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Method Article

Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells

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DOI:

10.3791/64216

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August 2nd, 2022

 ,  ,  ,  ,  ,  , 
1UQ Centre for Clinical Research, The University of Queensland, 2Centre for Advanced Imaging, Australian Institute for Bioengineering and Nanotechnology and ARC Training Centre for Innovation in Biomedical Imaging Technology, The University of Queensland, 3School of Biomedical Sciences, The University of Queensland, 4Mater Research and Mater Research Institute, The University of Queensland, 5Pathology Queensland, Royal Brisbane Women's Hospital

* These authors contributed equally

In This Article

Summary

Brain metastasis is a cause of severe morbidity and mortality in cancer patients. Most brain metastasis mouse models are complicated by systemic metastases confounding analysis of mortality and therapeutic intervention outcomes. Presented here is a protocol for internal carotid injection of cancer cells that produces consistent intracranial tumors with minimal systemic tumors.

Abstract

Brain metastasis is a cause of severe morbidity and mortality in cancer patients. Critical aspects of metastatic diseases, such as the complex neural microenvironment and stromal cell interaction, cannot be entirely replicated with in vitro assays; thus, animal models are critical for investigating and understanding the effects of therapeutic intervention. However, most brain tumor xenografting methods do not produce brain metastases consistently in terms of the time frame and tumor burden. Brain metastasis models generated by intracardiac injection of cancer cells can result in unintended extracranial tumor burden and lead to non-brain metastatic morbidity and mortality. Although intracranial injection of cancer cells can limit extracranial tumor formation, it has several caveats, such as the injected cells frequently form a singular tumor mass at the injection site, high leptomeningeal involvement, and damage to brain vasculature during needle penetration. This protocol describes a mouse model of brain metastasis generated by internal carotid artery injection. This method produces intracranial tumors consistently without the involvement of other organs, enabling the evaluation of therapeutic agents for brain metastasis.

Introduction

Brain metastasis is a prevalent malignancy associated with a very poor prognosis1,2. The standard of care for brain metastasis patients is multimodal, consisting of neurosurgery, whole brain radiotherapy and/or stereotactic radiosurgery depending on the patients' general health status, extracranial disease burden, and the number and location of tumors in the brain3,4. Patients with up to three intracranial lesions are eligible for surgical resection or stereotactic radiosurgery, while whole-brain radiation therapy is recommended for patients with multiple lesions to avoid the risk of surgery-related infection and edema5. However, whole brain radiotherapy can inflict damage on radiosensitive brain structures, contributing to poor quality of life6.

Systemic therapy is a non-invasive alternative and logical approach to treat patients with multiple lesions7. However, it is less considered due to the long standing notion that systemic therapies have poor efficacy because the passive delivery of cytotoxic drugs via the bloodstream cannot achieve therapeutic levels in the brain without the risk of unsafe toxicity8. This paradigm is starting to change with the recently U.S. Food and Drug Administration (FDA)-approved systemic therapy (tucatinib with trastuzumab and capecitabine indicated for metastatic HER2+ breast cancer brain metastasis)9,10,11,12 and the update in treatment guidelines to include consideration of systemic therapy options for brain metastasis patients13,14.

In this context, developments in the field of molecular targeted therapy, immunotherapy, and alternative drug-delivery systems, such as a targeted nano-drug carrier, can potentially overcome the challenges of brain metastasis treatment15,16,17,18. In addition, chemical and mechanical approaches to improve drug delivery via permeabilization of the brain-tumor barrier are also being investigated19,20. To study and optimize such approaches to be fit for purpose, it is crucial to use preclinical models that not only mirror the complex physiology of brain metastasis but also allow for objective analysis of intracranial drug response.

Broadly, the current approaches to model brain metastasis in vivo involve intracardiac (left ventricle), intravenous (usually tail vein), intracranial, or intracarotid (common carotid artery) injection of cancer cells in mice21,22,23,24,25,26,27. Apart from tumor engraftment strategies, genetically engineered mouse models where tumor formation is triggered by the removal of tumor suppressor genes or activation of oncogenes are useful for tumor modeling. However, only a few genetically engineered mouse models are reported to produce secondary tumors and even fewer that reliably produce brain metastases28,29,30.

Engraftment methods such as intracardiac (left ventricle) and intravenous (usually tail vein) injection mimic the systemic dissemination of cancer. These models typically produce lesions in multiple organs (e.g., brain, lungs, liver, kidneys, spleen) depending on the capillary bed that traps most tumor cells during their circulatory 'first pass'31. However, inconsistent rates of brain engraftment will require more animals to achieve the sample size for the desired statistical power. The number of tumor cells that eventually get established in the brain via these intracardiac and intravenous injection methods is variable. Hence, brain metastasis tumor burden can vary between animals and the difference in progression can make standardizing the experimental timeline and interpretation of results a challenge. The extracranial tumor burden can lead to non-brain metastasis mortality, rendering these models unsuitable for evaluating intracranial efficacy. Brain-tropic cell lines have been established using artificial clonal selection processes to reduce extracranial establishment, but take rates have been inconsistent, and the clonal selection process can reduce the heterogeneity normally found in human tumors32.

Brain-specific engraftment methods such as the intracranial and intracarotid injection allow for more consistent and efficient brain metastasis modeling. In the intracranial method33, cancer cells are typically injected into the frontal cerebral cortex, which generates quick and reproducible tumor outgrowth with low systemic involvement. While the procedure is well tolerated with low mortality33, the caveats are that it is a relatively crude approach that rapidly introduces a (localized) bolus of cells in the brain and does not model early brain metastasis pathogenesis. The needle damages brain tissue vasculature, which then causes localized inflammation5,34. From experience, there is a tendency for tumor cell injectate to reflux during removal of the needle, leading to leptomeningeal involvement. Alternatively, the intracarotid method delivers cells into the common carotid artery with brain microvasculature as the first capillary bed to be encountered, modeling survival in circulation, extravasation, and colonization24. In agreement with others25, our experience with this method found that it can result in facial tumors due to unintentional delivery of cancer cells via the external carotid artery to capillary beds in these tissues (unpublished data). It is possible to prevent facial tumors by first ligating the external carotid artery before common carotid artery injection (Figure 1). In the rest of the article, this method is referred to as the 'internal carotid artery injection'. From experience, the internal carotid artery injection method consistently generates brain metastasis with very few systemic events and has been successful in generating brain metastasis models of different primary cancers (e.g., melanoma, breast, and lung cancers) (Figure 1). The disadvantages are that it is technically challenging, time-consuming, invasive, and requires careful optimization of cell numbers and a monitoring timeline. In summary, both the intracranial and internal carotid artery injection methods produce mouse models suitable for evaluating therapeutic impact on brain tumor-related survival benefit.

This protocol describes the internal carotid artery injection method to produce a mouse model of brain metastasis with almost no systemic involvement and therefore suitable for preclinical evaluation of drug distribution and efficacy of experimental therapeutics.

Mouse brain metastasis model diagram; cancer cells injected into carotid; pathology results show melanoma, lung, breast cancer brain mets.
Figure 1: Schematic representation of internal carotid artery injection protocol for brain metastasis. Internal carotid artery injection with external carotid artery ligation can reliably produce a brain metastasis model from various primary cancers. In this protocol, three ligatures are placed on the carotid artery (annotated as L1-L3 in the figure). Please click here to view a larger version of this figure.

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Protocol

All studies were conducted within the guidelines of the Animal Ethics Committee of The University of Queensland (UQCCR/186/19), and the Australian Code for the Care and Use of Animals for Science Purpose.

1. Preparation of cancer cells for injection

NOTE: In this study, the human breast cancer cell line, BT-474 (BT474), was used. BT474 was cultured in complete growth medium comprising RPMI 1640 medium supplemented with 10% fetal bovine serum and 1% insulin. The cells were maintained in an incubator at 37 °C with 5% carbon dioxide in air atmosphere. Authenticate the cell line by satellite tandem repeats testing35, confirm expression of the reporter protein (e.g., luciferase) if any, and check for mycoplasma infection.

  1. Seed BT474 cancer cells at a seeding density of 2.0 x 106 cells into a T75 flask using 10 mL of complete growth media and culture (at 37 °C with 5% CO2) to 70%-80% confluency prior to injection.
  2. On the day of injection, discard growth media and wash the cell monolayer with phosphate buffered saline (PBS) twice.
  3. Add 5 mL of prewarmed cell culture dissociation reagent (see Table of Materials) and incubate at 37 °C for 5 min or until cells have detached. After 5 min, gently tap the flask to aid cell detachment.
  4. Add 5 mL of complete growth media containing 10% fetal bovine serum to quench the dissociation reagent activity.
  5. Gently resuspend the cells by pipetting to reduce cell clumps.
  6. Transfer the cell suspension into a 50 mL tube and centrifuge at 180 x g for 3 min at room temperature.
  7. Decant the supernatant and resuspend the cell pellet in 10 mL of Hank's Balanced Salt Solution (HBSS) without calcium and magnesium to minimize cell clumping.
  8. Centrifuge the cell suspension at 180 x g for 3 min at room temperature.
  9. Decant the supernatant to remove residual serum/dissociation reagent and resuspend the cell pellet in 3 mL of HBSS.
  10. Place a 100 µm cell strainer on a fresh 50 mL conical tube and pass the cell suspension through to remove cell clumps.
    NOTE: A single cell suspension is essential to minimize blood vessel occlusion and the risk of stroke on injection.
  11. Calculate the number of viable cells using Trypan Blue exclusion and a hemocytometer with standard methods.
  12. Dilute the cell suspension with HBSS to a cell concentration of 2.5 x 106 cells/mL.
  13. Keep the tube horizontal on ice and gently rock the tube periodically to minimize clumping. The cell suspension can be stored on ice for a maximum of 6 h.
    ​NOTE: Rocking was done manually but this can also be done using a shaker at low rpm.

2. Preparation of the mouse for the procedure

NOTE: In this study, 4-5 weeks old, female NOD scid mice were used. Introduce soft-diet recovery food (e.g., diet gel, hydrogel, mashed mouse chow) to mice 3 days before the procedure to encourage feeding after the procedure.

  1. Autoclave surgical tools. Spray and wipe down the surgical area and equipment with surface disinfectant, followed by 70% ethanol.
  2. Place an animal heat mat under the surgical board to prevent hypothermia. Switch this on 30 min prior to surgery. Spray and wipe down the surgical board with surface disinfectant followed by 70% ethanol.
  3. Prepare a clean animal housing cage and a warm heating pad for recovery.
  4. Put on clean personal protective equipment (gown, mask, hairnet, and gloves). Maintain sterility throughout the procedure by using clean exam gloves and a 'instruments tips-only' technique.
  5. Anesthetize the mouse with an anesthetic chamber using 5% isoflurane with an oxygen flow of 2 L/min until the mouse loses pedal reflex.
  6. Take the animal out of the chamber and place it in a nose cone delivering 2% isoflurane at an oxygen flow of 2 L/min for the remaining surgical procedure.
  7. Ear punch mouse for identification and use electric clippers to shave the fur from the neck region. Clean excess hair from exposed skin using tape.
  8. Weigh the mouse for calculating the anesthetic and analgesic drug doses required. Administer buprenorphine and meloxicam at 50 µg/kg and 1 mg/kg, respectively, via subcutaneous injection.
  9. Transfer the mouse to a warm surgical board and secure the nose cone with tape.
  10. Apply ocular lubricant to the eyes to prevent drying.
  11. Secure the mouse gently by first hooking the upper incisor teeth using thread taped to the surgical board, followed by taping the front and hind legs. This step extends the body and keeps the neck straight during the procedure.
  12. Perform preoperative skin preparation as described below.
    1. Wipe the neck with topical antiseptic (povidone-iodine) to reduce skin microflora load and remove loose hair. Clean from the center of the skin, working outward to prevent recontamination of the incision site. Repeat the process using 70% ethanol. Perform three alternating rounds of iodine and ethanol for disinfection.
    2. Lay a surgical drape over the animal. This is cut and fashioned from a piece of sterile paper towel or autoclave bag.
  13. Lay out sterile paper towels or autoclave bags for surgical tools.
  14. Check for reflex via 'Pinch test' to ensure sufficient anesthesia before continuing with the procedure.

3. Internal carotid injection

NOTE: In this experiment, a 31 G infusion cannula and foot-activated syringe-driver setup was utilized to facilitate the injection procedure (Supplementary Figure 1). This setup is optional and the user can use a 31 G insulin syringe and skip steps 3.11 and 3.12. To prepare the infusion cannula, pull and separate the needle portion from the syringe fitting portion of a 31 G needle using two pairs of suture clamps. Next, attach the needle portion to one end of a fine infusion tubing approximately 10 cm in length.

  1. Position the dissection microscope over the mouse.
  2. Using scissors, make a vertical 15 mm incision along the midline at the neck region starting from 5 mm below the jaw to the thoracic inlet.
  3. Using two pairs of angled forceps, part the skin and underlying salivary glands, and apply retractors to keep the trachea exposed. The next step will expose the carotid sheath that lies parallel to the trachea.
  4. Using two pairs of fine angled forceps, bluntly dissect the muscle and fat tissue adjacent to the trachea to expose the right carotid sheath. The carotid sheath is the fibrous layer covering the common carotid artery, vein, and vagus nerve, and this bundle can be visualized by the bright red common carotid artery. In this study, the injection was performed on the right carotid artery.
  5. Clear a segment of the common carotid artery caudal to the carotid bifurcation of the surrounding fascia and separate it from the vagus nerve and veins.
  6. Isolate and clear the carotid bifurcation (the junction that joins the external and internal carotid arteries) from the surrounding nerves and fascia. Position fine forceps under the external carotid artery and pass a silk suture (5-0 thickness) under the artery. Knot and tighten the suture and cut excess line.
    NOTE: This ligature (L1) will prevent injectate from transiting through the external carotid artery.
  7. Position fine forceps under the common carotid artery and pass a silk suture (5-0 thickness) under the artery. Tie a knot and tighten the suture at a position proximal to the proposed injection site. Cut the excess suture leaving about 10 mm of line.
    NOTE: This second ligature (L2) will restrict blood flow and bleeding after injection. It is also used to position and hold the carotid artery during the injection.
  8. Cut and moisten a strip of (autoclaved) low-lint disposable wipers (see Table of Materials) about 10 mm x 5 mm. Fold the strip into 4 mm x 5 mm, 2-3 mm thick, and place it underneath the carotid artery at the proposed site of injection. This will support the vessel during the injection.
  9. On the common carotid artery rostral to the proposed injection site, place a third ligation (L3) with a loose knot. This is tightened only after the injection (at step 3.16).
  10. Gently agitate the cell suspension and draw 200 µL of cell suspension into an insulin syringe (with 31 G needle).
  11. Load the syringe into the syringe driver that is connected to an activating foot pedal.
  12. Attach a fine cannula with a 31 G needle to the syringe and prime the line.
  13. Check whether the carotid artery is well-positioned and pressurized.
  14. Using two fine angled forceps, one gently tensioning onto the end of the first ligature and the other holding the 31 G needle, slowly insert the needle with bevel up into the lumen of the blood vessel taking care not to puncture it.
  15. Slowly inject 100 µL of cell suspension (from step 1.13) into the common carotid artery at 10 µL/s. This will deliver 2.5 x 105 cells into the blood vessel. Successful injection is visualized via the clearing of blood from the carotid blood vessel.
  16. Gently lift and tighten the loose ligature (L3) (from step 3.9) immediately after withdrawing the needle to prevent backflow and bleeding. Trim excess suture.
    NOTE: It is normal to observe a small amount of blood spurting after the withdrawal of the needle. However, there must not be any active bleeding after the third ligature is tightened.
  17. Remove the piece of moistened low-lint disposable wipers.
  18. Using a P200 pipette, rinse the surgical cavity twice with 150-200 µL of sterile water or saline.
  19. Check again for bleeding, and then remove the retractors.
  20. Reposition the soft tissue, salivary glands, and skin over the carotid artery and trachea.
  21. Close the skin layer of the incision using a suture needle holder, forceps, and an absorbable or non-absorbable 6/0 monofilament suture in a continuous pattern.
  22. Discard the cannula needle syringe and prepare a new setup for the next mouse. Using a new syringe will ensure that the number of cells injected into each mouse is consistent.
    NOTE: Representative snapshots of the procedure are in Supplementary Figure 2. If the vessel is punctured or torn by the needle, indicated by the leakage of injectate or bleeding, the procedure is deemed unsuccessful. Following this, the needle must be withdrawn, and the third ligature must be immediately tightened to prevent further bleeding. If bleeding is persistent after tightening of the suture, the animal must be euthanized with pentobarbital.

4. Post-injection recovery

  1. Inject buprenorphine (50 µg/kg) and meloxicam (1 mg/kg) via subcutaneous injection as post-surgical pain relief.
  2. Move the animal to a warm and clean cage to recover from anesthesia. It is normal for an animal to have subdued activity (huddled and inactive or moving around slowly) after waking up.
  3. After 30-45 min, transfer mice to a long-term holding facility.
  4. Provide mice with a soft diet (diet gel, hydrogel, mash) for at least a week post-surgery and check physical status daily with special attention for signs of stroke, infection, bleeding around the wound site.
  5. Two days post-surgery, it is typical for the animal to have reduced activity, mild ruffled fur and lost 15% of pre-operative body weight. Administer analgesic (meloxicam) daily for 2-3 days post-surgery to manage pain and aid recovery.
  6. From day 3 onwards, animals must have regained activity, increased in feeding and grooming frequency, and regaining weight. Euthanize animals with persisting physical condition deficits (pain, huddled, inactive, weight loss) after 4 days post-surgery by injecting sodium pentobarbital at 200 mg/kg via intraperitoneal injection.
  7. Tumor engraftment and progression can be monitored using anesthesia and imaging modality of choice such as bioluminescent imaging, MRI or PET/MRI. The requirement and ability to undertake such imaging will be inherently dependent on the individual project aims and facility within which it is undertaken, and depending on the type of reporter tag the cell lines used, accessibility of relevant radiochemistry and nuclear imaging facilities36,37
    NOTE: Some animals may not respond well and experience a stroke despite a successful procedure. After the procedure, animals that present with any symptoms of neurological distress (turning head and pulling to one side, circling behavior, rolling, thrashing, loss of motor function) must be euthanized immediately.

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Results

Comparing common carotid artery injection with or without external carotid artery ligation
When cancer cells were injected via the common carotid artery without first ligating external carotid artery24, facial tumors were found in 77.8% of the grafted mice (n = 7/9 animals). An example of facial tumor is illustrated in Supplementary Figure 3. The method described in this protocol prevents unintended facial metastasis by ligating the external ...

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Discussion

Brain metastasis is a complex process of cancer cells spreading from their primary site to the brain. Different animal models are available that mirror certain stages of this multi-step process and there are physiological and practical considerations to designing preclinical metastasis studies41,42. Most published studies investigating the use of nanomedicine for brain metastasis treatment have used intracardiac43,

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Disclosures

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the paper.

Acknowledgements

This research was funded by The Australian National Health and Medical Research Council (NHMRC), grant number APP1162560. ML was funded by a UQ postgraduate research scholarship. We would like to thank everyone who assisted with animal husbandry and in vivo imaging of the animals. We thank the Royal Brisbane and Women's Hospital for donating aliquots of zirconium for this study.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
100µm cell strainerCorningCLS431752
30G Microlance needleBD23748
31G Ultra-Fine II insulin syringeBD326103
Angled forcepsProscitechT67A-SSFine pointed, angled without serrations, 18mm tip, length 128 mm
Animal heat mat
Antibiotic and antimycoticThermoFisher Scientific15240062
Autoclave bags
BT-474 (HTB-20) breast cancer cell lineATCCHTB-20
Buprenorphine (TEMGESIC)
Countess cell counterThermoFisher ScientificC10227
Diet-76AClearH2O72-07-5022
Dissection microscope
Ear puncher
Electric clippers
Fine angled forcepsProscitechDEF11063-07Angled 45°, Tip smooth, Tip width: 0.4 mm, Tip dimension: 0.4 x 0.3 mm, length 9cm
Fine tubing for cannula, Tubing OD (in) 1/32, Tubing ID (in) 1/100inCole ParmerEW-06419-00
Foetal bovine serumThermoFisher Scientific26140079
Hank's Balanced Salt Solution without calcium and magnesiumThermoFisher Scientific14170120
HydrogelClearH2O70-01-5022
Isoflurane
Kimwipes Low lint disposable wipersKimberly Clark- KimwipesZ188964
Mashed mouse chow
Meloxicam (METACAM)
Nose coneFashioned out of a microfuge tube
PAA ocular lubricant (Carbomer 2mg/g) Bausch and lomb
Povidone-iodine solutionBetadine2505692
PPE (glove, mask, gown, hairnet)
RetractorsKent ScientificSURGI-5001
RPMI 1640 MediaThermoFisher Scientific11875093
Silk suture 13mm 5-0, P3, 45cmEthiconJJ-640G
Sterile normal salineThermoFisher ScientificTM4469
Sticky tape
Surgical boardA chopping board wrapped with autoclavable bag.
Surgical scissorsProscitechT104Tip Dimensions (LxD): 38x7mm, Length 115mm
Suture forcep/ Curved Brophy forcepsProscitechT113CCurved, Rounded narrow 2 mm tip, with serrations, length 165 mm
Suture needle holder (Olsen Hegar needle holder)ProscitechTC1322-180length 190 mm, ratchet clamp
Syringe driver with foot pedal/ UMP3 Ultra micro pumpWorld Precision InstrumentsUMP3-3
T75 tissue culture flaskThermoFisher Scientific156499
Thread
Trigene II surface disinfectantCeva
Trypan Blue and Cell Counting Chamber SlidesThermoFisher ScientificC10228
TrypLE Express dissociating mediumThermoFisher Scientific12605010

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