Renal Capsule Xenografting and Subcutaneous Pellet Implantation for the Evaluation of Prostate Carcinogenesis and Benign Prostatic Hyperplasia

Summary

We describe the manufacture of compressed hormone pellets, as well as subcutaneous surgical implantation into mice. This strategy can be combined with the growth of cell and tissue xenografts under the renal capsule of athymic nude mice to evaluate hormonal carcinogenesis and regulation of benign prostate growth.

Abstract

New therapies for two common prostate diseases, prostate cancer (PrCa) and benign prostatic hyperplasia (BPH), depend critically on experiments evaluating their hormonal regulation. Sex steroid hormones (notably androgens and estrogens) are important in PrCa and BPH; we probe their respective roles in inducing prostate growth and carcinogenesis in mice with experiments using compressed hormone pellets. Hormone and/or drug pellets are easily manufactured with a pellet press, and surgically implanted into the subcutaneous tissue of the male mouse host. We also describe a protocol for the evaluation of hormonal carcinogenesis by combining subcutaneous hormone pellet implantation with xenografting of prostate cell recombinants under the renal capsule of immunocompromised mice. Moreover, subcutaneous hormone pellet implantation, in combination with renal capsule xenografting of BPH tissue, is useful to better understand hormonal regulation of benign prostate growth, and to test new therapies targeting sex steroid hormone pathways.

Introduction

Prostate cancer (PrCa) and benign prostatic hyperplasia (BPH) are significant health burdens. PrCa is the second most prevalent solid organ cancer in men and a leading cause of cancer related death ¹. BPH is also highly prevalent among older men, and it is estimated that the clinical manifestation of BPH, lower urinary tract symptoms (LUTS), will affect 50-90% of men ². While androgens and estrogens are known to be important in PrCa and BPH, our understanding of the hormonal mechanisms that underlie carcinogenesis and growth remains incomplete ^3,4. Simple and genetically tractable animal models underpin the studies that will address these priorities in prostate research. In hormone responsive diseases such as PrCa and BPH, the use of subcutaneous, slow release hormone pellets, in isolation or in combination with renal capsule xenografting (the transfer of cells, tissues or organs from one species into another) into the immunocompromised mouse host, provides a simple and reproducible method for studying hormonal regulation of growth and carcinogenesis.

Subcutaneous hormone pellet implantation is a simple and reproducible technique to study hormonal regulation of carcinogenesis and benign growth in the prostate ⁵. Subcutaneous implantation of adult male mice with 25 mg testosterone (T) and 2.5 mg 17β-estradiol (E₂), causes an increase in serum E₂ and gradual decrease in T, recreating the dynamic hormonal environment of aging men ^5,6. In addition, this model recapitulates many of the clinical features of BPH-LUTS ^5,7. Alternative methods of hormone administration include oral/gavage administration or intraperitoneal injection, which cause distress to the rodent, are less consistent in delivering the same amount of drug over time, and are more labor-intensive than a one-time surgical implantation. Other subcutaneous modes for hormone and/or drug delivery include Silastic capsules ⁸. While we also have experience with use of Silastic capsules, compressed pellets are favored for their ease of use and reproducibility. Furthermore, release rates of compressed hormone pellets better mimic the dynamic sex steroid hormone ratios observed in aging men ^5,7.

Since the development of genetically immunocompromised mice, numerous in vivo model systems incorporating xenografting techniques have been developed for the study of a wide variety of normal and diseased tissues. There are several basic categories of xenografting. Tissue xenografts consist of a small piece of intact tissue, which can be a normal structure, malignant tumor, or benign growth. Szot et al. (2007) have recently illuminated renal capsule xenografting of pancreatic islets ⁹. Aamdal et al. (1985) described renal capsule xenografting of 27 human cancer cell lines in immunocompromised mouse hosts ¹⁰. Grafts can also be composed of a single immortalized cell line (cancerous or non-tumorigenic), or can consist of an immortalized cell line combined with cells isolated from mesenchyme (cell recombinant graft).

We favor xenografting of cell recombinants to probe the respective roles of stroma and epithelium in the development of PrCa and BPH. Cunha et al. (1980) was the first to report that when adult murine bladder epithelium is combined with embryonic urogenital sinus mesenchyme (UGM) and xenografted under the renal capsule of male mice hosts, the tissue develops into structures resembling prostate acini ¹¹. Norman et al. (1986) showed that adult mouse prostatic ductal epithelium and UGM, when combined in tissue recombinants, undergo ductal growth and branching morphogenesis ¹². Hayward et al. (1988) showed that human prostate epithelium responds to inductive fetal mesenchyme in a similar fashion ¹³. A hormonal model of carcinogenesis in the prostate, by combining the immortalized human prostate epithelial cell line BPH-1 with UGM, was first reported by Wang et al. ¹⁴ (2001) and has been used extensively in our laboratory ¹⁵. This model is well suited for understanding PrCa progression because benign prostatic epithelium transforms to metastatic disease, modeling advanced PrCa in humans ⁵. Because specific components can be genetically manipulated, cell recombinant grafting is particularly useful as an approach to evaluate stromal-epithelial interactions.

Other sites suitable for xenografting are intradermal, subcutaneous and orthotopic locations ¹⁶. Depending on the tissue and disease process of interest, these may be suitable alternative approaches. For the study of prostate hormonal carcinogenesis and benign growth regulation, we choose the renal capsule site for xenografting due to its higher graft take rate, abundant blood supply, and ability to implant a greater number of xenografts into one confined site ¹⁶. Moreover, hormonal manipulation of the host mouse that results in prostatic atrophy limits the use of prostate orthotopic grafting ¹⁶.

This protocol outlines techniques of compressed hormone/drug pellet manufacture and surgical implantation, as well as renal capsule xenografting of human prostate cell and tissue grafts. Together, these techniques provide a sensitive assay to determine if a genetically modified mouse is more susceptible to PrCa and/or BPH initiation than control strains. Xenografting is a powerful technique for in vivo evaluation of the potential of experimental cells to develop histologic and molecular features of malignancy, as well as an assay for novel treatment strategies for a wide variety of benign and malignant diseases.

Protocol

1. Preparation of Compressed Hormone/Drug Pellet

1. This procedure should be done in a chemical safety hood while wearing a laboratory coat, gloves, mask, safety glasses, and bonnet. To prevent cross-contamination, it is critical that the pellet making equipment is thoroughly cleaned with 70% ethanol prior to and after manufacture of each type of pellet.
2. Using an analytical scale and weigh paper, measure desired amount of hormone/drug powder. Approximately 5% extra material is recommended to accommodate loss during pressing.
3. Some hormones/drugs require a binding agent or filler, which is mixed prior to pellet manufacture. For example, to manufacture a 2.5 mg pellet of 17β-estradiol, combine the hormone with 22.5 mg of cholesterol, which results in a 25 mg pellet.
4. Carefully transfer into die set, place under the press and push lever firmly down to compress pellet. Use consistent pressure to maintain constant surface area to volume ratio.
5. Next reverse the die holder and push the lever down again to release the pellet.
6. Inspect pellet for integrity and determine if final mass is within desired range. An acceptable range is a ± 5% difference in the final mass (for example, an acceptable range for a 25 mg pellet is 23.8-26.3 mg).
7. Pellets can be made prior to surgery and stored (conditions and length of storage depends on stability of the hormone or drug used in pellet manufacture). Hormone pellets can be purchased as an alternative to pellet manufacture. Purchasing pellets avoids the use of non-pharmaceutical grade drugs since they will be formulated and compounded.

2. Preparation of Cell Recombinant Xenografts

1. Harvest and culture primary mouse urogenital mesenchyme (UGM) stromal cells for less than 4 weeks before being collected for counting.
2. Culture immortalized prostate epithelial cell lines in the usual fashion and collect for counting. For this example, prepare cell recombinant grafts by mixing 100,000 BPH-1 epithelial cells per graft with 250,000 UGM stromal cells per graft in suspension.
3. Pellet cells together and re-suspend in enough neutralized type 1 rat tail collagen to make desired number of grafts, each in a volume of 25 μl to 50 μl.
4. Set recombinant grafts at 37 °C for 15 minutes and then cover with growth medium for storage at 37 °C for up to 48 hours before grafting.

3. Preparation of Tissue Xenografts

1. Maintain universal precautions for blood borne pathogens during tissue procurement and surgical procedure. Following institutional review board approval and informed consent, obtain fresh prostate tissue from surgical resection. For studies of BPH growth, tissues can be harvested from transurethral resection of the prostate or simple prostatectomy procedures. Primary xenografts of PrCa or benign prostate tissue can be obtained from radical prostatectomy specimens.
2. Store harvested tissue in isotonic buffered cell media or normal saline on ice. Depending on the tissue and storage conditions, specimens may be stored for up 24 hours prior to preparation of tissue grafts. Common practice is to divide the specimen into three pieces, with one section fixed in formalin and routinely processed for histology, one piece snap frozen for molecular analysis and one piece further divided into tissue xenografts (1-3 mm in size).

4. Preparation of Fire Polished Glass Pipette

1. Wearing eye protection, laboratory coat and heat resistant gloves, place glass Pasteur pipette tip into Bunsen burner flame at approximately a 60° angle to start.
2. Keep a constant, slight movement to the pipette so as to avoid hot spots.
3. Draw the tip thinly and fire-polish with the goal being a slightly curved end with a rounded, closed tip.

5. Preparation of Instruments and Surgical Suite

1. Autoclave all instruments. Prior to surgery, inspect all instruments, taking care to maintain sterile field.
2. Set up sterile surgical area with absorbent pads, dissection scope, anesthesia nose cone, surgical instruments and heating lamp directed toward the working area. To sterilize tools in between mice, prewarm bead sterilizer.
3. Assemble and test anesthesia apparatus; weigh gas scavenger.
4. Using the microwave, heat wax pads for up to 5 min (in 1 min increments, kneading the wax pad in order to combine melted and unmelted wax) at 70-80% power, until all the wax is evenly melted. By touch, ensure the pad is not too hot and place under the empty rodent recovery cage.
5. Don personal protective equipment for rodent surgical procedures, including surgical mask, bonnet, gloves and gown.

6. Surgical Procedure: Renal Capsule Xenografting

1. All procedures involving research animals should be performed with approval of institutional animal care and use committees. Prior to induction of anesthesia, observe the mouse to ensure health and well-being.
2. Place the mouse into chamber for induction of isoflurane anesthesia at 3-5%. When the animal has stopped moving and respiratory rate has decreased, transfer to nose cone in the prone position, and maintain anesthesia at 1-3%.
3. If necessary, use clippers to shave the back of the mouse. This is not needed when utilizing athymic nude (nu/nu) mice.
4. Apply firm pressure to the webbing of the extended hind foot to evaluate adequacy of anesthesia. If the mouse responds to pressure, more time is needed for the anesthesia to take effect. If necessary, adjust anesthesia flow slightly to achieve adequate anesthesia.
5. When the mouse is adequately anesthetized, disinfect the surgical site with surgical iodine (Betadine) solution followed by 70% alcohol.
6. Don sterile gloves and gown; apply sterile drapes to the surgical site. Lift the back skin with a pair of toothed forceps, and using the coarse scissors make a 2-3 cm dorsal midline incision.
7. Using blunt scissors or a probe, separate the underlying dermis from the body wall (on both sides of the incision for bilateral grafting or on one side for unilateral grafting).
8. Reposition the mouse into lateral position, and identify the location of the kidney by viewing the renal profile through the muscle wall. Applying gentle manual pressure with the thumb and index finger on the abdomen may assist with visualizing internal organs.
9. Using fine iris scissors, and taking care to avoid major vessels and spinal nerves, make a 1 cm incision in the body wall parallel to the spine. Widen this incision to 1.5-2.0 cm (slightly longer than the long axis of the kidney) by gently opening the scissors wider after placing them in the initial incision.
10. Exteriorize the kidney by applying gentle pressure outside the muscle wall on either side of the kidney using the index finger and thumb. Tuck the skin edges below the exteriorized kidney, which will rest on the body wall. While the kidney is exteriorized, maintain hydration of the renal capsule by applying sterile saline.
11. Using fine #5 forceps gently lift the kidney capsule from the parenchyma of the kidney and with a fine scalpel, make a 2 -4 mm incision in the capsule. The size of the incision is determined by the size of the graft, but should be minimized in order to maintain integrity of the capsule.
12. Manipulate a glass Pasteur pipette that has been drawn thin and fire-polished with a rounded closed end under the capsule tangential to the surface of the kidney. Gently open a small capsule pocket for the grafts, using great care not to damage the kidney parenchyma.
13. The cut edge of the kidney capsule is lifted with the fine forceps, and the graft is inserted into the pocket under the capsule using the fire-polished glass pipette. Several grafts can be placed under the kidney capsule and evenly spaced on the kidney surface.
14. If, during the course of grafting, the capsule becomes dehydrated, it should be moistened by applying sterile saline.
15. When grafting is complete, gently lift the sides of the muscle wall incision to replace the kidney back into the body cavity. Observe that the grafts do not slip out from under the capsule.
16. Close the muscle wall with a single suture (4-0, FS-2 vicryl suture). The surgical procedure can be repeated on the contralateral kidney by repositioning the mouse.
17. When grafting is complete, the mouse may undergo subcutaneous pellet implantation during the same surgical procedure (See Protocol step 7).
18. Using toothed forceps, align the skin incision edges and apply 2-3 surgical wound clips to close the incision. Administer analgesia (we use 5-10 mg/kg carprofen using a subcutaneous injection) and place the mouse in the lateral recumbent position in the recovery cage. Ensure maintenance of body temperature with heat from a lamp or heating pad. Observe for full recovery of the mouse, which should occur in less than 15 minutes.
19. Mice should be observed for the next 24 hr for signs of post-operative pain, bleeding and/or other complications. Regularly inspect surgical incision for signs of infection.
20. Remove wound clips with surgical wound clip removal device 7-14 days after surgery.

7. Surgical Procedure: Subcutaneous Pellet Implantation

1. Repeat steps 6.1-6.5 from renal capsule xenografting surgical procedure.
2. Lift the back skin with a pair of skin forceps, and using the coarse scissors make a 1-2 cm dorsal midline incision.
3. Using blunt scissors or a probe, separate the underlying dermis from the body wall in a cranial direction.
4. Using skin forceps, gently lift the cranial aspect of the skin incision, and holding the hormone pellet gently with straight, serrated forceps, insert the pellet and release at the scruff of the neck, in the pocket you created with the blunt probe.
5. Repeat steps 6.18-6.20 from renal capsule xenografting surgical procedure. If performing subcutaneous pellet implantation only, use 1 surgical wound clip to close the surgical incision.

Results

Depending on the hypothesis to be tested, renal capsule xenografting of tissue or cell recombinants can be performed in isolation, or in combination with subcutaneous hormone pellet implantation. A schematic of a hypothetical experiment combining renal capsule xenografting experiment with subcutaneous hormone pellet implantation is illustrated in Figure 1. A variety of powdered and/or crystalline substances can be used in the manufacture of compressed pellets with the pellet press. Figure 2

Discussion

This paper and the protocol outlined herein describe the manufacture of compressed hormone and/or drug pellets, and the surgical procedure for subcutaneous pellet implantation of mice. Depending on the research question to be addressed, this technique can be performed singly, or in combination with renal capsule xenografting of prostate cell recombinant grafts or prostate tissue xenografts, which are techniques widely used in prostate research ^17-19. Taken together, these techniques offer a powerful approach t...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Pellet press	PARR Industries	2815	3mm Punch & Die set
Analytical Balance	Ohaus	1918302	Voyager or other
Bunsen burner	Fisher Scientific	03-962Q
Dissecting microscope	Leica	L2
Deltaphase Pad Kit (reheatable wax)	Braintree Scientific	39DP	Alternative to an electric heating pad
Hot Bead sterilizer	F.S.T.	18000-45
Isoflurane Vaporizer	Supera Anesthesia Innov	VAP3000
Induction Chamber	Supera Anesthesia Innov	RES644
F/AIR Canister	Supera Anesthesia Innov	80120
4 port Manifold	Supera Anesthesia Innov	RES536
Rebreathing Circuits	Supera Anesthesia Innov	CIR529
Inlet/Outlet Fittings	Supera Anesthesia Innov	VAP203/4
Pressure Reg/Gauge	Supera Anesthesia Innov	OXY508
Oxygen Flowmeter	Supera Anesthesia Innov	OXY660
21mm Clear Tubing	Supera Anesthesia Innov	301-150
Small Mice Nose Cone	Supera Anesthesia Innov	ACC526
Sterile surgical gown	Midwest Vet Supply	350.79866.2
Surgical mask	Midwest Vet Supply	350.50111.2	2-ply w/ earloops
Sterile gauze	Midwest Vet Supply	001.14100.2	2 x 2 inches
Sterile Gloves	Kimberly Clark	55092
Bouffant	Midwest Vet Supply	001.27100.2
Cotton Tip Applicator	Midwest Vet Supply	001.06220.2
Surgical Scrub Wash	Midwest Vet Supply	733.80010.3
Sterile Fields (Fenestrated)	General Econopak, Inc	88VCSTF
REAGENTS
Cholesterol	Sigma-Aldrich	C-3292
Testosterone Proprionate	Sigma-Aldrich	T-1875
17β-estradiol	Sigma-Aldrich	E-2758
Isoflurane	Midwest Vet Supply	193.33161.3
Carprofen	Midwest Vet Supply	193.70200.3	Injectable (Rimadyl)
Betadine	Webster Veterinary	07-836-3379
Sterile saline	Midwest Vet Supply	193.74504.3	NaCl 0.9%, Injectable
SURGICAL INSTRUMENTS
Straight Sharp/Blunt Scissors	Fine Scientific Tools (F.S.T.)	14054-13
Graefe forceps (Serrated, Toothed, Curved)	F.S.T.	11055-10
Fine Iris scissors	F.S.T.	14090-09
Graefe Scalpel	F.S.T.	10071-12
Dumont #5 forceps	F.S.T.	11251-20
Glass Pasteur pipets	Fisher Scientific	S67050	5" length
Graefe forceps (Serrated, Curved)	F.S.T.	11052-10
Graefe forceps (Serrated, Straight)	F.S.T.	11050-10
Vicryl Suture	Midwest Vet Supply	295-92100.2	4-0, FS-2, Absorbable
Needle Holder w/ scissor action	F.S.T.	12002012
Wound Clips	Braintree Scientific	ACS CS	May use 7mm or 9mm clips
Reflex Wound Clipper	F.S.T.	12031-09	Correspond w/ wound clip size
Would Clip Remover	F.S.T.	12033-00	Universal
Sterilization Container	F.S.T.	20810-01	Any autoclavable container will do
Syringe w/ needle	BD	148292C	1cc, 25Gx1
Ear Tag Applicator	National Band Tag Co.	1005s1	Alternative to ear notching
Small Animal Ear Tags	National Band Tag Co.	1005-1