A Murine Model of Cervical Spinal Cord Injury to Study Post-lesional Respiratory Neuroplasticity

Summary

Respiratory failure is the leading cause of death following a cervical spinal cord injury. Having a reproducible, quantifiable, and reliable pre-clinical animal model of respiratory failure induced by a partial cervical injury will help to understand the subsequent respiratory and non-respiratory neuroplasticity and allow testing putative repair strategies.

Abstract

A cervical spinal cord injury induces permanent paralysis, and often leads to respiratory distress. To date, no efficient therapeutics have been developed to improve/ameliorate the respiratory failure following high cervical spinal cord injury (SCI). Here we propose a murine pre-clinical model of high SCI at the cervical 2 (C2) metameric level to study diverse post-lesional respiratory neuroplasticity. The technique consists of a surgical partial injury at the C2 level, which will induce a hemiparalysis of the diaphragm due to a deafferentation of the phrenic motoneurons from the respiratory centers located in the brainstem. The contralateral side of the injury remains intact and allows the animal recovery. Unlike other SCIs which affect the locomotor function (at the thoracic and lumbar level), the respiratory function does not require animal motivation and the quantification of the deficit/recovery can be easily performed (diaphragm and phrenic nerve recordings, whole body ventilation). This pre-clinical C2 SCI model is a powerful, useful, and reliable pre-clinical model to study various respiratory and non-respiratory neuroplasticity events at different levels (molecular to physiology) and to test diverse putative therapeutic strategies which might improve the respiration in SCI patients.

Introduction

Spinal cord trauma is a common injury observed in the human population with dramatic incidences, such as permanent paralysis. However, the severity of the injury depends on the level and the extent of the initial trauma. Respiratory failure is the leading cause of mortality following upper cervical spinal cord injury (SCI)¹. Currently, the only therapeutic treatment is to place the patient under ventilatory assistance. Since few patients can be weaned off the ventilatory assistance², due to spontaneous recovery which occurs with post-lesional delay, the need to develop new innovative non-invasive therapeutics is urgent³. Having a good standardized pre-clinical model to investigate the effect of a cervical SCI on respiratory insufficiency and therefore, to study the application of putative therapeutic strategies, is essential.

In this technical article, we describe a specific pre-clinical murine model of respiratory impairment induced by a partial cervical SCI at the C2 level. This model is currently used by several laboratories around the world (for reviews: ^4-13). However, slight differences in the surgical procedure can be observed among the different investigators to generate this particular cervical injury murine model. The effect of a C2 SCI on the respiratory output was first described in 1895 by Porter¹⁴. A cervical hemisection induces a deafferentation of the phrenic motoneurons from their central drive (located in the rVRG in the brainstem, Figure 1A) on the ipsilateral side of injury, leading to a silent phrenic nerve activity and the subsequent diaphragm paralysis. The contralateral side remains intact and allows the animal to survive. Unlike different SCI located in a lower spinal segment (for example a contusive injury at C4 level¹⁵), the integrity of the phrenic motoneuron nucleus on both side is preserved. After a cervical C2 injury, some spontaneous activity can be observed on the ipsilateral side (phrenic and diaphragm) due to an activation of contralateral silent synaptic pathways which crossed the spinal midline at the segmental level C3-C6 (Crossed phrenic pathways, CPP, Figure 1B). The activation of the CPP, which is, by definition, a C2 hemisection combined with a contralateral phrenicotomy which induce an ipsilateral partial phrenic nerve recovery, can occur from hours to weeks post-injury^16-18. The real beneficial effect of this CPP pathway on the respiratory recovery is limited¹⁹ and further investigation and treatment should be developed to improve the magnitude of spontaneous restoration³.

This protocol provides a powerful type of pre-clinical murine model to study respiratory post-lesional plasticity at various levels (respiratory physiology from pre and phrenic motoneurons, interneurons, molecular and cellular, locomotion of the front limb for example) as well as a model to test invasive and non-invasive therapeutic strategies aimed to improve the respiratory and locomotor recovery following C2 partial cervical spinal cord injury.

Protocol

This protocol was approved by the Ethics committee of the RBUCE-UP chair of Excellence (University of Paris Sud, grant agreement No. 246556) and the Université de Versailles Saint-Quentin-en-Yvelines.

1. Preparation of Sterilized Surgical Instruments

1. Clean the surgical instruments with laboratory detergent.
2. Autoclave the instruments prior to surgery.
3. In a surgical session, sterilize the tools by placing the tips in a hot bead sterilizer for 10 min at 180 °C between 2 surgeries.

2. Preparation of Drugs

1. Prepare 2 x 1 ml syringes for pre-anesthetic drugs cocktail and post-operation drugs.
2. According to the weight of the rat, prepare a syringe with pre-anesthetic drugs: Carprofen (5 mg/kg), Buprenorphine (50 μg/kg), Baytril (5 mg/kg) and Dexmedetomidine (0.5 mg/kg). Complete the volume to 1 ml with Lactated Ringers.
3. Prepare in another syringe the reversal for the pre-anesthetic drugs: Atipamezole (500 μg/kg).

3. Anesthesia of the Rat

1. Administer subcutaneously to the animal the solution of pre-anesthetic drugs described in step 2.2. Then, put the animal back in cage and wait until the sedative effect appears.
2. Place the rat into a closed chamber filled up with 5% isoflurane in 100% O₂, and wait until the respiratory rhythm slows down (around 30 sec). Then, remove the rat from the chamber and place it onto the intubation table.

4. Orotracheal Intubation

1. Lie the animal on his back, then secure the head by placing a strap attached to his front teeth to the table.
2. With a fiber optic light, light up the thoracic space. Then, place a laryngoscope (or a custom made one, Jou et al.²⁰ for details) in the animal’s mouth. Visualize the vocal cords.
3. Slide and place an orotracheal guide into the trachea (between the vocal cords). Slide the orotracheal tube (16 G catheter size) on the guide.
4. Remove the guide and check with a laryngeal mirror placed at the end of the orotracheal tube for the presence of moisture, confirming the appropriate position of the tube in the trachea and not in the esophagus.
5. Connect the tube to a rodent ventilator (683 rodent ventilator, Harvard Apparatus) and adjust the concentration of isoflurane to 2% (in 100% O₂).
6. Secure the orotracheal tube with surgical tape.

5. Spinal Surgery

1. Place the animal in ventral decubitus position on a heated surgical plate, with the nose pointing at 90° angle to the surgeon. Maintain the body temperature around 37.5 °C throughout the surgery.
2. Shave the hair with clippers between the scapulas and remove the hair with gauze.
3. Clean the skin with betadine, then with 70% alcohol. Repeat this step 3x. 
4. A toe pinch is performed before beginning of surgery to ensure proper anesthetic depth. Then, perform a lateral skin incision rostro-caudally with scissors between the scapulas.
5. Cut the acromiotrapezius muscle rostro-caudally by following the tendon to prevent any bleeding. Then dissociate the rhomboid muscle to access the spinalis muscles (surrounding the vertebra).
6. Retract the spinalis muscle from C1 to C3 vertebra. The C2 vertebra is the one with a prominent apophysis.
7. Clean the muscle around the dorsal part of the vertebra by using sterile cotton swabs.
8. Start to remove carefully the apophysis of C2 with a rongeur. Then, continue meticulously until the dorsal spinal cord is exposed. Ensure that laminectomy is a dorsal hemi laminectomy. Pay careful attention to the dura which encloses the spinal cord, and arteries in the vicinity of this area.
9. With a #55 forceps, dissect rostro-caudally the dura along C2, next continue laterally on each rostral and caudal side.
10. Sponge up the cerebrospinal fluid.
11. Make a lateral section under the cervical dorsal root number 2 with the microscissors. Verify with a micro scalpel that the extent of the lesion is close enough to reach the midline of the spinal cord (see Figure 2A for a dorsal view of the injury). If not, then another cut could be made to complete the injury. In case of bleeding, use sterile cotton swabs. Be careful not to go to the contralateral side, otherwise, the animal will not recover from the injury and will have a respiratory failure.
12. Suture the muscles as a protective layer and suture back the skin. Clean the wound with Betadine saturated sterile gauze.
13. Turn-off the isoflurane vaporizer and inject the reversal drugs (Atipamezole [500 μg/kg, i.m.]), check the body temperature.
14. When the animal starts to breathe against the ventilator, disconnect the tracheal tube from the ventilator, and then remove the orotracheal tube. Place the animal in a heated cage for recovery.

6. Post-surgical care

Following surgery, the animals are monitored continuously to ensure the best environment possible for recovery. Antibiotics (Baytril, 5 mg/kg), anti-inflammatory (Carprofen, 5 mg/kg) and Buprenorphine (50 μg/kg) drugs are given every 12 hr for the first 2 days post-surgery to prevent infections and reduce the occurrence of post-operative pain. Rats have access ad libitum to soft food and water (or jellified water for the 1st post-operative day). Subcutaneous fluids can be used to prevent dehydration on the first few post-operative days. The body weight and the food intake are monitored daily. Their environment is enriched throughout the experiment and time post-injury (dual housing, tubing in their cages).

Results

Extent of Injury

The success and the reproducibility of this particular experimental model are dependent on the experience of each manipulator/surgeon. The subsequent amount of respiratory recovery (phrenic nerve activity and diaphragm activity) following a C2 injury is correlated with the remaining ventrolateral spared white matter²¹. Since the injury is “handmade” and requires some practice from the surgeon, the extent of each injury has to be checked by histologi...

Discussion

Technical Difficulties of Making the C2 Injury Model

The C2 injury murine model is an interesting tool to study respiratory post-lesional neuroplasticity. However, the steps needed to produce a reproducible and reliable model are numerous and each one could impact on the outcome of the study. For example, during the intubation process, extreme care is to be taken since the orotracheal tube can produce an inflammation of the trachea, which can lead to diverse complications such...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Animal
Male Sprague Dawley Rat	Janvier		225-250 g
Surgical Instruments
Student Dumont #5 forceps	Fine Science Tool	91150-20
Student Standard Pattern Forceps	Fine Science Tool	91100-12
Mayo-Stille Scissors	Fine Science Tool	14013-15	Curved
Student Vannas Spring Scissors	Fine Science Tool	91500-09	Straight
Spring Scissors - 8 mm Blades	Fine Science Tool	15025-10	Straight Blunt/Blunt
Friedman Pearson Rongeur	Fine Science Tool	16121-14	Curved
Dissecting Knife - Fine Tip	Fine Science Tool	10055-12	Straight
Olsen-Hegar Needle Holder	Fine Science Tool	12002-14	Serrated
Weitlaner-Locktite Retractor	Fine Science Tool	17012-11	2x3 Blunt
Absorbable surgical sutures	Centravet	BYO001	Suture size 4-0
Equipment
Hot Bead Steriliser	Fine Science Tool	18000-45
Catheter	Centravet	CAT188	16 G
Laryngoscope
Guide wire
Laryngeal mirror	Centravet	MIR011
Lactated Ringers	Centravet	RIN020
Syringe	Centravet
Needle	Centravet
O2	Air Liquid	I1001M20R2A001
683 RodentT Ventilator 115/230V	Harvard Apparatus	55-0000
Stand-Alone Vaporizer	WPI	EZ-155
Thin line heated bed	WPI	EZ-211
Air canister	WPI	EZ-258
Drugs
Carprofen	Centravet
Rimadyl	Centravet	RIM011
Buprenorphine	Centravet	BUP001
Baytril	Centravet	BAY001
Dexmedetomidine	Centravet	DEX010
Atipamezole	Centravet	ANT201
Betadine solution	Centravet	VET002
Isoflurane	Centravet	VET066