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1Discovery Neuroscience and 2Chemical and Screening Sciences, Wyeth Research, Princeton, New Jersey
Submitted 10 August 2006; accepted in final form 18 March 2007
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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, 2002). Local injection of loperamide and its analogs resulted in antipruritic activity in a mouse model of itch, and caused potent antinociception, inhibiting late phase formalin-induced flinching (DeHaven-Hudkins et al. 2002). However, in the formalin test in rats, 10-fold higher doses of naloxone administered subcutaneously were needed to antagonize loperamide compared with the doses needed to antagonize morphine when the agonists were administered subcutaneously, suggesting that the effects of loperamide might be mediated in part by receptors different from those that mediate the effects of morphine (Shannon and Lutz 2002). In a thermal injury–induced rat model, loperamide was able to block thermal hyperalgesia in morphine-tolerant rats, indicating a nonopioid mechanism of action (Nozaki-Taguchi and Yaksh 1999). Loperamide-induced analgesia could be explained, at least in part, by its inhibitory action on ion channels involved in nociception. Loperamide is known to block several types of voltage-gated ion channels, including L-type Ca2+ channels (Church et al. 1994; Hagiwara et al. 2003; Reynolds et al. 1984), delayed-rectifier potassium channels (Yang et al. 2005), and N-methyl-D-aspartate (NMDA) receptors (Church et al. 1994).
In an effort to expand our knowledge of hyperpolarization-activated cyclic nucleotide–gated (HCN) channel pharmacology, we screened a focused compound library composed of 175 known ion channel modulators (Wyeth Research, unpublished data). We found that loperamide produced a potent block of HCN channels at low micromolar concentrations. This observation was particularly interesting in the light of the involvement of HCN channels in regulating neuronal excitability, sensory processing, and the pathophysiology of pain (Chaplan et al. 2003; Hutcheon and Yarom 2000; Pape 1996; Yao et al. 2003). Indeed, prostaglandin E2–induced depolarization of membrane potential in dorsal root ganglion (DRG) neurons and dorsal horn neurons of the spinal cord involves cAMP-dependent induction of Ih (Baba et al. 2001; Ingram and Williams 1994). Positive regulation of Ih by prostaglandins produced during inflammation may lead to membrane depolarization and facilitation of repetitive activity, thus contributing to the sensitization to painful stimuli. Additionally, HCN channels were shown to drive the frequency of ectopic discharges in A
- and C-fibers that is commonly associated with mechanical allodynia in rat models of neuropathic and postoperative pain. Peripheral administration of the HCN channel blocker ZD7288 significantly attenuated mechanical allodynia induced by partial sciatic nerve injury and hind-paw incision (Chaplan et al. 2003; Dalle and Eisenach 2005; Yao et al. 2003).
Thus considering the important role of HCN channels in nociception and pathological pain, the observed loperamide-induced inhibition of HCN channels could provide an additional molecular mechanism for its analgesic action. To further characterize this novel finding, we studied loperamide pharmacology on Ih in rat DRG neurons.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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All protocols involving animals were in accordance with National Institute of Health guidelines and approved by the Wyeth IACUS. Two- to 3-wk-old Wistar rats were anesthetized with halothane before death by decapitation. L4–L6 DRGs were quickly dissected from animals and incubated in DMEM (Sigma) buffer containing 0.5 mg/ml collagenase (Worthington, Lakewood, NJ) and 0.5 mg/ml trypsin for 25–30 min at 35°C. Subsequently, ganglia were washed for 10–20 min and dissociated by triturating with a fire-polished Pasteur pipette. Isolated neurons were plated on poly-D-lysine (Sigma)–coated glass coverslips and cultured at 37°C in a humidified 5% CO2/air atmosphere in serum-free Neurobasal/B27 medium (Invitrogen, Carlsbad, CA).
Electrophysiology
Ih was recorded from the soma of visually identified small (22–30 µm), medium (32–40 µM), and large (50–65 µM soma diam) DRG neurons at 3–10 days in culture using standard whole cell recordings. Electrodes were pulled from borosilicate glass capillaries (TW150F, World Precision Instruments, Sarasota, FL). Pipette resistance ranged between 2–3 (medium-to-large neurons) and 3–4 M
(small neurons) when filled with the intracellular solution (in mM): 100 K-gluconate, 30 KCl, 1.5 MgCl2, 10 HEPES, 3 K2-ATP, and 0.5 cAMP; pH adjusted to 7.3 with Tris-OH. The series resistance (Rs) ranged between 5 and 11 M and was compensated by 50–70%. Recordings with >25% Rs change were excluded from the analysis. Unless specified otherwise, electrophysiological recordings were made 10–15 min after establishing whole cell configuration to allow saline equilibration. Pipette solution was aliquoted and stored at –20°C; during voltage-clamp recordings the pipette solution was kept on ice. The extracellular solution was HBSS (14025–092, Gibco) (in mM): 137.9 NaCl, 1.3 CaCl2, 0.5 MgCl2, 0.4 MgSO4, 4.2 NaHCO3, 0.3 Na2HPO4, 5.3 KCl, 0.4 KH2PO4, and 5.6 dextrose. Data were collected using a MultiClamp 700A amplifier and digitized using DigiData 1322A and pClamp9 software (all from Molecular Devices, Union City, CA). Current traces were filtered at 1–2 kHz and digitized at 4 kHz. Membrane voltages were not corrected for the +10-mV junction potential between the pipette and bath solutions. Recordings were made at room temperature (21–23°C). All drugs were obtained from Sigma.
Data were analyzed using Clampfit (Molecular devices) and Origin 6.0 (OriginLab, Northampton, MA). Concentration-response data sets were fitted with Hill's equation of the form I/I0 = 1/{1+([C]/IC50)k}, where [C] is the loperamide concentration, IC50 is the loperamide concentration producing 50% block of Ih, and k is Hill's coefficient. The G-V relationships were fitted with a Boltzmann equation of the form G/Gmax = 1/{1+exp[(V – V1/2)/k]}, where V1/2 is voltage for half-maximal activation, and k is the slope coefficient. The percent reduction of Ih steady-state activation was calculated for each cell according to the formula 100 x {1 – ([G/Gmax]loperamide/[G/Gmax]control) } and subsequently averaged for every membrane voltage. Statistical significance was evaluated by Student's t-test using Origin 6.0. Statistical significance is indicated as follows: *P < 0.05; **P < 0.01; and ***P < 0.001. Unless specified otherwise, all data are means ± SE.
Immunofluorescence
Immunofluorescence detection of HCN proteins was performed following previously published procedures (Vasilyev and Barish 2002) with minor modifications. In brief, DRGs were dissected and fresh frozen (–20°C on dry ice) in optimal cutting temperature compound (OCT, Sakura Finetek, Tokyo, Japan); 20-µm horizontal sections were cut on a cryostat (Leica, Nussloch, Germany). Sections were fixed with 4% paraformaldehyde in PBS, pH 7.4, for 20–30 min at 4°C, and rinsed (3 times, 15 min each) in PBS. Sections were permeabilized with 0.03% Triton X-100 (Sigma) in PBS containing 3% BSA and 5% normal goat serum for 1 h at room temperature. DRGs were incubated for 12–14 h at 4°C in primary antibody (Chemicon, Temecula, CA) diluted at appropriate concentrations (2 µg/ml for anti-HCN1 and anti-HCN2 and 4.0 µg/ml for anti-HCN4 antibodies) in PBS with 3% BSA. After rinsing in PBS (3 times, 20 min each), sections were incubated in fluorescein-conjugated goat antirabbit IgG (Zymed, South San Francisco, CA; diluted 1:100 in PBS containing 5% normal goat serum) for 1 h at room temperature. Finally, sections were rinsed in PBS (3 times, 20 min each) and mounted in Vectashield (Vector Laboratories, Burlingame, CA). Controls included omission of primary antibody. Processed sections were visualized with a Zeiss Axiovert 135 microscope under a x32 air objective and imaged using a Zeiss AxioCam MRm digital monochrome camera. To facilitate qualitative comparison of immunoreactivity, tissues were stained simultaneously using an identical protocol, and images were acquired using the same AxioCam digital camera parameters.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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We used subunit-selective affinity-purified antisera and immunofluorescence to study HCN subunit immunoreactivity in DRG neurons (Fig. 1A). HCN1 immunoreactivity was highest in large DRG somata sections, present at a significant level in a subset of somata sections of medium size, and was generally low in small somata sections. HCN2 immunofluorescence was more broadly distributed in DRGs. The highest levels of HCN2 immunofluorescence were observed in large somata sections, with significant levels of immunofluorescence present in a certain subpopulation of small to medium somata sections. HCN4 immunoreactivity in DRGs was generally low.
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) while blocking inward-rectifier potassium current. Isolated in this way, Ih was 90% blocked by 100 µM ZD7288 (Fig. 1B) at test voltages positive to –100 mV. An apparent time-independent component was still present; however, it was not sensitive to 100 µM ZD7288, and thus was not included into analysis. A representative recording of Ih from a large-diameter DRG neuron is shown in Fig. 1B. Loperamide inhibits Ih in a concentration-dependent manner
Ih was evoked by hyperpolarizing voltage steps to –90 mV delivered from a holding potential of –50 mV in the absence and presence of sequentially increasing concentrations of loperamide (in µM): 0, 1.2, 3.7, 11, 33, and 100 (Fig. 2, top). Loperamide blocked Ih in a concentration-dependent manner, with >90% of Ih being blocked at 100 µM. Ih steady-state current amplitudes recorded at different loperamide concentrations were normalized to the amplitude of Ih in control conditions, and the averages were plotted as a function of time (Fig. 2, middle) or drug concentration (Fig. 2, bottom). Fitting the concentration-response plots with Hill's equation resulted in IC50 = 4.9 ± 0.6 µM, k = 1.7 ± 0.2 (n = 7) and IC50 = 11.0 ± 0.5 µM, k = 1.6 ± 0.1 (n = 6) for large and small neurons, respectively. The statistical analysis shows that the mean loperamide IC50 in small neurons was significantly different (P < 0.05) from the mean loperamide IC50 in large cells.
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). Thus the relative contribution of opioid receptor–dependent versus opioid receptor–independent components to loperamide-induced Ih inhibition in DRG neurons in vivo needs further study. Extracellular loperamide shifts Ih steady-state activation
The percent inhibition of Ih current amplitude superimposed on the percent reduction of Ih steady-state activation at the corresponding membrane voltages is shown in Fig. 3. Ih current-voltage (I-V) relationships in control and after application of 3.7 µM loperamide were normalized to the maximal current amplitude recorded at a test voltage of –120 mV in control saline (Fig. 3A, bottom). Bath-applied loperamide blocked Ih amplitude in a voltage-dependent manner with a percent inhibition (at 3.7 µM) ranging from 88.7 ± 4.2% at –60 mV to 4.1 ± 8.7% at –120 mV (Fig. 3, A and B) and was accompanied by an apparent deceleration of Ih activation kinetics (Fig. 3A, top). The percent reduction of Ih amplitude and steady-state activation caused by bath application of loperamide was calculated from a single pool of recordings, and these averages were plotted against the corresponding test voltages (Fig. 3B). The loperamide-induced reduction of Ih amplitude was generally equivalent to the reduction of Ih steady-state activation (see METHODS for details) at all studied membrane voltages (Fig. 3B; P > 0.05 at all membrane voltages).
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50% inhibition of Ih) were calculated from tail current amplitudes measured 50–70 ms after repolarization (Fig. 4, top, inset), because deactivation of Ih is relatively slow. The conductance ratios G/Gmax show that loperamide shifted the threshold for Ih activation about –10 to –15 mV (–50 mV in control vs. –60 to –65 mV in the presence of 3.7 µM loperamide, respectively) as presented in Fig. 4 (bottom). The following results of the G-V fitting by the Boltzmann equation were obtained: V1/2 = –73.2 ± 0.8 mV, k = 8.5 ± 0.7 mV and V1/2 = –83.3 ± 0.4 mV, k = 7.9 ± 0.4 mV in control and after application of 3.7 µM loperamide, respectively. Bath-applied loperamide (3.7 µM) produced a statistically significant shift in the average V1/2 (–10 mV, P < 0.001) of Ih in large neurons and did not significantly change the slope coefficient (P > 0.05).
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fast) and slow (slow) time constants for Ih activation were voltage dependent, becoming progressively faster toward negative voltages (Fig. 5), and differed by almost an order of magnitude (in control at –110 mV, fast was 91.4 ± 10.8 ms and slow was 605.9 ± 105.9 ms; n = 9). These time constants were significantly slower in the presence of loperamide compared with control (in the presence of 3.7 µM loperamide: fast, at –110 mV was 188.7 ± 31.3 ms and slow was 1,131.3 ± 152.2 ms; n = 9). A direct, model-independent comparison of the rate of Ih activation (by measurement of Ih half-activation time) in control and after loperamide exposure confirmed the respective results for the Ih kinetics deceleration obtained using the two-exponential model for Ih activation. A 2.4-fold change (272.5 ± 20.1 ms in control vs. 646.7 ± 77.5 ms in the presence of 3.7 µM loperamide) of Ih half-activation time (at a test voltage of –90 mV) in small neurons was not significantly different from the 2.9-fold change (181.4 ± 21.5 ms in control and 500.7 ± 64.4 ms in the presence of 3.7 µM loperamide) measured in large cells (Fig. 6). To address the molecular mechanism of Ih inhibition and identify the position of the loperamide binding site on the cellular membrane, we studied the efficacy of loperamide applied to the internal versus external side of the membrane. Furthermore, we studied whether the effect was occurring through a cAMP-dependent (Ingram and Williams 1994) or cAMP-independent mechanism.
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If loperamide directly binds to the channel, the loperamide-induced slowing of Ih activation kinetics could be explained by a number of molecular mechanisms, for example, by binding to the intracellular S6 bundle or by interaction with the cAMP-binding domain, two regions closely involved with HCN channel gating. Alternatively, loperamide might act extracellularly by binding to the S1-S2 extracellular loop, which has been shown to be an important determinant of the rate of HCN channel activation. Because binding to sites located in the pore-forming region might result in a use-dependent block, we first elucidated whether loperamide-induced Ih inhibition depends on the application frequency of test pulses.
Ih amplitude was monitored for 6 min using a standard voltage protocol (see Fig. 2). Subsequently, the holding potential was changed from –50 to 0 mV (threshold for Ih activation was about –55 mV, thus at 0 mV holding potential, Ih channels are fully deactivated), and 3.7 µM loperamide was added to the bath saline. During the first 2 min of loperamide application, cells were held at 0 mV, and no test voltages were applied, thus insuring a complete Ih deactivation. Incubation with 3.7 µM loperamide for 2 min resulted in the reduction of Ih amplitude by 40 ± 8% (n = 5), a value similar to that observed with repeated activation (Ih amplitude was reduced by 38 ± 7%, n = 6, with 3.7 µM loperamide; Figs. 2 and 7A). Subsequently re-establishing the voltage-stimulating protocol did not induce any additional use-dependent component of Ih block (Fig. 7A). Control recordings obtained with the same voltage protocol showed no change of Ih in the absence of loperamide (Fig. 7A).
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10 min. Concurrent bath application of 10 µM loperamide produced a rapid and reversible Ih inhibition (Fig. 7B). Additionally, when applied through the patch pipette, loperamide did not show any substantial effect on Ih steady-state activation or activation kinetics as shown by a population analysis of V1/2 and gating kinetics in loperamide-filled versus control cells (Figs. 4 and 5), pointing to the extracellular location of the loperamide binding site. The effect of loperamide on Ih was measured in the presence of naloxone to test for the possible involvement of opioid receptors (Fig. 8). Application of 10 µM loperamide reduced Ih amplitude by 77 ± 5% (n = 6) and was not significantly different from the effect of equimolar loperamide when cells were preincubated for 5 min with 10 µM naloxone (loperamide-induced Ih inhibition in the presence of naloxone was 78 ± 2%; n = 3).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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This study revealed a novel pharmacology for the known analgesic loperamide that might provide an additional molecular mechanism of its analgesic action. Loperamide, originally known as an antidiarrheal drug, later was found to be active in several models of inflammatory, bone cancer, and acute pain (DeHaven-Hudkins et al. 1999, 2002; Menendez et al. 2003, 2005; Nozaki-Taguchi and Yaksh 1999; Sevostianova et al. 2005). This effect seems to be manifested by activating peripheral µ-opioid receptors (Menendez et al. 2003, 2005; Sevostianova et al. 2005; Shannon and Lutz 2002); however, the existence of an additional mechanism is supported by several findings. First, loperamide-induced analgesia could be antagonized with naloxone only at doses 10-fold higher than those needed to antagonize morphine administered by the same route (Sevostianova et al. 2005; Shannon and Lutz 2002); second, a significant antinociceptive effect of loperamide was found both ipsi- and contralateral to the thermal injury paw, with the effect at the contralateral site not being reversible with naloxone (Nozaki-Taguchi and Yaksh 1999); third, loperamide antagonized thermal hyperalgesia in morphine-tolerant rats, indicating a nonopioid mechanism of action (Nozaki-Taguchi and Yaksh 1999). Additionally, only high doses of loperamide were efficacious in reversal of the first phase of formalin-induced acute pain (paw liking and biting), but failed to produce antinociceptive effects in a model of acute thermal pain (Sevostianova et al. 2005).
Loperamide-induced inhibition of voltage-gated calcium channels (Church et al. 1994; Hagiwara et al. 2003; Reynolds et al. 1984) and NMDA receptors (Church et al. 1994) present both in CNS and in sensory fibers (Carlton et al. 1995; Coggeshall and Carlton 1998; Davidson et al. 1997; Kinkelin et al. 2000) is broadly consistent with its analgesic properties, considering the well-documented efficacy of selective antagonists of voltage-gated calcium channels and NMDA receptors in treating inflammatory and neuropathic pain symptoms (Chizh and Headley 2005; McGivern 2006). Our finding of loperamide-induced inhibition of HCN channels is also consistent with HCN channels involvement in the pathophysiology of pain (Chaplan et al. 2003; Hutcheon and Yarom 2000; Pape 1996; Yao et al. 2003). Because PGE2-induced depolarization of the membrane potential in DRG neurons and dorsal horn neurons of the spinal cord involves cAMP-dependent induction of Ih (Baba et al. 2001; Ingram and Williams 1994), the mechanism of lowering pain threshold by prostaglandins released in the area of inflammation is thought to involve, at least in part, activation of HCN channels. Thus opioid inhibition of adenylyl cyclase and subsequent inhibition of Ih might represent a mechanism by which opioids inhibit primary afferent excitability and relieve pain. The latter is supported by the fact that forskolin- and PGE2-induced Ih upregulation is inhibited by opioids acting through µ- and/or -receptors (Ingram and Williams 1994; Svoboda et al. 1999). However, our study showing a direct inhibition of HCN channels by loperamide does not support an exclusive mechanism of loperamide-induced analgesia through opioid receptor-induced Ih inhibition, but is consistent with the possibility of both opioid receptor-dependent and -independent mechanisms. Moreover, additional mechanisms for the peripheral antihyperalgesic action of loperamide might include inhibition of voltage-gated calcium channels in sensory neurons and the inhibition of tetrodotoxin-resistant sodium channels, both shown to be involved in mechanisms of central and peripheral opioid antinociception (Gold and Levine 1996; Schroeder et al. 1991).
Loperamide blocked Ih in a concentration-dependent manner with an IC50 = 4.9 ± 0.6 and 11.0 ± 0.5 µM for large- and small-diameter neurons, respectively. These values are compatible to the reported IC50 of loperamide-induced inhibition of voltage-gated calcium channels (IC50 = 2.5 µM) (Church et al. 1994; Hagiwara et al. 2003; Reynolds et al. 1984) but are an order of magnitude smaller then the IC50 reported for NMDA receptors (IC50 = 73 µM) (Church et al. 1994). The reported efficacious doses of loperamide in vivo were as high as 3–10 mg/kg (DeHaven-Hudkins et al. 1999, 2002), which in principle could result in a high micromolar range of loperamide in plasma. However, the pharmacokinetics of loperamide has not been reported in the previously mentioned studies; thus drawing any conclusions about the efficacious loperamide concentration in blood plasma would be purely speculative. Our finding of loperamide-induced inhibition of Ih channels is consistent with their involvement in the pathophysiology of pain (Chaplan et al. 2003; Hutcheon and Yarom 2000; Pape 1996; Yao et al. 2003).
HCN channels protein and functional expression
Four HCN channel subunits have been identified (Biel et al. 1999; Gauss and Seifert 2000; Kaupp and Seifert 2001; Ludwig et al. 1999; Monteggia et al. 2000; Santoro and Tibbs 1999) and expressed in heterologous systems. The observed immunofluorescence showing somewhat overlapping but independent expression patterns of HCN1 and HCN2 protein suggest the coexistence of different HCN channel isoforms in DRG neurons.
On the functional level, Ih kinetics and pharmacology were close to those reported previously (Cardenas et al. 1999; Scroggs et al. 1994; Yagi and Sumino 1998). The threshold for Ih activation ranged between –55 and –60 mV; voltage for half-maximal activation and the slope coefficient were V1/2 = –73.2 ± 0.8 mV and k = 8.5 ± 0.7 mV, respectively, which was similar to values reported by Cardenas et al. (1999) (V1/2 = –73.3 mV, k = 7.0 mV).
Mechanism of Ih inhibition by loperamide
We showed that the mechanism of loperamide-induced Ih inhibition is unrelated to the activation of opioid receptors and is reversible, voltage-dependent, use-independent, and is associated with a negative shift of V1/2 for Ih steady-state activation. The voltage dependence of Ih activation has been shown to be modulated by forskolin, PGE2, and opioids through a cAMP-dependent mechanism (Ingram and Williams 1994; Svoboda et al. 1999). Opioids had no effect on Ih alone, but were shown to reverse the effect of forskolin on Ih. This effect was antagonized by a broad-spectrum opioid receptor antagonist naloxone (Ingram and Williams 1994). Involvement of opioid receptors in the reported loperamide-induced Ih inhibition is unlikely because we did not observe any substantial Ih inhibition by 1.2 µM loperamide, considering a low-nanomolar affinity of loperamide for the opioid receptors. Additionally, the loperamide effect was not antagonized by naloxone. Therefore we suggest a direct inhibition of HCN channel activity by loperamide, probably by binding to the extracellular region of the channel. Alternatively, lipid-soluble drugs such as loperamide (clogP = 4.9) can bind to the channel site embedded in the lipid bilayer; however, in this scenario, intracellularly applied loperamide should also block Ih.
Slowing the rate of Ih activation by loperamide
The observed shift of Ih steady-state activation accompanied by the slowing of its activation kinetics in the presence of loperamide could be explained, at least in part, based on the preferential block of fast-HCN1 versus slow-gating HCN2-4 homomeric channels and/or heteromeric channels with a slow-gating (HCN2/4) stoichiometry (Biel et al. 1999; Kaupp and Seifert 2001; Ludwig et al. 1999; Santoro and Tibbs 1999; Vasilyev and Barish 2002). This idea is supported by our observation of preferential block of Ih channels in large versus small DRG neurons, considering a slower gating kinetics of Ih in small cells; however, equimolar loperamide reduced Ih activation rate in small and in large neurons to a similar extent. The pharmacology of loperamide on recombinant HCN channels would answer this question in more detail.
The mechanism of Ih activation kinetics slowing by bath-applied loperamide caused by opioid receptor–induced reduction of the intracellular cAMP (Ingram and Williams 1996) is unlikely because the cAMP level in this experiment was clamped by dialysis through the patch pipette (the measurements involved were made 10–15 min after establishing the whole cell configuration, thus allowing time for stabilization of pipette solutions with a saturation level of cAMP and the cytoplasm). Alternatively, loperamide might affect Ih kinetics by binding to domains principally involved in regulating the rate of HCN channel activation. Two regions affecting HCN channel activation kinetics have been identified, one being S1 and S1–S2 and the other being S6–CNBD. The reciprocal replacements of the whole S1 and S1–S2 region between recombinant HCN1 and HCN4 channels affected the activation kinetics about 16- and 3-fold, respectively (Ishii et al. 2001). Thus it is reasonable to suggest that slowing of the Ih activation rate by extracellularly applied loperamide may be caused by its interaction with HCN channel extracellular domain between S1 and S2, an observation supported by our finding that the loperamide binding site seems to be extracellular, located outside of the lipid bilayer. Additionally, the loperamide-induced reduction of Ih activation rate could not be explained in terms of a simple two-state model (by loperamide affecting the forward and backward rate constants) because accounting for the –10 mV shift of V1/2 was not sufficient to explain the loperamide-induced shift (about –20 mV) for the slow and fast time constants determined from a two-exponential model for Ih activation. The latter observation is also consistent with the S1–S2 hypothesis proposed earlier (Ishii et al. 2001) for HCN1 and HCN4 channel gating (2 channels with 2 orders of magnitude difference in their activation rates, yet a similar V1/2). The hypothesis of the loperamide binding site could be explored further with side-directed mutagenesis of the S1–S2 linker region in future studies.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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FOOTNOTES |
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Address for reprint requests and other correspondence: D. Vasilyev, Discovery Neuroscience, Wyeth Research, CN 8000, Princeton, NJ 08543-8000 (E-mail: vasylyd{at}wyeth.com)
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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, mean ± SE, n = 6) recorded from small neurons. A single representative recording from a small cell (
, n = 1) is shown superimposed with the main graph. Arrows represent bath application of cumulative concentrations of loperamide (in µM): 0.04, 1.2, 3.7, 11.1, 33.3, and 100. Bottom: normalized concentration-current relationships obtained from large (
, n = 6) for loperamide. Amplitudes of Ih recorded in the presence of loperamide were normalized to the amplitude of Ih recorded in control saline at the same test voltage. The asterisks represent the level of statistical significance between 2 data sets obtained from small vs. large cells at the respective loperamide concentration.
-slow time constant, n = 9), but were not significantly different from the respective time constants when loperamide was included in the pipette (
, n = 5, P > 0.05 for all membrane voltages). Asterisks indicate level of statistical significance between values of slow and fast time constants in control vs. value of corresponding time constant in presence of loperamide. All recordings were made from large neurons.
