Inducing White Matter Stroke in a Murine Model Using a Nitric Oxide Inhibitor

Protokół

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

Note: Begin by identifying the target murine population. In prior studies, only male wild-type C57/Bl6 mice have been used. However, various transgenic or knockout mice can also be used. Note that stereotactic coordinates are based on C57/Bl6 anatomy. It is recommended that each user initially verify the localization of the stroke to white matter.

 1. White Matter Stroke Induction - Medial Angled Approach

1. Begin by preparing a pulled glass pipette using a 0.5 mm capillary tube with a distal diameter between 15-25 µm.
2. Prepare a sterile 10 µl aliquot of L-Nio (N(5)-(1)-iminoethyl-L-ornithine hydrochloride) at 27.4 mg/ml (130 µM) in sterile 0.9% normal saline.
3. Pre-fill the pulled glass pipette with a small volume of L-Nio (2-5 µl) by affixing the glass pipette to tubing connected to a vacuum line. Lay the pipette flat on the bench top and insert the pulled end into the L-Nio solution.
   1. Apply the vacuum until at least 2 mm of the 0.5 mm portion of the pipette is filled. Turn off the vacuum and withdraw the pipette. Place it aside until Step 1.12.
4. Place the mouse in an induction chamber and induce anesthesia of the mouse using standard 32% isoflurane flowed through a vaporizer (5 L/min inhaled with 5 L/min oxygen and 0.5 L/min nitrogen) for 1 min or until deeply anesthetized. Transfer the mouse to a stereotactic apparatus equipped with a stereotactic microscope. Provide maintenance anesthesia using 32% isoflurane flowed through a vaporizer (2 L/min inhaled with 5 L/min oxygen and 0.5 L/min nitrogen ) and a nose cone. Check the depth of anesthesia with a toe pinch.
5. Adjust the injection arm to 36°.
6. Affix a pulled glass pipette holder to the distal end of a low-volume pressure injection system and attach it to the injection arm of the stereotactic setup.
7. Coat the anesthetized animal's whiskers with petroleum jelly and place artificial tear ointment over both eyes. Prepare a sterile surgical field by placing a sterile drape over the animal's head with a 5-10 cm opening over the head. Prepare an aseptic surgical surface by shaving the fur overlying the skull. Clean the scalp with alternating betadine and 70% alcohol swabs.
8. Make a 1.5 cm midline scalp incision with sterile fine scissors to expose the skull surface. Dry the skull with a sterile cotton swab, and using a stereotactic microscope at 1-3X magnification, remove any overlying periosteal tissue using a sterile micro-point tool.
9. Mark the Bregma as a reference point using a fine point marker.
10. Drill a 2 mm elliptical craniotomy using a sterile fine stip surgical drill bit beginning posteriorly at the Bregma and extending anteriorly just left of the midline. Remove bone fragments and overlying soft tissue so that the cerebral cortex can be visualized.
11. Keep the surgical field and cortical surface moist by intermittently applying drops of sterile saline.
12. Affix a pulled glass pipette to the injector arm of the stereotactic apparatus. Align the distal end of the pipette with the Bregma and zero the stereotactic coordinates.
13. Advance the pipette to the first anterior/posterior (A/P) and medial/lateral (M/L) coordinates provided in Table 1.
14. Advance the pipette to the cortical surface and zero the dorsal/ventral (D/V) measurement.
15. Slowly pass the pipette into the brain until reaching the first D/V coordinate in Table 1.
16. Using a low-volume pressure injection system set at 20 psi for 20 msec pulses, inject 100 nl of L-Nio into the brain and wait 5 min to prevent reflux up the pipette track.
    1. Use a calibrated reticle in the eyepiece of the stereotactic microscope and a magnification of 3X.
    2. Accordingly, displace a total of 0.100 mm³ (0.5 mm length in a 0.5 mm diameter pipette, corresponding to 100 nl) from the pulled glass pipette for each set of coordinates. By using a reticule, measure and standardize since each set up varies depending on the magnification and scales used.
    3. For accurate volume measurement during each injection, approach the angled pipette with the microscope from the side so that the air-fluid meniscus has a sagittal view. The meniscus should appear in the same focal plane of both the inner and outer walls of the pipette.
17. Slowly withdraw the pipette and repeat steps 1.13-1.16.3 at the second and third set of coordinates provided in Table 1.
18. After the final injection, remove the pipette and place enough bone wax to fill the craniotomy site. Approximate the edges of the scalp wound and bind with dermal adhesive.
19. Inject 0.1 ml of 0.5% Marcaine into the wound margins using a sterile 30 G needle to prevent local pain associated with the scalp incision.
20. Return the animal to housing and supply post-operative antibiotics (0.48 mg/ml trimethoprim-sulfamethoxazole or 0.5 mg/ml Levofloxacin) in the drinking water for 5 days.

Materiały

Name	Company	Catalog Number	Comments
L-N5-(1-Iminoethyl)ornithine, dihydrochloride	Calbiochem	400600-20MG
Isoflurane	Phoenix Pharmaceutical, Inc.	NDC 57319-559-06
Capillary tubes	World Precision Instruments	50-821-807
Picospritzer	Parker Instrumentation	Picospritzer II
Stereotactic setup	Kent Scientific	KSC51725
Pipette puller	KOPF	Model 720
Stereomicroscope SZ51	Olympus	88-124
Fine scissors	Fine Scientific Tools	14084-08
Forceps	Harvard Apparatus	PY2 72-8547
Curved forceps	Harvard Apparatus	PY2 72-8598
Blunt dissection tool	Fine Scientific Tools	10066-15
Drill	Dremel	8220-1/28
Drill bits	Fine Scientific Tools	19007-05
Vetbond	3M	1469SB
Marcaine	HOSPIRA	NDC 0409-1610-50
Trimethoprim-sulfamethaxole	STI Pharmacy	NDC 54879-007-16