objective (Zeiss, ×40). Slices were continuously superfused (2 ml min−1) with ACSF (32°C). Neurons located in the LC were visualized using infra-red Nomarski optics and recordings of IK made using standard whole cell voltage clamp techniques. Recordings were made with borosilicate pipettes of resistance 3–7 MΩ when filled with intracellular solution of the following composition (mM): potassium gluconate 140; NaCl 15; MgCl2 1; HEPES 10; EGTA 11; MgATP 2; Na2GTP 0.25; adjusted to a pH of 7.3 with KOH. Neurons were voltage clamped at −60 mV (Axopatch 1D, Axon Instruments, Foster City, CA, U.S.A.), series resistance (<12 MΩ) was compensated by 80% and continuously monitored during experiments. Currents were sampled at 50 Hz for subsequent analysis (Axograph 4.0, Axon Instruments). Liquid junction potentials of −11 mV were corrected. Drugs were applied to the slice by changing the perfusion buffer to one that differed only in the content of drug.

For recordings of currents through calcium channels, cells were dissociated using procedures based on those outlined in Ingram et al. (1997). After a 30 min incubation in ACSF (35°C) slices were transferred to a dissociation buffer of composition (mM) Na2SO4 82, K2SO4 30, HEPES 10, MgCl2 5, glucose 10, containing 20 units ml−1 papain, pH 7.3 and incubated for 2 min at 35°C. The slices were then placed in fresh dissociation buffer containing 1 mg ml−1 bovine serum albumin (BSA) and 1 mg ml−1 trypsin inhibitor. The LC region was subdissected from each slice with a fine tungsten wire and the cells dissociated from the slices by very gentle trituration in a pasteur pipette with a fire polished tip. The cells were plated onto plastic culture dishes and kept at room temperature in dissociation buffer. Cells remained viable for up to 6 h after dissociation.

Recordings of currents through Ca2+ channels were made using standard whole cell patch clamp techniques (Hamill et al., 1981) at room temperature (22–24°C). Immediately prior to recording, cells were superfused with a buffer of composition (mM) NaCl 140, KCl 2.5, CaCl2 2.5, MgCl2 1.5, HEPES 10, glucose 10, pH 7.3 in order to wash off the dissociation buffer. For calcium channel current recordings, cells were superfused in solution containing (mM) tetraethylammonium chloride 140, BaCl2 2, MgCl2 1, CsCl 2.5, HEPES 10, glucose 10, BSA 0.05%, pH 7.3. Recordings were made with fire polished borosilicate pipettes of resistance approximately 2 MΩ when filled with intracellular solution of the following composition (mM): CsCl 110, MgATP 5, Na2GTP 0.2, EGTA 10, CaCl2 2 and HEPES 10, pH 7.3. The peak calcium channel current in each cell was determined by stepping the membrane potential from a holding potential of −90 mV to potentials between −60 and +60 mV, usually for 30 ms, in 10 mV increments. The test current evoking a peak calcium channel current was then evoked every 30 s, and monitored for at least a further 2 min before drugs were applied. The inhibition of drugs was quantified by measuring the current amplitude isochronically with the peak of the control calcium channel current. Cells in which the calcium channel current declined in the absence of drug treatment were discarded. Whole cell capacitance and series resistance were compensated manually by nulling the capacitive transient evoked by a 20 mV pulse from −90 mV. The series resistance was between 1.5 and 5 MΩ; series resistance compensation of at least 80% was used in all experiments. An approximate value of whole cell capacitance was read from the amplifier capacitance compensation circuit (Axopatch 1D). Leak current was subtracted on line using a P/8 protocol, unless otherwise noted, typically the leak conductance was less than 1 nS. Evoked calcium channel currents were sampled at 5–10 kHz and recorded on hard disk for later analysis. Data was collected and analysed off line with the PCLAMP suite of programs (Axon Instruments). Cells were exposed to drugs via a series of flow pipes positioned above the cells. Drugs were applied after at least 2 min of control currents were collected, subsequent drug applications were made after the effects of the first drug application had fully reversed, or in the case of drug co-applications, 2 min into the application of the first drug. All data are expressed as mean±s.e.mean, unless otherwise indicated. Statistical significance was determined by using an unpaired students t-test unless otherwise stated.

Drugs and chemicals

Buffer salts were from BDH Australia or Sigma Australia. Papain was from Worthington Biochemical Corporation (Freehold, NJ, U.S.A.). BSA and trypsin inhibitor (Type II-O) were from Sigma Australia. Nociceptin (Phe-Gly-Gly-Phe-Thr-Gly-Ala-Arg-Lys-Ser - Ala - Arg - Lys - Leu - Ala-Asn-Gln) was synthesized and purified by Chiron Mimotopes (Clayton, Victoria, Australia). Nocistatin, (rat prepronociceptin125–132, mPNP-3-8P, Glu-Val-Glu-Gln-Lyn-Gln-Leu-Gln) was synthesized and purified by Auspep (Parkville, Victoria, Australia). Rat prepronociceptin154–181 (mouse prepronociceptin160–187) (Phe-Ser-Glu-Phe - Met - Arg - Gln -Tyr-Leu-Val-Leu-Ser-Met-Glu-Ser-Ser-Glu-Arg -Arg -Thr -Leu -His -Gln -Asn -Gly -Asn -Val) was made by solid phase synthesis by Research Genetics, Huntsville, Alabama, U.S.A.). [Phe1ψ(CH2-NH)Gly2]-nociceptin-(1–13)NH2 was a kind gift of Dr G. Calo.

Results

Dissociated LC neurons were identified as large (mean membrane capacitance 32±1 pF, n=86), usually multipolar neurons with somata shapes characteristic of LC neurons described in fixed tissue preparations (e.g. Swanson, 1976). In our dissociations the only cells of comparable size were large, spherical cells with a single small process, which were presumably MeV neurons. When LC neurons were stepped from a holding potential of −90 mV to potentials between −60 and +60 mV the inward currents in most cells began to activate at about −40 mV and were invariably greatest at membrane potentials between −10 and 0 mV. The peak current density did not differ between cells from male (125±7 pA pF−1, n=48) and female rats (116±6 pA pF−1, n=49). The peak inward current could be abolished by Cd2+ (30 μM, data not shown).

Nociceptin inhibited the peak inward IBa in all LC neurones when applied at a concentration of 300 pM or more (n=89). The effects of nociceptin reversed on washout (Figure 1a). A concentration response relationship for nociceptin inhibition of IBa was determined by application of one or more concentrations of nociceptin to cells stepped repetitively from −90 mV to the membrane potential that evoked the largest IBa in each neuron (either −10 or 0 mV). A logistic function fitted to the concentration-response relationship for nociceptin inhibition of IBa, gave a pD2 for nociceptin of 8.9±0.1 with a slope factor the curve of 0.8±0.1 (Figure 1b). The maximum inhibition of IBa by nociceptin was about 50% (Figure 1b).

Figure 1.

Figure 1

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Modulation of LC calcium channel currents by nociceptin. IBa was elicited by repetitively stepping the membrane potential from −90 mV to −10 mV. (a) (i) A time plot of the peak amplitude of IBa illustrating the effects of repeated applications of nociceptin. (ii) Selected traces from the same experiment, showing the inhibition of IBa by nociceptin. (b), Concentration-response relationship for nociceptin (EC50, 2 nM), and Phe1ψ-nociceptin1–13 (EC50, 30 nM) inhibition of IBa in LC neurons. Each point represents at least five cells tested. Also shown is the response of LC neurons to Phe1ψ-nociceptin1–13 applied after application and washout of nociceptin (1–30 nM) (n=5 for 100 nM, n=9 for 1 μM Phe1ψ-nociceptin1–13).

The inhibition of IBa by nociceptin was evident at a wide range of membrane potentials (Figure 2a) and was associated with a pronounced slowing of the activation of IBa (Figures 1a and 2b). When applied at a concentration of 3 nM, nociceptin increased the 0–95% risetime of IBa from 2.9±0.2 ms to 5.9±0.8 ms (P<0.003, paired t-test, n=9), the slowing reversed on washout of nociceptin (risetime was 2.8±0.2 ms after wash P>0.7, paired t-test versus predrug risetime).

Figure 2.

Figure 2

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Characteristics of nociceptin modulation of IBa. IBa were elicited by stepping the membrane potential from −90 mV to potentials between −60 and +60 mV in 10 mV increments. Nociceptin inhibited IBa over a range of membrane potentials. (a) A plot of the peak inward current at each test potential, before, during and after an application of nociceptin, 100 nM. This cell is a typical example of six experiments. (b) Families of traces from the same experiment, showing the IBa elicited by the steps from a holding potential of −90 mV to a test potential between −60 and +60 (i) in the absence of drug, (ii) in the presence of nociceptin and (iii) following wash of nociceptin.

The inhibition of IBa by nociceptin could be attenuated by a strong positive depolarizing step shortly before the test step. In the experiments illustrated in Figure 3a, LC steps were stepped twice to −10 mV, with an 80 ms depolarizing step to +80 mV between the test steps. These experiments were performed without leak subtraction. In control conditions, the amplitudes of the first (T1) and second (T2) test step did not differ from each other (P>0.8, unpaired t-test), the ratio of T2 : T1 was 0.97±0.02 (n=9). However, in the presence of nociceptin (100 nM), the amplitudes of the first step and the second step were significantly different from each other (P<0.006, unpaired t-test), the ratio of T2 : T1 was 1.6±0.05 (n=9). In the presence of nociceptin the amplitude of the first step was reduced by 52±2% (P<0.0002, paired t-test) compared to the first test step in the absence of drug, while the amplitude of the second test step was reduced by only 21±1% (P<0.0002, paired t-test) compared to second test step in the absence of nociceptin. The inhibition of the first test pulse (T1) by nociceptin was significantly greater (P<0.002) than the inhibition of the test pulse (T2) following the 80 ms depolarization to +80 mV.

Figure 3.

Figure 3

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Nociceptin inhibition of IBa is relieved by a positive prepulse. An LC neuron was voltage clamped at −90 mV and stepped twice to a test potential of −10 mV, with 90 ms between the test pulses. In (a), and 80 ms positive step to +80 mV was applied to the cell immediately after the first test pulse. In (b), the cell was held at −90 mV for the 90 ms between test pulses. The resulting raw current traces for steps in the absence of drug and in the presence of nociceptin are shown. Traces in (a) and (b) are from two separate applications of nociceptin about 10 min apart. The dashed line represents the zero current line, because of the complex step paradigm leak subtraction was not used. The break in the current traces represent a section of about 50 ms that has been omitted for clarity. Note that in (a), the amplitude of the second step is facilitated compared with the first, while in (b) there is no facilitation. Note also that in (a), nociceptin inhibits the outward current through the calcium channels at +80 mV. The experiment in (a) is typical of nine cells, the experiment in (b) of five cells.

While the step to +80 mV partly reversed the nociceptin inhibition of the amplitude of IBa, it completely reversed the kinetic slowing of IBa activation caused by nociceptin. The 0–95% risetime of the first step to −10 mV (T1) was 2.7±0.1 ms, in the presence of nociceptin (100 nM) the risetime was 4.9±0.3 ms (P<0.0002, paired t-test, n=9). The 0–95% risetime of the test step (T2) after the depolarizing step was 2.9±0.1 ms, in the presence of nociceptin the risetime was also 2.9±0.1 ms, which is not different from that in the absence of drug (P>0.8, paired t-test, n=9).

When there was no depolarizing step between the two test pulses, there was not relief from the effect of nociceptin on the amplitude or kinetics of IBa (Figure 3b). When cells were stepped twice to −10 mV, separated by 90 ms at the holding potential of −90 mV, the amplitudes of the first (T1) and second (T2) test step did not differ from each other (P>0.8, unpaired t-test), the ratio of T2 : T1 was 0.95±0.01 (n=5). In the presence of nociceptin (100 nM), the amplitude of the first test pulse was reduced by 47±4% (P<0.03, paired t-test, n=5) and the amplitude of the second test pulse was reduced by 45±4% (P<0.04, paired t-test). In the presence of nociceptin the ratio of T2 : T1 was 0.98±0.1 (n=5). The 0–95% risetime of the first test step to −10 mV was 2.6±0.1 ms, in the presence of nociceptin (100 nM) the risetime was 4.2±0.5 ms (P<0.02, paired t-test, n=5). The 0–95% risetime of the second test step was 2.7±0.1 ms, in the presence of nociceptin the risetime was 4.9± 0.5 ms (P<0.002, n=5).

When applied to LC neurons before nociceptin, Phe1ψ-nociceptin1–13 inhibited IBa in a concentration dependent manner, however, unlike the effects of nociceptin, the inhibition of IBa by Phe1ψ-nociceptin1–13 did not readily reverse on washout (Figure 4b). Equations describing concentration response relationships for agonists assume agonist/effector reactions are freely reversible, and the interaction of Phe1ψ-nociceptin1–13 with IBa is apparently not. However, a logistic function was fitted to the concentration response data from cells in which Phe1ψ-nociceptin1–13 was applied first, to obtain a notional pD2 of 7.6±0.2, with a maximum inhibition of about 18% (Figure 1b). In cells where nociceptin (1–30 nM) had been applied before Phe1ψ-nociceptin1–13, the agonist effects of Phe1ψ-nociceptin1–13 on IBa were strongly attenuated (Figure 1b).

Figure 4.

Figure 4

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Phe1ψ-nociceptin1–13 but not nocistatin blocks the effects of nociceptin on IBa. IBa was elicited by repetitively stepping the membrane potential from −90 mV to 0 mV in each cell. (a) A time plot of the peak amplitude of IBa illustrating the effects of nociceptin, followed by the application of the Phe1ψ-nociceptin1–13, which has little effect on IBa itself but antagonizes the effects of a subsequent co-application of nociceptin (3 and 30 nM). (b) A time plot of the peak amplitude of IBa illustrating the effects of nociceptin and ME, followed by an application of Phe1ψ-nociceptin1–13, which has little effect on a subsequent co-application of ME. (c) A time plot of the peak amplitude of IBa illustrating the effect of nocistatin, followed by a co-application of nociceptin and nocistatin.

When LC cells were stepped twice to −10 mV, with an 80 ms depolarizing step to +80 mV between the test steps, application of Phe1ψ-nociceptin1–13 (3 μM) inhibited the amplitude of first step (T1) by 24±9%, (P<0.05, paired t-test, n=7) but did not significantly inhibit the amplitude of the second step (T2, inhibition was 10±5%, P=0.08, paired t-test, n=7). In these cells the ratio of T2 : T1 before the application of Phe1ψ-nociceptin1–13 (3 μM) was 1.01±0.04, during the application of Phe1ψ-nociceptin1–13 (3 μM) the ratio was 1.25±0.09 (P=0.06 paired t-test, n=7).

Superfusion of Phe1ψ-nociceptin1–13 occluded the effects of a co-application of nociceptin (3–30 nM nociceptin, n=9, Figure 4a). In the presence of Phe1ψ-nociceptin1–13 (1 μM), IBa was inhibited by 7±2%, application of nociceptin (3 nM) in the continued presence of Phe1ψ-nociceptin1–13 (1 μM) did not inhibit IBa any further (inhibition was 1±1%, n=5, Figure 4a). In parallel control experiments nociceptin inhibited IBa by 26±4% (P<0.01 vs total inhibition of IBa by Phe1ψ-nociceptin1–13 (1 μM)+nociceptin (3 nM), which was 8±3%). Because of the partial agonist-like actions of Phe1ψ-nociceptin1–13 in LC cells, we did not systematically examine the potency of Phe1ψ-nociceptin1–13 to inhibit the actions of nociceptin. Application of Phe1ψ-nociceptin1–13 also partly occluded the inhibition of IBa by a maximally effective concentration of the opioid agonist methionine-enkephalin (ME, Figure 4b). ME alone (10 μM) inhibited IBa by 47±2% (n=7); in the presence of Phe1ψ-nociceptin1–13 (1 μM) and ME (10 μM) the inhibition of IBa was also 47±2% (n=6). In these experiments Phe1ψ-nociceptin1–13 (1 μM) alone inhibited IBa by 12±2% and ME inhibited the remaining current by 39±2% (significantly less than control, P<0.02).

Nociceptin increases IK in LC neurons in brain slices (Connor et al., 1996). Phe1ψ-nociceptin1–13 also produced a reversible, concentration-dependent outward current when applied to LC neurons voltage clamped at −60 mV in slices (Figure 5). The current activated by Phe1ψ-nociceptin1–13 was examined by determining the steady state current-voltage relationships for LC neurons in the presence and absence of Phe1ψ-nociceptin1–13. The current activated by Phe1ψ-nociceptin1–13 reversed polarity at −112±4 mV (n=5), and was accompanied by an increase in membrane conductance. The current activated by Phe1ψ-nociceptin1–13 showed pronounced inward rectification. The cord conductance of the Phe1ψ-nociceptin1–13-activated current measured at −60 mV was 0.8±0.3 nS, the cord conductance measured at −130 mV was 2.1±0.5 nS (n=5). The conductance activated by a high concentration of ME (10 μM) in the same cells reversed polarity at −116±7 mV (n=5). The cord conductance of the ME-activated current measured at −60 mV was 2.1±0.3 nS, and 4.7±0.6 nS when measured at −130 mV (n=5).

Figure 5.

Figure 5

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Modulation of LC IK by Phe1ψ-nociceptin1–13. LC neurons in slices were voltage clamped at −60 mV. (a) A continuous record of the membrane currents caused by superfusion of ME and Phe1ψ-nociceptin1–13. (b) Concentration-response relationship for Phe1ψ-nociceptin1–13 (EC50 75 nM) activation of IK in LC neurons, plotted as a percentage of the current resulting from the application of a high concentration of ME (10 μM) in each cell. Each point represents between four and eight cells tested. (c) A continuous record of the membrane currents caused by nociceptin, including a co-application of Phe1ψ-nociceptin1–13, which partially antagonizes the effects of nociceptin.

A logistic function fitted to the concentration-response relationship for Phe1ψ-nociceptin1–13 activation of IK gave a pD2 of 7.10±0.05 with a slope factor for the curve of 1.8±0.05 (Figure 5b). The highest concentrations of Phe1ψ-nociceptin1–13 examined caused an outward current that was about 50% of that caused by application of a high concentration of ME (10 μM, Figure 5). We have previously shown that nociceptin produces a maximal current similar to that caused by high concentrations of ME.

Application of Phe1ψ-nociceptin1–13 (1 μM) in the continued presence of a submaximally effective concentration of nociceptin (300 nM) reversibly reduced the outward current caused by nociceptin to 70±7% of the pre-Phe1ψ-nociceptin1–13 value (n=7, Figure 5c).

Application of nocistatin (10 μM) did not affect IBa in any LC neuron tested (Figure 4c, n=8), nor did it affect the inhibition of IBa by a subsequent co-application of nociceptin. The inhibition of IBa produced by nociceptin (3 nM) in the presence of nocistatin (10 μM) was similar to that produced in the absence of nocistatin (29±3%, n=7, versus 27±4%, n=6, respectively). Application of nocistatin (1 μM) did not change the membrane current or conductance of LC neurons voltage clamped at −60 mV in slices (n=5).

Application of rat prepronociceptin154–181 (1 μM) did not affect IBa in any LC neuron tested (IBa was 100±1% of control after 2 min in prepronociceptin154–181, n=8), nor did it affect the inhibition of IBa by a subsequent co-application of nociceptin. The inhibition of IBa produced by nociceptin (3 nM) in the presence of prepronociceptin154–181 (1 μM) was similar to that produced in the absence of prepronociceptin154–181 (20±3%, n=5, versus 22±5%, n=4, respectively). Application of prepronociceptin154–181 (1 μM) did not change the membrane current or conductance of LC neurons voltage clamped at −60 mV in slices (n=5).

Discussion

This study demonstrates that nociceptin, the endogenous ligand for the ORL1 receptor, inhibits IBa in all rat locus coeruleus neurons. Nociceptin has been previously shown to increase IK in the same neurons (Connor et al., 1996). The functional significance of modulation of IBa in LC neurons is not established, however, nociceptin is known to reduce noradrenaline release from mouse brain cortex in vitro, which presumably reflects inhibitory actions of nociceptin on the nerve terminals of LC cells (Schlicker et al., 1998). It is possible that this inhibition of noradrenaline release occurs via inhibition of LC nerve terminal calcium channels, although opening of nerve terminal (or somatic) K+ channels would be expected to have a similar inhibitory effect. The potency for nociceptin inhibition of noradrenaline release (about 30 nM, Schlicker et al., 1998) is between that for nociceptin inhibition of IBa (2 nM), and activation of IK (90 nM, Connor et al., 1996). The finding that nociceptin was considerably more potent at inhibiting IBa in dissociated LC cells than in activating IK in LC cells in slices is similar to previous findings in PAG neurons, where nociceptin inhibited IBa in dissociated cells with an EC50 of 5 nM (Connor & Christie, 1998), but activated IK with an EC50 of 42 nM (Vaughan et al., 1997). These differences in potency may reflect stronger coupling of ORL1 receptor to IBa than IK, as previously demonstrated for the μ-opioid receptor in acutely isolated neonatal LC neurons (Ingram et al., 1997). Alternatively, these differences could be due to degradation of nociceptin in slice preparations. It would have been of interest to determine the effects of nociceptin on IBa in LC slices, however the size and geometry of LC neurons make it impossible to achieve a semblance of a voltage clamp of IBa in these cells (M. Connor and M.J. Christie unpublished observations 1987–1999).

The inhibition of IBa in LC by nociceptin was probably mediated by activation of heterotrimeric guanine nucleotide binding proteins (G proteins), as has been demonstrated for nociceptin inhibition of high voltage activated IBa in hippocampus (Knoflach et al., 1996), periaqueductal grey (Connor & Christie, 1998) and sensory neurons (Abdulla & Smith, 1997). In the present study the inhibition of IBa by nociceptin was rapid, reversible, evident across a range of membrane potentials and was associated with a pronounced slowing of the activation of the currents, all characteristic features of the ubiquitous G protein βγ-subunit mediated pathway for inhibition of IBa (Herlitze et al., 1996; Ikeda, 1996). Further, the nociceptin-induced inhibition of the amplitude of IBa could be significantly reversed by a depolarizing prepulse to +80 mV. The relief of inhibition by the depolarizing prepulse is thought to reflect a voltage-dependent dissociation of G protein βγ subunits from the calcium channels (Herlitze et al., 1996; Ikeda, 1996; Zamponi & Snutch, 1998). Intriguingly, in contrast to the incomplete reversal of the nociceptin-induced inhibition of IBa amplitude, the depolarizing prepulse completely reversed the nociceptin-induced kinetic slowing of IBa. Although the reason for this is not known, the observation suggests that there may be a voltage-independent component of nociceptin modulation of IBa in LC neurons. It should be noted that the present study utilized Ba2+ as a charge carrier and strong intracellular Ca2+ buffering to maintain stable calcium channel currents. Thus, any modulation of IBa by Ca2+-dependent process would be likely to be suppressed in the present experiments, and it is possible that nociceptin may act via such additional mechanisms to modulate IBa in vivo.

The peptide analogue of nociceptin, Phe1ψ-nociceptin1–13, appears to act as a partial agonist both at inhibiting IBa and activating IK. The maximal effect of Phe1ψ-nociceptin1–13 was less than that of nociceptin on both conductances (see Connor et al., 1996, for comparison of nociceptin and ME on IK) while high concentrations of Phe1ψ-nociceptin1–13 occluded the effects of a co-application of nociceptin. Previous studies have reported Phe1ψ-nociceptin1–13 to be either a pure antagonist (Bigoni et al., 1999; Guerrini et al., 1998; Meis & Pape, 1998), a partial agonist (Allen et al., 1999; Bigoni et al., 1999; Okawa et al., 1999; Schlicker et al., 1998; Shah et al., 1998) or a full agonist (Butour et al., 1998; Calo et al., 1998b; Grisel et al., 1998; Okawa et al., 1999; Xu et al., 1998) in various assays of putative ORL1 function. The agonist activity of Phe1ψ-nociceptin1–13 appears to be strongest in whole animals experiments (Calo et al., 1998b; Grisel et al., 1998; Kapusta et al., 1999; Xu et al., 1998) or cells overexpressing ORL1 (Butour et al., 1998; Okawa et al., 1999).

The most parsimonious explanation of the different actions of Phe1ψ-nociceptin1–13 in various bioassays is that the receptor/effector coupling for nociceptin/ORL1 differs in various parts of the nervous system. In a cell line where ORL1 is a heterologously expressed to a high level, Phe1ψ-nociceptin1–13 is full agonist, as may be expected in a situation when coupling efficiency is artificially high. Further, agents that are partial agonists in in vitro bioassays, such as morphine, (e.g. Alt et al., 1998; Lemaire et al., 1978; Ingram et al., 1997) can demonstrate full agonist activity in in vivo assays of more complex functions such as analgesia, presumably because of the larger receptor reserve apparently available in vivo (e.g. Adams et al., 1990; Mjanger & Yaksh, 1990).

The perceived lack of agonist activity of Phe1ψ-nociceptin1–13 in peripheral assays of nociceptin action has also been suggested to reflect a heterogeneity of nociceptin receptors, with Phe1ψ-nociceptin1–13 only being an agonist at those in the central nervous system (e.g. Butour et al., 1998; Calo et al., 1998b). Although a potential molecular basis for ORL1 receptor heterogeneity has been established with the identification of a number of splice variants of the receptor, there is no evidence that these receptors are functionally different (Pan et al., 1998; Wang et al., 1994). Studies of nociceptin receptor binding in rodent brain have generally found only one high affinity nociceptin binding site (Foddi & Mennini, 1997; Makman et al., 1997; Albrecht et al., 1998; Varani et al., 1998, but see Mathis et al., 1997), although direct comparisons between central and peripheral nervous system nociceptin binding have not been performed.

The high activity of Phe1ψ-nociceptin1–13 in whole animal studies has led to the suggestion that it may be converted into an active compound in vivo (Kapusta et al., 1999). The studies reported here on dissociated cells demonstrate that Phe1ψ-nociceptin1–13 retains agonist activity in a situation where metabolism is unlikely.

The effects of Phe1ψ-nociceptin1–13 were reversible in slices, but did not reverse within the timecourse of experiments on IBa. The reason for this is unknown, but it could reflect either a very slow dissociation rate of Phe1ψ-nociceptin1–13 from ORL1 receptors under conditions used to record IBa, or a non-specific interaction of Phe1ψ-nociceptin1–13 with either IBa or the G-proteins which presumably transduce the signal between ORL1 and IBa. The effects of Phe1ψ-nociceptin1–13 on IBa were probably mediated via a G protein dependent mechanism, because Phe1ψ-nociceptin1–13 inhibition of IBa could be partially relieved by a depolarizing step to +80 mV, similar to the effects of nociceptin itself. Phe1ψ-nociceptin1–13 did not prevent the inhibition of IBa by maximally effective concentration of ME (M. Connor, unpublished observations 1999), although the inhibition of IBa by ME was partly occluded by Phe1ψ-nociceptin1–13. The lack of additivity between ME and Phe1ψ-nociceptin1–13 likely reflects the fact that IBa in LC neurons can only be inhibited to a maximum of about 50% (e.g. by high concentrations of nociceptin, as reported above).

The precursor polypeptide for nociceptin has been suggested to encode up to four additional potential peptides (Houtani et al., 1996; Mollereau et al., 1996; Nothacker et al., 1996). The 28 amino acids carboxy-terminal to the nociceptin sequence in the precursor are strictly conserved across mouse (prepronociceptin160–187), rat (prepronociceptin154–181), human and cow (Houtani et al., 1996; Mollereau et al., 1996; Nothacker et al., 1996; Okuda-Ashitaka et al., 1998). The carboxy-terminal peptide appears to be a necessary product of nociceptin production. Immunoreactivity for the octacosapeptide has been detected in mouse hypothalamus and amygdala (R.G. Allen, manuscript in preparation) and a fragment comprising the first 17 amino acids of this peptide has been shown to both stimulate locomotion (NocII, Florin et al., 1997) and produce naloxone-sensitive analgesia (orphanin FQ2, Rossi et al., 1998) in mice. There is no direct evidence that the 17 amino fragment of the C-terminal octacosapeptide is produced in vivo, so we examined the effects of the complete peptide on LC neurons. The peptide had no direct effects on LC IBa or the membrane properties of LC neurons in slices, and it did not occlude the effects of nociceptin. These results are consistent with previously reported lack of effects of the octacosapeptide or its 17 amino acid fragment on nociceptin binding to ORL1 (Nothacker et al., 1996). It is not known if rat prepronociceptin154–181 is released in the region of the LC, nor is there any information as to the location or nature of the receptors with which it interacts.

The other product of the nociceptin precursor that has been shown to have biological activity has been named nocistatin, because it prevents or reverses the activity of nociceptin in several in vivo and in vitro assays (e.g. Minami et al., 1998; Nicol et al., 1998; Okuda-Ashitaka et al., 1998). The nocistatin sequence is not as clearly conserved across species as nociceptin or the C-terminal octacosapeptide, however, a minimal core sequence required for nocistatin activity has been identified, Glu-Gln-Lys-Gln-Leu-Gln. In this study we used an extended octapeptide sequence conserved in rat and mouse, Glu-Val-Glu-Gln-Lys-Gln-Leu-Gln (rat prepronociceptin125–132, mouse prepronociceptin131–138, Okuda-Ashitaka et al., 1998). Nocistatin alone had no effect on LC neurons, nor did it prevent the effects of a subsequent co-application of nociceptin. These findings suggest that, perhaps not unexpectedly, the interactions of nociceptin and nocistatin may occur at an inter-neuronal level, rather than at the level of a single cell. Although there appears to be a high affinity binding site for nocistatin in brain, nothing is known about its localization, pharmacology or molecular identity (Okuda-Ashitaka et al., 1998).

This study demonstrates that nociceptin potently inhibits IBa in all LC neurons, which provides a further possible mechanisms for nociceptin modulation of noradrenaline release in brain (Connor et al., 1996; Schlicker et al., 1998). The putative nociceptin receptor antagonist Phe1ψ-nociceptin1–13 is a partial agonist at inhibiting IBa and opening IK in the LC, which makes it of limited use in investigating the function of ORL1 in this brain region. Nocistatin and rat prepronociceptin154–181 had no direct effects on LC neurons, suggesting that the receptors for these putative neuropeptides are either not located on LC neurons, or that the receptors couple to changes in cellular excitability via mechanisms not readily detectable using conventional electrophysiological techniques. The ORL1/nociceptin system is likely to be an important regulator of a range of functions within the LC and its projection fields.

Acknowledgments

This study was supported by the University of Sydney Medical Foundation. M. Connor was the recipient of a Rolf Edgar Lake Fellowship from the Faculty of Medicine, University of Sydney. C.W. Vaughan is the recipient of an R.D. Wright Fellowship from the National Health and Medical Research Council of Australia. E.A. Jennings was supported by The Wellcome Trust. R.G. Allen was supported by NIH NIDA Grant #11282. We thank Dr G. Calo for his kind gift of [Phe1ψ(CH2-NH)Gly2]-nociceptin-(1–13)NH2.

Abbreviations

ACSF

physiological saline

BSA

bovine serum albumin

CHO

chinese hamster ovary cells

G-protein

heterotrimeric guanine nucleotide-binding protein

IBa

calcium channel current

IK

potassium current

LC

locus coeruleus

ME

methionine enkephalin

ORL1

opioid receptor-like protein

Phe1ψ-nociceptin1–13

[Phe1ψ(CH2-NH)Gly2]-nociceptin-(1–13)NH2

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