The authors acknowledge the critical contribution of Dr. Beth Chaffee, DVM, PhD, DACVP to the development of this technique.&#160;&#160;

All described animal experiments were approved by the institutional animal care and use committee of University of Cambridge, Cambridge, UK.
1. Preparation of cell lines
<ol>
	<li>Grow cell lines in accordance with the laboratory&#8217;s standard cell culture protocols for traditional cell culture or injection into mice. Standard protocols used here are growth in Dulbecco&#8217;s modified Eagle&#8217;s medium containing 10% fetal bovine serum (FBS), L-glutamine, and penicillin/streptomycin (hereafter known as complete growth medium). 
	NOTE: In this experiment, Abrams osteosarcoma cells are used in Balb/c Foxn1 nu/nu mice. For breast cancer studies, 4T1 cells in Balb/c mice and EO771 cells in C57BL/6 mice are used.</li>
	<li>Grow cells in either vented tissue culture flasks or 6-well tissue culture plates at 37 &#176;C in 5% CO2.</li>
	<li>Passage the cell line of interest and prepare the cells for injection when the cells reach a confluency commonly used with injection of these cells into mice.</li>
</ol>
2. Animals
<ol>
	<li>Use Balb/c Foxn1 nu/nu mice at least 6-8 weeks of age for subcutaneous tumor generation to ensure that the animals are beyond the rapid growth phase and have achieved adulthood and skeletal maturity.</li>
	<li>Use either male or female mice. Make exceptions when selecting hormone-responsive cell lines (e.g., breast cancer cells in female mice and prostate cancer cells in male mice).</li>
	<li>For xenograft experiments, use immunodeficient athymic nude mice based on the cell line being incompatible with an intact mouse immune system under normal conditions.</li>
	<li>For allograft experiments using murine cell lines, this is also recommended based on dissimilar mouse genetic and immune backgrounds. However, for syngeneic experiments, use animals of the same strain as the cell line of interest.</li>
	<li>House animals at standard densities depending on the institution&#8217;s husbandry policies.</li>
</ol>
3. Subcutaneous tumors
<ol>
	<li>Harvest cell lines from culture by trypsinization and resuspend in sterile phosphate-buffered saline (PBS).</li>
	<li>Assess cell viability and determine the cell density by the trypan blue exclusion method. Use a hemocytometer or an automated cell counter to count the cells. A minimum cell viability of 90% is to be used for injection into mice to create subcutaneous tumors.</li>
	<li>Adjust the cell density to inject 1-2 x 105 cells in a final volume of 0.1 to 0.15 mL (100 to 150 &#181;L) of sterile PBS. Keep the cells on ice until injection.</li>
	<li>Alternatively, pellet cells by centrifuging at 800 x g for 5 min. Discard the supernatant and re-suspend the pelleted cells in undiluted sterile basement membrane matrix medium to obtain 1-2 x 105 cells in a final volume of 0.1 to 0.15 mL (100 to 150 &#181;L). Keep the cells on ice until further use.</li>
	<li>Anesthetize mice to be used for subcutaneous tumor growth with isoflurane in oxygen anesthesia. Use an induction dose of 5% isoflurane in 2 L/min oxygen and a maintenance dose of 2-3% isoflurane in 2 L/min oxygen. Check for the lack of blink or pedal reflexes before proceeding further. 
	NOTE: Isoflurane is an inhalational anesthetic. Use isoflurane in a well-ventilated area with appropriate scavenging and free gas collection systems. Please consult with the institutional veterinary staff to develop a plan for anesthesia induction, maintenance, and monitoring, and ensure that the laboratory staff have appropriate training in anesthesia monitoring and the handling of inhalant anesthetic agents.</li>
	<li>Remove hair from the dorsal region of the thorax or abdomen of anesthetized mice with depilating solution or with an electric clipper. Depilating solution is preferred to minimize potential trauma to the skin. Skip this step if using athymic nude mice.</li>
	<li>Clean the prepared area with a 70% ethanol swab prior to injection of the cell suspension.</li>
	<li>Use a 1 mL tuberculin syringe with a 27 G needle to inject cells subcutaneously over the dorsal region of the thorax or abdomen, not to be impacted by movement of the shoulder blades. Alternatively, inject the cells subcutaneously as a suspension in commercially available extracellular matrix. 
	NOTE: Injection in commercially available extracellular matrix will limit migration of the cell suspension in the subcutaneous space because these matrices solidify at room temperature.</li>
	<li>Recover the mice on a heating pad in individual cages until ambulatory. Mice can then be placed in their normal cages with clean, dry bedding.</li>
	<li>Monitor the size of the subcutaneous tumor overlying the dorsal thorax or abdomen with a caliper and measure body weights weekly to ensure that the subcutaneous tumors do not ulcerate or mice meet early removal criteria as established by the institutions&#8217; animal care and use committee. A maximum tumor size of 15 mm in any dimension is recommended to reduce the risk of skin ulceration or central tumor necrosis. 
	NOTE: Consult local guidelines to determine maximum permissible tumor size/volume.</li>
	<li>Euthanize mice bearing subcutaneous tumors after three to four weeks by CO2 inhalation followed by cervical dislocation. Follow institution&#8217;s acceptable policies for mouse euthanasia.</li>
	<li>Harvest subcutaneous tumors using aseptic surgical technique. Sterilize the skin overlying the tumor as before with 70% ethanol after removal of the hair (if applicable). Incise through the skin overlying the tumor with a #15 scalpel blade (with or without a scalpel blade handle). Sharply dissect the tumor from the surrounding attached soft tissues with a pair of sterile surgical scissors.</li>
	<li>Place the tumor in 6-well tissue culture plates containing complete growth medium and mince into multiple small fragments of pre-determined size (~ 0.6 mm x 0.6 mm x 0.6 mm &#8211; 0.25 mm3 up to 1 mm x 1 mm x 1 mm &#8211; 1 mm3) with a #15 scalpel blade (with or without a scalpel blade handle).</li>
	<li>Maintain tumor fragments in sterile complete growth medium at room temperature until the time of intratibial implantation. For cell lines that carry luciferase or fluorescent reporter genes, use ex-vivo bioluminescent or fluorescent imaging to confirm tumor viability ahead of intratibial implantation into mice.</li>
	<li>For cryopreservation, place multiple fragments in the same cryovial in complete growth medium supplemented with 20% FBS and 10% dimethyl sulfoxide (DMSO). Freeze gradually using a commercial cryopreservation system at &#8211;80 &#176;C and store long-term in liquid nitrogen. Preserve tumor fragments for subsequent analysis, but not for future implantation, by snap freezing using liquid nitrogen immersion. Store these frozen tumor fragments for long-term at &#8211;80 &#176;C. 
	NOTE: It has been previously reported that snap frozen tumors will not engraft and grow in vivo8.</li>
</ol>
4. Surgical implantation of subcutaneous tumor fragments
<ol>
	<li>Bring fresh or cryopreserved fragments of subcutaneous tumor to room temperature in complete growth medium prior to surgical implantation.</li>
	<li>Anesthetize mice of the strain of interest using isoflurane in oxygen anesthesia as described in Section 3. Check for the lack of pedal reflexes before proceeding. Administer subcutaneous buprenorphine at a dose of 0.02-0.05 mg/kg to provide peri-operative analgesia. This can be repeated every 6-8 h in the post-operative period, if needed.</li>
	<li>Remove hair on the right knee joint and proximal tibia of the hindlimb with depilating solution to minimize the potential trauma to the skin.</li>
	<li>Scrub the prepared area with surgical antiseptic. Scrub first with a 70% ethanol swab and then scrub with alternating chlorhexidine and saline scrub.</li>
	<li>Visualize the proximal tibia as the region just distal to the knee joint while flexing and extending the joint.</li>
	<li>Create a 3-4 mm incision at the level of the proximal tibia on the medial aspect of the limb with a #15 scalpel blade (with or without a scalpel blade handle). Incise through the skin and subcutaneous tissue to expose the medial cortex of the proximal tibia.</li>
	<li>Apply gentle pressure with the tip of a 25 G needle, while also rotating the tip, to create a small hole in the medial cortex of the proximal tibia. Make this hole approximately 2 mm distal to the knee joint at a point equidistant between the cranial and caudal tibial cortexes. Select the needle size depending on the size of the tumor fragments.</li>
	<li>Use sterile forceps to pick up and insert the tumor fragments into the medullary cavity of the proximal tibia. Use a 27 to 30 G needle to manipulate the tumor fragment into the medullary canal. Depending on the size of the tumor fragments, implant a minimum of 0.5 mm3 total tumor volume into each tibia. This may require implantation of 1 or more tumor fragments depending on the size of tumor fragments created. 
	NOTE: Modifications to prevent or limit displacement of the graft outside of the bone would be placement of bone wax or bone cement in the bone defect or either gel foam or a subcutaneous fat graft over the hole in the bone.</li>
	<li>Appose the skin edges with sterile liquid tissue adhesive or a single skin suture. Do not use wound clips in this site. Take caution if using fluorescence imaging in the post-operative period, since both tissue adhesives and suture have the potential to fluoresce.</li>
	<li>Recover the mice on a heating pad in individual cages until ambulatory.</li>
</ol>
5. Serial and end point assessment
<ol>
	<li>Anesthetize mice using isoflurane in oxygen anesthesia as described previously.</li>
	<li>Evaluate tibial tumor growth non-invasively by either weekly digital radiography, bioluminescence, or fluorescence imaging (if using cells expressing luciferase or a fluorescent reporter gene). Caliper measurements of the limb at the site of implantation can also be performed in awake mice.</li>
	<li>In addition to the traditional monitoring of tumor-bearing mice (body weight, activity level, respiratory rate, grooming, posture, mentation, and behavior) monitor mice weekly for signs of hind limb lameness, swelling, and surgical site infection.</li>
	<li>Monitor the skin surgical wound for the first 10-14 days for excessive redness, swelling, draining, and wound dehiscence until the skin wound is healed. After 4-5 weeks, evaluate mice in accordance with the study outcome evaluation either alive or following euthanasia.</li>
</ol>

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Dr. Hildreth was funded by the&#160;NIH under Award Number&#160;K01OD026527.&#160; Content is solely&#160;the responsibility of the authors&#160;and does not necessarily&#160;represent the official views of the&#160;NIH.

This report documents our model to create primary bone tumors or bone metastasis following the intratibial implantation of a tumor allograft. We believe that there are several critical steps in this process. A safe anesthetic plane should be established for both subcutaneous injection of the tumor cell suspension and intratibial placement of the resultant tumor fragments. There should be sterile preparation of the surgical site for both removal of the subcutaneous allograft and intratibial placement of the allograft. Tum...

Mouse models of human and animal disease are becoming increasingly popular in biomedical research. The utility of using mice in this context is that their anatomy and physiology are very similar to humans. They have a relatively short gestation period and time in post-natal life to achieve maturity, and are largely associated with a relatively low cost and ease of housing, albeit increasing costs of development or purchase are associated with greater degrees of genetic modification, immunodeficiency, and/or humanization1. Use of inbred strains results in a largely uniform animal population prior to study inclusion. A complete knowledge of their genome suggests a high degree of similarity to humans. Orthologous molecular targets for many disease processes have been identified in the mouse genome and there is now an extensive library of mouse-specific reagents that are easily obtainable. Therefore, they provide the opportunity for relatively high-throughput analysis in a more rapid and less expensive manner when compared to larger animal models1. In addition, with the advent of genetic editing strategies that allows for the overexpression or deletion of certain genes either globally or in a cell type specific manner and/or constitutively or in an inducible manner, they represent a very biologically useful model system for the investigation of human and animal diseases2.
Cancer is one field in which mouse models have great utility. Genetic mouse models of cancer rely on modulation of the expression of either oncogenes or tumor suppressor genes, alone or in combination, for cells to undergo oncogenic transformation. The injection of primary or established tumor cell lines into mice is also performed. The introduction of either cell lines or tissues from humans or other animal species, including mice, remains the most widely used model of cancer in vivo. The use of cells and tissues from dissimilar species (xenografts) in immunocompromised mice is most commonly performed2. However, the use of allograft tumor cells or tissues where both the host and recipient are of the same species allows for the interaction with an intact immune system when combined with the same host mouse strain in syngeneic systems3.
Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions3,4. These tumors compromise bone structure, increasing the risk of pathologic fracture, and leave patients with limited treatment options. Primary bone tumors metastasize to distant organs, with some types capable of spreading to other skeletal sites. In breast cancer patients, bone is the most common site of first metastasis and the most frequent first site of presentation of metastatic disease5,6. In addition, disseminated tumor cells (DTCs) are present in the bone marrow prior to the diagnosis of, and predict the development of, metastasis in other organs7. Therefore, it is believed that cancer cells present in bone are the source of cells that ultimately metastasize to other organ systems. Many mouse models of solid tumor metastasis exist that develop metastasis predominantly in the lung and lymph nodes, and depending on the tumor type and injection technique, potentially other organ systems3. However, mouse models of bone metastasis are lacking that dependably, reproducibly produce site specific skeletal metastasis and develop bone metastasis before mice reach early removal criteria from primary tumor burden or metastasis to other organs. We have reported a model of the primary bone tumor osteosarcoma that relies on the surgical implantation of a solid tumor allograft into the proximal tibia of mice8. Bone tumors formed in 100% of mice and 88% developed pulmonary metastasis. This incidence of metastasis exceeds what is commonly reported clinically in people (~20-50%), but is of great interest since the lung is the most common site of metastasis for osteosarcoma9,10,11. While this model is advantageous in modeling primary bone tumors, it also has great utility in modeling bone metastasis from other osteotropic solid tumors such as breast, lung, prostate, thyroid, hepatic, renal, and gastrointestinal tumors.
The rationale for the development of this model was to develop an alternative to the traditional intra-osseous injection typically into the proximal tibia or distal femur to model primary bone tumors or bone metastasis12. Our primary goal was to alleviate a known limitation of this technique i.e., tumor embolization of the lung. This results in the engraftment of these embolic tumor cells and &#8220;artifactual metastasis&#8221; that do not recapitulate the complete metastatic cascade from an established primary bone tumor that metastasizes to the lungs8,13. This would also be the situation when an established bone metastasis spreads to a distant site. In addition, this technique was, also, developed to produce a model of bone metastasis that would ensure a greater incidence of engraftment and growth of tumors in bone and at a uniform site when compared with orthotopic or intravascular injection techniques. This model has distinct advantages over these described techniques. This model involves controlled, consistent delivery of tumor cells into the bone. It, also, avoids artifactual lung metastasis following pulmonary embolization and establishes a baseline uniform study population. There is the benefit of site-specific tumors with this model without the risk of early removal criteria resulting from primary tumors or metastasis to other organs. Lastly, this model has great utility for modification, including the use of patient-derived xenografts.
The model presented has similarities to direct cell suspension injection into bone following a surgical approach followed by either injection through the cortex or delivery into the marrow cavity after making a small defect in the cortex (with or without reaming out the medullary cavity)8,14,15,16,17. However, the implantation of a tumor allograft makes this technique distinctly different. Therefore, the purpose of this report was to demonstrate this model of primary bone tumors and bone metastasis from solid tumors, which overcomes many limitations of previously described models. Research groups with experience in cell culture, mouse models, mouse anesthesia and surgery, and mouse anatomy are well equipped to reproduce our technique to model primary bone tumors or bone metastasis in mice.

<table>
	<tbody>
		<tr>
			<td>#15 scalpel blade</td>
			<td>Henry Schein Ltd.</td>
			<td>75614</td>
			<td>None</td>
		</tr>
		<tr>
			<td>6-well tissue culture plates</td>
			<td>Thermo Fisher Scientific</td>
			<td>10578911</td>
			<td>Used for mincing tumor pieces. Can also be used for cell culture</td>
		</tr>
		<tr>
			<td>Abrams osteosarcoma cell line</td>
			<td>Not applicable</td>
			<td>Not applicable</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Anesthesia machine with isoflurane vaporiser and oxygen tank(s)</td>
			<td>VetEquip</td>
			<td>901805</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Animal weighing scale</td>
			<td>Kent Scientific</td>
			<td>SCL- 1015</td>
			<td>None</td>
		</tr>
		<tr>
			<td>BALB/c nude mouse (nu/nu)</td>
			<td>Charles River Ltd.</td>
			<td>NA</td>
			<td>6-8 weeks of age. Male or female mice</td>
		</tr>
		<tr>
			<td>Bone cement</td>
			<td>Depuy Synthes</td>
			<td>160504</td>
			<td>Optional use instead of bone wax</td>
		</tr>
		<tr>
			<td>Bone wax</td>
			<td>Ethicon</td>
			<td>W31G</td>
			<td>Optional</td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Animalcare Ltd.</td>
			<td>N/A</td>
			<td>Buprecare 0.3 mg/ml Solution for Injection for Dogs and Cats</td>
		</tr>
		<tr>
			<td>Carbon dioxide euthanasia station</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Should be provided within animal facility</td>
		</tr>
		<tr>
			<td>Cell culture incubator set at 37 &#176;C and 5% carbon dioxide</td>
			<td>Heraeus</td>
			<td>Various</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Chlorhexidine surgical scrub</td>
			<td>Vetoquinol</td>
			<td>411412</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Cryovials (2 ml)</td>
			<td>Thermo Scientific Nalgene</td>
			<td>5000-0020</td>
			<td>Optional if cryopreserving tumor fragments</td>
		</tr>
		<tr>
			<td>D-luciferin (Firefly), potassium salt</td>
			<td>Perkin Elmer</td>
			<td>122799</td>
			<td>Optional if cell line of interest has a bioluminescent reporter gene</td>
		</tr>
		<tr>
			<td>Digital caliper</td>
			<td>Mitutoyo</td>
			<td>500-181-30</td>
			<td>Can be manual</td>
		</tr>
		<tr>
			<td>Digital microradiography cabinet</td>
			<td>Faxitron Bioptics, LLC</td>
			<td>MX-20</td>
			<td>Optional to evaluate bone response to tumor growth</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma Aldrich</td>
			<td>1371171000</td>
			<td>Optional if cryopreserving tumor fragments</td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s modified Eagle&#8217;s medium</td>
			<td>Thermo Fisher Scientific</td>
			<td>11965092</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Ethanol (70%)</td>
			<td>Sigma Aldrich</td>
			<td>2483</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Fetal bovine serum</td>
			<td>Thermo Fisher Scientific</td>
			<td>26140079</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Forceps, Dumont</td>
			<td>Fine Science Tools, Inc.</td>
			<td>11200-33</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Freezer (&#8211; 80 &#176;C)</td>
			<td>Sanyo</td>
			<td>MDF-794C</td>
			<td>Optional if cryopreserving or snap freezing tumor fragments</td>
		</tr>
		<tr>
			<td>Hemocytometer</td>
			<td>Thermo Fisher Scientific</td>
			<td>11704939</td>
			<td>Can also use automated cell counter, if available</td>
		</tr>
		<tr>
			<td>Hypodermic needles (27 gauge)</td>
			<td>Henry Schein Ltd.</td>
			<td>DIS55510</td>
			<td>May also use 25G (DIS55509) and 30G (Catalog DIS599) needles</td>
		</tr>
		<tr>
			<td>Ice</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Ideally small pieces in a container for syringe and cell suspension storage</td>
		</tr>
		<tr>
			<td>Iris scissors</td>
			<td>Fine Science Tools, Inc.</td>
			<td>14084-08</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Henry Schein Ltd.</td>
			<td>1182098</td>
			<td>None</td>
		</tr>
		<tr>
			<td>IVIS Lumina III bioluminescence/fluorescence imaging system</td>
			<td>Perkin Elmer</td>
			<td>CLS136334</td>
			<td>Optional if cell line of interest has bioluminescent or fluorescent reporter genes</td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>Thermofisher scientifc</td>
			<td>25030081</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Liquid nitrogen</td>
			<td>British Oxygen Corporation</td>
			<td>NA</td>
			<td>Optional if cryopreserving or snap freezing tumor fragments</td>
		</tr>
		<tr>
			<td>Liquid nitrogen dewar, 5 litres</td>
			<td>Thermo Fisher Scientific</td>
			<td>TY509X1</td>
			<td>Optional if cryopreserving tumor fragments</td>
		</tr>
		<tr>
			<td>Matrigel&#174; Matrix GFR, LDEV-Free, 5 ml</td>
			<td>Corning Life Sciences</td>
			<td>356230</td>
			<td>Optional. Also available in 10 ml size (354230)</td>
		</tr>
		<tr>
			<td>Microcentrifuge</td>
			<td>Thermo Fisher Scientific</td>
			<td>75002549</td>
			<td>Pellet cells at 1200 rpm for 5-6 minutes</td>
		</tr>
		<tr>
			<td>Mr. Frosty freezing containiner</td>
			<td>Fisher Scientific</td>
			<td>10110051</td>
			<td>Optional if cryopreserving tumor fragments</td>
		</tr>
		<tr>
			<td>NAIR Hair remover lotion/oil</td>
			<td>Thermo Fisher Scientific</td>
			<td>NC0132811</td>
			<td>Can alternatively use an electric clipper with fine blade</td>
		</tr>
		<tr>
			<td>Penicillin/streptomycin</td>
			<td>Sigma-Aldrich</td>
			<td>P4333</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Scalpel handle, #7 Short</td>
			<td>Fine Science Tools, Inc.</td>
			<td>10007-12</td>
			<td>User preference as long as it accepts #15 scalpel blade</td>
		</tr>
		<tr>
			<td>Small animal heated pad</td>
			<td>VetTech</td>
			<td>HE006</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>GT Vision Ltd.</td>
			<td>H600BV1</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Sterile phosphate-buffered saline (PBS)</td>
			<td>Thermo Fisher Scientific</td>
			<td>10010023</td>
			<td>Use for injections and also as part of the surgical scrub, alternating with chlorhexidine</td>
		</tr>
		<tr>
			<td>Tissue adhesive (sterile)</td>
			<td>3M Corporation</td>
			<td>84-1469SB</td>
			<td>Can alternatively use non-absorbable skin suture (6-0 size)</td>
		</tr>
		<tr>
			<td>Trypan blue</td>
			<td>Thermo Fisher Scientific</td>
			<td>5250061</td>
			<td>None</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA</td>
			<td>Thermo Fisher Scientific</td>
			<td>25300054</td>
			<td>Use 0.05%-0.25%</td>
		</tr>
		<tr>
			<td>Tuberculin syringe (1 ml with 0.1 ml gradations)</td>
			<td>Becton Dickinson</td>
			<td>309659</td>
			<td>Slip tip preferred over Luer</td>
		</tr>
		<tr>
			<td>Vented tissue culture flasks, T-75</td>
			<td>Corning Life Sciences</td>
			<td>CLS3290</td>
			<td>Can also use smaller or larger flasks, as needed</td>
		</tr>
	</tbody>
</table>

modeling primary bone tumors and bone metastasis with solid tumor graft implantation into bone

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

A positive result would be associated with tumor engraftment and progressive tumor growth over time. Depending on the tumor type, intraosseous tumor growth may be associated with progressive hind limb lameness, but many tumors do not cause lameness despite signs of attendant bone disease. Successful engraftment was documented with advanced imaging, whereby there would be progressive radiographic, &#181;CT, or &#181;MRI changes in the proximal tibia associated with the bone phenotype of the cell line of interest (osteolyt...

Watch this Scientific Journal Video about Modeling Primary Bone Tumors and Bone Metastasis with Solid Tumor Graft Implantation into Bone at JoVE.com

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

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

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

modeling-primary-bone-tumors-bone-metastasis-with-solid-tumor-graft

Research

JoVE Journal

Cancer Research

2.6K Views.  University of Alabama at Birmingham. 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.

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

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

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

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

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

Dual Effects of Melanoma Cell-derived Factors on Bone Marrow Adipocytes Differentiation

The crosstalk between bone marrow adipocytes and tumor cells may play a critical role in the process of bone metastasis. A variety of methods are available for studying the significant crosstalk; however, a two-dimensional transwell system for coculture remains a classic, reliable, and easy way for this crosstalk study. Here, we present a detailed protocol that shows the coculture of bone marrow adipocytes and melanoma cells. Nevertheless, such a coculture system could not only contribute to the study of cell signal transductions of cancer cells induced by bone marrow adipocytes, but also to the future mechanistic study of bone metastasis which may reveal new therapeutic targets for bone metastasis.

Here, we present a reliable and straightforward two-dimensional (2D) coculture system for studying the interaction between tumor cells and bone marrow adipocytes, which reveals a dual effect of melanoma cell-derived factors on the bone marrow adipocytes differentiation and also poses a classic method for the mechanistic study of bone metastasis.

The crosstalk between bone marrow adipocytes and tumor cells may play a critical role in the process of bone metastasis. A variety of methods are ...

Multimodal Bioluminescent and Positronic-emission Tomography/Computational Tomography Imaging of Multiple Myeloma Bone Marrow Xenografts in NOG Mice

Multiple myeloma (MM) tumors engraft in the bone marrow (BM) and their survival and progression are dependent upon complex molecular and cellular interactions that exist within this microenvironment. Yet the BM microenvironment cannot be easily replicated in vitro, which potentially limits the physiologic relevance of many in vitro and ex vivo experimental models. These issues can be overcome by utilizing a xenograft model in which luciferase (LUC)-transfected 8226 MM cells will specifically engraft in the mouse skeleton. When these mice are given the appropriate substrate, D-luciferin, the effects of therapy on tumor growth and survival can be analyzed by measuring changes in the bioluminescent images (BLI) produced by the tumors in vivo. This BLI data combined with positronic-emission tomography/computational tomography (PET/CT) analysis using the metabolic marker 2-deoxy-2-(18F)fluoro-D-glucose (18F-FDG) is used to monitor changes in tumor metabolism over time. These imaging platforms allow for multiple noninvasive measurements within the tumor/BM microenvironment.

Here we use bioluminescent, X-ray, and positron-emission tomography/computed tomography imaging to study how inhibiting mTOR activity impacts bone marrow-engrafted myeloma tumors in a xenograft model. This allows for physiologically relevant, non-invasive, and multimodal analyses of the anti-myeloma effect of therapies targeting bone marrow-engrafted myeloma tumors in vivo.

Multiple myeloma (MM) tumors engraft in the bone marrow (BM) and their survival and progression are dependent upon complex molecular and cellular ...

Measuring Bone Remodeling and Recreating the Tumor-Bone Microenvironment Using Calvaria Co-culture and Histomorphometry

Bone is a connective tissue constituted of osteoblasts, osteocytes, and osteoclasts and a mineralized extracellular matrix, which gives it its strength and flexibility and allows it to fulfill its functions. Bone is continuously exposed to a variety of stimuli, which in pathological conditions can deregulate bone remodeling. To study bone biology and diseases and evaluate potential therapeutic agents, it has been necessary to develop in vitro and in vivo models.
This manuscript describes the dissection process and culture conditions of calvarias isolated from neonatal mice to study bone formation and the bone tumor microenvironment. In contrast to in vitro and in vivo models, this ex vivo model allows preservation of the three-dimensional environment of the tissue as well as the cellular diversity of the bone while culturing under defined conditions to simulate the desired microenvironment. Therefore, it is possible to investigate bone remodeling and its mechanisms, as well as the interactions with other cell types, such as the interactions between cancer cells and bone.
The assays reported here use calvarias from 5-7 day old BALB/C mice. The hemi-calvarias obtained are cultured in the presence of insulin, breast cancer cells (MDA-MB-231), or conditioned medium from breast cancer cell cultures. After analysis, it was established that insulin induced new bone formation, while cancer cells and their conditioned medium induced bone resorption. The calvarial model has been successfully used in basic and applied research to study bone development and cancer-induced bone diseases. Overall, it is an excellent option for an easy, informative, and low-cost assay.

The ex vivo culture of bone explants can be a valuable tool for the study of bone physiology and the potential evaluation of drugs in bone remodeling and bone diseases. The presented protocol describes the preparation and culture of calvarias isolated from newborn mice skulls, as well as its applications.

Bone is a connective tissue constituted of osteoblasts, osteocytes, and osteoclasts and a mineralized extracellular matrix, which gives it its strength ...

Three-Dimensional Bone Extracellular Matrix Model for Osteosarcoma

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with diagnostic or prognostic difficulties. OS comprises genotypically and phenotypically heterogeneous cancer cells. Bone microenvironment elements are proved to account for tumor heterogeneity and disease progression. Bone extracellular matrix (BEM) retains the microstructural matrices and biochemical components of native extracellular matrix. This tissue-specific niche provides a favorable and long-term scaffold for OS cell seeding and proliferation. This article provides a protocol for the preparation of BEM model and its further experimental application. OS cells can grow and differentiate into multiple phenotypes consistent with the histopathological complexity of OS clinical specimens. The model also allows visualization of diverse morphologies and their association with genetic alterations and underlying regulatory mechanisms. As homologous to human OS, this BEM-OS model can be developed and applied to the pathology and clinical research of OS.

The bone extracellular matrix (BEM) model for osteosarcoma (OS) is well established and shown here. It can be used as a suitable scaffold for mimicking primary tumor growth in vitro and providing an ideal model for studying the histologic and cytogenic heterogeneity of OS.

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with ...

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

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

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

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

Establishment and Analysis of Three-Dimensional (3D) Organoids Derived from Patient Prostate Cancer Bone Metastasis Specimens and their Xenografts

Three-dimensional (3D) culture of organoids from tumor specimens of human patients and patient-derived xenograft (PDX) models of prostate cancer, referred to as patient-derived organoids (PDO), are an invaluable resource for studying the mechanism of tumorigenesis and metastasis of prostate cancer. Their main advantage is that they maintain the distinctive genomic and functional heterogeneity of the original tissue compared to conventional cell lines that do not. Furthermore, 3D cultures of PDO can be used to predict the effects of drug treatment on individual patients and are a step towards personalized medicine. Despite these advantages, few groups routinely use this method in part because of the extensive optimization of PDO culture conditions that may be required for different patient samples. We previously demonstrated that our prostate cancer bone metastasis PDX model, PCSD1, recapitulated the resistance of the donor patient&rsquo;s bone metastasis to anti-androgen therapy. We used PCSD1 3D organoids to characterize further the mechanisms of anti-androgen resistance. Following an overview of currently published studies of PDX and PDO models, we describe a step-by-step protocol for 3D culture of PDO using domed or floating basement membrane (e.g., Matrigel) spheres in optimized culture conditions. In vivo stitch imaging and cell processing for histology are also described. This protocol can be further optimized for other applications including western blot, co-culture, etc. and can be used to explore characteristics of 3D cultured PDO pertaining to drug resistance, tumorigenesis, metastasis and therapeutics.

Establishment and Analysis of Three-Dimensional 3D Organoids Derived from Patient Prostate Cancer Bone Metastasis Specimens and their Xenografts

Three-dimensional cultures of patient BMPC specimens and xenografts of bone metastatic prostate cancer maintain the functional heterogeneity of their original tumors resulting in cysts, spheroids and complex, tumor-like organoids. This manuscript provides an optimization strategy and protocol for 3D culture of heterogeneous patient derived samples and their analysis using IFC.

Three-dimensional (3D) culture of organoids from tumor specimens of human patients and patient-derived xenograft (PDX) models of prostate cancer, referred ...

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

Machine learning (ML) algorithms permit the integration of different features into a model to perform classification or regression tasks with an accuracy exceeding its constituents. This protocol describes the development of an ML algorithm to predict the growth of breast cancer bone macrometastases in a rat model before any abnormalities are observable with standard imaging methods. Such an algorithm can facilitate the detection of early metastatic disease (i.e., micrometastasis) that is regularly missed during staging examinations.
The applied metastasis model is site-specific, meaning that the rats develop metastases exclusively in their right hind leg. The model&#8217;s tumor-take rate is 60%&#8211;80%, with macrometastases becoming visible in magnetic resonance imaging (MRI) and positron emission tomography/computed tomography (PET/CT) in a subset of animals 30 days after induction, whereas a second subset of animals exhibit no tumor growth.
Starting from image examinations acquired at an earlier time point, this protocol describes the extraction of features that indicate tissue vascularization detected by MRI, glucose metabolism by PET/CT, and the subsequent determination of the most relevant features for the prediction of macrometastatic disease. These features are then fed into a model-averaged neural network (avNNet) to classify the animals into one of two groups: one that will develop metastases and the other that will not develop any tumors. The protocol also describes the calculation of standard diagnostic parameters, such as overall accuracy, sensitivity, specificity, negative/positive predictive values, likelihood ratios, and the development of a receiver operating characteristic. An advantage of the proposed protocol is its flexibility, as it can be easily adapted to train a plethora of different ML algorithms with adjustable combinations of an unlimited number of features. Moreover, it can be used to analyze different problems in oncology, infection, and inflammation.

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 (ML) algorithms permit the integration of different features into a model to perform classification or regression tasks with an accuracy ...

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

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

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

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

Isolation and Characterization of Bone Marrow Microenvironment in Myeloid Neoplasms

Identifying Bone Marrow Microenvironmental Populations in Myelodysplastic Syndrome and Acute Myeloid Leukemia

The bone marrow microenvironment consists of distinct cell populations, such as mesenchymal stromal cells, endothelial cells, osteolineage cells, and fibroblasts, which provide support for hematopoietic stem cells (HSCs). In addition to supporting normal HSCs, the bone marrow microenvironment also plays a role in the development of hematopoietic stem cell disorders, such as myelodysplastic syndromes (MDS) and acute myeloid leukemia (AML). MDS-associated mutations in HSCs lead to a block in differentiation and progressive bone marrow failure, especially in the elderly. MDS can often progress to therapy-resistant AML, a disease characterized by a rapid accumulation of immature myeloid blasts. The bone marrow microenvironment is known to be altered in patients with these myeloid neoplasms. Here, a comprehensive protocol to isolate and phenotypically characterize bone marrow microenvironmental cells from murine models of myelodysplastic syndrome and acute myeloid leukemia is described. Isolating and characterizing changes in the bone marrow niche populations can help determine their role in disease initiation and progression and may lead to the development of novel therapeutics targeting cancer-promoting alterations in the bone marrow stromal populations.

Author Spotlight: Analyzing Bone Marrow Microenvironment in Murine Hematological Malignancies 

Here a detailed protocol to isolate and characterize bone marrow microenvironmental populations from murine models of myelodysplastic syndromes and acute myeloid leukemia is presented. This technique identifies changes in the non-hematopoietic bone marrow niche, including the endothelial and mesenchymal stromal cells, with disease progression.

The bone marrow microenvironment consists of distinct cell populations, such as mesenchymal stromal cells, endothelial cells, osteolineage cells, and ...