In Vivo Glioblastoma Tumor Modeling via Stereotaxic Injection in Mice for Tumor Progression Studies

Summary

Glioblastoma research involves challenges in the understanding of its progression and assessment of the tumor microenvironment. This protocol uses stereotaxic injection of brain cancer cells in mice to establish precise in vivo modeling of brain cancer, enabling detailed studies of tumor biology and therapeutic responses.

Abstract

Glioblastoma (GB) is a highly aggressive brain cancer with a median survival of under 2 years despite conventional therapy. Research efforts frequently use murine in vivo models to study cancer progression and tumor growth, as these models recapitulate critical aspects of the tumor microenvironment, immune responses, and the appearance of classical GB pathology. Here, we describe a detailed protocol for performing stereotaxic injections to establish GB tumor models in immunocompetent C57BL/6 mice. Stereotaxic surgery enables rapid and precise delivery of GB cells grown as 2D cultures or spheroids into the brain using anatomical landmarks without the need for image guidance. Specifically, GL261 murine cell lines are injected into the striatum, allowing subsequent assessment of behavior and physiology. On day 21 post surgery, animals are euthanized following perfusion and their brains extracted and fixed. Hematoxylin-eosin staining on brain slices enables measurement of tumor area and volume, while immunohistochemistry enables imaging of tumor proliferation markers. This paper provides detailed protocols for a reliable approach to generate in vivo GB models for studying tumor growth and cancer progression. Additionally, this method is adaptable for other GB cell lines, transgenic mice, or patient-derived xenografts broadening its applicability for GB research.

Introduction

GB is a highly aggressive brain tumor and the most frequent malignant neoplasm of the central nervous system, distinguished by its rapid proliferation and deep infiltration into adjacent brain regions¹. Despite the implementation of standard therapeutic approaches, including surgery, radiotherapy, and chemotherapy, the prognosis for GB remains dismal². Patients have a median survival of less than 2 years³, with limited advancements in treatments or therapeutic options over the past decades⁴. This highlights the need for a more comprehensive understanding of the molecular and cellular mechanisms driving GB progression to identify effective treatments to improve patient outcomes.

In vitro studies are used for investigating tumor biology, but they are limited and fail to recapitulate many aspects of GB tumor characteristics, such as interactions with the tumor microenvironment and the immune response. Therefore, the use of preclinical models that accurately replicate the complexity of GB is needed for translational research. The use of C57BL/6 mice has become a cornerstone as a reliable and reproducible in vivo model for GB research, providing clear advantages compared to in vitro approaches for studying tumor biology and testing therapies⁵.

Early methods for inducing brain tumor formation in mice, such as chemical carcinogens or viral agents, were limited since they lack specificity and can lead to off-target effects, thus limiting their use in targeted studies^5,6. Genetically engineered models (GEMs) have significantly advanced the field by replicating mutations frequently present in human GB or expression of specific oncogenes⁷ allowing more specific studies of GB biology and therapeutic development⁸. Nevertheless, GEMS strongly rely on the ability of specific oncogenes to drive cell transformation, have low penetrance and high variability, and require access to genetically modified animals that include logistical and financial challenges^5,8.

Grafted/injected models offer a feasible, accessible and versatile method for generating GB tumor in vivo⁹. These models involve the administration of cultured tumorigenic cells, derived from immortalized cell lines or patient-derived tumors, directly into the mouse brains using stereotaxis guidance¹⁰. This approach allows researchers to increase the range of experimental possibilities by using cell lines or mouse models genetically modified to express oncogenes, mutations, or gain-or loss-of function manipulations. This ability to manipulate the experimental setup before or during tumor implantation makes grafted/injected models a valuable tool in GB research¹⁰.

Here, we describe an effective method for generating an in vivo tumor model using stereotaxic GL261 murine glioma cells into immunocompetent C57BL/6 mice. This approach provides an isogenic model that preserves immune system function, allowing the study of tumor-immune interactions with the tumor microenvironment, which is a key factor determining tumor growth and is a current target for GB therapies¹¹. The advantage of stereotaxic surgery lies in visible anatomical landmarks to precisely target the brain striatum for tumor cell injection, ensuring controlled tumor formation and localization and reducing variability between animals^10,11. Additionally, this method can be adapted to implant tumors in different brain regions depending on the experimental goals, such as implantation of pediatric GB cells into the hippocampus to study electrical activity, among other effects^12,13. Animals are closely monitored throughout the experimental timeline for postoperative care and manifestations of tumor implantation in the brain. This protocol not only facilitates the study of GB progression but also enables a wide range of downstream analyses. Simple hematoxylin-eosin staining (H&E) of brain sections is useful for quantification of tumor size¹⁴. Tumor proliferation markers, such as Ki67, can be assessed by immunohistochemistry (IHC) staining¹⁵ or by performing incorporation of thymidine analogs followed by chemical detection of DNA synthesis, which offers valuable insights into tumor development.

We used the GL261 cell line, a well-established model with a wide range of applications, to study GB progression in vivo¹⁶. GL261 are murine GB cells that harbor mutations in Kras and p53 and express known GB cancer drivers, as evidenced by genetic characterization¹⁷. Additionally, this model is reproducible and generates aggressive and infiltrative tumors in C57BL/6 mice while maintaining an intact immune microenvironment, with validation in preclinical studies involving radiotherapy, chemotherapy, and immunotherapy⁷. Although GL261 does not capture the full genetic heterogeneity of human GB, it remains a useful model for studying GB biology and avoids the use of immunocompromised mice. Overall, this method provides a reliable and reproducible in vivo GB model, enabling detailed studies of tumor biology and therapeutic development.

Protocol

All animal experiments were carried out according to the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The animal ethics protocol for the use of animals in this study was approved by the Committee on the Ethics of Animal Experiments of the University of Chile. This protocol used 8-12-week-old C57BL/6 immunocompetent mice housed at 23 ± 2 °C with a humidity of 35 ± 5% under a 12:12 h light-dark cycle with access to food and water ad libitum. This section outlines the protocol for establishing the in vivo GB model. Figure 1 provides an overview of the experimental workflow, including the estimated time required for each step.

1. Culture and preparation of GB cells for stereotaxis injection

1. Culture and preparation of GL261 cells in 2D
   1. Thaw GL261 cells in 5 mL of warm complete medium consisting of DMEM (10% fetal bovine serum, 1x non-essential amino acids, 2 mM L-Glutamine, and 2x Penicillin-Streptomycin) at 37 °C and 5% CO₂ in a humidified incubator.
   2. Centrifuge the cell suspension at 200 × g for 5 min and discard the supernatant.
   3. Wash the cells with PBS 2x, repeating steps 1.1.2 and 1.1.3.
   4. Resuspend the pellet of cells in 10 mL of complete medium and transfer to a 100 mm cell culture plate. Grow GL261 cells to 80% confluency.
      NOTE: Ensure that tumor cells do not exceed 80% confluency to maintain optimal viability and experimental consistency. Tumor cells in this example are GL261 cells, but other cell lines can be used based on specific research objectives and availability. Cells can be used for implantation after two passages. Use cells with up to a maximum of 8-10 passages post thawing to avoid potential alterations in proliferation, tumorigenicity, or genetic drift associated with prolonged culture.
   5. Wash 2x with PBS and add 1 mL of trypsin/EDTA. Incubate for 5 min at 37 °C.
   6. Resuspend in 5 mL of complete medium and disaggregate the cells.
   7. Transfer to a 15-mL tube and centrifuge at 200 × g for 3 min.
   8. Resuspend the cells by adding 5 mL of warm PBS, and centrifuge again at 200 × g for 3 min.
   9. Repeat step 1.1.8 twice to eliminate all traces of serum and dead cells.
   10. Resuspend the cells in 200 µL of warm PBS. Count the cells using automatic or manual counting using the hemocytometer.
       NOTE: Cell counting should be performed at least 6x to ensure maximum accuracy and maintain a variation coefficient below 5%.
   11. Prepare a cell suspension of 5,000 cells/µL in PBS (5 × 10⁶ cells/mL). Transport and keep the cell suspension on ice. Mix the cell suspension constantly to avoid aggregation of cells.
       NOTE: Cell viability is not affected by at least 2 h on ice, which is sufficient time to inject 6-8 animals by two independent operators. It is recommended to prepare a fresh cell suspension every 2 h to minimize variability between experiments.
2. Culture and preparation of GL261 cells in 3D neurospheres
   1. Start with a 10 cm plate of GL261 cells at 80% confluence. Wash the plate with 5 mL of PBS.
   2. Add 1 mL of trypsin-EDTA. Incubate for 5 min at 37 °C.
   3. Once the cells are detached, add 10 mL of serum-free DMEM F12 to the plate. Transfer the suspension to a 15 mL tube and centrifuge at 300 × g for 5 min.
   4. Discard the supernatant. Resuspend the cells in 10 mL of DMEM F12. Repeat centrifugation and resuspension of the cells.
   5. Prepare a cell suspension at a density of 50,000 cells/mL in neurosphere medium (DMEM F12 supplemented with 1x B27, 1x N2, 20 µg/mL epidermal growth factor (EGF), and 20 µg/mL fibroblast growth factor (FGF)).
   6. Seed 8 mL into a 50 mL flask. Supplement the medium every 3 days with 10% of the initial volume using 2x neurosphere medium.
   7. After 7 days of neurosphere formation, transfer the neurospheres to a 15 mL tube and centrifuge at 100 × g for 5 min. Discard the supernatant.
   8. Add 2x trypsin-EDTA and incubate for 5 min at 37 °C. Add 5 mL of neurosphere medium to suspend the cells, transfer to a 15 mL tube, and centrifuge at 200 × g for 3 min.
   9. Resuspend the cells in 5 mL of warm PBS and centrifuge again at 200 × g for 3 min.
   10. Repeat the centrifugation and resuspension 2x to eliminate all traces of medium and dead cells.
   11. Resuspend the cells in 200 µL of warm PBS. Count the cells using automatic or manual counting with a hemocytometer.
       NOTE: Cell counting should be performed at least 6x to ensure maximum accuracy and maintain a coefficient of variation below 5%.
   12. Prepare a cell suspension at a concentration of 5,000 cells/µL in PBS (5 × 10⁶ cells/mL). Transport and keep the cell suspension on ice.

2. Stereotaxis injection and implantation of GB cells

1. Administer the analgesic solution to each animal prior to starting the stereotaxic procedure.
   1. Prepare Ketoprofen analgesic solution by diluting 100 µL of 1% ketoprofen (10 mg/mL) with 900 µL of sterile saline, protecting the solution from light.
      NOTE: Alternative analgesics may be employed to minimize any potential influence on tumor growth, depending on specific experimental requirements.
   2. Inject subcutaneously the required volume of 0.1% Ketoprofen at a dose of 5 mg/kg using a 1 mL syringe with a 27 G needle.
2. Anesthesia and positioning in the stereotaxic apparatus
   1. Weigh the animal before surgery to monitor postsurgery variations.
   2. Place the animal in the isoflurane induction chamber (5% of isoflurane with 20% of O₂) for 3 min.
   3. While the animal is in the chamber, heat the stereotaxic apparatus with a heating blanket during the entire surgery procedure to avoid a decrease in animal temperature
   4. Confirm anesthesia by assessing the response of animals to nociceptive stimuli by gently pinching the plantar surface of the paw and observing the whisker reflex.
   5. Secure the mouse by positioning the ear bars at the bone above the ear canal and fix and stabilize the animal in the stereotaxic apparatus. Once the animal is fixed in the stereotaxic frame, decrease the isoflurane concentration to 2-3%.
      NOTE: It is important to secure the animal properly to prevent any damage to the ears during the surgery and to ensure that the head is positioned parallel to the stereotaxic frame for accuracy during the procedure.
3. Craniotomy and injection of GL261 cells
   1. Disinfect the incision area using a swab soaked in 10% hydrogen peroxide.
   2. Make a 0.6 mm incision on the dorsal part of the head of the mouse with a universal no. 15 scalpel to expose the skull.
   3. Gently remove the periosteum using the scalpel and surgical forceps.
   4. Locate bregma on the skull where the coronal suture intersects with the sagittal suture.
   5. From bregma, drill a 2 mm diameter hole at the coordinates: anteroposterior (AP) 0 mm; lateral (LL) -2.2 mm, using a veterinary-use Dremel mounted on the stereotaxic apparatus, based on the Paxinos Mouse Brain Atlas.
   6. Fill and install a 33 G Hamilton syringe in the stereotaxic frame. Position the tip of the syringe at the entrance of the hole in the skull.
      NOTE: Prior to loading the syringe, the cell suspension needs to be gently vortexed to homogenize the cells and prevent cell aggregation and clumps.
   7. Carefully lower the syringe to the dorsoventral (DV) coordinate of 3 mm at a constant rate of 1 mm/min.
   8. Inject a total volume of 2 µL of cells (1 × 10⁴ cells) at a rate of 1 µL/min. After the injection, wait for 2 min before proceeding. Carefully withdraw the needle at a rate of 1 mm/min.
   9. Suture the incision with three simple stitches along its entire length using sterile, braided, and silicon-coated black silk.
   10. Apply antibacterial-antifungal-antiinflammatory ointment in the wounded area.
   11. Remove the rodent from the stereotaxic apparatus and place it in a temperature-controlled observation box until full recovery from anesthesia is confirmed.

3. Postoperative care and animal monitoring

NOTE: The experimental timeline involves monitoring animals post surgery for 21 days and is divided into three different periods for supervising: post surgery care and monitoring phases. Nociceptive signs and symptoms will be evaluated using the modified Morton and Griffiths scale, as outlined in the publication "Guidelines on the Recognition of Pain, Distress, and Discomfort in Experimental Animals and a Hypothesis for Assessment"^17,18. This scale will be employed to systematically quantify pain and distress throughout the study. The progression of tumor formation in C57BL/6 mice involves tumor growth starting from day 7, which persists throughout the experiment.

1. Postsurgery care-Days 1-7
   1. Conduct daily monitoring of the animals. For the first 3 days, administer oral analgesia (ketoprofen at 5 mg/kg every 24 h).
   2. Evaluate the following parameters: appearance, weight, and the surgical site. Additionally, evaluate pain management using the Grimace Scale, focusing on orbital opening and nose bulging.
      NOTE: Humane endpoint criteria include the observation of signs associated with infection or local inflammation, such as redness in the manipulated area, the presence of tremors, or weight loss exceeding 20% of the animal's body weight during the experiment¹⁷.
2. Monitoring of Phases 2 and 3
   NOTE: The nociceptive monitoring system evaluates spontaneous behavior, appearance, weight, and motor impairments in mice, assigning scores from 0 to 3 per category.
   1. Between days 8 and 14 post surgery, conduct animal monitoring 3x during the week.
   2. During days 15 to 21, conduct daily monitoring of the animals.
      NOTE: The tumor growth curve shows that between days 17 and 21, the tumor volume begins to affect the motor functions unilaterally of mice. Until that point, the mice do not display any perceptible organ impairment or vital signs compromise.
   3. Score animals according to the scoring system based on spontaneous behavior, appearance, weight, and motor impairments in mice, assigning scores from 0 to 3 per category.
      1. Assess spontaneous behavior as normal (0), minor changes (1), inactivity (2), or extreme restlessness/immobility (3).
      2. Score appearance based on fur condition and distress signs: normal (0), ruffled fur (1), ocular/nasal secretions (2), or abnormal posture (3).
      3. Classify weight loss as absent (0), <10% (1), 10-20% (2), or >20% (3).
      4. Assess motor impairments as ranging from normal mobility (0) to slight difficulty (1), limb lethargy (2), or complete inactivity (3).
         NOTE: A total score out of 12 points guides interventions, ensuring standardized evaluation of animal well-being and experimental consistency. A score of 0-4 indicates a normal state, requiring no intervention. Scores between 5 and 8 suggest moderate distress, necessitating careful monitoring and the administration of analgesics. If the score reaches 9-12, the animal is in severe distress, requiring immediate euthanasia to prevent unnecessary suffering.
         ​NOTE: Humane endpoint criteria include the interruption of the experiment if any mouse exhibits motor impairments in its limbs earlier than expected for this model, or if there is weight loss exceeding 20% of their body weight. After 21 days, the tumor volume reaches the maximum ethical limit in WT animals, at which point intracranial pressure increases, leading to physiological changes associated with discomfort and pain in the animal.
3. Injection of 5-ethynyl-2'-deoxyuridine (EdU)
   1. Dilute the EdU compound in sterile saline and load into a 1 mL syringe with a 27 G needle.
   2. Restrain the animal by holding the mouse with the abdomen exposed, ensuring gentle but firm restraint.
   3. Insert the needle at a 30° angle into the lower right abdominal quadrant, avoiding midline structures. Slowly perform intraperitoneal injection of the EdU solution at 10 µg/g observing for resistance or leakage.
   4. Withdraw the needle and wait 2 h before proceeding with the experimental procedures.
      NOTE: If performing proliferation analysis using EdU Labeling, EdU must be injected at the end of the experiment for a short period of time (2 h). Timing and concentration of EdU injections are optimized for the GL261 model and must be optimized for other models.
4. Euthanasia and tissue collection
   1. Once the experimental timeline is completed, euthanize the mice using CO₂ inhalation followed by exsanguination by aortic perfusion.
      1. Perform euthanasia using a chamber connected to a CO₂ tank. First, clean the CO₂ chamber with 70% ethanol. Then, open the valve to allow CO₂ into the chamber and fill it completely or to about 70%, keeping the valve open for 1 min.
      2. Place the mouse in the chamber and observe its vital signs for 3 min. When the animal is unconscious (absence of paw reflex, tail reflex, and respiratory rate), proceed with aortic perfusion.
         NOTE: Isoflurane can be used to anesthetize the mice prior to the introduction of CO₂ into the euthanasia chamber. This ensures refined euthanasia and minimizes mice suffering.
      3. Once the mouse is anesthetized, expose the thoracic cavity by grasping the chest skin with forceps to make an incision. Cut laterally beneath the ribcage to expose the diaphragm and liver, then make two lateral cuts along the rib cage up to the clavicle. Lift or remove the sternum to fully expose the heart and lungs.
      4. Insert a blunt needle through the left ventricle at an angle, ensuring it enters the ascending aorta smoothly. Secure it with a vascular clamp or hemostatic forceps.
      5. Partially sever the right atrium to allow for the outflow of blood. Perfuse with saline solution using constant and low flow. Continue until the outflow is clear and the liver is pale (~20 mL for adult mice).
      6. Perfuse with fixation buffer (4% paraformaldehyde in phosphate buffer) using 15-20 mL.
      7. Once the animal exhibits rigid tissues such as body stiffening and mild tail flicking, remove the needle.
   2. Decapitate the animal, expose the skull with a midline incision, and trim the occipital bones. Carefully lift the skull cap and remove the brain from the cranial cavity.
   3. Postfix for 24 h using 4% paraformaldehyde at neutral pH.
   4. Transfer the tissue to a 30% sucrose solution for dehydration for at least 48 h.

4. Tissue processing and staining

1. Tissue brain section
   1. Remove the region starting 3 mm interaural of the brain, as the striatum lies between the interaural regions of 4 mm and 6 mm.
      NOTE: Removing this region also serves as support for proper tissue orientation during sectioning.
   2. Place O.C.T. compound on the cryostat tissue holder, which should already be cooled to -20 °C, and position the brain with the olfactory bulb facing upwards.
   3. Let the O.C.T. freeze completely on the holder. Once frozen, cover the brain completely with O.C.T. and allow the brain to freeze thoroughly before proceeding.
   4. Mount the holder containing brains onto the cryostat arm and adjust the position to begin sectioning.
   5. Make the initial sections until reaching the striatum region and prepare 25 µm serial sections of brain tissue.
   6. Collect the serial sections in a 24-well plate containing PBS and 0.02% sodium azide, starting from the top left corner. Continue sectioning until the tumor is completely sectioned or go beyond the hippocampus.
   7. Store the collected plates at 4 °C to preserve the integrity of the samples.
2. Hematoxylin-eosin (H&E) staining
   ​NOTE: H&E staining provides a quick alternative for assessing tumor volume and area. This staining can be completed within a day and evaluated the following day using standard microscopy techniques.
   1. Mount the brain sections onto microscope slides previously silanized with 10% cold fish skin gelatin. Once mounted, dry the sections at a temperature between 60 °C and 90 °C for 1 h and re-hydrate the brain sections in distilled water for 1 min.
   2. Stain the sections with Gill hematoxylin or Harris hematoxylin for 15 min.
   3. Differentiate the sections with acid alcohol (1 dip or 1 s) and perform bluing of slides with either warm tap water or a 2% sodium tetraborate solution for 5 min.
   4. Stain the sections with eosin for 2 min.
   5. Wash sections for 2 x 2 min with 90% ethanol.
   6. Wash the sections 2x with 100% ethanol for 2 min.
   7. Wash the sections for 2 x 2 min with 100% xylene and then repeat once more for an additional 5 min.
   8. Mount the sections with mounting medium and add coverslips.
      NOTE: All procedures can be performed at room temperature without affecting the H&E stain.
3. Ki67 staining by immunohistochemistry
   ​NOTE: Ki67 staining through immunohistochemistry provides a quick alternative for assessing tumor proliferation. Ki67 is a nuclear protein expressed during the active phases of the cell cycle involving G1, S, G2, and M; therefore, it provides a general proliferation index of the proportion of cells actively cycling. This staining can be completed within 2 days and evaluated the following day using standard microscopy techniques.
   1. Wash the brain sections with PBS for 5 min and permeabilize the sections with 0.5% Triton X-100 prepared in 1x PBS for 15 min.
   2. Wash the sections for 3 x 3 min with PBS
   3. Block peroxidase with 0.3% hydrogen peroxide in PBS for 30 min at room temperature and wash as in step 3.3.2.
   4. Incubate the sections with blocking solution (5% bovine serum albumin and 0.5% Triton in PBS) for 2 h and wash (as in step 3.3.2).
   5. Incubate the sections with Ki67 primary antibody (1:500 in blocking solution); overnight at 4 °C.
   6. Wash the sections for 3 x 3 min with PBS.
   7. Incubate the sections with secondary antibody (1:1,000 in PBS) for 2 h or overnight at 4 °C.
   8. Wash the sections for 3 x 3 min with PBS
   9. Prepare the ABC kit by mixing reagent A (Avidin) and reagent B (Biotin) in a 1:1 ratio depending on the required volume. Dilute the mixed solution to 1:1,000 by adding 1 part of the A-B mixture to 1,000 parts of PBS. Incubate the sections with the prepared ABC solution for 1 h.
   10. Incubate the sections with 3,3'-diaminobenzidine (DAB) for 5 min or until the DAB precipitate is visible in the tumor area.
   11. Mount the sections onto microscope slides previously silanized with 10% cold fish skin gelatin. Dry the mounted sections at 60-90 °C for 1 h.
   12. Hydrate the sections in distilled water for 2 min.
   13. Stain the sections with Mayer's hematoxylin for 2 min.
   14. Blue the hematoxylin by rinsing the sections with running tap water or by using a 2% sodium tetraborate solution for 5 min.
   15. Allow the sections to dry either at room temperature or in an oven at 60-90 °C. Mount the sections on a glass slide using rapid, anhydrous mounting medium.
4. EdU imaging
   NOTE: This method is used at the end of the experiment by injecting EdU in tumor-bearing animals for 2 h before euthanasia and tissue collection (see step 3.3).
   1. Mount the brain sections onto microscope slides as indicated in step 4.2.11. Wash the slides with PBS for 5 min.
   2. Permeabilize the sections with 0.5% Triton X-100 in PBS for 15 min. Wash the slides for 3 x 5 min with PBS.
      ​NOTE: Resuspend the reaction cocktail according to the manufacturer's recommendations. Use fresh aliquots each time.
   3. In a light-resistant case, add the reaction cocktail evenly to the slides. Incubate the tissue sections for 30 min at room temperature.
   4. Wash the slides for 3 x 5 min with PBS.
   5. Stain the nuclei with Hoechst 33342 or an alternative nuclear counterstain for 10 min.
   6. Wash slides for 1 x 5 min with PBS.
   7. Mount the sections with aqueous mounting medium. Allow the slides to dry in the dark before imaging.

5. Quantification and data analysis

1. Tumor volume quantification
   1. Visualize and scan the H&E-stained brain sections with a tissue scanner to obtain high-resolution images or use an equivalent microscope.
   2. Mark tumors from high-resolution images by outlining their contours in each individual scan of the tissue sections (e.g., with ImageJ software).
   3. Measure tumor area manually within each section to determine tumor length (L), width (W), and total Area (A).
   4. Estimate tumor volume by applying the formula (L × W² × 0.5)¹⁵.
2. Ki67 quantification
   1. Visualize and scan brain sections with the tissue scanner to obtain high-resolution images or use an equivalent microscope.
   2. Manually outline the regions of interest within the tumor to focus the analysis using the referenced software.
   3. Run positive cell detection method using the Positive Cell Detection tool to classify cells as Ki67-positive (brown) or negative (blue) based on DAB staining intensity.
   4. Optimize detection settings by adjusting parameters like Optical Density Sum to improve detection accuracy and minimize errors, such as fragmented nuclei or missed cells.
   5. Analyze the percentage of Ki67-positive cells within the tumor regions, representing the proliferation rate of tumor cells.
3. Edu-positive (EdU⁺) cell quantification
   1. Visualize and scan brain sections with a tissue scanner fluorescence microscope or equivalent microscope that contains the following filters: Hoechst 33342 (Excitation at 350 nm, emission at 461 nm) and EdU-Click-iT 647 (Excitation at 647 nm, emission at 665 nm).
   2. Split the channels and select regions of interest (ROI) at the tumor core and periphery, ensuring consistent ROIs across all images for accurate comparisons.
   3. For total cell count, count Hoechst-stained nuclei using the Cell Counter plugin.
   4. For EdU⁺ cell count, switch to the EdU (647 nm channel) and count all EdU⁺ nuclei using the Cell Counter.
   5. Calculate the percentage of EdU⁺ cells by dividing the number of EdU⁺ cells by the total number of nuclei in each ROI.

Results

The protocol provided here successfully establishes an in vivo GB model evidenced by the presence of unilateral motor problems and the presence of highly proliferative and infiltrative tumors. The following representative results confirm the effectiveness of the method and highlight the possible sources of variability.

GB tumor growth is associated with an increase in the monitoring score during the last 4-5 days with no evidence of major weight loss (Figure 2A

Discussion

GB remains highly lethal and resistant to treatment due to its ability to evade chemotherapy and radiotherapy¹⁹. The lack of effective therapies for GB has sustained interest in developing new treatments and uncovering new pathogenic pathways. This underscores the need for animal models that accurately reproduce the characteristics of these brain tumors, particularly their treatment resistance, tissue invasion, and patient anticancer immune responses. Therefore, it is crucial to use models that mi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3,3′-Diaminobenzidine (DAB)	Merck	D5905
ABC-HRP	VECTOR	PK-4000
Alcoholic eosin Y	Merck	102439
Automatic cell counter Countess II	Thermofisher Scientific	AMQAF1000
B-27 supplement 50x	Thermofisher Scientific	17504044
Biotinylated Goat Anti-Rabbit IgG (H+L)	Santa Cruz	(BA-1000-1.5)
C57BL/6 mice	Jackson Lab	664
Click-iT EdU Cell Proliferation Kit for Imaging, Alexa Fluor 647 dye	Thermofisher Scientific	C10340
Coverlips	Fisher Scientific	12323128
DMEM/F-12 supplement	Gibco	10565018
Dulbecco's modified Eagle's medium (DMEM)	Gibco	1965092
EdU (5-ethynyl-2′-deoxyuridine)	Thermofisher Scientific	E10187
Entellan	Merck	107961
Gel boost	Clear	72-04-5022
GL261	N/A	N/A	Gift from Dr.Eric Chevet
Glass slides	Fisher Scientific	11562203
Harris hematoxilyn	Merck	HHS32
Heating mantle	N/A	N/A
hiFBS	Gibco	A5209501
Hydrochloric acid	Merck	01-01-7646
Hydrogen peroxide	DifemCare	2402069
ImageJ software
Isofluorane	Lunan Pharma	N/A
Leica CM1510s cryostat	RANKIN	CM1510S
L-Glutamine	Gibco	25030081
Low-Profile disposable blades	N/A	N/A
Mayer’s hematoxylin	Merck	51275
Microsrynge	Hamilton	87930
Mixantin Plus	Drag Pharma	N/A
Mouse Recombinant  Epidermal Growth Factor (EGF)	PeproTech	315-09-500UG
Mouse recombinant Fibroblast Growth Factor (FGF)	PeproTech	450-33-50UG
N2 supplement 100x	Thermofisher Scientific	A1370701
Naxpet Ketoprophen	Drag Pharma	N/A
Nicotears gel	Nicolich	N/A
Non essential aminoacids (MEM-NEA)	Gibco	11140050
Paraformaldehyde	Merck	30525-89-4
PBS	N/A	N/A
Penicilin/streptomycil	Gibco	15140122
Positive charged slides	Bioland	BNAO1
ProLong Gold Antifade Mountant	Thermofisher Scientific	P36930
QuPath software
Rabbit Anti-Ki67	Abcam	ab15580
Scissors	N/A	N/A
Sodium Azide	Merck	S2002
Sodium tetraborate	Merck	221732
Stereotaxic frame	Stoelting	51730
Surgical blades N°15
Surgical scalpel	ChannelMED
Sutures	TAGUM	N/A
Syringes 1 mL and 27G Needle	Cranberry	AAJECR01
Tissue-TEK O.C.T	Sakura Finetek	25608-930
Triton X-100	Merck	9036-19-5
Trypsin/EDTA	Gibco	25200056
Tweezers	N/A	N/A
V Fraction Bovine Serum Albumin	Merck	0604-29-8
Veterinary dremel
Xylene	Merck	95-47-6