Cisterna Magna Injection Approach for the Treatment of Leptomeningeal Metastatic Brain Tumors

Summary

This protocol presents a straightforward procedure that enables direct intrathecal delivery of agents to cerebrospinal fluid without operative damage to the skull or brain parenchyma. This approach provides a solution to deliver molecules that could not otherwise cross the blood-brain barrier or blood-cerebrospinal fluid barrier to treat leptomeningeal metastatic disease.

Abstract

Leptomeningeal metastases (LM) in the setting of pediatric brain tumors are associated with particularly poor prognosis. Current therapies for pediatric brain tumors include surgery, radiation, and chemotherapy. However, LM are not amenable to surgical resection and frequently do not respond to radiation. Moreover, most chemotherapeutic drugs are unable to cross the blood-brain barrier or blood-cerebrospinal fluid (CSF) barrier to reach LM. Here, we describe a mouse model of pediatric brain tumors that develop LM and an easy, reproducible approach to deliver therapeutic agents to treat LM. We intrathecally injected a CSF tracer via the cisterna magna in mice. After a single injection, the CSF tracer influx was observed along the leptomeninges throughout the entire central nervous system and persisted for at least 6 hours. Daily injection did not result in apparent toxicity or impact animal weight. The results of this study highlight the utility of a mouse model of metastatic pediatric brain tumors as well as the potential application of a repeated intrathecal injection technique to deliver therapeutic agents to treat metastatic brain tumors.

Introduction

Leptomeningeal metastases (LM) are common in patients with advanced brain cancer^1,2,3, breast cancer^4,5, lung cancer⁶, melanoma^7,8, and leukemia⁹. The presence of LM uniformly confers a poor prognosis and negatively impacts patient quality of life. Because meaningful surgical resection is impossible due to the diffuse nature of LM, the standard of care for leptomeningeal metastatic disease involves radiation and chemotherapy¹⁰. However, extensive irradiation of the entire central nervous system often causes severe treatment-related side effects, particularly for young patients. Moreover, many chemotherapeutic drugs cannot readily cross the blood-brain barrier and blood-cerebrospinal fluid (CSF) barrier, limiting their distribution to LM upon intraperitoneal, intravenous or oral administration¹¹. For drugs that are capable of permeating these barriers, achieving effective pharmacological concentrations through simple diffusion from the blood to the LM remains a challenge^12,13. Intrathecal injection, which allows direct drug delivery to the CSF, can circumvent these barriers and is widely used for the treatment of LM¹⁴. Intrathecal injection can be performed through either lumbar puncture or implantation of catheters/cannulas into the ventricles or cisterna magna^15,16,17,18. However, the surgery that is required to perform these procedures in preclinical models is inconvenient and may cause undue stress to animals^{16,18,19,20,21,22}. These procedures also carry the risk of post-operative injection injury^16,23. Thus, the goal of this study was to develop a non-invasive approach to repeatedly and accurately deliver agents directly to the CSF to facilitate the treatment of LM in preclinical mouse models.

Medulloblastoma, one of the most common malignant pediatric brain tumors, is a significant cause of childhood morbidity and mortality^24,25. Approximately one-third of patients with malignant medulloblastoma present with metastases and nearly all have metastatic disease at the time of relapse^26,27. Medulloblastoma metastases are found almost exclusively along the leptomeninges on the surface of the brain and spinal cord^5,28,29. Therefore, intrathecal drug delivery shows promise as an effective technique for the treatment of metastatic medulloblastoma. Here, we describe a protocol to generate a metastatic medulloblastoma mouse model as well as a simple and reliable intrathecal injection approach to deliver agents to the CSF in mice. Because surgery is not required, this approach is less invasive than other methods and allows animals to recover more rapidly.

Protocol

All animal procedures were performed according to the NIH guidelines and were approved by the Research Animal Facility Committee, Children's Research Institute (Protocol #30351).

1. Generation of a mouse model of metastatic medulloblastoma

NOTE: Because this is a recovery surgery, procedures should be performed under sterile conditions. All surgical tools should be sterilized by autoclaving or with a hot bead sterilizer.

1. Stereotactic implantation of medulloblastoma tumor cells
   1. Transfect tumor cells with mCherry and luciferase reporters. To transfect cells, culture 2 x 10⁵ patient-driven xenograft medulloblastoma cells in a 24 well plate with 360 μL of neural stem cell medium and proliferation supplement. Add 40 μL of lentivirus encoding both mCherry and luciferase in one vector to the culture medium without polybrene, and culture overnight at 37 °C.
   2. After the overnight culture, collect cells into a microcentrifuge tube by pipetting.
   3. Centrifuge cells at 220 x g for 5 min at room temperature. Remove the supernatant carefully with a micropipette and discard the supernatant into a 5% bleach solution.
   4. Wash once by resuspending the pelleted cells in 500 μL of fresh neural stem cell medium and centrifuging at 220 x g for 5 min at room temperature.
   5. Remove the supernatant and resuspend cells in 16 μL of neural stem cell medium. Keep the tube on ice for the duration of the transplantation procedure.
   6. Anesthetize an immunodeficient mouse (4-6 weeks old) by placing the animal in an isoflurane induction chamber (3% isoflurane, 97% air, 250 mL/min) until the toe pinch reflex ceases.
   7. Load the mouse in the prone position on a stereotactic frame over a heating pad connected to an anesthesia gas mask to keep the mouse under 1.5-2% isoflurane anesthesia.
   8. Disinfect the head of the mouse with 70% ethanol and apply non-antibiotic ophthalmic ointment to keep the eyes moist while the mouse is anesthetized.
   9. Using a sterile #10 scalpel, make an approximately 12 mm incision in the skin at the midline of the scalp over the position of the cerebellum. Expose the skull by pushing aside the muscle tissue over the lambdoid suture with sterile fine forceps.
   10. Determine the coordinates for injection into the cerebellum using a mouse brain atlas (1 mm lateral to the midline). Use the beveled end (sharp point) of a sterile 18 G needle to puncture the skull and create a small hole with a diameter of approximately 0.5 mm in the skull. Use a dissection microscope to visualize the injection site.
   11. Load cells (5 x 10⁴ in 4 µL of neural stem cell media) into a 5 μL microliter syringe with an unbeveled 24 G needle. Mount the syringe on a micromanipulator and introduce the needle through the hole in the skull into the brain at a depth of 2.2 mm.
   12. Inject cells using an electronic pump at a flow rate of 3 μL/min. After injection, leave the needle in place for 1 min to avoid the backflow of the cells or fluid.
   13. Withdraw the needle slowly and remove the mouse from the frame.
   14. Close the skin with tissue adhesive or sutures.
   15. Place the mouse in a new cage containing bedding. Place the cage under a heat lamp to keep the animal warm. Repeat steps 1.1.6 -1.1.15 for each mouse.
   16. Monitor the animals until they have completely recovered from anesthesia and are fully mobile (approximately 2-3 min). Continue to monitor these animals for an additional 30 min post-operatively.
   17. Administer 0.05-0.1 mg/kg buprenorphine subcutaneously as postoperative analgesia. Continue to administer buprenorphine every 12 h for 48 h after the procedure to manage any pain from the injection.
       NOTE: Guidelines for postoperative pain management vary between institutions. Please consult with the institutional guidelines to ensure proper pain management.
   18. Monitor the tumor growth once a week using a bioluminescent imaging system.
2. Irradiation of tumor-bearing mice
   1. Irradiate the tumor-bearing animals when the luciferase signals reach ~1 x 10⁶-1 x 10⁷ radiance per second.
      NOTE: The length of time for animals to develop tumors depends on the number of tumor cells, but typically animals develop tumors within 3-4 weeks.
   2. Pre-warm the irradiator following the manufacturer's instructions.
   3. Restrain the mouse in a lead shielding without anesthesia. The shielding is composed of two lead boxes with a narrow open space between, allowing radiation exposure to only the brain and spine while shielding the rest of the body.
   4. Place the restrained mouse into the irradiator.
   5. Irradiate the mouse with 2 Gy per dose at a rate of approximately 120 rads/min.
   6. After irradiation, return the mouse to its original cage.
   7. Repeat irradiation every other day for a total cumulative dose of up to 18 Gy.
   8. Image mice once per week with a bioluminescent imaging system to monitor the development of metastases. Metastases typically develop within 3-4 weeks following the first dose of radiation.
3. Identifying metastases
   1. Perfuse mice intracardially with 0.01 M phosphate buffer saline (PBS) followed by 4% paraformaldehyde (PFA) in 0.01 M PBS at a rate of 5 mL/min.
   2. Use fine forceps to remove the brain and spinal cord and place in cold PBS.
      NOTE: Alternatively, the spinal cord can be extracted using the method described by Richner et al (2017 Jove)³⁰.
   3. Use a fluorescence microscope to examine the location of mCherry⁺ tumor cells and acquire images.
   4. Fix the tissues in 4% PFA at 4 °C overnight. Then dehydrate the tissues with 25% sucrose overnight and embed in tissue embedding medium. Freeze blocks in a dry ice bath with 100% ethanol and store embedded tissue at -80 °C.
   5. Use a cryostat to cut 12 μm sections of brain and spinal cord.
   6. Stain the sections with 4′,6-diamidino-2-phenylindole (DAPI). Examine the metastatic tumor cells in each section under a fluorescence microscope and acquire images.

2. Intrathecal injection

NOTE: Because this is a recovery surgery, procedures should be performed under sterile conditions.

1. Intrathecal injection of CSF tracer
   1. Prepare a 10 µL Neuros syringe by attaching a 33 G beveled needle and stopper. Set the stopper to 3 mm and wash the needle 10 times with sterile deionized water, and then 10 times with sterile PBS.
   2. Anesthetize a mouse (4-6 weeks old) by placing the animal in an isoflurane induction chamber (2.5-3% isoflurane, 97% air, 250 mL/min) until the toe pinch reflex ceases.
   3. Load the mouse in the prone position on a stereotactic frame over a heating pad connected to an anesthesia gas mask to keep the mouse under 1.5-2% isoflurane anesthesia.
   4. Restrain the head of the mouse between the two ear bars. Use a tooth bar to align and support the front of the head.
   5. Adjust the height of the ear bars and the tooth bar to make sure the line connecting the most prominent aspects of the cranium and the spine forms an angle of 15° with the horizontal line. In this position, the cisterna magna is nearly the highest point of the mouse body.
   6. Shave the head and neck of the mouse, remove fur, disinfect the head with alcohol scrubs, and apply sterile ophthalmic ointment to keep the eyes moist during anesthesia.
   7. Prepare CSF tracer by dissolving fluorescein isothiocyanate (FITC)-dextran-500 in artificial CSF (aCSF) at a concentration of 0.25% w/v. Load 3 µL of CSF tracer into the 10 µL Neuros syringe.
   8. Place the syringe onto the syringe holder in the micromanipulator connected to the stereotactic fame at an angle of 35°.
   9. Palpate the space between the occiput and C1 vertebrae with either an index finger or a cotton-tipped swab. Use these two anatomical landmarks to determine the midline of the cisterna magna. An indentation is observed, which is used to define the puncture site. Mark the puncture site with a marker.
   10. Insert the needle into the puncture site and move the needle down slowly. The moment the needle pierces the skin should be clearly visible under the microscope. After the needle pierces the skin, continue moving down the needle for 0.2-0.6 mm. Inject 3 µL CSF tracer at a rate of 3 µL/min using an electronic infusion pump.
   11. Gently remove the needle and allow the mouse to recover in a fresh cage with new bedding under a heat lamp.
   12. At 30 min or 6 h after injection,perfuse mice intracardially with 0.01 M PBS followed by 4% PFA in 0.01 M PBS at a rate of 5 mL/min.
   13. Remove the entire brain and spinal cord and check for any evidence of brain tissue injury. Acquire images of the brain and spinal cord under a fluorescence microscope.
   14. Post-fix the brain and spinal cord in 4% PFA overnight, and cryoprotect with 25% sucrose in 0.01 M PBS at 4 °C for 24 h.
   15. Section the brain and spinal cord tissue into 12 μm thick coronal or sagittal sections using a cryostat. Stain the slides with DAPI, and examine the intracranial distribution of the CSF tracer.
   16. To determine whether mice can tolerate multiple intrathecal injections, intrathecally inject aCSF for 3 consecutive days, followed by a rest period of 4 days; then inject for another 3 consecutive days.
   17. Monitor the mice and record their weight daily.

Results

We first evaluated metastases development after irradiation in a patient-derived xenograft mouse model. Without irradiation, animals developed large tumors in the cerebellum (Figure 1 A-E). In contrast, irradiation ablated the majority of the tumor cells in the cerebellum but facilitated the development of metastases along the leptomeninges (Figure 1 F-J). We then characterized the movement of the CSF fluorescent tracer along th...

Discussion

In this report, we present a detailed protocol to generate a metastatic brain tumor mouse model and perform intrathecal injection into the cisterna magna. This injection approach provides a straightforward method to deliver labeled molecules to CSF compartments. By injecting CSF tracer, we observed a tracer influx into different regions of the brain and spinal cord (see Figure 2). Agents with differing molecular sizes may have distinct diffusion properties; molecules that are smaller in size...

Materials

Name	Company	Catalog Number	Comments
33-gauge Neuros syringe, 10uL	Hamilton Company	65460-05	10uL glass syringe with 33-gauge beveled needle and stopper
4% Paraformaldehyde solution in PBS	Alfa Aeser	J19943	4% PFA
5 µL Microliter Syringe Model 7105 KH, Knurled Hub Needle, 24 gauge, 2.75 in, point style 3	Hamilton Company	88000	Microliter syringe
aCSF	Tocris Bioscience (Fisher Scientific)	35-252-5ML	artificial cerebrospinal fluid
Bard-Parker Sterile Disposable Scalpels #10	VWR	89176-380	Scalpels
Buprenorphine HCL injectable, 0.3MG/ML, C3	Covetrus	059122
Butler Schein Paralube Non-antibiotic opthalmic ointment	Fisher Scientific	NC1865385	eye ointment
Cryostat	Leica	CM1950
DAPI, FluoroPure grade	Thermo Fisher Scientific	D21490
Deltaphase Isothermal Pad	Braintree Scientific	BP-PAD	heating pad
Digital 3-axes Manipulator Arm, left	Stoelting	51904	Manipulator
Digital Conversion for Stoelting Manipulator Arm, left	Stoelting	51914	Digital Display for manipulator
Digital Just for Mouse Stereotaxic	Stoelting	51730D	Stereotaxic frame
DPBS, no calcium, no magnesium	Thermo Fisher Scientific	14190250	Phosphate buffered saline
Dumont #5 Forceps	Fine Science Tools	11295-00	forceps
Dumont #7 Forceps	Fine Science Tools	11297-00	forceps
Ethanol	Fisher Scientific	04-355-226
Extra Narrow Scissors	Fine Science Tools	14088-10	scissors
Fisherbrand Sterile Alcohol Prep Pad	Fisher Scientific	22-363-750
Fluorescein isothiocyanate–dextran	Sigma Aldrich	FD500S	FITC dextran 500
Fluorescence microscope	Olympus	BX53
Fluorescence stereo microscope	Leica	M205 FA	Dissection microscope
Gas Anesthesia Mask for Stereotaxis	Stoelting	51609M
Germinator 500	Fisher Healthcare	NC9956482	Hot bead sterilizer
Graefe Forceps, 10.2 cm	Stoelting	52102-60P	forceps
Heat Lamp with table clamp	Braintree Scientific	HL-1 120V
Induction chamber, anesthesia accessories, small animlal, 5.25 liter	Medvet International	93805107	induction chamber
Isoflurane solution	Covetrus	029405
IVIS Lumina III In Vivo Imaging System	Perkin Elmer	CLS136334	IVIS Imager
Liquivet (Rapid) Tissue Adhesive	World Precision Instruments	504561	Tissue adhesive
Neurocult stem cell medium and proliferation supplement	Stem Cell Technologies	Cat# 05751	cell culture medium
Quintessential Stereotaxic Injector (QSI)	Stoelting	53311	electronic pump for injection
Restrainer & Split Shield	Braintree Scientific	MHS2-HE set	Restrainer
RS 2000 Biological System Irradiator	Rad Source Technologies		Irradiator
Sterile Cotton Tipped Applicators	Medline Industries	MDS202000
Sucrose	VWR International	EM-8510
Surgical Clipper, Cordless, Rechargeable w/#40 blade	Stoelting	51465	Electric shaver
Tissue-Plus OCT Compound	Fisher Scientific	23-730-571	Tissue embedding medium
Stereomicroscope	Leica	M60
XenoLight D-Luciferin Potassium Salt	Perkin Elmer	1227799	Luciferin