Intracranial Cannular Implantation in a Mouse to Deliver Therapeutic Agents to Brain Tumors

Overview

This video presents a procedure for intracranial cannula implantation in a brain tumor mouse model, to facilitate the administration of an immunotherapeutic agent. The delivery of glypican-2-specific chimeric antigen receptor T cells in the tumor microenvironment triggers T cell activation, resulting in tumor lysis and tumor regression.

Protocol

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

1. Preparing the mouse for surgery

1. Anesthetize the mouse in an induction chamber with isoflurane (2-4%) at an oxygen flow rate of 1 L/min until an adequate plane of anesthesia is reached (approximately 5 min).
2. Weigh the mouse using a scale to the nearest 0.1 g and administer subcutaneous slow-release (SR) buprenorphine (1 mg/kg) or other analgesic.
   NOTE: SR buprenorphine provides analgesia for 72 h.
3. Shave the fur on the top of the mouse's head using electric clippers or depilatory agents.
4. Use a spatula to gently open the bottom of the stereotactic arm and insert the guide cannula using forceps. Tighten the screw on the arm to secure the cannula so that approximately 1/2 to 2/3 of the white plastic portion of the cannula is protruding from the bottom of the opening, along with the entire 5 mm metal length of the cannula.
5. Insert and secure the mouse's top teeth in the bite bar of the stereotaxic apparatus. Pull the nose cone forward and tighten it, ensuring the mouse is inhaling isoflurane.
6. Mount the mouse on the warmed stereotaxic apparatus using ear cuffs or ear bars, avoiding excessive pressure.
   NOTE: Warmed tray should have a rectal thermometer inserted, and the warming tray should adjust to maintain a normal body temperature of the mouse during the procedure.
7. Apply sterile ophthalmic ointment to both eyes using a cotton-tipped applicator.
8. Wipe the surgical site with povidone-iodine on a pad or applicator, followed by an alcohol pad. Perform this step three times in total.
9. Before beginning the procedure, perform a toe pinch with forceps to assess for adequate sedation.

2. Surgical procedure

NOTE: All aspects of the surgical procedure utilize sterilized instruments and aseptic techniques. The mice continue under anesthesia with isoflurane (2-4%) throughout the duration of the procedure, approximately 10-20 min.

1. Gently pick up the scalp between the ears with forceps. Using sterile scissors, cut the lifted scalp parallel to the skull and remove an oval flap of skin (0.75-1 cm in length) to expose the skull.
   NOTE: Scissors are preferred over a scalpel to provide a clean, oval-shaped opening and to prevent unnecessary damage to surrounding skin and tissue.
2. Push away fascia using a scalpel or cotton-tipped swabs and a hemostatic cotton pellet to help slow excessive bleeding as needed.
   NOTE: Using the wooden side of a sterile cotton tip can also push away fascia and help avoid excessive bleeding.
3. Identify the landmarks bregma and lambda, the respective anterior and posterior marks on the skull where the cranial plates meet.
   NOTE: Identification can be augmented by wiping the top of the exposed skull with hydrogen peroxide.
4. Gently score the skull using a scalpel to create a surface for the acrylic to attach. Scoring should include multiple linear lines approximately 0.5-1 cm in length at 90° angles to each other.
5. Using the stereotaxic arm, localize the cannula to the landmark of interest (bregma or lambda). Once localized, raise the cannula tip 1-2 mm above the skull surface and move to the desired coordinates. For intratumoral injections, this uses the same A/P and M/L coordinates as the tumor placement.
6. On the exposed skull, away from the area where the cannula will enter, make two screw holes with an 18 G needle or a surgical drill. Ensure that the holes are spaced out to include enough room for the cannula. Using a drill bit, twist through the screw holes until they catch on the skull. Insert and fasten two screws into the holes using a flat-tipped screwdriver. Then, gently pull the screws up to ensure they are secured.
   NOTE: Do not insert the screws until they are flush with the skull, or they may damage the mouse brain underneath. Leave at least a 1-2 mm gap between the screw and the skull.
7. Using an 18 G needle or surgical drill, drill through the skull at the identified coordinates to create a hole for the cannula to be inserted.
8. Using the stereotactic arm, lower the cannula to the desired D/V coordinate.
   NOTE: The D/V coordinate of cannula implantation needs to account for the projection length of the dummy and treatment cannulas, and may be more superficial than the orthotopic injection of tumor cells (Figure 1).
9. In a porcelain 12-well plate, fill one well with acrylic resin powder (approximately 0.3 g) and 10-15 drops (approximately 0.5-0.75 mL) of acrylic resin liquid. This produces a viscous white-colored material. Draw up the mixture into a 1 mL syringe and use it to coat and cover the skull, filling in the spaces around the cannula and screws.
   NOTE: The viscous material hardens into cement over time, so this step should be completed promptly after mixing.
10. While the cement is still pliable, loosen the screw on the stereotactic arm and use a spatula in the opening at the bottom to gently release the cannula from the holder and slowly retract the stereotactic arm upward away from the mouse.
11. Once the cement is completely dry, insert the dummy cannula into the guide cannula and tightly secure it by turning clockwise.
12. Once the procedure is complete, place the mouse back in its warmed home cage to carefully monitor, ensuring adequate recovery and recording any post-procedure observations, including that the mouse has fully regained consciousness, before returning to the colony.
    NOTE: It is generally recommended to place only half of the cage on a heating pad to allow for the animal to move to the cooler side to avoid overheating.
13. Administer additional analgesics as needed if the mice display behaviors indicative of pain post-operatively, such as meloxicam 5 mg/kg subcutaneously delivered once daily for up to 3 days.

3. Preparing chimeric antigen receptor (CAR) T cells

1. Measure the pre-transfected CAR T cell concentration using a cell counter.
2. Centrifuge pre-transfected T cells at 200 x g for 5 min at room temperature (RT).
3. Aspirate the supernatant using a sterile Pasteur pipet on a vacuum aspiration system and resuspend the pellet in phosphatae buffered saline (PBS) to a desired concentration. Typical delivery volumes are 2-5 µL. Typical cell doses are 0.5-5 x 10⁶ cells.

4. CAR T cell infusions

1. Prepare the treatment cannula by feeding the top through a small piece of PKG tubing.
2. Fill the treatment syringe with the CAR T cell suspension and insert it through the other end of the PKG tubing, enough to cover the top of the treatment cannula.
3. Anesthetize the mouse with isoflurane (2-4%) at an oxygen flow rate of 1 L/min.
4. Stabilize the guide cannula using forceps at the base, and then carefully unscrew and remove the dummy cannula, allowing for access into the guide cannula.
   NOTE: A stereotaxic setup is not necessary, but can be used to stabilize the head for treatment.
5. Infuse the CAR T cells for 1 min and hold the treatment cannula in place for an additional 1 min following the cessation of infusion.
6. Remove the treatment cannula and screw the dummy cannula tightly back in place.
7. Administer subcutaneous meloxicam (5 mg/kg) for optional pain control.

Representative Results

Figure 1: Guide cannula with projection dummy and treatment cannulas. (A) Guide cannula with 0.5 mm projection and 2 mm projection dummy cannulas in place. (B) Guide cannula with 0.5 mm projection and 2 mm projection treatment cannulas in place.

Materials

Name	Company	Catalog Number	Comments
18 G needles	BD	511097	1 1/2 inch metal hub
Acrylic resin liquid	Lang Dental	B1323
Acrylic resin powder	Lang Dental	B1323
Alcohol wipes	BD	326895
Centrifuge 5240	Eppendorf	5420000040	Centrifuge
Cotton tipped swabs	Puritan	826-WC	Handle Width = 2.11 mm (0.083), Head Width = 1.27 mm (0.050), Handle Length = 147.62 mm (5.812), Overall Length = 152.4 mm (6), Head Length = 12.7 mm (0.500)
Drill bit holder	P1 Technologies	DH-1	Drill bit holder for D56-D70
Drill bit	P1 Technologies	D58	1.07 mm
Dummy cannula	P1 Technologies	C315DCS-5/SPC	Configuration: Small cap;  Length: Cut 5.00 mm below pedestal; Projection: 0.50 mm
Flat tip screwdriver	P1 Technologies	SD-80	Screwdriver
Graefe forceps	Fine Science Tools	11051-10	Forceps
Guide cannula	P1 Technologies	C315GS-5/SPC	Configuration: 5.00 mm pedestal height; Length: Cut 5.00 mm below pedestal
Hemostatic cotton pellets with racemic epinephrine	Pascal	1151602
MOXI Z Mini automated cell counter Kit	Moxi	MXZ001	Cell counter
NOD scid gamma (NSG) mice	Jackson Laboratory	5557	6 to 12-week-old males and females
Pasteur pipet	VWR	14673-043
PKG tubing	P1 Technologies	C313CT	Diameter: 0.58 mm x 1.27 mm
Porcelain 12 well plate	Flinn Scientific	AP6064
Povidone iodine	Medline	MDS093943
Scalpel	World Precision Instrument	50-822-457	Disposable Scalpel, no.10, sterile, 10/box, Plastic Handle with 6" Ruler
Screws	P1 Technologies	0-80 X 3/32	2.4 mm
Stereotaxic Frame	David Kopf Instruments	940	Model 940 Small Animal Stereotaxic Instrument with Digital Display Console
Student fine scissors	Fine Science Tools	91460-12	Scissors
Treatment cannula	P1 Technologies	C315IS-5/SPC	33GA; Configuration: Standard internal; Length: Cut 5.00 mm below pedestal; Projection: 0.50 mm
Treatment syringes	Hamilton	87908	5 µL, Model 75 Cemented Needle Special (SN) Syringe, 75SN/22/0.5"/PT3
Vactrap XL	Foxx Life Sciences	305-4401-FLS	Vacuum System