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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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>&#946;-actin</td>
			<td>SANTA CRUZ BIOTECHNOLOGY</td>
			<td>sc-47778</td>
			<td></td>
		</tr>
		<tr>
			<td>&#946;2-AR antibody</td>
			<td>SANTA CRUZ BIOTECHNOLOGY</td>
			<td>sc-569</td>
			<td>replaced by &#946;2-AR (E-3): sc-271322</td>
		</tr>
		<tr>
			<td>Bis&#8211;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 &#181;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

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...

Watch this Scientific Journal Video about A Syngeneic Orthotopic Osteosarcoma Sprague Dawley Rat Model with Amputation to Control Metastasis Rate at JoVE.com

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

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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Cancer Research

3.5K Views.  The Johns Hopkins University School of Medicine. 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.

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

Fluorescent Orthotopic Mouse Model of Pancreatic Cancer

Pancreatic cancer remains one of the cancers for which survival has not improved substantially in the last few decades. Only 7% of diagnosed patients will survive longer than five years. In order to understand and mimic the microenvironment of pancreatic tumors, we utilized a murine orthotopic model of pancreatic cancer that allows non-invasive imaging of tumor progression in real time. Pancreatic cancer cells expressing green fluorescent protein (PANC-1 GFP) were suspended in basement membrane matrix, high concentration, (e.g., Matrigel HC) with serum-free media and then injected into the tail of the pancreas via laparotomy. The cell suspension in the high concentration basement membrane matrix becomes a gel-like substance once it reaches room temperature; therefore, it gels when it comes in contact with the pancreas, creating a seal at the injection site and preventing any cell leakage. Tumor growth and metastasis to other organs are monitored in live animals by using fluorescence. It is critical to use the appropriate filters for excitation and emission of GFP. The steps for the orthotopic implantation are detailed in this article so researchers can easily replicate the procedure in nude mice. The main steps of this protocol are preparation of the cell suspension, surgical implantation, and whole body fluorescent in vivo imaging. This orthotopic model is designed to investigate the efficacy of novel therapeutics on primary and metastatic tumors.

A procedure to implant green fluorescent protein-expressing pancreatic cancer cells (PANC-1 GFP) orthotopically into the pancreas of Balb-c Ola Hsd-Fox1nu mice to assess tumor progression and metastasis is presented here.

Pancreatic cancer remains one of the cancers for which survival has not improved substantially in the last few decades. Only 7% of diagnosed patients will ...

Murine Experimental Model of Original Tumor Development and Peritoneal Metastasis via Orthotopic Inoculation with Ovarian Carcinoma Cells

Epithelial ovarian carcinoma (EOC) is associated with a poor prognosis because it shows peritoneal dissemination. To improve the prognosis, it is important to control peritoneal dissemination. However, it is still unclear how tumor cells detach from primary lesions and attach to the mesothelium. The establishment of an appropriate animal model is needed to gain an understanding of the mechanism of peritoneal dissemination in vivo. In the current study, we introduce the process from the local injection of EOC cells into the murine ovarian surface to the development of metastasis, including the peritoneum and distant organs. Female nude mice (BALB/c nu/nu) at 8 weeks of age were used. Under a microscopic field of view, EOC cells (1 x 105 cells/&#181;l of medium-extracellular matrix (ECM)-based hydrogel/unilateral ovary/mouse) were injected into murine ovaries through a retroperitoneal approach from the dorsal flank. This proposed method is a less invasive procedure for the mouse and minimizes damage to the ovary. Here, we describe the methodological steps in the development of the original and metastatic tumor formation of EOC.

Murine Experimental Model of Original Tumor Development and Peritoneal Metastasis via Orthotopic Inoculation with Ovarian Carcinoma Cells

To clarify the multistep process of peritoneal dissemination of ovarian carcinoma, the current study presents a murine experimental model of original tumor development and peritoneal metastasis via orthotopic inoculation with these tumor cells.

Epithelial ovarian carcinoma (EOC) is associated with a poor prognosis because it shows peritoneal dissemination. To improve the prognosis, it is ...

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

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

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

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

Practical Considerations in Studying Metastatic Lung Colonization in Osteosarcoma Using the Pulmonary Metastasis Assay

The pulmonary metastasis assay (PuMA) is an ex vivo lung explant and closed cell culture system that permits researchers to study the biology of lung colonization in osteosarcoma (OS) by fluorescence microscopy. This article provides a detailed description of the protocol, and discusses examples of obtaining image data on metastatic growth using widefield or confocal fluorescence microscopy platforms. The flexibility of the PuMA model permits researchers to study not only the growth of OS cells in the lung microenvironment, but also to assess the effects of anti-metastatic therapeutics over time. Confocal microscopy allows for unprecedented, high-resolution imaging of OS cell interactions with the lung parenchyma. Moreover, when the PuMA model is combined with fluorescent dyes or fluorescent protein genetic reporters, researchers can study the lung microenvironment, cellular and subcellular structures, gene function, and promoter activity in metastatic OS cells. The PuMA model provides a new tool for osteosarcoma researchers to discover new metastasis biology and assess the activity of novel anti-metastatic, targeted therapies.

The goal of this article is to provide a detailed description of the protocol for the pulmonary metastasis assay (PuMA). This model permits researchers to study metastatic osteosarcoma (OS) cell growth in lung tissue using a widefield fluorescence or confocal laser-scanning microscope.

The pulmonary metastasis assay (PuMA) is an ex vivo lung explant and closed cell culture system that permits researchers to study the biology of lung ...

A Syngeneic Pancreatic Cancer Mouse Model to Study the Effects of Irreversible Electroporation

Pancreatic cancer (PC), a disease which kills approximately 40,000 patients each year in the US, has successfully evaded several therapeutic approaches including the promising immunotherapeutic strategies. Irreversible electroporation (IRE) is a non-thermal ablation technique that induces tumor cell death without destruction of adjacent collagenous structures, thus enabling the procedure to be performed in tumors very close to blood vessels. Unlike thermal ablation techniques, IRE results in gradual apoptotic cell death, along with immediate ablation induced necrosis, and is currently in clinical use for selected patients with locally advanced PC. An ablative, non-target specific procedure like IRE can induce a myriad of responses in the tumor microenvironment. A few studies have addressed the effects of IRE on tumor growth in other tumor types, but none have focused on PC. We have developed a syngeneic mouse model of PC in which subcutaneous (SQ) and orthotopic tumors can be successfully treated with IRE in a highly controlled setting, facilitating various longitudinal studies post procedure. This animal model serves as a robust system to study the effects of IRE and ways to improve the clinical efficacy of IRE.

Irreversible electroporation (IRE) is a non-thermal ablation technique used for the treatment of locally advanced pancreatic cancer. Being a relatively new technique, the effects of IRE on the tumor growth are poorly understood. We have developed a syngeneic mouse model that facilitates studying the effects of IRE on pancreatic cancer.

Pancreatic cancer (PC), a disease which kills approximately 40,000 patients each year in the US, has successfully evaded several therapeutic approaches ...

A Bioluminescent and Fluorescent Orthotopic Syngeneic Murine Model of Androgen-dependent and Castration-resistant Prostate Cancer

Orthotopic tumor modeling is a valuable tool for pre-clinical prostate cancer research, as it has multiple advantages over both subcutaneous and transgenic genetically engineered mouse models. Unlike subcutaneous tumors, orthotopic tumors contain more clinically accurate vasculature, tumor microenvironment, and responses to multiple therapies. In contrast to genetically engineered mouse models, orthotopic models can be performed with lower cost and in less time, involve the use of highly complex and heterogeneous mouse or human cancer cell lines, rather that single genetic alterations, and these cell lines can be genetically modified, such as to express imaging agents. Here, we present a protocol to surgically injecting a luciferase- and mCherry-expressing murine prostate cancer cell line into the anterior prostate lobe of mice. These mice developed orthotopic tumors that were non-invasively monitored in vivo and further analyzed for tumor volume, weight, mouse survival, and immune infiltration. Further, orthotopic tumor-bearing mice were surgically castrated, leading to immediate tumor regression and subsequent recurrence, representing castration-resistant prostate cancer. Although technical skill is required to carry out this procedure, this syngeneic orthotopic model of both androgen-dependent and castration-resistant prostate cancer is of great use to all investigators in the field.

The goal of this protocol is to demonstrate the intra-prostatic injection of prostate cancer cells, with subsequent castration. Orthotopic pre-clinical models of androgen-dependent and castration-resistant prostate cancer are critical to study the disease in the context of a clinically relevant tumor microenvironment and an immunocompetent host.

Orthotopic tumor modeling is a valuable tool for pre-clinical prostate cancer research, as it has multiple advantages over both subcutaneous and ...

Immunophenotyping of Orthotopic Homograft (Syngeneic) of Murine Primary KPC Pancreatic Ductal Adenocarcinoma by Flow Cytometry

Homograft (syngeneic) tumors are the workhorse of today&#39;s immuno-oncology (I/O) preclinical research. The tumor microenvironment (TME), particularly its immune-components, is vital to the prognosis and prediction of treatment outcomes, especially those of immunotherapy. TME immune-components are composed of different subsets of tumor-infiltrating immune cells assessable by multi-color FACS. Pancreatic ductal adenocarcinoma (PDAC) is among the deadliest malignances lacking good treatment options, thus an urgent and unmet medical need. One important reason for its non-responsiveness to various therapies (chemo-, targeted, I/O) has been its abundant TME, consisting of fibroblasts and leukocytes that protect tumor cells from these therapies. Orthotopically implanted PDAC is believed to more accurately recapture the TME of human pancreatic cancers than conventional subcutaneous (SC) models.
Homograft tumors (KPC) are transplants of mouse spontaneous PDAC originating from genetically engineered KPC-mice (KrasG12D/+/P53-/-/Pdx1-Cre) (KPC-GEMM). The primary tumor tissue is cut into small fragments (~2 mm3) and transplanted subcutaneously (SC) to the syngeneic recipients (C57BL/6, 7&#8211;9 weeks old). The homografts were then surgically orthotopically transplanted onto the pancreas of new C57BL/6 mice, along with SC-implantation, which reached tumor volumes of 300&#8211;1,000 mm3 by 17 days. Only tumors of 400&#8211;600 mm3 were harvested per approved autopsy procedure and cleaned to remove the adjacent non-tumor tissues. They were dissociated per protocol using a tissue dissociator into single-cell suspensions, followed by staining with designated panels of fluorescently-labeled antibodies for various markers of different immune cells (lymphoid, myeloid and NK, DCs). The stained samples were analyzed using multi-color FACS to determine numbers of immune cells of different lineages, as well as their relative percentage within tumors. The immune profiles of orthotopic tumors were then compared to those of SC tumors. The preliminary data demonstrated significantly elevated infiltrating TILs/TAMs in tumors over the pancreas, and higher B-cell infiltration into orthotopic rather than SC tumors.

Immunophenotyping of Orthotopic Homograft Syngeneic of Murine Primary KPC Pancreatic Ductal Adenocarcinoma by Flow Cytometry

The experimental procedure on the immunophenotyping of murine orthotopic PDAC homografts aims at profiling the tumor immuno-microenvironment. Tumors are orthotopically implanted via surgery. Tumors of 200&#8211;600 mm3 in size were harvested and dissociated to prepare single-cell suspensions, followed by multi-immune marker FACS analysis using different fluorescently-labeled antibodies.

Homograft (syngeneic) tumors are the workhorse of today&#39;s immuno-oncology (I/O) preclinical research. The tumor microenvironment (TME), particularly ...

Orthotopic Transplantation of Syngeneic Lung Adenocarcinoma Cells to Study PD-L1 Expression

The use of mouse models is indispensable for studying the pathophysiology of various diseases. With respect to lung cancer, several models are available, including genetically engineered models as well as transplantation models. However, genetically engineered mouse models are time-consuming and expensive, whereas some orthotopic transplantation models are difficult to reproduce. Here, a non-invasive intratracheal delivery method of lung tumor cells as an alternative orthotopic transplantation model is described. The use of mouse lung adenocarcinoma cells and syngeneic graft recipients allows studying tumorigenesis under the presence of a fully active immune system. Furthermore, genetic manipulations of tumor cells before transplantation makes this model an attractive time-saving approach to study the impact of genetic factors on tumor growth and tumor cell gene expression profiles under physiological conditions. Using this model, we show that lung adenocarcinoma cells express increased levels of the T-cell suppressor programmed death-ligand 1 (PD-L1) when grown in their natural environment as compared to cultivation in vitro.

Here we describe a minimally invasive syngeneic orthotopic transplantation model of mouse lung adenocarcinoma cells as a time- and cost-reducing model to study non-small cell lung cancer.

The use of mouse models is indispensable for studying the pathophysiology of various diseases. With respect to lung cancer, several models are available, ...

Acellular and Cellular Lung Model to Study Tumor Metastasis

It is difficult to isolate tumor cells at different points of tumor progression. We created an ex vivo lung model that can show the interaction of tumor cells with a natural matrix and continual flow of nutrients, as well as a model that shows the interaction of tumor cells with normal cellular components and a natural matrix. The acellular ex vivo lung model is created by isolating a rat heart-lung block and removing all the cells using the decellularization process. The right main bronchus is tied off and tumor cells are placed in the trachea by a syringe. The cells move and populate the left lung. The lung is then placed in a bioreactor where the pulmonary artery receives a continual flow of media in a closed circuit. The tumor grown on the left lung is the primary tumor. The tumor cells that are isolated in the circulating media are circulating tumor cells and the tumor cells in the right lung are metastatic lesions. The cellular ex vivo lung model is created by skipping the decellularization process. Each model can be used to answer different research questions.

Here, we present a protocol for an ex vivo lung cancer model that mimics the steps of tumor progression and helps to isolate a primary tumor, circulating tumor cells, and metastatic lesions.

It is difficult to isolate tumor cells at different points of tumor progression. We created an ex vivo lung model that can show the interaction of tumor ...

Patient-derived Orthotopic Xenograft Models for Human Urothelial Cell Carcinoma and Colorectal Cancer Tumor Growth and Spontaneous Metastasis

Cancer patients have poor prognoses when lymph node (LN) involvement is present in both high-grade urothelial cell carcinoma (HG-UCC) of the bladder and colorectal cancer (CRC). More than 50% of patients with muscle-invasive UCC, despite curative therapy for clinically-localized disease, will develop metastases and die within 5 years, and metastatic CRC is a leading cause of cancer-related deaths in the US. Xenograft models that consistently mimic UCC and CRC metastasis seen in patients are needed. This study aims to generate patient-derived orthotopic xenograft (PDOX) models of UCC and CRC for primary tumor growth and spontaneous metastases under the influence of LN stromal cells mimicking the progression of human metastatic diseases for drug screening. Fresh UCC and CRC tumors were obtained from consented patients undergoing resection for HG-UCC and colorectal adenocarcinoma, respectively. Co-inoculated with LN stromal cell (LNSC) analog HK cells, luciferase-tagged UCC cells were intra-vesically (IB) instilled into female non-obese diabetic/severe combined immunodeficiency (NOD/SCID) mice, and CRC cells were intra-rectally (IR) injected into male NOD/SCID mice. Tumor growth and metastasis were monitored weekly using bioluminescence imaging (BLI). Upon sacrifice, primary tumors and mouse organs were harvested, weighed, and formalin-fixed for Hematoxylin and Eosin and immunohistochemistry staining. In our unique PDOX models, xenograft tumors resemble patient pre-implantation tumors. In the presence of HK cells, both models have high tumor implantation rates measured by BLI and tumor weights, 83.3% for UCC and 96.9% for CRC, and high distant organ metastasis rates (33.3% detected liver or lung metastasis for UCC and 53.1% for CRC). In addition, both models have zero mortality from the procedure. We have established unique, reproducible PDOX models for human HG-UCC and CRC, which allow for tumor formation, growth, and metastasis studies. With these models, testing of novel therapeutic drugs can be performed efficiently and in a clinically-mimetic manner.

This protocol describes the generation of patient-derived orthotopic xenograft models by intra-vesically instilling high-grade urothelial cell carcinoma cells or intra-rectally injecting colorectal cancer cells into non-obese diabetic/severe combined immunodeficiency (NOD/SCID) mice for primary tumor growth and spontaneous metastases under the influence of lymph node stromal cells, which mimics the progression of human metastatic diseases.

Cancer patients have poor prognoses when lymph node (LN) involvement is present in both high-grade urothelial cell carcinoma (HG-UCC) of the bladder and ...