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

Summary

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.

Abstract

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.

Introduction

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 lesions^3,4. 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 disease^5,6. 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 osteosarcoma^9,10,11. 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 “artifactual metastasis” that do not recapitulate the complete metastatic cascade from an established primary bone tumor that metastasizes to the lungs^8,13. 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.

Protocol

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

1. Grow cell lines in accordance with the laboratory’s standard cell culture protocols for traditional cell culture or injection into mice. Standard protocols used here are growth in Dulbecco’s modified Eagle’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.
2. Grow cells in either vented tissue culture flasks or 6-well tissue culture plates at 37 °C in 5% CO₂.
3. 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.

2. Animals

1. 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.
2. 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).
3. For xenograft experiments, use immunodeficient athymic nude mice based on the cell line being incompatible with an intact mouse immune system under normal conditions.
4. 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.
5. House animals at standard densities depending on the institution’s husbandry policies.

3. Subcutaneous tumors

1. Harvest cell lines from culture by trypsinization and resuspend in sterile phosphate-buffered saline (PBS).
2. 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.
3. 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 µL) of sterile PBS. Keep the cells on ice until injection.
4. 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 µL). Keep the cells on ice until further use.
5. 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.
6. 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.
7. Clean the prepared area with a 70% ethanol swab prior to injection of the cell suspension.
8. 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.
9. 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.
10. 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’ 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.
11. Euthanize mice bearing subcutaneous tumors after three to four weeks by CO₂ inhalation followed by cervical dislocation. Follow institution’s acceptable policies for mouse euthanasia.
12. 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.
13. 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 – 0.25 mm³ up to 1 mm x 1 mm x 1 mm – 1 mm³) with a #15 scalpel blade (with or without a scalpel blade handle).
14. 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.
15. 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 –80 °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 –80 °C.
    NOTE: It has been previously reported that snap frozen tumors will not engraft and grow in vivo⁸.

4. Surgical implantation of subcutaneous tumor fragments

1. Bring fresh or cryopreserved fragments of subcutaneous tumor to room temperature in complete growth medium prior to surgical implantation.
2. 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.
3. Remove hair on the right knee joint and proximal tibia of the hindlimb with depilating solution to minimize the potential trauma to the skin.
4. Scrub the prepared area with surgical antiseptic. Scrub first with a 70% ethanol swab and then scrub with alternating chlorhexidine and saline scrub.
5. Visualize the proximal tibia as the region just distal to the knee joint while flexing and extending the joint.
6. 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.
7. 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.
8. 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.
9. 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.
10. Recover the mice on a heating pad in individual cages until ambulatory.

5. Serial and end point assessment

1. Anesthetize mice using isoflurane in oxygen anesthesia as described previously.
2. 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.
3. 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.
4. 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.

Results

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, µCT, or µMRI changes in the proximal tibia associated with the bone phenotype of the cell line of interest (osteolytic, osteob...

Discussion

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...

Materials

List of materials used in this article

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