The present address of Chris P. Bailey is the Department of Pharmacy and Pharmacology, University of Bath, Bath BA2 7AY, UK.
C-terminal splice variants of the μ-opioid receptor: existence, distribution and functional characteristics
Version of Record online: 18 AUG 2008
© 2007 The Authors Journal Compilation © 2007 International Society for Neurochemistry
Journal of Neurochemistry
Volume 104, Issue 4, pages 937–945, February 2008
How to Cite
Oldfield, S., Braksator, E., Rodriguez-Martin, I., Bailey, C. P., Donaldson, L. F., Henderson, G. and Kelly, E. (2008), C-terminal splice variants of the μ-opioid receptor: existence, distribution and functional characteristics. Journal of Neurochemistry, 104: 937–945. doi: 10.1111/j.1471-4159.2007.05057.x
- Issue online: 18 AUG 2008
- Version of Record online: 18 AUG 2008
- Received August 23, 2007; accepted September 24, 2007.
- G-protein-coupled receptor;
- splice variant;
- μ-opioid receptor
The distribution of the mRNA of different C-terminal splice variants of the μ-opioid receptor in rat CNS was assessed by RT-PCR. The mRNA species for MOR1, MOR1A and MOR1B were readily detectable and distributed widely throughout the rat CNS, with levels of MOR1 and MOR1A mRNA being overall greater than for MOR1B. We did not find convincing evidence that significant levels of MOR1C, MOR1C1, MOR1C2 and MOR1D are present in rat CNS. To examine possible differences in the agonist-induced regulation of MOR1, MOR1A and MOR1B, we expressed these constructs in HEK293 cells along with G-protein-coupled inwardly rectifying K+ channel subunits and measured the rate and extent of desensitisation of (d-Ala2,N-Me-Phe4,glycinol5)-enkephalin (DAMGO)- and morphine-induced G-protein-coupled inwardly rectifying K+ currents. Morphine-induced desensitisation was rapid for all three splice variants (t½: 1.2–1.7 min) but DAMGO-induced desensitisation was significantly slower for MOR1B (t½ 4.2 min). Inhibition of endocytosis by expression of a dynamin-dominant negative mutant increased the rate of DAMGO-induced desensitisation of MOR1B. These data show that some splice variants of μ-opioid receptor are widely expressed in rat CNS but question the existence of others that have been reported in the literature. In addition, whereas the rate of desensitisation of MOR1 and MOR1A is agonist-independent, that for MOR1B is agonist-dependent.
dorsal root ganglia
G-protein-coupled inwardly rectifying K+ channel
guanosine 5′(γ-thio) triphosphate
Morphine and related opioids exert their powerful biological effects; analgesia, euphoria and respiratory depression through the μ-opioid receptor (MOPr), a G-protein-coupled, plasma membrane receptor protein. Identical forms of the rat receptor, originally termed MOR1, were cloned by several groups (Chen et al. 1993; Fukuda et al. 1993; Thompson et al. 1993). It is the product of a single gene, Oprm1, spanning approximately 250 kb of chromosome 1 (Rat Genome Sequencing Project Consortium 2004). Shortly after the initial cloning, Zimprich et al. (1995) identified a splice variant, MOR1B in which the last 12 amino acids of the C-terminal tail are replaced by five others (Table 1). Subsequently, several more splice variants, differing only in the C-terminal portion of the cytoplasmic tail, have been described in the rat (Pasternak et al. 2004; Schnell and Wessendorf 2004), mouse and human (reviewed by Pasternak 2004). Pasternak’s group (Pan et al. 2000) also describe an additional series of splice variants in the mouse giving mRNAs that differ at their 5′ ends. The putative C-terminal, rat splice variants, which form the subject of this paper, have exons 1, 2 and 3 in common but different exons or combinations of exons at the 3′ end leading to sequences of 1–82 amino acids after the splice point (Table 1).
However, the biological relevance of these reported splice variants is unclear; many of them appear to be of very low abundance and may not in fact be mature mRNAs. Furthermore, data on the distribution of the reported splice variants and functional differences between them are limited. MOR1, MOR1B and MOR1C have all been reported to be widely distributed throughout the brain (Zimprich et al. 1995; Schnell and Wessendorf 2004) although, using a different method, MOR1B was reported to be more localised (Schulz et al. 1998). Zimprich et al. (1995) found that functional coupling of MOR1B to adenylyl cyclase desensitised at a slower rate than did that of MOR1 during exposure to (d-Ala2,N-Me-Phe4,glycinol5)-enkephalin (DAMGO) and this was subsequently shown to be due to faster resensitisation and recycling of MOR1B back to the plasma membrane (Koch et al. 1998). It has been reported that in guanosine 5′-(γ-thio) triphosphate (GTPγS) binding assays agonists may exhibit different potencies and efficacies at different C-terminal splice variants (Pasternak et al. 2004). In the present study we have revisited the subject of C-terminal MOPr splice variants and have examined carefully the question of their existence, relative expression and functional characteristics.
Total RNA was isolated from the brain, spinal cord and dorsal root ganglia of male, 150 g Wistar rats, using TriZol, following the manufacturer’s instructions (Invitrogen, Paisley, UK). Single-strand cDNA was transcribed using M-MLV RT (Promega, Southampton, UK) and random hexanucleotide primers (Roche, Burgess Hill, UK). cDNAs were amplified using Biotaq polymerase (Bioline, London, UK) and primers as indicated. The sequences of the primers are given in Table 2. The forward primers, MOR1F1 and MOR1F2, hybridise to coding sequence common to all splice variants. Sequences of reverse primers were chosen to give good discrimination between splice variants and were either within the variant coding sequence or within the 3′ untranslated region as convenient. Generally, annealing was at 55°C for 1 min, unless otherwise indicated, and extension at 72°C for 30 s. The resulting products were subjected to electrophoresis on a 1.5% agarose gel containing ethidium bromide (0.8 μg/mL) and photographed under UV-illumination. Band density was quantified using Kodak ID3.6 software (Kodak, Rochester, NY, USA).
The PCR products were filled, phosphorylated and cloned into the SmaI site of pBluescript KSII+ using standard methods (Sambrook et al. 1989). The mammalian expression constructs, pcDNA3/T7MOR1 and pcDNA3/T7MOR1B were gifts from Dr Volker Höllt, Otto von Guericke University, Magdeburg, Germany. To construct pcDNA/T7MOR1A, the 3′ section of MOR1B was excised with XhoI and XbaI and replaced with a XhoI/XbaI fragment of the MOR1A PCR product which had been cloned into pBluescript. The K44A dominant negative (DN) mutant dynamin construct is as described previously (Damke et al. 1994; Mundell et al. 2001).
Cell culture and transfection
Cells were maintained at 37°C in 95% O2, 5% CO2, in Dulbecco’s modified Eagle’s medium supplemented with 10% foetal calf serum, 10 U/mL penicillin and 10 mg/mL streptomycin.
To produce a stable HEK293 cell line that expressed the G-protein-coupled inwardly rectifying K+ (GIRK) channel subunits Kir3.1 and Kir3.2, cDNAs encoding the two subunits (originally supplied by Dr Andrew Tinker, University College London) were cloned into pcDNA3.1 hygro. An internal ribosome entry site was placed between the two cDNAs to allow expression of both subunits from the single transcript under the control of the cytomegalovirus (CMV) promoter. HEK293 cells were transfected with this construct, using lipofectamine and stable transfectants were selected by the addition of hygromycin to the growth medium (125 μg/mL). Clones were screened by western blotting, probing with rabbit anti-GIRK 1 (Chemicon, Millipore, Watford, UK). A positive clone was selected, grown up and used for subsequent transient transfections with MOPr splice variants and electrophysiological recordings.
For transient transfections, cells were plated out at ∼40% confluence on glass coverslips, in 24-well plates, 24 h before co-transfection with peGFP-N (0.3 μg) and pcDNA3 into which the cDNA encoding MOR1, MOR1A or MOR1B had been inserted (0.5 μg), using calcium phosphate precipitation (Sambrook et al. 1989). To reduce the rate of recycling of MOR1B, some were transfected with a pcDNA3 construct encoding a DN mutant of dynamin (0.6 μg) in addition to peGFP-N and the MOR1B construct. Cells were incubated for 48 h before use.
The binding of [35S]GTPγS to membranes of HEK293 cells transiently expressing the rat MOPr splice variants was based on the assay described previously (Traynor and Nahorski 1995). In brief, cells were resuspended in ice-cold membrane buffer (0.2 mmol/L MgSO4, 0.38 mmol/L KH2PO4 and 0.61 mmol/L Na2HPO4, pH 7.4) and lysed in a hand-held homogeniser. The homogenates were centrifuged at 50 000 g for 20 min at 4°C and the pellets were resuspended in 50 mmol/L Tris–HCl, pH 7.4. Aliquots of membrane (50 μg of protein) were then incubated with 50 pmol/L [35S]GTPγS in assay buffer (50 mmol/L Tris–HCl, 3 mmol/L MgCl2, 0.2 mmol/L EGTA, 100 mmol/L NaCl, pH 7.4, 50 μmol/L GDP, 1 mmol/L dithiothreitol and containing DAMGO or morphine as indicated (final volume 500 μL) for 1 h at 22°C. Non-specific binding in all cases was determined by the addition of 10 μmol/L GTPγS to the assay. Binding was stopped by the addition of 4 mL of ice-cold 50 mmol/L Tris–HCl, pH 7.4, and the samples were rapidly filtered through glass fibre filters using a Brandel cell harvester, washing with 50 mmol/L Tris–HCl, pH 7.4. The amount of [35S]GTPγS bound to membranes on individual filters was then determined by liquid scintillation counting.
Whole cell patch-clamp recordings
Non-confluent monolayers of cells co-transfected with peGFP-N (to allow identification of transfected cells) and a MOPr construct, were mounted in a perfusion chamber (1.9 mL) on an inverted fluorescence microscope and superfused (4 mL/min, 25°C) with extracellular solution (160 mmol/L NaCl, 5 mmol/L KCl, 2 mmol/L CaCl2, 1 mmol/L MgCl2, 11 mmol/L glucose, 5 mmol/L HEPES pH 7.4). Cells expressing GFP were selected for whole cell voltage-clamp recordings. Electrodes (3–5 MΩ) were filled with 122 mmol/L KCl, 11 mmol/L EGTA, 1 mmol/L CaCl2, 2 mmol/L MgCl2, 10 mmol/L HEPES, 4 mmol/L MgATP, 0.25 mmol/L Na2GTP, 5 mmol/L NaCl, pH 7.2.
To enhance the amplitude of the MOPr-evoked GIRK currents, the K+ and Na+ concentrations of the extracellular solution were changed to 50 and 115 mmol/L respectively, at the start of recording, and inward current through these inwardly rectifying channels recorded. Furthermore, to ensure that currents evoked by prolonged exposure to agonists did not decline because of the inward current raising the intracellular K+ concentration, thereby reducing the electrochemical drive for further entry of K+ into the cell, a protocol that minimised the amount of K+ entry into the cell was used (Johnson et al. 2006). Cells were initially held at a membrane potential of −60 mV and when the extracellular solution was changed from low [K+] to high [K+], the membrane potential was changed to −25 mV, the reversal potential for the GIRK channel under these conditions. To measure GIRK channel activation in response to morphine or DAMGO, the membrane potential was then stepped from −25 to −60 mV for 60 ms every 5 s allowing current flow into the cell. Barium chloride (1 mmol/L), a blocker of Kir channels, abolished the DAMGO- and morphine-evoked currents. No opioid-evoked current was observed in HEK293-Kir cells that were not transfected with the MOPr splice variants.
Desensitisation of the MOPr-evoked GIRK current was quantified by expressing the current amplitude as a percentage of the initial peak current. The decay of current was fitted to a single exponential using Prism software (GraphPad Software Inc., San Diego, CA, USA).
All data are expressed as means ± SEM. Unpaired, two-tailed Student’s t-test was used to assess statistical significance of functional differences between the splice variants. To analyse the PCR data, the ratio of the intensity of the MOPr band to that of the glyceraldehyde-3-phosphate dehydrogenase (GAPdH) band was taken to compensate for differences in the amount of cDNA between samples. PCR samples from all tissues from a single rat, for a particular splice variant were run on the same gel. However, when data from several rats was pooled, the MOPr : GAPdH ratio for the individual brain regions was compared with that from hippocampus to compensate for inherent differences between individual gels. To assess statistical significance of differences in mRNA expression between tissues, anova was used; where significance was found, this was followed by a post hoc Dunnett’s test to identify significantly different groups. Statistical significance of differences in desensitisation data was assessed by an unpaired Student’s t-test. In all cases the null hypothesis was rejected at p < 0.05.
Detection and distribution of splice variants
An RT-PCR approach was used to determine the mRNA distribution of previously described C-terminal splice variants of the MOPr within the rat CNS. A forward primer, MOR1F1, corresponding to a sequence within exon 3 and hence common to all the known splice variants was used in conjunction with reverse primers specific for the alternative 3′ sequences. The latter were designed using sequences published on the EMBL nucleotide sequence database (see Table 2 for primer sequences). As they correspond to sequences in distinct, downstream exons, only cDNA can be amplified to give products of the expected size; any contaminating genomic DNA would not be amplified. The presence of the various postulated MOPr isoforms was assayed in spinal cord, dorsal root ganglia (DRG), whole brain and selected brain regions [cortex, hippocampus, locus coeruleus (LC) and ventral tegmental area]. Other work in our group uses the LC as a relatively homogenous source of noradrenergic neurons that express the MOPr but not δ- or κ-opioid receptors (Bailey et al. 2003, 2004), hence it was of interest to identify the MOR1 splice variants present in this nucleus and the ventral tegmental area was chosen as it is involved in the rewarding response to morphine (Williams et al. 2001).
MOR1 and MOR1A mRNA were detectable but not saturating, in all tissues tested after 26 cycles of PCR. Primers MOR1F1 and MOR1R or MOR1AR gave single bands of ∼320 and ∼260 bp respectively (Fig. 1). Cloning of these PCR products into pBluescript and subsequent sequencing confirmed their identity, giving C-terminal sequences that were identical to the published sequences for MOR1 (Chen et al. 1993; Fukuda et al. 1993) and MOR1A (Pasternak et al. 2004) (Table 1). MOR1B was more difficult to detect; PCR with primers MOR1F1 and MOR1BR required more than 35 cycles and, in many samples, gave several other products in addition to the expected band of ∼430 bp (Fig. 2), including a band that was difficult to resolve from the MOR1B product. Raising the annealing temperature resulted in loss of all bands, so nested PCR was used to increase the specificity. An aliquot of the reaction mix was re-amplified using MOR1F2, a primer still within exon 3 but nearer the splice site, and MOR1BR. This gave a predominant band of ∼390 bp (Fig. 2) which, on cloning and sequencing, was found to be identical to part of the published sequence for MOR1B (Zimprich et al. 1995). For the semi-quantitive experiments (Fig. 1), the number of cycles in the first round was reduced to 10 and care was taken to ensure that the second round did not reach saturation.
The splice variants MOR1C, MOR1C1, MOR1C2 and MOR1D have previously been described in the rat (Pasternak et al. 2004; Schnell and Wessendorf 2004). These putative splice variants include various combinations of exons designated 7, 8, 9a and 9b (Pasternak 2004). MOR1C1 and MOR1C2 contain exons 7, 8 and 9a or 9b respectively; MOR1D lacks exon 7, containing exons 8 and 9b and MOR1C has exon 7 followed by a sequence described by other workers (Schnell and Wessendorf 2004). Amplification with MOR1F1 and the exon-specific primers gave several faint bands after a high number of cycles but no predominant bands of the expected sizes. However, using nested PCR of 30 cycles with MOR1F1 and MOR1ex8R followed by 26 cycles with MOR1F2 and MOR1ex8R, a band of 255 bp, the expected size for the MOR1C variants was occasionally seen (data not shown). Sequencing confirmed that this band was indeed a MOR1C fragment. It was only detected after high numbers of cycles and then erratically between different tissues and different rats. Under these conditions, PCR would amplify sequences from just a few molecules of template and so it is likely that we are detecting incompletely spliced intermediates and not mature mRNA.
To obtain an estimate of the relative concentrations of the mRNAs of MOR1, MOR1A and MOR1B splice variants in each tissue, a fragment of GAPdH was also amplified and the densitometric ratio of each MOR band to that of GAPdH was quantified, the number of cycles of amplification being limited to within the linear range. MOR1, MOR1A and MOR1B mRNAs were present at each level of the spinal cord, in DRG and in all brain regions assayed in each of the three animals tested. Figure 1a shows the results from a representative animal and Fig. 1b the pooled results from three rats. MOR1 and MOR1A bands were clearly visible after 26 cycles, suggesting that they are present at similar concentrations. However, MOR1B mRNA required several more PCR cycles to be detectable and hence appears to be present at a much lower concentration. The expression of MOR1 mRNA, relative to that in the hippocampus, was greatest in the LC and only in this tissue did the difference reach statistical significance. No statistically significant differences were seen in the expression of either MOR1A or MOR1B mRNA in any tissue when compared with hippocampus although there is a trend towards higher concentrations of MOR1B in the DRG and LC.
MOPr splice variant functional coupling and agonist-induced desensitisation
We studied the function of the three MOPr splice variants for which mRNA was found at significant levels in the rat CNS; MOR1, MOR1A and MOR1B. Following expression of these MOPr isoforms in HEK293 cells, receptor-saturating concentrations of DAMGO (10 μmol/L) and morphine (30 μmol/L) increased binding of [35S]GTPγS to membrane preparations of the cells (data not shown). In each case, the stimulation because of morphine was less than that of DAMGO, in line with the former being a partial agonist at MOPr. To determine whether the different splice variants displayed different regulatory properties, we next investigated the ability of the receptors to undergo desensitisation by measuring the decay of the opioid-activated GIRK current in cells stably transfected with GIRK subunits, Kir3.1 and Kir3.2 and transiently transfected with a plasmid encoding a MOR1 isoform. For each splice variant, the DAMGO-stimulated GIRK currents were similar, as were those stimulated by morphine (data not shown), and similar to that which we have previously reported for MOR1 (Johnson et al. 2006). The desensitisation induced by receptor-saturating concentrations of morphine (30 μmol/L) and DAMGO (10 μmol/L) is shown in Fig. 3 and numerical values given in Table 3. In each case, morphine induced approximately 80% desensitisation with a t½ of between 1.2 and 1.8 min. There was no statistically significant difference between the three splice variants. DAMGO also induced desensitisation to a similar extent and with similar kinetics in cells expressing MOR1 or MOR1A. However, DAMGO-induced desensitisation was significantly slower in MOR1B-expressing cells although the extent of desensitisation was unaffected.
|DAMGO (10 μmol/L)||Morphine (30 μmol/L)|
|t½ (min)||% Desensitisation||t½ (min)||% Desensitisation|
|2.0 ± 0.3 (n = 3)||85 ± 4% (n = 3)||1.6 ± 0.5 (n = 5)||74 ± 7% (n = 5)|
|1.6 ± 0.1 (n = 6)||79 ± 3% (n = 6)||1.2 ± 0.2 (n = 5)||80 ± 2% (n = 5)|
|4.2 ± 0.9* (n = 4)||64 ± 6% (n = 4)||1.7 ± 0.3 (n = 6)||78 ± 3% (n = 6)|
It has previously been suggested that the apparent slow rate of desensitisation of MOR1B is due to rapid internalisation, resensitisation and recycling of this receptor (Koch et al. 1998). To examine this possibility, we over-expressed a DN mutant form of dynamin (dynamin K44A) to prevent receptor internalisation through clathrin coated pits (Damke et al. 1994). In cells over-expressing the DN dynamin and MOR1B, the desensitisation induced by morphine was unaffected but the rate of desensitisation induced by DAMGO was markedly enhanced (Fig. 4). However, DN dynamin had no significant effect on either the morphine- or DAMGO-induced desensitisation of MOR1 or MOR1A.
The mRNA species for three of the splice variants of MOPr, MOR1, MOR1A and MOR1B were consistently found throughout the rat CNS. As different exon-specific primers were, of necessity, used in each case, it is impossible to make accurate measurements of the relative amounts of the three isoforms. However, the fact that MOR1 and MOR1A mRNA were readily detectable and MOR1B mRNA required more cycles to be detected, suggests that MOR1 and MOR1A are the predominant isoforms and that, overall, MOR1B is present in lesser amounts. Our data indicate the average expression throughout one area of the CNS and it may be that MOR1B is concentrated in a particular cell type where its different properties have an important role. On investigating the distribution of each splice variant across the CNS, MOR1 was found to show a statistically significant concentration within the LC but otherwise, each isoform was fairly evenly distributed between tissues: a tendency for MOR1B to be concentrated in the DRG and LC did not reach statistical significance. A previous study, also using PCR, (Zimprich et al. 1995) also found MOR1 and MOR1B mRNAs to have similar, widespread distributions in the brain. The authors described some variations in expression between brain areas, with the lowest levels of both mRNA isoforms in the olfactory bulb. Although we were unable to detect the previously reported individual MOR1C variants, by using a primer pair designed to amplify a sequence common to all three of these variants, we were able to amplify a MOR1C fragment. However, a PCR band was only obtained after very many cycles and then only in a few samples. Its presence was erratic, appearing in different tissues in different rats. The conditions required for amplification would allow detection from a very few molecules of template and it is possible that the MOR1C variants derive from incompletely spliced intermediates and may not be present in the mature mRNA population. Using in situ hybridisation, Schnell and Wessendorf (2004) reported that MOR1C was widely distributed in the rat brain. However, the riboprobe used contained 47 bases of exon 3 sequence which may have allowed some cross-reaction with other C-terminal MOR splice variants. The reported nucleotide sequence of MOR1C which Schnell and Wessendorf (2004) amplified from rat is identical to that from mouse apart from a single base change. Part of this sequence is present in the rat genome but the 3′ portion cannot be found on a Blast search of the rat genome. Abbadie et al. (2000a,b) have used this mouse-specific sequence to design a 20-residue peptide antigen to raise a polyclonal antiserum for an immunohistochemical study of the distribution of MOR1C in the rat brain and spinal cord. As much of this epitope is not encoded by the rat Oprm1 gene and is not present in either of the rat MOR1C variants, MOR1C1 and MOR1C2, (Pasternak et al. 2004), but several unrelated proteins contain parts of it, the specificity of this antibody for MOR1C may be less than previously considered.
We were unable to detect MOR1D mRNA and, again, it is possible that in previous reports, very sensitive PCR conditions have allowed amplification of incompletely or erroneously spliced RNA. Although Pasternak et al. (2004) reported the cloning of rat MOR1D cDNA, which encoded a single amino acid, threonine, after the splice point, this is entirely different from the mouse MOR1D sequence described by the same group (Pan et al. 1999) which has the seven amino acid sequence, RNEEPSS, after the splice point. Furthermore, Abbadie et al. (2000b) raised an antibody to the mouse sequence and obtained immunohistological staining in rat CNS. Again, this epitope is not encoded by the rat Oprm1 gene but very similar amino acid sequences are found in several other proteins, raising doubts about the specificity of the antibody.
For many G-protein-coupled receptors, different splice variants have been found to display important functional differences (e.g. Mundell et al. 2002; reviewed in Kilpatrick et al. 1999) and the MOPr is likely to be no exception. Differences in the C-terminal tails would be expected to affect the interaction of the receptor with components of signalling, regulatory and recycling pathways. Drug-induced GIRK current decays on continued application of the drug because of desensitisation of the receptor (Johnson et al. 2006). In the present study MOR1 and MOR1A showed similar kinetics of desensitisation, irrespective of whether desensitisation was induced by morphine or DAMGO. MOR1B, however, desensitised more slowly in response to DAMGO than it did to morphine. MOR1B has previously been reported to be resistant to agonist-induced desensitisation because it internalises and recycles rapidly (Koch et al. 1998). To test whether rapid recycling was the cause of the slower desensitisation in response to DAMGO, we co-expressed a mutant form of dynamin (DN dynamin; Damke et al. 1994), which we have previously shown to inhibit clathrin-mediated internalisation in HEK293 cells (Mundell et al. 2001). Under these conditions, DAMGO-induced desensitisation was now faster, and indeed similar to that seen for the other splice variants without DN dynamin expression. This confirms that even under conditions of rapid (< 5 min) agonist-induced desensitisation, MOR1B trafficking greatly influences the rate of desensitisation. In addition these studies show that for MOR1B, unlike the other two splice variants studied, the rate of receptor desensitisation is agonist dependent. Furthermore, the fact that the rate of morphine-induced desensitisation for MOR1B was unchanged following inhibition of clathrin-mediated internalisation indicates that receptor trafficking plays little apparent role in the regulation of the morphine-induced response. This is consistent with other studies where it has been proposed that morphine-induced desensitisation of MOPr is extensive and prolonged primarily because the morphine-activated receptor undergoes protein kinase C-dependent desensitisation (Bailey et al. 2006) but does not undergo extensive internalisation and recycling (Schulz et al. 2004). DN dynamin has no significant effect on the rate of DAMGO-induced desensitisation of MOR1 and MOR1A hence it is only MOR1B that is recycled (and hence resensitised) rapidly enough to escape the rapid net desensitisation of the receptor. A previous study has reported marked differences in the agonist-induced regulation of the C-terminus splice variants of the mouse MOPr (Koch et al. 2001), but the mouse splice variants studied were either quite different from the rat splice variants (mouse MOR1D), have not been reported in rat (mouse MOR1E), or included mouse MOR1C (the rat form of which we were unable to detect convincingly).
In conclusion, we have found that mRNA for the MOPr isoforms MOR1, MOR1A and MOR1B are widely expressed throughout the rat CNS, with mRNA for MOR1 and MOR1A being generally expressed at higher levels than MOR1B. Each of these three splice variants underwent rapid desensitisation in the presence of morphine, however DAMGO-induced desensitisation of MOR1B was slower than for the other two splice variants, because of the ability of this spice variant to undergo rapid recycling. Importantly, we were unable to detect significant levels of mRNA for MOR1C, MOR1C1, MOR1C2 or MOR1D in rat CNS, which leaves open the question of their existence in vivo and consequently their physiological relevance in neuronal function.
This work was funded by the Wellcome Trust.
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