Modulation of opioid analgesia, tolerance and dependence by Gs-coupled, GM1 ganglioside-regulated opioid receptor functions

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Modulation of opioid analgesia, tolerance and dependence by Gs-coupled, GM1 ganglioside-regulated opioid receptor functions Stanley M. Crain and Ke-Fei Shen Studies of direct excitatory effects elicited by opioid agonists on various types of neurone have been confirmed and expanded in numerous laboratories following the initial findings reviewed previously by Stanley Crain and Ke-Fei Shen. However, the critical role of the endogenous glycolipid GM1 ganglioside in regulating Gs-coupled, excitatory opioid receptor functions has not been addressed in any of the recent reviews of opioid stimulatory mechanisms. This article by Stanley Crain and Ke-Fei Shen focuses on crucial evidence that the concentration of GM1 in neurones might, indeed, play a significant role in the modulation of opioid receptor-mediated analgesia, tolerance and dependence.

S. M. Crain, Professor Emeritus, and K-F. Shen, Principal Associate, Department of Neuroscience, Albert Einstein College of Medicine, Yeshiva University, Bronx, NY 10461, USA.

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Gangliosides, defined as sialic acid-containing glycolipids, are abundantly distributed on the surface of most neurones, where they constitute about 3–5% of the phospholipids in the outer half of the membrane bilayer1. Nearly 100 different gangliosides have been characterized and increasing evidence suggests that many of these glycolipids interact selectively with specific membrane protein receptors so as to modulate signal transduction2. Specific gangliosides have been shown to bind to allosteric sites on several growth-factor receptors and thereby regulate the binding of the growth factor to the receptor recognition site as well as coupling to second messenger systems2. This article focuses on the remarkably specific regulatory effects of GM1 ganglioside on excitatory opioid receptor functions. A number of recent reviews have noted our evidence and hypothesis3 that opioid-induced activation of protein kinase A (PKA) via Gs-coupled opioid receptors might play a significant role in counteracting opioid receptor-mediated inhibitory effects4–6. However, emphasis has generally been placed on the roles of other signal transduction systems4–6 that might account for opioid stimulatory effects, such as Gi–Go-mediated increases in cAMP, protein kinase C (PKC) and phospholipases. On the other hand, the studies reviewed

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T below suggest that GM1 regulation of Gs-coupled opioid receptor functions provides a surprisingly effective and hitherto unrecognized mechanism, which could underlie the unusually plastic excitatory activities associated with the development of tolerance and dependence during chronic exposure of nerve cells to opioid agonists. Electrophysiological studies of the effects of opioids on nociceptive types of dorsal root ganglion (DRG) neurones in culture have suggested that Gi–Go-coupled inhibitory effects mediated by opioid receptors (e.g. shortening of the Ca2+-dependent component of the action potential duration and inhibition of transmitter release) provide a useful cellular model of opioid analgesia7. By contrast, Gs-coupled excitatory effects mediated by opioid receptors (e.g. prolongation of the action potential duration3,8–14 and stimulation of transmitter release15), elicited by lower concentrations of morphine and other bimodally acting opioid agonists in these cells, might provide insights into mechanisms underlying opioid hyperalgesia3,8,14 and anti-analgesia9,13,16,17 (Fig. 1). Co-treatment of DRG neurones with agents that selectively block these excitatory, but not inhibitory, opioid receptor functions [such as cholera toxin (CTX)] not only enhances the inhibitory (antinociceptive) potency of morphine or other bimodally acting opioid agonists but also attenuates tolerance or dependence during chronic opioid exposure11–13,18–21.

Interconversion of opioid receptors between inhibitory and excitatory modes depends on GM1 levels In an earlier review based primarily on electrophysiological analyses of the effects of opioids on DRG neurones in culture, Crain and Shen3 suggested that naloxone (NLX)-reversible excitatory effects (i.e. those that prolong the action potential duration), which are directly evoked in these neurones by low (nM) concentrations of opioid agonists, could be accounted for by the activation of putative subtypes of m-, d- and k-opioid receptors that are coupled to the stimulatory, regulatory G protein, Gs. However, no biochemical evidence was available to determine if molecularly distinct Gs-coupled receptors mediate these excitatory opioid effects (see below). Gscoupled, excitatory effects mediated by opioid receptors on DRG neurones are selectively blocked by low concentrations of the A subunit of CTX (Ref. 11) (which ADP-ribosylates Gs), whereas Gi–Go-coupled, inhibitory effects mediated by opioid receptors [generally elicited by higher (mM) opioid concentrations] are selectively blocked by pertussis toxin (PTX) (which ADP-ribosylates Gi and Go)8,22. These results suggest that the activation of Gs-coupled opioid receptors on sensory neurones elicits stimulatory effects via transduction systems mediated by adenylate cyclase (AC), cAMP and PKA, which thereby counteract the inhibitory effects mediated by the activation of Gi–Go-coupled opioid receptors on these cells3,10,22 (Fig. 1). Nevertheless, electrophysiological studies of DRG neurones have not precluded the

Copyright © 1998, Elsevier Science Ltd. All rights reserved. 0165 – 6147/98/$19.00 PII: S0165-6147(98)01241-3

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Fig. 1. GM1 ganglioside-induced interconversion of opioid receptors between the Gi–Go-coupled inhibitory mode (right) and the Gs-coupled excitatory mode (left) in sensory dorsal root ganglion (DRG) neurones. Note the sharply contrasting linkages of these Gi–Go-coupled receptors compared with Gs-coupled receptors to K+ and Ca2+ conductances (gK and gCa), which control action potential duration (APD) and transmitter release in presynaptic terminals of nociceptive neurones involved in opioid analgesic systems3. The positive-feedback phosphorylation cycle involving adenylate cyclase (AC), cAMP, protein kinase A (PCA) and GM1 might mediate sustained sensitization of excitatory, Gscoupled opioid receptor functions, resulting in tolerance and dependence during chronic opioid treatment10. By contrast, inhibitory, Gi–Go-coupled opioid receptor functions might be progressively desensitized by the activation of G protein-coupled receptor kinases (GRK) and arrestins, as well as by activation of protein kinase C (PKC) (see text), thereby contributing to the development of tolerance. Sustained opioid-induced activation of PKC could also increase the activity of Ca2+ channels, providing an additional mechanism for modulating opioid tolerance or dependence4–6,60. CTX-A, CTX-B; A and B subunits of cholera toxin. Modified figure, reproduced, with permission, from Ref. 10.

possibility that opioid receptors might be able to switch their coupling from Gi–Go to Gs under certain conditions8,22. Subsequent studies with cloned m-, d- and k-opioid receptors transfected into various cell types devoid of intrinsic opioid receptors have shown that they become coupled via Gi–Go regulatory proteins to PTX-sensitive, inhibitory transducer systems, whereas no evidence of coupling via Gs to excitatory systems has been reported23,24. This paradox has now been clarified by recent studies demonstrating that a homogeneous population of cloned d-opioid receptors transfected into CHO cells (which normally show only opioid inhibition of forskolin-stimulated AC via PTX-sensitive Gi coupling) can be rapidly converted to an excitatory mode by treatment with GM1 ganglioside (10 nM–1 mM for 30 min)25,26. Application of the same opioid agonist to the GM1-

treated cells then results in stimulation of AC activity and an increase in the concentration of cAMP, suggesting that these cloned opioid receptors had undergone a conformational change that switched their coupling from Gi to Gs, thereby converting these receptors from an inhibitory to excitatory mode (Fig. 1; see below). This effect of GM1 is remarkably specific: comparative tests using GM2, GM3 and other gangliosides do not alter opioid inhibition of AC. Furthermore, replacing the positively charged Arg192 residue of the cloned d-opioid receptor (located near the external membrane surface) with an uncharged Ala residue does not alter opioid inhibitory effects when transfected into HEK293 cells; however, this mutation eliminates GM1-induced switching of coupling from Gi to Gs, so that GM1 treatment no longer results in the stimulation of AC by opioids26. The failure to detect opioid stimulation of AC in the initial

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series of studies of cloned opioid receptors transfected into CHO or other cells23,24 appears to have been due to the paucity of endogenous GM1 ganglioside in the cell types used in those experiments25, in contrast to the abundance of GM1 on neuronal cell surfaces1. Interestingly, in recent studies of HEK cells transfected with cloned m-opioid receptors, low concentrations (10–100 pM) of a m-opioid peptide DAMGO {[D-Ala2, MePhe4, Gly(ol)5]enkephalin} are, in fact, able to stimulate cAMP accumulation; morphine (10 nM) can elicit similar excitatory effects after the blockade of inhibitory opioid receptor functions by PTX (see Fig. 2 in Ref. 27). These results from the study of cloned m-opioid receptor functions27 are remarkably consistent with opioid excitatory effects on the action potential duration of naive and PTX-treated DRG neurones3,8, suggesting that HEK293 cells might contain low levels of GM1. In another recent study of cloned m-opioid receptors transfected into CHO cells, the acute application of low concentrations of morphine or m-opioid peptide was not able to stimulate AC activity in naive or PTX-treated cells28. Nevertheless, after treatment with morphine for 4 h, the application of naloxone resulted in marked

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increases in cAMP accumulation (i.e. ‘AC supersensitization’ occurred)28. These results are consistent with the naloxone-precipitated excitatory effects that occur in DRG neurones after acute treatment with GM1 or chronic treatment with opioids9,10,18,29 (see below). The experimental paradigm used by Wu et al.25,26 to demonstrate the conversion by GM1 of cloned d-opioid receptors from a Gi-linked, inhibitory to a Gs-linked, excitatory state was based upon previous electrophysiological evidence. This evidence was that after brief treatment (for 1–8 min) of DRG neurones with low concentrations of GM1 (10 nM) (but not GM2, GM3, or other gangliosides or glycolipids) the threshold concentration of the opioid peptide dynorphin A(1–13) that is required to prolong the action potential duration in many DRG neurones is markedly decreased from nanomolar to picomolar or femtomolar levels12,30. Furthermore, the opioid antagonist naloxone, which blocks excitatory as well as inhibitory opioid effects when applied at nanomolar concentrations to naive DRG cells, paradoxically prolongs the action potential duration of GM1treated cells9. This resembles naloxone-precipitated, excitatory withdrawal effects after chronic morphine

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V treatment in vitro10 (see also Refs 28 and 29) and in vivo31. Conversely, brief treatment of naive DRG neurones with the B subunit of CTX [which binds specifically and with high affinity (Kd = 10–10 M) to GM1 sites on neuronal membranes1] or with affinity-purified, anti-GM1 antibodies selectively blocks opioid-induced but not forskolin-induced prolongation of the action potential duration12. By contrast, the same treatments do not attenuate opioid-induced shortening of the action potential duration. These electrophysiological studies suggest that the increased sensitivity of GM1-treated DRG neurones to the excitatory effects of opioid agonists and the partial agonist properties of opioid antagonists is due to GM1 binding to an allosteric regulatory site on opioid receptors12, which increases the efficacy of the coupling of opioid receptors to Gs. The mechanism underlying GM1induced increases in the efficacy of excitatory opioid receptor functions is unknown, but might involve GM1mediated enhancement of the access of the receptors to larger numbers of compartmentalized Gs proteins, resulting in a much higher-gain AC–cAMP transduction cascade10 (see also Ref. 32). Under these conditions, even the very weak partial agonist property of naloxone becomes effective in eliciting excitatory responses (which prolong the action potential duration) in GM1treated neurones at nanomolecular concentrations (whereas >10 mM naloxone is required to prolong the action potential duration of naive cells)9. However, the results do not preclude a GM1-induced increase in the binding affinity of naloxone to opioid receptors (see below).

GM1 treatment increases efficacy and proportions of Gs-coupled opioid receptors

The GM1-regulated bimodal properties of opioid receptors are in sharp contrast to most monoaminergic and other G protein-coupled receptors where molecularly distinct receptor subtypes are coupled to either Gs or Gi–Go (e.g. Gs-coupled b-adrenoceptors and Gi–Gocoupled a2-adrenoceptors33). We are unaware of any studies indicating that GM1 or other specific gangliosides can regulate the G protein-coupling functions of adrenoceptor or other neuronal receptors. However, it should be noted that treating neuroblastoma cells with GM1 does, in fact, modulate Gs-coupled 5-HT receptors34 and Gs-coupled prostaglandin E1 receptors35 by enhancing agonist-induced elevation of cAMP levels. A small, but significant, fraction of the opioid receptors in nociceptive types of mouse DRG neurone appear to be coupled to Gs under normal culture conditions. This is shown by the marked excitatory effects, such as the prolongation of the action potential duration and a decrease in K+ conductance in the membrane, that can be elicited in these cells using low (fM–nM) concentrations of opioids without adding exogenous GM1 (Refs 3, 8 and 36). These data suggest that DRG neurones are endowed with a significant amount of endogenous GM1, in contrast to the absence of GM1 from CHO cells25,26. Gs-

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coupled opioid receptors have a much greater efficacy in altering membrane conductance than Gi–Go-coupled opioid receptors. This is because of the amplification resulting from the link between the Gs protein and the second-messenger cascade involving AC, cAMP and PKA (as occurs with b-adrenergic receptors37), in contrast to the direct link between Go and ion channels38. Therefore, following the application of low (lower than nanomolar) concentrations of bimodally acting opioid agonists to DRG neurones, both Gs-coupled, excitatory as well as Gi–Go-coupled, inhibitory receptors are activated. However, the inhibitory effects are generally masked by the activation of high-efficacy, Gs-coupled excitatory receptor functions, which results in the prolongation of the action potential duration8 or decreases in membrane K+ conductance (Fig. 1)36,39. Patch-clamp recordings in dissociated DRG neurones show that the application of low (pM) concentrations of opioids does, in fact, result in a rapid onset of increased K+ conductance (an inhibitory effect). However, these effects are generally masked within a few minutes by the delayed onset of a sustained decrease in K+ conductance (an excitatory effect) via the more slowly acting, cAMP-mediated transducer cascade39. By contrast, at higher (mM) opioid concentrations, DRG neurones generally display dosedependent, inhibitory effects (Fig. 1), which are apparently due to the summation of activation of the much larger fraction of opioid receptors that are normally in the inhibitory mode. On the other hand, following acute application of exogenous GM1 or chronic opioid treatment (which elevates endogenous GM1 levels29), opioid-induced excitatory effects on DRG neurones become much more prominent, and neurones that have been chronically treated with opioids generally display a prolongation of the action potential duration – even when acutely tested with high (0.1–10 mM) opioid concentrations. Inhibitory effects mediated by Gi–Gocoupled opioid receptors are evidently attenuated or masked by activation of the larger proportions of opioid receptors that become coupled to Gs in these treated DRG neurones, following conversion of many opioid receptors from the inhibitory to excitatory mode (as occurs in CHO cells25). These mechanisms could underlie the development of tolerance during chronic opioid treatment (see below).

Opioid antagonists at very low concentrations selectively block the excitatory mode of opioid receptors The conformational change induced by GM1 binding to opioid receptors in naive DRG neurones appears to involve not only a switch in coupling from Gi–Go to Gs but also a remarkably selective enhancement in the efficacy of opioid antagonists in blocking Gs-coupled opioid receptor functions. Very low (pM) concentrations of the opioid receptor antagonists naloxone, naltrexone and diprenorphine selectively block excitatory effects (which prolong the action potential duration) elicited by low

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(pM–nM) concentrations of morphine (and other m, d or k agonists) in naive DRG neurones13,19. These antagonists thereby unmask inhibitory effects (which shorten the action potential duration) that generally require higher (mM) concentrations of morphine or other opioid agonists (Fig. 2a,b). Much higher (nM) concentrations of naloxone, naltrexone or diprenorphine are required to block inhibitory opioid receptor functions (Figs 1 and 2a,b). Similarly, acute co-treatment of mice with morphine (1–3 mg per kg body mass) and remarkably low doses of naltrexone (1–100 ng per kg body mass) enhances the antinociceptive potency of morphine, as measured by hot-water, tail-flick assays19,40 (Fig. 2c). The mechanism underlying the efficacy of these very low doses of naloxone, naltrexone and other opioid antagonists in selectively blocking Gs-coupled opioid receptor functions in DRG neurones is unknown. Perhaps the binding affinity of these antagonists is enhanced following GM1-induced conformational changes in a way that is analogous to the more than tenfold increase in binding affinity of agonists to Gs-coupled 5-HT receptors after ganglioside treatment of neuroblastoma cells34. These correlative studies in vitro and in vivo provide novel insights into cellular mechanisms that could underlie many clinical and animal reports of ‘paradoxical’ analgesic effects of low doses of naloxone and naltrexone that have been published during the past three decades (see Refs 19 and 40). Several other types of alkaloid and peptide have been shown to have remarkably potent blocking actions on excitatory, Gs-coupled opioid receptor functions when tested at picomolar concentrations on DRG neurones13,41,42. These include opioids that have unexpectedly strong analgesic potencies in vivo, such as etorphine43 and the peptide biphalin44. Co-treatment of DRG neurones with nanomolar concentrations of morphine (which prolongs the action potential duration when applied alone) plus picomolar concentrations of etorphine or biphalin, markedly enhances the inhibitory potency of morphine13,42; the same result occurs with the application of nanomolar levels of morphine plus picomolar naltrexone or naloxone (Refs 19, 40). Furthermore, the acute application of etorphine or biphalin to DRG neurones elicits dose-dependent shortening of the action potential duration at all concentrations ranging from picomolar to micromolar13,42, in contrast to the characteristic excitatory effects evoked by low (pM–nM) concentrations of morphine and most m-, d- and k-opioid alkaloid and peptide agonists3,8. These electrophysiological studies of the unique effects of etorphine on DRG neurones might help to account for unexplained biochemical actions of this opioid alkaloid on HEK cells transfected with m-opioid receptors. The acute application of etorphine to these HEK293 cells elicits dose-dependent inhibition of cAMP accumulation at all concentrations ranging from picomolar to micromolar, whereas a m-opioid peptide at a concentration of 10–100 pM increases cAMP levels27. 362

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Furthermore, in HEK293 cells transfected with k-opioid receptors, pretreatment with etorphine but not with a kopioid agonist results in a marked increase in the efficacy of k-opioid inhibition of cAMP accumulation45. Studies of DRG neurones in vitro indicate that etorphine and biphalin have potent agonist actions at inhibitory Gi–Go-coupled opioid receptors concomitant with potent antagonist actions at excitatory Gs-coupled opioid receptors13,42, which could account for the fact that their analgesic potency is 1000 times stronger than morphine43,44. Acute co-treatment of mice with morphine (1 mg per kg body mass) and a very low subthreshold dose of etorphine (10 ng per kg body mass) does, in fact, enhance the antinociceptive potency of morphine40, as occurs when morphine and low-dose naltrexone are administered together19. The remarkably similar effects of co-treatment with very low doses of etorphine or naltrexone in enhancing morphine’s antinociceptive potency (notwithstanding their opioid ‘agonist’ compared with ‘antagonist’ effects, respectively, when administered alone at higher, conventional, clinical doses) would be difficult to account for without recognizing the shared, high-efficacy, selective antagonist action of both alkaloids on excitatory opioid receptor functions, as revealed by studies of DRG neurones in vitro13,19.

Alterations of bimodal opioid receptor functions during chronic opioid treatment The block of sustained activation of excitatory opioid receptor functions in DRG neurones in culture treated chronically with micromolar concentrations of morphine by co-treatment with either the B subunit of CTX (Ref. 12) or picomolar concentrations of naltrexone (Ref. 19) has two effects. It prevents both the development of the usual tolerance to the inhibitory effects (which shorten the action potential duration) of morphine and supersensitivity to the excitatory effects (which prolong the action potential duration) of extremely low concentrations of opioid agonists, as well as of nanomolar concentrations of naloxone, on these treated DRG neurones10,18. These studies in vitro indicate that sustained activation of Gs-coupled, GM1-regulated opioid receptor functions is required for the development of cellular manifestations of tolerance and physical dependence. It should be emphasized that opioid receptors in the Gscoupled excitatory mode appear to become progressively sensitized during chronic exposure of DRG neurones to bimodally acting opioid agonists10. This is in sharp contrast to the marked desensitization that occurs during sustained exposure to agonists of Gs-coupled badrenoceptors33, some types of Gi–Go-coupled opioid receptors33,46,47 and many other G protein-coupled receptors48, following the activation of G protein-coupled receptor kinases33, arrestins33,48, PKA (Ref. 49) and PKC (Ref. 50) (see Fig. 1). Interestingly, chronic morphine treatment increases the levels of b-adrenoceptor kinase and b-arrestin in the rat locus coeruleus51. These

V alterations might contribute to the homologous desensitization of inhibitory opioid receptors that underlies the tolerance observed in locus coeruleus neurones chronically treated with opioids46,47. The excitatory partial agonist effects of naloxone and other opioid antagonists in tests on DRG neurones that have been chronically treated with morphine or that have been acutely treated with GM1 (Refs 9, 10) is in sharp contrast to the potent excitatory antagonist action of naloxone and naltrexone during acute or chronic cotreatment of naive DRG neurones with morphine19,20. Opioid excitatory supersensitivity also occurs after acute treatment of DRG neurones with forskolin, which rapidly elevates cAMP levels8. These studies in vitro led to the proposal10 that opioid tolerance or dependence is mediated not only by upregulation of the Gs–AC–cAMP–PKA system22,28,52,53 but also by elevation of the concentration of GM1 ganglioside, following activation of the cAMP–PKA-dependent glycosyltransferase that synthesizes GM1 (Refs 29, 54, 55). Coordination of these processes provides a positive feedback phosphorylation cycle (dotted loop in Fig. 1) that could amplify the sensitivity of GM1-regulated, Gs-coupled, excitatory opioid receptors to low levels of endogenous opioids10, and thereby account for the protracted dependence (e.g. naloxone supersensitivity) observed for months after the withdrawal of chronic exogenous opioids in vitro20 as well as in vivo56. Elevation of GM1 by sustained activation of Gs-coupled opioid receptors might have two major effects: (1) an increase in the conversion of opioid receptors from the inhibitory Gi–Go-coupled mode25,26, thereby increasing the number of receptors in the Gscoupled mode (Fig. 1); and (2) an increase in the efficacy of coupling9,10,32 of these receptors to the AC–cAMP transducer system. These alterations in excitatory opioid receptor function during chronic morphine treatment of DRG neurones appear to play important roles in the attenuation of opioid inhibitory potency (tolerance) and the development of opioid excitatory supersensitivity (dependence). Interestingly, treating DRG neurones that had previously been chronically supersensitized to morphine with a low (nM) concentration of etorphine for one week after morphine had been withdrawn blocked the protracted dependence (e.g. naloxone supersensitivity) that is otherwise observed for months after neurones are returned to normal culture conditions20. Presumably, this post-treatment with etorphine blocks the sustained activation of the supersensitized excitatory opioid receptors by the low levels of endogenous opioid agonists present in the serum or embryo extract culture medium20. Remarkably similar results have recently been reported following the treatment of monkeys that were chronically dependent on morphine with a low subanalgesic dose of the etorphine analogue dihydroetorphine for several days after morphine had been withdrawn (60 ng per kg body mass; see below)57. Acute administration of naloxone at a dose that would otherwise have precipi-

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tated a full-blown withdrawal syndrome in morphinedependent monkeys failed to evoke withdrawal signs in animals that had been post-treated with dihydroetorphine57 (M. D. Aceto, pers. commun.).

Roles of Gs-coupled opioid receptor functions in vivo Intracerebroventricular injection of antibodies directed against Gsa has been reported to enhance the supraspinal antinociception elicited by morphine and bendorphin in mice58. These results are consistent with the hypothesis that selective blockade of Gs-coupled, excitatory opioid receptor functions by low doses of naltrexone or etorphine enhances the antinociceptive potency of morphine19,40 (Fig. 2c). Similarly, selective blockade of excitatory opioid receptor functions during chronic cotreatment of mice with morphine (1–3 mg per kg body mass) and low doses of naltrexone or etorphine (10–100 ng per kg body mass) does, indeed, prevent the development of tolerance (measured by hot-water, tail-flick assays) and of dependence (measured by naloxoneprecipitated, withdrawal-jumping assays)19,40. These studies of the effects of etorphine on mice in vivo, together with correlative evidence that etorphine and dihydroetorphine antagonize excitatory opioid receptor functions in DRG neurones in vitro13, suggest a cellular mechanism to account for recent unexpected findings that the chronic treatment of monkeys with supra-analgesic doses of dihydroetorphine for six weeks does not result in naloxone-precipitated withdrawal symptoms59. By contrast, characteristic withdrawal has been observed in parallel tests on monkeys that had been chronically treated with morphine59. The remarkable agreement between these results in vivo and studies in vitro on DRG neurones following acute as well as chronic co-treatment with morphine and low doses of naltrexone or etorphine provides strong support for the hypothesis that excitatory, Gs-coupled, GM1-regulated opioid receptor functions play important roles in modulating opioid analgesia, tolerance and dependence. Studies in vitro showing that the elevation of endogenous GM1 enhances excitatory opioid receptor functions that might mediate anti-analgesia, tolerance and dependence appear to be in disagreement with reports that the administration of exogenous GM1 in vivo attenuates pain, tolerance and dependence60 (see Ref. 4). However, GM1 was administered intrathecally or systemically in these reports on rats. Therefore, the actual GM1 concentrations reaching the nociceptive neurones in the spinal cord were not clearly defined and might have been substantially higher than the GM1 concentrations (as low as 10 nM) that enhance the efficacy of Gs-coupled opioid receptors and convert opioid receptors from inhibitory to excitatory modes in cultured cells9,25,30. High concentrations of GM1 have been shown to block the translocation of PKC from the cytosol to neuronal surface membranes, thereby interfering with the activation by PKC of increases in Ca2+ influx, mediated by NMDA

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receptors, and associated anti-analgesic or excitatory effects60. Furthermore, high GM1 concentrations (10 mM) decrease forskolin-stimulated cAMP levels in some cells35, perhaps by inhibiting certain subtypes of AC directly61, thereby counteracting the dose-dependent GM1 enhancement of Gs-coupled opioid receptor functions. Excitatory, as well as inhibitory, modes of opioid receptors might be present on various types of central, as well as peripheral, neurones in nociceptive and other CNS networks that regulate opioid analgesia and the effects of tolerance or dependence, depending on the concentration of GM1 in the surface membrane of each cell. For example, enhancement of the antinociceptive potency of morphine by co-treating mice with very low doses of naltrexone19,40 appears to be mediated by the selective block of excitatory opioid receptors located on presynaptic DRG terminals in the spinal cord (see also Ref. 15). Furthermore, attenuation of naloxone-precipitated, withdrawal-jumping behaviour in mice that are chronically dependent on morphine by co-treatment with low concentrations of naltrexone19,40 suggests that this effect is due to selective blockade of supersensitized, excitatory opioid receptors located not only on DRG neurones and dorsal horn networks but also on locus coeruleus and related brainstem neurones which appear to play primary roles in mediating physical behavioural signs of opioid withdrawal46,51,62.

Concluding remarks Opioid receptors can be interconverted rapidly between inhibitory (Gi/Go-coupled) and excitatory (Gscoupled) modes following physiological alterations in the concentration of a specific cAMP–PKA-dependent glycolipid, GM1 ganglioside, in the neuronal cell membrane. This dynamic plasticity provides a unique cellular mechanism that might modulate opioid analgesia, tolerance and dependence, as well as functions mediated by opioid receptors involved in brain-reward circuits63,64 and other emotional states of the nervous system.

Note added in proof Since the submission of this article, Gan et al. [(1997) Anesthesiology 87, 1075–1081] reported a clinical study on 60 post-hysterectomy patients indicating that an infusion of a low concentration of naloxone [0.25 mg per kg body mass per hour, intravenous (i.v.)] during a 24 h test period significantly reduced the cumulative, patientcontrolled use of morphine (i.v.) from about 60 mg down to 40 mg. Furthermore, the differences between the cumulative morphine use in the naloxone-co-treated versus placebo groups began to occur only after 4–8 h and became increasingly prominent after 20–24h of naloxone treatment. These data suggest that the patients using morphine alone were becoming progressively tolerant to the analgesic effects of morphine during the 24 h test period, in contrast to the stable rate of morphine use by the patients also receiving a low dose of naloxone. The 364

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‘surprising and intriguing opioid-sparing effect seen with low-dose naloxone’ in this study by Gan et al. was predicted by our preclinical studies in vitro and in vivo demonstrating that co-treatment with low doses of naloxone or naltrexone results in the sustained enhancement of morphine’s antinociceptive potency by selectively antagonizing its excitatory, anti-analgesic side-effects19,40. Selected references 1 Ledeen, R. W. (1989) in Neurobiology of Glycoconjugates (Margolis, R. V. and Margolis, R. K., eds), pp. 43–83, Plenum Press 2 Bremer, E. G. and Hakomori, S. (1984) in Ganglioside Structure, Function and Biomedical Potential (Ledeen, R. W., Yu, R. K., Rapport, M. M. and Suzuki, K., eds), pp. 381–394, Plenum Press 3 Crain, S. M. and Shen, K-F. (1990) Trends Pharmacol. Sci. 11, 77–81 4 Smart, D. and Lambert, D. G. (1996) Trends Pharmacol. Sci. 17, 264–269 5 Huang, L-Y. M. (1995) in The Pharmacology of Opioid Peptides (Tseng, L. F., ed.), pp. 131–149, Harwood Academic Publishers 6 Sarne, Y., Fields, A., Keren, O. and Gafni, M. (1996) Neurochem. Res. 21, 1353–1361 7 North, R. A. (1986) Trends Neurosci. 9, 114–117 8 Shen, K-F. and Crain, S. M. (1989) Brain Res. 491, 227–242 9 Crain, S. M. and Shen, K-F. (1992) J. Pharmacol. Exp. Ther. 260, 182–186 10 Crain, S. M. and Shen, K-F. (1992) Brain Res. 575, 13–24 11 Shen, K-F. and Crain, S. M. (1990) Brain Res. 525, 225–231 12 Shen, K-F. and Crain, S. M. (1990) Brain Res. 531, 1–7 13 Shen, K-F. and Crain, S. M. (1994) Brain Res. 636, 286–297 14 Shen, K-F. and Crain, S. M. (1994) J. Neurosci. 14, 5570–5579 15 Suarez–Roca, H. and Maixner, W. (1993) J. Pharmacol. Exp. Ther. 264, 648–653 16 Fujimoto, J. M. and Rady, J. J. (1989) J. Pharmacol. Exp. Ther. 251, 1045–1052 17 Arts, K. S., Fujimoto, J. M. and Crain, S. M. (1993) Pharmacol. Biochem. Behav. 46, 623–628 18 Shen, K-F. and Crain, S. M. (1992) Brain Res. 597, 74–83 19 Crain, S. M. and Shen, K-F. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 10540–10544 20 Crain, S. M. and Shen, K-F. (1995) Brain Res. 694, 103–110 21 Crain, S. M. and Shen, K-F. (1996) Brain Res. 741, 275–283 22 Cruciani, R. A., Dvorkin, B., Morris, S. A., Crain, S. M. and Makman, M. H. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 3019–3023 23 Uhl, G. R., Childers, S. and Pasternak, G. (1994) Trends Neurosci. 17, 89–93 24 Zaki, P. A., Bilsky, E. J., Vanderah, T. W., Lai, J., Evans, C. J. and Porreca, F. (1996) Annu. Rev. Pharmacol. Toxicol. 36, 379–401 25 Wu, G., Lu, Z. and Ledeen, R. W. (1997) Mol. Brain Res. 44, 341–346 26 Wu, G. et al. (1998) Ann. New York Acad. Sci. 845, 126–138 27 Blake, A. D., Bot, G., Freeman, J. C. and Reisine, T. (1997) J. Biol. Chem. 272, 782–790 28 Avidor-Reiss, T., Bayewitch, M., Levy, R., Matus-Leibovitch, N., Nevo, I. and Vogel, Z. (1995) J. Biol. Chem. 270, 29732–29738 29 Wu, G., Fan, S. F., Ledeen, R. W. and Crain, S. M. (1995) J. Neurosci. Res. 42, 493–503 30 Shen, K-F., Crain, S. M. and Ledeen, R. E. (1991) Brain Res. 559, 130–138 31 Wikler, A. (1980) Opioid Dependence, Plenum Press 32 Crain, S. M. and Shen, K-F. (1998) Ann. New York Acad. Sci. 845, 106–125 33 Freedman, N. J. and Lefkowitz, R. J. (1996) Recent Prog. Hormone Res. 51, 319–351 34 Berry-Kravis, E. and Dawson, G. (1985) J. Neurochem. 45, 1739–1747 35 Wu, G., Lu, Z-H. and Ledeen, R. W. (1996) Glycoconjugate J. 13, 235–239 36 Fan, S.F., Shen, K-F. and Crain, S. M. (1991) Brain Res. 558, 166–170 37 Lefkowitz, R. J. and Caron, M. G. (1990) Adv. Sec. Mess. Phosph. Res. 24, 1–8 38 Miyake, M. J., Christie, M. J. and North, R. A. (1989) Proc. Natl. Acad. Sci. U. S. A. 86, 3419–3422 39 Fan, S. F. and Crain, S. M. (1995) Brain Res. 696, 97–105

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Shen, K-F. and Crain, S. M. (1997) Brain Res. 757, 176–190 Shen, K-F. and Crain, S. M. (1995) Brain Res. 673, 30–38 Shen, K-F. and Crain, S. M. (1995) Brain Res. 701, 158–166 Blane, G. F., Boura, A. L. A., Fitzgerald, A. E. and Lister, R. E. (1967) Br. J. Pharmacol. Chemother. 30, 11–22 Horan, P. J. et al. (1993) J. Pharmacol. Exp. Ther. 265, 1446–1454 Blake, A. D., Bot, G., Li S., Freeman, J. C. and Reisine, T. (1997) J. Neurochem. 68, 1846–1852 Aghajanian, G. K. (1978) Nature 276, 186–188 Harris, G. C. and Williams, J. T. (1991) J. Neurosci. 11, 7025–7029 Chuang, T. T., Iacovelli, L., Sallese, M. and De Blasi, A. (1996) Trends Pharmacol. Sci. 17, 416–421 Freedman, N. J., Liggett, S. B., Drachman, D. E., Pei, G., Caron, M. G. and Lefkowitz, R. J. (1995) J. Biol. Chem. 270, 17953–17961 Ueda, H. et al. (1995) J. Neurosci. 15, 7485–7499 Terwilliger, R. Z., Ortiz, J., Guitart, X. and Nestler, E. J. (1994) J. Neurochem. 63, 1983–1986 Terwilliger, R. Z., Beitner-Johnson, D., Sevarino, K. A., Crain, S. M. and Nestler, E. J. (1991) Brain Res. 548, 100–110 Makman, M. H., Dvorkin, B. and Crain, S. M. (1988) Brain Res. 445,

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Ligand-binding studies: old beliefs and new strategies

analysis. This wide literature notwithstanding, it is not infrequent to find a faulty interpretation of binding protocols or data for convenience, simplicity, misinterpretation between theory and practice, or just for the sake of tradition.

G. Enrico Rovati

Equilibrium-binding experiments

Ligand-binding studies remain a very popular technique among many experimentalists. As far as equilibrium experiments are concerned, saturation and displacement curves are commonly performed for simplicity, convenience or for the sake of tradition. However, alternative protocols, such as ‘mixed’-type protocols or multiligand experiments, are also possible. Indeed, there are cases where kinetic experiments, usually considered a ‘second-choice’ experiment, might have a superior resolving power compared to equilibrium ones. A combination of equilibrium and kinetic experiments might be a powerful solution to overcome limits and shortcomings of each specific technique and is discussed in this issue by G. Enrico Rovati. Thus, a careful choice of the design, a protocol optimization and a computerized analysis of the data can yield a dramatic improvement in the precision of the parameter estimation over more conventional approaches. Despite the advent of molecular biology, ligand-binding studies remain a very popular technique among many experimentalists. A great number of papers have been published on the design, analysis and interpretation of ligand-binding data, using a number of different approaches from graphical interpretations to computer

Conventionally, two distinct types of equilibrium experiments are most commonly performed: the ‘saturation experiment’ and the ‘competition experiment’. The latter might involve the same chemical species of the tracer (homologous displacement), or a different chemical species (heterologous displacement). Indeed, from a mathematical point of view, both saturation and homologous displacement curves contain exactly the same type of information1,2 and the choice between them is not a theoretical problem, but rather a practical one. In fact, both experimental protocols have advantages and drawbacks (Table 1). For example, with saturation designs, one cannot usually span more than a 100-fold concentration range, thus limiting the overall precision of parameter estimates and reducing the ability to distinguish multiple binding sites. This is especially true if, as is often found in the literature, a tenfold concentration range or even less is spanned. A second problem that is overlooked with saturation protocols is that the affinity estimate strongly depends upon knowledge of the specific activity of the labelled species3. An error computed at this level will be carried over all the concentrations used4, thus leading to a wrong estimation of Kd. Conversely, a competitiontype experiment has different limitations. To estimate the lowest possible Kd values, one must use the smallest feasible concentration of label, otherwise part of the binding curve will become unavailable owing to the self displacement of the tracer1,3. This will lead to the problem of counts that are too close to the counter background, especially when using tritiated ligands, which usually have a low specific activity.

Copyright © 1998, Elsevier Science Ltd. All rights reserved. 0165 – 6147/98/$19.00 PII: S0165-6147(98)01242-5

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G. E. Rovati, Assistant Professor of Pharmacology, Laboratory of Molecular Pharmacology, Institute of Pharmacological Sciences, University of Milan, Via Balzaretti 9, 20133 Milan, Italy.

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