Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

Summary

This manuscript describes a consistent way to quickly perform survival rodent orchiectomies and ovariectomies.

Abstract

Sex hormone signaling plays a critical role in multiple organ systems as well as in the progression of various diseases, including neurodegenerative disease. The manipulation of sex hormone levels in the murine model system allows for the study of their impact on organs/tissues and within disease progression. Orchiectomy - the surgical removal of the testes - and ovariectomy - the surgical removal of the ovaries - provide a method to deplete the endogenous sex hormones so that the precise hormone levels can be provided through drug or other delivery methods. Here, we provide rapid and minimally invasive methods for both orchiectomy and ovariectomy in the murine model system for the reduction of sex hormones. This protocol details the surgical preparation and excision of the testes through the scrotal sac, and excision of the ovaries via two incisions in the right and left lateral dorsum.

Introduction

The testes and ovaries are the primary organs responsible for sex hormone production. The cascade of hormonal communication leading to the production of testosterone and estrogen is a well-characterized process that begins in the hypothalamus with the release of gonadotropin-releasing hormone (GnRH)¹. The release of GnRH causes the release of luteinizing hormone (LH) and follicle-stimulating hormone (FSH) from the pituitary gland. As these hormones enter the bloodstream, they then affect other tissues in the body. The primary target of LH is the testes (in males) and the ovaries (in females)². In response to LH, the testes produce and release testosterone³. Similarly, the ovaries produce estrogen⁴. While the intended effects of these hormones are to prepare the cells and body for fertilization and ensure a functioning reproductive system, many other bodily systems can be affected.

Sex hormones have been linked to several physiological functions. For instance, estrogen helps maintain bone homeostasis by preventing the resorption of bone by osteoclasts. For this reason, ovariectomized mouse models can be used to study the physiology of bone diseases such as osteoporosis^5,6,7. Testosterone and estrogen are also research targets for many cardiovascular and neurodegenerative diseases. Recently, elevated testosterone production coupled with high-fat diet has been linked to vascular oxidative stress⁸. In the brain, changes in LH after ovariectomy have caused alterations in spatial memory⁹. Reduction of estrogens following ovariectomy has also become a model system for studying cell death in the hippocampus, as this can induce apoptosis, resulting in memory deficits¹⁰. Testosterone has also shown a role in the growth of kidneys both in mouse models and humans following kidney transplantation¹¹.

The creation of a hormone-deprived murine model allows for the study of sex hormones and their hormone cascades on various diseases or tissues. This can be accomplished by surgical removal of the testes (orchiectomy) or ovaries (ovariectomy). This procedure can be performed in mice of any strain when they are as young as weaning age (twenty-one days) or any adult age. Ovariectomy is performed in female mice, while orchiectomy is performed in male mice. By removing these organs, the levels of estrogen and testosterone, and many of their derivatives, such as progesterone, can be greatly reduced^12,13. The process of performing orchiectomies or ovariectomies in mice can be rapid and minimally invasive with the proper technique. Rapid excision of these organs in a safe and efficient manner can allow for quick surgical processing while keeping mouse numbers minimal by having a 100% survival rate when performed correctly. Here, we detail a protocol for the rapid excision of the testes and ovaries and demonstrate the proper post-surgical monitoring to enable researchers to perform this surgery quickly and safely. We also include visual examples of the sex organs and surrounding tissues to provide the surgeon with anatomical landmarks when performing this procedure.

Protocol

All animal experiments were approved by the Institutional Animal Care and Use Committee at UTSW (APN#2019-102840).

1. Murine orchiectomy

1. Prepare an aseptic working field and ensure that necessary surgical instruments have been sterilized and are readily available. Follow aseptic practices for rodent survival surgery.
2. Record the weight of the male mouse and administer the preferred method of anesthesia according to the institutional guidelines. In this protocol, 2% isoflurane was delivered via a precision vaporizer to maintain anesthesia. Before the procedure, administer pain medicine according to the institutional guidelines. In this protocol, 1.0 mg/kg Buprenorphine SR and 5 mg/kg Meloxicam were delivered subcutaneously.
3. Once the mouse is under anesthesia, cover the animal's eyes with lubricating eye gel to avoid the formation of eye injury, corneal desiccation, or ulcer.
4. Before proceeding, ensure the proper plane of anesthesia by performing a toe pinch response test.
5. Prepare the animal's surgical area before placing it on the aseptic, draped surgical field.
   1. Next, shave the inguinal and scrotal areas of the mouse using a pair of clippers.
      NOTE: Shaving the lower abdomen superior to the penis will allow for better visualization during the surgery.
   2. Using a cotton-taped applicator, apply a thin layer of depilatory cream (see Table of Materials) to the animal's skin, covering the scrotum and surrounding area that was just shaved. Wait for 30 s and remove the cream with a piece of clean gauze.
   3. Use 70% ethanol-soaked gauze sponges to wipe any remaining cream and hair off the area.
      NOTE: 70% ethanol in this step is to aid in the complete removal of depilatory cream. Antiseptic preparation of the surgical site is carried out in step 1.8.
6. Transfer the mouse to the surgical area and make sure a heat source is available to maintain the animal's body temperature during the operation. Place the animal in a supine position on a surgical drape and use soft adhesive tape to adhere the mouse's feet to the surgical mat.
7. If the testes ascend into the abdomen, carefully palpate the abdomen to have the testes descend. Use a gloved hand to apply gentle rolling pressure downward on the abdomen.
   NOTE: It is easier to perform this surgery under a dissection scope or using a pair of loupes for enhanced magnification. If performing this under a dissection scope, take a moment to position the mouse's scrotum under the visual field of the scope. A soft tape can be used to adhere the mouse's legs or abdomen to the surgical field to ensure it stays in the correct placement.
8. Scrub the skin with Betadine surgical scrub (or a similar antiseptic skin preparation scrub), followed by 70% alcohol scrub at least three times to ensure proper skin disinfection. For each scrub, scrub in a radial motion outward, so the middle of the incision site is scrubbed first and ending with the outer border of the shaved surgical area being scrubbed last.
9. At this time, put on sterile surgical gloves. For the next steps of the procedure, use aseptic techniques. Cover the mouse with a sterile surgical drape with a small incision (cut to approximately 0.5-1 inch square to fit over the surgical site) over the animal to cover the body in the drape material.
10. Create a ventral midline incision in the scrotum approximately 1 cm -1.5 cm in length using a surgical scalpel blade or similar instruments following your approved institutional animal welfare guidelines.
11. Grasping the cut border of the skin, use a blunt tipped instrument to separate the skin from the underneath tissue. Take time to do this on the lateral, superior, and inferior borders of the incision.
12. Isolate a testicle by using the spatula and forceps to move the incision in the skin to be centered atop one of the testicles. If this is difficult, go back to 1.11 and further separate the skin from the underneath layers.
13. Take a pair of curved forceps and place them on either side of the testicle. Apply a gentle downward pressure to exteriorize the testicle. Grab the thin, transparent layer of muscle that is on top of the testicle using forceps. This is the cremaster muscle. Verification of the correct muscle layer is made by observing this layer and its circulation, moving independently from the testicle beneath it.
    NOTE: Identification of correct tissue is confirmed by its transparency and ability to move independently from the underlying tissues.
    1. Manipulate the muscle tissue using forceps until the apex of the muscle is grasped. This should be the lowest portion, with the testes and cauda epididymis located underneath.
14. Grasp the cremaster muscle with forceps and move gently away from the testes and epididymis. Create a 0.5 cm incision through the cremaster muscle of the first testicle at its apex using a small pair of Vannas spring scissors.
15. Place a locking clamp on the posterior edge of the cut cremaster muscle. Use a narrow pair of hemostats or a pair of locking micro needle holders. Once the tissue is clamped, gently lay the instrument on the lateral side of the mouse to keep the tissue clamped down during the following procedural steps.
16. Use forceps to grab the superior border of the incision made in step 1.14. While the two ends of the cremaster muscle are being retracted, use another pair of forceps to reach inside the cavity and gently grasp the testicle.
    1. Pull it outward through the hole in the muscle. Be careful to watch for any signs of nicked blood vessels and blood in the surgical field.
    2. The testicle, epididymis, attached spermatic cord, and blood vessel will be apparent when the tissues are properly externalized.
    3. Near the caudal-dorsal end of the epididymis and testicle, there is a fibrous point of attachment of the testicle to the cremaster muscle. Roll the exteriorized tissue laterally to locate this insertion point. Sever this to avoid damage to the muscle and allow for further isolation and visualization of tissues. Use two pairs of forceps to perform a clamp and tear technique.
17. Look for fatty tissue around the testicle from the inguinal fat pad. Use a pair of forceps to grab onto the fat pad and gently pull to exteriorize it.
    1. Do not grab the blood vessel that travels along the fat pad, as pulling on this can lead to hemorrhage.
18. Locate the spermatic cord, blood vessels, and remaining fat pad that is proximal to the exteriorized tissues and use a pair of hemostats to clamp it. While holding slight tension on the distal testicle, cauterize the spermatic cord and blood vessels distal to the hemostat clamp.
    1. Once complete, slowly release the clamp proximal to the cauterized end and check for signs of bleeding. If present, repeat this step. It is helpful to use a pair of forceps to hold gentle tension on the distal end of the tissue being cauterized.
19. Allow the severed stump of the spermatic cord to retract back into the body. Grab the two cut ends of the cremaster muscle and bring them together. Assess the size of the cut in the tissue, and close the tissue using 4-0 or 5-0 absorbable sutures. Depending on the incision size, 1 or 2 sutures will be needed. Trim the suture ends to 0.5 cm using suture scissors.
20. Repeat steps 1.13-1.20 on the other side of the scrotum to remove the second testicle.
21. Prepare to close the skin incision.
    1. Ensure good hemostasis of both tissue stumps. Poor hemostasis will result in residual bleeding in the surgical field. If this occurs, locate the bleeding and cauterize. This is avoided by slow and careful manipulation of the tissues with blunt instruments.
    2. If blood is seen in the surgical field, use a sterilized cotton-tipped applicator to dry the area for better visualization. Use a syringe to drip saline onto the area to irrigate any blood or fluid away from the surgical field.
22. Pull the two sides of the skin incision together to prepare it for closure. Ensure the sutured ends of the cremaster incisions are not protruding into the skin incision. If needed, further trim the ends of the suture.
23. Close the skin.
    1. If using wound clips, use forceps to evert the skin and pull it away from the underlying tissue. Place one wound clip centered over the incision. Check that the skin is well clipped. If the incision is slightly too large for one wound clip to sufficiently close it, surgical skin glue can be applied to the superior and inferior portions of the incision.
    2. If using sutures, place the needed number of sutures through the skin, without grabbing the underlying tissue. This type of incision will require 2-3 single interrupted sutures using 4-0 non-absorbable suture material.
24. Clean skin around the surgical site gently with saline and a sterilized cotton-tipped applicator to remove any dried blood or residual antiseptic scrub.

2. Murine ovariectomy

1. Prepare an aseptic working field and ensure that necessary surgical instruments have been sterilized and are readily available. Follow aseptic practices for rodent survival surgery.
2. Record the weight of the female mouse and administer the preferred method of anesthesia according to your institutional guidelines. In this protocol, 2% isoflurane was delivered via a precision vaporizer to maintain anesthesia. Before the procedure, administer pain medicine according to your institutional guidelines. In this protocol, 1.0 mg/kg Buprenorphine SR and 5 mg/kg Meloxicam were delivered subcutaneously.
3. Administer lubricating eye gel on the animal's eyes to avoid the formation of eye injury, corneal desiccation, or ulcer. Make sure a heat source is available to maintain the animal's body temperature during the operation. Ensure the proper plane of anesthesia by checking for a toe pinch response.
4. Prepare the animal's surgical area before placing it in the surgical field.
   1. Gently shave the hair on the dorsolateral region of the animal's back using a pair of clippers. Remove hair from the region above the superior border of the hips and below the inferior border of the rib cage, as well as the areas between these landmarks.
   2. Using a cotton-tipped applicator, apply a layer of depilatory cream to the animal's skin, covering the area that was just shaved. Wait for 30 s and remove the cream with a piece of gauze, removing the fine hairs that remain.
   3. Use a 70% ethanol-soaked gauze sponge to wipe any remaining cream and hair from the surgical area.
      NOTE: 70% ethanol in this step is to aid in completely removing depilatory cream. Antiseptic preparation of the surgical site is performed in step 2.5.
5. Place the animal in the lateral position (for incision to remove one ovary at a time) in the surgical field and make sure that a source of heat is available. Scrub the operative area with Betadine surgical scrub (or a similar antiseptic skin preparation scrub), followed by 70% alcohol at least three times to ensure proper skin disinfection. For each scrub, scrub in a radial motion outward, so the middle of the incision is scrubbed first and the outer border of the shaved surgical area is scrubbed last.
6. At this time, put on sterile surgical gloves. For the next steps of the procedure, use aseptic techniques. Place a surgical drape with a small incision in it (cut an approximately 0.5-1 inch square in the drape to fit the surgical site) over the animal so that the body is covered in drape material.
7. Locate the ideal location for the surgical incision by finding the halfway point between the hips and ribs of the mouse (see Figure 2). Grasp the skin with forceps and make a 1.0 cm incision into the skin at this location.
8. Use a blunt tipped instrument (probe, surgical spatula, or blunt tipped hemostats) to separate the skin from the underneath muscle tissue. Take time to do this on the lateral, superior, and inferior borders of the incision.
9. Locate the dorsolateral muscle wall of the abdomen. If needed, remove fat tissue that lies between the skin and muscle layers. In mature mice, this fat tissue is more prominent. Move this fat tissue toward the caudal end of the mouse, exposing the muscle wall.
   1. Fat tissue is differentiated from the abdominal wall by color: fatty tissue appears pale white and lies more superficial, while the abdominal wall appears pink and lies deeper than the fatty tissue.
10. Grasp the abdominal wall with rat tooth forceps and make a 0.5 cm incision into it.
11. Grab one border of this incision with one pair of forceps, use a second pair to reach inside the body cavity, and find the ovary, uterine horn, and fat pad. Gently pull this through the incision in the muscle wall to separate the muscle from the underlying tissue and exteriorize it. Locate the transition between the uterine horn and the ovary.
12. Sever the ovary by performing a crush and tear technique.
    1. Place one clamp on the distal end of the uterine horn and place the second clamp just distal to the first. While applying firm pressure with both clamps, move the more distal one away from the body, tearing the connection between them. To minimize bleeding and tissue damage, keep the proximal clamp stationary during this process and be careful not to also pull it away from the body.
    2. Alternatively, use a cautery. Use a pair of forceps or hemostats to clamp the uterine horn just proximal to the desired cautery point. Use a cautery tool to sever the tissues.
13. Gently and slowly release the proximal clamp and check for signs of bleeding.
14. Allow the tissue stump to retreat into the body cavity. Grab the abdominal wall and gently pull upwards. Do not force the tissue stump inwards, as this increases the likelihood of bleeding.
15. Locate the borders of the incision into the muscle wall and bring them together to prepare to suture it.
16. Place 1-2 single interrupted sutures into the abdominal muscle wall using 4-0 absorbable suture material. Trim the suture ends to 0.5 cm using suture scissors.
17. Close the incision by allowing the skin to return to its natural resting state and then evert it upwards, being careful not to grab underlying tissue or the ends of the suture material. Place 1-2 wound clips on the everted skin. Based on your institutional guidelines, the skin can also be closed using 4-0 non-absorbable suture material.
18. To remove the other ovary, switch the mouse's position to have the other lateral side facing up. Be gentle when switching the mouse's position so as not to put too much pressure on the wound clipped skin from step 2.17.
19. On this side, repeat steps 2.5 - 2.17.
20. Clean skin around the surgical site gently with saline and a sterilized cotton-tipped applicator to remove any dried blood or residual antiseptic scrub.

3. Post-operative care

1. Per your institutional guidelines, administer pain medicine for up to 72 h post-operation. Document the animals' surgical procedure, noting the date, time, anesthetic, and analgesic used. Here, Buprenorphine slow-release and meloxicam is administered pre-operatively.
2. After the completion of the animals' surgery, transfer them to a clean cage lined with a dry paper towel. Place this cage 2/3 on a heat source, leaving the other 1/3 of the cage off heat. Do not place mice together in a cage until they have recovered from anesthesia.
3. Allow the animal to regain the ability to freely walk around the cage before placing them back into a clean cage with normal animal bedding. At this time, mice can be housed together if all of them have recovered from anesthesia.
4. For the first several days following surgery, ensure moist food and water is easily accessible. Place some moistened food on the bottom of the cage. Frequently check on the mouse surgical site and look for any signs of infection or bleeding.
5. Following surgery, ensure mice are either housed alone or with other animals that have undergone the same procedure simultaneously. A mouse recovering from surgery located in the same cage as one without surgery can introduce a hazard and should be avoided.
6. For mice who have undergone ovariectomy and have wound clips on the dorsolateral sides of the back, remove any rodent igloo habitats from the cage to reduce the chance of the mouse's wound clip getting caught on the igloo during the first week of healing.
7. Keep monitoring the animal for signs of infection, pain, or surgical complications every 12 h through the first 72 h of the healing time.
8. Remove wound clips using a wound clip remover tool 10-14 days after surgery.

Results

The procedure presented here is performed in one- to three-month-old mice in the C57BL/6J background. Male mice weighed 16-28 g, and female mice weighed 14-24 g at the time of the procedure. This procedure has been optimized to be applicable for mice of many ages, from weaning through adulthood.

Surgical orchiectomy involves a single skin incision in the ventral scrotal sac, as depicted in Figure 1A. Both testes are removed one at a time and are severed through th...

Discussion

Surgical removal of testes and ovaries allows for studying murine physiology under controlled hormone deprivation. This technique is important for many fields of science, including neurodegeneration, mineral metabolism, cardiovascular, and reproductive health^{15,16,17,18,19,20,21}. Here, we de...

Materials

Name	Company	Catalog Number	Comments
1mL Syringe	BD	309659
30G 1/2" Needle	BD	305106
AutoClip System	Fine Science Tools	12020-00
Betadine Solution	Fisher Scientific	NC0158124
Cotton-Tipped Applicators	Fisher Scientific	10-000-692
Double -ended Micro Spatula	Fine Science Tools	10091-12
Galilean Loupes	Fine Science Tools	28050-30	Optional, can provide better clarity during procedure
Gauze Sponges, 4"x4"	Fisher Scientific	13-761-52
Graefe Forceps	Fine Science Tools	11150-10
High Temp Cautery Kit	Fine Science Tools	18010-00	Using the fine tip attachment
Needle Holders	Fine Science Tools	12001-13
PGA Absorbable Suture:4-0 / NFS-2 Reverse Cutting 19MM / 30 IN	Covetrus	29242	4-0 or 5-0 Absorbable sutures are best
Rodent Warming pad	Kent Scientific	RT-0515
Sterile Alcohol Prep Pads	Fisher Scientific	22-363-750
Straight Locking Micro Needle Holders	Fine Science Tools	12060-01
Surgical Scissors	Fine Science Tools	140-60-09
Vannas Spring Scissors - 2.5mm Cutting Edge	Fine Science Tools	15000-08
Veet Sensitive Hair Remover Gel Cream	Amazon	N/A
Wahl Professional Animal Compact Trimmer and Grooming Kit, Blue	Amazon	#9861-900