Generation of a Liver Orthotopic Human Uveal Melanoma Xenograft Platform in Immunodeficient Mice

Podsumowanie

Orthotopic human liver metastatic uveal melanoma xenograft mouse models were created using surgical orthotopic implantation techniques with patient-derived tumor chunk and needle injection techniques with cultured human uveal melanoma cell lines.

Streszczenie

In recent decades, subcutaneously implanted patient-derived xenograft tumors or cultured human cell lines have been increasingly recognized as more representative models to study human cancers in immunodeficient mice than traditional established human cell lines in vitro. Recently, orthotopically implanted patient-derived tumor xenograft (PDX) models in mice have been developed to better replicate features of patient tumors. A liver orthotopic xenograft mouse model is expected to be a useful cancer research platform, providing insights into tumor biology and drug therapy. However, liver orthotopic tumor implantation is generally complicated. Here we describe our protocols for the orthotopic implantation of patient-derived liver-metastatic uveal melanoma tumors. We cultured human liver metastatic uveal melanoma cell lines into immunodeficient mice. The protocols can result in consistently high technical success rates using either a surgical orthotopic implantation technique with chunks of patient-derived uveal melanoma tumor or a needle injection technique with cultured human cell line. We also describe protocols for CT scanning to detect interior liver tumors and for re-implantation techniques using cryopreserved tumors to achieve re-engraftment. Together, these protocols provide a better platform for liver orthotopic tumor mouse models of liver metastatic uveal melanoma in translational research.

Wprowadzenie

Uveal melanoma is the most common intraocular malignant tumor among adults in the western world. During the past 50 years, the incidence of uveal melanoma (5.1 cases per million) has remained stable in the United States^1,2. Uveal melanoma arises from melanocytes in the iris, ciliary body, or choroid, and it is an extremely lethal disease when it develops metastasis. The death rate of patients with uveal melanoma metastasis was 80% at 1 year and 92% at 2 years after initial diagnosis of the metastases. The time between diagnosis of metastases and death is typically short, less than 6 months, without regards to therapy^3,4. The cancer spreads through the blood and tends to dominantly metastasize to the liver (89-93%)^4,5. An effective mouse model is urgently needed for further investigation of liver-metastatic uveal melanoma. For translational research, there is a clear demand to generate a liver-localized metastatic uveal melanoma mouse model.

Patient-derived tumor xenograft (PDX) mouse models are expected to provide individualized medicine strategies. These models might be predictive of clinical outcomes, be useful for preclinical drug evaluation, and be used for biological studies of tumors⁶. Representative PDX models are ectopically tumor-implanted xenograft mice, which have tumor at subcutaneous sites. Most researchers can do surgery for subcutaneous implantation without special practice^7,8. They can also monitor subcutaneous tumors easily. Although subcutaneous PDX models became popular in the research phase, they have some hurdles in moving to practical use. Subcutaneous implantation forces patient-derived tumors to engraft at a different microenvironment from the tumor origin, so that it leads to engraftment failure and slow tumor growth ^{9,10,11,12,13,14}. Orthotopic engraftment may be a more ideal and rational approach for a PDX model because it uses the same organ as the original tumor^15,16.

Recently, we developed protocols for surgical orthotopic implantation techniques of patient-derived liver-metastatic uveal melanoma tumors and needle injection techniques with a cultured human liver-metastatic uveal melanoma cell line in NOD.Cg-Prkdc^scidIl2rg^tm1Wjl/SzJ (NSG) mice^17,18. The protocols result in consistently high technical success rates. We also established CT scanning techniques that are useful to detect interior liver tumors, and we developed re-implantation of cryopreserved tumors in the PDX platform. We found that uveal melanoma tumor xenograft models maintain the characteristics of the original patient liver tumor, including their histopathological and molecular features. Together, these techniques provide a better platform for liver orthotopic tumor models for uveal melanoma in translational research.

Protokół

Patients enrolled in the study should provide written consent allowing the use of discarded surgical samples for research purposes and genetic studies, according to an Institutional Review Board-approved protocol. This protocol was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health and approved by the Institutional Animal Care and Use Committee (IACUC).

1. Collection of Fresh Patient-derived Tumor Tissue

1. Obtain patient-derived tumor tissue from surgery or a needle biopsy in a hospital operating room.
2. Put the tumor tissue in a 100 mL container containing Hanks' balanced salt solution (HBSS) solution on ice.
3. Transfer the tissue into a sterile hood (Biosafety level 2) in a laboratory.
4. Proceed to step 2 as soon as possible.
   NOTE: For safety reasons, exclude patients with known HIV or Hepatitis B or C infections.

2. Processing of Fresh Patient-derived Tumor Tissue

1. Put the tissue in a 50 mL tube containing phosphate-buffered saline (PBS) on ice. For washing the tissue, add PBS in the tube and discard PBS from the tube twice.
2. Transfer the tissue into a Petri dish containing PBS on ice.
3. Using sterile forceps and scissors, remove the necrotic parts of the tissue. Keep the tissue moist and cold during steps 2.3 to 2.5. For needle biopsy samples, skip step 2.3 and 2.5, and do not cut the samples.
   NOTE: The necrotic tissue often breaks apart easily when touched.
4. Cut the tissue into 1 mm³ cubes for surgical liver implantation.
5. Cut the rest of the tissue into 2 mm cubes in the Petri dish.
6. Transfer them to a 1.7 mm microtube with 4% formalin for histological analysis and to another tube for genomic and proteomic analysis.
7. Put the microtubes in a liquid nitrogen jar with liquid nitrogen. Transfer the tubes to a -80 °C freezer for permanent storage.
   NOTE: The time between sample removal from the patient and tissue processing should not exceed 30 min.

3. Surgical Liver Implantation with Patient-derived Tumor Tissue

1. Spray all objects coming into the hood for surgery with 70 % ethyl alcohol.
   NOTE: This includes surgical instruments, heating pads, and anesthesia machines.
2. Measure the weight of a cotton swab and fabric sheet.
3. Anesthetize a mouse with a 3–5% isoflurane vaporizer by placing it in the induction chamber.
4. Once the mouse is fully anesthetized, place it in supine position on a heating pad. Place the isoflurane cone on the mouse's snout to inhale 1.5–3% isoflurane for maintenance of anesthesia.
   NOTE: The mouse needs to be on the heating pad during the entire procedure. Lack of heating may cause hypothermia.
5. Confirm proper anesthesia by no reaction when the foot of the mouse is pricked with ultrafine forceps.
6. Inject buprenorphine (0.6 mg/kg) subcutaneously on the flank using a 27 G needle on a micro syringe before surgery.
7. Apply 70% ethyl alcohol to the abdomen and spread the fur upwards and downwards. After spreading the fur, confirm easier visualization of the skin below the left subcostal area for an easier cut. Do not shave off the fur from the abdomen.
   NOTE: The fur will hide the incision site after surgery and prevent the mouse from scratching the incision post operation. However, you can shave the fur to prevent infection of the incision site according to institutional standards.
8. Apply iodine and let it be absorbed into the skin.
9. Place a sterile surgical drape with a 2 cm hole on the mouse.
10. Lift abdominal skin with curved ultrafine forceps and make a 1 cm transverse left subcostal skin incision with curved scissors.
11. Insert the tip of the curved scissors beneath the skin of the incision and slightly open them to separate peritoneum from skin. Retract the scissors from the incision with closed blades.
    NOTE: Opening and closing scissors inside the mouse can cause damage and bleeding.
12. Locate the liver under the peritoneum. Confirm a dark reddish color through the peritoneum.
13. With curved scissors, make a 1 cm transverse incision in the peritoneum. If a peritoneal artery bleeds from the cutting edge, immediately stop the bleeding with cautery.
14. Grab fat tissue using curved ultrafine forceps with one hand, insert the edge of a cotton swab beneath the left liver lobe and roll the swab downward with the other hand to bring out the liver.
    NOTE: Grabbing fat tissue is important to keep the fat tissue from sticking to the cotton swab.
15. Exteriorize the liver on the cotton swab and place the liver on a non-woven absorbent fabric sheet.
    NOTE: The fabric sheet plays two essential roles in stabilizing the liver and absorbing hemorrhage.
16. Make an incision 5 mm in width and depth using a sterile No. 11 scalpel blade to form a pocket in the parenchyma while softly pressing the incision site with the cotton swab.
    1. Insert the blade in parallel with the surface of the liver and cut horizontally.
    2. Press the incision site with the cotton swab to stop any hemorrhage.
       NOTE: Do not keep the blade vertical, otherwise you will break through the liver and injure large vessels in the middle of the liver.
17. Roll the cotton swab upward to open the incision site and implant a 1 mm³ cube of tumor tissue into the pocket with curved ultrafine forceps. Retract the forceps while rolling the cotton swab in reverse rotation and pressing down.
    NOTE: Pressing down on the incision site with the cotton swab while retracting the forceps helps to prevent displacement of the tumor inside the pocket.
18. Gently take the cotton swab off the incision site after implantation. Proceed to step 3.19 as soon as possible.
19. Put an absorbable hemostat on the incision site.
20. Confirm hemostasis. If bleeding continues, add more hemostat on the incision site.
21. Peel the liver off the fabric sheet with forceps (preferably blunt-ended) and put the liver back into the abdominal cavity.
22. Suture peritoneum with double ligature using 5-0 absorbable suture.
23. Suture skin with triple ligature using 5-0 absorbable suture.
    NOTE: Triple ligature helps to prevent surgical incision dehiscence.
24. Observe the mouse until fully awake and put it back in the cage.
25. Measure the weight of the cotton swab and the fabric sheet with blood for bleeding volume during the surgery. Compare them with their original weights before surgery. Reduce bleeding during the surgery to less than 10% of circulating blood volume in mouse.

4. Collecting and Processing of Cultured Human Liver Metastatic Uveal Melanoma Cell Line

1. Prepare cultured cells.
2. Collect cells and calculate the cell number using a cell counter.
3. Prepare an appropriate amount of cell suspension for 10.0 x 10⁶ cells in a 15 mL tube.
4. Spin the tube at 300 x g for 5 min in a centrifuge at room temperature.
5. Remove the supernatant in the 15 mL tube. Leave the cell pellet at the bottom of the tube.
6. Add 50 μL of RPMI 1640 medium into a 1.7 mL tube.
7. Cut the tip of a 200 μL tip with scissors to enlarge the tip opening.
8. Add 60 μL of basement membrane matrix using a pipette with the cut tip into the 1.7 mL tube that has RPMI.
9. Mix RPMI and matrix in the 1.7 mL tube. Vortex it.
10. Add 110 μL of the mixture into the cell pellet in the 15 mL tube. Transfer the cell suspension into a new 1.7 mL tube.
11. Keep the tube on ice before needle injection.

5. Surgical Needle Implantation of Cultured Human Liver Metastatic Uveal Melanoma Cell Line into Liver

1. Follow the above protocol from steps 3.1 to 3.15.
2. Collect the cell suspension with a microsyringe with a 27 G needle.
3. Insert the needle along the surface of the liver and advance the tip of the needle 5 mm deeper.
4. Inject 20 μL of cell suspension into the liver.
5. Cauterize the insertion point of the liver to prevent the injected cells from leaking out. Confirm hemostasis.
6. Follow the above protocol from steps 3.21 to 3.24.

6. CT Scan

1. Place the mouse into a restrainer in the awake state.
2. Wipe the tail with a sterile alcohol pad for disinfection and vasodilation.
3. Inject 100 µL of CT contrast agent through the tail vein with a 27 G needle on a 1 mL syringe.
4. Wait for 4 h after injection before taking the CT scan.
   NOTE: It takes 4 h until the agent is taken up by liver Kupffer cells.
5. Four hours after injection, anesthetize the tumor-bearing mouse with 3–5% vaporized isoflurane by placing it in the induction chamber.
6. Once the mouse is fully anesthetized, place it in the prone position on a CT. Place the isoflurane cone on the mouse's snout to inhale 1.5–3% isoflurane for maintenance of anesthesia.
7. Confirm proper anesthesia by no reaction when the foot of the mouse is pricked with ultrafine forceps.
8. Take a CT scan for 15 min.
9. Ensure that the mouse until it is fully awakened after the CT scan and put it back into the cage.
10. Evaluate for the existence of tumor and measure the tumor size on the CT images.
    NOTE: The contrast agent enhances normal liver parenchyma so that it is easy to recognize unenhanced tumor. Do not misinterpret the gallbladder and stomach as tumor.

7. Harvesting and Processing Tissue

1. Euthanize mice using CO₂ followed by cervical dislocation by placing the index finger and thumb behind the skull and pulling the body by the base of the tail. Proceed to step 7.2 as soon as possible.
2. Place the mouse in a supine position and spray the abdomen with 70% ethyl alcohol.
3. Use sterile forceps and sterile scissors to make a 3-cm transverse incision below the xiphoid process to expose the abdominal organs.
4. Excise the tumor tissue and perform steps 2.1 to 2.2.
5. Cut the rest of the tumor into 2 mm cubes in the Petri dish.
6. Transfer them to a cryogenic tube with cryomedium for re-implantation after cryopreservation.
7. Put the tubes in a cryogenic freezing container filled with isopropanol.
8. Transfer the container to a -80 °C freezer for temporary storage. Do not put the cryotubes with cryomedium directly into a liquid nitrogen tank. Freeze them slowly at a cooling rate of -1 °C/min to preserve tumor tissue.
9. On the next day, transfer the tubes into a liquid nitrogen tank for permanent storage.

8. Re-implantation

1. Keep tubes frozen in a liquid nitrogen jar with liquid nitrogen until ready to implant tissue. Minimize exposure of the tissue to room temperature to maintain viability and enhance chances of engraftment.
2. Thaw cryopreserved tube in a 37 °C water bath.
3. Perform steps 2.2–2.4.
4. Implant the thawed tumor into mice as described in steps 3.1–3.24.

Wyniki

Surgical orthotopic implantation using the liver pocket method can transplant human liver metastatic uveal melanoma tumor into the mouse liver with a high success rate of 80% within six months. The xenograft tumor engrafts in the liver as a solitary tumor without daughter nodules (Figure 1 and Figure 3A). The surgical orthotopic injection technique into the liver using microneedles successfully engrafted cultured human liver-meta...

Dyskusje

The current orthotopic xenograft models are labor-intensive, time-consuming, and expensive to create. Orthotopic tumor xenograft mouse models for liver cancer were established more than two decades ago^19,20,21. However, this technique is complicated and requires use of special equipment, such as a micro-needle holder and 6-0 to 8-0 fine sutures under a microscope. Tumor and normal liver tissue must be sewn up carefully so that t...

Materiały

Name	Company	Catalog Number	Comments
Materials, tissues and animals
Buprenorphine
CO2 tank
Cryomedium
Exitron nano 12000 (Alkaline earth metal-based nanoparticle contrast agent)	Miltenyl Biotec	130-095-700
HBSS 1x, with calcium & magnesium	Corning	21-020-CM
Human liver metastatic uveal melanoma cell line
Human uveal melanoma tissue in the liver			All tissue handling should be done in a Biosafety Level 2 hood. Be careful when working with human tissue; always use gloves and avoid direct skin contact. Assume patients may have been infected with HIV or other highly transmissible organisms. Do not process samples known to carry infections.
Iodine
Isoflurane	Purdue Products	67618-150-17
Isopropanol	Fisher scientific	A416-1	Avoid direct contact to skin and eye and inhalation of anesthetic agent.
Liquid nitrogen
Matrigel HC	BD	354248
NOD.Cg-PrkdcscidIl2rgtm1Wjl/SzJ (NSG) mice	Jackson Lab	5557	4 to 8 weeks old
PBS 1x, without calcium and magnesium	Corning	21-031-CM
RPMI 1640	Corning	10-013-CV
Sterile alcohol prep pad (70% isopropyl alcohol)	Nice-Pak products	B603
4% paraformaldehyde phosphate buffer solution	Wako	163-20145
70% Ethyl alcohol solution	Fisher Scientific	04-355-122
Name	Company	Catalog Number	Comments
Equipments
Absorbable hemostat	Johnson and Johnson	63713-0019-61
Autoclave
Body weight measure
Cautery	Bovie Medical	MC-23009
Cell counter
Centrifuzer
Cotton swab
Cryo freezing container	NALGENE	5100-0001
Cryotube	SARSTEDT	72.379
Curved scissors	World Precision Instruments	503247
Curved ultrafine forceps	World Precision Instruments	501302
Fabric sheet
Freezer
F/AIR Filter Canister	Harvard Apparatus	600979
Heating pad
Isoflurane vaporizer	Artisan Scientific	66317-1
Liquid nitrogen
Liquid nitrogen jar	Thermo Fisher Scientific	2123
Micro-CT scan	Siemens
Needle holder	World Precision Instruments	501246
Petri dishes	Fisher Scientific	FB0875713
Pipette
Spray bottle
Sterile hood			Biosafety level 2 cabinet
Sterile No.11 scalpel	AD Surgical	A300-11-0
Straight forceps	World Precision Instruments	14226
Surgical drape
Tail vein restrainer	Braintree Scientific	TV-150-STD
Water bath
1 mL TB syringe with 27 G needle	BD	309623
1.7 mL tube	Bioexpress	C-3260-1
5-0 PDO Suture	AD Surgical	S-D518R13
15 mL conical tubes	AZER SCIENTIFIC	ES-9152N
27 G needle	BD	780301
27 G needle	Hamilton	7803-01
50 mL conical tubes	AZER SCIENTIFIC	ES-9502N
50 µL micro syringe	BD	80630
50 µL micro syringe	Hamilton	7655-01
100 mL container	Fisher Scientific	12594997
200μL tip