Name: A Syngeneic Orthotopic Osteosarcoma Sprague Dawley Rat Model with Amputation to Control Metastasis Rate (Video) | JoVE Journal
Uploaded: 2021-05-03T13:00:00.000+00:00
Duration: 7 min 31 s
Description: Explore a syngeneic orthotopic osteosarcoma rat model using Sprague-Dawley rats with limb amputation to control metastasis. This protocol enables preclinical testing of novel cancer therapies in an immunocompetent setting.

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5 minutes

NIH funding through National Cancer Institute, grant # CA228582. Shun Ishiyama is currently receiving a grant from Toray Medical Co., Ltd.

All the procedures and experiments involving rats were performed according to protocols approved by Johns Hopkins Animal Care and Use Committee.
1. The SD rat OS cell line UMR-106 cell culture protocol
<ol>
	<li>Grow cells in DMEM, supplemented with 10% (v/v) FBS, penicillin (10 U/mL)-streptomycin (10 U/mL) at 37 &#176;C in humidified 5% CO2 atmosphere. Perform experiments using cells with passages of 2-812.</li>
</ol>
2. Intratibial injection of OS cells protocol
NOTE: Time-mated pregnant SD rats give birth in the animal facility and at 3 weeks of age, litters are used (since UMR 106 cell line is syngeneic to SD rats, no irradiation is needed).
<ol>
	<li>Induction
	<ol>
		<li>Place the rat in a medium-sized induction chamber and induce anesthesia with 2%-3% isoflurane. Monitor the animal continuously for the depth of anesthesia by reflex to toe pinch, respiratory rate, and character.</li>
		<li>Insert the nose into the nosecone. Secure with tape, if necessary.</li>
		<li>Remove the hair on the right leg up to the ventral and the dorsal lower abdomen with clippers or use depilatory agent. Place the rat in a supine position.</li>
		<li>Scrub the surgical area aseptically using 70% ethanol and dilute chlorhexidine acetate or dilute betadine. Begin around the area of the knee and scrub in a circular motion both proximally and distally. Repeat this step three times. No drape is used for tumor implantation.</li>
		<li>Apply eye lubricant in both the eyes of the rat to prevent corneal desiccation caused by anesthesia. Place the rat on a low heat setting heating pad. Ensure that the rat has normal body temperature (37 &#176;C) and normal respiratory rate.</li>
	</ol>
	</li>
	<li>Surgery
	<ol>
		<li>Turn on isoflurane at ~1.5%-2% (for maintenance). Ensure that the animal is at an adequate plane of anesthesia via lack of a toe-pinch reflex. If not, increase the isoflurane percentage to 2.5%.</li>
		<li>Mark a sterile needle (approximately 22 G) at 10 mm from the tip for guidance of depth to insert.</li>
		<li>Insert the needle 10 mm down into the diaphysis of the tibia by entering the bent knee in the middle of the tibial plateau, extending the needle through the metaphysis into the diaphysis using a light drill-like motion to make an opening. Remove the needle.</li>
		<li>Load the cell suspension into the Hamilton syringe directly before injection into the tibia. To do this, gently mix the cells before drawing into syringe as gravity makes the cells settle in the bottom of the tube. 
		NOTE: Cells can be stored in a 1.5 to 2 mL tube (at room temperature) before drawing into the Hamilton syringe (100 &#181;L) after careful mixing. Cells can be kept at room temperature for 2-3 h during the implantation procedure. Always check the cell viability of cells in the tube before and then after the implantation session. Trypan blue exclusion is the easiest method for cell viability assessment.</li>
		<li>Once the bone is traversed with the first needle, insert a second (smaller diameter needle also marked at 10 mm) needle attached to the 100 &#181;L Hamilton syringe loaded with cells. Ensure to insert the needle up to the 10 mm mark into the same hole extending into the diaphysis.</li>
		<li>Gently discharge 20 &#181;L of 75,000 OS cells suspension in PBS into the diaphysis and marrow cavity. 
		NOTE: The needle should not wobble in the bone and should feel secure. If the needle easily moves, the cortex may have been traversed at the diaphysis. Repeat the insertion again to get a firmer placement before the injection of cells.</li>
		<li>Remove the Hamilton needle from the bone. 
		NOTE: If a small drop of blood forms, apply light pressure. If clear fluid drop forms at the puncture site, the needle may not have been extended far enough in the bone and the tumor cell suspension may have leaked back through the hole. Record this in the notes but generally, tumor implantation will be successful. With experience, the tumor implantation procedure should take 5 min per rat. With experience, the tumor implantation of cells in the bone will become easier. Accidental injection of cells in the muscle around the bone, may not lead to the tumor microenvironment needed for pulmonary metastasis.</li>
	</ol>
	</li>
	<li>Recovery
	<ol>
		<li>Ensure that the rat is normothermic. Place the rat in a cage with a heating pad placed under the cage for recovery.</li>
		<li>After fully awake, mobile, and breathing well, inject the rats with Buprenorphine (1.0-1.2 mg/kg SC). 
		NOTE: To get access to Buprenorphine, check with the institution to see options for approval through the veterinary staff to place the order.</li>
		<li>Move the rats back to clean cages and monitor once a day, each week.</li>
	</ol>
	</li>
</ol>
3. Measurement and monitoring
<ol>
	<li>Measure the tumor size 9-10 days post-implantation and then every 2 days until 3 weeks post-implantation to establish a growth rate. Measure the maximum diameter of the tibia using an electronic or manual caliper. Store the data in a spreadsheet with a formula to calculate tumor volume. Measure the contralateral (not implanted tibia) as the baseline. 
	NOTE: The implanted hindlimb diameters are used as a surrogate for tumor size. The hind limbs are measured perpendicular to the tibia long axis at the largest diameter for two measurements, ventral/dorsal and media/ lateral on the limb. The estimated tumor volume is calculated by the formula11: Tumor volume (mm3) = largest diameter (mm) x smallest diameter (mm)2/2.</li>
	<li>Consider the rats for survival amputation or chemotherapy treatment when tumors approach 15 mm in the largest dimension or 3 weeks post tumor implantation. The contralateral limb measures about 7-9 mm in most rats at this age.</li>
</ol>
4. Doxorubicin intravenous administration
<ol>
	<li>Anesthetize the rats with 2% isoflurane. Prepare the skin over jugular vein with three surgical washes using betadine and alcohol as described13.</li>
	<li>Under careful dissection, visualize the right or left jugular vein. Insert the needle into the overlying muscle and then direct toward the rat&#39;s head into the jugular vein lumen as it is visualized in the jugular vein anterior to the pectoralis muscle.
	<ol>
		<li>When the needle is inserted into the jugular vein, gently draw blood into the syringe to ensure proper insertion. It is possible to believe that the needle is through the vein, but the needle is under the vein and not in the lumen. If the jugular vein becomes too small for injections when blunt dissecting to expose the jugular vein, use the other jugular vein for the injection.</li>
	</ol>
	</li>
	<li>Inject doxorubicin (2 mg/kg) slowly over 1 min in a volume of 100-150 &#181;L. The solution can be visualized intravenously in the jugular vein during delivery.</li>
	<li>Inject similar volumes of normal saline in control rats.</li>
	<li>Remove the needle and apply gentle pressure on the vein with a sterile gauze.</li>
	<li>Close the skin incision using 3-4 wound clips. Remove the clips at 7-10 days post injection. 
	NOTE: Rats do not usually try to remove metal clips but will bite and remove sutures. The jugular injection method is preferable to tail vein injections for doxorubicin since any drug that leaks outside the vessel causes necrosis of tail that may require tail amputation.</li>
</ol>
5. Hind limb amputation protocol
<ol>
	<li>Induction
	<ol>
		<li>Place the rat in a medium-sized induction chamber and induce anesthesia with 2%-4% isoflurane. Monitor the rat continuously for the depth of the anesthesia. 
		NOTE: The induction chamber is scavenged to a charcoal canister and any other gases are removed by a down-draft table used for surgery.</li>
		<li>Insert the nose into a nosecone. Secure with tape, if necessary. 
		NOTE: This procedure is done on a down-draft table to scavenge excess volatile gases (i.e., isoflurane).</li>
		<li>Remove the hair on the right leg up to the ventral and the dorsal lower abdomen with clippers or use depilatory agent. Place the rat in a supine position.</li>
		<li>Scrub the surgical area aseptically using 70% ethanol and dilute chlorhexidine acetate or dilute betadine. Prepare the skin for surgery from middle of the calf to skin area just above the hip joint of the right lower abdomen. Scrub the leg proximal and the distal area circumferentially. Repeat this step three times.</li>
		<li>Apply the eye lubricant in both the eyes of the rat. Ensure that the rat has normal body temperature (37 &#176;C) and normal vital signs. Monitor and regulate body temperature using a heating pad that is connected to a rectal temperature probe monitor.</li>
	</ol>
	</li>
	<li>Surgery
	<ol>
		<li>Turn on isoflurane at 1.5%-3% (maintenance). Monitor for anesthetic depth, including reaction to toe pinch, respiratory rate, and character. Adjust the rate of isoflurane as needed to maintain an appropriate plane of anesthesia. 
		NOTE: Respiratory rate while under anesthesia should be between 50-100 breaths per minute. Deep, infrequent breaths are an indication that the rat is too deeply anesthetized.</li>
		<li>Open sterile instrument packs and drapes and don sterile gloves. Take care to maintain sterility through the duration of the procedure. Basic sterile surgical instruments needed include, scalpel blade holder, forceps, hemostats, needle holder, scissors, and wound clip applier.</li>
		<li>Pull the leg of the rat through the vent of the sterile surgical drape. Ensure that the animal is at an adequate plane of anesthesia via toe-pinch reflex.</li>
		<li>Using a scalpel blade or surgical scissors, make a circumferential, cutaneous incision just proximal to the stifle (knee joint).</li>
		<li>Deglove the hind limb using gauze or blunt dissection to expose the femoral artery and vein on the ventral-medial surface of the hind limb.</li>
		<li>Ligate the vessels using 4-0 absorbable sutures at the level of the mid-femur and transect distally.</li>
		<li>Clamp the vein distally to reduce leakage during muscle dissection. 
		NOTE: Circumferential musculature will be transected distal to the level of femoral artery vessel ligation and muscles elevated from the femur to the level of the coxofemoral joint.</li>
		<li>Using blunt dissection, abduct the hip joint with lateral outward rotation.</li>
		<li>Find the head of the femur and disarticulate it from the acetabulum. Cut any remaining tissue keeping the leg attached to the body.</li>
		<li>Give a splash block to the acetabulum and sciatic nerve with approximately 6 mg/kg Ropivacaine.</li>
		<li>Close the musculature over the acetabulum using a simple interrupted suture (4-0, absorbable suture). 
		NOTE: Additional 0.5% bupivacaine or lidocaine (0.15-0.2 mg total) may be injected in several sites along the closed muscle layer (local infiltration/splash block).</li>
		<li>Oppose and close the edges of the skin using wound clips placed apart every 5-10 mm.</li>
	</ol>
	</li>
	<li>Recovery
	<ol>
		<li>Place the rat in a clean recovery cage with heat support using a heating pad placed under the cage. 
		NOTE: To prevent hyperthermia, the heating pad should not be in direct contact with the animal and should not exceed 40 &#176;C.</li>
		<li>Monitor the animal until it fully recovers and is normothermic (37.5-39 &#176;C). 
		NOTE: The animal should not be left unattended until fully conscious and sternally recumbent and moving easily around the cage.</li>
		<li>After fully awake, mobile, and breathing well, inject the rats with Buprenorphine (1.0-1.2 mg/kg SC). 
		NOTE: Giving Buprenorphine in an anesthetized rat may impair recovery.</li>
		<li>Administer 10 mL of warm lactated Ringer&#39;s solution subcutaneously between the shoulder blades.</li>
		<li>Move the rats back to the clean cage and reunite with conspecifics.</li>
		<li>Monitor all the animals twice daily for the next month for signs of pain and distress, including piloerection, hunched posture, or inappetence or signs of incision site infection, including erythema, purulent discharge, or wound dehiscence. 
		NOTE: To date we have not observed clinical signs (such as infections) following post-operative recovery in any of the animals. Only one rat had a dehiscence with sutures, after which wound clips were used to close wounds with no further problems.</li>
		<li>Administer buprenorphine or meloxicam to animals presenting clinical signs of pain at published doses in consultation with a laboratory animal veterinarian.</li>
		<li>Administer antibiotics (i.e., cephalosporin) to animals presenting clinical signs of pain at published doses in consultation with a laboratory animal veterinarian. 
		NOTE: Any animals exhibiting protracted signs of pain or distress that do not improve with analgesics or animals exhibiting signs of infection that do not respond to antibiotics need to be humanely euthanized.</li>
	</ol>
	</li>
</ol>
6. Imaging with X-ray
<ol>
	<li>After tumor implantation, image the tibias and lungs non-invasively to detect tumor growth using X-ray with a machine designed for rodents.</li>
	<li>Anesthetize the rats as previously done.</li>
	<li>Take images at 3x magnification for 6 s at 25 kV.</li>
	<li>Process the film using the X-ray processor. Radiographs can be digitally scanned as well.</li>
</ol>
7. Necropsy procedure
<ol>
	<li>Euthanize the rats with CO2. Confirm death via lack of heartbeat and immediately draw 3 mL blood from the heart for serum or plasma samples.</li>
	<li>Open the thorax and abdomen for examination.</li>
	<li>Isolate the trachea and cannulate with a (18 G) catheter. To secure the catheter in the trachea, tie a silk suture around both the trachea with the catheter.</li>
	<li>Connect the infusion catheter to a 3 or 5 mL syringe. Infuse formalin or saline to gently inflate the lung lobes for better histology specimens. On infusion, the lungs will puff up and enable a better visualization of pulmonary metastases.</li>
	<li>Examine, dissect, and weigh all the selected thoracic and abdominal organs (such as liver, kidneys).</li>
	<li>Fix the organs in formalin for histopathology or freeze on dry ice, 2-methylbutane, or liquid nitrogen.</li>
	<li>For evaluation of protein expression using western blot, make lysates of frozen tissues. Antibodies that react with rat tissues are detailed13.</li>
</ol>
8. Immunohistochemistry
<ol>
	<li>Process the primary OS tissue, embed in paraffin, and section it for immunohistochemical staining.</li>
	<li>Retrieve the antigen after deparaffinization using citrate buffer (pH 6.0). Incubate in 0.3% H2O2 in methanol for 30 min to quench endogenous peroxidase.</li>
	<li>Block the 5 &#181;m thick paraffin sections using normal serum.</li>
	<li>Incubate with primary anti-CD68 and CD3 antibodies (see table) overnight at 4 &#176;C.</li>
	<li>Rinse the sections in PBS and incubate them in HRP polymer using a detection kit. 
	NOTE: Immunostaining was developed with diaminobenzidine as a chromogen.</li>
</ol>
9. Western blotting
<ol>
	<li>Lyse the UMR-106 cells in 200-300 &#181;L of lysis buffer14 to perform standard gel electrophoresis and western blotting.</li>
	<li>Use 4% to 12% Bis-Tris gels.</li>
	<li>Incubate in anti-ErbB2, anti-ErbB4, anti-EGFR, anti-ERK, &#946;-actin or anti-mouse &#946;2-AR primary antibody (see table) and horseradish peroxidase-linked secondary antibody.</li>
	<li>Add the chemiluminescent substrate. Expose the membranes to X-ray film. 
	NOTE: &#946;-actin levels are used as loading controls.</li>
</ol>

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<li>Khanna, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+orthotopic+model+of+murine+osteosarcoma+with+clonally+related+variants+differing+in+pulmonary+metastatic+potential%5BTitle%5D)+AND+%22Clinical+%26+Experimental+Metastasis%22%5BJournal%5D)">An orthotopic model of murine osteosarcoma with clonally related variants differing in pulmonary metastatic potential.</a> Clinical & Experimental Metastasis. 18 (3), 261-271 (2000).</li>
<li>Martin, T. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Parathyroid+hormone-responsive+adenylate+cyclase+in+induced+transplantable+osteogenic+rat+sarcoma%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Parathyroid hormone-responsive adenylate cyclase in induced transplantable osteogenic rat sarcoma.</a> Nature. 260 (5550), 436-438 (1976).</li>
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<li>Yu, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Establishment+of+reproducible+osteosarcoma+rat+model+using+orthotopic+implantation+technique%5BTitle%5D)+AND+%22Oncology+Reports%22%5BJournal%5D)">Establishment of reproducible osteosarcoma rat model using orthotopic implantation technique.</a> Oncology Reports. 21 (5), 1175-1180 (2009).</li>
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</ol>

No disclosure to declare.

Rats with OS tibial implants develop measurable tumors by 3 weeks post-implantation. If limbs with tumors are amputated 3 weeks post-implantation, the incidence of lung metastasis is reduced significantly. OSs are both osteolytic and osteoblastic. Rats without amputation develop lung metastases that are multiple and variably sized, observed by radiography or at necropsy by 7 weeks post-implantation.EGFR, ErbB2, and ErbB4 are expressed in rat UMR106 OS, similar to human OS16,<sup class="...

Osteosarcoma (OS) is the most common primary bone tumor in children, adolescents, and young adults. The most recent advance in the treatment of OS occurred in the 1980s when multi-agent chemotherapy was shown to improve overall survival compared to surgery alone1. OS develops during rapid bone growth, typically occurring in long tubular bones such as femur, tibia, and humerus. They are characterized by an osteolytic, osteoblastic, or mixed appearance with notable periosteal reaction2. Chemotherapy and surgical resection can improve the outcome for patients with a 5-year survival for 65% of patients2,3. Unfortunately, high grade OS patients with metastatic disease have 20% survival. OS invades regionally and metastasizes primarily to the lungs or other bones and is more prevalent in males. The most compelling need for these young patients is a novel therapy that prevents and eliminates viability of distant metastases.
OS pre-clinical models have been reviewed4,5,6,7 and few available immunocompetent models using amputation of orthotopic OS have been developed. In 2000, an important model was developed using BALB/c mice with orthotopic syngeneic OS and amputation8. Compared to this mouse model, the rat model is based on genetically outbred and 10 times larger animals leading to some advantages. The rat UMR106 model was developed from a 32P induced OS in a Sprague Dawley (SD) rat, which was derived into a cell line9. In 2001, orthotopic implantation of UMR106-01 was first described in implanted tibias of athymic mice with rapid, consistent primary tumor development and radiological, histologic features in common with OS in humans. Pulmonary metastases developed and were dependent on orthotopic placement of UMR106 into the bone microenvironment10. In 2009, Yu et al.11 established a reproducible orthotopic femur OS rat model using UMR106 cells in larger male SD rats. The successful tumor implantations and lung metastasis rate in rats without amputation were similar to the data presented here. In this study, an added amputation to the model using young rats was performed, which suggested that the timing of primary tumor removal is crucial in modeling OS, especially related to metastatic progression. With this refinement, amputation and in vivo imaging improve this model for pre-clinical studies for novel drug assessment for OS.

<table><tbody><tr><td>AKT</td><td>Cell Signaling TECHNOLOGY</td><td>4685S</td><td></td></tr><tr><td>absorbable suture</td><td>Ethicon</td><td>J214H</td><td></td></tr><tr><td>&amp;beta;-actin</td><td>SANTA CRUZ BIOTECHNOLOGY</td><td>sc-47778</td><td></td></tr><tr><td>&amp;beta;2-AR antibody</td><td>SANTA CRUZ BIOTECHNOLOGY</td><td>sc-569</td><td>replaced by &amp;beta;2-AR (E-3): sc-271322</td></tr><tr><td>Bis&amp;ndash;Tris gels</td><td>Thermo Fisher</td><td>NP0321PK2</td><td></td></tr><tr><td>Buprenorphine SR Lab</td><td>ZooPharm</td><td>IZ-70000-201908</td><td></td></tr><tr><td>CD3 antibody</td><td>Dako</td><td>#A0452</td><td></td></tr><tr><td>CD68 antibody</td><td>eBioscience</td><td>#14-0688-82</td><td></td></tr><tr><td>Chemiluminescent substrate</td><td>cytiva</td><td>RPN2232</td><td></td></tr><tr><td>CL-Xposure film</td><td>Thermo Fisher</td><td>34089</td><td></td></tr><tr><td>Complete Anesthesia System</td><td>EVETEQUIP</td><td>922120</td><td></td></tr><tr><td>diaminobenzidine</td><td>VECTOR LABORATORIES</td><td>SK-4100</td><td></td></tr><tr><td>Doxorubicin</td><td>Actavis</td><td>NDC 45963-733-60</td><td></td></tr><tr><td>EGFR antibody</td><td>SANTA CRUZ BIOTECHNOLOGY</td><td>sc-03</td><td>replaced by EGFR (A-10): sc-373746</td></tr><tr><td>ERBB2 antibody</td><td>SANTA CRUZ BIOTECHNOLOGY</td><td>sc-284</td><td>replaced by Neu (3B5): sc-33684</td></tr><tr><td>ERBB4 antibody</td><td>SANTA CRUZ BIOTECHNOLOGY</td><td>sc-283</td><td>replaced by ErbB4 (C-7): sc-8050</td></tr><tr><td>ERK antibody</td><td>SANTA CRUZ BIOTECHNOLOGY</td><td>sc-514302</td><td></td></tr><tr><td>eye lubricant</td><td>PHARMADERM</td><td>NDC 0462-0211-38</td><td></td></tr><tr><td>Hamilton syringe (100 &amp;micro;L)</td><td>Hamilton</td><td>Model 1710 SN SYR</td><td></td></tr><tr><td>horseradish peroxidase-linked secondary antibody</td><td>cytiva</td><td>NA934</td><td></td></tr><tr><td>HRP polymer detection kit</td><td>VECTOR LABORATORIES</td><td>MP-7401</td><td></td></tr><tr><td>HRP polymer detection kit</td><td>VECTOR LABORATORIES</td><td>MP-7402</td><td></td></tr><tr><td>isoflurane</td><td>BUTLER SCHEIN</td><td>NDC 11695-6776-2</td><td></td></tr><tr><td>isoflurane vaporizer</td><td>EVETEQUIP</td><td>911103</td><td></td></tr><tr><td>UMR-106 cell</td><td>ATCC</td><td>CRL-1661</td><td></td></tr><tr><td>X-ray</td><td>Faxitron</td><td>UltraFocus</td><td></td></tr><tr><td>X-ray processor</td><td>Hope X-Ray Peoducts Inc</td><td>MicroMax X-ray Processor</td><td>Hope Processors are not available in USA anymore</td></tr><tr><td>wound clips</td><td>BECTON DICKINSON</td><td>427631</td><td></td></tr></tbody></table>

a syngeneic orthotopic osteosarcoma sprague dawley rat model with amputation to control metastasis rate

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.

Immunocompetent SD outbred rats are used for these OS studies, which offers an animal model with an intact immune system. We have used the UMR106 cell line from ATCC, developed from cells that were initially isolated from an OS from a SD rat. We implanted the cells into SD rats, thus providing a syngeneic model for OS. UMR106 cells are implanted into the tibia of 3-week-old male and female SD rats, simulating a pediatric OS model. Moreover, the orthotopic implantation of UMR106 cells directly into the tibia metaphysis/di...

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 ...

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Research

JoVE Journal

Cancer Research

To study in vivo tumor growth and tumor microenvironment, we used a syngeneic and orthotopic mouse model of ovarian cancer in immunocompetent animals. We transduced a mouse tumor cell line (MOV1) with Katushka fluorescent protein (MOV1KAT) and here we show its orthotopic implantation in ovary and in vivo imaging.

An Orthotopic Model of Serous Ovarian Cancer in Immunocompetent Mice for in vivo Tumor Imaging and Monitoring of Tumor Immune Responses

The generation of an orthotopic rat model of pleural malignant mesothelioma by implantation of II-45 mesothelioma cells into the pleural cavity of immune competent rats is presented. A flow cytometric method to analyze seven immune cell subsets in these animals from a 25 µl blood sample is also described.

Orthotopic Implantation and Peripheral Immune Cell Monitoring in the II-45 Syngeneic Rat Mesothelioma Model

An orthotopic breast cancer primary tumor model and surgical removal of primary tumor to extend mouse life to generate spontaneous metastasis are described. The tumor growth and progression are monitored and quantified by luciferase fluorescence imaging.

An Orthotopic Mouse Model of Spontaneous Breast Cancer Metastasis

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.

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

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

This protocol was designed to train a machine learning algorithm to use a combination of imaging parameters derived from magnetic resonance imaging (MRI) and positron emission tomography/computed tomography (PET/CT) in a rat model of breast cancer bone metastases to detect early metastatic disease and predict subsequent progression to macrometastases.

Machine Learning Algorithms for Early Detection of Bone Metastases in an Experimental Rat Model

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

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

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

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

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

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

This current protocol outlines a procedure for creating a rat model of osteosarcopenia using ovariectomy.

The Creation of a Rat Model for Osteosarcopenia via Ovariectomy

Explore a syngeneic orthotopic osteosarcoma rat model using Sprague-Dawley rats with limb amputation to control metastasis. This protocol enables preclinical testing of novel cancer therapies in an immunocompetent setting.

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

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