Inducing Fibular Nerve Injury in a Mouse to Study Neuromuscular Junction Regeneration

Samenvatting

Source: Dalkin, W., et al. The Fibular Nerve Injury Method: A Reliable Assay to Identify and Test Factors That Repair Neuromuscular Junctions. J. Vis. Exp. (2016)

This video demonstrates a surgical procedure to induce fibular nerve injury in a mouse to study neuromuscular junction (NMJ) regeneration. The procedure involves exposing the fibular nerve, applying controlled pressure to induce injury, and ensuring the epineurium remains intact to support axonal regeneration. Over time, macrophages clear axonal debris, facilitating nerve repair and the reestablishment of NMJs.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparing Animals for Surgery

1. Anesthetize mice with a mixture of ketamine (90 mg/kg) and xylazine (10 mg/kg) via subcutaneous inguinal injection with a sterile 1 ml insulin syringe. The carrier solution contains a mixture of 0.9% saline, 17.4 mg/ml ketamine, and 2.6 mg/ml xylazine. Place animals back in cages while waiting for medication to take effect.
   NOTE: If the loading dose does not provide sufficient anesthesia for the duration of the procedure, an additional 25% of the loading dose may be injected.
2. Monitor animals post-injection to check for steady respiratory rates and appropriate depression of arousal levels. Check arousal level with a hind foot pinch, which should elicit no response when sufficiently anesthetized.
   NOTE: This usually takes 3-5 min for a young adult mouse averaging 25 to 30 g. If the animal is still responsive after 10 minutes post-injection, an additional 25% of the anesthetic loading dose may be injected.
3. Apply petrolatum and light mineral oil ophthalmic ointment to the animal's eyes to prevent dryness. Remove animals from the cage and place them on a clean, flat surface. Shave the desired hind limb from foot to pelvis using an electric hair trimmer, exposing only the lateral aspect of the limb.
4. Apply a chemical hair remover to the shaved site for 1 min. Manually remove the hair using laboratory wipes. Clean the depilated area with a laboratory wipe soaked in ethanol.

2. Surgical Procedure

1. Sterilize surgical instruments via autoclave or other appropriate method. Clean the surgical site and surgical board with 80% ethanol/H₂O. Disinfect the surgical site with providone. Place the mouse on the surgical board and align it with the limb restraints. Keep the target hind limb in an anatomically natural position with the knee joint slightly extended without internal or external rotation.
2. Place the animal and board under the surgical microscope. Orient to the proper incision site via palpation of superficial landmarks, specifically the bony knee joint and the ridge between the tibialis anterior and gastrocnemius muscles.
3. Make an approximately 3 cm incision through the skin using a scalpel or spring scissors while using general forceps for gripping. Make the incision perpendicular to the underlying course of the common fibular nerve.
4. Continue the incision through the superficial fascia, exposing the biceps femoris and vastus lateralis muscles. Separate these muscles by cutting through the connecting deep fascia. A 1-2 cm cut should be sufficient.
5. Retract the biceps femoris muscle caudally using mechanical retractors, revealing the common fibular nerve.
6. Trace the nerve proximally until its intersection with the tendon of the lateral head of the gastrocnemius muscle is found.
   Note: Exposure may require additional manipulation of the retracted skin and muscle. This intersection is used as a stable landmark for nerve injury.
7. Grasp the nerve with a fine forceps, aligning the tips in a parallel fashion to the lateral border of the gastrocnemius tendon. Crush the common fibular nerve by applying steady, hard pressure for 5 sec.
8. Corroborate the full crush of the nerve by visual inspection through the surgical scope. It will appear translucent at the site of injury. If using mice expressing fluorescence proteins in peripheral axons, the fluorescence will disappear from the site of injury.
9. Remove retractors and realign muscles in their anatomic positions. Close the incision site with 6-0 silk sutures. 1-3 simple interrupted sutures is sufficient. Place the recuperating mouse on a heating pad in a clean cage.
10. Monitor all animals for 2 hr post-operation to check for breathing and any adverse reactions to the anesthesia. Administer an initial dose of buprenorphine 0.05-0.10 mg/kg via subcutaneous inguinal injection immediately following recovery from surgery. Give 3 additional doses every 12 hr over the next 48 hr. After full recovery, return the mice to the animal care facility.

Materialen

Lijst van materialen gebruikt in dit artikel

Naam	Bedrijf	Catalogusnummer	Opmerkingen
Ketamine	VetOne	501072
Xylazine	Lloyd Inc.	3437
Buprenorphine	Zoopharm	1Z-73000-150910
Nair	Nair
Kim-wipes	Kimtech	34155
Electric Razor	Braintree Scientific	CLP-64800
80% EtOH/H₂O
10% Proviodine
1 ml Insulin Syringe
Spring Scissors	Vannas	91500-09
No. 15 scalpel	Braintree Scientific	SSS 15
#5 Forceps	Dumont	11252-00
6-0 silk suture on reverse cutting needle	Suture Express	752B
Rodent Heating Pad	Braintree Scientific	AP-R-18.5
Alexa 555 conjugated alpha-BTX	Molecular Probes	B35451
Vectashield	Vector Labs	H-1000
Olympus Stereo Zoom Microscope	Olympus	562037192
Zeiss 700 Confocal Microscope	Zeiss
Variable-flow peristaltic perfusion pump	Fisher Scientific	13-876-3