Determining Competitive Potential of Bone Metastatic Cancer Cells in the Murine Hematopoietic Stem Cell Niche

Abstract

The ability of cancer cells to compete with hematopoietic stem cells (HSCs) to target the bone marrow microenvironment, or the HSC niche, during the dissemination process is critical for the development of bone metastasis. Here, we describe the methods for testing the relative potential of cancer cells to compete with HSCs for occupancy of the HSC niche by measuring the peripheral blood level of engrafted HSCs by flow cytometry in mice after bone marrow transplantation and tandem cancer cell inoculation. This method is useful for determining the molecular mechanisms for the roles of HSCs in the regulation of bone metastases.

1. Introduction

Hematopoietic stem cells (HSCs) give rise to mature blood cells (e.g., myeloid and lymphoid lineages) during a process known as hematopoiesis. Besides differentiation potential, to maintain the homeostasis of hematopoiesis HSCs must be capable of self-renewal, similar to other stem cell types. Although most HSCs reside within the bone marrow, it is known that a portion of HSCs circulate throughout the peripheral blood. These circulating HSCs eventually home to the bone marrow [1]. It has been suggested that the specific microenvironment in the marrow, or the “niche,” is an indispensable component for controlling HSC differentiation, self-renewal, and homing [2]. Growing evidence suggests that osteoblasts and their progenitor cells are an important element of the HSC niche, although bone marrow stromal cells (endothelial cells, macrophages, fibroblasts, pericytes, and adipocytes) are also known to serve as HSC niche components [3, 4]. Indeed, chemotactic factors released from osteoblasts and other niche components [e.g., C-X-C motif chemokine 12 (CXCL12), and stem cell factor (SCF)] help to regulate essential HSC functions through the receptors expressed on HSCs [e.g., C-X-C chemokine receptor type 4 (CXCR4), c-Kit] [2, 5-7].

Interestingly, it has recently been revealed that disseminated tumor cells (DTCs) use bone homing mechanisms similar to HSCs to gain access to the marrow [8]. Moreover, once in the marrow, DTCs directly compete with HSCs to occupy the HSC niche and survive within the bone microenvironment [8]. The HSC niche also induces DTCs to undergo cellular dormancy, a state similar to HSC quiescence. It has been suggested that these dormant DTCs acquire treatment resistance and escape from immune surveillance, eventually developing into full-blown bone metastases [9, 10]. Therefore, the HSC niche is a critical player in bone metastatic progression. However, little is known as to the molecular mechanisms whereby DTCs interact with HSCs to establish bone metastases.

In this protocol, we describe the methods necessary to determine the potential of cancer cells to compete with HSCs for the bone marrow HSC niche by combining cancer cell inoculation techniques with bone marrow transplantation. The workflow of these methods to address specific questions in HSC niche competition is outlined in Fig. 1. These models can be further utilized to understand the mechanisms that control bone marrow stem cell niche homing of highly competitive cancer cells in order to promote the development of novel therapeutic strategies for bone metastasis.

Fig. 1.

Hematopoietic stem cell (HSC) niche competition assay workflow. (a) In the first model, a cancer cell line is implanted subcutaneously, grown for 3 weeks and then resected. Five days later, a bone marrow transplant is performed. (b) In the second model, a bone marrow transplant is performed after total body irradiation, and then a cancer cell line is inoculated systemically with intracardiac injection. In both models, the readout for HSC niche competition is HSC engraftment analysis by flow cytometry of peripheral blood. Additionally, a survival outcome can be used in the second model

2. Materials

2.1. Bone Marrow Transplant

1. Five- to Eight-week-old mice whose leukocytes express the cluster of differentiation (CD)45.2 or Ly5.2, such as C57BL/6J (Jackson Laboratory)

2. Five- to Eight-week-old mice whose leukocytes express CD45.1 or Ly5.1, such as B6.SJL-Ptprc^a Pepc^b/BoyJ (Jackson Laboratory)

3. CO₂ euthanasia chamber

4. Biosafety cabinet

5. 70% Ethanol solution

6. Sterile surgical instruments (scissors, scalpel, and forceps)

7. 3 mL Syringe with 25-g needle (Becton Dickinson)

8. Dulbecco’s phosphate-buffered saline without calcium or magnesium (DPBS): 20 mg/L KCl, 200 mg/L KH₂PO₄, 8.00 g/L NaCl, 2.16 g/L Na₂HPO₄, and pH 7.2–7.4

9. Fetal bovine serum (FBS)

10. Fluorescence activated cell sorting (FACS) buffer: 5% FBS, DPBS

11. 10-cm Tissue culture (TC) dish

12. 40-μm Mesh strainer

13. 50-mL Conical tube

14. Hemocytometer and trypan blue solution, 0.4%

15. 0.9% Sterile saline solution

16. Isoflurane vaporizer

17. A gamma cell 40 cesium source

18. 28-g Insulin syringe

2.2. Cancer Cell Inoculation

1. Cancer cell of interest

2. DPBS

3. Biosafety cabinet

4. 40-μm Mesh strainer

5. 50-mL Conical tube

6. Hemocytometer and trypan blue solution, 0.4%

7. 1.5-mL Microcentrifuge tubes

8. 0.9% Sterile saline solution

9. Five- to Eight-week-old mice

10. Electrical clippers or a depilatory agent

11. Isoflurane vaporizer

12. Povidone-iodine, 5%

13. Sterile alcohol prep pads

14. 28-g Insulin syringe

15. Gelfoam® gelatin sponges (Pfizer)

16. 10-cm TC plate

17. Sterile surgical gauze

18. Sterile surgical instruments (scissors and forceps)

19. Surgical wound clips

20. Heating pad

2.3. HSC Engraftment Analysis

1. Heating pad

2. Sterile razor blades

3. 5-mL Round bottom tubes

4. Peripheral blood collection buffer: 5 μM ethylenediaminete-traacetic acid (EDTA), DPBS

5. Red blood cell (RBC) lysis buffer (BioLegend)

6. DPBS

7. FACS buffer

8. Anti-CD45.1 and CD45.2 antibodies (BioLegend)

9. Flow cytometer

3. Methods

3.1. Bone Marrow Transplant

1. Euthanize and douse the skin of the donor mice in 70% ethanol (see Note 1). Remove the hind limbs and store in a 10-cm TC dish containing ice cold sterile DPBS on ice.

2. In a biosafety cabinet, separate the femur from the tibia and remove the excess tissue thoroughly using a scalpel. Be careful to maintain the integrity of the bone in order to maintain sterility of the bone marrow. Store the bare bones in a 10-cm TC dish containing ice cold sterile DPBS.

3. Remove the epiphyses from the femurs and tibias using a scalpel.

4. Flush the marrow from the diaphysis into a new sterile 10-cm TC dish with ice cold sterile FACS buffer using a 25-g needle and 3-mL syringe.

5. Pass the marrow several times through the syringe to dissociate the marrow pieces, and then pass the marrow suspension through a 40-μm mesh strainer into a 50-ml conical tube.

6. Determine the bone marrow cell number using a hemocytometer and the trypan blue exclusion assay and then pellet the bone marrow cells by centrifugation (350 × g for 5 min).

7. Aspirate the supernatant and resuspend the bone marrow cells in ice cold sterile 0.9% saline at a concentration of 2 × 10⁵ cells/100 μL and store on ice (see Note 2).

8. If the intracardiac injection model is to be used, the recipient mice must be irradiated prior to bone marrow transplantation. Radiation dose is animal model dependent (see Note 3).

9. Anesthetize the recipient mouse and perform retro-orbital sinus injection of 2 × 10⁵ bone marrow cells/100 μL per each mouse.

10. Allow the animal to recover from anesthesia in a clean cage placed on a heating pad.

11. The engraftment of donor cells will be followed using flow cytometry as described in Subheading 3.3.

3.2. Cancer Cell Inoculation

1. Wash and harvest the cells to be tested under sterile conditions (see Note 4).

2. Pass the cells through a 40-μm cell strainer into a 50-mL conical tube and determine cell number as before with a hemocytometer and the trypan blue exclusion assay and then pellet the cells by centrifugation (180 × g for 5 min).

3. Aspirate the supernatant and resuspend the cells in ice cold sterile 0.9% saline to an appropriate concentration and volume and transfer to a sterile microcentrifuge tube and store on ice (see Note 5).

4. If performing subcutaneous implant of the inoculum, cut the gelatin foam into 3 mm³ squares (scaffolds) using surgical scissors and with forceps place them in a 10-cm TC dish containing ice cold 0.9% saline. This must be performed under sterile conditions.

5. Anesthetize the recipient mouse using isoflurane inhalation and prepare it for the surgery to be performed, either subcutaneous implant and resection (Subheading 3.2.1) or intracardiac injection (Subheading 3.2.2) (see Note 6).

3.2.1. Subcutaneous Implant and Resection

1. If the recipient mouse has fur on its back, it must first be removed using electrical clippers or a depilatory agent.

2. Place the anesthetized mouse in the prone position and disinfect the skin of the back using a povidone-iodine scrub and alcohol prep pad.

3. Lift up a tent of skin midway up the back using forceps and make a 1-cm horizontal incision using blunt-tipped surgical scissors.

4. Use blunt-tipped surgical scissors to create a 1–2-cm subcutaneous pouch rostral to the incision site.

5. Blot a 3-mm³ gelatin scaffold on sterile surgical gauze to remove excess liquid and then saturate it with inoculum and place it deep into the subcutaneous pouch using forceps.

6. Release the implant and withdraw the forceps without disturbing the placement of the implant.

7. Close the incision using a surgical wound clip.

8. Allow the animal to recover from anesthesia in a clean cage placed on a heating pad.

9. Remove the wound clip after 7 days or when the incision has healed.

10. Three weeks after implantation, anesthetize the mouse using isoflurane inhalation. If the animal has fur on its back, it must first be removed using electrical clippers or a depilatory agent.

11. Place the anesthetized mouse in the prone position and disinfect the skin of the back using a povidone-iodine scrub and alcohol prep pad.

12. Make an incision near the subcutaneous implant with blunt-tipped surgical scissors.

13. Lift up a tent of skin near the incision/implant and use blunt-tipped surgical scissors to reopen the subcutaneous pouch.

14. Dissect the implant away from the skin and surrounding tissue and remove it entirely.

15. Close the incision using a surgical wound clip.

16. Allow the animal to recover from anesthesia in a clean cage placed on a heating pad.

17. Remove the wound clip after 7 days or when the incision has healed.

3.2.2. Intracardiac Injection

1. If the animal has fur on its chest, it must first be removed using electrical clippers or a depilatory agent.

2. Place the anesthetized mouse in the supine position and disinfect the skin of the chest using a povidone-iodine scrub and alcohol prep pad.

3. Load 100 μL of inoculum into an insulin syringe.

4. Use the index and middle finger of your nondominant hand to secure the limbs of the mouse (see Note 7).

5. Using your dominant hand, insert the needle perpendicular about 7–8 mm into the injection point, 1–2 mm left of the midline halfway between the clavicle and xiphoid process.

6. Using your nondominant hand, pull up very slightly on the plunger being careful not to move the needle. If the needle is in the correct position (left ventricle), you will see a pulse of bright red blood flush into the inoculum.

7. Inject the inoculum slowly over about 30 s. During the injection, periodically confirm that the needle is still in the left ventricle by pulling up on the plunger slightly to check for the signature pulse of bright red blood (see Note 8).

8. Gently remove the needle.

9. Allow the animal to recover from anesthesia in a clean cage placed on a heating pad.

3.3. HSC Engraftment Analysis

1. Place the cage of animals to be analyzed on the heating pad and allow it to warm up for 10 min.

2. Place a mouse on top of the wire food rack and thread its tail through to the bottom.

3. Make a small nick in the tail vein perpendicular to the tail with a razor blade and collect the blood droplets in a 5-mL round bottom tube containing 4-mL peripheral blood collection buffer (see Note 9). Use the lip of the tube to catch the blood droplets and be sure that they mix with the collection buffer by tapping the tube gently in order to prevent coagulation.

4. Apply pressure to the wound with a piece of gauze and confirm hemostasis of the tail wound before placing the mouse back in its cage.

5. Centrifuge the collection tubes (350 × g for 5 min).

6. Decant the supernatant and resuspend the pellets in 500 μL RBC lysis buffer and incubate at room temperature for 5 min (see Note 10).

7. Fill each tube with 4 mL DPBS and then centrifuge them (350 × g for 5 min).

8. Decant the supernatant and resuspend the pellets in 500 μL FACS buffer containing anti-CD45.1 and CD45.2 antibodies (see Note 11).

9. Incubate on ice in the dark for 1 h.

10. Fill each tube with 4 mL DPBS and then centrifuge them (350 × g for 5 min).

11. Decant the supernatant and resuspend the pellets in 500 μL FACS buffer.

12. Analyze the samples for expression of host hematopoietic cells versus expression of donor hematopoietic cells by distinguishing CD45.1 and CD45.2 positive cells by flow cytometry (see Note 12).

4. Notes

1. The donor mice must express a different CD45 allele than the recipient mice in order to determine HSC engraftment. We recommend performing the HSC engraftment analysis (Subheading 3.3) on representative naïve donor and recipient mice before transplantation to verify distinguishability by flow cytometry.

2. Instead of mouse donor cells, it is possible to use human HSCs at the same concentration, as long as the recipient animal is immunocompromised. In this case, we would recommend using NOD.Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ (NSG) mice as a recipient.

3. If greater HSC engraftment is desired, the recipient mice can be irradiated immediately before transplantation. For lethal irradiation levels in immunocompetent mice, we recommend a total body dose of 1140 cGy (two 570 cGy doses, 3 h apart). For lethal irradiation levels in immunocompromised mice, we recommend a total body dose of 700 cGy (two 350 cGy doses, 3 h apart). If human HSC transplant is to be performed in immunocompromised mice, we recommend a sublethal total body dose of 300 cGy (two 150 cGy doses, 3 h apart).

4. In order to remove the variable of implanted cell growth after intracardiac injection, it is possible to irradiate the cell line to be tested immediately before harvest and implantation. In our studies using prostate cancer cells, we first irradiated them with 8 Gy (two 4 Gy doses, 3 h apart). However, irradiation dose will be cell dependent. In this case, a dose that causes cell growth arrest but not apoptosis should be used.

5. The concentration of the cell suspension depends on the aggressiveness of the cell line and the inoculation to be performed. As a general rule, the subcutaneous implant will require 10 μL of inoculum whereas the intracardiac injection requires 100 μL. In our studies using prostate cancer cells, we used 2 × 10⁵ cells/10 μL for the subcutaneous implant and 2 × 10⁶ cells/100 μL for the intracardiac injection. Very aggressive cell lines may not require as many cells, but these are good starting points. For ease of use, we recommend creating individual tubes containing the inoculum required for each subcutaneous implant.

6. The model chosen depends on the hypothesis to be tested. The subcutaneous implant model relies on the ability of the cell line to disseminate from the primary implant to the bone on its own. While this closely models what happens clinically, the number of disseminated cells in the bone is often quite low and is entirely cell line dependent. The intracardiac model floods the circulation with many cells which increases “dissemination” to the bone, with the caveat that it does not accurately model the dissemination process. Also, using the intracardiac model it is possible to employ an engraftment readout using animal survival if the recipient animals are first lethally irradiated. In this model, longer survival times signify greater HSC engraftment.

7. It is essential that the mouse is flat on its back with its internal organs centered. It is common for the mouse to shift its weight naturally to favor one side over the other, but if this happens it is difficult to reliably locate the left ventricle with the needle. When the mouse is in the correct position, use your hand to secure it in place without using excessive pressure. Pressure on the abdomen causes shifting of the internal organs, again making the injection more challenging.

8. If the action of pulling up on the plunger creates a vacuum and not a pulse of blood, the needle has either drifted too far up or too deep and is no longer in the left ventricle. If this happens, slightly push the needle deeper or withdraw it until the pulse of blood is observed before proceeding with the injection.

9. Create a nick about halfway down the tail so that if the initial cut fails to produce a blood droplet it is possible to make another nick rostral to the initial cut. Do not squeeze or attempt to milk the tail, as this can cause contamination with lymphatic fluid. In our experience, five drops (~10–50 μL) is sufficient for analysis.

10. This process can be repeated until the pellet is no longer red, or can be skipped entirely if there is concern for lost cells during RBC lysis.

11. It is recommended to use antibodies conjugated to fluorophores that have emission spectrums that do not overlap much, such as fluorescein isothiocyanate (FITC) and allophycocyanin (APC). Concentrations and multiplexing of antibodies require in lab optimization.

12. It is useful to perform HSC engraftment analysis many times over a period of several weeks. We have found that hematopoietic cells of transplanted origin can be detected up to 16 weeks after transplant.