In rodents oxytocin produces antinociception in normal animals and reduces hypersensitivity after neuropathic injury (see Ref. 40 for review). In humans, oxytocin produces analgesia in some studies after intracerebroventricular,32 intranasal,39,41,53 and intravenous30 administration. In 2 large prospective trials,11,14 we observed a lower than anticipated incidence of persistent pain after cesarean delivery. Rats recover more rapidly after neuropathic injury when this occurs at the time of delivery, an effect which is reversed by intrathecal injection of antagonists of oxytocin signaling,23 suggesting a role for oxytocin in speeding recovery from pain after injury.45
Oxytocin may act in the periphery to produce analgesia. Systemically administered oxytocin, which does not to cross the blood–brain barrier, produces antinociception and reduces responses to formalin in mice44 and intraplantar oxytocin in rats reduces C- and Aδ-fiber responses in spinal cord and reduces responses to formalin.20 Oxytocin exposure in acutely dissociated primary sensory afferent neurons decreases membrane potential, increases potassium currents, and reduces excitability in normal rats19 and in those after neuropathic injury.27 However, procedures used in primary sensory neuron culture result in loss of most medium and large diameter cells16 which are presumptively A-neurons and can alter neuronal properties, questioning their relevance to in vivo conditions.29 In addition, anatomical studies focus on colocalizing oxytocin-binding receptors with markers of C-neurons including isolectin B419 and calcitonin gene–related peptide.20 Thus, little is known about oxytocin's effects on A-neurons.
The outcome measure most commonly used for experimental neuropathic injury in rodents is mechanical hypersensitivity, yet neuropathic pain in humans is as often accompanied by hyposensitivity as hypersensitivity to mechanical stimuli.18 Using intracellular recording of primary sensory afferent neurons in the anesthetized, intact animal, we recently showed in an injury of the L5 spinal nerve that results in hypersensitivity in the area innervated by the L4 dermatome, that high-threshold mechanoreceptors (HTMRs) are indeed sensitized, with larger receptive fields than controls. The opposite is true for the tactile, low-threshold mechanoreceptors (LTMRs), which are desensitized, with smaller than normal receptive fields.3 As such, the distinction of afferent subtypes activated by mechanical stimuli or light to moderate intensity is lost. There have been no previous studies of the effect of oxytocin on physiologically characterized mechanosensitive afferents.
The primary goal of this study was to describe the effects of acute oxytocin exposure on physiologically characterized LTMRs and HTMRs after peripheral nerve injury compared with a sham surgical control without nerve injury. We hypothesized, based on studies from dissociated neurons, that oxytocin would hyperpolarize neurons and reduce their excitability.
Oxytocin may produce antinociception by actions at either oxytocin and vasopressin 1a receptors or both of these receptors24,42,44,45 or potentially by other mechanisms including alteration in chloride channel function.50 A secondary goal if the current study was to determine the distribution of these receptors in primary sensory afferent neurons and the effect of nerve injury on their expression.
Forty-eight female Sprague–Dawley rats (4-6 weeks of age) were obtained from Harlan Industries, Indianapolis, IN. Animals were housed together in pairs in a climate-controlled room under a 12-hour light/dark cycle and were studied during the light cycle. The use and handling of animals were in accordance with guidelines provided by the National Institutes of Health and the International Association for the Study of Pain, and the procedures and experiments were approved by the Institutional Animal Care and Use Committee of Wake Forest University Health Sciences.
2.1. L5 partial spinal nerve ligation
Rats were deeply anesthetized with isoflurane (TEVA Pharmaceutical USA, North Wales, PA), and under aseptic conditions, the skin was incised at the midline over the lumbar spine. In 24 animals, the right L5 spinal nerve was identified, and approximately 1/3 to 1/2 thickness of the L5 spinal nerve was ligated with 9-0 nylon suture under a dissecting microscope as previously described.22 Care was taken not to pull the nerve or contact the intact L4 spinal nerve. After hemostasis was achieved, the muscle layer was approximated with 4-0 synthetic absorbable suture (Look, Reading, PA), and the skin closed with absorbable suture. In 24 other animals, sham surgery was performed as described above, except that the right L5 spinal nerve was identified but not injured. After the surgery, rats were returned to their cages, kept warm under a heat lamp, and monitored during recovery.
2.2. Behavioral testing
Animals were placed on a mesh surface in a plastic cage and were acclimated for 20 minutes before testing. Withdrawal threshold was assessed by application through the mesh flooring of calibrated von Frey filaments on the footpad until the filaments bent. This was performed by a person blinded to the surgical treatment. The von Frey filaments used were 3.84, 4.08, 4.31, 4.56, 4.74, 4.93, 5.18, 5.46, and 5.88, corresponding to 0.5, 0.9, 1.7, 3.7, 5.5, 8.0, 12.4, 21.5, and 53.0 g, respectively. The filament was applied 3 times with a response considered positive if it resulted in a brisk withdrawal of the paw on any of the 3 applications. The force resulting in a 50% probability of withdrawal (withdrawal threshold) was determined using the up–down method as previously described.13 Withdrawal thresholds were determined before partial spinal nerve ligation (pSNL) or sham surgery and 1 week after surgery. All animals were included in the data analysis, and no animal in the study had a wound dehiscence or infection during the study.
Animals were deeply anesthetized with isoflurane 3%. The trachea was intubated and the lungs ventilated using pressure-controlled ventilation (Inspira PCV; Harvard Apparatus, Holliston, MA) with isoflurane in humidified oxygen. Heart rate by electrocardiography was monitored throughout as a guide to depth of anesthesia. Anesthetized animals were immobilized with pancuronium bromide (2 mg/kg) and inspired, and end tidal isoflurane concentration maintained at 2% throughout the study. A dorsal incision was made in the lumbar midline, and the L4 dorsal root ganglion (DRG) and adjacent spinal cord were exposed by laminectomy as previously described.5 The exposed nervous tissue was continuously superfused with oxygenated artificial cerebrospinal fluid (aCSF [in mM]: 127.0 NaCl, 1.9 KCl, 1.2 KH2PO4, 1.3 MgSO4, 2.4 CaCl2, 26.0 NaHCO3, and 10.0 D-glucose). The spinal column was secured using custom clamps, and the anesthetized animal was transferred to a preheated (32-34°C) recording chamber where the superfusate was slowly raised to 37°C (MPRE8; Cell MicroControls, Norfolk, VA) and its flow rate kept at 1 mL per minute (bath exchange time of ∼5 seconds). Pool temperature adjacent to the DRG was monitored with a thermocouple (IT-23; Physitemp, Clifton, NJ). Rectal temperature (RET-3; Physitemp) was maintained at 36 ± 1°C with radiant heat.
The electrophysiological recording session from each animal was limited to a maximum duration of 75 minutes to diminish the likelihood that surgery and experimental manipulation would result in primary sensory afferent sensitization. Dorsal root ganglion neuronal somata were impaled with quartz micropipettes (80-250 MΩ) containing 1-M potassium acetate. DC output from an Axoclamp 2B amplifier (Axon Instruments/Molecular Devices, Sunnyvale, CA) was digitized and analyzed off-line using Spike2 (CED, Cambridge, United Kingdom). Sampling rate for intracellular recordings was 21 kHz throughout (MicroPower1401; CED).
2.4. Cell inclusion criteria
Because of concern for sensitization from repeated high-threshold stimuli within the same dermatome,8 we limited the number of nociceptive cell recordings per animal. For this reason, only one nociceptive afferent was recorded per animal with the exception of 2 animals in which only 2 nociceptive afferents were recorded.
Inclusion criteria for analysis were the skin innervated by the cell had not previously been stimulated or manipulated in any way that could activate nociceptive afferents; resting membrane potential was more negative than −40 mV; action potential (AP) amplitude was ≥30 mV, after hyperpolarization (AHP) was present; and the neuronal receptor field (RF) was in an appropriate location. Although cellular RFs were found across the entire dermatome in these intact preparations, only those along the medial portion of the dermatome were used in this study (posterior-lateral aspect of the animal leg and paws (Fig. 1A). The study was limited to these areas to avoid effects from the DRG exposure surgery toward the midline and/or the inability to access the RF at this location. Only mechanosensitive cells were included, and their responses and properties were evaluated after threshold and suprathreshold activation. Cellular electrical excitability was documented before, during, and after oxytocin exposure as described below.
2.5. Cellular classification protocol
The procedure used in this study to classify primary sensory afferents in vivo has been described in detail.6 Briefly, the RF was located with the aid of a stereomicroscope using increasing mechanical stimulation progressing from light touch with a fine sable hair paint brush to searching with blunt probes and ultimately with fine-tipped forceps, progressing from gentle to noxious pinch. To further avoid repeated noxious stimulation of the dermatome, the RF area was not mapped. Mechanical threshold was determined using calibrated von Frey filaments. Adaptation rate was evaluated using suprathreshold probes mounted in a micromanipulator. Skin stretch and vibratory stimuli to application of tuning forks of 256 and 512 Hz (SKLAR Instruments, West Chester, PA) were also tested in many cells if the RF was sufficiently accessible. Based on the combination of their mechanical threshold, conduction velocity, and dynamic response (phasic on-off or tonic), cells were classified into 2 groups: LTMR or HTMR (A- and C-fiber HTMRs). LTMRs were subclassified as hairs, mechanical, and cold-sensitive (MC) or rapid adapting by standard criteria.9 Specific cellular subtypes excluded from study were slow adapting tactile afferents (SA-I and SA-II), C-polymodal nociceptors (nociceptors which saturate their responses well below the mechanical nociceptive thresholds in humans9,10,17,21,48 and mechanoinsensitive afferents.
2.6. Somatic electrical properties
Active membrane properties of all excitable neurons were analyzed before, during, and after DRG perfusion with oxytocin in aCSF as described below (Fig. 1B). Active electrical properties examined were amplitude and duration of the AP and of the AHP as well as the maximum rates of spike depolarization and repolarization (MRD and MRR, respectively). Action potential and AHP durations were measured at half amplitude (D50 and AHP50, respectively) to minimize hyperpolarization-related artifacts. Passive electrical properties examined were membrane resting potential (Em), input resistance (Ri), and time constant (Tau: τ), determined by injecting incremental hyperpolarizing current pulses (Ic pulses) (≤0.1 nA, 500 ms) through balanced electrodes (Fig. 1B).
To calculate conduction velocity, spike latency was obtained by stimulating the RF at the skin surface using a bipolar electrode (0.5 Hz, current range: 0.1-1.2 mA) and a stimulus isolator (A360LA, WPI, Sarasota, FL). This was performed after all natural stimulation to prevent potential alterations in RF properties by electrical stimulation (Fig. 1B). All measurements were obtained using the absolute minimum intensity required to excite neurons consistently without jitter. This variability (jitter) in the AP generation latency (particularly at significantly shorter latencies), seen at traditional (2-3-fold threshold) intensity has been presumed to reflect spread to more proximal sites along axons. Stimuli ranged in duration from 50 to 100 µs. Utilization time was not taken into account. Conduction distances were measured for each afferent on termination of the experiment by inserting a pin through the RF (marked with ink at the time of recording) and carefully measuring the distance to the DRG along the closest nerve.
2.7. Somatic exposure to oxytocin
After characterization with the initial aCSF perfusion (aCSFpre), the cells were exposed to aCSF containing oxytocin, 1 nM, and their cellular properties (RF and somatic) retested 5 minutes at the end of this exposure. The perfusion was returned to aCSF postoxytocin exposure (aCSFpost) and cellular properties tested ∼10 minutes later (Fig. 1B). The constituents of the aCSF were identical in all solutions except for oxytocin presence. In all cases, temperature, aCSF flow rate (±1 mL per minute), and oxygenation were kept constant. A maximum of 2 cells were recorded per experiment with a resting period of ∼20 minutes of constant perfusion with normal aCSF between recordings (Fig. 1B).
2.8. Tissue preparation for immunocytochemistry
Before use in tissues from animals in this experiment, immunostaining experiment conditions were determined from recently prepared tissue in other animals. The antivasopressin 1A receptor antibody (rabbit, 1:100, #AVP1A11-P; Alpha Diagnostics, San Antonio, TX) showed selective immunostaining in brain regions of vasopressin 1A receptor location in the rat brain and consistent immunostaining in a subset of DRG neurons and was used for tissues from animals in the experiment. Details of use of this antibody in experimental tissue are presented after the following paragraph regarding oxytocin receptor immunostaining.
We screened 3 commercially antibodies for the oxytocin receptor in DRG sections. The first 2, obtained from Santa Cruz Biotechnology, Dallas, TX, are goat antioxytocin receptor antibodies (catalog numbers sc-8102 and sc-8103) and were tested in dilutions of 1:50, 1:100, 1:200, 1:500 on freshly prepared DRG 20-μ sections from adult rats. Sections were preblocked with 3% normal donkey serum containing 0.3% Triton X-100 in 0.01-M phosphate-buffered saline [PBS]), for 1 hour at room temperature, then incubated with one of the primary antibodies overnight at 4°C and finally with a donkey anti-goat Cy3-conjugated IgG (1:200) secondary antibody (Jackson ImmunoResearch Labs, West Grove, PA) in 0.01-M PBS for 2 hours at room temperature. In other experiments, we used another secondary antibody (rabbit anti-goat [Jackson ImmunoResearch], 1:200 in PBS. The third antibody, obtained from Alomone Labs, Jerusalem, IL, is a rabbit antioxytocin receptor antibody [catalog number AVR-013] and was at 1:200, 1:500, and 1:1000 on freshly prepared DRG 16-μ sections from adult rats. Sections were preblocked with 3% normal donkey serum containing 0.3% Triton X-100 in 0.01 M PBS) for 1 hour at room temperature then incubated overnight with the primary antibody overnight at 4°C and finally with a donkey anti-rabbit Cy3-conjugated IgG (1:600) secondary antibody (Jackson ImmunoResearch Labs) in 0.01-M PBS for 2 hours at room temperature. We also examined another secondary antibody (donkey anti-rabbit biotin 1:500/streptavidin-Cy3 1:4000; Jackson ImmunoResearch Labs). In all cases, we saw diffuse immunostaining in nearly all DRG neurons (representative images in supplemental data 1, available at http://links.lww.com/PAIN/A736). We previously had screened other commercially available antibodies for the oxytocin receptor in brain (LifeSpan Biosciences, Inc, Seattle, WA and Alpha Diagonistics International, Inc, San Antonio, TX), but the location of immunostaining did not correspond to radioligand binding of oxytocin receptors using a highly selective radiolabeled peptide,49 so their use was not pursued in DRG sections.
To examine vasopressin 1a receptors, following the electrophysiological experiments in 3 animals with pSNL and 3 animals with sham surgery, the thorax was opened and fixative (4% paraformaldehyde in 0.1-M phosphate buffer, pH 7.4) was perfused through the left ventricle with a peristaltic pump at 20 mL/minute for 15 minutes. L4 and L5 DRGs ipsilateral to surgery were then identified, removed, and immersed in fixative (postfixation) for 2 hours at 4°C. Afterwards, ganglia were immersed in 30% sucrose at 4°C for cryoprotection until sectioned on a cryostat. Sections (20 μm) were collected on slides and stored at −80°C until processed. Sections from 3 animals per group (sham and pSNL) were processed simultaneously, and antibodies for vasopressin 1A receptor and neuronal nuclei were used to examine the expression of this receptor in DRG neurons. The sections were washed, blocked, and incubated overnight at 4°C with primary antibodies against the vasopressin 1A receptor (rabbit, 1:100, #AVP1A11-P; Alpha Diagnostics) and against NeuN (mouse, 1:100; # Millipore MAB 377, Temecula, CA) followed by the corresponding donkey anti-rabbit Alexa fluor 488 (1:200) and anti-mouse Cyanine Cy3 (1:400) secondary antibodies (Jackson ImmunoResearch Labs) in 0.01-M PBS. Vasopressin 1A receptor antibody specificity was tested using a blocking peptide (#AVP1A11-P), following the manufacturer instructions. Finally, the sections were washed thoroughly in PBS, mounted on plus-slides, air-dried, dehydrated in ethanol, cleared in xylene, and cover slipped with DPX-mounting media.
Sections were examined on a Nikon E600 epifluorescence microscope, and images were captured with a CCD digital camera attached to the microscope using a 20X objective. Images of the ipsilateral L4/5 DRGs were captured. Quantification for each image was performed, and 3 randomly selected sections were examined per rat. The images were quantified using Image J (U. S. National Institutes of Health, Bethesda, Maryland, http://imagej.nih.gov/ij/, 997-2011). Upper and lower thresholds were adjusted to contain and match the immunoreactivity, generating an image with immunoreactive elements appearing as red pixels. These same thresholds were applied to all sections.
To conduct the analysis of the DRGs, the 15 to 20 neurons with clear nuclear staining were selected, processing in a rough row from left to right on the slide, then to the next row inferior to the first. Cell diameter was calculated using the average of the major and minor orthogonal axes. The border of each included cell was manually traced, and, using a macro, the number of pixels of immunoreactivity above threshold was determined in each cell. The experimenter performing image analysis was blinded to group.
2.9. Statistical analysis
Electrophysiologic properties of all LTMR subtypes and of the A- and C-HTMR subtypes were combined for analysis. Before analysis, parametric assumptions were evaluated for all variables using histograms, descriptive statistics, and the Shapiro–Wilk test for normality. Data that were normally distributed are reported as mean ± SE and those not normally distributed as median (first and 3ed quartiles). Student t test and repeated-measures analysis of variance were used for normally distributed data, and Friedman test and Mann–Whitney U test were used for not normally distributed data. Contrasts of interest in electrophysiologic properties were between sham an pSNL groups as a function of type of afferent and, within afferents, between oxytocin-containing aCSF perfusion and aCSFpre and aCSFpost. Comparison of distribution of cell diameter in populations positive and negative for vasopressin 1A receptor immunoreactivity were analyzed by χ2. Analyses were performed using OriginPro 9.5 (Northampton, MA).
2.9.1. Sample size determination
The study was powered with a minimum number of 12 cells within HTMR and LTMR classes to observe a difference in Em of 6 mV with α = 0.05 and 1 − β = 0.80.
One week after surgery, withdrawal threshold ipsilateral to pSNL (median: 4.8 g [4.2-11.2]) was significantly lower than that in sham surgery animals (median: 19.2 g [11.7-58.2]; P < 0.01). By contrast, withdrawal threshold contralateral to surgery did not differ between groups and did not differ from presurgery baseline (data not shown).
3.2. Electrophysiology: effects of partial spinal nerve ligation
Intracellular recordings were obtained in 63 well-characterized mechanosensitive neurons innervating the L4 dermatome from 48 animals (Fig. 1C). Recordings of 28 cells were obtained from sham animals and were classified as LTMRs (16 total: 8 hairs and 8 LTMR-RAs) and HTMRs (12 total, all A-HTMRs). The remaining 35 cells were obtained from pSNL animals and were classified as LTMRs (21 total: 7 hairs, 2 LTMR-MCs, and 12 LTMR-RAs) and HTMRs (14 total: 12 A-HTMRs and 2 C-HTMRs; Fig. 1A). Receptive fields were located on the lateral aspect of the hindlimb and the plantar aspect of the paw (Fig. 1C).
Low-threshold mechanoreceptor afferents recorded from pSNL animals were significantly desensitized compared with those recorded from sham animals as evidenced by a higher mechanical threshold in pSNL vs sham (MT 0.295 mN [0.07-39] vs 0.078 mN [0.07-3.9], respectively. P < 0.01). By contrast, nociceptive afferents recorded from pSNL animals were significantly sensitized compared with those from sham animals (MT 26.5 mN [1.6-588] vs 147 mN [98-980] in pSNL vs sham, respectively. P < 0.01).
3.3. Passive electrical cellular properties
For LTMRs, pSNL resulted in more negative Em, with no changes in Ri or tau, whereas pSNL resulted in no change for HTMRs in Em or Ri, but an increase in tau (Table 1 [values in bold], Fig. 2A). Oxytocin had opposite effects on Em in the 2 classes of afferents in pSNL animals, increasing Em in LTMRs and decreasing it in HTMRs, but had no effect on Em in either class of afferents in sham controls (Table 1 [values in italic], Fig. 2A). Oxytocin increased tau in LTMR and HTMR afferents in both pSNL and sham animals but had no effect on Ri (Table 1 [values in italic]). These effects on Em and tau were fully reversed with perfusion of nonoxytocin-containing aCSF. Two significant differences did occur in sham animals at the end of this last perfusion with aCSF—a reduction in Em in LTMRs and a reduction in tau (Table 1 [values in italic]).
In addition to these effects on passive electrical properties, exposure to oxytocin resulted in spontaneously propagated spikes followed by development of spontaneous APs in 6 of 21 LTMRs in pSNL animals (Fig. 2B). By contrast, oxytocin exposure to HTMRs in pSNL animals resulted only in membrane hyperpolarization and electrical silence. These effects in spontaneous activity resolved after perfusion with nonoxytocin-containing aCSF, and none of these effects were observed with oxytocin exposure in sham animals.
3.4. Active electrical cellular properties
Two components of the evoked AP were evaluated in sham and pSNL animals before and after oxytocin: AP amplitude and duration (D50) and AHP amplitude and duration (AHP50). As expected, LTMRs differed from HTMRs in the control condition, with smaller amplitude of both AP and AHP but with similar duration of these components of the AP (Fig. 2C and Tables 2 and 3). Partial spinal nerve ligation did not affect active electrical properties of LTMRs or HTMRs (Fig. 2C and Tables 2 and 3).
Oxytocin had no effect on active electrical properties with the single exception of an increase in after-hyperpolarization duration in HTMRs in the sham group only (Fig. 2C and Tables 2 and 3 [value in italic]). The contrast between LTMRs and HTMRs in sham and pSNL groups described above was not affected by oxytocin exposure.
3.5. V1a receptor immunocytochemistry
A total of 337 NeuN-positive neurons from L4 DRGs in 6 animals were analyzed for V1a receptor immunoreactivity. Immunoreactivity was restricted to neurons and, as has been previously described, presented as punctate labeling without obvious outer membrane localization (Figs. 3A and B). Vasopressin 1A receptor immunopositive cells were larger than immunonegative cells in diameter in the entire study population (Fig. 3C left panel; mean 34 ± 6.3 vs 26 ± 3.4 μ, respectively, P < 10−6). In the 2-way analysis of variance on cell diameter, there was no main effect of surgical group (P = 0.2), a highly significant effect of immunoreactivity (P < 10−6), and a marginally significant interaction (P = 0.02; Fig. 3C right panel). The proportion of cells meeting criteria for vasopressin 1A receptor immunopositivity using our sampling approach was decreased in the pSNL group (65/164 cells [40%]) compared with the sham group (99/173 cells [57%]; P = 0.0036 by χ2).
Key findings of the current study are (1) replication of sensitization of HTMRs and desensitization of LTMRs following nerve injury and (2) novel observations that sensitized HTMR neurons are hyperpolarized whereas LTMR neurons are desensitized by oxytocin. (3) In addition, vasopressin 1a receptors are present in large as well as small- and medium-sized DRG neurons. These data fill gaps in our knowledge regarding the mechanism of peripherally mediated analgesia from oxytocin.
4.1. Effects of injury and oxytocin on high-threshold mechanical nociceptors
Several lines of evidence point to the relevance of HTMR dysfunction in pain states. A-HTMRs project more extensively in the spinal cord than do C-fibers,10 which may be responsible for large areas of mechanical hypersensitivity in some patients with neuropathic pain18. A-HTMRs show decreased mechanical threshold and increased receptive field area after acute (incision) or chronic (nerve ligation) injury, with quantitatively smaller effects on C-HTMRs.3,4 After nerve injury, rats avoid a floor designed to stimulate sensitized HTMRs, whereas normal rats do not.7 A-HTMR response to sustained receptive field stimulation in normal rats is rapidly adapting but is shifted to a sustained response after intermittent, repeated high-intensity stimuli. After nerve injury, only sustained responses are observed.8 Finally, selective, transcutaneous optogenetic silencing of A-HTMRs increases withdrawal threshold to mechanical stimulation after nerve injury.6
Oxytocin-induced A-HTMR hyperpolarization in vivo after nerve injury in the current study is similar to oxytocin's effects in vitro in trigeminal nucleus neurons from animals with nerve injury.27 Oxytocin induces hyperpolarization in vitro in DRG neurons in normal rats,19 whereas we saw no such effect in vivo in A-HTMRs in animals without nerve injury, perhaps due to effects of axotomy in vitro. Hyperpolarization could reduce excitability at peripheral terminals and reduce neurotransmitter release at central terminals.
4.2. Effects of injury and oxytocin on low-threshold tactiles
There is also support for LTMR dysfunction in neuropathic pain. Areas of tactile hyposensitivity are common in patients and cannot solely be explained by denervation.18 Gate control theory35 posits a normal brake on nociceptive input by tactile input, a brake which would be reduced if LTMRs are poorly responsive or unresponsive. Strategies which presumably drive LTMR input such as transcutaneous electrical nerve stimulation are commonly applied in chronic pain syndromes.51
We confirmed in the current study that mechanical threshold for LTMRs increased after nerve injury compared with controls3; although in the current study, this was accompanied by a reduced Em, whereas in the previous study, it was not. We speculate that repeated testing with strong mechanical stimuli within the dermatome in the previous study may have acutely depolarized Em in LTMR units.
Oxytocin depolarized LTMRs in pSNL animals but not in sham animals. The mechanisms by which oxytocin produces divergent effects on LTMRs and HTMRs after injury but has no effect in controls are unclear (see brief speculation in the next section), but oxytocin's net effect was to partially restore the nerve injury–induced divergent perturbations in HTMR and LTMR passive electrical properties towards normal. The relevance of the spontaneous LTMR activity is unclear, given that neither intranasal31 nor intrathecal15 oxytocin induces paresthesias or phantom sensations in humans.
4.3. Speculation on mechanisms for oxytocin's effects
Oxytocin-induced depolarization of LTMRs in pSNL animals is consistent with excitatory G protein signaling through oxytocin or vasopressin 1a receptors through inositol triphosphate and rise in intracellular calcium in uterine myometrium1 and in some observations in spinal cord12 and DRG2 neurons. By contrast, oxytocin inhibits APs from current pulses in transient receptor potential for vanilloid-1 (TRPV-1)–positive DRG neurons in vitro, associated with increased potassium conductance,24 consistent with oxytocin-induced activation of KATP channels leading to membrane hyperpolarization.19,54 Taken together, these studies suggest that, depending on neuronal expression of ATP- and calcium-dependent potassium channels by afferent class and injury status,25,43,55 oxytocin-mediated increases in intracellular calcium can depolarize and excite or hyperpolarize and inhibit neuronal activity and activation.
4.4. What was not studied
Based on both clinical and laboratory observations, we focused on fast-conducting mechanosensitive neurons rather than C-HTMRs and C-polymodals that are less affected by nerve injury than myelinated afferents and have an unclear role in mediating altered mechanosensation after nerve injury.17,48 Certainly, oxytocin affects unmyelinated sensory afferents, including desensitization of TRPV-1 ion channels through actions on oxytocin37 or vasopressin 1a24 receptors and reduced heat-evoked pain and nociception in some studies in animals44 and humans.39 We do not infer that injury-induced plasticity of C-afferents or oxytocin's actions on them is irrelevant.
Because of technical limitations in the duration of stable cell recording, we did not test the effects of oxytocin or vasopressin 1a receptor antagonists on oxytocin's actions, so the receptor(s) upon which oxytocin acts is not addressed in these studies. Future studies using genetic knockouts of these receptors in mice28,44 could address this issue. Finally, we were unable to assess oxytocin receptors anatomically using immunostaining, although they are known to be present in rat24 and human46 dorsal root ganglia. Specifically, we were unable to replicate selective immunostaining with a commercially available antibody36 that could reflect difference in batches or in conditions not discussed in that report.
4.5. Vasopressin 1a receptors are present in small, medium, and large diameter sensory afferent neurons
The receptors upon which oxytocin acts to produce analgesia are unclear, it stimulates both oxytocin and vasopressin 1a receptors. Messenger RNA for both receptors is expressed in DRG neurons, and divergent results have been obtained in different studies. For example, studies in knockout mice suggest oxytocin acts primarily through vasopressin 1a receptors in the periphery and centrally,44 whereas pharmacologic studies suggest intraplantar oxytocin acts at least through oxytocin receptors20 while intraplantar vasopressin acts through oxytocin and vasopressin 1a receptors.34 We recently showed reversal of recovery from nerve injury–induced hypersensitivity in rats by intrathecal injection of either vasopressin 1a or oxytocin receptor selective antagonists, without a sex difference.45
We observed punctate immunoreactivity for vasopressin 1a receptors in large and small diameter DRG neurons in this study, whereas in the trigeminal nucleus, such immunoreactivity is limited to small–medium cells.27 Fully 22% of immunopositive neurons in the current study were >40 μm in diameter, compared with only 5% in that study of trigeminal nucleus neurons.27 In mouse DRG neurons, 97% of cells expressing vasopressin 1a messenger RNA were small or medium size (<30 μm), but it is unclear whether large cells were lost in the dissociation method.44 A recent study identified immunoreactivity for both oxytocin and vasopressin 1a in thin fibers in skin,34 but DRG tissue was not examined. As such, this is the first report of vasopressin 1a in rat DRG to demonstrate immunostaining in large as well as small- and medium-sized neurons.
4.6. Clinical translation
Whether systemically administered, oxytocin produces analgesia in humans is unclear with positive30,53 and negative33,38 results in clinical pain states as well as positive39,41 and negative results in normal volunteers.26,47,56,57 Nearly all these studies used intranasal administration and small sample sizes and have been criticized for extremely low positive predictive value.52 The current study predicts that systemic or locally administered oxytocin is most likely to affect sensation in the presence of injury, and that it would increase perception of stimuli transduced by LTMRs (eg, vibration) and decrease intensity and/or duration of stimuli transduced by HTMRs (eg, pinch or punctate stimulation).
Interpretation of this study is limited by the study conditions: single sex (female), study within a short period (1 week) after nerve injury, focus on only 2 classes of sensory afferents, nonrandomized presentation of oxytocin perfusion, administration of oxytocin surrounding the neuronal soma rather than into its receptive field, lack of assessment of oxytocin on receptive field area or of mechanical threshold and systematic, but not stereologic selection of cells in immunohistochemical studies. Many of these electrophysiologic limitations reflect the extraordinary technical difficulties to stably record from single neurons in vivo in the live animal. In addition, future studies could more completely characterize these neurons by intracellular injection of dye after physiologic study.
Intracellular recording in vivo in rats shows that peripheral nerve injury desensitizes tactile afferents and sensitizes mechanical nociceptors. Perfusion of the DRG with oxytocin acutely hyperpolarizes these sensitized nociceptors while simultaneously depolarizing these desensitized tactile afferents. Vasopressin 1a receptor immunoreactivity is present in small to medium and large afferent neurons consistent with their possible site of oxytocin action. These data are consistent with existing behavioral studies in rodents suggesting an antihypersensitivity effect of peripheral oxytocin after injury and predict specific psychophysical effects of peripherally administered oxytocin in normal humans and those with injury-induced pain.
Conflict of interest statement
J.C. Eisenach consults to Adynxx (San Francisco, CA) and TEVA Pharmaceutical Industries, Ltd (Petah Tikva, Israel) regarding preclinical and clinical analgesic development of analgesics. Neither company uses the methods described in this manuscript and did not participate in this work or the manuscript. The remaining authors have no conflicts of interest to declare.
Supported in part by grant R37 GM48085 to J.C. Eisenach from the National Institutes of Health, Bethesda, MD.
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Oxytocin; Neurophysiology; Neuropathic pain; Vasopressin Receptor; Sensory Afferent
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