A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease

The Murine Ommaya, which mimics the Ommaya reservoir used clinically, allows researchers to test a variety of direct targeted therapies for brain metastases and universally fatal leptomeningeal disease in a preclinical model. The advantage of this technique is that researchers can deliver microliter volume of drugs directly into the cerebrospinal fluid to treat central nervous system metastases, bypassing the blood-brain barrier. This Murine Ommaya implantation protocol is designed for researchers or technicians who have stereotactic surgery experience and aseptic technique training, which are essential to ensure the success of this procedure.

Demonstrating the procedure will be Margie Baldwin and Michelle Danielson from Comparative Medicine at the University of South Florida, and Vincent Law, a senior research associate from Dr.Peter Forsyth's lab at the Moffitt Cancer Center and Research Institute. Begin by injecting the mouse subcutaneously with one milligram per kilogram sustained release buprenorphine. After anesthetizing the mouse, prepare it for surgery.

Clip the surgical site with enough border area to keep fur from contaminating the incision site. Shave the fur of the entire surface of the head, and prepare the skin using sterile technique. Then saturate the site with germicidal skin antiseptic, working from the center of the site to the periphery, and allow it to dry.

Position the body to ensure the spine is kept level with the cisterna magna. While applying slight traction to the tail, place tape at the tail base to secure it. Apply either a sterile drape or a sterile adhesive-backed plastic drape material to protect the surgical site from contamination.

With the neck in full extension, run the surgical scissor tips downward with slight pressure across the occipital bone, beginning just between the pinnae. Make a small three to five millimeter midline incision, slightly above the palpated concavity. Draw five microliters of the cell suspension into a 30 gauge Hamilton syringe.

Use blunt tipped forceps with a one to two millimeter tip to gently press down on the cisterna magna. Introduce the tips in a closed position, and open them while applying downward pressure on the dura. Repeat the blunt dissection until the dural membrane is easily identified and the associated blood vessels are visible in the exposed area.

While holding the forceps open to retract surrounding musculature, introduce a 30 gauge non-coring needle under the dura to visualize the bevel, making sure that the needle is only introduced just beyond the bevel itself. Slowly deploy a syringe plunger and deliver cells just below the dura. If leakage at the injection site is noted, apply gentle pressure with a cotton-tipped applicator.

Close the skin by applying a wound clip or a micro drop of skin adhesive. Monitor the mice daily after the surgery for the first week. If a mouse appears to be in pain or distress, treat it with a subcutaneous injection of 10 milligrams per kilogram carprofen once every 12 to 24 hours for up to five days based on veterinary consultation and directive.

Assemble the Murine Ommaya injection device using a 25 gauge miniature injection port and a one millimeter spacer disc. Use a cyanoacrylate sterile adhesive to ensure the penetration of approximately 2.5 millimeters of the metal cannula into the right cerebral hemisphere, and prepare the skin according to sterile technique. Make a small skin incision followed by blunt dissection of the underlying subcutaneous tissues to expose the skull.

Dry the skull using hydrogen peroxide soaked cotton-tipped applicator sticks. Drill a 0.9 millimeter burr hole in the skull 0.5 millimeters posterior and 1.1 millimeters lateral of the bregma to expose the dura matter. Move the microdrill aside and gently score the bone immediately surrounding the burr hole.

Affix an injection port to the skull using a cyanoacrylate sterile adhesive, and insert it to a depth of approximately 2.5 millimeters. Suture the incision using 4-0 non-absorbable nylon sutures in an interrupted stitch pattern or pursestring suture. House post-surgery mice in individual cages for recovery.

To dose the mouse, access the Murine Ommaya using a port injection adapter and a Hamilton syringe. Using forceps, hold the top of the miniature injection port and gently insert the port injector adapter fully into the port septum. Once the injection is made, detach the Murine Ommaya from the port injection adapter with forceps.

For the purpose of visualizing the route of the injection, 2%Evans Blue was injected via the Murine Ommaya model. The dye successfully infiltrated the ventricles and the brain in 15 minutes. After 30 minutes, the dye became visible on the spinal cord.

As a proof of concept, BALB/c mice were injected with a luciferase-labeled Her2+TUBO breast cancer cell line intracisternally, and the Murine Ommayas were implanted. Approximately one week after the injection, the mice began to develop LMD. These mice were treated once a week for up to four weeks with Her2 antibody immunotherapy, either through systemic therapy via intraperitoneal injection or intrathecally via the Murine Ommaya.

While untreated mice died by day 19, all mice that retrieved intrathecal therapy through the Murine Ommaya survived. By week four, a complete regression of tumors was observed. In comparison to mice treated with systemic therapy, mice that received intrathecal therapy had a much longer overall survival.

When attempting this procedure, it is important that the skull is dry before placing the Murine Ommaya. Use a small amount of glue and position the Murine Ommaya for about 10 seconds to adhere to the skull. With this technique, researchers can explore a wide array of cancer drugs in mouse models that are clinically relevant to design rational therapeutic strategies for patients with central nervous system metastasis.