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The Journal of Neuroscience, December 1, 1998, 18(23):9685-9694
Axotomy Reduces the Effect of Analgesic Opioids Yet Increases the
Effect of Nociceptin on Dorsal Root Ganglion Neurons
Fuad A.
Abdulla1 and
Peter A.
Smith2
1 Department of Physical Therapy, Tennessee State
University, Nashville, Tennessee 37290, and 2 Department of
Pharmacology and Division of Neuroscience, University of Alberta,
Edmonton, Alberta, Canada T6G 2H7
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ABSTRACT |
There is some doubt as to the effectiveness of opioids in the
management of neuropathic pain. We therefore examined the actions of
morphine and the opioid-like peptide nociceptin (both 1 µ) on dorsal
root ganglion (DRG) neurons that were isolated from control or from
nerve-injured rats. Both substances reduced  -conotoxin (CTX)
GVIA-sensitive, N-type Ca2+ channel current
and small persistent nifedipine/ CTX-insensitive (non-N, non-L type)
current. Nifedipine-sensitive L-type current was unaffected. The effect
of nociceptin was antagonized by naloxone benzoylhydrazone (nalbzoh)
but not by naloxone. Sciatic nerve section (axotomy) profoundly
reduced the effects of morphine and the µ-receptor
agonist D-ala2,
N-Me-Phe4,Gly-ol5
enkephalin (DAMGO). The effect of the  -agonist
[(+)-(5 ,7,8 )-N-methyl-N-(7-(1-pyrrolidinyl)-1-oxaspiro(4,5)dec-8-yl)-benzeneacetamide] (U69593) was unchanged, whereas the effect of nociceptin was
increased. All agonists produced their strongest effects
on the small, putative nociceptive cells and their weakest effects on
the largest cells. The  -receptor agonist, enkephalin
D-pen2,5 (DPDPE), was without effect on
control or on axotomized cells. These and other data suggest that the
functional downregulation of µ-opioid receptors on sensory nerves
contributes to the poor efficacy of opioids in neuropathic pain. Also,
the increased effectiveness of nociceptin after axotomy supports the
hypothesis that its actions are mediated via a "non-opioid"
receptor. Pronounced suppression of Ca2+ channel
current in axotomized DRG neurons by nociceptin led to a reduction in
Ca2+-dependent K+ conductance and
a marked increase in excitability. Despite this, the spinal
administration of nociceptin or agonists that activate ORL1
(opioid-like orphan receptor) may prove to be of clinical interest in
the management of neuropathic pain.
Key words:
sympathetic pain; spinal analgesia; substantia
gelatinosa; superficial dorsal horn; C-fiber; causalgia; chronic
constriction injury
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INTRODUCTION |
The spinal analgesic action of
opioids involves reduction of neurotransmitter release from primary
afferent fibers (Jessell and Iversen, 1977 ), an effect attributed to
suppression of Ca2+ channel current/conductance
(ICa/gCa)
in sensory nerve terminals (Hori et al., 1992). This hypothesis is
supported by the observation that opioids suppress
high-voltage-activated (HVA) ICa or
Ca2+-dependent action potentials in the cell bodies
of dorsal root ganglion (DRG) neurons (Werz and MacDonald, 1982; Werz
et al., 1987; Moises et al., 1994a,b; Taddese et al., 1995; Womack and McCleskey, 1995). Despite the widespread use of opioids in the management of "nociceptive" pain, several clinical studies attest to their poor efficacy in "neuropathic" pain that is sometimes invoked by nerve injury (Arners and Megerson, 1988; Iadarola and Caudle, 1997). Also, immunohistochemical studies have shown that peripheral nerve section (axotomy) reduces the expression of µ- and
-opioid receptors in the cell bodies of sensory neurons and in their
terminal projections in the dorsal horn (De Groot et al., 1997; Zhang
et al., 1998a,b). Thus, limited clinical efficacy in neuropathic pain
may involve a reduction in the ability of opioids to suppress
ICa in sensory neurons. We therefore examined whether sciatic nerve section (axotomy) reduces the effect of morphine,
the selective µ-opioid ligand D-ala2,
N-Me-Phe4,Gly-ol5
enkephalin (DAMGO), the selective -opioid agonist -receptor agonist enkephalin D-pen2,5 (DPDPE), or
the -opioid agonist
[(+)-(5,7,8)-N-methyl-N-(7-(1-pyrrolidinyl)-1-oxaspiro(4,5)dec-8-yl)-benzeneacetamide] (U69593) on HVA ICa
(IBa) in the cell bodies of DRG neurons.
The limited clinical efficacy of morphine (Arners and Megerson,
1988) is reflected by a weak spinal analgesic effect in animal models
of neuropathic pain (Nichols et al., 1995; Ossipov et al., 1995;
Yamamoto and Nozaki Taguchi, 1996; Wegert et al., 1997). By contrast,
intrathecal injection of the opioid-like peptide nociceptin (orphanin
FQ) (Meunier et al., 1995; Reinscheid et al., 1995; Nothacker et al.,
1996) attenuates the thermal hyperalgesia produced by partial sciatic
nerve injury (Yamamoto et al., 1997). Nociceptin does not interact with
µ-, -, or -receptors but is instead an endogenous ligand for
the so-called "opioid-like orphan receptor"
(ORL1) (Meunier et al., 1995; Reinscheid et al.,
1995; Nothacker et al., 1996). Effects of nociceptin at
ORL1 are antagonized by the peptide
[Phe1 (CH2-NH)Gly2]
nociceptin (1-13) NH2 (Guerrini et al., 1998) but not by
naloxone (Calo et al., 1997). Naloxone benzoylhydrazone (nalbzoh) is a nonselective nociceptin antagonist that also blocks µ-receptors (Dunnill et al., 1996). Like morphine, nociceptin attenuates HVA gCa in DRG neurons (Abdulla and Smith 1997b). It
also attenuates glutamatergic transmission in the spinal cord (Faber et
al., 1996) and suppresses glutamatergic EPSPs in substantia gelatinosa
neurons (Lai et al., 1997), probably by a presynaptic action on
Ca2+ channels in primary afferent terminals (Liebel
et al., 1997). Because morphine analgesia is clearly attenuated after
peripheral nerve injury (Yamamoto and Nozaki Taguchi, 1996; Wegert et
al., 1997) whereas nociceptin continues to be effective [Yamamoto et al. (1997), but see also Hao et al. (1998)], we also examined the
effect of axotomy on nociceptin-induced suppression of HVA gCa in DRG neurons.
A preliminary report has been published previously (Abdulla and Smith,
1997c).
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MATERIALS AND METHODS |
Axotomy. The left sciatic nerve of adult male Sprague
Dawley rats (120-170 gm) under sodium pentobarbital anesthesia (50-55 mg/kg) was sectioned proximal to its bifurcation into the tibial and
the peroneal divisions. As described previously (Abdulla and Smith,
1997a), a 5-10 mm segment of the sciatic nerve was removed to prevent
regeneration. Animals were housed in separate cages and examined twice
daily for the first 3 d after surgery and then once daily for any
signs of postoperative stress. All protocols were approved by the
University of Alberta Health Sciences Animal Welfare Committee.
Electrophysiology. Rats were decapitated, and DRG neurons
from L4 and L5 were dissociated as described by White et al. (1989). These ganglia receive the majority of fibers from the sciatic nerve
(Swett et al., 1991). Cells were plated into 3 cm plastic Petri dishes
and used for recording within 2-10 hr. The cells were superfused with
various extracellular solutions at ~2 ml/min. For action potential
(AP) recording, external solution contained (in mM):
150 NaCl, 5 KCl, 2.5 CaCl2, 1 MgCl2, 10 HEPES-NaOH, pH 7.4, and 10 D-glucose; (osmolarity, 330-340 mOsm). Internal solution contained (in mM): 130 potassium gluconate, 2 Mg-ATP, 0.3 Na-GTP, 11 EGTA, 10 HEPES-KOH, pH 7.2, and 1 CaCl2,
(osmolarity 310-320 mOsm). Single APs were generated using a 2 msec
pulse of depolarizing current, and spike width was measured at 50% of
maximum amplitude. Excitability was measured by counting the number of
APs that discharged in response to 1 sec pulses of current at threshold
strength. Ba2+ (IBa)
was used as the charge carrier to record ICa.
For these experiments, external solution contained (in mM):
160 TEA-Cl, 10 HEPES, 2 BaCl2, 10 glucose, and 200 nM TTX, adjusted to pH 7.4 with TEA-OH; internal solution
contained (in mM): 120 CsCl2, 5 Mg-ATP,
0.4 Na2-GTP, 10 EGTA, 20 HEPES-CsOH, pH 7.2. IBa was evoked at  10 mV from a holding
potential of 90 mV. Leak subtraction procedures were deemed
unnecessary (Abdulla and Smith, 1997a). "Maximal outward"
(K+) current was determined form the
I-V relationship and recorded at +70 mV from a
Vh of 90 mV using an external solution that contained (in mM): 145 N-methyl-D-glucamine chloride (NMG-Cl), pH 7.4, 10 KCl, 2.5 CaCl2, 10 HEPES, 1.0 MgCl2, and 10 D-glucose; internal
solution contained (in mM): 100 potassium gluconate, 40 NMG-Cl, pH 7.2, 2 Mg-ATP, 0.3 Na-GTP, 11 EGTA, 10 HEPES, and 1.0 CaCl2. For recording Na+ currents
(INa), external solution contained (in
mM): 100 NaCl, 5 KCl, 4 MgCl2, 10 HEPES,
and 60 D-glucose, adjusted to pH 7.4 with NaOH; internal
solution contained (in mM): 140 CsCl2,
10 NaCl, 2 Mg-ATP, 0.3 Na-GTP, 2 EGTA, 10 HEPES, and 2 MgCl2, adjusted to pH 7.2 with NaOH.
Whole-cell recordings were made at 22°C using an Axoclamp 2A
amplifier in discontinuous voltage-clamp or bridge-balance
current-clamp mode. Borosilicate glass patch electrodes had DC
resistance of 4-6 M for AP recording or 1-3 M for current
recording. Sampling rates of 30-60 kHz and clamp gains >25 nA/mV
could be attained with these low-resistance electrodes. Because the
discontinuous voltage-clamp method allows the cell to be clamped to the
measured membrane voltage, it circumvents some of the series resistance problems that could be encountered if a conventional patch-clamp amplifier had been used. Because we did not use leak subtraction and
appropriate compensation circuitry is not available in the Axoclamp 2A
amplifier, the illustrated data records, which were filtered to 3 dB
at 1 or 3 kHz, display large capacitance transients. Under
current-clamp, input capacitance (Cin)
was calculated from the input resistance
(Rin) and the membrane time constant
( m) using the equation m = Cin · Rin. Under
voltage clamp, Cin was measured by integrating
the area of capacitative current transients that were generated by 10 mV commands ( V). This yielded the charge Q that is
related to Cin by Q = V · Cin. Data were acquired and analyzed using
Pclamp 5.5.1. software, and final records were produced using Microcal
Origin 4.1 or 5.0. Drugs were applied by superfusion, and all data are
presented as means ± SEM. Statistical significance was assessed
by Student's unpaired t test or  2 as
appropriate. Agonist-induced percentage changes in
IBa and outward current are calculated only from
those cells that responded. Thus, nonresponding cells that would have
0% change in IBa or maximal outward current are
excluded from the statistical analysis. This approach was not feasible
for description of changes in excitability because the smallest
possible change, i.e., a change from one spike to two spikes, would be
recorded as a 100% increase in excitability. Data for excitability
changes are therefore collected from all cells, and nonresponders are
entered into the averages as exhibiting a 0% increase in excitability.
Data on the percentage of cells responding to agonists in each category
are presented separately. DRG cells were classified into "large,"
"medium," and "small" subtypes on the basis of their AP shape
and/or Cin according to previously defined
criteria (Abdulla and Smith 1997a). The large cells were defined as
those with AP duration <3 msec and Cin of >90
pF; medium cells had an AP duration of 3-5 msec,
Cin of 70-90 pF, and an "inflection" on the
falling phase of AP; and the small cells had an AP duration >5 msec,
Cin of <70 pF, and a large inflection or
"shoulder" on the falling phase.
Drugs and chemicals. Nociceptin (rat or human;
Phe-Gly-GlyPhe-Thr-Gly-Ala-Arg-Lys-Ser-Ala-Arg-Lys-Leu-Ala-Asn-Gln)
was from Peptide Institute (Louisville, KY); naloxone hydrochloride,
nalbzoh, -conotoxin (CTX) GVIA, DAMGO, DPDPE, and U69593 were from
Research Biochemicals International (Natick, MA), morphine sulfate was from British Drug Houses (Toronto, Canada), and all other chemicals and
drugs were from Sigma (St. Louis, MO).
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RESULTS |
Effects of morphine and nociceptin on IBa
before and after axotomy
In neurons from control rats, morphine (1 µM) was
less effective on large cells than on medium or small cells. It
inhibited IBa in only 4 of 32 control large
cells by 22.6 ± 2.1%, in 13 of 36 control medium cells by
26.9 ± 1.6%, and in 21 of 41 small cells by 35.5 ± 2.0%.
Two to 4 weeks after axotomy, the inhibitory effects of morphine on
IBa were reduced. Therefore, morphine reduced IBa in 3 of 40 axotomized large cells by
21.2 ± 1.9%, in 6 of 47 medium cells (p < 0.02) by 17.8 ± 1.8% (p < 0.002), and
in 16 of 61 small cells (p < 0.01) by 23.2 ± 1.9% (p < 0.001).
Like morphine, nociceptin (1 µM) exerted weak effects on
large control cells and stronger effects on medium and small cells. It
inhibited IBa in 3 of 30 control large cells by
23.3 ± 3.9%, in 14 of 36 medium cells by 26.2 ± 1.3%, and
in 21 of 41 small cells by 34.7 ± 1.9%. By contrast with the
reduced effect of morphine seen after axotomy, the effects of
nociceptin were increased. Thus, nociceptin reduced
IBa in 6 of 40 axotomized large cells by
29.0 ± 1.5%, in 29 of 46 medium cells (p < 0.03) by 37.9 ± 2.1% (p < 0.001), and
in 45 of 60 small cells (p < 0.02) by 48.4 ± 2.0% (p < 0.001).
In most experiments, morphine and nociceptin were applied to the same
cell, and their relative effects were examined. Both drugs (at 1 µM) produced similar amounts of
IBa suppression in small, medium, and large
control cells. After axotomy, however, the response of any one cell to
nociceptin was larger than its response to morphine. Some typical data
records are illustrated in Figure
1A. Figure
1A1 illustrates superimposed recordings
of IBa obtained from the same small cell before
and after superfusion of morphine or nociceptin (both 1 µM). The amount of IBa suppression induced by morphine is very similar to that induced by nociceptin. Figure 1A2 illustrates recordings from an
axotomized small cell. The effect of morphine is small compared with
the effect of nociceptin. The data from these two cells are summarized
in Figure 1B. This shows the time course of the
suppression of IBa by both drugs in both cells.
The enhanced effect of nociceptin and the reduced effect of morphine in
the axotomized cell can be clearly seen. Figure 1C
summarizes the percentage suppression of IBa
induced by morphine or nociceptin in small (S),
medium (M), and large (L) cells
before and after axotomy, and Figure 1D shows the
percentage of cells responding in each category. These graphs
reemphasize the preferential effects of the drugs on the small cells
and the decrease in effectiveness of morphine and the increased
effectiveness of nociceptin after axotomy.
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Figure 1.
Effects of morphine and nociceptin on
IBa in DRG cells. A, Sample
records from small cells from control
(A1) and axotomized
(A2) animals. Currents were recorded
at 10 mV from a Vh of 90 mV. Tails were recorded at
40 mV, and voltage records were omitted for clarity. Both morphine
and nociceptin decreased IBa in the cells
illustrated both before and after axotomy. Note that the
IBa suppression induced by 1 µM morphine was similar to that induced by nociceptin in
the control cell, whereas in the axotomized cell the effect of morphine
was decreased and that of nociceptin was increased. B,
Time course of the effects of morphine or nociceptin on cells
illustrated in A1 and
A2 (data normalized). C,
Graphs to show percentage suppression of IBa
in large (L), medium
(M), and small (S)
neurons from control or axotomized animals in response to 1 µM morphine or nociceptin. D, Graphs show
percentage of cells in each category responding to 1 µM
morphine or to 1 µM nociceptin before or after
axotomy.
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Effects of µ-, -, and -agonists before and
after axotomy
To better characterize the effect of axotomy on the morphine
response, we examined changes in IBa suppression
by the selective µ-agonist DAMGO, the selective -agonist DPDPE,
and the selective -agonist U69593. The results are shown in Table
1. The -agonist DPDPE (1 µM) was without effect on IBa on
control DRG cells. It also failed to affect small, medium, or large
cells after axotomy. By contrast, the µ-agonist DAMGO and the
-agonist U69593 (both 1 µM) produced pronounced
IBa suppression in small control cells and
moderate effects on medium and large cells. After axotomy, however, the
effects of the µ-agonist, DAMGO were significantly reduced, whereas
those of the -agonist U69593 were essentially unchanged. Typical
experiments are illustrated in Figure 2.
Figure 2A show superimposed recordings of
IBa from the same control small cell before and
during application of DAMGO, U69593, or DPDPE (all 1 µM).
The current is suppressed by the µ- and -agonists but not by the
-agonist. Figure 2B illustrates a similar series of records obtained from an axotomized small cell. DAMGO produces very
modest suppression of IBa, the effect of
U69593 is comparable to that seen in a control cell, and the
-agonist DPDPE is without effect.
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Figure 2.
Effects of µ-, -, and -agonists on
IBa in control and axotomized DRG cells.
A, Effects of the µ-agonist DAMGO and the -agonist
U69593 and the lack of effect of the -agonist DPDPE (all 1 µM) on a small cell from a control animal.
B, Effects of the same three agonists on a small cell
from an axotomized animal. Note small response to DAMGO and lack of
effect of DPDPE. The response to U69593 is similar to that seen in the
control cell. Current and time calibrations refer to all records.
Currents evoked at 10 mV from Vh = 90
mV. Each panel shows superimposed recordings of
IBa evoked before and during superfusion of
drugs. Voltage command trace was omitted for clarity.
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Effects of -conotoxin GVIA and dihydropyridines on responses to
nociceptin and morphine
Agonists at µ-opioid receptors such as PL017
(Tyr-Pro-NMePhe-D-Pro-NH2)
inhibit N-type ICa
(ICa,N) in rat DRG neurons. In some
cells, this agonist also attenuates a sustained component of the
current that is insensitive to both -CTX GVIA and nifedipine (Moises
et al., 1994b; Rusin and Moises, 1995). To characterize which
components of HVA gBa in small cells were
affected by nociceptin and morphine, we examined the effect of -CTX
GVIA (1 µM; n = 5) and nifedipine (2 µM; n = 4). As might be expected, the
effects of morphine, which acts primarily on µ-opioid receptors
(Martin, 1983; Paternak, 1993), were similar to those of PL017.
The effects of nociceptin were indistinguishable from those of
morphine. Typical experiments are illustrated in Figure
3. Those illustrated in Figure
3A,C show that morphine and nociceptin do not affect L-type IBa (IBa,L).
Morphine or nociceptin (1 µM) reduced
IBa at 10 mV by a similar amount in the
presence or absence of 2 µM nifedipine. The experiments
illustrated in Figure 3B,D show that morphine and nociceptin
inhibit IBa,N as well as another small,
noninactivating, -CTX GVIA-resistant current. The marked reduction
of IBa at 10 mV induced by 1 µM
morphine or nociceptin was attenuated by -CTX GVIA. Because
suppression of current was still seen under these conditions, morphine
and nociceptin must act on a current other than
IBa,N. This current is not
IBa,L (Fig. 3A,C), so it is concluded that both agonists act on a nifedipine/-CTX GVIA-resistant current (Rusin and Moises, 1995). The lack of complete blockade of the effect
of effects of morphine and nociceptin by -CTX GVIA cannot be
attributable to use of a submaximal concentration of toxin, because 1 µM CTX GVIA blocks all effects of noradrenaline on
IBa in axotomized DRG neurons (Abdulla and
Smith, 1997a).
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Figure 3.
Effects of -CTX GVIA and nifedipine on
IBa suppression induced by morphine and
nociceptin. Superimposed current records in all parts of the figure are
IBa responses to a step to 10 mV from a
Vh of 90 mV in the presence and absence of
morphine or nociceptin (both 1 µM) and/or nifedipine (2 µM) or -CTX GVIA (1 µM). Time and
current calibration in first record applies to all traces; voltage
records were omitted for clarity. A, Left-hand data
records show suppression of current by morphine. Right-hand data
records comprise superimposed wash response (Wash) and
current recorded in the presence of nifedipine and again in the
presence of morphine plus nifedipine. The amount of IBa suppression produced is
seen most clearly from the time course graph. This shows amplitudes of
successive IBa responses recorded once every
20 sec in the presence of morphine, nifedipine, and nifedipine plus
morphine. Nifedipine reduces the total current but does not impair the
action of morphine. B, Left-hand data records are from
another small cell, again showing suppression of current by morphine.
Right-hand data records comprise superimposed wash response
(Wash) and current recorded in the presence of -CTX
GVIA and again in the presence of morphine plus -CTX GVIA. The time
course graph shows that morphine is much less effective in attenuating
the total IBa in the presence of -CTX
GVIA. However, a clear reduction of the current is still observed.
C, Data records and time course of an experiment
performed with nociceptin and nifedipine. The ability of nociceptin to
inhibit total IBa is not impaired by
nifedipine. D, Data records and time course of an
experiment performed with nociceptin and -CTX GVIA. Although the
toxin impairs nociceptin-induced IBa
suppression, the effect is not completely blocked.
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Effects of morphine and nociceptin on the excitability of
DRG cells
Noradrenaline and neuropeptide Y (NPY) suppress
gBa,N in DRG neurons (Abdulla and Smith,
1997a,b, 1999) in much the same way as nociceptin and morphine. In the
case of noradrenaline, this effect is only seen after axotomy, and in
the case of NPY the effect is much greater after axotomy. Suppression
of gCa,N leads to attenuation of
Ca2+-sensitive K+ conductance(s)
(gK,Ca) and increases in
excitability that may contribute to spontaneous activity in injured
sensory nerves (Abdulla and Smith, 1997a, 1999). We therefore examined
whether nociceptin and morphine can also increase excitability of DRG
cells and how these effects might be changed after axotomy.
AP discharge was invoked using a 1 sec pulse of current at the
threshold strength in the presence or absence of morphine or nociceptin
(both 1.0 µM). Figure
4A1
illustrates typical increases in excitability induced by morphine and
nociceptin on the same small control cell. Morphine and nociceptin
produced comparable effects in control cells at this concentration, and
the relative sensitivity of cell types paralleled that seen with
effects on IBa. Thus, large cells were affected
less frequently and to a lesser extent than medium or small cells.
Morphine increased the firing rate of only 1 of 26 control large cells,
whereas nociceptin increased the firing rate of 2 of 24 cells. Morphine
increased the excitability of 17 of 36 medium control cells by 42.3%
and of 17 of 36 small cells by 50.9%. Similarly, nociceptin increased the excitability of 16 of 36 medium cells by 41.7% and of 17 of 35 small cells by 49.5%. Two to 4 weeks after axotomy, the effects of
morphine on excitability were reduced. Morphine increased
the excitability of 1 of 25 large cells by 2.6%, 7 of 32 medium cells (p < 0.03) by 31.3%, and 10 of 41 small cells
(p < 0.04) by 17.3% (p
values from 80 2 tests to compare number of cells of
each type affected before or after axotomy).
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Figure 4.
Effects of morphine or nociceptin on the
excitability of DRG cells. A, Sample records from a
control small cell (A1) and an
axotomized small cell (A2)
illustrating the effects of 1 µM morphine and 1 µM nociceptin on excitability. To generate APs, both
cells were depolarized with a 1 sec pulse of current at threshold
strength (current trace omitted for clarity). Note that both morphine
and nociceptin increase the excitability of the control cell in a
similar way; however, in the axotomized cell, the response to morphine
was much less than the response to nociceptin. B, Time
course of the effects of morphine and nociceptin on cells illustrated
in A1 and A2
(data normalized). C, Graphs summarize effects of
morphine or nociceptin on the percentage change in numbers of spikes
discharged by 1 sec pulses of current at threshold strength in large
(L), medium (M), and
small (S) neurons from control or axotomized
animals. D, Graphs show percentages of large, medium,
and small neurons from control or axotomized animals that respond to
morphine or nociceptin.
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By contrast, the effects of nociceptin on excitability were
increased after axotomy. At 1 µM it increased
the excitability of 5 of 25 large cells by 24.0%, 23 of 32 medium
cells (p < 0.025) by 89.8%, and 30 of 40 small
cells (p < 0.02) by 111.7%
(p values from 80 2 tests to compare
number of cells of each type affected before or after axotomy). This
finding is consistent with the observed increased effectiveness of
nociceptin in suppressing IBa after axotomy
(Fig. 1).
Figure 4A2 illustrates the small effect
of morphine on the excitability of a small axotomized cell and the
profound increase in excitability produced by nociceptin in the same
cell. The data from these two cells are summarized in Figure
4B, which shows the time course of the increase in
excitability induced by both drugs. The enhanced effect of nociceptin
and the reduced effect of morphine in the axotomized cell can be
clearly seen. Figure 4C summarizes the percentage increase
in number of APs (spikes) induced by morphine or nociceptin in control
small (S), medium (M), and large
(L) cells before and after axotomy, and Figure 4D shows the percentage of cells responding in each
category. These graphs reemphasize the preferential effects of the
drugs on the small cells and the decrease in effectiveness of morphine and the increased effectiveness of nociceptin after axotomy.
Pharmacology of the effects of morphine and nociceptin
The effect of morphine on IBa and on
excitability of DRG cells excitability was antagonized by the broad
spectrum µ-, -, and -opioid antagonist naloxone (1 µM; n = 4 for both). Effects mediated via
ORL1 receptors are unaffected or relatively insensitive to
inhibition by naloxone (Connor et al., 1996a,b; Faber et al., 1996;
Calo et al., 1997). In agreement with this, naloxone (up to 1 µM) failed to antagonize the effects of nociceptin on
IBa (n = 4) or on excitability
(n = 4). On the other hand, the
3-agonist nalbzoh has been reported to exert an
antagonist action at ORL1 receptors (Dunnill et al., 1996).
Nalbzoh (1 µM) antagonized the effects of nociceptin on
excitability (n = 4) and on IBa
(n = 5). Nalbzoh also has significant antagonist action
at µ-opioid receptors (Dunnill et al., 1996), and we found that it
antagonized the actions of morphine on IBa
(n = 4) and on excitability (n = 4). We
have not yet tested whether the peptide
[Phe1(CH2-NH)Gly2]
nociceptin (1-13) NH2 (Guerrini et al., 1998) can
selectively antagonize nociceptin responses on DRG neurons.
Effects of morphine and nociceptin on K+ and
Na+ currents
It has been assumed that the increase in excitability induced by
nociceptin and morphine result from suppression of
gK,Ca as a consequence of their action on
gCa,N. To test this, we examined their effect on
Ca2+-sensitive and Ca2+
-insensitive components of the maximal outward current at +70 mV. This
current reflects outward movement of K+ through
delayed-rectifier (IK), A-type
K+ channels and through gK,Ca
channels (Akins and McCleskey, 1993). Both morphine and nociceptin (1 µM) decreased the maximal outward current (at +70 mV from
Vh = 90 mV). In control cells, morphine decreased the
total outward current in 2 of 23 large cells by 18.8 ± 2.2%, 13 of 33 medium cells by 23.4 ± 1.7, and in 17 of 32 small cells by
25.5 ± 1.4%. Similarly, nociceptin reduced the maximal outward
current in 2 of 21 control large cells by 19.6 ± 4.4%, 12 of 31 medium cells by 23.8 ± 2.0%, and 16 of 32 small cells by
25.8 ± 1.6%. After axotomy, the effects of morphine on maximal
outward current were decreased, and the effects of nociceptin were
increased. Thus morphine decreased the total outward current in 2 of 27 large cells by 16.4 ± 2.0%, in 5 of 30 medium cells (p < 0.05) by 18.0 ± 1.2%
(p < 0.02), and in 9 of 35 small cells by
19.0 ± 1.6% (p < 0.007), whereas
nociceptin reduced it in 4 of 45 large cells by 22.0 ± 3.0%, 20 of 30 medium cells (p < 0.03) by 30.5 ± 1.4% (p < 0.01), and in 26 of 35 small cells
(p < 0.04) by 33.6 ± 1.7%
(p < 0.002).
The Ca2+ channel blocker Cd2+
(0.5 or 1.0 mM) reduced total outward current recorded at
+70 mV and occluded the effects of morphine and nociceptin (both at 1 µM) on the current that remained
(Vh = 90 mV; n = 10-15 for
each cell type, for both morphine and nociceptin from control and
axotomized animals). This shows that the actions of morphine and
nociceptin are exerted on gK,Ca because these
currents that depend on Ca2+ influx are blocked by
the extracellular application of Cd2+.
Some typical experiments are illustrated in Figure
5A. Illustrations of maximal
outward current in a small cell recorded before and during application
of 1 µM morphine or nociceptin are shown in Figure
5A1. Both agonists produce similar amounts of
suppression of the current. In the presence of 0.5 mM
Cd2+, the effects of both agonists are blocked.
Figure 5A2 illustrates a similar experiment
performed on an axotomized small cell. Nociceptin is clearly more
effective than morphine, and the effects of both drugs are prevented by
Cd2+. Figure 5B summarizes the time
course data from the experiments performed on the two cells shown in
Figure 5A1,A2. The similar effects of both drugs on the control cell is clearly seen as is the
large effect of nociceptin and the small effect of morphine after
axotomy. Figure 5C summarizes the data from all cells that responded. Again the decreased effectiveness of morphine and the increased effectiveness of nociceptin after axotomy can be seen. This
trend is also apparent from Figure 5D, which illustrates the
percentage of cells responding to each drug before and after axotomy.
As with IBa and with excitability changes, both
nociceptin and morphine exert their strongest effects on small cells
and their weakest effects on large cells. Also the differential effects of axotomy on responses of outward current to morphine and nociceptin parallel those seen with IBa and with
excitability.
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Figure 5.
Effects of morphine or nociceptin (both 1 µM) on maximal outward current recorded at +70 mV
(Vh = 90 mV). A,
Superimposed data records of suppression of maximal outward current by
1 µM morphine or nociceptin in a control
(A1) and in an axotomized
(A2) small cell (voltage commands
omitted for clarity). After initial suppression and recovery of current
by morphine or nociceptin, the current is again suppressed after
superfusion of 1 mM Cd2+ to block
gK, Ca responses. Note that in the control
cell morphine and nociceptin produced similar effects on the outward
current; however, after axotomy the effect of morphine was much smaller
than that of nociceptin. Neither morphine nor nociceptin has effects on
outward current recorded in the presence of Cd2+.
B, Time course of the effects of morphine and nociceptin
on cells illustrated in A1 and
A2 (current is normalized for convenient
comparison). C, Graphs summarizing effects of morphine
and nociceptin (1 µM) on maximal outward current
amplitude in large (L), medium
(M), and small (S)
neurons from control or axotomized animals. D, Graphs
show percentages of large, medium, and small neurons from control or
axotomized animals that respond to morphine or nociceptin (both 1 µM).
|
|
Neither morphine nor nociceptin (1 µM) affected
INa in any of the cell types either before or
after axotomy. Experiments were performed on 61 cells from control
animals and on 54 cells from axotomized animals.
| |
DISCUSSION |
The primary finding of this work is that morphine becomes less
effective and nociceptin becomes more effective in suppressing ICa,N in DRG cell bodies after axotomy. These
findings contribute to the understanding of the weak clinical efficacy
of morphine in neuropathic pain. In addition, they lend support to the
idea that spinal administration of nociceptin may prove of use in the management of nerve injury-induced pain, which is typically chronic and
often intractable.
Because effects mediated via -receptors are unchanged by axotomy and
neither control nor axotomized cells respond to -agonists, attenuation of the effect of morphine results from reduction of effects
mediated through µ-receptors. This conclusion, which is supported by
the observed decreased effectiveness of DAMGO, presumably reflects
downregulation of µ-receptor expression (Zhang et al., 1998b).
Because axotomy reduces µ-receptor-like immunoreactivity both in DRG
cell bodies and in the dorsal horn of the spinal cord (De Groot et al.,
1997; Zhang et al., 1998b), we suggest that the limited effectiveness
of morphine as a spinal analgesic in nerve injury models results from
impairment of its ability to suppress ICa in
sensory nerve terminals. This possibility now needs to be examined more
directly by testing whether nerve injury impedes opioid-induced
suppression of excitatory synaptic transmission in the superficial
laminae of the dorsal horn (Hori et al., 1992).
The lack of the effect of the -agonist DPDPE on
IBa in control neurons confirms previous
findings from other laboratories (Schroeder et al., 1991; Moises et
al., 1994b) and fits with observation that -opioid receptor-like
immunoreactivity is localized intracellularly to the Golgi complex and
to vesicular membranes in DRG neurons (Zhang et al., 1998a).
The observation that axotomy decreases or fails to alter responsiveness
to µ-, -, or -opioids yet selectively increases responsiveness
to nociceptin strongly supports the hypothesis that it acts at a
"nonopioid" receptor (Guerrini et al., 1998), presumably
ORL1 (Meunier et al., 1995; Reinscheid et al., 1995; Nothacker et al., 1996; Henderson and McKnight, 1997). Our findings also raise the possibility that injury increases the
effectiveness of nociceptin at primary afferent terminals. Two recent
reports are consistent with this. Jia et al. (1998) found increased
nociceptin binding in dorsal horn in inflammatory pain (complete
Freund's adjuvant-rat hindpaw model), and Yamamoto et al. (1997)
reported that nociceptin attenuates thermal hyperalgesia induced by
unilateral sciatic nerve constriction in rats. It is therefore now
necessary to test directly whether nerve injury increases the ability
of nociceptin to attenuate synaptic transmission in the dorsal horn (Lai et al., 1997; Liebel et al., 1997). Such experiments may also
resolve the findings of Yamamoto et al. (1997) and Jia et al. (1998)
with those of Hao et al. (1998) who reported attenuation of
nociceptin-induced analgesia in two different neuropathic pain models.
Despite this inconsistency, which may devolve from the different types
of pain models used, the possible use of nociceptin as a spinal
analgesic in neuropathic pain is clearly worthy of further investigation.
The finding that morphine, DAMGO, U69593, and nociceptin exert their
strongest effects on small and medium cells confirms and extends the
previous results of Taddese et al. (1995) who showed that µ-opioids
have a preferential action on gCa of cells that
are involved in the transfer of nociceptive information [also see
Bessou and Perl (1969)]. Another factor that may contribute to the
limited effectiveness of morphine in neuropathic pain is its inability
to produce effects on large cells after axotomy (Fig. 1C,D).
This is because nerve injury promotes reorganization of non-nociceptive
primary afferents so that they make contact with ascending nociceptive
pathways in the dorsal horn (Woolf et al., 1982). Thus,
innocuous sensory activity may be transmitted via pathways that
normally carry nociceptive information. This may account for the
phenomenon of mechanical allodynia, which sometimes accompanies
peripheral nerve injury. The fact that nociceptin becomes more
effective on all cell types, including large cells, after axotomy
supports the idea that spinal administration of ORL1
agonists may be useful in the management of neuropathic pain. Interestingly, Hao et al. (1998) recently reported that intrathecal nociceptin dose-dependently alleviates mechanical and cold
allodynia-like behavior in two different models of neuropathic pain.
Intrathecal (spinal) administration of nociceptin thus
produces analgesia in various pain models (Henderson and McKnight, 1997; Hao et al., 1998), and some reports attest to its efficacy in
neuropathic pain (Yamamoto et al., 1997; Jia et al., 1998). Despite
this, there are conflicting reports as to the effect of intracerebroventricular administration. Although initial
results suggested that intracerebroventricular nociceptin
induced hyperalgesia (Meunier et al., 1995; Reinscheid et
al., 1995), more recent studies (for review, see Henderson and
McKnight, 1997) report analgesic actions, anti-opioid actions, and
biphasic hyperalgesic/analgesic effects. Any capacity of nociceptin to
produce hyperalgesia presents something of a paradox because the vast
majority of its cellular neurophysiological actions resemble those of
classic, analgesic opioids, i.e., it decreases transmitter release,
activates an inwardly rectifying K+ conductance
(gKIR), and suppress HVA
gCa in various types of spinal cord, brain, or
DRG neurons (Connor et al., 1996a,b; Faber et al., 1996; Vaughan and
Christie 1996, 1997; Abdulla and Smith, 1997b; Liebel et al., 1997;
Vaughan et al., 1997). Such effects are indistinguishable from those of
morphine (Jessell and Iversen, 1977; Cherubini and North, 1985;
Hescheler et al., 1987; Williams et al., 1988; Surprenant et al., 1990;
Schroeder et al., 1991; Hori et al., 1992; Moises et al., 1994a,b;
Taddese et al., 1995; Womack and McCleskey, 1995). Unlike morphine,
however, nociceptin suppresses T-type Ca2+ current
in DRG neurons (Abdulla and Smith, 1997b), but any relevance of this
observation to its reported, central hyperalgesic action remains to be
established. Apart from the fact that sensitivity of DRG neurons to
nociceptin was increased after axotomy whereas their sensitivity to
morphine was decreased, we found no differences in the action of
morphine and nociceptin. Both agonists affected gBa,N and -CTX GVIA/nifedipine-resistant
current to the same extent. Our findings therefore illustrate more
similarity than difference between the cellular actions of morphine and
nociceptin. Receptors for both agonists are preferentially expressed on
small DRG cells, and each individual neuron that expresses
µ-receptors usually also expresses ORL1. Moreover, these
two receptors seem to couple to exactly the same effectors. The
observations that morphine suppresses the N- and non-N, non-L
components of HVA gCa are similar to those
previously reported with the selective µ-receptor agonist PL017
(Rusin and Moises, 1995).
Spontaneous activity in damaged primary afferent fibers can contribute
to chronic neuropathic pain (Devor et al., 1994). Such activity is
enhanced by noradrenaline (Abdulla and Smith, 1997a) that is released
from sympathetic nerves that sprout into axotomized ganglia (McLachlan
et al., 1993). Although nociceptin suppresses gCa,N, reduces
gKCa, and thereby excites axotomized DRG
cells in a manner similar to noradrenaline (Fig.
4A2), it still acts as an effective
analgesic when it is administered intrathecally to nerve-injured rats
(Yamamoto et al., 1997). This presumably reflects suppression of
gCa,N in primary afferent terminals (Liebel et
al., 1997) and hyperpolarization of substantia gelatinosa neurons (Lai
et al., 1997). Because its intracerebroventricular actions also remain
to be resolved (Meunier et al., 1995; Reinscheid et al., 1995;
Henderson and McKnight, 1997), nociceptin appears to exert only a clear
analgesic effect at the spinal level. Any possible clinical use of
ORL1 agonists in neuropathic pain would therefore be
restricted to spinal routes of administration. This is not altogether
infeasible because various techniques for chronic spinal administration
of drugs in humans are now available (Du Pen, 1998).
| |
FOOTNOTES |
Received June 4, 1998; revised Sept. 16, 1998; accepted Sept. 17, 1998.
This work was supported by the Alberta Paraplegic Foundation, The Rick
Hansen Man-in-Motion Foundation, and the Medical Research Council of
Canada. Dr. Abdulla gratefully acknowledges fellowship support from the
Alberta Heritage Foundation for Medical Research. We thank Dr. W. F. Colmers and Mr. Tim Moran for their comments on an early version of
this manuscript.
Correspondence should be addressed to Dr. Peter A. Smith, Department of
Pharmacology, 9.75 Medical Sciences Building, University of Alberta,
Edmonton, Alberta, Canada, T6G 2H7.
| |
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Copyright © 1998 Society for Neuroscience 0270-6474/98/18239685-10$05.00/0
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Abstract
Introduction
