The authors would like to thank all members of the Ricke lab, past and present. We thank Calvin Patten, Jr., DVM and Brigitte Raabe, DVM, for helpful comments regarding the protocol. We would like to acknowledge the NIDDK, NCI, and NIEHS for their financial support for these studies: R01DK093690, R01CA123199, RC2ESO18764. TMN is a trainee in the Medical Scientist Training Program at the University of Rochester funded by NIH T32 GM07356; the NIH under Ruth L. Kirschstein National Research Service Award F30DK093173 also supports this project. CDV is supported by T32CA157322 and ABT is supported by T32ES007015 at the University of Wisconsin-Madison. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of General Medical Sciences or NIH.

1. Preparation of Compressed Hormone/Drug Pellet
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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 17&beta;-estradiol, combine the hormone with 22.5 mg of cholesterol, which results in a 25 mg pellet.</li>
	<li>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.</li>
	<li>Next reverse the die holder and push the lever down again to release the pellet.</li>
	<li>Inspect pellet for integrity and determine if final mass is within desired range. An acceptable range is a &plusmn; 5% difference in the final mass (for example, an acceptable range for a 25 mg pellet is 23.8-26.3 mg).</li>
	<li>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.</li>
</ol>
2. Preparation of Cell Recombinant Xenografts
<ol>
	<li>Harvest and culture primary mouse urogenital mesenchyme (UGM) stromal cells for less than 4 weeks before being collected for counting.</li>
	<li>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.</li>
	<li>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 &mu;l to 50 &mu;l.</li>
	<li>Set recombinant grafts at 37 &deg;C for 15 minutes and then cover with growth medium for storage at 37 &deg;C for up to 48 hours before grafting.</li>
</ol>
3. Preparation of Tissue Xenografts
<ol>
	<li>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.</li>
	<li>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).</li>
</ol>
4. Preparation of Fire Polished Glass Pipette
<ol>
	<li>Wearing eye protection, laboratory coat and heat resistant gloves, place glass Pasteur pipette tip into Bunsen burner flame at approximately a 60&deg; angle to start.</li>
	<li>Keep a constant, slight movement to the pipette so as to avoid hot spots.</li>
	<li>Draw the tip thinly and fire-polish with the goal being a slightly curved end with a rounded, closed tip.</li>
</ol>
5. Preparation of Instruments and Surgical Suite
<ol>
	<li>Autoclave all instruments. Prior to surgery, inspect all instruments, taking care to maintain sterile field.</li>
	<li>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.</li>
	<li>Assemble and test anesthesia apparatus; weigh gas scavenger.</li>
	<li>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.</li>
	<li>Don personal protective equipment for rodent surgical procedures, including surgical mask, bonnet, gloves and gown.</li>
</ol>
6. Surgical Procedure: Renal Capsule Xenografting
<ol>
	<li>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.</li>
	<li>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%.</li>
	<li>If necessary, use clippers to shave the back of the mouse. This is not needed when utilizing athymic nude (nu/nu) mice.</li>
	<li>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.</li>
	<li>When the mouse is adequately anesthetized, disinfect the surgical site with surgical iodine (Betadine) solution followed by 70% alcohol.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>If, during the course of grafting, the capsule becomes dehydrated, it should be moistened by applying sterile saline.</li>
	<li>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.</li>
	<li>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.</li>
	<li>When grafting is complete, the mouse may undergo subcutaneous pellet implantation during the same surgical procedure (See Protocol step 7).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Remove wound clips with surgical wound clip removal device 7-14 days after surgery.</li>
</ol>
7. Surgical Procedure: Subcutaneous Pellet Implantation
<ol>
	<li>Repeat steps 6.1-6.5 from renal capsule xenografting surgical procedure.</li>
	<li>Lift the back skin with a pair of skin forceps, and using the coarse scissors make a 1-2 cm dorsal midline incision.</li>
	<li>Using blunt scissors or a probe, separate the underlying dermis from the body wall in a cranial direction.</li>
	<li>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.</li>
	<li>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.</li>
</ol>

<ol>
	<li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Cancer Facts Figures. 4, (2010).</li><li>McVary, K. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BPH:+epidemiology+and+comorbidities.">BPH: epidemiology and comorbidities.</a> Am. J. Manag. Care. 12, S122-S128 (2006).</li><li>Nicholson, T. M., Ricke, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Androgens+and+estrogens+in+benign+prostatic+hyperplasia:+past,+present+and+future.">Androgens and estrogens in benign prostatic hyperplasia: past, present and future.</a> Differentiation. 82, 184-199 (2011).</li><li>Ricke, W. A., Wang, Y., Cunha, G. 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W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interactions+between+adult+human+prostatic+epithelium+and+rat+urogenital+sinus+mesenchyme+in+a+tissue+recombination+model.">Interactions between adult human prostatic epithelium and rat urogenital sinus mesenchyme in a tissue recombination model.</a> Differentiation. 63, 131-140 (1998).</li><li>Wang, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+human+prostatic+epithelial+model+of+hormonal+carcinogenesis.">A human prostatic epithelial model of hormonal carcinogenesis.</a> Cancer Res. 61, 6064-6072 (2001).</li><li>Ricke, E. 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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+regeneration+of+murine+prostate+from+dissociated+cell+populations+of+postnatal+epithelia+and+urogenital+sinus+mesenchyme.">In vivo regeneration of murine prostate from dissociated cell populations of postnatal epithelia and urogenital sinus mesenchyme.</a> Proc. Natl. Acad. Sci. U.S.A. 100, 11896-11903 (2003).</li><li>Priolo, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+and+genomic+characterization+of+mouse+xenografts+of+human+primary+prostate+tumors.">Establishment and genomic characterization of mouse xenografts of human primary prostate tumors.</a> Am. J. Pathol. 176, 1901-1913 (2010).</li><li>Pritchett-Corning, K. R., Mulder, G. B., Luo, Y., White, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+of+Rodent+Surgery+for+the+New+Surgeon.">Principles of Rodent Surgery for the New Surgeon.</a> J. Vis. Exp. (47), e2586 (2011).</li></ol>

The authors declare that they have no competing financial interests.

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...

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 17&beta;-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.

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

renal capsule xenografting and subcutaneous pellet implantation for the evaluation of prostate carcinogenesis and benign prostatic hyperplasia

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.

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</stro...

Watch this Scientific Journal Video about Renal Capsule Xenografting and Subcutaneous Pellet Implantation for the Evaluation of Prostate Carcinogenesis and Benign Prostatic Hyperplasia at JoVE.com

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

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.

New therapies for two common prostate diseases,  prostate cancer (PrCa) and benign prostatic hyperplasia (BPH), depend  critically on experiments ...

renal-capsule-xenografting-subcutaneous-pellet-implantation-for

Research

JoVE Journal

Medicine

24.9K Views.  University of Wisconsin-Madison. 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.

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

Evaluation of Nanoparticle Uptake in Tumors in Real Time Using Intravital Imaging

Current technologies for tumor imaging, such as ultrasound, MRI, PET and CT, are unable to yield high-resolution images for the assessment of nanoparticle uptake in tumors at the microscopic level1,2,3, highlighting the utility of a suitable xenograft model in which to perform detailed uptake analyses. Here, we use high-resolution intravital imaging to evaluate nanoparticle uptake in human tumor xenografts in a modified, shell-less chicken embryo model. The chicken embryo model is particularly well-suited for these in vivo analyses because it supports the growth of human tumors, is relatively inexpensive and does not require anesthetization or surgery 4,5. Tumor cells form fully vascularized xenografts within 7 days when implanted into the chorioallantoic membrane (CAM) 6. The resulting tumors are visualized by non-invasive real-time, high-resolution imaging that can be maintained for up to 72 hours with little impact on either the host or tumor systems. Nanoparticles with a wide range of sizes and formulations administered distal to the tumor can be visualized and quantified as they flow through the bloodstream, extravasate from leaky tumor vasculature, and accumulate at the tumor site. We describe here the analysis of nanoparticles derived from Cowpea mosaic virus (CPMV) decorated with near-infrared fluorescent dyes and/or polyethylene glycol polymers (PEG) 7, 8, 9,10,11. Upon intravenous administration, these viral nanoparticles are rapidly internalized by endothelial cells, resulting in global labeling of the vasculature both outside and within the tumor7,12. PEGylation of the viral nanoparticles increases their plasma half-life, extends their time in the circulation, and ultimately enhances their accumulation in tumors via the enhanced permeability and retention (EPR) effect 7, 10,11. The rate and extent of accumulation of nanoparticles in a tumor is measured over time using image analysis software. This technique provides a method to both visualize and quantify nanoparticle dynamics in human tumors.

We present a novel approach to quantify nanoparticle localization in the vasculature of human xenografted tumors using dynamic, real-time intravital imaging in an avian embryo model.

Current technologies for tumor imaging, such as ultrasound, MRI, PET and CT, are unable to yield high-resolution images for the assessment of nanoparticle ...

Cell Population Analyses During Skin Carcinogenesis

Cancer development is a multiple-step process involving many cell types including cancer precursor cells, immune cells, fibroblasts and endothelial cells. Each type of cells undergoes signaling and functional changes during carcinogenesis. The current challenge for many cancer researchers is to dissect these changes in each cell type during the multiple-step process in vivo. In the last few years, the authors have developed a set of procedures to isolate different cell populations during skin cancer development using K14creER/R26-SmoM2YFP mice. The procedure is divided into 6 parts: 1) generating appropriate mice for the study (K14creER+ and R26-SmoM2YFP+ mice in this protocol); 2) inducing SmoM2YFP expression in mouse skin; 3) preparing mouse skin biopsies; 4) isolating epidermis from skin; 5) preparing single cells from epidermis; 6) labeling single cell populations for flow cytometry analysis. Generation of sufficient number of mice with the right genotype is the limiting step in this protocol, which may take up to two months. The rest of steps take a few hours to a few days. Within this protocol, we also include a section for troubleshooting. Although we focus on skin cancer, this protocol may be modified to apply for other animal models of human diseases.

Carcinogenesis is a process involving interactions between cancer cells and the cancer microenvironment. To dissect the molecular events, one needs to isolate different cell populations at different stages during cancer development. Using a mouse model for basal cell carcinoma, we describe a protocol for cell analyses during carcinogenesis.

Cancer development is  a multiple-step process involving many cell types including cancer precursor  cells, immune cells, fibroblasts and endothelial ...

Evaluation of Tumor-infiltrating Leukocyte Subsets in a Subcutaneous Tumor Model

Specialized immune cells that infiltrate the tumor microenvironment regulate the growth and survival of neoplasia.&#160; Malignant cells must elude or subvert anti-tumor immune responses in order to survive and flourish. Tumors take advantage of a number of different mechanisms of immune &#8220;escape,&#8221; including the recruitment of tolerogenic DC, immunosuppressive regulatory T cells (Tregs), and myeloid-derived suppressor cells (MDSC) that inhibit cytotoxic anti-tumor responses. Conversely, anti-tumor effector immune cells can slow the growth and expansion of malignancies: immunostimulatory dendritic cells, natural killer cells which harbor innate anti-tumor immunity, and cytotoxic T cells all can participate in tumor suppression. The balance between pro- and anti-tumor leukocytes ultimately determines the behavior and fate of transformed cells; a multitude of human clinical studies have borne this out. Thus, detailed analysis of leukocyte subsets within the tumor microenvironment has become increasingly important. Here, we describe a method for analyzing infiltrating leukocyte subsets present in the tumor microenvironment in a mouse tumor model. Mouse B16 melanoma tumor cells were inoculated subcutaneously in C57BL/6 mice. At a specified time, tumors and surrounding skin were resected en bloc and processed into single cell suspensions, which were then stained for multi-color flow cytometry. Using a variety of leukocyte subset markers, we were able to compare the relative percentages of infiltrating leukocyte subsets between control and chemerin-expressing tumors. Investigators may use such a tool to study the immune presence in the tumor microenvironment and when combined with traditional caliper size measurements of tumor growth, will potentially allow them to elucidate the impact of changes in immune composition on tumor growth. Such a technique can be applied to any tumor model in which the tumor and its microenvironment can be resected and processed.

This protocol describes a method for the detailed evaluation of leukocyte subsets within the tumor microenvironment in a mouse tumor model. Chemerin-expressing B16 melanoma cells were implanted subcutaneously into syngeneic mice. Cells from the tumor microenvironment were then stained and analyzed by flow cytometry, allowing for detailed leukocyte subset analyses.

Specialized immune cells that infiltrate the tumor microenvironment regulate the growth and survival of neoplasia.&#160; Malignant cells must elude or ...

Balloon-based Injury to Induce Myointimal Hyperplasia in the Mouse Abdominal Aorta

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine balloon denudation model, which is comparable with established vessel injury models in large animals. The aorta denudation model with balloon catheters mimics the clinical setting and leads to comparable pathobiological and physiological changes. Briefly, after performing a horizontal incision in the aorta abdominalis, a balloon catheter will be inserted into the vessel, inflated, and introduced retrogradely. Inflation of the balloon will lead to intima injury and overdistension of the vessel. After removing the catheter, the aortic incision will be closed with single stiches. The model shown in this article is reproducible, easy to perform, and can be established quickly and reliably. It is especially suitable for evaluating expensive experimental therapeutic agents, which can be applied in an economical fashion. By using different knockout-mouse strains, the impact of different genes on MH development can be assessed.

This article demonstrates a murine model to study the development of myointimal hyperplasia (MH) after aortic balloon injury.

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine ...

Surgical Treatment for Benign Prostatic Hyperplasia: Holmium Laser Enucleation of the Prostate (HoLEP).

Benign prostatic hyperplasia (BPH) commonly occurs among elderly males. The condition causes symptoms in the lower urinary tract, thereby reducing the quality of life. BPH includes 2 major treatments: medication and surgery. Surgical treatment is often the most effective and final intervention. Holmium laser enucleation of the prostate (HoLEP), one of the most efficient surgical procedures for BPH, is conducted transurethrally. During surgery, the most important and most difficult part is to locate the surgical capsule of the prostate, which is consistent to the idea of anatomic surgery. Different skills may be needed in treating different parts of the prostate. HoLEP tends to be efficient and safe with good hemostatic properties, and able to treat bladder calculus, which may be the complications of BPH. The technique is particularly suitable for prostate of varying volumes and sizes. Indeed, HoLEP also presents certain disadvantages, such as a long learning curve and costly equipment. Regardless, this method may be the "new standard" and the excellent method for the surgical treatment of BPH.

Surgical Treatment for Benign Prostatic Hyperplasia: Holmium Laser Enucleation of the Prostate HoLEP.

Here we present a safe and efficient protocol, holmium laser enucleation of the prostate, to treat benign prostatic hyperplasia.

Benign prostatic hyperplasia (BPH) commonly occurs among elderly males. The condition causes symptoms in the lower urinary tract, thereby reducing the ...

Enucleation of the Prostate for the Treatment of Benign Prostatic Hyperplasia Using a 980 nm Diode Laser

In the aging male population, the occurrence of lower urinary tract symptoms (LUTS) caused by benign prostatic hyperplasia (BPH) is a common problem. Here, we introduce a new technique called 980 nm diode laser enucleation (DiLEP) to treat BPH1. Diode lasers can absorb both water and hemoglobin at the same time, so they are good for cutting and hemostasis2. The diode laser was approved by the FDA in 2007, and has been used in the treatment of BPH because of its effective cutting and hemostasis effect3. DiLEP presents several advantages over other techniques, such as TURP, HoLEP, and PVP. During the procedure, we define the boundary of a high-volume prostate and separate it into three lobes with a diode laser by burning two rings and one groove (like a Cupid&#39;s arrow). Compared to other procedures, mDiLEP has fewer intraoperative complications, a shorter learning curve, and achieves more tissue resection.

Here, we present a protocol for modified 980 nm diode laser enucleation to treat large volume benign prostatic hyperplasia.

In the aging male population, the occurrence of lower urinary tract symptoms (LUTS) caused by benign prostatic hyperplasia (BPH) is a common problem. ...

Direct Drug Delivery to Kidney via the Renal Artery

There is a need for directed injections to enable increased and specific renal exposure for efficient evaluation of drug targets in the renal research field. Accumulation of drugs in certain organs may give rise to adverse and unwanted effects, depending on the nature of injectate. To minimize spillover and/or accumulation in other tissues, the herein described method directs the formulation into the renal artery bloodstream by inserting a catheter in the infra renal aorta, just below where it branches into the renal artery, resulting in the kidney as first reached organ and distributing of formulation throughout the kidney.
This manuscript provides a detailed description of the method, as well as its challenges and difficulties. It guides the experimenter to become skillful with this type of microsurgery that requires accuracy under sterile conditions. Speed is crucial for minimizing the ischemia and practicing the procedure will increase the chance of successful injections without adverse effects. By modulating the time between injection and reperfusion as well as the injected volume, the risk of spillover to other organs is mitigated. 
Note that this technique is suitable for single dosing strategies.

This manuscript describes a method for targeted delivery to a single kidney via a catheter placed in the infrarenal abdominal aorta in the mouse.

There is a need for directed injections to enable increased and specific renal exposure for efficient evaluation of drug targets in the renal research ...

Repetitive Blood Sampling from the Subclavian Vein of Conscious Rat

Rats are widely used in pharmacokinetics (PK) and toxicokinetic (TK) studies that need to collect a certain amount of blood at specific time points to detect drug exposure. The rat blood sampling method determines the quality of the plasma and further affects the precision of the test results. The subclavian vein blood collection method described in this protocol collects blood samples repeatedly in the conscious state of animals to meet the needs of PK and TK tests. The skills of restraint handling and appropriate procedure of needle incision ensure the success rate of blood collection. It is easy to operate while ensuring the quality of plasma and at the same time catering to animal welfare. However, this method requires skilled operation, and an improper one may cause animal weakness, pain, lameness, and even mortality. The current method has been used in the testing facility for a 4-week oral toxicity study in Sprague Dawley (SD) rats with TK. The maximum amount of blood collected within 24 h did not exceed 20% of the animal's total blood. The animals' body weight was more than 200 g for males and females. The data showed that the animals' bodyweight increased steadily every week, and the clinical observation was normal after repetitive sample collection.

The present protocol describes a simple and efficient method for collecting blood from the subclavian vein in rats. It enables rapid, timely, and easily identifiable sampling without anesthesia and obtains high-quality blood through repetitive sample collection.

Rats are widely used in pharmacokinetics (PK) and toxicokinetic (TK) studies that need to collect a certain amount of blood at specific time points to ...

A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice

Murine models are employed to probe various aspects of peritoneal dialysis (PD), such as peritoneal inflammation and fibrosis. These events drive peritoneal membrane failure in humans, which remains an area of intense investigation due to its profound clinical implications in managing patients with end-stage kidney disease (ESKD). Despite the clinical importance of PD and its related complications, current experimental murine models suffer from key technical challenges that compromise the models&#39; performance. These include PD catheter migration and kinking and usually warrant earlier catheter removal. These limitations also drive the need for a greater number of animals to complete a study. Addressing these drawbacks, this study introduces technical improvements and surgical nuances to prevent commonly observed PD catheter complications in a murine model. Moreover, this modified model is validated by inducing peritoneal inflammation and fibrosis using lipopolysaccharide injections. In essence, this paper describes an improved method to create an experimental model of PD.

This article describes modifications of a procedure to implant a peritoneal dialysis catheter in a murine model to avoid major technical issues observed with the conventional techniques.

Murine models are employed to probe various aspects of peritoneal dialysis (PD), such as peritoneal inflammation and fibrosis. These events drive ...

Systematic Hearing Performance Evaluation Process for Adolescents with Cochlear Implantation at Early Ages

Cochlear implant (CI) provision is the most effective clinical treatment to restore hearing performance in individuals with profound sensorineural hearing loss (SNHL). It has been successful in providing improved speech perception outcomes, especially in quiet environments. However, speech perception performance within complex environments, lexical tone recognition, and music perception have been shown to only improve with newer fine structure coding strategies or related techniques. Therefore, the methods used to assess hearing performance in noisy environments, lexical tone recognition, and music perception are of vital importance. These assessments must reflect the postoperative outcomes and also provide guidance for the programming, rehabilitation, and application of new coding strategies. In this study, hearing performance in simple and complex situations was evaluated before and after upgrading to a fine structure strategy. The participants were a cohort of Mandarin-speaking adolescents, who were experienced CI users. The comprehensive clinical workflow involved assessments of speech in quiet conditions, speech in noisy conditions, lexical tone recognition, and music perception. This battery of tests is explained in detail, from the coding strategy to the test methods, including the test process, environment, device, material, and order. The details that require special attention are discussed, such as the position of the participants, the angle of the loudspeaker, the intensity of the sound, the noise type, the practice test, and the way of answering questions. Each test step, method, and material for speech, lexical tone, and music perception is presented in detail. Finally, the clinical results are discussed.

This paper describes a battery of tests used to clinically assess the hearing performance of adolescent, Mandarin-speaking, experienced cochlear implant users upgraded to a new fine structure coding strategy. The battery of tests include speech in quiet conditions, speech in noisy conditions, lexical tone, and music perception.

Cochlear implant (CI) provision is the most effective clinical treatment to restore hearing performance in individuals with profound sensorineural hearing ...