Patient-derived xenograft model for uterine leiomyoma by sub-renal capsule grafting
- Ketamine hydrochloride (100 mg/ml) (Hospira, Lake Forest, IL, NDC 0409-2051-05)
- Xylazine (20 mg/ml) (Akorn Animal Health, Lake Forest, IL, NADA 139-236)
- Buprenorphine (0.3 mg/ml) (Reckitt Benckiser, Hull, England, NDC 12496-0757-1)
- Meloxicam (Eloxiject) (5 mg/ml) (Henry Schein, Dublin, OH, NDC 11695-6925-2)
- Decon CiDehol 70% Isopropyl alcohol, 16 oz spray bottles (ThermoFisher Scientific [TFS], Waltham, MA, 04-355-63)
- Surgical Scrub and Handwash 2% Chloroxylenol (Vetoquinol, Ft. Worth, TX, 1703000826) or 10% Povidone iodine prep pads (TFS 06-669-70)
- 17β-Estradiol (Millipore-Sigma [Sigma], St. Louis, MO, E2758)
- Progesterone (Sigma P0130)
- Cholesterol (Sigma C3045)
- Diethyl Ether (Sigma 309958)
- FBS (TFS, 16000044)
- DMEM/F-12 (TFS 11320082)
- Hanks’s Balanced Salt solution (TFS 14175095)
- Collagenase Type I (Sigma C0130)
- DNase I (Sigma D5025)
- Gibco Antibiotic-Antimycotic (100×) (TFS 15240096)
- 0.25% Trypsin-EDTA (TFS 25200114)
- Collagen I High Concentration, Rat Tail (Corning, New York, NY, 354249)
- Cell culture grade water
- 1N NaOH
- 10× PBS
- Culture medium: DMEM/F-12 containing 10% FBS and Gibco Antibiotic-Antimycotic
- Tissue digestion medium: Hanks’s Balanced Salt solution containing 1.5 mg/ml collagenase Type I supplemented and Gibco Antibiotic-Antimycotic
- Collagen stock solution: Dilute collagen I to 3–5 mg/ml with sterile water. Snap-freeze aliquots (1 ml/tube) in liquid nitrogen, and store at –80°C. Thaw overnight at 4°C prior to use.
- Collagen setting solution: Mix 100 µl 10× PBS, 23 µl 1N NaOH and 877 µl cell culture grade water. Sterilize with a 0.2 µm syringe filter, and store at 4°C in an airtight small-void-volume container.
- Ketamine/Xylazine Anesthetic (dosage 90/8 mg/kg ketamine/xylazine): Mix 0.9 ml Ketamine (100 mg/ml) and 0.14 ml xylazine (20 mg/ml), and dilute it to 10 ml with saline. Inject 100 µl per 10 grams body weight.
- Buprenorphine (dosage 0.1 mg/kg): Dilute 300 µl Buprenorphine (0.3 mg/ml) to 9 ml with saline. Inject 100 µl per 10 grams body weight.
- Meloxicam (dosage 2.0 mg/kg): Dilute 800 µl Meloxicam (5 mg/ml) to 10 ml with saline. Inject 100 µl per 20 grams body weight.
- Hormone pellets: Because murine hormone levels are far below that of humans, the host must be supplemented with both E2 and P4 to achieve the robust growth of xenografts. Dissolve cholesterol, progesterone (P4) and/or estradiol (E2) in ethyl ether in the following ratio by weight, 1% estradiol and/or 5% progesterone with cholesterol to 100 %. Evaporate solution overnight with stirring under the fume hood. Compress powder into 3 mm diameter pellets with the Parr 2811 Pellet Press. Cut pellets to ~30 mg (~2 mm length). Subcutaneous implantation of two 30 mg E2 + P4 pellets sustains serum E2 and P4 levels of 374.5 pg/ml (N = 20, 95% CI 291.5 to 457.5) and 50 ng/ml (N = 20, 95% CI 37 to 63), respectively, for 2 months. These are comparable to the serum hormone levels of cycling women . Pellets should be replaced every 2 months.
- Disposable Scalpel #22 (World Precision Instruments, Sarasota, FL, 500354)
- Surgical instruments (Sterilize prior to procedure): Moria MC32/B Iris forceps (Fine Science Tools, Foster City, CA (FST), 11373-22), needle holder with suture cutter (FST 12002-14), scissor (FST 14054-13), hardened fine scissor (FST14090-09), student Vanna spring scissor (FST 91500-09), forceps (FST 11064-07), extra-fine Graefe forceps (FST 11150-10)
- Glass rod with balled tip (< 1 mm3 ball) made from 9 inch Pasteur pipets. Flame pipets ~20 mm below the tip, pull the tip and further burn the closed end to make a ball .
- Supplementary surgical tools: small animal clipper, cautery kit (FST 18010-00), wound clip applier, remover, and clips (Fisher 01-804, 01-804-15, 01-804-5)
- Coated Vicryl Suture 4-0 27'' (Ethicon, Cincinnati, OH, J310H)
- Exel International Tuberculin Syringes (FST 14-840-50)
- Dissecting microscope with LED illumination (e.g., Leica, DMS1000, Buffalo Grove, IL)
- MultiSample BioPulverizer or single BioPulverizers (BioSpec Products Inc, Bartlesville, OK, 59012MS or 59012N)
- The Parr 2811 Pellet Press (Parr Instrument Company, Moline, IL)
- Biological Safety Cabinet
- Magnetic stirrer and stirrer bar
- 500 ml beaker
- 125 ml or 250 ml baffled flask
Isolation of UL cells
1.Place UL tissues in cold DMEM/F-12 medium containing antibiotic-antimycotic soon after extraction (Fig. 1A).
2.Using a #22 surgical blade, cut the UL tissues into pieces of < ~9 mm3 in a 10 mm plastic petri dish, excluding any calcified or necrotic portions as well as myometrium. Keep tissue covered with tissue digestion medium (Fig. 1B).
3.Suspend minced tissues into tissue digestion medium of a volume greater than five times tissue volume and transfer to 125 ml baffled flasks (< 50 ml) or 250 ml baffled flasks (< 80 ml) (Fig. 1C).
4.Add 10 µl DNase I stock per 10 ml tissue digestion medium.
5.Digest tissues at 37°C on a shaker 200–250 rpm until the pieces of tissue disappear. When digestion takes longer than 3 h, stop the shaker, wait for large tissue pieces settle on the bottom, and then transfer the digestion medium containing cells into 50 ml polypropylene tubes on ice. Add fresh digestion medium to the flask. Restart the shaker to digest tissues until large pieces disappear. Digestion time < 6 h is recommended for the best result.
6.Filter the cell suspension through a 100 µm Falcon cell strainer (BD Falcon) into new 50 ml polypropylene centrifuge tubes.
7.Centrifuge at 220× g for 5 min at 4°C.
8.Remove supernatant. Gently resuspend cells in 45 ml culture medium.
9.Centrifuge at 220× g for 5 min at 4°C again.
10.Remove supernatant. Gently resuspend cells in 10–20 ml of culture medium.
12.Plate cells on 10 cm culture dish at ~2–4 × 106 cells per plate in the culture medium, and incubate for 1–3 d (Fig. 1D).
Cell pellet preparation
13.Collect cells from culture dish.
13.1.Aspirate culture medium. Gently wash cells with ~8 ml of pre-warmed PBS (Mg++ and Ca++ free) twice.
13.2.Rinse the surface of cells with 1 ml of 0.25% EDTA-Trypsin, and aspirate.
13.3.Add 2 ml of 0.25% EDTA-Trypsin, and incubate at 37°C until cells detach. If a majority of cells are still attached after 15 min, transfer the trypsin solution into a 15 ml centrifuge tube with 5 ml culture medium. Add 2 ml of fresh 0.25% EDTA-Trypsin solution to the culture dish. Incubate at 37°C for an additional 15 min.
13.4.Dispense 5 ml of culture medium to the culture dish. Collect detached cells by pipetting several times.
13.5.Transfer the cell suspension to a 15 ml centrifuge tube. Centrifuge at 220× g for 5 min.
13.6.Remove the supernatant. Gently resuspend the cell pellet into ~5 ml culture medium.
13.7.Count the cells in this suspension.
14.Collect a volume of cell-suspension that contains the number of cells required to make the desired number of grafts. Transfer it into a centrifuge tube.
15.Centrifuge at 220× g for 5 min. Remove the supernatant. Calculate the volume of the collagen gel solution required to prepare desired number of cell pellets.
16.Prepare collagen gel solution: mix one volume of collagen stock solution with one volume of setting solution, all on ice.
17.Resuspend cells collected in step15 in collagen gel solution using a pipette with a wide-orifice tip.
18.With a wide-orifice tip, dispense droplets of desired pellet volume onto the surface of a cell culture plate (6 well-plate is recommended).
19.Incubate the plate containing the droplets at 37°C in a CO2 incubator for 15–30 min. After confirming the droplets have solidified, add pre-warmed (37°C) DMEM/F-12 to a depth of 0.5–1.0 cm (~6 ml per well for 6 well plate), and gently detach the cell pellet from the bottom with the tip of a P20 micropipette (Fig. 1E).
20.Transfer cell pellets to surgery suite for grafting.
Ovariectomy and xenografting
21.Prepare clean post-surgical cage on a heating pad set to low.
22.Intraperitoneally inject ketamine/xylazine anesthetic. When the mouse is unresponsive to foot pinching and whiskers unmoving, continue.
23.Apply analgesics at this time as required by your institution.
24.Apply eye lubricant, and shave dorsal caudal half of mouse.
25.Treat the entire dorsal caudal half of the mouse with a disinfectant such as chloroxylenol, followed by 70% alcohol. Repeat thrice.
26.Transfer mouse to a heating pad set to low.
27.Make a 1.0 cm incision through the skin of the dorsal midline parallel to the spine. Separate the skin and muscle wall laterally by probing with scissors (Fig. 2A).
28.Identify the ovarian fat pad through the muscle (Fig. 2B). Make a 2 mm incision perpendicular to the spine just rostral to the ovarian fat, avoiding blood vessels.
29.Reach through both the skin and muscle incisions with forceps, grasp the ovarian fat, and pull the ovary through the incision to the exterior of the body. Hold the ovary away from the kidney, separating connective tissue if necessary.
30.Remove the ovary by cauterizing arteries and connective tissues. Or clamp arteries and uterus just below the oviduct with a hemostat (Fig. 2C), then remove the ovary and oviduct with a fine scissors on the distal side of the hemostat. To avoid bleeding, wait for > 1 min before removing hemostat.
31.Return the uterus into the peritoneal cavity.
32.With gentle pressure from thumbs and forefingers positioned on the muscle wall, gently push the kidney out through the muscle wall incision (Fig. 2D).
33.Using the soft rounded tips of the Moria iris forceps, gently grip the kidney capsule and lift it away (1 mm) from the kidney parenchyma. Ideally the capsule is gripped at the outer edge of the kidney to optimize space for grafts. Pierce the capsule with one blade of the opened spring scissor (Fig. 2E), and move the blade into the capsule for the length of the blade. Make a single straight incision (~2 mm) along the longitudinal axis of the kidney.
34.Dip the tip of the glass rod into DMEM/F12 in the dish of cell pellet. Holding one edge of the capsule at the incision with the Moria iris forceps, gently insert the wet tip of the glass rod through the incision between the capsule and kidney parenchyma to form a pocket (Fig. 2F). The pocket with a diameter of the pellet should extend far from the capsule incision but not to the hilus.
35.While holding open the incision with the Moria iris forceps in one hand, gently pick up a single cell pellet from medium with forceps and insert it into the pocket.
36.Use the glass rod to gently push the pellet deep into the pocket (Fig. 2G), and then release the capsule. To force the pellet deeper, lightly compress the capsule from the outside with the glass rod.
37.Holding the muscle incision open with forceps, gently guide the kidney back into the peritoneal cavity.
38.Close the muscle wall incision with absorbable sutures in the simple interrupted pattern (Fig. 2H).
39.Repeat steps 28–38 on the contralateral side.
40.Grasp the skin just rostral to the skin incision site with forceps. Probe apart the skin and muscle wall to form a tunnel from the incision site to the nape of the neck, using another forceps.
41.With forceps holding the hormone pellet, push the pellet through the tunnel, and deposit the pellet at the nape of the neck (Fig. 2I).
42.Release the skin and close the incision with wound clips or suture.
43.Return the mouse to a clean cage on a heating pad. Once the mouse is ambulatory, monitoring can be discontinued and the cage returned to the rack. The mouse should be given analgesic in subsequent days and wound clips removed according to the approved IACUC protocol.
Assessing tumor growth in live mice
44.Follow steps 21–28, looking for the kidney rather than an ovarian fat pad. Visually check the growth of PDXs on the kidney through this incision, or expose the kidney following step 32 (Fig. 3A).
45.Follow steps 37 and 38 if the kidney was exposed.
46.Repeat on the contralateral side
47.Then follow 42–43.
48.Euthanize mouse. To obtain blood, anesthetize mouse with ketamine and xylazine. When full anesthetic depth is reached, perform cardiac puncture; or more preferably, obtain blood through enucleation, as there is less cell lysis.
49.Spray mouse with 70% ethanol to minimize hair contamination. Cut through the skin and muscle walls, and expose the kidneys.
50.Grasp the renal vessels at the hilus, and cut the kidneys away from the body.
51.Transfer the kidney bearing PDXs to a small petri dish with PBS, still grasping by the vessels so as not to damage the kidneys.
52.Image the grafts on the kidneys with a ruler in the viewer along the x-and y-axes (Fig. 3B). So long as the axes are perpendicular, one axis is set on the greatest diameter available.
53.For an accurate assessment of height, cut through the center of the graft and kidney parenchyma to obtain a cross section which displays the entire height of the graft (Fig. 3C).
54.Trim excess kidney tissues using a surgical knife, and process the PDX and surrounding kidney tissues for desirable histological analyses. For the extraction of nucleic acid and protein, remove kidney tissues completely, and freeze PDXs in liquid nitrogen.
55.Collect host female reproductive tract, and fix for future reference.
Tumor volume measurement
56.Calculate tumor volume (Fig. 3B and 3C). Although the PDX grows as an ellipsoid as revealed upon complete extraction, the tumor volume has been assessed as a hemi-ellipsoid, the portion of tumor that extends above the surface of the kidney. Tumor volume = ⅔ πabc: a and b = radius on x and y axes, c = height = (h1 + h2)/4.
Protein/RNA extraction method
57.Place a clean BioPulverizer in a shallow container. Cool thoroughly with liquid nitrogen. Then place it on bench top.
58.Place pre-frozen tissue in the well of the mortar, and insert pestle.
59.Pound pestle with hammer multiple times to pulverize the tissue.
60.Scrape the powder into a microcentrifuge tube with a sterile spatula.
61.Clean the pulverizer between samples, and repeat as needed.
62.Proceed with a standard protein and RNA extraction protocol.
|5||Substantial tissue is undigested||
|20||Pellet is too soft||
|32||Kidney slips back in||
|34||Capsule tears despite gentle handling||
|49||Tumor didn’t grow in hosts with E2 + P4||
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