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&#39;s Hospital for donating aliquots of zirconium for this study.

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 &#176;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.
<ol>
	<li>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 &#176;C with 5% CO2) to 70%-80% confluency prior to injection.</li>
	<li>On the day of injection, discard growth media and wash the cell monolayer with phosphate buffered saline (PBS) twice.</li>
	<li>Add 5 mL of prewarmed cell culture dissociation reagent (see Table of Materials) and incubate at 37 &#176;C for 5 min or until cells have detached. After 5 min, gently tap the flask to aid cell detachment.</li>
	<li>Add 5 mL of complete growth media containing 10% fetal bovine serum to quench the dissociation reagent activity.</li>
	<li>Gently resuspend the cells by pipetting to reduce cell clumps.</li>
	<li>Transfer the cell suspension into a 50 mL tube and centrifuge at 180 x g for 3 min at room temperature.</li>
	<li>Decant the supernatant and resuspend the cell pellet in 10 mL of Hank&#39;s Balanced Salt Solution (HBSS) without calcium and magnesium to minimize cell clumping.</li>
	<li>Centrifuge the cell suspension at 180 x g for 3 min at room temperature.</li>
	<li>Decant the supernatant to remove residual serum/dissociation reagent and resuspend the cell pellet in 3 mL of HBSS.</li>
	<li>Place a 100 &#181;m 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.</li>
	<li>Calculate the number of viable cells using Trypan Blue exclusion and a hemocytometer with standard methods.</li>
	<li>Dilute the cell suspension with HBSS to a cell concentration of 2.5 x 106 cells/mL.</li>
	<li>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. 
	&#8203;NOTE: Rocking was done manually but this can also be done using a shaker at low rpm.</li>
</ol>
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.
<ol>
	<li>Autoclave surgical tools. Spray and wipe down the surgical area and equipment with surface disinfectant, followed by 70% ethanol.</li>
	<li>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.</li>
	<li>Prepare a clean animal housing cage and a warm heating pad for recovery.</li>
	<li>Put on clean personal protective equipment (gown, mask, hairnet, and gloves).&#160;Maintain sterility throughout the procedure by using clean exam gloves and a &#39;instruments tips-only&#39; technique.</li>
	<li>Anesthetize the mouse with an anesthetic chamber using 5% isoflurane with an oxygen flow of 2 L/min until the mouse loses pedal reflex.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Weigh the mouse for calculating the anesthetic and analgesic drug doses required. Administer buprenorphine and meloxicam at 50 &#181;g/kg and 1 mg/kg, respectively, via subcutaneous injection.</li>
	<li>Transfer the mouse to a warm surgical board and secure the nose cone with tape.</li>
	<li>Apply ocular lubricant to the eyes to prevent drying.</li>
	<li>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.</li>
	<li>Perform preoperative skin preparation as described below.
	<ol>
		<li>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.</li>
		<li>Lay a surgical drape over the animal. This is cut and fashioned from a piece of sterile paper towel or autoclave bag.</li>
	</ol>
	</li>
	<li>Lay out sterile paper towels or autoclave bags for surgical tools.</li>
	<li>Check for reflex via &#39;Pinch test&#39; to ensure sufficient anesthesia before continuing with the procedure.</li>
</ol>
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.
<ol>
	<li>Position the dissection microscope over the mouse.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>Gently agitate the cell suspension and draw 200 &#181;L of cell suspension into an insulin syringe (with 31 G needle).</li>
	<li>Load the syringe into the syringe driver that is connected to an activating foot pedal.</li>
	<li>Attach a fine cannula with a 31 G needle to the syringe and prime the line.</li>
	<li>Check whether the carotid artery is well-positioned and pressurized.</li>
	<li>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.</li>
	<li>Slowly inject 100 &#181;L of cell suspension (from step 1.13) into the common carotid artery at 10 &#181;L/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.</li>
	<li>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.</li>
	<li>Remove the piece of moistened low-lint disposable wipers.</li>
	<li>Using a P200 pipette, rinse the surgical cavity twice with 150-200 &#181;L of sterile water or saline.</li>
	<li>Check again for bleeding, and then remove the retractors.</li>
	<li>Reposition the soft tissue, salivary glands, and skin over the carotid artery and trachea.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
4. Post-injection recovery
<ol>
	<li>Inject buprenorphine (50 &#181;g/kg) and meloxicam (1 mg/kg) via subcutaneous injection as post-surgical pain relief.</li>
	<li>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.</li>
	<li>After 30-45 min, transfer mice to a long-term holding facility.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>

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.

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,<sup class="xr...

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&#39; 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 &#39;first pass&#39;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 &#39;internal carotid artery injection&#39;. 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.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64216/64216fig01.jpg" /> 
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). <a href="https://www.jove.com/files/ftp_upload/64216/64216fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100&#181;m cell strainer</td>
			<td>Corning</td>
			<td>CLS431752</td>
			<td></td>
		</tr>
		<tr>
			<td>30G Microlance needle</td>
			<td>BD</td>
			<td>23748</td>
			<td></td>
		</tr>
		<tr>
			<td>31G Ultra-Fine II insulin syringe</td>
			<td>BD</td>
			<td>326103</td>
			<td></td>
		</tr>
		<tr>
			<td>Angled forceps</td>
			<td>Proscitech</td>
			<td>T67A-SS</td>
			<td>Fine pointed, angled without serrations, 18mm tip, length 128 mm</td>
		</tr>
		<tr>
			<td>Animal heat mat</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic and antimycotic</td>
			<td>ThermoFisher Scientific</td>
			<td>15240062</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclave bags</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BT-474 (HTB-20) breast cancer cell line</td>
			<td>ATCC</td>
			<td>HTB-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine (TEMGESIC)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Countess cell counter</td>
			<td>ThermoFisher Scientific</td>
			<td>C10227</td>
			<td></td>
		</tr>
		<tr>
			<td>Diet-76A</td>
			<td>ClearH2O</td>
			<td>72-07-5022</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ear puncher</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electric clippers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fine angled forceps</td>
			<td>Proscitech</td>
			<td>DEF11063-07</td>
			<td>Angled 45&#176;, Tip smooth, Tip width: 0.4 mm, Tip dimension: 0.4 x 0.3 mm, length 9cm</td>
		</tr>
		<tr>
			<td>Fine tubing for cannula, Tubing OD (in) 1/32, Tubing ID (in) 1/100in</td>
			<td>Cole Parmer</td>
			<td>EW-06419-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Foetal bovine serum</td>
			<td>ThermoFisher Scientific</td>
			<td>26140079</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#39;s Balanced Salt Solution without calcium and magnesium</td>
			<td>ThermoFisher Scientific</td>
			<td>14170120</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogel</td>
			<td>ClearH2O</td>
			<td>70-01-5022</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipes Low lint disposable wipers</td>
			<td>Kimberly Clark- Kimwipes</td>
			<td>Z188964</td>
			<td></td>
		</tr>
		<tr>
			<td>Mashed mouse chow</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Meloxicam (METACAM)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nose cone</td>
			<td></td>
			<td></td>
			<td>Fashioned out of a microfuge tube</td>
		</tr>
		<tr>
			<td>PAA ocular lubricant (Carbomer 2mg/g)</td>
			<td>&#160;Bausch and lomb</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone-iodine solution</td>
			<td>Betadine</td>
			<td>2505692</td>
			<td></td>
		</tr>
		<tr>
			<td>PPE (glove, mask, gown, hairnet)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Retractors</td>
			<td>Kent Scientific</td>
			<td>SURGI-5001</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640 Media</td>
			<td>ThermoFisher Scientific</td>
			<td>11875093</td>
			<td></td>
		</tr>
		<tr>
			<td>Silk suture 13mm 5-0, P3, 45cm</td>
			<td>Ethicon</td>
			<td>JJ-640G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile normal saline</td>
			<td>ThermoFisher Scientific</td>
			<td>TM4469</td>
			<td></td>
		</tr>
		<tr>
			<td>Sticky tape</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical board</td>
			<td></td>
			<td></td>
			<td>A chopping board wrapped with autoclavable bag.</td>
		</tr>
		<tr>
			<td>Surgical scissors</td>
			<td>Proscitech</td>
			<td>T104</td>
			<td>Tip Dimensions (LxD): 38x7mm, Length 115mm</td>
		</tr>
		<tr>
			<td>Suture forcep/ Curved Brophy forceps</td>
			<td>Proscitech</td>
			<td>T113C</td>
			<td>Curved, Rounded narrow 2 mm tip, with serrations, length 165 mm</td>
		</tr>
		<tr>
			<td>Suture needle holder (Olsen Hegar needle holder)</td>
			<td>Proscitech</td>
			<td>TC1322-180</td>
			<td>length 190 mm, ratchet clamp</td>
		</tr>
		<tr>
			<td>Syringe driver with foot pedal/ UMP3 Ultra micro pump</td>
			<td>World Precision Instruments</td>
			<td>UMP3-3</td>
			<td></td>
		</tr>
		<tr>
			<td>T75 tissue culture flask</td>
			<td>ThermoFisher Scientific</td>
			<td>156499</td>
			<td></td>
		</tr>
		<tr>
			<td>Thread</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trigene II surface disinfectant</td>
			<td>Ceva</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue and Cell Counting Chamber Slides</td>
			<td>ThermoFisher Scientific</td>
			<td>C10228</td>
			<td></td>
		</tr>
		<tr>
			<td>TrypLE Express dissociating medium</td>
			<td>ThermoFisher Scientific</td>
			<td>12605010</td>
			<td></td>
		</tr>
	</tbody>
</table>

modeling brain metastasis by internal carotid artery injection of cancer cells

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.

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 externa...

Watch this Scientific Journal Video about Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells at JoVE.com

Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells

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.

Brain metastasis is a cause of severe morbidity and mortality in cancer patients. Critical aspects of metastatic diseases, such as the complex neural ...

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5.8K Views.  The University of Queensland. 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.

Video: Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells

Intra-iliac Artery Injection for Efficient and Selective Modeling of Microscopic Bone Metastasis

Intra-iliac artery (IIA) injection is an efficient approach to introduce metastatic lesions of various cancer cells in animals. Compared to the widely used intra-cardiac and intra-tibial injections, IIA injection brings several advantages. First, it can deliver a large quantity of cancer cells specifically to hind limb bones, thereby providing spatiotemporally synchronized early-stage colonization events and allowing robust quantification and swift detection of disseminated tumor cells. Second, it injects cancer cells into the circulation without damaging the local tissues, thereby avoiding inflammatory and wound-healing processes that confound the bone colonization process. Third, IIA injection causes very little metastatic growth in non-bone organs, thereby preventing animals from succumbing to other vital metastases, and allowing continuous monitoring of indolent bone lesions. These advantages are especially useful for the inspection of progression from single cancer cells to multi-cell micrometastases, which has largely been elusive in the past. When combined with cutting-edge approaches of biological imaging and bone histology, IIA injection can be applied to various research purposes related to bone metastases.

This manuscript provides the detailed procedure of intra-iliac artery (IIA) injection, a technique to deliver cancer cells specifically to hind limb tissues including bones to establish experimental bone metastases. Although initially established with breast tumor models, this protocol can be easily extended to other cancer types.

Intra-iliac artery (IIA) injection is an efficient approach to introduce metastatic lesions of various cancer cells in animals. Compared to the widely ...

A Portal Vein Injection Model to Study Liver Metastasis of Breast Cancer

Breast cancer is the leading cause of cancer-related mortality in women worldwide. Liver metastasis is involved in upwards of 30% of cases with breast cancer metastasis, and results in poor outcomes with median survival rates of only 4.8 - 15 months. Current rodent models of breast cancer metastasis, including primary tumor cell xenograft and spontaneous tumor models, rarely metastasize to the liver. Intracardiac and intrasplenic injection models do result in liver metastases, however these models can be confounded by concomitant secondary-site metastasis, or by compromised immunity due to removal of the spleen to avoid tumor growth at the injection site. To address the need for improved liver metastasis models, a murine portal vein injection method that delivers tumor cells firstly and directly to the liver was developed. This model delivers tumor cells to the liver without complications of concurrent metastases in other organs or removal of the spleen. The optimized portal vein protocol employs small injection volumes of 5 - 10 &#956;l, &#8805; 32 gauge needles, and hemostatic gauze at the injection site to control for blood loss. The portal vein injection approach in Balb/c female mice using three syngeneic mammary tumor lines of varying metastatic potential was tested; high-metastatic 4T1 cells, moderate-metastatic D2A1 cells, and low-metastatic D2.OR cells. Concentrations of &#8804; 10,000 cells/injection results in a latency of ~ 20 - 40 days for development of liver metastases with the higher metastatic 4T1 and D2A1 lines, and &#62; 55 days for the less aggressive D2.OR line. This model represents an important tool to study breast cancer metastasis to the liver, and may be applicable to other cancers that frequently metastasize to the liver including colorectal and pancreatic adenocarcinomas.

A surgical procedure was developed to deliver mammary tumor cells to the murine liver via portal vein injection. This model permits investigation of late stages of liver metastasis in a fully immune competent host, including tumor cell extravasation, seeding, survival, and metastatic outgrowth in the liver.

Breast cancer is the leading cause of cancer-related mortality in women worldwide. Liver metastasis is involved in upwards of 30% of cases with breast ...

Intracarotid Cancer Cell Injection to Produce Mouse Models of Brain Metastasis

Metastasis, the spread and growth of malignant cells at secondary sites within a patient&#39;s body, accounts for &#62; 90% of cancer-related mortality. Recently, impressive advances in novel therapies have dramatically prolonged survival and improved quality of life for many cancer patients. Sadly, incidence of brain metastatic recurrences is fast rising, and all current therapies are merely palliative. Hence, good experimental animal models are urgently needed to facilitate in-depth studies of the disease biology and to assess novel therapeutic regimens for preclinical evaluation. However, the standard in vivo metastasis assay via tail vein injection of cancer cells produces predominantly lung metastatic lesions; animals usually succumb to the lung tumor burden before any meaningful outgrowth of brain metastasis. Intracardiac injection of tumor cells produces metastatic lesions to multiple organ sites including the brain; however, the variability of tumor growth produced with this model is large, dampening its utility in evaluating therapeutic efficacy. To generate reliable and consistent animal models for brain metastasis study, here we describe a procedure for producing experimental brain metastasis in the house mouse (Mus musculus) via intracarotid injection of tumor cells. This approach allows one to produce large number of brain metastasis-bearing mice with similar growth and mortality characteristics, thus facilitating research efforts to study basic biological mechanisms and to assess novel therapeutic agents.

Brain metastasis has become an urgent unmet medical need as its incidence has increased while therapeutic options have remained palliative. Creating experimental animal models of brain metastasis via intracarotid arterial injection of cancer cells facilitates mechanistic studies of the disease biology and evaluation of novel intervention regimens.

Metastasis, the spread and growth of malignant cells at secondary sites within a patient&#39;s body, accounts for &#62; 90% of cancer-related mortality. ...

Micromanipulation of Circulating Tumor Cells for Downstream Molecular Analysis and Metastatic Potential Assessment

Blood-borne metastasis accounts for&#160;most cancer-related deaths and involves circulating tumor cells (CTCs) that are successful in establishing new tumors at distant sites. CTCs are found in the bloodstream of patients as single cells (single CTCs) or as multicellular aggregates (CTC clusters and CTC-white blood cell clusters), with the latter displaying a higher metastatic ability. Beyond enumeration, phenotypic and molecular analysis is extraordinarily important to dissect CTC biology and to identify actionable vulnerabilities. Here, we provide a detailed description of a workflow that includes CTC immunostaining and micromanipulation, ex vivo culture to assess&#160;proliferative and survival capabilities of individual cells, and in vivo metastasis-formation assays. Additionally, we provide a protocol to achieve the dissociation of CTC clusters into individual cells and the investigation of intra-cluster heterogeneity. With these approaches, for instance, we precisely quantify survival and proliferative potential of single CTCs and individual cells within CTC clusters, leading us to the observation that cells within clusters display better survival and proliferation in ex vivo cultures compared to single CTCs. Overall, our workflow offers a platform to dissect the characteristics of CTCs at the single cell level, aiming towards the identification of metastasis-relevant pathways and a better understanding of CTC biology.

Here, we present an integrated workflow to identify phenotypic and molecular features that characterize circulating tumor cells (CTCs). We combine live immunostaining and robotic micromanipulation of single and clustered CTCs with single cell-based techniques for downstream analysis and assessment of metastasis-seeding ability.

Blood-borne metastasis accounts for&#160;most cancer-related deaths and involves circulating tumor cells (CTCs) that are successful in establishing new ...

Pathological Analysis of Lung Metastasis Following Lateral Tail-Vein Injection of Tumor Cells

Metastasis, the primary cause of morbidity and mortality for most cancer patients, can be challenging to model preclinically in mice. Few spontaneous metastasis models are available. Thus, the experimental metastasis model involving tail-vein injection of suitable cell lines is a mainstay of metastasis research. When cancer cells are injected into the lateral tail-vein, the lung is their preferred site of colonization. A potential limitation of this technique is the accurate quantification of the metastatic lung tumor burden. While some investigators count macrometastases of a pre-defined size and/or include micrometastases following sectioning of tissue, others determine the area of metastatic lesions relative to normal tissue area. Both of these quantification methods can be exceedingly difficult when the metastatic burden is high. Herein, we demonstrate an intravenous injection model of lung metastasis followed by an advanced method for quantifying metastatic tumor burden using image analysis software. This process allows for investigation of multiple end-point parameters, including average metastasis size, total number of metastases, and total metastasis area, to provide a comprehensive analysis. Furthermore, this method has been reviewed by a veterinary pathologist board-certified by the American College of Veterinary Pathologists (SEK) to ensure accuracy.

Intravenous injection of cancer cells is often used in metastasis research, but the metastatic tumor burden can be difficult to analyze. Herein, we demonstrate a tail-vein injection model of metastasis and include a novel approach to analyze the resulting metastatic lung tumor burden.

Metastasis, the primary cause of morbidity and mortality for most cancer patients, can be challenging to model preclinically in mice. Few spontaneous ...

Modeling Brain Metastases Through Intracranial Injection and Magnetic Resonance Imaging

Metastatic spread of cancer is an unfortunate consequence of disease progression, aggressive cancer subtypes, and/or late diagnosis. Brain metastases are particularly devastating, difficult to treat, and confer a poor prognosis. While the precise incidence of brain metastases in the United States remains hard to estimate, it is likely to increase as extracranial therapies continue to become more efficacious in treating cancer. Thus, it is necessary to identify and develop novel therapeutic approaches to treat metastasis at this site. To this end, intracranial injection of cancer cells has become a well-established method in which to model brain metastasis. Previously, the inability to directly measure tumor growth has been a technical hindrance to this model; however, increasing availability and quality of small animal imaging modalities, such as magnetic resonance imaging (MRI), are vastly improving the ability to monitor tumor growth over time and infer changes within the brain during the experimental period. Herein, intracranial injection of murine mammary tumor cells into immunocompetent mice followed by MRI is demonstrated. The presented injection approach utilizes isoflurane anesthesia and a stereotactic setup with a digitally controlled, automated drill and needle injection to enhance precision, and reduce technical error. MRI is measured over time using a 9.4 Tesla instrument in The Ohio State University James Comprehensive Cancer Center Small Animal Imaging Shared Resource. Tumor volume measurements are demonstrated at each time point through use of ImageJ. Overall, this intracranial injection approach allows for precise injection, day-to-day monitoring, and accurate tumor volume measurements, which combined greatly enhance the utility of this model system to test novel hypotheses on the drivers of brain metastases.

Intracranial brain metastasis modeling is complicated by an inability to monitor tumor size and response to treatment with precise and timely methods. The presented methodology couples intracranial tumor injection with magnetic resonance imaging analysis, which when combined, cultivates precise and consistent injections, enhanced animal monitoring, and accurate tumor volume measurements.

Metastatic spread of cancer is an unfortunate consequence of disease progression, aggressive cancer subtypes, and/or late diagnosis. Brain metastases are ...

Modeling Primary Bone Tumors and Bone Metastasis with Solid Tumor Graft Implantation into Bone

Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions. These lesions compromise bone structure, increase the risk of pathologic fracture, and leave patients with limited treatment options. Primary bone tumors metastasize to distant organs, with some types capable of spreading to other skeletal sites. However, recent evidence suggests that with many solid tumors, cancer cells that have spread to bone may be the primary source of cells that ultimately metastasize to other organ systems. Most syngeneic or xenograft mouse models of primary bone tumors involve intra-osseous (orthotopic) injection of tumor cell suspensions. Some animal models of skeletal metastasis from solid tumors also depend on direct bone injection, while others attempt to recapitulate additional steps of the bone metastatic cascade by injecting cells intravascularly or into the organ of the primary tumor. However, none of these models develop bone metastasis reliably or with an incidence of 100%. In addition, direct intra-osseous injection of tumor cells has been shown to be associated with potential tumor embolization of the lung. These embolic tumor cells engraft but do not recapitulate the metastatic cascade. We reported a mouse model of osteosarcoma in which fresh or cryopreserved tumor fragments (consisting of tumor cells plus stroma) are implanted directly into the proximal tibia using a minimally invasive surgical technique. These animals developed reproducible engraftment, growth, and, over time, osteolysis and lung metastasis. This technique has the versatility to be used to model solid tumor bone metastasis and can readily employ grafts consisting of one or multiple cell types, genetically-modified cells, patient-derived xenografts, and/or labeled cells that can be tracked by optical or advanced imaging. Here, we demonstrate this technique, modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone.

Bone metastasis models do not develop metastasis uniformly or with a 100% incidence. Direct intra-osseous tumor cell injection can result in embolization of the lung. We present our technique modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone, leading to reproducible engraftment and growth.

Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions. These ...

Modeling Brain Metastasis Via Tail-Vein Injection of Inflammatory Breast Cancer Cells

Metastatic spread to the brain is a common and devastating manifestation of many types of cancer. In the United States alone, about 200,000 patients are diagnosed with brain metastases each year. Significant progress has been made in improving survival outcomes for patients with primary breast cancer and systemic malignancies; however, the dismal prognosis for patients with clinical brain metastases highlights the urgent need to develop novel therapeutic agents and strategies against this deadly disease. The lack of suitable experimental models has been one of the major hurdles impeding advancement of our understanding of brain metastasis biology and treatment. Herein, we describe a xenograft mouse model of brain metastasis generated via tail-vein injection of an endogenously HER2-amplified cell line derived from inflammatory breast cancer (IBC), a rare and aggressive form of breast cancer. Cells were labeled with firefly luciferase and green fluorescence protein to monitor brain metastasis, and quantified metastatic burden by bioluminescence imaging, fluorescent stereomicroscopy, and histologic evaluation. Mice robustly and consistently develop brain metastases, allowing investigation of key mediators in the metastatic process and the development of preclinical testing of new treatment strategies.

We describe a xenograft mouse model of breast cancer brain metastasis generated via tail-vein injection of an endogenously HER2-amplified inflammatory breast cancer cell line.

Metastatic spread to the brain is a common and devastating manifestation of many types of cancer. In the United States alone, about 200,000 patients are ...

Portal Vein Injection of Colorectal Cancer Organoids to Study the Liver Metastasis Stroma

Hepatic metastasis of colorectal cancer (CRC) is a leading cause of cancer-related death. Cancer-associated fibroblasts (CAFs), a major component of the tumor microenvironment, play a crucial role in metastatic CRC progression and predict poor patient prognosis. However, there is a lack of satisfactory mouse models to study the crosstalk between metastatic cancer cells and CAFs. Here, we present a method to investigate how liver metastasis progression is regulated by the metastatic niche and possibly could be restrained by stroma-directed therapy. Portal vein injection of CRC organoids generated a desmoplastic reaction, which faithfully recapitulated the fibroblast-rich histology of human CRC liver metastases. This model was tissue-specific with a higher tumor burden in the liver when compared to an intra-splenic injection model, simplifying mouse survival analyses. By injecting luciferase-expressing tumor organoids, tumor growth kinetics could be monitored by in vivo imaging. Moreover, this preclinical model provides a useful platform to assess the efficacy of therapeutics targeting the tumor mesenchyme. We describe methods to examine whether adeno-associated virus-mediated delivery of a tumor-inhibiting stromal gene to hepatocytes could remodel the tumor microenvironment and improve mouse survival. This approach enables the development and assessment of novel therapeutic strategies to inhibit hepatic metastasis of CRC.

Portal vein injection of colorectal cancer (CRC) organoids generates stroma-rich liver metastasis. This mouse model of CRC hepatic metastasis represents a useful tool to study tumor-stroma interactions and develop novel stroma-directed therapeutics such as adeno-associated virus-mediated gene therapies.

Hepatic metastasis of colorectal cancer (CRC) is a leading cause of cancer-related death. Cancer-associated fibroblasts (CAFs), a major component of the ...

In Vivo Immunogenicity Screening of Tumor-Derived Extracellular Vesicles by Flow Cytometry of Splenic T Cells

Immunogenic cell death of tumors, caused by chemotherapy or irradiation, can trigger tumor-specific T cell responses by releasing danger-associated molecular patterns and inducing the production of type I interferon. Immunotherapies, including checkpoint inhibition, primarily rely on preexisting tumor-specific T cells to unfold a therapeutic effect. Thus, synergistic therapeutic approaches that exploit immunogenic cell death as an intrinsic anti-cancer vaccine may improve their responsiveness. However, the spectrum of immunogenic factors released by cells under therapy-induced stress remains incompletely characterized, especially regarding extracellular vesicles (EVs). EVs, nano-scale membranous particles emitted from virtually all cells, are considered to facilitate intercellular communication and, in cancer, have been shown to mediate cross-priming against tumor antigens. To assess the immunogenic effect of EVs derived from tumors under various conditions, adaptable, scalable, and valid methods are sought-for. Therefore, herein a relatively easy and robust approach is presented to assess EVs' in vivo immunogenicity. The protocol is based on flow cytometry analysis of splenic T cells after in vivo immunization of mice with EVs, isolated by precipitation-based assays from tumor cell cultures under therapy or steady-state conditions. For example, this work shows that oxaliplatin exposure of B16-OVA murine melanoma cells resulted in the release of immunogenic EVs that can mediate the activation of tumor-reactive cytotoxic T cells. Hence, screening of EVs via in vivo immunization and flow cytometry identifies conditions under which immunogenic EVs can emerge. Identifying conditions of immunogenic EV release provides an essential prerequisite to testing EVs' therapeutic efficacy against cancer and exploring the underlying molecular mechanisms to ultimately unveil new insights into EVs' role in cancer immunology.

In Vivo Immunogenicity Screening of Tumor-Derived Extracellular Vesicles by Flow Cytometry of Splenic T Cells

This manuscript describes how to assess in vivo immunogenicity of tumor cell-derived extracellular vesicles (EVs) using flow cytometry. EVs derived from tumors undergoing treatment-induced immunogenic cell death seem particularly relevant in tumor immunosurveillance. This protocol exemplifies the assessment of oxaliplatin-induced immunostimulatory tumor EVs but can be adapted to various settings.

Immunogenic cell death of tumors, caused by chemotherapy or irradiation, can trigger tumor-specific T cell responses by releasing danger-associated ...