Translational Orthotopic Models of Glioblastoma Multiforme

Summary

Here, we describe a preclinical orthotopic mouse model for GBM, established by intracranial injection of cells derived from genetically engineered mouse model tumors. This model displays the disease hallmarks of human GBM. For translational studies, the mouse brain tumor is tracked by in vivo MRI and histopathology.

Abstract

Genetically engineered mouse (GEM) models for human glioblastoma multiforme (GBM) are critical to understanding the development and progression of brain tumors. Unlike xenograft tumors, in GEMs, tumors arise in the native microenvironment in an immunocompetent mouse. However, the use of GBM GEMs in preclinical treatment studies is challenging due to long tumor latencies, heterogeneity in neoplasm frequency, and the timing of advanced grade tumor development. Mice induced via intracranial orthotopic injection are more tractable for preclinical studies, and retain features of the GEM tumors. We generated an orthotopic brain tumor model derived from a GEM model with Rb, Kras, and p53 aberrations (TRP), which develops GBM tumors displaying linear foci of necrosis by neoplastic cells, and dense vascularization analogous to human GBM. Cells derived from GEM GBM tumors are injected intracranially into wild-type, strain-matched recipient mice and reproduce grade IV tumors, therefore bypassing the long tumor latency period in GEM mice and allowing for the creation of large and reproducible cohorts for preclinical studies. The highly proliferative, invasive, and vascular features of the TRP GEM model for GBM are recapitulated in the orthotopic tumors, and histopathology markers reflect human GBM subgroups. Tumor growth is monitored by serial MRI scans. Due to the invasive nature of the intracranial tumors in immunocompetent models, carefully following the injection procedure outlined here is essential to prevent extracranial tumor growth.

Introduction

Glioblastoma (GBM; grade IV glioma) is the most common and malignant brain tumor, and current therapies are ineffective, resulting in a median survival of 15 months¹. Reliable and accurate preclinical models that represent the complex signaling pathways involved in brain tumor growth and pathogenesis are essential to expedite the progress in evaluating new therapeutic regimens for GBM. Mouse models in which human brain tumor cell lines are implanted subcutaneously in immunocompromised mice do not reflect the native immune environment of brain tumors, nor can they be used to evaluate the ability of therapeutics to cross the blood-brain barrier². Ideally, preclinical mouse models should also reproduce closely the human GBM histopathology, including the high level of invasiveness into the surrounding parenchyma³. Although genetically engineered mouse (GEM) models develop tumors in the context of an intact immune system, complicated breeding schemes are often required, and tumors may develop slowly and inconsistently⁴. GEM-derived allograft models are better suited for preclinical therapeutic studies, where large cohorts of tumor-bearing mice are needed in a shorter time frame.

In a previous report, we described an orthotopic GBM mouse model derived directly from GEM tumors. Tumorigenesis in the GEM is initiated by genetic events in cell populations (primarily astrocytes) expressing glial fibrillary acidic protein (GFAP), that result in progression to GBM. These TRP GEMs harbor a TgGZT121 transgene (T), which expresses T121 after exposure to the GFAP-driven Cre recombinase. T121 protein expression results in the suppression of Rb (Rb1, p107, and p103) protein activity. Co-expression of a GFAP-driven Cre transgene (GFAP-CreERT2) targets expression to adult astrocytes after induction with tamoxifen. TRP mice also harbor a Cre-dependent mutant Kras (Kras^G12D; R) allele, to represent activation of the receptor tyrosine kinase pathway, and are heterozygous for the loss of Pten (P)^5,6. Concurrent gene aberrations in the receptor tyrosine kinase (RTK), PI3K, and RB networks are implicated in 74% of GBM pathogenesis⁷. Therefore, the primary signaling pathways altered in human GBM are represented by the engineered mutations in TRP mice, in particular GBM tumors, in which shared downstream targets of RTKs are activated⁵.

The GEM-derived syngeneic orthotopic model was validated as a model that recapitulates features of human brain tumors, including invasiveness and the presence of subtype biomarkers, for use as a platform to evaluate cancer therapeutics targeting aberrant pathways in GBM. Cells were cultured from tumors harvested from TRP brains and re-implanted in the brain of strain-matched mice, using stereotactic equipment for intracranial injection in the cortex. This preclinical orthotopic mouse model developed GBM tumors that were highly cellular, invasive, pleomorphic with a high mitotic rate, and displayed linear foci of necrosis by neoplastic cells and dense vascularization, as observed for human GBM. Tumor volumes and growth were measured by in vivo magnetic resonance imaging (MRI).

In this report, we describe the optimal technique for the intracranial injection of primary GBM cells or cell lines into the wild-type mouse brain, using TRP tumors as an example. The same protocol may be adapted for immunocompromised mice and other GBM cell lines. Crucial tips are given for avoiding common pitfalls, such as suboptimal cell preparation or cell leakage at the injection site, and for using the stereotactic equipment correctly to ensure model reproducibility and reliability. For translational purposes, we validate the model by MRI detection of brain tumor growth in live animals, histological characterization, and present an example of treatment in tumor-bearing mice.

Protocol

The study protocol described here was approved by the NCI at Frederick Animal Care and Use Committee. NCI-Frederick is accredited by AAALAC International and follows the Public Health Service Policy for the Care and Use of Laboratory Animals. Animal care was provided in accordance with the procedures outlined in the “Guide for Care and Use of Laboratory Animals (National Research Council, 2011; The National Academies Press, Washington D.C.).

1. Preparation of cells for injection

NOTE: Mouse brain tumor primary cells (MBRs) used for this model were originally isolated from tamoxifen induced TRP GEM mice, as described in El Meskini et al.⁵. Details on the cell preparation can be found in this reference.

1. Perform the following steps using sterile technique in a biosafety cabinet.
2. Culture primary brain tumor cells in vitro until they reach the exponential growth phase.
3. Harvest the cells in 0.25% trypsin, and once they have detached, dilute them with growth medium to inactivate the trypsin.
4. Pellet the cells by centrifugation at 400 x g and wash with serum-free media. Repeat once prior to counting.
5. Count the cells manually or with an automated cell counter. Resuspend the cells in 5% sterile methylcellulose in 1x phosphate buffered saline (PBS) at the desired concentration, based on a final injection volume of 2 µL. The desired cell concentration may vary based on the cell line and brain tumor model. For GBM GEM TRP cells, we inject 2 µL of a 25 x 10⁶ cells/ml solution, or 50,000 cells.

2. Mouse strain

1. Breed or purchase an appropriate mouse strain that matches the strain background of the brain tumor cells. For TRP cells, use 9-week-old B6D2F1/J mice (from a cross between C57BL/6J [B6] females and DBA/2J [D2] males) as tumor cell recipients.

3. Setting up the surgical area

NOTE: All surgical steps are conducted using aseptic technique in a clean, sanitized environment. Scrubs and personal protective equipment, including a mask, should be worn by the surgeon. Surgical tools must be heat-sterilized prior to use.

1. Place surface protectors under the stereotaxic apparatus and on the work surface.
2. Attach the vinyl tubing from the anesthesia machine to the IN port of the gas anesthesia platform of the stereotaxic apparatus, and another tube to the OUT port.
3. Connect the digital display to a power source and to the apparatus.
4. Connect the micropump controller (Figure 1A) to a power source and attach the micropump (Figure 1A) to the manipulator arm of the apparatus (also see the manufacturer instructions).
5. Set the micropump to 5.6 nL/s for the 2,000 nL volume injection.
6. Turn on the hot bead sterilizer.
7. Plug the mouse heating pad into the temperature controller and connect to a power source. Place the plate on the stage platform. Set the plate temperature to 37 °C.
8. Backload a 30 G precision syringe, being careful not to introduce bubbles. Insert the plunger and attach the needle. Make sure to grip the needle by the hub only. Depress the plunger until a drop of the methylcellulose cell mixture dispenses through the needle. Clean the needle with a 70% alcohol preparation pad by carefully wiping the sides of the needle, or by placing a sterile gauze pad on the work surface and blotting it. Take care not to bend or break the needle.
9. Attach the precision syringe to the micropump and rotate manipulator arm away from the stage, to prevent syringe-needle displacement prior to placing the mouse on the stage.

4. Preparing the mouse for surgery

1. Anesthetize the mouse by placing it in the induction chamber with the isoflurane vaporizer set to 2.5%, or according to institutional guidelines. Turn on the isoflurane flow to the nosecone.
2. Transfer the mouse to the stereotaxic apparatus and secure to the nosecone, with the top teeth positioned onto the nosecone support (Figure 1B, 9). Tighten the knob on the nosecone to secure (Figure 1B). Ensure an appropriate level of anesthesia by performing a toe pinch to check for reflex.
   1. Monitor the ears, feet, and mucous membranes for color (pink) to ensure adequate oxygenation. Also, monitor the respiratory rate (an increase or decrease could indicate the need to adjust isoflurane levels).
3. Insert ear bars (Figure 1B) into both ears and tighten the knob to secure the head.
4. Apply eye ointment to lubricate the eyes while the mouse is under anesthesia.
5. Use curved forceps to pluck the hair on the mouse's head to an area about 150% larger than the planned incision, to ensure adequate aseptic conditions.
6. Inject 0.5-1 mg/kg buprenorphine SR analgesia subcutaneously, or use another protocol-approved analgesic.
7. Position the mouse rectal probe to monitor internal temperature and prevent hypothermia due to anesthesia. Keep the mouse body temperature between 36.5 and 38.5 °C.
8. Sanitize the surgical field using outward circles, alternating between a surgical scrub and ethanol three times.
9. Using forceps to pull the skin taut, make an incision of approximately 1 cm with the scalpel blade, starting between the eyes. The bregma should be visible through the incision.
   NOTE: For proper sterile technique, touch the surgical site only with the tips of sterilized tools, and place tool tips down on a sterile surface only (such as the inside of an autoclaved tool pack).
10. Use the wooden end of cotton-tipped applicator to scrape away excess connective tissue, and then the cotton end of another applicator to dry.

5. Cell injection

1. Return the manipulator arm with the syringe attachment over the mouse, tightening the knob to secure. Use the X and Y knobs in the horizontal plane to move the syringe mount over bregma. Lower the needle using the Z knob to confirm the bregma position. Set the digital readout console to zero.
2. Use the X and Y knobs and the corresponding digital readout to move the needle to the desired position. For the cortical cortex location, the appropriate coordinates are 3 mm posterior, 2 mm lateral right to the bregma, and 2 mm deep from the dura mater. Use the Z knob to move the needle to surface of skull.
3. Puncture a hole in the skull using a 1 mL syringe with an attached 25 G needle. Place the bevel of the needle toward the 30 G precision syringe and needle, and carefully rotate the manipulator arm to the side. Using thumb and finger, roll the needle back and forth slowly with gentle pressure until the needle tip just pierces the skull cap.
4. Use a cotton tipped applicator to dab any blood away from the needle hole. Replace the manipulator arm to the appropriate position with the loaded precision syringe needle above the hole. Align the tip of the needle with the hole, using the Z knob to lower the needle down to the brain dura, and set the digital readout console to zero.
5. Using the Z knob, lower the needle 1 mm, and then wait 1 min. Repeat until the desired depth is reached (2 mm as indicated).
   NOTE: The needle is lowered slowly to prevent additional damage to the surrounding brain tissue and back-flow of the cell solution.
6. Start the micropump, and then monitor to ensure that the pump stops when 2 µL has been injected. This process should take about 6 min at the indicated speed. Then, wait 1 min prior to moving the needle.

6. Removal of the needle and wound closure

1. Raise the needle 1 mm and then wait 1 min. Repeat until the needle is completely free from the skull.
2. If necessary, use a cotton-tipped applicator to dab any blood away from the injection site.
3. Using the wooden end of a cotton-tipped applicator, take a small piece of bone wax (~1 mm) and shape it into a cone. Place it into the opening in the skull, pushing the wax into the hole.
4. Heat forceps using a bead sterilizer and use them to melt remaining wax on the skull and smooth it.
5. Place roughly two drops of bupivacaine (anesthetic) solution into the incision and use forceps to pull the edges of skin together.
6. Pull the skin taut and place one or two wound clips to close the skin.
7. Place the mouse in a clean recovery cage on a heating pad in a draft-free area, and closely observe the mouse. Allow the mouse to wake up fully from the anesthesia, resuming normal activity, before returning it to regular housing.
8. Check on the mouse daily following the surgical procedure and administer pain management, according to institutional guidelines.
   1. If buprenorphine SR (Step 4.6) is used for analgesia, the sustained-release formula lasts for 72 h. Repeat the injection only if visible pain or discomfort is present after 72 h. When performed correctly, the mice recover well from the intracranial injections and additional analgesia is not needed.
   2. Remove the staples 7-10 days post-surgery.
9. Monitor tumor growth by live animal imaging (MRI).
   1. Euthanize the mice when humane endpoints are reached (i.e., if the animal loses 20% body weight or becomes hypothermic).
   2. Due to the invasive nature of these brain tumors, observe the mice for neurological symptoms, such as uneven gait, partial paralysis, spinning, or head tilt. Euthanize a mouse if any of these clinical signs of advanced tumor growth are observed.

Results

Mice injected with brain tumor cells should be monitored daily for signs of tumor growth such as seizures, ataxia, or weight loss. Brain tumor growth may also be monitored by MRI scanning at regular intervals. Weekly MRI scans allow the visualization of increasing tumor burden within the brain and tumor volume measurements (Figure 1C). In particular, TRP tumors exhibit aggressive growth, and 3D tumor volumes are measurable by MRI within 2 to 3 weeks post-intracranial injection (with an avera...

Discussion

Preclinical models are essential for the evaluation of new therapeutic targets and novel treatment strategies in GBM. Genetically engineered mouse models for GBM have the advantage of tumor occurrence in the autochthonous site, but often with a long latency and unpredictable tumor growth¹³. The GEM model tumors exhibit a latency of 4-5 months, and the ideal time window for imaging, recruitment, and treatment is variable among individual mice. The orthotopic model has a well-established and tractab...

Materials

Name	Company	Catalog Number	Comments
5% methylcellulose in 1X PBS, autoclaved	Millipore Sigma	M7027
1mL Tuberculin Syringe, slip tip	BD	309659
6" Cotton Tipped Applicators	Puritan	S-18991
Adjustable stage platform	David Kopf Instruments	Model 901
Aerosol Barrier Tips	Fisher Scientific	02-707-33
Alcohol Prep Pads Sterile, Large - 2.5 x 3 Inch	PDI	C69900
B6D2  mouse strain (C57Bl/6J x DBA/2J)	Jackson Laboratory	Jax #10006
Bone Wax	Surgical Specialties	901
Bupivacaine 0.25%	Henry Schein	6023287
BuprenorphineSR	ZooPharm	n/a
Clear Vinyl Tubing 1/8ID X 3/16OD	UDP	T10004001
CVS Lubricant Eye Ointment	CVS Pharmacy	247881
Disposable Scalpels, #10 blade	Scalpel Miltex	16-63810
Gas anesthesia machine with oxygen hook-up and anesthesia box	Somni Scientific	n/a	Investigator may use facility standard equipment
Gas anesthesia platform for mice	David Kopf Instruments	Model 923-B
GraphPad Prism	Graphpad	Prism      9      version 9.4.1
Hamilton 30 g needle, ½ “, small hub, point pst 3	Hamilton	Special Order
Hamilton precision microliter syringe, 1701 RN, no needle 10 µL	Hamilton	7653-01
Hot bead sterilizer with beads	Fine Science Tools	18000-45
Invitrogen Countess 3 Automated Cell Counter	Fisher Scientific	AMQAX2000
IsoFlurane	Piramal Critical Care	29404
Isopropyl Alcohol Prep Pads	PDI	C69900
ITK_SNAP (Version 36.X, 2011-present)	Penn Image Computing and Science Laboratory (PICSL) at the University of Pennsylvania, and the Scientific Computing and Imaging Institute (SCI) at the University of Utah
KOPF Small Animal Stereotaxic Instrument with digital readout console	David Kopf Instruments	Model 940
Masterflex Fitting, PVDF, Straight, Hose Barb Reducer, 1/4" ID x 1/8" ID	Masterflex	HV-30616-16
Mouse Heating Plate	David Kopf Instruments	PH HP-4M
Mouse Rectal Probe	David Kopf Instruments	PH RET-3-ISO
Nalgene Super Versi-Dry Surface Protectors	ThermoFisher Scientific	74000-00
P20 pipette	Gilson	F123600
Povidone Iodine Surgical Scrub	Dynarex	1415
Reflex 9 mm Wound Clip Applicator	Fine Science Tools	12031-09
Reflex 9 mm Wound Clip Remover	Fine Science Tools	12033-00
Reflex 9 mm Wound Clips	Fine Science Tools	12032-09
Semken forceps, curved	Fine Science Tools	11009-13
Temperature Controller	David Kopf Instruments	PH TCAT-2LV
Trypsin-EDTA (0.25%)	ThermoFisher Scientific	25200056
Tuberculin Syringe with 25g needle, slip tip	BD	309626
UltraMicroPump 3 with Micro2T Controller	World Precision Instruments	Model UMP3T