Chronicpain affects 11% of the American population and places an economic burden of $560 billion on the United States.1Specifically, neuropathic pain results from conditions such as nerve injury, metabolic dysregulation, and drug toxicity.2,3Various rodent models have been developed to study treatments for chronic pain,4,5but translation to human application during phase I and II clinical trials has been largely unsuccessful. In fact, only 0.3% of phase I clinical trials result in new treatments for chronic pain.6
Peripheral nerve injuries cause neuronal hypersensitivity and activate dorsal root ganglion (DRG) neurons, resulting in neuropathic pain.7,8Previous work has demonstrated that in vivo electrical DRG stimulation, and the resulting inhibition of DRG neurons, can alleviate allodynic behaviors in a rodent model.9Surgically implanted electrical DRG stimulators are currently the most effective form of treatment for medically refractory dermatome-specific neuropathic pain in the clinic.10–12We have demonstrated that low intensity focused ultrasound (liFUS) treatment at the DRG has similar efficacy for neuropathic pain in a rodent model of common peroneal nerve injury (CPNI).13,14
A positive correlation has been identified between swine models and the subsequent translation of treatments to successful clinical trials.15However, the use of swine models in the pain literature has been limited. Domestic swine have been employed to develop models of inflammation, muscular and bone pain,16–18and mixed neuritis.19The mixed neuritis model used both complete Freund’s adjuvant (an immune-related neuropathic model) and ligation, thus creating a mixed etiology for chronic pain. Here, we describe a CPNI model of neuropathic pain in domestic swine. This is a modification of the well-validated sciatic nerve injury model used in rodents and involves ligation of a branch of the sciatic nerve, specifically, the common peroneal nerve (CPN).13,14,20,21
In order to help move neuropathic pain treatments toward clinical implementation, we have translated our CPNI model from rodents13,14国内猪。国内猪CPNI疼痛莫del allows positioning of the incision and nerve ligation site near the patella, thus minimizing damage during surgery at targeted upstream treatment sites such as the nerve roots and the DRG. This model is advantageous because, compared with other animal models, domestic swine are readily available, are more economical, and allow for better modeling of human anatomy.22The purpose of the current study was to validate a new model of neuropathic pain in domestic swine. We demonstrated that CPNI ligation induced significant thermal and mechanical allodynia and significant motor deficits. Further, we showed that CPNI ligation decreased in the myelin protein level upstream of the ligation site.
开云体育世界杯赔率
Animals
This study was approved by our Institutional Animal Care and Use Committee. Twelve male domestic swine, aged 7–8 weeks, were used (13–18 kg; Parsons Farm). Animals were housed in open pens (188 × 88 cm), with lights on from 7amto 7pm. Feeding occurred twice a day (miniswine diet, Envigo Teklad; 3% body weight/feeding) with free access to water. An enrichment object was placed in the pen (plastic ball or pin).
Animals remained in pens for 24 hours after arrival to acclimate. The following day, animals were released from pens to socialize and exercise for 20–30 minutes. Researchers spent 20 minutes daily sitting with the animals to habituate them to human interaction. Continuity of the research team, time of day, and testing techniques remained consistent throughout the study to minimize animal stress levels during the study. Acclimation to the sling apparatus began on day 2 after arrival. Animals were lifted into a Lomir Biomedical sling 2 times a day for 5 days. Animals were placed in slings and baseline mechanical, thermal, and motor/social responses were assessed prior to surgery. Animals were on site for 1 week before surgical interventions began. A schema of the experimental timeline is shown inFig. 1.
Schema of experimental timeline.
CPNI Surgery
CPNI surgery was performed using a modification of a method previously validated in rats.13,14Anesthesia was induced with a mixture of xylazine (2.2 mg/kg), ketamine (2.2 mg/kg), and tiletamine and zolazepam (4.4 mg/kg), administered subcutaneously (s.c.), as well as 5% isoflurane. Carprofen (2.2 mg/kg) and cefazolin (4.4 mg/kg) were given (s.c.) for pain and infection control. Following induction, isoflurane was reduced to 1.6%–2% for the duration of the surgery. Animals were intubated and placed in the prone position, and breathing was maintained with a ventilator. Aseptic technique was used throughout the procedure. An oral thermometer was utilized to monitor body temperature, and a heating blanket was adjusted to keep the animal normothermic at 38.6°C ± 0.5°C.23Lactated Ringer’s solution was given intravenously throughout the surgery (7 ml/kg/hr; Baxter Healthcare Corporation). The left hind leg was shaved and marked 2 cm posterior to the fibular head and 1 cm above and below. Bupivacaine (2 mg/kg, s.c.) was injected at the incision site for perioperative pain control. Sterile draping was placed to allow access to the left hip, knee, and ankle, and the area was covered with Ioban (3M). A skin incision was made at the previously marked site, and the muscle was transected using blunt dissection to reveal the CPN. Two 2.0 silk sutures were tightly placed 1–2 cm apart on the CPN to create the injury. Both layers of muscle overlying the nerve were reapproximated using interrupted 4.0 Vicryl sutures. The subcutaneous tissue was then closed with 4.0 Vicryl sutures. Skin closure evolved from staples to nylon suture over the course of the study. Temperature, pulse, and respiration were monitored throughout until animals were standing, eating, and drinking. Oral carprofen (Rimadyl, Zoetis) was given once a day for 3 days postoperatively, and animals were allowed to recover for 7 days prior to postsurgical behavioral testing.
Sensory Threshold and Behavior Testing
Von Frey filaments (VFFs) and thermal tests were utilized to determine mechanical and thermal allodynia, respectively. Both procedures required adjustment as we transitioned from rats to swine.13Initially, we placed animals in slings and performed VFF testing using the skin on the ventral surface (back side) of the hind hoof directly above the hoof padding (n = 2). However, the hoof padding prevented accurate assessment. For the remaining animals (n = 10), the skin on the dorsal surface (front side) of the hind foot, just above the hoof, was tested and these data were included in analyses.
The VFF test measures mechanical sensitivity using a series of calibrated nylon filaments ranging from 1 to 284.28 g (Stoelting). Responses to fibers of 4.0 g or lower indicated enhanced mechanical sensitivity and were deemed allodynic.13,24,25An upper cutoff of 284.28 g was used to avoid tissue damage or actively lifting the foot. During testing, animals were poked 3 times with a filament for 3–5 seconds, progressing from the smallest to largest fiber. A response was defined by withdrawal, guarding, or kicking of the hind hoof. Filaments were applied in increasing order of force until the animal responded. After a response was recorded, 6 subsequent fibers were applied and responses were recorded, according to the Chaplan up-down method.26If the swine reacted to a fiber, the next lower weight (and smallest diameter) fiber was used. If no response occurred, the next largest fiber was employed. If the swine did not respond to the 284.28-g fiber, the test was terminated.
Thermal sensitivity thresholds were measured using the Medoc Pathway (Advanced Medical Systems). Initially, a test paradigm used in human subjects was implemented (maximum temperature of 45°C), for which a series of responses to cool, warm, hot, and cold were recorded. However, this required 40 minutes per test in the sling. Based on excessive vocalization and attempts to escape, animals were determined to be unable to tolerate restraint in the sling for the amount of time required. The protocol was modified for swine, and each site was tested once (maximum temperature of 45°C). The probe was placed in the same area of the hoof used for VFF testing and the temperature was increased from 25°C to 45°C at a rate of 0.5°C per second. The test was terminated once the animal responded, and the temperature was recorded. A response was defined with the same withdrawal, guarding, or kicking as in the VFF test. If the animal had no reaction to 45°C, the test was considered completed. In an attempt to raise the ceiling effect of this test at baseline, we iteratively increased the maximum temperature to 50°C. Data with this technique are available from 4 animals.
Social behavior and motor scores were monitored for 10-minute periods before sling testing. Motor scores were based on three categories: weight-bearing, animal appearance, and vocalization.19If weight-bearing was equal on both legs, animals received a score of 0, and if animals were carrying weight mainly on their intact leg, animals received a score of 1. Similarly, appearance was scored from 0 to 1 based on guarding behavior in the injured leg, and from 0 to 2 based on the level of vocalization. Higher scores indicate greater pain behavior. Social behavior was scored based on interactions with other animals in the cohort during the same 10-minute window. Restlessness was based on pacing and jumping, agitation was scored via interaction with other animals upon being approached, aggression was based on attacking and biting, and isolation was determined by the amount of interaction swine had with one another. Scores were 0–2 for all behaviors except isolation (0–1).19Again, higher scores indicate higher levels of distress. Social and motor scores were determined per animal. Therefore, no comparison was made between the surgery and contralateral hooves.
Immunohistochemical Analysis
Immunohistochemical analysis was performed to assess the presence of myelin protein zero (P0), a structural component of the myelin sheath in the peripheral nervous system in both the injured CPN and the uninjured contralateral CPN. After behavioral testing was complete, animals were euthanized. Transcardiac perfusion was performed using heparinized saline (4 L) followed by 4% paraformaldehyde (PFA; 5 L), and both CPNs were extracted and placed in PFA. Nerves were stored at 4°C in PFA for 24 hours, dehydrated with 5% sucrose, and then stored overnight in 20% sucrose. Tissue was next incubated in 20% sucrose/optimal cutting temperature compound solution (OCT; Fisher Scientific; 2:1 ratio), and finally, liquid nitrogen vapor was used to freeze the nerve in OCT. The samples were sectioned at 10 μm and mounted on slides. Cross sections were permeabilized with 100% cold methanol. Sections were rinsed in 1× phosphate-buffered saline (PBS) and blocked for 1 hour in 30% fetal bovine serum, 1% bovine serum albumin, and 0.1% Triton X-100 in 1× PBS. The primary antibody, anti-P0 antibody 1/1000 (P0; Aves Labs), was diluted in 1× PBS. Sections were rinsed in 1× PBS, incubated for 1 hour with goat anti–chicken Alexa Fluor 555 secondary antibody (Thermo Fisher) 1/200, and stained with DAPI and mounted with Vectashield (Vector Laboratories). Images were acquired at ×20 and ×40 with a Zeiss ApoTome microscope (Axio Imager A2). Images were quantified for intensity and location of P0 signal in both the injured and the contralateral CPN. Changes in P0 expression can be used to assess disruptions or changes in myelin structure and function in the injured nerve as well as the degree of damage at the ligation site.
Statistical Analysis
Statistical analyses and graphs were created using GraphPad Prism version 6 (GraphPad Software). Mean ± SEM was used to represent all data. One-way ANOVA was used for behavioral and electrophysiological data, and a Tukey post hoc test was applied where appropriate; p < 0.05 determined significance.
Results
Sensory Thresholds and Behavior in CPNI Model
All animals received baseline VFF testing 24 hours prior to CPNI surgery. The scores for baseline VFF were 281.84 g in both the right and left hind hooves; none of the animals displayed allodynia or responded to the highest weighted fiber in either the left or right hind hoof (n = 10). After the CPNI surgery, all animals displayed allodynic mechanical responses to fibers of less than 4 g on the left side. Post CPNI VFF scores were significantly lower than presurgical baseline responses (3.28 ± 1.19 g, p < 0.001, paired t-test;Fig. 2）.A similar increase in mechanical sensitivity was seen after CPNI induction in our previous rat model.14The right hind hoof was used as an internal control, and VFF scores remained unchanged at 281.84 g in the right hind hoof following CPNI surgery.
Mechanical thresholds of the dorsal aspect of the left hind hoof. VFFs were used to measure mechanical thresholds (force, in grams) at baseline and after CPNI. A paired t-test revealed a significant decrease in mechanical thresholds following CPNI surgery (***p < 0.001; n = 10). Mean ± SEM values were 281.80 ± 0.00 g at baseline and 3.70 ± 1.47 g after CPNI surgery.
Animals showed no response to thermal stimuli (up to the 50°C cutoff) prior to CPNI surgery. Post CPNI surgery, the left hind foot responded at 33.17°C ± 2.00°C, suggesting increased thermal response and allodynia (p < 0.01;Fig. 3）.正确的后蹄又作为实习生al control following CPNI surgery and showed no change from the 50°C baseline response.
Thermal thresholds of the dorsal aspect of the left hind hoof. Thermal thresholds were determined with the Medoc Pathway quantitative sensory testing stimulator. CPNI significantly decreased thermal thresholds compared to baseline values (**p < 0.01, paired t-test; n = 4). Mean ± SEM values were 49.67°C ± 0.33°C at baseline and 33.17°C ± 2.31°C post-CPNI surgery.
All animals exhibited normal motor scores prior to CPNI (score of 0; n = 12), meaning that animals were not displaying signs of uneven weight-bearing, odd appearance, or abnormal vocalization prior to CPNI surgery. Ten of the swine had a score of 0 for social scores, indicating no increased signs of restlessness, agitation, aggression, or isolation from the other animals in their cohort. However, at the same time point, two animals had a social score of 1, based on signs of agitation toward the other animals. This was determined by the animals’ tendency to move away from other pigs and handlers upon approach. One week after CPNI, motor scores increased from 0 to 1 in 11 animals due to leg guarding (a submeasure of weight-bearing), and from 0 to 2 in 1 animal due to leg guarding and difficulty with weight-bearing (p < 0.01, paired t-test;Fig. 4A）.After CPNI, 11 of 12 animals demonstrated normal social behavior, with one animal displaying signs of agitation (p > 0.05;Fig. 4B）.Compared to baseline, 1 less animal was showing signs of agitation toward the other animals. This result was most likely due to habituation. Further, none of the animals were displaying signs of aggression, isolation, or restlessness after CPNI surgery. These data indicate that CPNI surgery results in leg-guarding behavior, but did not affect social interactions.
Social and motor scores of swine.A:Motor scores were determined on a scale of 0–4, with 4 representing the most severe deficit. After CPNI surgery, motor scores significantly increased (***p < 0.001, paired t-test; n = 12). Mean ± SEM values were 0.00 ± 0.00 at baseline and 1.08 ± 0.08 after CPNI surgery.B:Social scores were based on a scale of 0–7, with 7 representing the most severe deficit. Social scores did not significantly change between the baseline and post-CPNI time points (p > 0.05; n = 12).
Immunohistochemistry and Morphology
In a subset of CPN samples (n = 4) taken upstream of the ligation site itself, tissue was stained and imaged for the presence of P0. This protein is essential for the formation and structure of the myelin sheath,27and nerve injury results in decreases of P0. We viewed the uninjured control right CPN and ligated left CPN samples, taken 2 weeks after the ligation. The right uninjured CPN showed robust P0 expression (Fig. 5A）.In contrast, nerves that underwent ligation displayed only small traces of positive P0 immunofluorescence (Fig. 5B上游的损伤部位。丧失P0 this tissue indicates that P0, essential for proper myelin formation, is significantly decreased as a direct result of the nerve injury in this model. This finding corroborates previous studies that show that significant decreases in P0 have been associated with the development of neuropathic pain.28
Immunostaining of the CPN using anti-P0 antibody. Immunostaining was performed 2 weeks after CPNI.A:Representative immunostaining of the contralateral (right) CPN. Myelin staining for P0 can be observed inred. DAPI counterstaining (blue) shows nuclei of Schwann cells.B:Representative immunostaining of the injured (left) CPN. P0 immunostaining is absent. Images were acquired at the ×20 objective.
Discussion
Our results demonstrated that CPNI surgery modified both mechanical and thermal thresholds in domestic swine. VFF scores were significantly reduced from 281.84 to 4.00 g, or lower, after CPN ligation. Thermal thresholds also significantly decreased as a result of the ligation surgery. Behavioral limb guarding developed following the CPNI surgery and was concurrent with the mechanical and thermal allodynia. These behavioral responses seen in our neuropathic swine model closely mimic those seen in human neuropathic pain.29
Neuropathic pain models in rodents, especially rats, are well defined and allow for the use of sensory thresholds as a surrogate measurement of pain.29Other behavioral tests also afford supporting data.29These models are ideal for mechanistic studies. Peripheral nerve injury models include axotomies, chronic constriction injuries, partial and complete sciatic nerve ligations, and CPN ligations, to name a few.30The predictive validity of small-animal studies for the translation of novel pain treatments to the clinical setting has been low.31Thus, once proof of principle has been documented to test a new treatment option, a large-animal model offers certain advantages. Specifically, in our laboratory we focus on devices to modulate pain. Large-animal models have greater potential genetic and anatomical similarities to humans31but have been used less frequently.32–34
The large-animal models available for the study of neuropathic pain have, to date, been limited. Animal models of neuropathic pain exist in ovine, equine, and canine systems.32–34Ovine and equine options are limited by Q fever and size concerns, respectively, in many research departments. Dogs pose psychological challenges for some researchers and are susceptible to other diseases that may confound their use as study subjects.33,35,36Though farm pigs have been used to study inflammatory, muscular, and bone pain,16–18no model of neuropathic pain has been developed. Minipigs have been used to generate a partial neuropathic/partial inflammatory pain model19用完全弗氏佐剂和nerve ligation. To test noninvasive treatments for neuropathic pain with the hope of transitioning these treatments to the clinic, a neuropathic pain model is optimal.
农场的猪是理想的用于临床前experimentation as they can be readily introduced and housed in animal facilities and are more cost-effective than minipigs.34,37These animals have circadian rhythms, skin sensitivity, pharmacokinetic characteristics, and anatomy that more closely mimic those of humans.37Anatomically, the location of vertebra, anatomical landmarks, and distance from the skin surface to deeper structures are similar to those in humans. This new porcine system advances the field of preclinical studies by introducing a neuropathic pain model using an animal that has anatomical structure, size, and pharmacokinetic characteristics similar to those of humans without being a significant burden on animal facilities. Of note, there is a high correlation between the use of domestic swine models and the translation of novel treatments to a clinical setting.15In our work, we have also applied this model to a preclinical liFUS treatment directed at the L5 DRG. This novel treatment represents a noninvasive alternative to current neuromodulatory treatments that involve the surgical placement of electric stimulators.38
In the present study, we chose to measure neuropathic allodynia 7 days after CPNI induction. Rodent CPNI models based on VFF responses have shown that nerve ligation results in 180 days of neuropathic allodynia.39Threshold responses remain analogous between time points 7 days and 6 months after surgery. Similar effects have been found in peripheral nerve injury models in large animals. Specifically, previous ovine models of sciatic nerve injury have shown that neuropathic allodynia develops 1 week after surgery34,35,40and continues 6 months after the surgery.34,35,40The 7-day post-CPNI time point further allows us to directly compare our findings with both CPNI and treatments to our previous work in rodents.13,14
As noted in the开云体育世界杯赔率section, modifications were made to optimize the mechanical and thermal testing procedures in swine. The tissue on the ventral aspect of the hoof was too thick to accurately assess nociceptive sensitivity with VFF fibers. This issue was resolved by modifying the procedure to test the skin located on the dorsal surface of the hind foot just above the hoof, which is also innervated by the CPN. We used the modified technique for the final 10 swine. Further, we defined post-CPNI allodynia as a 4.0-g VFF response in this domestic swine model, with 281.4 g as the upper limit of the VFF response. This wide range in VFF response, between the lower allodynic point and the upper fiber limit, allows us to titrate future clinically relevant therapeutic treatments.
Originally, when assessing thermal sensitivity in our swine model, we attempted to implement the same testing paradigm that our laboratory uses on human subjects. This involved testing each site 8 times per session for cool, warm, hot, and cold sensitivity (between 25°C and 45°C). However, this process took 30–40 minutes per animal and we found that animals were unable to tolerate this much time in a sling. Therefore, the total time for each testing session was limited to 20 minutes for all tests. In addition, the Medoc Pathway cutoff temperature was adjusted to 50°C, as the study progressed, to increase the test window available for measuring the effect of therapeutic treatment. With these modifications, Medoc test results were reproducible and quantifiable in all animals and CPNI surgery animals displayed increased sensitivity to heat stimulus. Patients with neuropathic pain show similar increased sensitivity to heat.41Together, these modifications in both testing procedures help validate the allodynia produced by the swine CPNI model and provide a way forward to bridge the gap between preclinical treatments and the translation to clinical trials.
Immunostaining analysis confirmed changes in the ligated CPN. P0 expression was significantly reduced or absent upstream of the ligation site, 2 weeks following surgery. P0 is essential for the formation of the myelin sheath around nerves and is necessary for the normal action potential propagation in the central nervous system and in parts of the peripheral nervous system.27When myelin is disrupted, action potentials fire in an ectopic manner and nerve conduction velocities are lowered. Genetic modification, or knockout, of P0 in mouse models results in neuropathy.27,42Changes observed in P0 expression, in combination with allodynia as measured in VFF and thermal testing, confirm the neuropathic phenotype of this CPNI model in domestic swine.
One notable issue in working with pigs is their rapid growth, which may complicate interpretation of threshold data over time.43These animals gain 0.5 kg per day on average,43which allows researchers to quickly and cost-effectively develop a model that mimics humans. Yet, this rapid growth made tasks such as lifting the pig into the sling for behavioral testing difficult over time. This swine model worked well for our liFUS treatment, because the treatment alleviated neuropathic pain symptoms over a 1-month period.38However, measuring long-term change (such as during a 1-year period) would be difficult in this model, as animals weigh more than 300 kg at full size.43
The novel swine CPNI model of neuropathic pain presented here is a modified version of the sciatic nerve injury surgery commonly used in rats.14,19,25This new large-animal model presents a clinically relevant model for assessment, validation, and translation of research therapeutics for neuropathic pain. The model includes repeatable and quantifiable behavioral and histological measures that make it possible to reproduce quantitative tests for neuropathic pain. Our future work will focus on the implementation of this model to assess the efficacy of liFUS treatment for neuropathic pain.
Conclusions
Here we validate a new neuropathic pain model in domestic swine, bridging the gap between rodent models and future clinical human trials. CPNI resulted in impaired mechanical and thermal sensory thresholds and leg guarding. This neuropathic phenotype was further confirmed at the histological level through P0, which showed evident reductions in myelin structure. This new nerve injury–based neuropathic pain model in swine will aid in the development of treatments for pain that may translate into effective experimental treatments to clinical practice.
Acknowledgments
这项工作得到了国家卫生研究院的基金SBIR 1 R43 NS107076-01A1 (Soft-Focused HIFU [High-Intensity Focused Ultrasound] Treatment of Common Peroneal Nerve Injury) and RO1 NS110627 (Transduction of Mechanical Stimuli in Myelination and Peripheral Nerve Repair). Dr. Pilitsis receives non–study-related grant support from NIH (grants 2R01CA166379-06 and U44NS115111).
Disclosures
Dr. Pilitsis is a consultant for Boston Scientific, Nevro, TerSera, Medtronic, Saluda, and Abbott and receives grant support from Medtronic, Boston Scientific, Abbott, Nevro, and TerSera. She is a medical advisor for Aim Medical Robotics and Karuna and has stock equity.
Author Contributions
Conception and design: Pilitsis. Acquisition of data: Pilitsis, Hellman, Maietta, Clum, Byraju, Raviv, Staudt, Nalwalk, Belin, Poitelon. Analysis and interpretation of data: Hellman, Byraju. Drafting the article: Hellman. Critically revising the article: Pilitsis, Hellman, Nalwalk. Reviewed submitted version of manuscript: Pilitsis. Approved the final version of the manuscript on behalf of all authors: Pilitsis. Statistical analysis: Hellman. Administrative/technical/material support: Jeannotte. Study supervision: Pilitsis.
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40 ↑
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42 ↑
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43 ↑
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