Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve

Summary

We evaluated the effect of cervical sympathetic ganglion block on nerve repair using artificial nerve conduits. Male beagle dogs were each implanted with an artificial nerve across a 10-mm gap in the left inferior alveolar nerve; left cervical sympathetic ganglion was blocked by injecting 99.5% ethanol via lateral thoracotomy.

Abstract

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen films. Favorable clinical outcomes have been achieved when using these tubes for the treatment of damaged inferior alveolar nerve (IAN). A critical factor for the successful nerve regeneration using PGA-C tubes is blood supply to the surrounding tissue. Cervical sympathetic ganglion block (CSGB) creates a sympathetic blockade in the head and neck region thus increasing blood flow in the area. To ensure an adequate effect, the blockade must be administered with local anesthetics one to two times a day for several consecutive weeks; this poses a challenge when creating animal models for investigating this technique. To address this limitation, we developed an ethanol-induced CSGB in a canine model of long-term increase in blood flow in the orofacial region. We examined whether IAN regeneration via PGA-C tube implantation can be enhanced by this model. Fourteen Beagles were each implanted with a PGA-C tube across a 10-mm gap in the left IAN. The IAN is located within the mandibular canal surrounded by bone, therefore we chose piezoelectric surgery, consisting of ultrasonic waves, for bone processing, in order to minimize the risk of nerve and vessel injury. A good surgical outcome was obtained with this approach. A week after surgery, seven of these dogs were subjected to left CSGB by injection of ethanol. Ethanol-induced CSGB resulted in improved nerve regeneration, suggesting that the increased blood flow effectively promotes nerve regeneration in IAN defects. This canine model can contribute to further research on the long-term effects of CSGB.

Introduction

In many cases, traumatic injury of the inferior alveolar nerve (IAN) is iatrogenic, being frequently caused by the extraction of the third molar or the placement of dental implants^1,2,3. Injury of the IAN can lead to deficits in thermal and touch sensations as well as paresthesia, dysesthesia, hypoesthesia, and allodynia. Nerve injury is treated not only by conservative therapy but also by other methods, including suturing and autograft placement. However, these methods have drawbacks, which often include the lack of symptom improvement and neurological defects at the donor site^4,5,6.

The artificial nerve — polyglycolic acid-collagen (PGA-C) tube was originally developed in Japan. It is a bio-absorbable tube with its inner lumen filled with a spongiform collagen⁷. In animal experiments, this tube was used to enhance nerve regeneration in beagle dogs with peroneal nerve defect, and was shown to promote higher level of recovery than autologous nerve transplantation⁸. The clinical application of the PGA-C tube began in 2002 in patients with peripheral nerve injuries. Moreover, favorable clinical outcomes have been achieved in the treatment of trigeminal neuropathy (IAN and lingual nerve)^9,10,11. A critical factor for successful nerve regeneration using PGA-C tubes is blood supply to the surrounding tissue⁸. Cervical sympathetic ganglion block (CSGB) creates a sympathetic blockade in the head and neck region and increases blood flow to the respective innervated area¹²; thus, it has been used in the treatment of complex regional pain syndrome and circulatory insufficiency^13,14,15. However, there have been only a few experimental investigations on the efficacy of CSGB in increasing blood flow^16,17. To ensure adequate CSGB efficiency, the blockade must be applied together with local anesthetics once or twice daily for several weeks, thus posing a challenge when generating animal models to investigate this technique. To address this limitation, in a previous study, we developed a canine model of long-term increased blood flow in the orofacial region¹⁸. The model was generated by performing a CSGB by injecting 99.5% ethanol. We evaluated the oral mucosal blood flow and nasal skin temperature by laser Doppler flowmetry and infrared thermography once per week for 12 weeks. We found that the blood flow of the orofacial region was increased for 7 – 10 weeks in this model.

In the present study, we evaluated the effects of ethanol-induced CSGB on nerve regeneration.
The PGA-C tube was implanted into beagle dogs across a 10-mm gap in the left IAN. A week later, CSGB was performed by injecting ethanol. Three months after surgery, we performed a variety of electrophysiological, histological, and morphological studies to evaluate the effects of CSGB on nerve regeneration. We provide a detailed protocol for IAN reconstruction using a PGA-C tube and ethanol-induced CSGB.

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Protocol

This study was conducted in accordance with the Guiding Principles for the Care and Use of Animals and approved by the Committee for Animal Research of Kyoto University (Kyoto, Japan; authorization number: R-16-16). All efforts were made to minimize animal suffering, and all sections of this report adhere to the ARRIVE (Animal Research: Reporting of in Vivo Experiments) guidelines.

1. Fabrication of the PGA-C tube

1. To fabricate the artificial nerve conduit by means of an absorbable polyglycolic acid (PGA) tube, use a tubular braiding machine equipped with 48 spindles and five PGA fibers, comprised of bundles of 26 filaments (Figure 1)¹⁸.
2. To render the PGA tube surface hydrophilic, expose it to plasma discharge.
3. Use 1% v/w atelocollagen in hydrochloride solution⁷.
   NOTE: Atelocollagen is extracted from porcine skin via enzyme treatment and subjected to a virus check. It mainly consists of type I (70–80%) and type III collagen, the ratio of which is described in detail elsewhere⁷. Prepare the collagen solution by dissolving 1 g collagen in 100 mL hydrochloride solution (pH = 3.0). Since the density of the hydrochloride solution is approximately 1.0, the w/w collagen concentration is almost 1%.
4. Coat the tube with the collagen layers by repeatedly dipping it into the 1% collagen hydrochloride solution for 5 s each time.
   1. After dipping, dry the tube on a clean bench at room temperature. Perform next dipping after ensuring the tube is completely dry (about 6 h for air-drying).
   2. Repeat the coating process 10 times.
5. Subject the PGA-C tube to 140 °C for 24 h under vacuum (dehydrothermal treatment), in order to control bio-absorption and crosslinking of the collagen molecules. Perform the entire process under aseptic conditions.
   NOTE: This procedure generates a tube of 14-mm final length, 3-mm inner diameter, and 50-μm wall thickness.

2. Surgical Procedure Set-up

1. Use adult male beagles weighing 9.0 to 13.0 kg.
   1. House animals in separate cages, under controlled kennel conditions (12-h light and dark cycle).
   2. Provide solid food and water ad libitum.
2. Weigh the beagles.
3. Autoclave all surgical instruments.
4. Don sterilized gloves and disinfect all surfaces of the operating setting with an 80% ethanol solution. Discard the used gloves.
5. Perform surgical handwashing.
6. Put on a fresh mask, gown, and sterile gloves.

3. Anesthesia and Skin Preparation

1. Anesthetize the dog with a mixture of 5 mg/kg ketamine hydrochloride and 1 mg/kg xylazine by an intramuscular injection.
2. Intubate by a tracheal tube of 7.5 mm diameter and 25 cm length.
3. Place the dog on the right lateral position. Maintain general anesthesia with 3.2% sevoflurane with oxygen (1.0 L/min).
4. Use a heating pad to maintain body temperature at 37 °C.
5. Apply an ophthalmic gel over the anterior surface of the eyes to avoid corneal abrasion.
6. Carefully shave the surgical field (left side chest area) using surgical clippers. Spray a substantial amount of alcoholic solution over the operative site. Wait at least 15 sec. Repeat the application 3 times.
7. Record the heart rate and oxygen saturation during surgery.

4. Inferior Alveolar Nerve Reconstruction Using PGA-C tube: Development of the Reconstruction-only Model

1. Inject 3 mL of 1% lidocaine using a 27 G needle to the left mandibular gingiva as a local anesthetic and analgesic.
2. Perform a 5-cm transverse incision with a number 15 scalpel blade in the left mandibular gingiva, to expose the mandibles of the animal.
3. Use piezoelectric ultrasonic vibrations to grind the proximal aspect of the mandible into a 3-cm × 8-mm rectangle through the posterior mental foramen.
   NOTE: The vibration frequency was 28 ‒ 32 kHz.
4. Remove the frontal part of the mandibular bone plate (dimensions, 3 cm × 8 mm) to expose the left IAN (Figure 2A)¹⁸.
   NOTE: The reconstruction site corresponds to the root apex of the first molar
5. Transect the IAN with a scalpel to remove a 10-mm segment.
6. Insert the proximal and distal stumps of the severed nerve into the nerve tube to a depth of 2 mm.
7. Use 8-0 nylon sutures and a surgical microscope at 8X magnification to suture the tube to the proximal and distal nerve ends (Figure 2B)¹⁸.
8. Return the bone plate to its original site in the mandible.
9. Close the wound with 4-0 nylon sutures.
10. One day after surgery, confirm that the mandibular bone plate is in its proper position.
    1. 4.10.1 Perform computed tomography (CT) imaging of the facial bone under anesthesia. Set CT parameters as follows: 120 kVp, 200 mAs, 0.5 mm/s, 0.5-mm slice thickness.
       1. Administer anesthesia using a mixture of 5 mg/kg ketamine hydrochloride and 1 mg/kg xylazine (Figure 3).
11. Administer ampicillin (100 mg/day) as an antibiotic and acetaminophen (100 mg/day) as an analgesic for a week after surgery.

5. Ethanol-induced CSGB: Development of the Reconstruction + CSGB Model

1. Perform IAN reconstruction as described in section 4 and allow a week for recovery.
2. Anesthetize the animal with 1.5% sevoflurane in oxygen (4 L/min) and air (6 L/min). Shave and clean the intended surgical field, as described in section 3.
3. Mark the incision line with a surgical skin marker by drawing a line on the left side chest area (Figure 4, the incision line is 20 cm in length).
4. Inject 5 mL of 1% lidocaine using a 21 G needle to the left side chest area as a local anesthetic and analgesic.
5. Incise the left side chest skin with a number 10 scalpel blade.
6. Incise the fat layer with an electric scalpel to expose the muscle fascia.
7. Expose the serratus ventralis muscle and scalenus muscle.
8. Raise the serratus ventralis muscle and scalenus muscle from ventral to dorsal to expose the second and third ribs (Figure 5).
9. Perform a left lateral thoracotomy at the second and third intercostal space to expose the left cervical sympathetic ganglion (Figure 6).
10. Inject 0.2 mL of 99.5% ethanol into the cervical sympathetic ganglion using a 30 G needle under direct visualization (Figure 7).
11. Close the intercostal space with interrupted 1-0 absorbable stitches.
12. Close the skin with interrupted 3-0 nylon stitches.
13. Administer ampicillin (100 mg/day) as an antibiotic and acetaminophen (100 mg/day) as an analgesic for a week after surgery.
14. At 1 week after CSGB, measure facial skin temperature with infrared thermography to confirm the CSGB.

6. Electrophysiological Recordings

1. To measure sensory nerve action potential (SNAP) and sensory nerve conduction velocity (SCV) of the IAN three months after reconstruction, anesthetize animals as described in section 3.
   NOTE: SNAP and SCV should be measured on both the experimental and normal control sides for each dog in both treatment groups.
2. Make an incision in the left mandibular gingiva with a number 10 scalpel blade.
3. Carefully remove the mandibular bone plate to avoid physically damaging the regenerated nerve.
4. Stimulate the IAN using a pair of needle electrodes, to record the SNAP and SCV.
   1. Insert the electrodes proximally to the nerve conduit.
   2. Apply 10-kHz electrical stimulus 20 times.
5. Analyze the results.
   1. Determine SNAP by calculating the average response amplitude to the electrical stimulation.
   2. Measure the peak latency and peak amplitude from the chart recordings.
   3. Calculate the recovery index with the following equation: peak amplitude of the left IAN of the reconstruction-only or reconstruction + CSGB group / peak amplitude of the normal control (central segment of the right IAN in the reconstruction-only group)^19,20.

7. Histological Analysis

1. Section Preparation
   1. Three months after reconstruction, harvest the left IAN, including 1 cm of the nerve on either side of the reconstructed site.
   2. Harvest the right IAN at the level corresponding to the harvest site on the left side.
   3. Prefix the harvested nerves by immersion in 2.5% glutaraldehyde in a 0.1 M cacodylate buffer solution (pH 7.4, 48 °C, 24 h).
   4. Postfix with 2% osmium tetroxide solution (48 °C, 4 h) and potassium ferrocyanide in 0.1 M phosphate buffer solution (pH 7.4, 2 h).
   5. Dehydrate the nerves with a series of graded ethanol solutions.
   6. Embed in epoxy resin (paraffin).
   7. Section the specimens at a thickness of 0.5 ‒ 1.0 μm.
2. Toluidine Blue Staining and Morphological Analysis
   1. Stain sections with toluidine blue solution.
   2. Obtain microscopy images using an optical microscope, at 400X magnification at the following regions along the samples: left IAN, the center of the regenerated segment and 2 mm distally to the stump; right IAN, the center of the IAN segment corresponding to the harvest site on the left side.
   3. Select images of all regions with regenerated nerve fibers.
      1. Randomly select 8 ‒ 10 areas of 100 μm × 100 μm containing regenerated nerve fibers.
      2. Perform morphological analysis using an appropriate software to measure the following parameters: myelinated nerve fiber diameter (μm) and density (count/area), nerve tissue percentage, and G-ratio (myelinated axon diameter/myelinated nerve fiber diameter).
3. Immunostaining
   1. Follow standard protocols for paraffin section staining.
   2. Incubate with primary antibodies for 30 min at 25 °C.
   3. Wash with phosphate-buffered saline 3 times at 25 °C.
   4. Incubate with secondary antibodies labeled with horseradish-peroxidase for 30 min at 25 °C.
   5. Obtain images using a light microscope.
4. Transmission Electron Microscopy (TEM)
   1. Prepare nerves as described in step 7.1.
   2. Section nerves at a thickness of 70 ‒ 90 μm using an ultramicrotome.
   3. Stain sections with Reynold’s lead citrate and uranyl.
   4. Examine and image by transmission electron microscopy.

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Results

We observed an increase in the facial skin temperature of the blocked side 1 week after the left CSGB (Figure 8).

At 3 months post-reconstruction, the PGA-C tube at the reconstruction area was absorbed and regeneration of the inferior alveolar nerve was observed in the reconstruction-only and reconstruction + CSGB groups (Figure 9A, B)^...

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Discussion

We present an efficient method for IAN regeneration by using a bioabsorbable nerve tube in combination with ethanol-induced CSGB. For this study we used dogs, since other animal models, like mice, rats, and rabbits, have a short life expectancy and small body size, and hence cannot be used to perform the precise surgical procedures. As the IAN is located within the mandibular canal surrounded by bone, a surgical technique is necessary to avoid nerve and blood vessel damage when performing nerve reconstruction. An importa...

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Materials

Name	Company	Catalog Number	Comments
NMP Collagen PS	Nippon Meatpackers	301-84621	Atelocollagen extracted from young porcine skin by enzyme treatment
Surgical clippers	Roboz Surgical Instrument Company	RC-5903
Disposable scalpel (No.15)	Kai medical	219ABBZX00073000
VarioSurg3	Nakanishi	VS3-LED-HPSC, E1133	Piezoelectric surgery for bone processing
4-0 nylon sutures	Ethicon	8881H
8-0 nylon sutures	Ethicon	2775G
Isepamicin sulfate	Nichi-Iko	620005641
Disposable scalpel (No.10)	Kai medical	219ABBZX00073000
30-gauge needle	Nipro	1134
1-0 absorbable stitches	Ethicon	J347H
3-0 Nylon stitches	Ethicon	8872H
Neo Thermo	NEC Avio	TVS-700	Infrared thermography
Neuropack Σ	NIHON KOHDEN	MEB-5504	Orthodromic recorder for electrophysiological recording
Toluidine Blue	Sigma-Aldrich	T3260-5G
Light microscope	Keyence	BZ-9000
Mouse anti-human neurofilament protein monoclonal antibody	DAKO	N1591
Polyclonal rabbit anti-S100 antibody	DAKO	Z0311
Transmission electron microscopy	Hitachi High Technologies	Hitachi H-7000
Dynamic cell count	Keyence	BZ-H1C	Software for morphological evaluation