Demyelination and Remyelination of Mouse Spinal Cord Neurons

Protokol

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

1. Prepare Syringe for Injection

1. Dissolve lysolecithin in a 1% solution in phosphate-buffered saline (PBS; pH 7.4) and store at -20 °C in small aliquots (75 μl). Thaw a vial to room temperature (RT).
   NOTE: If lysolecithin is undissolved, sonicate the tube in an ultrasonic cleaner (40 kHz) for approximately 30 min to form a uniform solution.
2. Handle the pre-pulled glass capillary with extreme care to avoid damaging the delicate tip. Unscrew the nut of a 10 μl injecting syringe and thread it onto the flat end of the capillary, followed by two ferrules, ensuring that the mating ends of the ferrules align and the capillary is snug in the conical ferrule (Figure 1). Rinse the syringe with isopropyl alcohol and remove the plunger. Once dry, screw the needle assembly hand tight onto the injecting syringe.
3. Attach the metal hub needle to the priming syringe. Pierce a rubber disc with the needle and slide it down to the base. Fill the priming syringe with lysolecithin. Gently depress the priming syringe until liquid is visible at the tip of the needle. This ensures air bubbles will not be introduced into the injecting syringe.
4. Insert the metal hub needle of the priming syringe into the injecting syringe. Making a firm seal with the rubber disc, slowly depress the priming syringe until the capillary fills the tip with solution. Carefully remove the priming syringe while simultaneously depressing to fill the barrel of the injecting syringe without introducing air bubbles. Insert the plunger into the injecting syringe and ensure the solution flows from the tip of the capillary as the plunger is gently depressed.
   NOTE: If any air bubbles are visible in the capillary or injecting syringe, the syringe preparation must be repeated from the beginning. Continuous fluid in the injecting apparatus is critical to ensure accurate injection volumes.
5. Attach the completed injecting syringe to the arm of a stereotactic micromanipulator. This completed apparatus will be able to inject 15-20 animals before needing to be refilled.
6. Discard the remaining lysolecithin from the priming syringe. Withdraw and depress isopropyl alcohol several times, and detach the metal hub needle. Wait several hours for the remaining isopropyl alcohol in the priming syringe to evaporate before filling again.

2. Prepare the Animal for Surgical Procedure

NOTE: This procedure is described for female C57BL/6 mice aged 8-10 weeks.

1. Anesthetize the animal with an intraperitoneal injection of ketamine (200 mg/kg) and xylazine (10 mg/kg) or per institutional animal care regulations. Plan for the animal to be under anesthesia for approximately 1 hr if using injectable anesthesia.
2. Test that the animal is deeply anesthetized by firmly pinching the foot. A properly anesthetized animal will not respond to the pinch.
3. Using clippers, shave a 2-3 cm² area on the dorsal side of the animal, close to the ears. Be careful not to damage the ears.
4. Wipe the area clean with 70% ethanol applied to a gauze pad. Ensure all clipped hair has been removed from the area. Disinfect the area with iodine.
5. Apply petroleum jelly to the eyes to prevent drying throughout the procedure.
6. Keep the animal in a heated recovery chamber until ready to begin the procedure.

3. Perform the Surgical Procedure

NOTE: Ensure adequate aseptic technique for all steps of the procedure. This includes proper use of gloves, hairnets, masks, and drapes. All tools should be sterilized before coming in contact with the animal.

1. Move the animal to a stereotactic frame, dorsal side up, elevated at the mid-section by folded paper towels to exaggerate the curvature of the spine. Fasten the arms and tail with surgical tape and secure the head with a tooth clamp. Stabilizing ear bars are not necessary for this procedure.
2. Use a scalpel to make a 3 cm midline incision, starting just below the ears and cutting in the caudal direction.
3. Locate the divide between the two large adipose structures and use fine forceps in each hand to pull these apart. Spread retractors to open the surgical field.
4. Under a surgical microscope, locate the prominent outgrowth process of the thoracic vertebra 2 (T2) vertebra (NOTE: This feature is characteristic of the C57BL/6 mouse strain). Perform a blunt dissection with closed spring scissors through the overlaying musculature to better visualize T2. Using the forceps, feel for the hard surfaces of thoracic vertebra 3 (T3) and thoracic vertebra 4 (T4) to confirm proper anatomical location.
5. Using spring scissors, make shallow lateral cuts (2-3 mm deep) of the connective tissue between T3 and T4. Due to the natural spacing between vertebrae in the upper thoracic portion of the mouse vertebral column, a laminectomy is unnecessary to reveal the spinal cord. Be mindful that a cut that is too deep will pierce and damage the cord.
   NOTE: A small degree of bleeding is common during this step. If this occurs, hold a sponge spear into the area until the bleeding subsides (30-60 sec).
6. Visualize the spinal cord. It will be covered with a thick layer of visible dura if this meningeal layer is not yet cut while exposing the cord. A prominent blood vessel runs caudal/rostral through the approximate midline of the spinal cord.
   NOTE: This vasculature should not be used as a landmark for the midline. Instead, adequate lighting should reveal the grey-white matter boundaries flanking the dorsal column, which should be used to estimate the midline.
7. If the dura remains intact, make gentle lateral scrapes with a 32 G metal needle until cleared. The goal is to remove the dura while not cutting the remaining underlying meninges, which are not as thick and harder to see.
   NOTE: The release of cerebrospinal fluid indicates a breach of the arachnoid, and while this can occur without mechanical damage to the tissue, the accumulated cerebrospinal fluid should be removed with a sponge spear to visualize the surface of the cord better.
8. Move the injecting syringe in place and slowly lower it until the tip of the capillary just barely touches the spinal cord immediately lateral of either side of the midline. Lock the arm in place.
9. Use the graded measurements of the Z-direction stereotactic arm to make a baseline position measurement. From this reading, subtract 1.3 mm. Use a quick and shallow downward motion to pierce the tissue and carefully lower the capillary until the new measurement is reached. Optional: if desired, lesions in the dorsal column can be produced by the same piercing motion at the midline and a depth of 0.3 mm.
   NOTE: These values are specific for injecting between T3 and T4. If opting to perform the injection at any other location in the spinal cord, these values must be derived from any available mouse brain atlas.
10. Use the micromanipulator to depress lysolecithin into the ventral spinal cord white matter. Make one rotation of the micromanipulator every 5 sec for 2 min, resulting in a final volume of 0.5 μl. Leave the capillary in place for an additional 2 min to prevent backflow of solution, and then carefully remove the capillary.
11. Tie a single suture in the muscle/adipose tissue overlaying the spinal column. Use a non-interrupted suture to close the skin. Apply more iodine to the incision site.
12. Place the animal in a heated recovery chamber until it recovers, then return it to its cage. Apply analgesics post-operatively as per institutional animal care regulations. Additional post-operative care is usually not necessary as the animals are fully ambulatory and capable of self-feeding and drinking as soon as they recover from anesthesia.
13. Repeat the procedure for the remaining animals.
    NOTE: With proficiency, the operation can be completed in 10-15 min per animal, particularly with the help of a second person tying sutures. The same glass capillary can be used for approximately 15-20 surgeries before the tip becomes blunt and should be replaced. Control surgeries can be performed identically as described, with an injection of PBS into the spinal cord instead of lysolecithin. We do not recommend cleaning the capillaries for future use.

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Sonuçlar

Figure 1. Assembly of the injecting syringe. (A) The nut of the injecting syringe is threaded onto the flat end of the glass capillary, followed by the two ferrules such that their mating ends interlock. Once the capillary is firmly snug in the conical ferrule, the assembly is screwed hand-tight onto the end of the injecting syringe. (B) Piece the center of a rubber disc with the metal hub needle attached to the priming syringe and slide it down to the base. (C) Withdraw lysolecithin solution into the priming syringe. (D) Gently depress the priming syringe until the first drop of lysolecithin is visible at the tip of the needle. (E) Insert the priming syringe into the barrel of the injecting syringe, making a firm seal with the rubber disc. Gently depress the solution until it runs to the end of the capillary. Carefully withdraw the priming syringe while maintaining pressure on the plunger to remove the metal hub needle without introducing air bubbles into the injecting syringe.

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Malzemeler

Name	Company	Catalog Number	Comments
Spring scissors	Fine Science Tools	15004-08
Forceps	Fine Science Tools	11254-20
Retractor	Fine Science Tools	17003-03
Clippers	Philips	QG3330
Heating recovery chamber	Peco Services	V1200
Surgical tape	3M	1527-1
Scalpel handle	Fine Science Tools	10003-12
Scalpel blade	Feather	No. 15
Sponge spear	Beaver Visitec	581089
5-0 Vicryl sutures	Ethicon	J511G
Curved ToughCut spring scissors	Fine Science Tools	15123-12
32 gauge metal needle	BD	305106
Needle holders	Fine Science Tools	12002-14
Angled forceps	Fine Science Tools	11251-35
Cotton tipped applicator	Puritan	806-WC
Gauze pads	Safe Cross First Aid	3763
10 μL syringe	Hamilton	7635-01	Make sure to purchase the microliter, not gastight syringe
Compression fitting	Hamilton	55750-01	Contains the 2 ferrules and removable nut, but the nut that comes with the 10 μL syringe is a tighter fit
Priming kit	Hamilton	PRMKIT	Contains the priming syringe, removable hub needle and the rubber discs
Pre-pulled glass capillaries	WPI	TIP10TW1 (pack of 10)	Contains capillaries with 10 μm inner diameter. 30 μm inner diameter also work (TIP30TW1)
Stereotactic frame	David Kopf instruments	Model 900
Ultrasonic cleaner	Fisher Scientific	FS-20
Microscope slides	VWR	48311-703
Bright field microscope	Olympus	BX51
Lysophosphatidylchoine	Sigma	L1381
PBS	Oxoid	BR0014
Isopropyl alcohol	Sigma	109827
Ketamine	CDMV
Xylazine	CDMV
Iodine	West Penetone	2021
Vaseline petroleum jelly	VWR	CA05971