Implementation of Minimally Invasive Brain Tumor Resection in Rodents for High Viability Tissue Collection

Summary

The present protocol describes a standardized resection of brain tumors in rodents through a minimally invasive approach with an integrated tissue preservation system. This technique has implications for accurately mirroring the standard of care in rodent and other animal models.

Abstract

The present protocol describes a standardized paradigm for rodent brain tumor resection and tissue preservation. In clinical practice, maximal tumor resection is the standard-of-care treatment for most brain tumors. However, most currently available preclinical brain tumor models either do not include resection, or utilize surgical resection models that are time-consuming and lead to significant postoperative morbidity, mortality, or experimental variability. In addition, performing resection in rodents can be daunting for several reasons, including a lack of clinically comparable surgical tools or protocols and the absence of an established platform for standardized tissue collection. This protocol highlights the use of a multi-functional, non-ablative resection device and an integrated tissue preservation system adapted from the clinical version of the device. The device applied in the present study combines tunable suction and a cylindrical blade at the aperture to precisely probe, cut, and suction tissue. The minimally invasive resection device performs its functions via the same burr hole used for the initial tumor implantation. This approach minimizes alterations to regional anatomy during biopsy or resection surgeries and reduces the risk of significant blood loss. These factors significantly reduced the operative time (<2 min/animal), improved postoperative animal survival, lower variability in experimental groups, and result in high viability of resected tissues and cells for future analyses. This process is facilitated by a blade speed of ~1,400 cycles/min, which allows the harvesting of tissues into a sterile closed system that can be filled with a physiologic solution of choice. Given the emerging importance of studying and accurately modeling the impact of surgery, preservation and rigorous comparative analysis of regionalized tumor resection specimens, and intra-cavity-delivered therapeutics, this unique protocol will expand opportunities to explore unanswered questions about perioperative management and therapeutic discovery for brain tumor patients.

Introduction

Glioblastoma (GBM) is the most common and aggressive primary brain tumor in adults. Despite recent advances in neurosurgery, targeted drug development, and radiation therapy, the 5-year survival rate for GBM patients is less than 5%, a statistic that has not significantly improved in over three decades¹. Hence, there is a need for more effective treatment strategies.

To develop new therapies, it is becoming increasingly apparent that investigational protocols need to (1) utilize translatable preclinical models that accurately recapitulate the tumor heterogeneity and microenvironment, (2) mirror the standard therapeutic regimen used in patients with GBM, which currently includes surgery, radiotherapy, and chemotherapy, and (3) account for the difference between resected core and residual, invasive tumor tissues^2,3,4,5. However, most of the currently available preclinical brain tumor models either do not implement surgical resection or utilize surgical resection models that are relatively time-consuming, leading to a significant amount of blood loss or lack standardization. Furthermore, performing resection of rodent brain tumors can be challenging due to a lack of clinically comparable surgical tools or protocols and the absence of an established platform⁶ for systematic tissue collection (Table 1).

The present protocol aims to describe a standardized paradigm for rodent brain tumor resection and tissue preservation using a multi-functional non-ablative minimally invasive resection system (MIRS) and an integrated tissue preservation system (TPS) (Figure 1). It is expected that this unique technique will provide a standardized platform that can be utilized in various studies in preclinical research for GBM and other types of brain tumor models. Researchers investigating therapeutic or diagnostic modalities for brain tumors can implement this protocol to achieve a standardized resection in their studies.

Protocol

All animal studies were approved by the University of Maryland and the Johns Hopkins University Institutional Animal Care and Use Committee. C57BL/6 female mice, 6-8 weeks of age, were used for the present study. The mice were obtained from commercial sources (see Table of Materials). All Biosafety Level 2 (BSL-2) regulations were followed, including the usage of masks, gloves, and gowns.

1. Initial intracranial tumor implantation

1. At the initial phase of the study, intracranially inject each mouse with 100,000 cells (GL261 murine glioma cell line) suspended in 4 µL of phosphate-buffered saline (1x PBS) to a depth of 2.5 mm following the previously published report⁷.
2. Quantify the tumor signal in each mouse using the in vivo imaging system⁸ 9 days following tumor implantation.
   NOTE: If needed, stratify the mice into two groups based on tumor burden. In the present study, the two groups were: (1) mice with relatively small tumor burden (mean bioluminescent signal = 5.5e+006 ± 0.1e+006 photons/s, n = 10) and (2) mice with relatively large tumor burden (mean bioluminescent signal = 1.69e+007 ± 0.2e+007 photons/s, n = 10), (p < 0.05, Mann-Whitney test)⁹.
3. Divide each group into two comparable subgroups.
   NOTE: In this study, the two subgroups were: untreated mice (n = 5) and mice with tumors undergoing surgical resection using the MIRS (n = 5), (p > 0.05, Mann-Whitney test)⁹.
4. Starting from the day of the resection, track the tumor progression using the in vivo imaging system at a frequency based on the tumor growth pattern.
   NOTE: For the GL261 cell line, track the tumor progression on the day of the resection and then every 3-6 days.

2. Tumor resection using MIRS

1. Anesthetize the mouse using an isoflurane-O₂ gas mixture in an induction chamber or intraperitoneal injection of xylazine/ketamine solution.
   1. If using the gas anesthetic, set the gas flow rate to 1.0 mL/min and the vaporizer to 2.0% for anesthesia induction, typically requiring 3-5 min in the chamber (see Table of Materials).
   2. If using the injectable anesthetic, anesthetize the mice by injecting 0.2 mL of anesthetic solution (80-100 mg/kg of ketamine and 10-12.5 mg/kg of xylazine, see Table of Materials) intraperitoneally.
2. Assess the animal for adequate sedation by pinching the toe. Apply ophthalmic ointment to the eyes to avoid dryness of the cornea. Administer an analgesic (0.03-0.06 mg/kg buprenorphine subcutaneously) prior to the procedure.
3. Place the mouse onto the stereotactic frame (see Table of Materials) once full sedation has been confirmed.
   NOTE: If using the gas anesthetic, place the nose of the mouse in a nose cone where it will continue to receive the isoflurane-O₂ mixture during the procedure (1.5%).
4. Remove the previous staple, then remove the hair either with a depilatory cream or by shaving. Disinfect the skin with alternating cycles of a chlorhexidine/betadine-based scrub and alcohol. Then, using a sterile scalpel, create a 1 cm longitudinal midline incision along the previous surgical scar.
5. Attach the MIRS handpiece to the stereotactic arm through the stage adapter/handpiece holder for enhanced stability and precision.
6. Set up the MIRS machine (see Table of Materials) using the following settings (Figure 1).
   1. Insert the power cord set on the rear panel into the power cord receptacle. Turn the power to the system on or off by toggling (1 = ON, 0 = OFF).
   2. Insert one end of the nitrogen hose (supplied with the console) into the male fitting on the console's rear panel. Rotate the connection nut clockwise to tighten the connection.
      NOTE: The opposite end of the hose needs to be connected to the nitrogen supply.
   3. Before attaching the hose to the nitrogen supply, confirm that the supply pressure does not exceed 100 psig, which is the input supply pressure recommended for the console.
      NOTE: The console will generate its own aspiration when activated by the foot pedal once the nitrogen has been supplied to the console.
   4. Secure the lid of the vacuum port and seal it to avoid any leakage in the vacuum system. Any leaks in the aspiration system will affect the performance of the MIRS console.
   5. If the aspiration is below 17 during setup or priming, check that the aspiration knob on the front of the console is set at the maximum level (100), ensure there is no leakage in the aspiration system, and confirm that the nitrogen input supply pressure is correct.
   6. To connect the foot pedal to the console, insert the gray foot pedal connector into its gray receptacle until it clicks and fits into position.
      NOTE: The foot pedal connector connects to the console in one orientation, and it is keyed.
   7. To connect the handpiece to the console, insert the blue handpiece connector into its blue receptacle until it clicks and fits into position.
      NOTE: The handpiece connector connects to the console in one orientation, and it is keyed.
   8. Prime each handpiece before using the system by aspirating sterile fluid from a small bowl into the aperture, through the tubing and handpiece, and then into the canister to ensure that the inside of the tubing and handpiece are lubricated to reduce the tissue occlusions.
   9. Prepare for aspiration alone or aspiration with cutting by selecting the appropriate mode on the console's front panel. Initiate using the foot pedal.
7. Insert the 23 G MIRS cannula into the burr hole to a depth of 2.5 mm.
8. Initiate the resection process by depressing the foot pedal connected to the cannula. Perform one full cycle (360°) or more of resection using the control knob in the handpiece.
   NOTE: The more cycles performed, the more volume of tumor tissue resected.
9. Once the resection process is complete, withdraw the 23 G MIRS cannula from the burr hole and use 5 mL of 1x PBS to flush the tubing and dislodge any residual debris.
10. Remove the mouse from the stereotactic frame and close the wound with a stapler or 4-0 suturing material (see Table of Materials).
11. Place the mouse on a heating pad or under a warming light during recovery from anesthesia before returning it to its cage.
12. After the experiment is complete, purge the cannula via flushing. Alternate with chilled media and air to "push" all of the resected tissue back to the collection canister. Remove the collection canister from the system and cap off with the provided cap.
13. After completing step 2.12, place the distal tip of the cannula into 3% H₂O₂ and apply suction at 24-25 in Hg to fill the suction line back to the suction collection canister and let stand for 60-90 s. Flush with sterile media pulsing air and media intermittently.
14. Monitor the mice for any neurological signs (abnormal erratic movements or seizures) following the procedure.
15. Euthanize the mice with severe neurological impairments (become lethargic, have a gaunt appearance, hunched back, or have erratic movements).
    NOTE: For the present study, 200 mg/kg of a commercially available euthanasia solution (see Table of Materials) was used to euthanize each mouse.

3. Tissue collection via TPS

1. Immerse the tumor sample in a tissue culture dish containing an RBC lysis medium (see Table of Materials) for 5 min at room temperature.
   NOTE: The tissue harvested from the MIRS into the TPS will be primarily in the form of single cells along with small chunks of tissue.
2. Place a 70 µm filter (see Table of Materials) on a 50 mL conical tube and use a plunger of a 5 mL syringe to pass the tumor sample through the filter.
3. With a transfer pipette, use the RPMI-1640 media to facilitate the passing of cells and any tissue mass through the filter.
4. Centrifuge at 428 x g for 5 min at 4 °C. Discard the supernatant by a pipette.
5. Resuspend each sample in 5 mL of prepared RPMI-1640 medium.
6. To increase tissue viability, especially if large tissue chunks are visualized on initial impression, add the required volumes of the enzyme cocktail (containing DNAse I, collagenase IV, dispase, Papain, and EDTA, see Table of Materials) to each sample. Use the vortex to mix the solutions.
   NOTE: Step 3.6 is optional. The composition of the enzyme cocktail (for a total volume of 5 mL/sample): 300 µL of DNAse I Grade II, 150 µL of Collagenase/Dispase (cleaves fibronectin, collagenase IV, I, and nonpolar amino acids), 250 µL of Papain (nonspecific protease), and 6 µL of 0.5 M EDTA.
7. Place the samples in a shaker incubator set at 200 rpm, 37 °C for 20 min.
8. After 20 min, spin the samples at 428 x g for 5 min at 4 °C. Discard the supernatant.
9. Filter single cells through a 70 µm cell strainer and spin down at 274 x g for 3 min at 4 °C. Conduct cell viability analysis¹⁰ with Trypan Blue and Hemocytometer (see Table of Materials).
   NOTE: Day 0 viability ranges from 30%-70% and increases significantly within 2-3 days.
10. Proceed to Steps 4, 5, or 6, depending on the viability test needed.

4. Growing cells in adherent culture

1. In a certified laminar flow hood, resuspend the pellet in serum-containing adherent medium (such as DMEM, 10% fetal bovine serum (FBS), and 1% penicillin/streptomycin (P/S) solution) and plate cells in an adherent cell flask.
2. Maintain the cells in a controlled incubated environment (37 °C, 5% CO₂).

5. Growing cells in suspension culture (neurospheres)

1. In a certified laminar flow hood, resuspend the pellet in serum-free complete stem cell medium¹¹ and plate in a suspension flask.
2. Maintain the cells in a controlled incubated environment (37 °C, 5% CO₂) for 2-3 days to allow neurosphere formation.
3. After visualization of the neurospheres in the culture medium, use Trypsin-EDTA or Accutase (see Table of Materials) to obtain single-cell suspensions for passaging.
   NOTE: As long as special care is taken during harvest and appropriate media supporting neural stem cells is used, the stem cells in the harvested tissue must form neurospheres within a few days.

6. Preparing cells for reimplantation

1. Resuspend the pellet at a concentration of 100,000 live cells per 4 µL of 1x PBS.
2. Immediately inject into naïve mice using the intracranial tumor implantation method (step 1).

7. Histological analysis

1. Immediately following resection, extract and fix the brains in 4% paraformaldehyde (PFA) for 24 h¹².
2. Transfer the brains to a 30% sucrose solution until they are saturated with sucrose (sunken down to the bottom of the container).
3. Transfer the brains to a 70% ethanol solution.
4. Perform paraffin block embedding, sectioning, and standard hematoxylin and eosin (H&E) staining following previously published report¹³.
   NOTE: The thickness of each section taken for staining was 10 µm.

Results

Surgical resection using the MIRS results in a significant decrease in the tumor burden
In the group with a smaller tumor burden, the mean baseline bioluminescent signal was 5.5e+006 photons/s ± 0.2e+006 in the subgroup that underwent resection. Following resection, the mean bioluminescent signal decreased to 3.09e+006 photons/s ± 0.3e+006, (p <0.0001, Mann-Whitney test)⁹ (Figure 2). The bioluminescen...

Discussion

Tumor resection is a cornerstone of neurosurgical oncology treatment plans for both low-grade and high-grade brain tumors. Cytoreduction and debulking of the tumor correlate with improved neurological function and overall survival in patients with brain tumors^1,2,5,6. Although protocols for surgical resection have been previously described in rodent models, these protocols have suffered from se...

Materials

Name	Company	Catalog Number	Comments
1 mL syringes	BD	309628
15 mL conical tubes	Corning	430052
200 proof ethanol	PharmCo	111000200
5 mL pipettes	CoStar	4487
70 micron filter	Fisher	08-771-2
Accutase	Millipore Sigma	SIG-SCR005
Anased (Xylazine injection, 100 mg/mL)	Covetrus	33198
Anesthesia System	Patterson Scientific	78935903
Anesthesic Gas Waste Container	Patterson Scientific	78909457
Bench protector underpad	Covidien	10328
C57Bl/6, 6-8 week old mice	Charles River Laboratories	Strain Code 027
ChroMini Pro	Moser	Type 1591-Q
Collagenase-Dispase	Roche	#10269638001
Countess II Automated Cell Counter	Thermo Fisher
Countess II FL Hemacytometer	Thermo Fisher	A25750
Debris Removal Solution	Miltenyi Biotech	#130-109-398
D-Luciferin	Goldbio	LUCK-1G
DMEM F12 media	Corning	10-090-CV
DMEM media	Corning	10-013-CV
DNAse I	Sigma Aldrich	#10104159001
Eppendorf tubes	Posi-Click	1149K01
Euthanasia solution	Henry Schein	71073
FBS	Millipore Sigma	F4135
Fetal Bovine Serum	Thermo Fisher	10437-028
Formalin	Invitrogen	INV-28906
Gauze	Henry Schein	101-4336
hEGF	PeproTech EC	100-15
Heparin	Sigma	H-3149
hFGF-b	PeproTech EC	1001-18B
Induction Chamber	Patterson Scientific	78933388
Isoflurane	Covetrus	11695-6777-2
Isoflurane Vaporizer	Patterson Scientific	78916954
Ketamine	Covetrus	11695-0703-1
Kopf Stereotactic frame	Kopf Instruments	5001
Lightfield Microscope	BioTek	Cytation 5
Microinjection Unit	Kopf	5001
Micromotor drill	Foredom	F210418
MRI system	Bruker	7T Biospec Avance III MRI Scanner
NICO Myriad System	NICO Corporation
Ophthalmic ointment	Puralube vet ointment
Papain	Sigma Aldrich	#P4762
PBS	Invitrogen	#14190250
PenStrep	Millipore Sigma	N1638
Percoll solution	Sigma Aldrich	#P4937
Pipette controller	Falcon	A07260
Povidone-iodine solution	Aplicare	52380-1905-08
Progesterone	Sigma	P-8783
Putrescine	Sigma	P-5780
RPMI Media	Invitrogen	INV-72400120
Scalpel blade	Covetrus	7319
Scalpel handle	Fine Science Tools	91003-12
Skin marker	Time Out	D538,851
Staple remover	MikRon	ACR9MM
Stapler	MikRon	ACA9MM
Staples	Clay Adams	427631
Stereotactic Frame	Kopf Instruments	5000
Sucrose	Sigma Aldrich	S9378
Suture, vicryl 4-0	Ethicon	J494H
T-75 culture flask	Sarstedt	83-3911-002
TheraPEAKTM ACK Lysing Buffer (1x)	Lonza	BP10-548E
Trypsin-EDTA	Corning	MDT-25-053-CI