A "Patient-Like" Orthotopic Syngeneic Mouse Model of Hepatocellular Carcinoma Metastasis

Summary

The metastatic spread of cancer is the major cause of cancer-related deaths. We provide an in-depth description of our survival surgery methodology for establishing a “patient-like” orthotopic syngeneic mouse model system for studying the mechanisms of metastasis in solid organ tumors.

Abstract

The majority of cancer-related deaths are caused by the metastasis of the cancer rather than the primary tumor itself. Yet, the underlying mechanisms of cancer metastasis are still unclear. Animal models are essential for elucidating the mechanisms and for evaluating novel strategies for the treatment of metastatic cancers. Here, an in-depth description of a “patient-like” orthotopic syngeneic mouse model for exploring the mechanisms of metastasis of solid organ tumors is provided. The survival surgical implantation of BNL 1ME A.7R.1 mouse hepatocellular carcinoma cells directly into the liver (the organ of origin) of the inbred wild-type immune competent laboratory mouse strain, BALB/c is described. The success and reproducibility of this methodology recommends it for widespread use in elucidating the biological mechanisms of solid organ cancer metastasis.

Introduction

Hepatocellular carcinoma (HCC) is one of the most lethal cancers, with poor prognosis and low life expectancy. Nearly all cancer-related deaths are due to the metastatic spread of the disease from the originating organ to additional distant organs^2-6.HCC progression is a complex process. Therefore, being able to model the tumor microenvironment that is naturally found in metastatic HCC in animal models can prove to be a successful and useful way to reveal relevant mechanisms in humans.⁷Unfortunately, the mechanisms of cancer metastasis are still largely unclear. Therefore, there is a need to establish animal models that will enable us to elucidate the underlying molecular mechanisms of metastases of cancers such as HCC^3,5.

Mouse model systems are a very useful approach for delineating mechanisms and evaluating novel strategies for treatment of metastatic human cancers^5,8. The various mouse model systems that presently exist are a testament to efforts of researchers to correctly depict the complexity of the disease^5,8,9.

Ogunwobi and colleagues recently used a survival surgical approach to demonstrate establishment of a novel orthotopic syngeneic mouse model for the study of metastasis in HCC³. Their work established a “patient-like” mouse model that recapitulates features of aggressive and metastatic HCCs³. They further demonstrated that this mouse model system can be used to study the biology of circulating tumor cells, and that this holds potential for gaining novel insights into the mechanisms of cancer metastasis³.

The aim of this paper is to describe in detail the methodology used in establishing this “patient-like” orthotopic syngeneic mouse model of HCC metastasis³. The methodology of how to implant BNL 1ME A.7R.1 mouse HCC cells directly into the liver (the organ of origin) of the inbred wild-type immune competent laboratory mouse strain, BALB/c³ using survival surgery will be described. Unlike other mouse xenograft tumor models where human tumor cells are implanted into immune deficient mice, this system is syngeneic and is, therefore, suitable for studying the role of the immune system in tumor metastasis^3,5,8. This approach will likely gain widespread use for studying the mechanisms of metastasis in solid organ cancers.

Protocol

Ethics Statement
All animal studies were approved by the Institutional Animal Care and Use Committee (IACUC) of Hunter College of The City University of New York.

Note: Eight BALB/c mice were used in this experimental procedure. Five BALB/c mice were implanted with 5 X 10⁶ BNL 1ME A.7R.1 mouse hepatocellular carcinoma cells to produce the primary tumors. Three BALB/c mice with no implantations were used as controls. BNL 1ME A.7R.1 is a murine hepatocellular carcinoma cell line that was derived by chemical transformation from the murine non-tumorigenic hepatocyte cell line BNL CL.2. BNL 1ME A.7R.1 was obtained from the American Type Culture Collection (ATCC).

1. Pre-surgery

1. Ensure that mice for all survival surgeries are healthy. To ensure that mice are healthy, inspect signs of dehydration by tenting of the skin. Do not use mice with any evidence of disease or distress in these experiments.
2. Weigh mice to ensure they are not underweight. Ensure that the weight is at least 16-20 grams.
3. Prepare 5×10⁶ BNL 1ME A.7R.1 mouse HCC cells for implantation per mouse:
   1. Aspirate media from a 10 ml 60% - 70% confluent cell culture dish of BNL 1ME A.7R.1 mouse HCC cells. Wash twice with 5 ml PBS. Add 2 ml of trypsin to the dish and incubate at 37 °C in a humidified incubator with air and 5% CO₂ for 5 min.
   2. Subsequently, pipette 10 µl trypsinized cells into a manual hemocytometer for cell counting under a microscope at magnification of 10X. Count the total number of cells found in the four large corner squares and multiply the dilution factor by the total number of cells. Then, divide by the number of squares counted and multiply by 10⁶ for the cell concentration (cells per ml).
   3. Using the calculation, collect 5 million cells per mouse in sterile 1.5 ml microcentrifuge tubes. Label 1.5 ml microcentrifuge tubes.
4. Resuspend the 5×10⁶ BNL 1ME A.7R.1 mouse HCC cells in 100 µl of phosphate buffered saline (PBS). Keep these cells on ice till implantation in surgery.
5. Perform activities required for preparation of mice for surgery such as anesthesia induction and hair clipping away from sterile equipment and materials to avoid contamination.
6. Induce general anesthesia in mice by inhalation of isoflurane using an isoflurane vaporizer along with oxygen at a constant concentration flow rate of 1 L per min (approx. 2.1%). Note: The system is self-reliant, and includes a precision vaporizer, an oxygen supply and a charcoal canister that is linked to a breathing circuit needed for gas/anesthetic scavenging.
7. Ensure that anesthesia is adequate to prevent the animal from undergoing pain during the surgery. Perform toe-pinch and observe animal for non-responsiveness to confirm anesthesia.
8. At that time, switch the valve line, seal to the box and open to the nose cone.
9. As soon as the mouse is anesthetized, apply a tiny amount of an ophthalmic ointment to the corneas as a preventive measure from drying and damage.
10. Subsequently, move the animal to the preparation area, with the nose cone properly placed and connected (for maintenance anesthesia). Prepare the site of the surgical incision for surgery by depilation at this preparation area.
11. Ensure that the area of surgery is at a location free of all other activity.
12. Ensure that the work surface is sturdy and appropriately sanitized, unsoiled and clutter-free before the surgery. Place a sterile piece of linen or surgical tray over it.
13. Ensure that the operative field is available to only persons carrying out the surgery. Ensure that the surgical room is cleaned and swept with an appropriate cleaner and sprayed with an appropriate disinfectant.
14. Ensure that it is also positioned away from doors and windows as air currents can spread dust leading to contamination of the site.
15. Perform the survival surgery with sterile instruments. Either soak instruments in a disinfectant according to manufacturer’s recommendation or autoclave. Use basic sterile supplies that are ready for use.
16. Separate instruments according to function to help prevent contamination of sterile areas.
17. Shave approximately 1 inch around the site of incision (midline incision) to access the liver of the animal. Remove the hair from the animal away from the site of surgery.
18. After shaving, scrub the area with a soapy disinfectant (betadine surgical scrub) and wash the excess disinfectant away with sterile PBS solution. Perform this several times, with the entire scrub process lasting approximately 5 min. Remove and discarded the used gloves.
19. Afterward, change to protective clothing such as surgical gown, scrub top, and face mask. Next, sanitize hands with proper scrub solution before putting on surgical gloves. Note: This is a necessary step to prevent shedding and contaminating the surgical site with potential contaminants.

2. Surgery

1. Provide an external heat source for warmth throughout the entire period of anesthesia. The animal may experience some hypothermia because anesthesia causes disturbances of thermoregulation; therefore, maintain the body temperature.
2. Lay the mouse on its back throughout the surgery.
3. Fix the fore limbs, hind limbs and tail of the mouse to the surgery table with pieces of surgical tapes.
4. Make a straight line incision immediately inferior to the xyphisternum.
5. Use a sterile self-retaining surgical retractor to expose the liver.
6. With a syringe having a 20G needle, slowly and carefully inject 100 µl of PBS containing 5×10⁶ BNL 1ME A.7R.1 mouse HCC cells into the right lobe of the liver.
7. After removing the needle, apply a sterile cotton swab for at least 1 min to stop any potential bleeding due to high vascularity of the liver.
8. Close the subcutaneous tissue with an absorbable suture followed by skin closure using surgical clips. Subsequently, clean the wound once again with betadine scrub. Note: This will ensure the closure of the surgical wound.
9. Subcutaneously give 100 µl of normal saline.

3. Post-surgery

1. Immediately after surgery, transfer the mouse to a clean cage with a post-surgery identification card to notify caretakers of the mouse condition. Provide food and water. Note: In our experience, a mouse that has just undergone surgery will fare best if housed alone with wet food and water for the initial 24-hr period post-surgery.
2. Keep the animal warm with a heat lamp that generates heat not surpassing 38 ºC. Use a thermometer to measure the temperature until the mouse is fully recovered.
3. House the animal in a cage preferably with paper bedding to ensure comfort.
4. After initial recovery, monitor the animal for signs of discomfort, bleeding, pain and distress. Give buprenorphine (0.05 -0.1 mg/kg) to relieve pain. Closely monitor the activity, behavior and appearance, as well as water and food consumption for approximately 2 hr. If ambulatory, move the animal back to the area for regular normal housing of mice and continue to monitor at 6-12 hr intervals for 24 hr post-surgery. Perform a weight check if desired.
5. However if the animal is having issues recuperating, take appropriate supportive care actions. If mice are unable to recover, humanely euthanize mice via CO₂ asphyxiation and cervical dislocation. Follow approved institutional guidelines for euthanasia.
6. Post-surgery, check for clinical evidence of HCC which is usually observable as significant reduction in body condition score. Note: At this point, a primary tumor has likely developed in the liver and metastatic deposits have likely developed in the lungs.
7. At this point, sacrifice all mice (experimental and control) involved in the experiment by humane euthanasia. Subject the mice to carbon dioxide asphyxiation: very slowly release carbon dioxide into the chamber over a period of 5 -10 min. This will cause respiratory arrest.

Results

The livers of Balb/c mice were implanted with 5×10⁶ BNL 1ME A.7R.1 mouse HCC cells. Clinical evidence of HCC development was observable 63 days post-surgery. Consequently, mice were humanely euthanized. Necropsy was performed on the euthanized mice. The lungs and liver were resected and a careful gross examination was performed to identify macroscopic tumors. In one mouse, a superficial tumor was observed on the surface of the liver with some attachment to the abdominal wall (Figure 1A). In a ...

Discussion

In this article an in-depth description of the method is given that was recently reported by Ogunwobi and colleagues of successful establishment of an orthotopic syngeneic mouse model of HCC metastasis (Figure 2)³. The take rate of tumors for this procedure is generally high. We have previously observed a tumor take rate of 100%³. However, the take rate can be variable depending on the competence of the investigator. In a recent experiment performed by a new graduate student under t...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Micro Dissecting Tweezer	Roboz	(RS-5040)	Tip 0.20 x 0.12 mm
Graefe Micro Dissecting Forceps, serrated curved tip	Roboz	(RS-5111)	1 X 2 teeth, curved tip width 0.6 mm
Micro Dissecting Retractor-Agricola (3 by 3 prongs)	Roboz	(RS-6501)	Blunt 3 X 3 prongs, depth 4 mm, spread 25 mm
Micro Dissecting Retractor-Goldstein (3 by 3 prongs)	Roboz	(RS-6503)	Blunt, 3 X 3 prongs, depth 4 mm, spread 19 mm
Jameson Caliper (Measures tumors)	Roboz	(RS-6466)	80 mm/3 inch scale, chrome plated
Micro Dissecting Scissors, large ring sissors, straight and sharp	Roboz	(RS-5852)	23 mm blades, length 4 inches, flat shanks and large rings
Scalpel with blades, for delicate dissecting procedures	Roboz	(RS-9861-36)
Scalpel handle	Roboz	(RS-9884)	Solid
Littauer Stitch Sissors	Roboz	(RS-7074)	Length 4.5 inches
Brown needle holder, for easy suture tying	Roboz	(RS-7960)	Convex jaw, fine serrations
Reflex 7MM wound clips with reflex 7 clip applier	Roboz	(RS-9262)	safe, secure alternative method of wound closure
Instrument tray and lid	Roboz	RT-1350S
Mini-clipper with detachable blade	Roboz	(RS-5903)
Germinator 500 (the Germ Terminator) Dry Sterilizer	Roboz	Ds-400, Ds-401, DS-501	For fast decontamination of micro-dissecting instruments.   Instruments decontaminate within 15 sec.
Microinjection needles	VWR	BD305125	25G Needle