We would like to thank Michele L. Danielson, Tricia Favors-Watson, and the rest of the Comparative Medicine team at the University of South Florida for their technical support and maintaining our animals. We thank Instech Laboratories, Inc. for their effort working with us based on our request to develop an MRI-compatible Murine Ommaya. This work is supported by the National Institutes of Health (NIH) R21 CA216756 (to K.S.M. Smalley), Department of Defense (DOD) W81XWH1910675 (to B. Czerniecki and P. Kalinski), and the Moffitt Cancer Center CBMM Innovative Awards (to P. Forsyth and D. Duckett). Editorial assistance was provided by the Moffitt Cancer Center&#39;s Office of Scientific Writing by Dr. Paul Fletcher and Daley Drucker. No compensation was given beyond their regular salaries.

The protocol was approved by the University of South Florida Institutional Animal Care and Use Committee (IS00005974).
1. Injection of CTCs into CSF to generate a mouse LMD model
<ol>
	<li>Preparation of CTCs
	<ol>
		<li>Calculate the number of CTCs needed for injection, and prepare a single-cell suspension at 1.0 &#215; 104 cells/&#181;L in sterile phosphate-buffered saline (PBS). Place the cell suspension on ice or at 4 &#176;C throughout the procedure. 
		&#8203;NOTE: When using cell lines that require trypsinization, be sure to wash cells twice with sterile PBS to remove trypsin. Perform cell count using a hemocytometer or an automated cell counter. If a large cohort (&#62;50 mice) is used, recount the cells and validate cell viability between injections to ensure a consistent number of cells are administered per mouse.</li>
	</ol>
	</li>
	<li>Presurgical procedure
	<ol>
		<li>Inject the mouse subcutaneously with 1&#160;mg/kg buprenorphine sustained-release (Bup-SR). 
		NOTE: Do not inject the area of proposed incision; choose an injection site well away from the scapula.</li>
		<li>Anesthetize the mouse with 2-3% isoflurane until it shows no signs of the righting reflex. In addition, check for tail and/or paw pinch reflex to confirm the state of anesthesia.</li>
		<li>Prepare the mouse for surgery in a location remote from the operating area. Clip the surgical site (i.e., the dorsal surface of the skull) with enough border area to keep fur from contaminating the incision site; then, saturate the site with germicidal skin antiseptic, working from the center of the site to the periphery, and then allow to dry. Apply either sterile drape or a sterile adhesive-backed plastic drape material to protect the surgical site from contamination. 
		&#8203;NOTE: Instruments are to be autoclaved in advance and tips re-sterilized between animals using a glass bead sterilizer.</li>
		<li>Shave the fur of the entire ventral surface of the head, and prepare the skin using sterile technique.</li>
		<li>Situate the nose with a modified L-shaped nose cone of the stereotactic apparatus, ensuring the nares remain clear and open. Using tape across the ventral surfaces of both pinnae, gently pull the skin forward to secure it to the nose cone, and bend the neck at an approximately 90&#176; angle once secured. Administer 1.5% isoflurane to maintain anesthesia. 
		&#8203;NOTE: Proper positioning will result in the incision area being presented in a manner that allows for the easy identification of the caudal ridge of the occipital bone.</li>
		<li>Position the body to ensure the spine is kept level with the cisterna magna, and while applying slight traction to the tail, place tape at the tail base to secure.</li>
	</ol>
	</li>
	<li>Surgical cisterna magna injection
	<ol>
		<li>With the neck in full extension, and beginning just between the pinnae, run the surgical scissor tips downward with slight pressure across the occipital bone. 
		NOTE: While in this midline position, a small depression is noticeable when the scissor tips dip into the concave area over the cisterna magna.</li>
		<li>Make a small 3-5 mm midline incision just above the palpated concavity. Draw 5 &#181;L of cell suspension at 1.0 &#215; 104 cells/&#181;L (total 5.0 &#215; 104 cells) into a 30 G Hamilton syringe.</li>
		<li>Use blunt-tipped forceps with 1-2 mm tips to gently press down on the cisterna magna. Introduce tips in a closed position and open them while applying downward pressure on the dura.</li>
		<li>Repeat blunt dissection described in the previous step until the dural membrane is easily identified, and the associated blood vessels are visible in the exposed area. 
		&#8203;NOTE: The resulting injection window ensures blood vessels are undamaged during needle insertion.</li>
		<li>While holding the forceps open to retract surrounding musculature, introduce a 30 G non-coring needle under the dura to visualize the bevel. Ensure the needle is only introduced just beyond the bevel itself. Slowly deploy a syringe plunger, and deliver cells just below the dura.</li>
		<li>Properly place the bevel to observe the injection beneath the dura. Carefully administer the technique, and use the 30 G non-coring needle to prevent damage to the membrane and ensure minimal leakage. If leakage is noted, apply gentle pressure with a cotton-tipped applicator.</li>
		<li>Close the skin by applying a wound clip or micro-drop of skin adhesive. Once the injection is successfully performed, allow the mice to recover from anesthesia on a warm blanket, observing them continuously until they can maintain sternal position and demonstrate purposeful movement.</li>
		<li>Monitor the mice daily after the surgery for the first week. If a mouse appears to be in pain or distress, treat it with 10 mg/kg subcutaneous injection of carprofen once every 12 to 24 h for up to 5 days based on veterinary consultation and directive. Allow mice to recover and monitor them for at least 48 to 72 h before proceeding to the next step. 
		NOTE: Bup-SR, given preemptively, will last up to 72 h, so additional analgesics are not normally necessary. However, animals will be provided additional analgesic as necessary (if lethargic, not eating, ruffled). If a surgical site develops signs of postoperative infection (i.e., redness, swelling, tenderness, allodynia, hyperalgesia, hyperpathia, or suppuration), or if the mouse is guarding the affected area, euthanize the mouse. If cancer cells are successfully injected into the CSF, LMD and tumor progression will develop in the CNS within 1 or 2 weeks (depending on the types of CTCs or cell line used) (Figure 1).</li>
	</ol>
	</li>
</ol>
2. Murine Ommaya assembly and implantation
<ol>
	<li>Preparation of station
	<ol>
		<li>Disinfect the station surface. Place a blue covering over the surface, and keep ready all sterilized tools and supplies indicated in Figure 2.</li>
	</ol>
	</li>
	<li>Presurgical procedure
	<ol>
		<li>Apply analgesia (Bup-SR), and maintain sterile technique as described in section 1.2.</li>
	</ol>
	</li>
	<li>Surgical implantation of Murine Ommaya
	<ol>
		<li>Assemble the Murine Ommaya injection device using a 25 G (0.51 mm outer diameter) miniature injection port and a 1 mm spacer disc. Use a cyanoacrylate sterile adhesive to ensure the penetration of approximately 2.5 mm of the metal cannula into the right cerebral hemisphere (Figure 3A-C). 
		NOTE: An MRI-compatible Mouse Ommaya prototype was developed in this laboratory, which already has both parts (miniature injection port and spacer) 3D-printed together as a single unit (Figure 3D). The single unit version was tested by MRI and can save time by eliminating the assembly step.</li>
		<li>Anesthetize the mouse with 2-3% isoflurane until there are no signs of the righting reflex. Further, check for tail and/or paw pinch reflex to confirm the state of anesthesia.</li>
		<li>Shave the entire ventral surface of the head of fur, and prepare the skin according to the sterile technique, as previously described in step 1.2.3. Place the mouse in the stereotactic apparatus with a nose cone to continue isoflurane administration during the procedure; decrease the isoflurane to 1.5%. Gently tighten the ear bars to secure the head, and apply eye lubricant to to cover the eyes of the mouse.</li>
		<li>Make a small skin incision (3 mm), followed by blunt dissection of the underlying subcutaneous tissues to expose the skull. Dry the skull using hydrogen peroxide-soaked cotton-tipped applicator sticks.</li>
		<li>Drill a burr hole in the skull 0.5 mm posterior and 1.1 mm lateral of the bregma-the anatomical point on the skull where the coronal suture is intersected perpendicularly by the sagittal suture (Figure 3A)-as well as a 0.9 mm burr hole to expose the dura mater. Move the microdrill aside, and gently score the bone immediately surrounding the burr hole prior to inserting an injection port (depth of approximately 2.5 mm), affixed to the skull using a cyanoacrylate sterile adhesive. State suture around the injection spot&#160;using 4-0 no-absorbance nylon sutures in an interrupted stitch pattern or purse string suture11.</li>
		<li>House post-surgery mice in individual cages for surgery recovery. 
		&#8203;NOTE: It is possible that the Murine Ommaya may come off when multiple surgery-recovered mice are housed in the same cage, possibly due to constant tampering interactions. It is recommended that Murine Ommaya-implanted mice are housed individually (or no more than 2 mice per cage) over the course of the drug efficacy trial.</li>
	</ol>
	</li>
</ol>
3. Murine Ommaya treatment
<ol>
	<li>Dosing mice using the Murine Ommaya
	<ol>
		<li>Anesthetize the mouse with 2-3% isoflurane, until there are no signs of righting reflex. Further, check for tail and/or paw pinch reflex to confirm maintenance of anesthesia.</li>
		<li>Access the Murine Ommaya using a port injection adapter and a Hamilton syringe. Using forceps, hold the top of the miniature injection port and gently insert the port injector adaptor fully into the port&#39;s septum. 
		NOTE: The port injector pierces the septum of the port in a manner that reduces dead space to a minimum and allows for controlled injection via a motorized syringe pump (Figure 4A). Targeted/novel treatments are injected at a flow rate of 1 &#181;L/min while under anesthesia. The treatment is to be given at specified intervals (daily/weekly) for a set duration (weeks/months), according to the researcher&#39;s discretion. 
		&#8203;A volume between 3 and 7 &#181;L is optimal, as a volume &#60; 3 &#181;L is unreliable, and a volume &#62; 7 &#181;L may cause too much pressure. The size of the Hamilton syringe can range from 10 to 100 &#181;L, depending on the number of mice in each treatment arm. Preloading the appropriate volume into the Hamilton syringe will prevent repeated replacement of the syringe, thereby minimizing errors. It is best to dedicate 1 Hamilton syringe per treatment arm.</li>
		<li>Once the injection is made, detach the Murine Ommaya from the port injection adaptor using forceps, and return the mouse to the cage to recover from anesthesia, as described above in step 1.3.7.</li>
	</ol>
	</li>
	<li>Euthanasia
	<ol>
		<li>Euthanize the mice by inhalation of carbon dioxide (CO2) from a compressed tank source. Expose the mice to increasing concentrations of CO2 (i.e., a displacement rate from 10% to 30% of the chamber volume/min is to be used) to avoid or minimize discomfort or distress. Monitor the mice until assurance of cessation of cardiovascular and respiratory movements.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Chamberlain, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leptomeningeal+metastasis.">Leptomeningeal metastasis.</a> Current Opinion in Neurology. 22 (6), 665-674 (2009).</li><li>Nayar, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leptomeningeal+disease:+current+diagnostic+and+therapeutic+strategies.">Leptomeningeal disease: current diagnostic and therapeutic strategies.</a> Oncotarget. 8 (42), 73312-73328 (2017).</li><li>Shapiro, W. R., Johanson, C. E., Boogerd, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Treatment+modalities+for+leptomeningeal+metastases.">Treatment modalities for leptomeningeal metastases.</a> Seminars in Oncology. 36 (4), 46-54 (2009).</li><li>Davies, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prognostic+factors+for+survival+in+melanoma+patients+with+brain+metastases.">Prognostic factors for survival in melanoma patients with brain metastases.</a> Cancer. 117 (8), 1687-1696 (2011).</li><li>Znidaric, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Breast+cancer+patients+with+brain+metastases+or+leptomeningeal+disease:+10-year+results+of+a+national+cohort+with+validation+of+prognostic+indexes.">Breast cancer patients with brain metastases or leptomeningeal disease: 10-year results of a national cohort with validation of prognostic indexes.</a> Breast Journal. 25 (6), 1117-1125 (2019).</li><li>Glitza, I. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leptomeningeal+disease+in+melanoma+patients:+An+update+to+treatment,+challenges,+and+future+directions.">Leptomeningeal disease in melanoma patients: An update to treatment, challenges, and future directions.</a> Pigment Cell &amp; Melanoma Research. 33 (4), 527-541 (2020).</li><li>Raizer, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+and+leptomeningeal+metastases+from+cutaneous+melanoma:+survival+outcomes+based+on+clinical+features.">Brain and leptomeningeal metastases from cutaneous melanoma: survival outcomes based on clinical features.</a> Neuro-oncology. 10 (2), 199-207 (2008).</li><li>Taillibert, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leptomeningeal+metastases+from+solid+malignancy:+a+review.">Leptomeningeal metastases from solid malignancy: a review.</a> Journal of Neurooncology. 75 (1), 85-99 (2005).</li><li>DeVos, S. L., Miller, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+intraventricular+delivery+of+drugs+to+the+rodent+central+nervous+system.">Direct intraventricular delivery of drugs to the rodent central nervous system.</a> Journal of Visualized Experiments. (75), e50326 (2013).</li><li>Kodumudi, K. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sequential+anti-PD1+therapy+following+dendritic+cell+vaccination+improves+survival+in+a+HER2+mammary+carcinoma+model+and+identifies+a+critical+role+for+CD4+T+cells+in+mediating+the+response.">Sequential anti-PD1 therapy following dendritic cell vaccination improves survival in a HER2 mammary carcinoma model and identifies a critical role for CD4 T cells in mediating the response.</a> Frontiers in Immunology. 10, 1939 (2019).</li><li>Dunn, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+model+of+wound+healing.">Murine model of wound healing.</a> Journal of Visualized Experiments. (75), e50265 (2013).</li><li>Simon, M. J., Iliff, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+cerebrospinal+fluid+(CSF)+flow+in+neurodegenerative,+neurovascular+and+neuroinflammatory+disease.">Regulation of cerebrospinal fluid (CSF) flow in neurodegenerative, neurovascular and neuroinflammatory disease.</a> Biochimica et Biophysica Acta. 1862 (3), 442-451 (2016).</li><li>Shackleford, G. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continuous+and+bolus+intraventricular+topotecan+prolong+survival+in+a+mouse+model+of+leptomeningeal+medulloblastoma.">Continuous and bolus intraventricular topotecan prolong survival in a mouse model of leptomeningeal medulloblastoma.</a> PLoS One. 14 (1), 0206394 (2019).</li></ol>

Peter Forsyth serves on Advisory Boards for Abvie Inc., Bayer, Bristol Meyers Squib, BTG, Inovio, Novocure, Tocagen, and Ziopharm, outside the submitted work. All other authors have nothing to disclose.

Here, the Murine Ommaya has been described as a reliable model, which allows for repeated administration of anti-cancer agents into the CSF space in preclinical models of LMD and other CNS-related diseases. CSF was sampled from mice while the device was still attached without interruption. This direct-targeted therapy xenograft model is an important step in developing and testing rational treatment strategies for LMD. The time from cisterna magna injections to the initial sign of LMD development varies depending on the cancer cell type. LMD and CNS metastases begin to take form around 1 or 2 weeks after CTC inoculation. If a highly proliferative cancer cell line is used, it is possible that metastasis can occur in &#60;7 days. In this case, vascularization due to tumor growth can sometimes makes implantation of the Murine Ommaya challenging. One solution to this challenge is to reduce the number of cancer cells for injection into the CSF space, allowing for more time before tumor development. Furthermore, this protocol has been optimized to implant the Murine Ommaya no later than 72 h after the cisterna magna injection to ensure enough time for mice to recover from surgery before the first treatment. Researchers should calculate the growth rate of the CTCs in xenograft models before planning the treatment regimen.
Although there are other direct intraventricular delivery methods, such as using an osmotic pump system or an intracerebroventricular (ICV) bolus injection, as described previously9, there are several advantages to using the Murine Ommaya model. For example, an ICV bolus injection is done in a single delivery, whereas the Murine Ommaya allows for multiple doses of treatment at any time, whether given as a single agent or as combined therapies. The osmotic pump is designed to sustain for up to 14, 28, or 42 days before the pump needs replacement, and sometimes more frequently if a smaller mouse with a smaller pump is used. Changing the osmotic pump requires a surgical procedure, which adds stress to tumor-bearing mice. A Murine Ommaya replacement is not necessary for long-duration experiments as long as the device remains intact. It also minimizes potential variability that results from changing the pump9. Implanted Murine Ommayas in the experimental mice remained intact for longer than 42 days, and this duration allowed for long-lived treatment regimens.
Previous findings suggest that a pulsatile intermittent dosing into the CSF has better efficacy against LMD than a prolonged drug delivery process by infusion13. It would be impossible to perform repeated single-dose injections using the osmotic pump system. There is no easy way to flush out the remaining trapped liquid after each injection. The osmotic pump is also limited to delivering mixtures of compatible or single drugs and typically requires higher volumes of drug preparation for continuous infusion. In contrast, the Murine Ommaya is designed for accurate micro-injections as little as 3 to 7&#160;&#181;L, without having to account for dead space, and there is no limitation on the type of drugs researchers can use, including immune cell therapy. The Murine Ommaya also minimizes reagent waste if a particular sample is precious and maximizes the use of that resource. For any treatment regimen that requires multiple doses of anti-cancer therapies, the Murine Ommaya is easy to use, and there is minimal risk of infection or surgical misadventure, with the alternative approaches of repeatedly accessing the CSF surgically or via repeated delivery with a needle. The Murine Ommaya provides researchers with the flexibility to adjust drug concentrations and dosing frequencies and to assess target modulation and duration of the study according to the research of interest.
A limitation of the Murine Ommaya is that researchers may find it difficult to implant the device in smaller mice. Hence, it is better to use mice that are at least 8 to 10 weeks of age. It is possible for the Murine Ommaya to come off during the treatment trial if the device is not secured to the skull during the implantation steps and the glue wears off, or if the mice have abrasively tampered with it. The latter scenario occurs more frequently when multiple mice were housed in the same cage. Hence, it is recommended to house no more than two Murine Ommaya-implanted mice per cage for the duration of the treatment schedule. This protocol was modified to apply cyanoacrylate sterile adhesive on the spacer, which was found to be the most effective glue for adhering the spacer to the skull surface and preventing the Murine Ommaya from coming off. The results showed that LMD mice benefited from direct intrathecal therapy via the Murine Ommaya, with increased overall survival. Single microliter volumes could be safely administered, bypassing the BBB, thereby reducing the amount of drug preparation. Most importantly, the mice that were cured of CNS metastases from the Her2-antibody immunotherapy study have remained healthy.
A collaboration with a manufacturer aimed to develop an MRI-compatible version of the Murine Ommaya for the PDX model of LMD. Because this prototype version has a spacer incorporated, no assembly is required, allowing for better adherence to the skull. A limitation of this prototype is that although the device is MRI-compatible, it generates a shadow where the device is inserted, which decreases the visibility of the image for quantification analyses. The MRI-compatible version is a good alternative tool when ex vivo CTC sampling is a limiting factor, and prelabeling cells is not feasible. The combination of an LMD xenograft model and the Murine Ommaya technique is beneficial for studying direct-targeted drug efficacy bypassing the BBB. The results from these in vivo&#160;studies are clinically relevant to design rational therapeutic strategies for patients with LMD.

Leptomeningeal disease (LMD) is an aggressive late-stage metastasis of the CNS, in which tumor cells access the CSF and infiltrate the surface of the brain and spinal cord1. The most common cancers that cause LMD include those of the breast and lung as well as melanoma2. LMD results in a number of neurologic symptoms and signs such as headaches, cranial nerve palsies, stiff neck, and radiculopathies. The prognosis for patients with LMD is generally very poor (average survival is measured in weeks) and is universally fatal3,4,5,6,7. Treatment with surgery, radiation, and systemic chemotherapy is palliative. Systemic therapy for LMD may fail because of inadequate drug penetration into the CSF across an intact BBB or blood-CSF barrier across the choroid plexus1.
Therefore, administering cancer therapeutics (e.g., drugs and antibody-based treatments including checkpoint inhibitors and cellular therapies) directly into the CSF may overcome this limitation8. Accessing and sampling CSF from patients is possible through an Ommaya reservoir that is implanted beneath the scalp. This device allows for the administration of cancer agents (e.g., methotrexate and trastuzumab) as well as the sampling of CSF for diagnostic studies (e.g., the cytological diagnosis of LMD to monitor for responses to treatment) without performing a spinal tap. A murine Ommaya reservoir was designed to mimic those used clinically. The reservoir requires the assembly of an accessing port and spacer parts and a modification of the mouse cannulation technique, which allows the device to remain permanently intact throughout the duration of the drug study. This device has been designated as the &#34;Murine Ommaya&#34;.
In contrast to the osmotic infusion pump technique, which requires the preparation of excess liquid volumes to prefill the empty space in the tubing and continuous infusion over frequent injections9, the Murine Ommaya minimizes the wastage of drug solutions. It permits effective administration of multiple single doses of treatments at any given time in small quantities (3-7 &#181;L) into the CSF using a Hamilton syringe, a miniature access port, and an automatic injector. In real time, the efficacy of test drugs against LMD can be determined by imaging. Using this approach, a variety of chemotherapies, antibodies, and cell immunotherapies (as single or combined agents) can be tested against LMD to translate in vivo&#160;findings into rational treatment strategies for patients. To further improve the imaging capacity for a patient-derived xenograft (PDX) model of LMD, a collaboration was undertaken with a manufacturer to develop a magnetic resonance imaging (MRI)-compatible version of the Murine Ommaya, which requires no assembly and is ready to use. MRI capability is beneficial, especially for PDX models in which the quantity of circulating tumor cells (CTCs) from CSF is sometimes the limiting factor, and often when prelabeling CTCs is infeasible.
This paper describes a detailed protocol that begins with the injection of CTCs to render mice with LMD . The Murine Ommaya is then surgically implanted, and multiple drug treatment steps via the Murine Ommaya are performed. As a proof of concept for demonstration, an in vivo side-by-side comparison was performed, in which the murine human epidermal growth factor receptor 2 (Her2) antibody called clone 7.16.4 (the human equivalent of trastuzumab) was delivered10. The antibody targets Her2+ breast cancer cells either via the Murine Ommaya (direct-targeted or intrathecal therapy) or by intraperitoneal injection (systemic therapy). The results showed that mice with LMD that received direct intrathecal immunotherapy lived significantly longer than those treated with the same therapy systemically. The CNS metastases in mice treated via the Murine Ommaya were almost completely regressed by the third dose of the third week of treatment, resulting in improved overall survival.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mm spacer disc</td>
			<td>Alzet, Durect Corporation</td>
			<td>#0008670</td>
			<td>Spacer disc only</td>
		</tr>
		<tr>
			<td>4-0 ethilon nylon suture</td>
			<td>Any vendor</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Automatic syringe pumps</td>
			<td>Harvard Syringe Pumps (or any vendor)</td>
			<td>#70-4505</td>
			<td>Pump 11 Elite</td>
		</tr>
		<tr>
			<td>Bead sterilizer</td>
			<td>Braintree Scientific Inc. (or any vendor)</td>
			<td>#GER 5287-120V</td>
			<td>Germinator 500</td>
		</tr>
		<tr>
			<td>Buprenorphine Sustained-Release (Bup-SR)</td>
			<td>Zoopharm</td>
			<td></td>
			<td>DEA controlled</td>
		</tr>
		<tr>
			<td>Cyanoacrylate sterile adhesive</td>
			<td>Any vendor</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gas inhalation anestehsia system</td>
			<td>VeteEquip</td>
			<td>#901812</td>
			<td>COMPAC5</td>
		</tr>
		<tr>
			<td>Hamilton microliter syringes</td>
			<td>Hamilton</td>
			<td>10, 25, 50, and 100 &#956;L</td>
			<td>30 G for cisterna magna injection</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide</td>
			<td>Any vendor</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>IVIS 200 imaging system</td>
			<td>Caliper Life Sciences</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnifying glass with light</td>
			<td>Any vendor</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Microdrill</td>
			<td>Stoelting (or any vendor)</td>
			<td>#51555M</td>
			<td></td>
		</tr>
		<tr>
			<td>MRI imaging</td>
			<td>Bruker</td>
			<td>BioSpec series</td>
			<td>Optional</td>
		</tr>
		<tr>
			<td>Murine Ommaya (MRI-compatible) prototype</td>
			<td>Instech Laboratories, Inc.</td>
			<td>#VAB620-25MRI-3.3</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS)</td>
			<td>Any vendor</td>
			<td>n/a</td>
			<td>0.1 mm Sterile-Filtered</td>
		</tr>
		<tr>
			<td>PinPort injector</td>
			<td>Instech Laboratories, Inc.</td>
			<td>#PNP3M-50</td>
			<td></td>
		</tr>
		<tr>
			<td>PinPort</td>
			<td>Instech Laboratories, Inc.</td>
			<td>#1-PNP3F28-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Rodent Surgical Instruments (Scissors, Forceps)</td>
			<td>Roboz Surgical Instrument (or any vendor)</td>
			<td></td>
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		</tr>
		<tr>
			<td>Stereotaxic device</td>
			<td>Stoelting (or any vendor)</td>
			<td>#51730M</td>
			<td></td>
		</tr>
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			<td>Sterile blue paper/ drape covering</td>
			<td>Any vendor</td>
			<td>n/a</td>
			<td>n/a</td>
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			<td>Sterile cotton sticks</td>
			<td>Any vendor</td>
			<td>n/a</td>
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	</tbody>
</table>

a murine ommaya xenograft model to study direct-targeted therapy of leptomeningeal disease

Leptomeningeal disease (LMD) is an uncommon type of central nervous system (CNS) metastasis to the cerebral spinal fluid (CSF). The most common cancers that cause LMD are breast and lung cancers and melanoma. Patients diagnosed with LMD have a very poor prognosis and generally survive for only a few weeks or months. One possible reason for the lack of efficacy of systemic therapy against LMD is the failure to achieve therapeutically effective concentrations of drug in the CSF because of an intact and relatively impermeable blood-brain barrier (BBB) or blood-CSF barrier across the choroid plexus. Therefore, directly administering drugs intrathecally or intraventricularly may overcome these barriers. This group has developed a model that allows for the effective delivery of therapeutics (i.e., drugs, antibodies, and cellular therapies) chronically and the repeated sampling of CSF to determine drug concentrations and target modulation in the CSF (when the tumor microenvironment is targeted in mice). The model is the murine equivalent of a magnetic resonance imaging-compatible Ommaya reservoir, which is used clinically. This model, which is affixed to the skull, has been designated as the &#34;Murine Ommaya.&#34; As a therapeutic proof of concept, human epidermal growth factor receptor 2 antibodies (clone 7.16.4) were delivered into the CSF via the Murine Ommaya to treat mice with LMD from human epidermal growth factor receptor 2-positive breast cancer. The Murine Ommaya increases the efficiency of drug delivery using a miniature access port and prevents the wastage of excess drug; it does not interfere with CSF sampling for molecular and immunological studies. The Murine Ommaya is useful for testing novel therapeutics in experimental models of LMD.

In mice, the total volume of CSF is approximately 35-40 &#181;L and is produced at a rate of about 350 nL/min; it turns over 12-13 times a day12. For the purpose of visualizing the route of the injection, 2% Evans Blue was injected via the Murine Ommaya model, following which 15 min and 30 min were allowed to elapse before harvesting the brains for analysis. The dye successfully infiltrated the ventricles and the brain in 15 min. Within 30 min, the dye became visible on the spinal cord (Figure 4).
As a proof of concept, BALB/c mice were injected with a luciferase-labeled Her2+ TUBO breast cancer cell line intracisternally, and the Murine Ommayas were implanted. Approximately 1 week after the injection of cancer cells, the mice began to develop LMD. These mice were treated once a week for up to 4 weeks with Her2-antibody immunotherapy, either through systemic therapy via intraperitoneal injection or intrathecally via the Murine Ommaya (Figure 5A).
Although untreated mice died by day 19, all mice that received intrathecal therapy through the Murine Ommaya survived (P = 0.004). By week 4, a complete regression of tumors was observed. In comparison to mice treated with systemic therapy, which had moderate success in treating LMD, mice that received intrathecal therapy had a much longer overall survival (Figure 5B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62033/62033fig01.jpg" /> 
Figure 1: Injection of circulating tumor cells into the cisterna magna in a murine xenograft model to study leptomeningeal disease and central nervous system metastases. (A) An illustration showing the location of the cisterna magna and the CSF accessing site, in which CTCs are injected using a Hamilton syringe. (B)&#160;A representative IVIS image of mice that had developed leptomeningeal disease and central nervous system metastases (brain and along the spinal cord) after 2 weeks of injection with circulating tumor cells. Cells were labeled with a luciferase reporter gene. Control animals injected with saline did not develop tumors (n = 3), and the experiment was performed in triplicate. Abbreviations: CTCs = circulating tumor cells; IVIS = in vivo imaging system. <a href="https://www.jove.com/files/ftp_upload/62033/62033fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62033/62033fig02.jpg" /> 
Figure 2: An example of a workstation setup for performing the Murine Ommaya implantation in mice. (1) Gas anesthesia machine/vaporizer. (2) Sterile blue paper drape covering a stereotaxic stand. (3) Stereotaxic device (stand/stage, ear bars, nose cone). (4) Microdrill. (5) Magnifying glass with light. (6) Sterile cotton tapped applicator sticks with sterile saline rinse container. (7) Hydrogen peroxide. (8) Bead sterilizer. <a href="https://www.jove.com/files/ftp_upload/62033/62033fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62033/62033fig03.jpg" /> 
Figure 3: The implantation of the Murine Ommaya device. (A)&#160;An illustration with an arrow pointing at the location of the bregma on the skull, and the approximate distance at which a burr hole is drilled in the skull (0.5 mm posterior/1.1 mm lateral) from the bregma using a microdrill. (B) A Murine Ommaya is assembled by combining a metal cannula and a 1 mm spacer as the base for glue attachment to the skull. (C)&#160;Representative images of mice that had Murine Ommayas implanted; these mice are monitored to ensure they are bright, alert, and reactive before receiving any injections. (D) An example of the prototype magnetic resonance imaging-compatible Murine Ommaya and representative brain magnetic resonance imaging images of Murine Ommaya implants. <a href="https://www.jove.com/files/ftp_upload/62033/62033fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62033/62033fig04.jpg" /> 
Figure 4: Intraventricular (central nervous system) injection using the Murine Ommaya. (A) An image of an injection, accessing the ventricle and central nervous system via the Murine Ommaya. Mice remain under anesthesia during the injection. In the example, the Murine Ommaya is connected to the miniature port that is attached to a prefilled Hamilton syringe. Injections are performed using an automatic injection set at an infusion rate of 1 &#181;L/min and a volume of 5-7 &#181;L. An image of a mouse brain injected with Evans Blue is shown. The circle shows where the Murine Ommaya was attached. No leakage of the dye was observed on the exterior of the brain. A cross section of the brain shows the lateral ventricles were filled with the dye; the dye did not penetrate brain parenchyma. (B) Images of mouse brains after 15 and 30 min following the injection of Evans Blue dye. The dye infiltrated the brain (15 min) and began to circulate on the spinal cord (30 min). Out of 5 mice, 4 received dye for visualization, and 1 served as control. The experiment was repeated in triplicate. <a href="https://www.jove.com/files/ftp_upload/62033/62033fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62033/62033fig05.jpg" /> 
Figure 5: Direct-targeted immunotherapy using the Murine Ommaya increases the overall survival of breast cancer-associated leptomeningeal disease mice. (A)&#160;BALB/c mice were injected with luciferase reporter-labeled human epidermal growth factor receptor 2-positive TUBO cells, a murine breast cancer cell line. Three days after cisterna magna injections, Murine Ommayas were implanted. Mice began to develop leptomeningeal disease (LMD) 1 week after injection. LMD mice were treated with either a human epidermal growth factor receptor antibody systemically via intraperitoneal injection or via intrathecal (Murine Ommaya) as a direct-targeted approach. Injections were given once a week for up to 4 weeks. Compared to untreated mice, mice receiving immunotherapy survived much longer. Murine Ommaya mice had complete disease regression by the fourth week, and these mice were eventually cured of disease. (B)&#160;These mice also had significantly better median survival (Mantel-Cox test; P = 0.004; n = 5 mice per treatment arm) and better overall survival than systematically treated LMD mice. Abbreviations: LMD = leptomeningeal disease; IP = intraperitoneal.&#160;<a href="https://www.jove.com/files/ftp_upload/62033/62033fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease at JoVE.com

A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease

Here, we describe a murine xenograft model that functionally resembles an Ommaya reservoir in patients. We developed the Murine Ommaya to study novel therapeutics for the universally fatal leptomeningeal disease.

Leptomeningeal disease (LMD) is an uncommon type of central nervous system (CNS) metastasis to the cerebral spinal fluid (CSF). The most common cancers ...

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Cancer Research

5.2K Views.  H. Lee Moffitt Cancer Center & Research Institute. Here, we describe a murine xenograft model that functionally resembles an Ommaya reservoir in patients. We developed the Murine Ommaya to study novel therapeutics for the universally fatal leptomeningeal disease.

Video: A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease

Development and Maintenance of a Preclinical Patient Derived Tumor Xenograft Model for the Investigation of Novel Anti-Cancer Therapies

Patient derived tumor xenograft (PDTX) models provide a necessary platform in facilitating anti-cancer drug development prior to human trials. Human tumor pieces are injected subcutaneously into athymic nude mice (immunocompromised, T cell deficient) to create a bank of tumors and subsequently are passaged into different generations of mice in order to maintain these tumors from patients. Importantly, cellular heterogeneity of the original tumor is closely emulated in this model, which provides a more clinically relevant model for evaluation of drug efficacy studies (single agent and combination), biomarker analysis, resistant pathways and cancer stem cell biology. Some limitations of the PDTX model include the replacement of the human stroma with mouse stroma after the first generation in mice, inability to investigate treatment effects on metastasis due to the subcutaneous injections of the tumors, and the lack of evaluation of immunotherapies due to the use of immunocompromised mice. However, even with these limitations, the PDTX model provides a powerful preclinical platform in the drug discovery process.

Utilizing patient-derived tumors in a subcutaneous preclinical model is an excellent way to study the efficacy of novel therapies, predictive biomarker discovery, and drug resistant pathways. This model, in the drug development process, is essential in determining the fate of many novel anti-cancer therapies prior to clinical investigation.

Patient derived tumor xenograft (PDTX) models provide a necessary platform in facilitating anti-cancer drug development prior to human trials. Human tumor ...

The Colon-26 Carcinoma Tumor-bearing Mouse as a Model for the Study of Cancer Cachexia

Cancer cachexia is the progressive loss of skeletal muscle mass and adipose tissue, negative nitrogen balance, anorexia, fatigue, inflammation, and activation of lipolysis and proteolysis systems. Cancer patients with cachexia benefit less from anti-neoplastic therapies and show increased mortality1. Several animal models have been established in order to investigate the molecular causes responsible for body and muscle wasting as a result of tumor growth. Here, we describe methodologies pertaining to a well-characterized model of cancer cachexia: mice bearing the C26 carcinoma2-4. Although this model is heavily used in cachexia research, different approaches make reproducibility a potential issue. The growth of the C26 tumor causes a marked and progressive loss of body and skeletal muscle mass, accompanied by reduced muscle cross-sectional area and muscle strength3-5. Adipose tissue is also lost. Wasting is coincident with elevated circulating levels of pro-inflammatory cytokines, particularly Interleukin-6 (IL-6)3, which is directly, although not entirely, responsible for C26 cachexia. It is well-accepted that a primary mechanism by which the C26 tumor induces muscle tissue depletion is the activation of skeletal muscle proteolytic systems. Thus, expression of muscle-specific ubiquitin ligases, such as atrogin-1/MAFbx and MuRF-1, represent an accepted method for the evaluation of the ongoing muscle catabolism2. Here, we present how to execute this model in a reproducible manner and how to excise several tissues and organs (the liver, spleen, and heart), as well as fat and skeletal muscles (the gastrocnemius, tibialis anterior, and quadriceps). We also provide useful protocols that describe how to perform muscle freezing, sectioning, and fiber size quantification.

Mice bearing the Colon-26 (C26) carcinoma represent a classical model of cancer cachexia. Progressive muscle wasting occurs in association with tumor growth, over-expression of muscle-specific ubiquitin ligases, and reductions in muscle cross-sectional area. Fat loss is also observed. Cachexia is studied in a time-dependent manner with increasing severity of wasting.

Cancer cachexia is the progressive loss of skeletal muscle mass and adipose tissue, negative nitrogen balance, anorexia, fatigue, inflammation, and ...

A Mouse Model of Fatigue Induced by Peripheral Irradiation

Cancer-related fatigue (CRF) is a distressing and costly condition that often affects patients receiving cancer treatments, including radiation therapy. Here we describe a method using targeted peripheral irradiation to induce fatigue-like behavior in mice. With appropriate shielding, the irradiation targets the lower abdominal/pelvic region of the mouse, sparing the brain, in an effort to model radiation treatment received by individuals with pelvic cancers. We deliver an irradiation dose that is sufficient to induce fatigue-like behavior in mice, measured by voluntary wheel-running activity (VWRA), without causing obvious morbidity. Since wheel running is a normal, voluntary behavior in mice, its use should have little confounding effect on other behavioral tests or biological measures. Hence, wheel running can be used as a feasible outcome measure in understanding the behavioral and biological correlates of fatigue. CRF is a complex condition with frequent comorbidities, and likely has causes related both to cancer and its various treatments. The methods described in this paper are useful for investigating radiation-induced changes that contribute to the development of CRF and, more generally, to explore the biological networks that can explain the development and persistence of a peripherally-triggered but centrally-driven behavior like fatigue.

We describe a method using targeted peripheral irradiation to induce fatigue-like behavior in mice. The selected non-lethal irradiation dose leads to a week-long reduction in voluntary wheel-running activity.

Cancer-related fatigue (CRF) is a distressing and costly condition that often affects patients receiving cancer treatments, including radiation therapy. ...

Tumor Engraftment in a Xenograft Mouse Model of Human Mantle Cell Lymphoma

B lymphocytes are key players in immune cell circulation and they mainly home to and reside in lymphoid organs. While normal B cells only proliferate when stimulated by T lymphocytes, oncogenic B cells survive and expand autonomously in undefined organ niches. Mantle cell lymphoma (MCL) is one such B cell disorder, where the median survival rate of patients is 4 - 5 years. This calls for the need of effective mechanisms by which the homing and engraftment of these cells are blocked in order to increase the survival and longevity of patients. Therefore, the effort to develop a xenograft mouse model to study the efficacy of MCL therapeutics by blocking the homing mechanism in vivo is of utmost importance. Development of animal recipients for human cell xenotransplantation to test early stage drugs have long been pursued, as relevant preclinical mouse models are crucial to screen new therapeutic agents. This animal model is developed to avoid human graft rejection and to establish a model for human diseases, and it may be an extremely useful tool to study disease progression of different lymphoma types and to perform preclinical testing of candidate drugs for hematologic malignancies, like MCL. We established a xenograft mouse model that will serve as an excellent resource to study and develop novel therapeutic approaches for MCL.

Mantle cell lymphoma (MCL) is a difficult to treat B cell disorder and it is equally difficult to establish a xenograft mouse model of primary MCL to study and develop therapeutics. Here, we describe the successful establishment of MCL xenografts in mice to help understand its underlying biology.

B lymphocytes are key players in immune cell circulation and they mainly home to and reside in lymphoid organs. While normal B cells only proliferate when ...

A Melanoma Patient-Derived Xenograft Model

Accumulating evidence suggests that molecular and biological properties differ in melanoma cells grown in traditional two-dimensional tissue culture vessels versus in vivo in human patients. This is due to the bottleneck selection of clonal populations of melanoma cells that can robustly grow in vitro in the absence of physiological conditions. Further, responses to therapy in two-dimensional tissue cultures overall do not faithfully reflect responses to therapy in melanoma patients, with the majority of clinical trials failing to show the efficacy of therapeutic combinations shown to be effective in vitro. Although xenografting of melanoma cells into mice provides the physiological in vivo context absent from two-dimensional tissue culture assays, the melanoma cells used for engraftment have already undergone bottleneck selection for cells that could grow under two-dimensional conditions when the cell line was established. The irreversible alterations that occur as a consequence of the bottleneck include changes in growth and invasion properties, as well as the loss of specific subpopulations. Therefore, models that better recapitulate the human condition in vivo may better predict therapeutic strategies that effectively increase the overall survival of patients with metastatic melanoma. The patient-derived xenograft (PDX) technique involves the direct implantation of tumor cells from the human patient to a mouse recipient. In this manner, tumor cells are consistently grown under physiological stresses in vivo and never undergo the two-dimensional bottleneck, which preserves the molecular and biological properties present when the tumor was in the human patient. Notable, PDX models derived from organ sites of metastases (i.e., brain) display similar metastatic capacity, while PDX models derived from therapy naive patients and patients with acquired resistance to therapy (i.e., BRAF/MEK inhibitor therapy) display similar sensitivity to therapy.

Patient-derived xenograft (PDX) models more robustly recapitulate melanoma molecular and biological features and are more predictive of therapy response compared to traditional plastic tissue culture-based assays. Here we describe our standard operating protocol for the establishment of new PDX models and the characterization/experimentation of existing PDX models.

Accumulating evidence suggests that molecular and biological properties differ in melanoma cells grown in traditional two-dimensional tissue culture ...

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 ...

Intra-Cardiac Injection of Human Prostate Cancer Cells to Create a Bone Metastasis Xenograft Mouse Model

As the most common male malignancy, prostate cancer (PC) ranks second in mortality, primarily due to a 65%-75% bone metastasis rate. Therefore, it is essential to understand the process and related mechanisms of prostate cancer bone metastasis for developing new therapeutics. For this, an animal model of bone metastasis is an essential tool. Here, we report detailed procedures to generate a bone metastasis mouse model via intra-cardiac injection of prostate cancer cells. A bioluminescence imaging system can determine whether prostate cancer cells have been accurately injected into the heart and monitor cancer cell metastasis since it has great advantages in monitoring metastatic lesion development. This model replicates the natural development of disseminated cancer cells to form micro-metastases in the bone and imitates the pathological process of prostate cancer bone metastasis. It provides an effective tool for further exploration of the molecular mechanisms and the in vivo therapeutic effects of this disease.

Here, we present a protocol for the intra-cardiac injection of human prostate cancer cells to generate a mouse model with bone metastasis lesions.

As the most common male malignancy, prostate cancer (PC) ranks second in mortality, primarily due to a 65%-75% bone metastasis rate. Therefore, it is ...

Spontaneous Murine Model of Anaplastic Thyroid Cancer

Anaplastic thyroid cancer (ATC) is a rare but lethal malignancy with a dismal prognosis. There is an urgent need for more in-depth research on the carcinogenesis and development of ATC, as well as therapeutic methods, since standard treatments are essentially depleted in ATC patients. However, low prevalence has hampered thorough clinical studies and the collection of tissue samples, so little progress has been achieved in creating effective treatments. We used genetic engineering to create a conditionally inducible ATC murine model (mATC) in a C57BL/6 background. The ATC murine model was genotyped by TPO-cre/ERT2; BrafCA/wt; Trp53ex2-10/ex2-10 and induced by intraperitoneal injection with tamoxifen. With the murine model, we investigated the tumor dynamics (tumor size ranged from 12.4 mm2 to 32.5 mm2 after 4 months of induction), survival (the median survival period was 130 days), and metastasis (lung metastases occurred in 91.6% of mice) curves and pathological features (characterized by Cd8, Foxp3, F4/80, Cd206, Ki67, and Caspase-3 immunohistochemical staining). The results indicated that spontaneous mATC possesses highly similar tumor dynamics and immunological microenvironment to human ATC tumors. In conclusion, with high similarity in pathophysiological features and unified genotypes, the mATC model resolved the shortage of clinical ATC tissue and sample heterogeneity to some extent. Therefore, it would facilitate the mechanism and translational studies of ATC and provide an approach to investigate the treatment potential of small molecular drugs and immunotherapy agents for ATC.

Here, we present a standard pipeline to obtain murine ATC tumors by spontaneous genetically engineered mouse models. Further, we present tumor dynamics and pathological information about the primary and metastasized lesions. This model will help researchers to understand tumorigenesis and facilitate drug discoveries.

Anaplastic thyroid cancer (ATC) is a rare but lethal malignancy with a dismal prognosis. There is an urgent need for more in-depth research on the ...

Patient&#45;Derived Hepatocarcinoma Cell Organoids for Liver Regeneration Study

A Hepatocellular Cancer Patient&#45;Derived Organoid Xenograft Model to Investigate Impact of Liver Regeneration on Tumor Growth

Recurrence poses a notable challenge after hepatocellular carcinoma (HCC) treatment, impacting more than 70% of patients who undergo surgical resection. Recurrence stems from undetected micro-metastasis or de novo cancer, potentially triggered by postsurgical liver regeneration. Prior research employed HCC cell lines in orthotopic models to study the impact of liver regeneration, but their limited validity prompted the need for a more representative model. Here, we introduce a novel approach utilizing patient-derived HCC organoids to investigate the influence of liver regeneration on HCC.
Patient tumor tissues are processed to create tumor organoids, embedded in a three-dimensional basement membrane matrix, and cultured in a liver-specific medium. One million organoids are injected into the right superior lobe (RSL) of immunodeficient mice, confirming macroscopic tumor growth through sonography. The intervention group undergoes resection of the left lateral lobe (LLL) (30% of total liver volume) or additionally, the middle lobe (ML) (65% of total liver volume) to induce liver regeneration within the tumor site. The control group experiences re-laparotomy without liver tissue resection. After 2 weeks, both groups undergo tumor and normal tissue explantation.
In conclusion, this patient-derived HCC organoid model offers a robust platform to investigate the impact of liver regeneration post-cancer resection. Its multi-cellular composition, genetic diversity, and prolonged culture capabilities make it an invaluable tool for studying HCC recurrence mechanisms and potential interventions.

Organoids generated from patient tumors are orthotopically injected into the mouse liver. The resection of non-tumorous liver tissue leads to a regenerative environment in the liver tissue where the tumor is located.

Recurrence poses a notable challenge after hepatocellular carcinoma (HCC) treatment, impacting more than 70% of patients who undergo surgical resection. ...

Protocol for Osteosarcoma PDX Mouse Models: Studying Growth and Drug Resistance

Establishment of Patient-Derived Xenograft Mouse Model with Human Osteosarcoma Tissues

Osteosarcoma is the most common primary malignant bone tumor in children and adolescents. Despite the development of new treatment plans in recent years, the prognosis for osteosarcoma patients has not significantly improved. Therefore, it is crucial to establish a robust preclinical model with high fidelity. The patient-derived xenograft (PDX) model faithfully preserves the genetic, epigenetic, and heterogeneous characteristics of human malignancies for each patient. Consequently, PDX models are considered authentic in vivo models for studying various cancers in transformation studies. This article presents a comprehensive protocol for creating and maintaining a PDX mouse model that accurately mirrors the morphological features of human osteosarcoma. This involves the immediate transplantation of freshly resected human osteosarcoma tissue into immunocompromised mice, followed by successive passaging. The described model serves as a platform for studying the growth, drug resistance, relapse, and metastasis of osteosarcoma. Additionally, it aids in screening the target therapeutics and establishing personalized treatment schemes.

Author Spotlight: Replicating Human Osteosarcoma Progression in Immunodeficient Mice for Cancer Study

The present protocol describes the method for establishing a patient-derived xenograft (PDX) mouse model using human osteosarcoma tissue.

Osteosarcoma is the most common primary malignant bone tumor in children and adolescents. Despite the development of new treatment plans in recent years, ...