Propofol (2,6-diisopropylphenol) is an IV anesthetic that is widely used for the induction and maintenance of general anesthesia. The enhancement of GABAergic and glycinergic neurotransmission by propofol is thought to be responsible for its general anesthetic properties.1 It has been debated whether propofol at subhypnotic concentrations has analgesic properties.2 Animal studies have reported systemic propofol to have either no effect3–5 or antinociceptive and/or antihyperalgesic effects.6–8 Systemic administration of propofol depressed noxious stimulus–evoked responses of neurons in the spinal cord dorsal9–13 and ventral14,15 horns and either reduced16 or had no effect5 on formalin-evoked spinal neuronal expression of c-fos, a marker of neuronal activity. Several studies of experimental pain in humans reported analgesic effects of subhypnotic doses of propofol.17–20 Moreover, some recent clinical studies have reported that surgical patients receiving propofol anesthesia experienced reduced postoperative pain21,22 (see also Ref. 23).
Based on 3-compartment models of drug pharmacokinetics,24,25 it would be expected that propofol, which is highly lipid-soluble, would be cleared more slowly from peripheral tissues when the infusion is stopped. If propofol exerted a peripheral antinociceptive effect, this might account for its postsurgical analgesic effect. Peripheral application of propofol has been reported to have antinociceptive effects in the formalin26 and bee venom tests.12 In the latter study, peripherally injected propofol also inhibited the responses of nociceptive dorsal horn neurons. In the present study, we hypothesized that propofol in peripheral tissue such as skin exerts an antinociceptive effect. To mimic the action of residual propofol in peripheral tissues after the cessation of propofol infusion, we applied propofol topically to the skin and investigated potential antinociceptive and antihyperalgesic actions using behavioral and electrophysiological approaches in rats.
All experiments were conducted using adult male Sprague-Dawley rats under protocols approved by the University of California Davis Institutional Animal Care and Use Committee. Rats were housed individually and had unrestricted access to food and water.
These studies used 8 rats (body weight: 510–696 g). Thermal and mechanical paw withdrawal tests were conducted as previously described.27 For thermal paw withdrawal (Hargreaves)28 testing, rats were first habituated over 3 successive daily sessions to stand on a glass surface heated to 30°C ± 1°C within a ventilated Plexiglas enclosure. Paw withdrawal latencies were measured by directing a light beam (Plantar Test 390; IITC Life Science, Woodland Hills, CA) onto the plantar surface of the hindpaw through the glass plate from below, and the latency from onset of the light to brisk withdrawal of the stimulated paw was measured. To prevent potential tissue damage, a cutoff of 18 seconds was imposed. For formal testing, either vehicle (10% intralipid; Fresenius Kabi, Uppsala, Sweden) or propofol (Sigma-Aldrich Co., St. Louis, MO) at a concentration of 1%, 10%, or 25% (dissolved in 10% intralipid) was applied to the ventral surface of one hindpaw using a cotton swab and allowed to dry for 2 minutes, after which the rat was placed onto the glass surface. Withdrawal latencies for both the treated and the untreated paws were measured at 5, 15, 30, 45, 60, and 120 minutes after application.
We also tested the effect of propofol on allyl isothiocyanate (AITC)-induced hyperalgesia. AITC (mustard oil, 50%, in mineral oil; Sigma) was applied topically to one hindpaw in 3 treatment groups: (a) AITC alone (no pretreatment); (b) AITC, followed 10 minutes later by intralipid; or (c) AITC, followed 10 minutes later by topical 10% propofol. All chemicals were applied topically to the hindpaw, and withdrawal latencies for the treated and untreated (contralateral) paws were measured as described above.
For mechanical paw withdrawal threshold measurements, rats were habituated over 3 successive days to standing on a wire mesh screen surface. Baseline mechanical withdrawal thresholds were assessed using an electronic von Frey filament (1601C; IITC Life Science) pressed against the plantar surface of one hindpaw. This device registered the force (g) at the moment that the hindpaw was withdrawn away from the filament. Vehicle or propofol was then applied to one hindpaw, and mechanical paw withdrawal thresholds were measured at the same postapplication time points as noted above. Topical application of propofol was used because of its permeation through rat skin29,30 and to avoid potential damage from an intraplantar injection.
Thermal withdrawal latencies and mechanical paw withdrawal thresholds were normalized to baseline averages. Treatment groups were compared using 2-way repeated-measures analysis of variance (ANOVA) followed by Tukey post hoc comparison tests.
Thirty-eight rats (body weight: 400–510 g) were used. Anesthesia was induced with sodium pentobarbital (65 mg/kg, intraperitoneally) and maintained by constant infusion of pentobarbital via a jugular vein catheter at a rate sufficient to maintain areflexia for the duration of the experiment (10–20 mg/kg/h). Oxygen was delivered via a tracheal cannula. Core body temperature was monitored and maintained by heating pad. A laminectomy exposed the lumbar spinal cord for single-unit recording as detailed previously.31 Briefly, a tungsten microelectrode (10 MΏ; FHC, Bowdoin, ME) was driven into the dorsal horn to record extracellular single-unit activity. We specifically searched for mechanoresponsive units that additionally responded to noxious skin heating. Thermal sensitivity was assessed using a Peltier thermode (0.5-in. diameter, NTE-2A; Physitemp, Clifton, NJ) programmed to deliver heat and cold stimuli to the cutaneous mechanical receptive field. The skin-thermode interface temperature was monitored using a thermocouple (IT-21; Physitemp) connected to a BAT-12 (Physitemp) thermometer and was displayed along with single-unit activity and electrocardiogram using Powerlab (ADInstruments, Colorado Springs, CO) and Spike2 (Cambridge Electronic Design, Cambridge, UK) interfaces. The heat stimulus was increased from an adapting temperature of 34°C to 52°C over 15 seconds, followed by recooling to 34°C. A cooling stimulus followed 2 minutes later (34°C–0°C over 45 seconds). Units were additionally tested for mechanical sensitivity using a series of von Frey monofilaments (bending force: 0.68–1258.9 mN) applied in ascending order: cotton wisp, touch, and pinch applied to the receptive field. All of the present units were classified as wide dynamic range (WDR), based on their thermal sensitivity and incrementally increasing responses to graded mechanical stimuli.
To assess effects of propofol on thermally evoked responses, the following protocol was used. The heat and cold stimuli sequence was first delivered before any chemical stimulation. Two minutes later, the Peltier thermode was withdrawn from the skin and 10% propofol (30 μL) or vehicle (30 μL) was applied topically to the hindpaw receptive field by pipette. The thermode was replaced at the same location 3 minutes after the chemical application, and the heat and cold stimuli sequence was delivered again 5, 10, 15, and 30 minutes after 10% propofol or vehicle. The thermode was mounted on a micromanipulator (World Precision Instruments, Sarasota, FL) to allow precise repositioning of the thermode surface at the same skin site for postchemical thermal testing. Additional concentrations of propofol (1%, 25%) were tested in the identical manner out to 15 minutes after application.
We additionally assessed the effect of topical propofol on sensitization of heat-evoked responses induced by topical application of AITC. The heat and cold stimuli sequence was applied first as above. Two minutes later, AITC (10 μL, 75% in mineral oil; Sigma) was applied topically to the ventral hindpaw. Five minutes after AITC application, either 10% propofol or vehicle was applied as above. The heat and cold stimuli sequence was applied 5, 10, 15, and 30 minutes after propofol or vehicle application. In many experiments, a second unit was subsequently recorded on the opposite side of the spinal cord and one of the protocols described above was followed on the opposite hindpaw. As previously reported,31 responses recorded from the second unit did not exhibit any indication of sensitization. At the conclusion of each unit recording session, the recording site was marked by electrolytic lesion, and the spinal cord was postfixed in 10% buffered formalin. Sections of the spinal cord were cut on a freezing microtome and examined under the light microscope.
Action potentials were recorded, stored, and analyzed using Spike2 (Cambridge Electronic Design). In a few instances, 2 action potentials were recorded simultaneously and were sorted by spike size and waveform. The number of thermally evoked action potentials was summed over 30 seconds for heat stimuli and 45 seconds for cold stimuli and baseline-corrected by subtracting the number of spontaneous action potentials counted over the same time period before the thermal stimulus. Thermal thresholds were defined as the temperature at which the unit activity changed by >2 SDs from the prestimulus firing rate. For mechanical testing, the firing rate 15 seconds immediately before each stimulus was summed and subtracted from the firing rate 15 seconds after application of the mechanical stimulus. In most cases, unit responses were normalized to the pre-propofol or pre-AITC baseline, and were analyzed using 1-way repeated-measures ANOVA with Tukey post hoc comparison tests between time points. Statistical analyses were conducted using GraphPad Prism software (GraphPad, La Jolla, CA) or SPSS 9.0 software (SPSS Inc., Chicago, IL), where P < 0.05 was considered significant. Error reported is the SEM.
The hindpaw receiving topical propofol (ipsilateral) exhibited a concentration-dependent increase in withdrawal latency (Fig. 1A). The group tested with 25% propofol was significantly different from vehicles and 1% propofol (Fig. 1A; P < 0.05, 2-way repeated-measures ANOVA, Tukey post hoc), with a significant increase in latency at 5 minutes after propofol (P < 0.05, 1-way ANOVA with Tukey post hoc). Withdrawal latencies for the hindpaw contralateral to the side of propofol application were not affected at any concentration of propofol (Fig. 1B; 2-way repeated-measures ANOVA, P > 0.1). There was no significant effect of propofol at any concentration on von Frey mechanically evoked withdrawal thresholds for the ipsilateral (Fig. 1C) or contralateral (Fig. 1D) hindpaw.
Topical hindpaw application of AITC resulted in a significant decrease in paw withdrawal latency that was not significantly affected by topical postapplication of intralipid (Fig. 2A). However, topical application of 10% propofol 10 minutes after AITC resulted in increased paw withdrawal latency that was significant at 15 minutes (Fig. 2A; P < 0.05, Tukey post hoc) and prevented hyperalgesia as evidenced by a significant difference between the AITC-only versus propofol + AITC group (P < 0.05, 2-way repeated-measures ANOVA). None of the treatments significantly affected contralateral paw withdrawal latency (Fig. 2B).
A total of 62 heat-responsive dorsal horn units were recorded. All units were of the WDR type, responding to innocuous mechanical stimulation as well as noxious heat. An example is shown in Figure 3A. Units were mainly located in lamina I, with some in the deeper dorsal horn (Fig. 3B), at a mean depth of 214 ± 37 μm (SEM) below the surface.
Heat-evoked responses of dorsal horn units were significantly reduced by topical application of 10% and 25% (but not 1%) propofol. Figure 3A shows an example of a unit for which topical application of 10% propofol resulted in a time-dependent decline in the magnitude of the noxious heat-evoked response over the first 10 to 15 minutes, followed by recovery 30 minutes later. Figure 3C plots the mean normalized heat-evoked responses of 11 units at various times relative to topical application of propofol, which resulted in a significant reduction at 15 minutes (P < 0.05, 1-way repeated-measures ANOVA with Tukey post hoc test). Topical application of vehicle (10% intralipid) did not affect heat-evoked responses at any time point (Fig. 3D). No unit was directly excited by application of 10% propofol. Figure 4 summarizes the data, showing that 1% propofol was ineffective whereas both 10% and 25% propofol concentrations significantly attenuated heat-evoked responses at the 15-minute time points after propofol, with significant inhibition also at 10 minutes after 25% propofol (P < 0.05, 1-way repeated-measures ANOVA with Tukey post hoc test). Neither 10% nor 25% propofol significantly affected thermal thresholds (10% propofol: pretreatment 42.1°C ± 0.3°C [SEM]; posttreatment 42.6°C ± 0.4°C; 25% propofol: pretreatment 41.4°C ± 0.4°C, posttreatment: 42.5°C ± 0.8°C).
Twenty-nine heat-responsive units also responded to the cold stimulus. Responses elicited by the cold stimulus were not significantly affected by topical hindpaw application of 1%, 10%, or 25% propofol. Likewise, none of the propofol concentrations significantly affected mechanically evoked responses of 29 WDR units tested (Fig. 5).
We additionally tested whether propofol affects AITC enhancement of heat-evoked responses. When AITC was applied topically to the hindpaw followed by topical application of vehicle, there was a subsequent significant increase in the magnitude of the heat-evoked response (Fig. 6A), consistent with our previous report.31 However, when AITC was applied topically followed by topical application of 10% propofol, there was no change in the magnitude of heat-evoked responses compared with the pre-AITC + propofol response (Fig. 6B), indicating that topical propofol reduced the hyperalgesic effect of AITC.
In the present study, topical application of propofol dose-dependently suppressed thermal paw withdrawals without affecting mechanical sensitivity, indicating that topical propofol acts as an analgesic but not as a local anesthetic. Propofol also suppressed noxious heat-evoked responses of WDR dorsal horn neurons, without affecting their responses to mechanical or cold stimuli. WDR neurons encode the intensity of noxious stimuli32 and seem to be sufficient for pain sensation.33 The correspondence between the inhibitory effect of propofol on nocifensive behavior and neuronal responses supports the argument that WDR neurons are involved in signaling heat pain. We additionally observed that topical propofol reduced or prevented AITC enhancement of thermal paw withdrawals and heat-evoked responses of WDR neurons, implying an antihyperalgesic effect. We believe that the observed analgesic and antihyperalgesic effects are mediated locally in skin tissue. Although we did not presently measure tissue or plasma propofol concentrations, it was previously reported that transdermal application of propofol at concentrations comparable to the highest concentration used presently (25%) resulted in plasma concentrations in the 50 to 300 ng/mL range,30 which is well below the range of plasma propofol concentrations for sedation (3–8 µg/mL). These findings are discussed below in relation to potential mechanisms underlying the antinociceptive and antihyperalgesic effects of topical propofol.
Our finding that topical application of propofol induced antinociception and inhibited spinal WDR neurons is consistent with a previous report that peripherally applied propofol reduced nocifensive behavior and spinal neuronal activity elicited by subcutaneous injection of bee venom.12 However, these findings are in marked contrast to the well-known algesic effects of propofol, which induces pain upon IV34 or intradermal35 injection in humans and elicits nocifensive behavior in mice in a TRPA1-dependent manner.36 Propofol directly activates TRPV1 and TRPA136–38 with a greater effect on TRPA1.35 Propofol also reversed desensitization of TRPV1 as manifested by resensitization of capsaicin-evoked responses in dorsal root ganglion cells and prolongation of capsaicin-evoked nocifensive behavior38 in a TRPA1-dependent manner.39 It is thus difficult to reconcile these pronociceptive actions of propofol with the presently observed antinociceptive effects. Repeated or continuous application of propofol led to desensitization of TRPV1 expressed in human embryonic kidney cells, and high concentrations of propofol blocked TRPA1 activation.35 Thus, it is possible that high concentrations of topically applied propofol may desensitize TRPV1 and/or TRPA1 expressed in cutaneous nociceptive nerve endings, thereby reducing their sensitivity to noxious heat, consistent with the presently observed heat analgesia and inhibition of spinal neuronal responses to noxious heat. TRPA1 is also expressed by skin cells including keratinocytes,40 and it is conceivable that propofol may act at TRPA1 or TRPV1 expressed in non-neural cells to indirectly inhibit nociceptive nerve endings in the skin.
There are several additional potential explanations for the local antinociceptive effect of propofol. Propofol inhibits fatty acid amide hydrolase, the enzyme that degrades the endocannabinoid anandamide.41 The antinociceptive effect of hindpaw injection of propofol in the formalin test was attenuated by antagonists of cannabinoid type 1 and 2 receptors,26 suggesting that propofol antinociception involved an increase in local skin levels of anandamide, which in turn inhibited nociceptors via a variety of possible actions (see Ref. 42 for recent review). The antinociceptive effect was considered to be local, because administration of an even larger dose of propofol to the contralateral hindpaw had no effect. In the present study, we did not observe any change in withdrawal latency for the hindpaw contralateral to the one receiving propofol (Fig. 1B), arguing against a systemically mediated antinociceptive effect.
Another possibility is that the antinociceptive effect of peripherally applied propofol is mediated via an interaction with γ-aminobutyric acid A (GABAA) receptors. Subunits of GABAA receptors are localized to unmyelinated fibers in glabrous skin, and intraplantar injection of the GABAA receptor agonist muscimol was antinociceptive in the formalin test.43 Propofol was shown to excite dorsal root ganglion neurons via GABAA receptors; unlike propofol, intradermal injection of GABA did not elicit pain in humans.35 We therefore speculate that propofol may activate peripheral nerve endings expressing GABAA receptors to induce a local antinociceptive, rather than pronociceptive, effect.
Propofol or its halogenated analog was reported to inhibit voltage-sensitive sodium channels44 and to reduce axonal excitability,45 potentially contributing to antinociception. Finally, the present study cannot exclude a possible central inhibitory mechanism triggered by peripheral propofol. Propofol excitation of nociceptor nerve endings via TRPV1 and TRPA1 could provide a barrage of nociceptive input to the central nervous system to engage antinociceptive mechanisms that might provide descending inhibition of heat-sensitive dorsal horn neurons. However, propofol, even at the highest concentration tested, was not presently observed to directly excite any spinal WDR neurons, arguing against this possibility.
In conclusion, our data show that topical application of propofol has analgesic and potentially antihyperalgesic effects via inhibition of nociceptive spinal WDR neurons. Importantly, propofol reduced responses of WDR neurons to noxious heat, but not to cold or innocuous mechanical stimuli, indicating a selective analgesic effect and not a generalized local anesthesia. These results contrast with previous reports of pronociceptive effects of propofol and might be explained by direct or indirect effects that reduce the excitability of nociceptive nerve endings in the skin. Additional preclinical studies are needed to clarify the mechanism by which peripheral propofol induces its antinociceptive effect. The antinociceptive effect of topical propofol is consistent with clinical reports of postoperative analgesia and reduced opiate requirements in surgical patients emerging from anesthesia induced by propofol infusion.
Name: Kenichi Takechi, MD.
Contribution: This author helped design and conduct the study, collected and analyzed data, and wrote the first draft of the manuscript.
Attestation: Kenichi Takechi attests to having approved the final manuscript.
Name: Mirela Iodi Carstens, BA.
Contribution: This author helped design and conduct the study and performed histology.
Attestation: Mirela Iodi Carstens attests to having approved the final manuscript.
Name: Amanda H. Klein, PhD.
Contribution: This author helped design and conduct the study and analyze data.
Attestation: Amanda H. Klein attests to having approved the final manuscript.
Name: E. Carstens, PhD.
Contribution: This author helped design and conduct the study, assisted with data collection and analysis, and edited the manuscript.
Attestation: E. Carstens is the archival author. E. Carstens attests to having approved the final manuscript.
This manuscript was handled by: Martin S. Angst, MD.
The authors thank Minh Trannguyen for excellent technical assistance.
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