This work is supported by grants from the Sock it to Sarcoma! Foundation, the Australian and New Zealand Sarcoma Association, the Children&#39;s Leukemia &#38; Cancer Research Foundation and Perpetual Philanthropy. W.J.L is supported by a Simon Lee Fellowship and a research fellowship from the National Health and Medical Research Council, and the Cancer Council WA.

Animals used in these experiments were obtained from the Animal Resource Centre (Perth, Western Australia). Animals were maintained under standard pathogen-free conditions at the Harry Perkins Institute of Medical Research Bioresources North Facility (Perth, Western Australia). All experiments were carried out following the protocol as approved by the Harry Perkins Institute of Medical Research Animal Ethics Committee. BALB/c mice of 8-12 weeks of age were used in these experiments. The WEHI 164 fibrosarcoma cell line was obtained from CellBank Australia (Westmead, NSW).
1. Inoculation of cells
<ol>
	<li>Preparation of cells and animals
	<ol>
		<li>Ensure that the cell line is maintained in the recommended media. For example, maintain WEHI 164 cell line in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 2 mM L-glutamine, 10% fetal bovine serum, 20 mM HEPES, 0.05 mM 2-mercaptoethanol, 100 U/mL penicillin, and 100 &#181;g/mL streptomycin. 
		NOTE: Passage cells at least 3 and up to 5 times after being removed from cryogenic storage. To ensure an optimum cell viability, cells should be split when they are between 70-80% confluent. Tumor cell lines should be tested for mycoplasma, as infection can alter the cell growth and influence the immune response in vivo.</li>
		<li>One day before inoculation, shave mice on the lower right flank using clippers. 
		NOTE: Female BALB/c mice, aged between 8-12 weeks, of normal weight (16 -22 grams) were used in this experiment.</li>
		<li>On the day of inoculation, harvest WEHI 164 cells when 70-80% confluent by trypsinization.
		<ol>
			<li>Aspirate the culture medium from the tissue culture flasks and then add sterile phosphate buffered solution (1x PBS), to remove remaining traces of fetal bovine serum (FBS).</li>
			<li>Aspirate the PBS from the tissue culture flasks. Add 3 mL of 0.05% trypsin (for a T75 flask) and then swirl the flask so that the whole surface of flask with cells is covered by trypsin.</li>
			<li>Incubate the flask at 37 &#176;C, 5% CO2 incubator for 3 min. Check cells periodically, by tapping on the sides of the flask to see if cells have dislodged.</li>
			<li>Remove flasks from cell culture incubator and add 5 mL of media supplemented with FBS to neutralize the trypsin. 
			NOTE: Do not leave cells in trypsin longer than necessary, as this can damage cells and lead to low cell viability.</li>
			<li>Pipet suspension multiple times to obtain a single cell suspension. Transfer cell suspension to a conical centrifuge tube.</li>
			<li>Pellet cells by spinning at 350 x g for 3 min.</li>
		</ol>
		</li>
		<li>Wash the cells three times in 1x PBS.
		<ol>
			<li>Resuspend cells in 50 mL of sterile 1x PBS and wash cells by pipetting cell suspension up and down. Pellet cells by spinning at 350 x g for 3 min.</li>
			<li>Aspirate the supernatant and resuspend cells in 15 mL of sterile 1x PBS. Wash cells by pipetting cell suspension up and down. Pellet cells by spinning at 350 x g for 3 min.</li>
			<li>Aspirate the supernatant and resuspend cells in exactly 10 mL of sterile 1x PBS. Wash cells as in step 1.1.4.2 and transfer a small amount (approximately 100 &#181;L) of cell suspension to an centrifuge tube for counting. Pellet cells by spinning at 350 x g for 3 min.</li>
		</ol>
		</li>
		<li>Determine the cell number using the Trypan blue exclusion method by either using a hemocytometer or an automated cell counter. Resuspend cells in sterile 1x PBS at a concentration of 5 x 106 cells/mL. Keep cell suspension on ice. 
		NOTE: The viability of tumor cells should be equal or above 80 % to ensure reproducible tumor growth.</li>
	</ol>
	</li>
	<li>Subcutaneous inoculation
	<ol>
		<li>Mix the cell suspension thoroughly and fill a syringe with a 26 G needle with 100 &#181;L of cell suspension (5 x 105 cells) in sterile 1x PBS. Repeat mixing of cells before loading the next syringe. 
		NOTE: Keep cells on ice throughout the procedure to maintain viability.</li>
		<li>Restrain the mouse appropriately, ensuring access to the lower-right flank. Inoculate the mouse subcutaneously on the shaved lower-right flank. 
		NOTE: Make sure the inoculation is not in the peritoneum by lifting the needle slightly, which should be visible under the skin. A bubble-like lump should form under the skin following inoculation.</li>
		<li>Monitor mice as required by the applicable ethics approval and perform surgical resection when the tumours have grown to a size of about 50 mm2.</li>
	</ol>
	</li>
</ol>
2. Partial surgical resection of the tumor
NOTE: This protocol requires TWO researchers; one for surgical procedures (SURGEON), and another for mouse monitoring (ASSISTANT).
<ol>
	<li>Surgery setup
	<ol>
		<li>On day 12 post inoculation, when tumors have reached a size of approximately 50 mm2, dose mice with 100 &#181;L (0.1 mg/kg) of buprenorphine s.c. in the scruff of the neck, 30 minutes prior to surgery.</li>
		<li>Set up the surgical area with a heat pad covered with bench coat and set up a nose cone for anaesthesia. Sterilize surgical tools prior to use, and between each animal using a heat bead sterilizer, allowing tools to cool before use.&#160;Have the following surgical equipment clean and within easy reach: chlorhexidine, swab, gauze, eye gel, two curved forceps, scissors, clip applicator, clip remover, clip refills (Figure 2A, 2B).</li>
		<li>Warm the heating chamber to 37 &#176;C and set up another heat pad for recovery (Figure 2C).&#160;Place sterilized tools on a sterile surface such as autoclaved pads.</li>
	</ol>
	</li>
	<li>Anesthesia
	<ol>
		<li>Place the mouse in the induction chamber and anesthetize the mouse with 4% isoflurane (4% in 100% oxygen at a flow rate of 1 L/min) until the breathing rate slows to approximately 60 breaths per minute (1 per second) (this usually takes &#60;1 min). 
		NOTE: Do not leave the mouse in the chamber for too long as that may lead to asphyxiation and death. Only have one mouse under anesthesia at a time.</li>
		<li>Transfer the mouse onto the heat pad on the surgery table, place the mouse with its nose in the nose cone and maintain the anesthetic state with 3-4% isoflurane in 100% oxygen at a flow rate of 0.5 L/min. Monitor the breathing rate to ensure that the depth of anesthesia is maintained. 
		NOTE: The ASSISTANT must monitor the breathing of the mouse throughout the surgery to ensure the correct level of anesthesia is maintained. Lower the anesthetic concentration if breathing becomes too slow or increase the concentration if the depth of anesthesia is too shallow. If the mouse begins gasping, remove mouse from the nose cone, decrease the anesthetic concentration, and wait until breathing normalises before placing on the nose cone again.</li>
		<li>Perform a &#34;pinch test&#34; and &#34;corneal reflex test&#34;14 to ensure that the mouse is fully anesthetized before commencing surgery. 
		NOTE: Movement of any part of the mouse is an indication that the mouse is not fully anesthetized. The animal should immediately be given additional anesthetic by increasing the anesthetic concentration.</li>
		<li>Cover the mouse&#39;s eyes with a small amount of ophthalmic gel to avoid eye dryness.</li>
	</ol>
	</li>
	<li>Surgical procedure (SURGEON)
	<ol>
		<li>Swab the surgical area 3 times with alcoholic chlorhexidine. Using forceps and a pair of scissors, make a 1 cm straight incision along the dorsal side, 3 mm away from the tumor (Figure 3A, 3B). 
		NOTE: Standardizing the incision to 1 cm in every mouse (using a ruler) allows for even assessment of wound healing between mice. Locating the incision 3 mm away from the tumor allows for subsequent intratumoral adjuvant therapy without leakage from the wound.</li>
		<li>Using tweezers, pull away the facia and subcutaneous fatty tissue between the tumor and peritoneum. The subcutaneous tumor is normally attached to the skin-side.</li>
		<li>Open the wound by gently holding the skin on the tumor bearing side using tweezers, and &#34;invert&#34; the tumor so that it is visible outside (Figure 3C, 3D). 
		NOTE: The section of tumor to be debulked should be closest to the opening, to have enough skin to close the wound. Be careful not to cut the skin when removing the tumor.</li>
		<li>Using a pair of scissors, cut away the tumor capsule from the half to remove, starting from the base of the tumor closest to the opening.</li>
		<li>For 50% debulk surgery, cut across the middle of the tumor. Using curved forceps, scoop up the section of the tumor to be removed (50%); scoop up any remnants from the debulked area.</li>
		<li>For 75% debulk, perform a 50% tumor debulk as in part 2.3.5 above. Then cut in half the remaining 50% of tumor and scoop up 25% of the tumor, using curved forceps as described above.</li>
	</ol>
	</li>
	<li>Closing the surgical site
	<ol>
		<li>Place the remaining tumor back underneath the skin, and using forceps, pull the skin flaps together and line up the skin along the wound.</li>
		<li>Hold the skin together 5 mm from the edge of the wound, and use surgical clips to close the wound, starting on the side closest to the forceps. Apply as many clips as needed to ensure no underlying tissue is exposed. Generally, three to four clips are applied with 2 mm gaps between clips. 
		NOTE: If any clips is not well applied, remove it using a clip remover and replace with new clips.</li>
	</ol>
	</li>
	<li>Recovery of mice (ASSISTANT)
	<ol>
		<li>Allow the mice to recover by putting them into the warm (37 &#176;C) heating chamber.</li>
		<li>Place the mouse&#39;s cage on the heat pad. Monitor the mice in the heating chamber until they have recovered from the anesthetic (awake and walking) and then put the mice back into the cage. Leave the cage on the heat pad for a further 10 minutes, until the mice have become more active.</li>
		<li>Give the mice wet and soft food. Monitor the mice 1 hour after surgery for recovery and ensure clips remain in place.&#160;Ensure the cage is half on/half off the heat pad to allow animals to self-regulate temperature while unattended.</li>
		<li>Dose mice with 0.1 mg/kg buprenorphine (100 &#181;L subcutaneously in the scruff of the neck), 6-8 hours after surgery (at the end of the day). Monitor mice early the following morning, and dose mice again with 0.1 mg/kg buprenorphine (100 &#181;L subcutaneously in the scruff of the neck). Give more wet food as needed.</li>
		<li>Monitor mice daily for the next seven days. Clips may be removed after seven days using the clip remover.</li>
	</ol>
	</li>
	<li>Adjuvant or neoadjuvant treatment
	<ol>
		<li>Treat mice peri-operatively with (neo)adjuvant therapy at any given time, depending on the treatment of interest.</li>
		<li>For example, treat mice with one dose of 100 &#181;g of anti-CTLA-4 intraperitoneally (i.p.) on day 15 after inoculation, or with three doses of 200 &#181;g anti-PD-1 i.p. on day 15, 17 and 19 after inoculation.</li>
	</ol>
	</li>
	<li>Experimental controls
	<ol>
		<li>When using this model to assess the effects of inflammation/wound healing, consider using the following control groups: 1) No-surgery control (treatments can still be administered intratumorally); 2) Sham surgery control: A surgical incision is made in the skin; the tumor is manipulated and exposed, but no tumor tissue is removed; the wound is closed with clips.</li>
	</ol>
	</li>
</ol>

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No disclosures.

We provide a protocol for a mouse model of incomplete surgical resection of soft tissue sarcoma to test peri-operative therapies. We also standardized the surgical incision to allow assessment of wound healing between mice following treatment.
Tumor placement is an important part of this protocol. We have opted for a subcutaneous tumor model to allow easy surgical access to the tumor site and administration of local therapies with minimal burden on the mice. It is also important to ensure that...

Surgery remains the main treatment option for many solid tumors1, including soft tissue sarcoma2,3. Despite improvements in cancer surgery techniques, and combinations with (neo)adjuvant therapies, there is still a high risk of cancer relapse and metastasis following primary tumor resection4,5. In soft tissue sarcoma, relapses occur particularly locoregionally, at the site of surgery, resulting in increased morbidity and mortality. In the clinical setting, it can be difficult to obtain wide enough margins (e.g., due to anatomical constraints), resulting in incomplete resection and subsequent tumor recurrence6. Surgical stress and the subsequent process of wound healing are known to create an immunosuppressive tumor microenvironment favorable for tumor recurrence7,8. Therefore, the discovery and development of new therapies for soft tissue sarcoma, particularly immunotherapies, should ideally take the surgical wound healing response into account.
Most preclinical studies for adjuvant therapies are initially carried out using subcutaneous syngeneic or xenotransplant mouse models, without incorporating the surgical stress and wound healing response9,10. Therefore, we developed a syngeneic subcutaneous mouse soft tissue sarcoma model incorporating incomplete surgical resection. WEHI 164 fibrosarcoma cells are inoculated subcutaneously, and once tumors are established, we remove 50-75% of the tumor bulk (Figure 1A-E). Tumors consistently re-grow from the remaining tumor. This model allows for testing adjuvant therapies while considering the effect of surgical stress and wound healing. Similar surgical models of incomplete resection have been used in a number of studies by several groups and found to be reproducible and effective11,12,13. Here, we provide a detailed description of this protocol.

<table><tbody><tr><td>26 gauge 0.5 mL insulin syringe</td><td>Becton Dickinson, Australia</td><td>326769</td><td>None</td></tr><tr><td>2-Mercaptoethanol</td><td>Life Technologies Australia Pty Ltd</td><td>21985023</td><td>None</td></tr><tr><td>Anaestetic gas machine</td><td>Darvall Vet, Australia</td><td>SKU: 2848</td><td>None</td></tr><tr><td>Anti-CTLA-4</td><td>BioXcell, USA</td><td>BE0164</td><td>None</td></tr><tr><td>Anti-PD-1</td><td>BioXcell, USA</td><td>BP0273</td><td>None</td></tr><tr><td>Buprenorphine Hydrochloride Injection, 0.3mg/mL</td><td>RB healthcare UK Limited, UK</td><td>55175</td><td>Prescription order</td></tr><tr><td>Chlorhexidine Surgical Scrub 4%</td><td>Perigo Australia, Australia</td><td>CHL01449F(scrub</td><td>None</td></tr><tr><td>Fetal Bovine serum</td><td>CellSera, Australia</td><td>AU-FBS-PG</td><td>None</td></tr><tr><td>Forceps Fine 10.5 cm</td><td>Surgical house, Western Australia</td><td>CC74110</td><td>None</td></tr><tr><td>Forceps Fine 12 cm Serrated</td><td>Surgical house, Western Australia</td><td>CC74212</td><td>None</td></tr><tr><td>Forceps Halsted 14 cm</td><td>Surgical house, Western Australia</td><td>CD01114</td><td>None</td></tr><tr><td>Heating chamber</td><td>Datesand Ltd, UK</td><td>Mini-Thermacage</td><td>None</td></tr><tr><td>HEPES (1M)</td><td>Life Technologies Australia Pty Ltd</td><td>15630080</td><td>None</td></tr><tr><td>Isoflurane</td><td>Henry Schein Animal Health, Australia</td><td>SKU: 29405</td><td>Prescription order</td></tr><tr><td>Lubricating Eye Ointment</td><td>Alcon</td><td>n/a</td><td>None</td></tr><tr><td>Penicillin/streptomycin 1000X</td><td>Life Technologies Australia Pty Ltd</td><td>15140122</td><td>None</td></tr><tr><td>Phosphate Buffered Solution 10x</td><td>Life Technologies Australia Pty Ltd</td><td>70013-032</td><td>None</td></tr><tr><td>Reflex 7mm Clips</td><td>Able scientific, Australia</td><td>AS59038</td><td>None</td></tr><tr><td>Reflex 7mm Wound Clip Applicator</td><td>Able scientific, Australia</td><td>AS59036</td><td>None</td></tr><tr><td>Reflex Wound Clip Remover</td><td>Able scientific, Australia</td><td>AS59037</td><td>None</td></tr><tr><td>Rodent Qube Anesthesia Breathing Circuit</td><td>Darvall Vet, Australia</td><td>#7885</td><td>None</td></tr><tr><td>Roswell Park Memorial Institute (RPMI) 1640 Medium + L-glutamine</td><td>Life Technologies Australia Pty Ltd</td><td>21870092</td><td>None</td></tr><tr><td>Scissors Iris STR 11 cm</td><td>Surgical house, Western Australia</td><td>KF3211</td><td>None</td></tr><tr><td>Scissors Iris STR 9 cm</td><td>Surgical house, Western Australia</td><td>JH4209</td><td>None</td></tr><tr><td>Small Induction Chamber</td><td>Darvall Vet, Australia</td><td>SKU: 9630</td><td>None</td></tr><tr><td>TrypLE express 1x</td><td>Life Technologies Australia Pty Ltd</td><td>12604-021</td><td>None</td></tr><tr><td>Germinator 500 Glass Bead Sterilizer</td><td>Cellpoint Scientific Inc., USA</td><td>5-1460-DK</td><td></td></tr></tbody></table>

a mouse model of incompletely resected soft tissue sarcoma for testing (neo)adjuvant therapies

Surgery is often the first treatment for many solid tumors. However, local relapses frequently occur following primary tumor resection, despite adjuvant or neo-adjuvant therapies. This occurs when surgical margins are insufficiently tumor-free, resulting in residual cancer cells. From a biological and immunological perspective, surgery is not a null event; the wound healing environment is known to induce both pro- and anti-tumorigenic pathways. As a consequence, preclinical models for drug development aimed at preventing local relapse should incorporate surgical resection when testing new (neo)adjuvant therapies, to model the clinical settings in patients treated with surgery.
Here, we describe a mouse model of incomplete surgical resection of WEHI 164 soft tissue sarcoma that allows testing of (neo)adjuvant therapies in the setting of a wound healing response. In this model, 50% or 75% of the tumor is removed, leaving behind some cancer tissue in situ to model gross residual disease after surgery in the clinical setting. This model allows testing therapies in the context of surgery while also considering the wound healing response, which may affect the efficacy of (neo)adjuvant treatments. The incomplete surgical resection results in reproducible regrowth of the tumor in all mice in the absence of adjuvant therapy. Adjuvant treatment with checkpoint blockade results in reduced tumor regrowth. This model is thus appropriate for testing therapies in the context of debulking surgery and its associated wound healing response and can be extended to other types of solid cancer.

Tumor growth to a size of 50 mm2 is an ideal size for partial debulk. The incomplete surgical resection of 50 mm2 tumors results in 100% (n=5) reproducible regrowth of the tumors in the absence of adjuvant immunotherapy (Figure 4A). We next used the model to test adjuvant immunotherapies using antibodies against checkpoint molecules Cytotoxic T Lymphocyte Associated Protein 4 (CTLA-4) and Programmed Death Receptor 1 (PD-...

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A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing Neoadjuvant Therapies

In this protocol, we describe a mouse model of incomplete surgical resection of soft tissue sarcoma for testing (neo)adjuvant therapies.

A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing (Neo)adjuvant Therapies

Surgery is often the first treatment for many solid tumors. However, local relapses frequently occur following primary tumor resection, despite adjuvant ...

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9.4K Views.  University of Western Australia. In this protocol, we describe a mouse model of incomplete surgical resection of soft tissue sarcoma for testing (neo)adjuvant therapies.

Video: A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing Neoadjuvant Therapies

Surgical Procedures and Methodology for a Preclinical Murine Model of De Novo Mammary Cancer Metastasis

A rate-limiting aspect of transgenic mouse models of mammary adenocarcinoma is that primary tumor burden in mammary tissue typically defines study end-points. Thus, studies focused on elucidating mechanisms of late-stage de novo metastasis are compromised, as are studies examining efficacy of anti-cancer therapies targeting mediators of metastasis in the adjuvant setting. Numerous murine mammary cancer models have been developed via targeted expression of dominant oncoproteins to mammary epithelial cells yielding models variably mimicking histopathologic and transcriptome-defined breast cancer subtypes common in women1. While much has been learned regarding the biology of mammary carcinogenesis with these models, their utility in identifying molecules regulating growth of late-stage metastasis are compromised as mice are typically euthanized at earlier time points due to significant primary tumor burden. Moreover, since a significant percentage of women diagnosed with breast cancer receive adjuvant therapy after surgical resection of primary tumors and prior to presence of detectable metastatic disease, preclinical models of de novo metastasis are urgently needed as platforms to evaluate new therapies aimed at targeting metastatic foci. To address these deficiencies, we developed a murine model of de novo mammary cancer metastasis, wherein primary mammary tumors are surgically resected, and metastatic foci subsequently develop over a 115 day post-surgical period. This long latency provides a tractable model to identify functionally significant regulators of metastatic progression in mice lacking primary tumor, as well as a model to evaluate preclinical therapeutic efficacy of agents aimed at blocking functionally significant molecules aiding metastatic tumor survival and growth.

Surgical Procedures and Methodology for a Preclinical Murine Model of De Novo Mammary Cancer Metastasis

Pre-clinical models evaluating adjuvant therapy targeting breast cancer metastasis are lacking. To address this, we developed a murine model of de novo pulmonary mammary adenocarcinoma metastasis, wherein therapies administered in the adjuvant setting (post surgical resection of primary tumors) can be evaluated for efficacy in impacting previously seeded pulmonary metastases.

A rate-limiting aspect of transgenic mouse models of mammary adenocarcinoma is that primary tumor burden in mammary tissue typically defines study ...

Cancer Research

A Preclinical Mouse Model of Osteosarcoma to Define the Extracellular Vesicle-mediated Communication Between Tumor and Mesenchymal Stem Cells

Within the tumor microenvironment, resident or recruited mesenchymal stem cells (MSCs) contribute to malignant progression in multiple cancer types. Under the influence of specific environmental signals, these adult stem cells can release paracrine mediators leading to accelerated tumor growth and metastasis. Defining the crosstalk between tumor and MSCs is of primary importance to understand the mechanisms underlying cancer progression and identify novel targets for therapeutic intervention. 
Cancer cells produce high amounts of extracellular vesicles (EVs), which can profoundly affect the behavior of target cells in the tumor microenvironment or at distant sites. Tumor EVs enclose functional biomolecules, including inflammatory RNAs and (onco)proteins, that can educate stromal cells to enhance the metastatic behavior of cancer cells or to participate in the pre-metastatic niche formation. In this article, we describe the development of a preclinical cancer mouse model that enables specific evaluation of the EV-mediated crosstalk between tumor and mesenchymal stem cells. First, we describe the purification and characterization of tumor-secreted EVs and the assessment of the EV internalization by MSCs. We then make use of a multiplex bead-based immunoassay to evaluate the alteration of the MSC cytokine expression profile induced by cancer EVs. Finally, we illustrate the generation of a bioluminescent orthotopic xenograft mouse model of osteosarcoma that recapitulates the tumor-MSC interaction, and show the contribution of EV-educated MSCs to tumor growth and metastasis formation. 
Our model provides the opportunity to define how cancer EVs shape a tumor-supporting environment, and to evaluate whether blockade of the EV-mediated communication between tumor and MSCs prevents cancer progression.

Direct injection of cancer-derived extracellular vesicles (EVs) leads to reprogramming of bone marrow supporting tumor progression; however, which cells mediate this effect is unclear. Herein, we describe a step-by-step protocol to investigate EV-mediated tumor-mesenchymal stem cell (MSC) interactions in vivo, revealing a crucial role for EV-educated MSCs in metastasis.

Within the tumor microenvironment, resident or recruited mesenchymal stem cells (MSCs) contribute to malignant progression in multiple cancer types. Under ...

An Orthotopic Resectional Mouse Model of Pancreatic Cancer

There is a lack of satisfactory animal models to study adjuvant and/or neoadjuvant therapy in patients being considered for surgery of pancreatic cancer (PC). To address this deficiency, we describe a mouse model involving orthotopic implantation of PC followed by distal pancreatectomy and splenectomy. The model has been demonstrated to be safe and suitably flexible for the study of various therapeutic approaches in adjuvant and neo adjuvant settings.
In this model, a pancreatic tumor is first generated by implanting a mixture of human pancreatic cancer cells (luciferase-tagged AsPC-1) and human cancer associated pancreatic stellate cells into the distal pancreas of Balb/c athymic nude mice. After three weeks, the cancer is resected by re-laparotomy, distal pancreatectomy and splenectomy. In this model, bioluminescence imaging can be used to follow the progress of cancer development and effects of resection/treatments. Following resection, adjuvant therapy can be given. Alternatively, neoadjuvant treatment can be given prior to resection.
Representative data from 45 mice are presented. All mice underwent successful distal pancreatectomy/splenectomy with no issues of hemostasis. A macroscopic proximal pancreatic margin greater than 5 mm was achieved in 43 (96%) mice. The technical success rate of pancreatic resection was 100%, with 0% early mortality and morbidity. None of the animals died during the week after resection.
In summary, we describe a robust and reproducible technique for a surgical resection model of pancreatic cancer in mice which mimics the clinical scenario. The model may be useful for the testing of both adjuvant and neoadjuvant treatments.

In the clinical context, patients with localized pancreatic cancer will undergo pancreatectomy followed by adjuvant treatment. This protocol reported here aims to establish a safe and effective method of modelling this clinical scenario in nude mice, through orthotopic implantation of pancreatic cancer followed by distal pancreatectomy and splenectomy.

There is a lack of satisfactory animal models to study adjuvant and/or neoadjuvant therapy in patients being considered for surgery of pancreatic cancer ...

A Syngeneic Orthotopic Osteosarcoma Sprague Dawley Rat Model with Amputation to Control Metastasis Rate

The most recent advance in the treatment of osteosarcoma (OS) occurred in the 1980s when multi-agent chemotherapy was shown to improve overall survival compared to surgery alone. To address this problem, the aim of the study is to refine a lesser-known model of OS in rats with a comprehensive histologic, imaging, biologic, implantation, and amputation surgical approach that prolongs survival. We used an immunocompetent, outbred Sprague-Dawley (SD), syngeneic rat model with implanted UMR106 OS cell line (originating from a SD rat) with orthotopic tibial tumor implants into 3-week-old male and female rats to model pediatric OS. We found that rats develop reproducible primary and metastatic pulmonary tumors, and that limb amputations at 3 weeks post implantation significantly reduce the incidence of pulmonary metastasis and prevent unexpected deaths. Histologically, the primary and metastatic OSs in rats were very similar to human OS. Using immunohistochemistry methods, the study shows that rat OS are infiltrated with macrophages and T cells. A protein expression survey of OS cells reveals that these tumors express ErbB family kinases. Since these kinases are also highly expressed in most human OSs, this rat model could be used to test ErbB pathway inhibitors for therapy.

Here, a syngeneic orthotopic implantation followed by an amputation procedure of the osteosarcoma with spontaneous pulmonary metastasis that can be used for preclinical investigation of metastasis biology and development of novel therapeutics is described.

The most recent advance in the treatment of osteosarcoma (OS) occurred in the 1980s when multi-agent chemotherapy was shown to improve overall survival ...

Intratibial Osteosarcoma Cell Injection to Generate Orthotopic Osteosarcoma and Lung Metastasis Mouse Models

Osteosarcoma is the most common primary bone cancer in children and adolescents, with lungs as the most common metastatic site. The five-year survival rate of osteosarcoma patients with pulmonary metastasis is less than 30%.Therefore, the utilization of mouse models mimicking the osteosarcoma development in humans is of great significance for understanding the fundamental mechanism of osteosarcoma carcinogenesis and pulmonary metastasis to develop novel therapeutics. Here, detailed procedures are reported to generate the primary osteosarcoma and pulmonary metastasis mouse models via&#160;intratibia injection of osteosarcoma cells. Combined with the bioluminescence or X-ray live imaging system, these living mouse models are utilized to monitor and quantify osteosarcoma growth and metastasis. To establish this model, a basement membrane matrix containing osteosarcoma cells was loaded in a micro-volume syringe and injected into one tibia of each athymic mouse after being anesthetized. The mice were sacrificed when the primary osteosarcoma reached the size limitation in the IACUC-approved protocol. The legs bearing osteosarcoma and the lungs with metastasis lesions were separated. These models are characterized by a short incubation period, rapid growth, severe lesions, and sensitivity in monitoring the development of primary and pulmonary metastatic lesions. Therefore, these are ideal models for exploring the functions and mechanisms of specific factors in osteosarcoma carcinogenesis and pulmonary metastasis, the tumor microenvironment, and evaluating the therapeutic efficacy in vivo.

The present protocol describes intratibia osteosarcoma cell injection to generate mouse models bearing orthotopic osteosarcoma and pulmonary metastasis lesions.

Osteosarcoma is the most common primary bone cancer in children and adolescents, with lungs as the most common metastatic site. The five-year survival ...

Magnetic Resonance-Guided High Intensity Focused Ultrasound Generated Hyperthermia: A Feasible Treatment Method in a Murine Rhabdomyosarcoma Model

Magnetic resonance-guided high intensity focused ultrasound (MRgHIFU) is an established method for producing localized hyperthermia. Given the real-time imaging and acoustic energy modulation, this modality enables precise temperature control within a defined area. Many thermal applications are being explored with this noninvasive, nonionizing technology, such as hyperthermia generation, to release drugs from thermosensitive liposomal carriers. These drugs can include chemotherapies such as doxorubicin, for which targeted release is desired due to the dose-limiting systemic side effects, namely cardiotoxicity. Doxorubicin is a mainstay for treating a variety of malignant tumors and is commonly used in relapsed or recurrent rhabdomyosarcoma (RMS). RMS is the most common solid soft tissue extracranial tumor in children and young adults. Despite aggressive, multimodal therapy, RMS survival rates have remained the same for the past 30 years. To explore a solution for addressing this unmet need, an experimental protocol was developed to evaluate the release of thermosensitive liposomal doxorubicin (TLD) in an immunocompetent, syngeneic RMS mouse model using MRgHIFU as the source of hyperthermia for drug release.

Presented here is a protocol to use controlled hyperthermia, generated by magnetic resonance-guided high intensity focused ultrasound, to trigger drug release from temperature-sensitive liposomes in a rhabdomyosarcoma mouse model.

Magnetic resonance-guided high intensity focused ultrasound (MRgHIFU) is an established method for producing localized hyperthermia. Given the real-time ...

Modeling Spontaneous Metastatic Renal Cell Carcinoma (mRCC) in Mice Following Nephrectomy

One of the key challenges to improved testing of new experimental therapeutics in renal cell carcinoma (RCC) is the development of models that faithfully recapitulate early- and late-stage metastatic disease progression. Typical tumor implantation models utilize ectopic or orthotopic primary tumor implantation, but few include systemic spontaneous metastatic disease that mimics the clinical setting. This protocol describes the key steps to develop RCC disease progression stages similar to patients. First, it uses a highly metastatic mouse tumor cell line in a syngeneic model to show orthotopic tumor cell implantation. Methods include superficial and internal implantation into the sub-capsular space with cells combined with matrigel to prevent leakage and early spread. Next it describes the procedures for excision of tumor-bearing kidney (nephrectomy), with critical pre- and post- surgical mouse care. Finally, it outlines the steps necessary to monitor and assess micro-and macro-metastatic disease progression, including bioluminescent imaging as well provides a detailed visual necropsy guide to score systemic disease distribution. The goal of this protocol description is to facilitate the widespread use of clinically relevant metastatic RCC models to improve the predictive value of future therapeutic testing.&#160;

Modeling Spontaneous Metastatic Renal Cell Carcinoma mRCC in Mice Following Nephrectomy

Models of spontaneous metastatic renal cell carcinoma (RCC) disease progression can be used for evaluating treatments in a clinically relevant setting. This protocol demonstrates different procedures for orthotopic kidney tumor cell implantation, proper nephrectomy, and finally outlines a necropsy guide for visual and bioluminescent scoring of metastatic burden and localization.

One of the key challenges to improved testing of new experimental therapeutics in renal cell carcinoma (RCC) is the development of models that faithfully ...

Generating a Murine Orthotopic Metastatic Breast Cancer Model and Performing Murine Radical Mastectomy

In vivo mouse models to assess breast cancer progression are essential for cancer research, including preclinical drug developments. However, the majority of the practical and technical details are commonly omitted in published manuscripts which, therefore, makes it challenging to reproduce the models, particularly when it involves surgical techniques. Bioluminescence technology allows for the evaluation of small amounts of cancer cells even when a tumor is not palpable. Utilizing luciferase-expressing cancer cells, we establish a breast cancer orthotopic inoculation technique with a high tumorigenesis rate. Lung metastasis is assessed utilizing an ex vivo technique. We, then, establish a mastectomy model with a low local recurrence rate to assess the metastatic tumor burden. Herein, we describe, in detail, the surgical techniques of orthotopic implantation and mastectomy for breast cancer with a high tumorigenesis rate and low local recurrence rates, respectively, to improve breast cancer model efficiency.

We introduce a murine orthotopic breast cancer model and radical mastectomy model with bioluminescence technology to quantify the tumor burden to mimic human breast cancer progression.

In vivo mouse models to assess breast cancer progression are essential for cancer research, including preclinical drug developments. However, the majority ...

Modeling Brain Tumor Metastasis via Internal Carotid Artery Injection of Cancer Cells

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/23284/modeling-brain-tumor-metastasis-via-internal-carotid-artery-injection'>Source: Lim, M., et al. Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells. J. Vis. Exp. (2022)</a>In this video, an anesthetized mouse undergoes a surgical procedure in which cancer cells are injected into the common carotid artery (CCA) to facilitate targeted delivery to the brain. The procedure involves precise suturing and manipulation of the carotid arteries to direct the injected cells toward the internal carotid artery (ICA), promoting tumor formation in the brain over time.

Source: Lim, M., et al. Modeling Brain Metastasis by Internal Carotid Artery Injection of Cancer Cells. J. Vis. Exp. (2022)In this video, an anesthetized ...

JoVE Encyclopedia of Experiments

Neuroscience

Neuropathology

Tumors

An Adjuvant Therapy in a Mouse Model with an Incompletely Resected Subcutaneous Tumor

Source: Rwandamuriye, F. X. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/60882">A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing (Neo)adjuvant Therapies.</a>&#160;J. Vis. Exp.&#160;(2020)
This video illustrates testing of adjuvant or additional therapy in a mouse model with incompletely resected subcutaneous tumors. The partially tumor-resected mouse is treated with an antibody targeting the anti-cytotoxic protein of T regulatory cells, potentially reducing tumor regrowth and suggesting effective adjuvant therapy.

Terapia uzupełniająca w modelu mysim z niecałkowicie wyciętym guzem podskórnym

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing (Neo)adjuvant Therapies

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Surgical Procedures and Methodology for a Preclinical Murine Model of De Novo Mammary Cancer Metastasis

A Preclinical Mouse Model of Osteosarcoma to Define the Extracellular Vesicle-mediated Communication Between Tumor and Mesenchymal Stem Cells

An Orthotopic Resectional Mouse Model of Pancreatic Cancer

A Syngeneic Orthotopic Osteosarcoma Sprague Dawley Rat Model with Amputation to Control Metastasis Rate

Intratibial Osteosarcoma Cell Injection to Generate Orthotopic Osteosarcoma and Lung Metastasis Mouse Models

Magnetic Resonance-Guided High Intensity Focused Ultrasound Generated Hyperthermia: A Feasible Treatment Method in a Murine Rhabdomyosarcoma Model

Modeling Spontaneous Metastatic Renal Cell Carcinoma (mRCC) in Mice Following Nephrectomy

Generating a Murine Orthotopic Metastatic Breast Cancer Model and Performing Murine Radical Mastectomy

Modeling Brain Tumor Metastasis via Internal Carotid Artery Injection of Cancer Cells

Terapia uzupełniająca w modelu mysim z niecałkowicie wyciętym guzem podskórnym