We thank the University of Texas Southwestern Medical Center Animal Resource Center for their help in surgical training and protocol review. We thank the Wert Lab support team for their invaluable assistance. This work has been supported by funds from the National Institute of Health (NIH P30EY030413). Biorender.com was used for the creation of cartoon schematics.

All animal experiments were approved by the Institutional Animal Care and Use Committee at UTSW (APN#2019-102840).
1. Murine orchiectomy
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Once the mouse is under anesthesia, cover the animal&#39;s eyes with lubricating eye gel to avoid the formation of eye injury, corneal desiccation, or ulcer.</li>
	<li>Before proceeding, ensure the proper plane of anesthesia by performing a toe pinch response test.</li>
	<li>Prepare the animal&#39;s surgical area before placing it on the aseptic, draped surgical field.
	<ol>
		<li>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.</li>
		<li>Using a cotton-taped applicator, apply a thin layer of depilatory cream (see Table of Materials)&#160;to the animal&#39;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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Transfer the mouse to the surgical area and make sure a heat source is available to maintain the animal&#39;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&#39;s feet to the surgical mat.</li>
	<li>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&#39;s scrotum under the visual field of the scope. A soft tape can be used to adhere the mouse&#39;s legs or abdomen to the surgical field to ensure it stays in the correct placement.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>The testicle, epididymis, attached spermatic cord, and blood vessel will be apparent when the tissues are properly externalized.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>Do not grab the blood vessel that travels along the fat pad, as pulling on this can lead to hemorrhage.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Repeat steps 1.13-1.20 on the other side of the scrotum to remove the second testicle.</li>
	<li>Prepare to close the skin incision.
	<ol>
		<li>Ensure good hemostasis of both tissue stumps. Poor hemostasis will result in residual bleeding in the surgical field. If this occurs, locate&#160;the bleeding and cauterize. This is avoided by slow and careful manipulation of the tissues with blunt instruments.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Close the skin.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Clean skin around the surgical site gently with saline and a sterilized cotton-tipped applicator to remove any dried blood or residual antiseptic scrub.</li>
</ol>
2. Murine ovariectomy
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Administer lubricating eye gel on the animal&#39;s eyes to avoid the formation of eye injury, corneal desiccation, or ulcer. Make sure a heat source is available to maintain the animal&#39;s body temperature during the operation. Ensure the proper plane of anesthesia by checking for a toe pinch response.</li>
	<li>Prepare the animal&#39;s surgical area before placing it in the surgical field.
	<ol>
		<li>Gently shave the hair on the dorsolateral region of the animal&#39;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.</li>
		<li>Using a cotton-tipped applicator, apply a layer of depilatory cream to the animal&#39;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.</li>
		<li>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&#160;is performed in step 2.5.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Grasp the abdominal wall with rat tooth forceps and make a 0.5 cm incision into it.</li>
	<li>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.</li>
	<li>Sever the ovary by performing a crush and tear technique.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Gently and slowly release the proximal clamp and check for signs of bleeding.</li>
	<li>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.</li>
	<li>Locate the borders of the incision into the muscle wall and bring them together to prepare to suture it.</li>
	<li>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.</li>
	<li>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&#160;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.</li>
	<li>To remove the other ovary, switch the mouse&#39;s position to have the other lateral side facing up. Be gentle when switching the mouse&#39;s position so as not to put too much pressure on the wound clipped skin from step 2.17.</li>
	<li>On this side, repeat steps 2.5 - 2.17.</li>
	<li>Clean skin around the surgical site gently with saline and a sterilized cotton-tipped applicator to remove any dried blood or residual antiseptic scrub.</li>
</ol>
3. Post-operative care
<ol>
	<li>Per your institutional guidelines, administer pain medicine for up to 72 h post-operation. Document the animals&#39; surgical procedure, noting the date, time, anesthetic, and analgesic used. Here, Buprenorphine slow-release and meloxicam is administered pre-operatively.</li>
	<li>After the completion of the animals&#39; 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>For mice who have undergone ovariectomy and have&#160;wound clips on the dorsolateral sides of the back, remove any rodent igloo habitats from the cage to reduce the chance of the mouse&#39;s wound clip getting caught on the igloo during the first week of healing.</li>
	<li>Keep monitoring the animal for signs of infection, pain, or surgical complications every 12 h through the first 72 h of the healing time.</li>
	<li>Remove wound clips using a wound clip remover tool 10-14 days after surgery.</li>
</ol>

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No conflicts of interest.

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 health15,16,17,18,19,20,21. Here, we detail a protocol for the fast, safe, and effective removal of murine ovaries and testis using survival surgery to deplete the sex hormones. When performed by an experienced surgeon, these procedures can take as little as 5 minutes, resulting in a very high survival rate in mice.
Some considerations when planning for a mouse surgery include the best incision site and the physiology of the animal. In the case of orchiectomy, the mouse testis can also be removed through the abdomen by a midline ventral incision13. This technique not only opens the peritoneal cavity, but also relies on the quick identification of the inguinal fat pads. For new surgeons, it is difficult to rapidly perform this technique. When mice are very young, the fat pads are also less developed and, therefore, more difficult to locate. The scrotal approach outlined in this protocol is a suitable option for researchers who are new to mouse surgery, operating on small, young mice, or who wish to minimize the chance of disturbing other organs in the peritoneal cavity by an abdominal incision.
Surgical ovariectomy involves the removal of one or both ovaries. As for most surgical procedures, safe and fast methods usually involve making the incision into the skin and underlying fascia as small as possible. This allows for reduced sutures and wound clips and faster healing. While being able to make the incisions smaller is a skill that comes with practice, being confident about the location of the incision is beneficial. Figure 2A demonstrates an example of how to locate the best incision points. Due to the small mouse anatomy, the ovary will lie near the liver, and on the left side of the mouse, near to the spleen. Accidentally probing and pulling on these organs can result in undesired bleeding or harm. We provide a detailed guide to the correct incision location and identification of the surrounding anatomical landmarks to aid in this technique and reduce the likelihood of adverse complications.
Here, we provide an example of how this procedure can be used to effectively knock down the circulating hormone levels in mice less than a week after the surgical procedure, as well as how the remaining gonadal tissue atrophies over time after the procedure. Previous methods and research detail how these can be done to generate menopause or andropause mouse models, but many of these are performed in rat models rather than mice12,22. While rats and mice have similar anatomy, the murine system has smaller anatomical structures. Here, we provide a method that reliably works in mouse models as young as 3 weeks of age. Overall, these two protocols represent a consistent way to quickly perform survival rodent orchiectomies and ovariectomies, recognize the mouse anatomy, and minimize surgical complications, creating a basis for consistent and effective results.

The testes and ovaries are the primary organs responsible for&#160;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 osteoporosis5,6,7. 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 reduced12,13. 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.

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murine orchiectomy and ovariectomy to reduce sex hormone production

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.

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 the vas deferens and spermatic blood vessels, resulting in the removal of the testis and attached epididymis (Figure 1B). The removed contents include the testis, epididymis, and inguinal fat pad, shown in Figure 1C. Successful removal of the testicles can be confirmed by visual observation of removed tissues and ensuring good hemostasis of the tissue stump prior to closing the body cavity. The success of orchiectomy can be measured by observation of well-recovered mice that demonstrate healed incision sites (Figure 1D). Mice that have undergone orchiectomy will also demonstrate a reduction in the level of testosterone measured in mouse serum as early as a week after the procedure (Figure 1E).
Surgical bilateral ovariectomy is done by incisions into the left and right dorsum of female mice (Figure 2A). The lateral incision approach can be used if only one ovary is to be removed. In this approach, the abdominal wall is incised, and the ovarian fat pad is located. The fat pad, ovary, ovarian duct, and distal uterine horn, as depicted in Figure 2B, is exteriorized, followed by severing the distal uterine horn. This technique results in the removal of the ovary and oviduct. The ovary and ovarian duct can be removed without including the distal uterine horn, but there is a risk that any remaining ovarian cells will continue to keep the mouse in estrous cycles. Including the distal uterine horn ensures that the full ovary and duct are removed, there will be a depletion of the sex hormones, and the mice will no longer cycle. Figure 2C depicts an example of the murine female reproductive organs dissected out of the body. It can be observed that the ovaries themselves are much smaller than the surrounding tissues. A dissection scope or loupes aids in the identification of these tissues. Figure 2D demonstrates the ovarian anatomy under a dissecting scope with tissue from the uterine fat pad removed, allowing for better visualization. The success of this procedure can be measured by the observation of well-recovered mice that demonstrate healed incision sites (Figure 2E).
Ovariectomy surgery is safe and efficient when performed rapidly and with minimal disturbance to surrounding tissues. Following the procedure outlined here accomplishes this by ensuring that the skin and abdominal wall incision are positioned correctly, allowing for quick location of the uterine tissues. Identifying the inferior border of the rib cage and the superior border of the leg bones ensures the incision is made near the tissue of interest. Making an incision that is about halfway between the inferior border of the ribs and superior border of the leg bones and is located about 1.5 cm lateral to the spine allows for good positioning of the surgical site when performing a lateral incision approach (Figure 2A).
Confirmation of surgical ovariectomy is shown by observation of uterine atrophy. Figure 2F shows a uterine horn dissected from a 6-month-old mouse, while Figure 2G shows a uterine horn from a 6-month-old mouse that underwent surgical ovariectomy at three months of age. The ovariectomized uterine horn appears thinner and lacks the attachment to the ovary on the distal end. This method of confirmation of the surgical technique effectively shows that the removal of the ovary resulted in atrophy of the uterine tissue.
Once tissues have been successfully identified and excised, the surgery can be completed by closure of the wounds and careful post-operative monitoring. Proper aseptic technique14 is important in ensuring good outcomes when performing survival surgery. As the mouse begins to heal from the incisions, monitoring is performed to look for signs of pain and surgical complications. A possible surgical complication is internal bleeding because of poking or nicking blood vessels while trying to incise or manipulate the tissue. This is denoted by a persistent red, flushed appearance underneath the skin around the surgical site. Avoiding complications such as these is achieved by proper localization of incision sites and aiding the surgeon's vision by use of a surgical microscope or loupes. Visual examination is recommended every 12 hours for the first three days post-surgery. Successful surgery will result in clean, healing wounds (Figure 1D, 2E).
<img alt="Orchiectomy process and outcomes; images of dissection technique and testosterone levels in mice." class="xfigimg" src="/files/ftp_upload/64379/64379fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Male murine orchiectomy. (A) A male mouse is shown in the supine position with the location of a surgical incision into the scrotal sac for the removal of the testis, which is shown in red. (B) Image taken during the orchiectomy surgical procedure depicting the point of detachment in green. (C) Dissected-out testis, fat pads, and epididymis from a male mouse. (D) A male mouse with a well-healed wound clip over the scrotal incision following orchiectomy. (E) Mouse serum testosterone concentration in ng/mL as determined by ELISA interpolation. Both groups represent 9-week-old mice. Surgical orchiectomies were performed at 8 weeks of age. N = 3 mice per group. Samples were run in triplicate. Serum samples were collected by cardiac puncture and frozen until the time of analysis. *, p &lt; 0.05. Error bars = SEM. <a href="https://www.jove.com/files/ftp_upload/64379/64379fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Mouse dissection for reproductive organ study, diagrams showing uterine horns, ovaries, oviduct." class="xfigimg" src="/files/ftp_upload/64379/64379fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Female murine ovariectomy. (A) A female mouse shown in the lateral position with the surgical incision site for the removal of a single ovary from the dorsal lateral side, shown in red. Other important anatomical landmarks are highlighted. (B) Image taken during surgery depicting the exteriorized ovary, oviduct, and distal uterine horn with the point of detachment shown in green. (C) Dissected uterine horns, ovaries, and fat pads from a female mouse. (D) A close-up view of the dissected ovary, ovarian duct, and distal portion of the uterine horn with fat from the fat pad removed, allowing for better visualization of the ovary. (E) A female mouse with a well-healed wound clip over the incision following ovariectomy. (F) Dissected mouse uterine horn. (G) Dissected mouse uterine horn 3 months post-ovariectomy procedure, showing uterine atrophy. <a href="https://www.jove.com/files/ftp_upload/64379/64379fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Sex hormones diagram: male and female mice with reproductive organs. Gender differences in biology." class="xfigimg" src="/files/ftp_upload/64379/64379fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Graphical abstract for murine ovariectomy and orchiectomy. A pictorial representation of the orchiectomy procedure for male mice involving the removal of the testis and ovariectomy procedure involving the removal of the ovaries, both of which result in a sex hormone depleted mouse model. <a href="https://www.jove.com/files/ftp_upload/64379/64379fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production at JoVE.com

Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

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

Sex hormone signaling plays a critical role in multiple organ systems as well as in the progression of various diseases, including neurodegenerative ...

murine-orchiectomy-and-ovariectomy-to-reduce-sex-hormone-production

Nanyang Technological University

Research

JoVE Journal

Medicine

8.5K Views.  UT Southwestern Medical Center. Het algemene doel van deze procedure is om de chirurgische verwijdering van muizentestikels en eierstokken op een snelle, veilige en effectieve manier aan te tonen. Muizen orchiectomie wordt uitgevoerd door een ventrale middellijnincisie in de scrotumzak, waar de teelballen worden geïsoleerd en verwijderd, gevolgd door sluiting van de huidincisie. De ovariëctomie van muizen wordt uitgevoerd door een dorsolaterale incisie in de buik waar de eierstok wordt geïsoleerd en verwijderd, gevolgd d...

Video: Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

An In Vivo Estrogen Deficiency Mouse Model for Screening Exogenous Estrogen Treatments of Cardiovascular Dysfunction After Menopause

Postmenopausal women are at greater risk of developing cardiovascular diseases than premenopausal women. Female mice ovariectomized (OVX) at weaning display increased atherosclerotic lesions in the aorta compared with female mice with intact ovarian function. However, laboratory models involving estrogen-deficient mice with atherosclerosis-prone status are lacking. This deficit is crucial because clinical estrogen deficiency in menopausal women may aggravate the incidence of pre-existing or ongoing lipid disruption and atherosclerosis. In this study, we establish an in vivo estrogen-deficient mouse model by bilateral ovariectomy via a double dorsal-lateral incision in apolipoprotein E (apoE)-/- mice. We then compare the effects of 17&#946;-estradiol and pseudoprotodioscin (PPD) (a phytoestrogen) perorally administered via hazelnut spread. We find that although PPD exerts some effect on reducing final body weight and plasma TG in OVX apoE-/- mice, it has anti-atherosclerotic and cardiac-protective capacities comparable with its 17&#946;-estradiol counterpart. PPD is a phytoestrogen that has been reported to exert anti-tumor properties. Thus, the proposed method is applicable for screening phytoestrogens via peroral administration to substitute for traditional hormone replacement therapy in postmenopausal women, which has been reported to have potentially deleterious tumorigenetic capacity. Peroral administration via hazelnut spread is noninvasive, rendering it widely applicable to many patients. This article contains step-by-step demonstrations of bilateral ovariectomy via the double dorsal-lateral incision in apoE-/- mice and peroral 17&#946;-estradiol or phytoestrogen hormone replacement via hazelnut spread. Plasma lipid and cardiovascular function analyses using echocardiography follow.

Clinically, estrogen deficiency in menopausal women may aggravate the incidence of lipid disruption and atherosclerosis. We established an in vivo estrogen deficiency model by bilateral ovariectomy via a double dorsal-lateral incision in apoE-/- mice. The mouse model is applicable for screening exogenous estrogen treatments of cardiovascular dysfunction after menopause.

Postmenopausal women are at greater risk of developing cardiovascular diseases than premenopausal women. Female mice ovariectomized (OVX) at weaning ...

Biochemistry

Orthotopic Ovarian Transplantation Procedures to Investigate the Life- and Health-span Influence of Ovarian Senescence in Female Mice

Ovarian transplantation was first conducted at Utah State University in 1963. In more recent work, heterochronic transplantation of mammalian ovaries is being used to investigate the health-protective effects of young ovaries in young females. The current procedures employ an orthotopic transplantation method, where allogenic ovaries are transplanted back to their original position in the ovarian bursa. This is in contrast to the more commonly used heterotopic transplantation of ovaries/ovarian tissue subcutaneously or under the kidney capsule. All three locations provide efficient revascularization of the transplanted tissues. However, orthotopic transplantation provides the ovary with the most natural signaling environment and is the only procedure that provides the opportunity for the animal to reproduce naturally post-operatively. One must take care to remove all endogenous ovarian tissue during the ovariectomy procedure. If any endogenous tissue remains or if only one ovary is removed, the transplanted tissue will remain dormant until the existing tissue becomes senescent. While revascularization of the transplanted ovaries occurs very quickly, the transplant recipient can take a considerable amount of time to adapt to a new hypothalamic/pituitary/gonadal/adrenal (HPG/A) axis signaling regime associated with the transplanted tissue. This normally takes about 100 days in the mouse. Therefore, transplantation experiments should be designed to accommodate this adaptation period. Typical results with ovarian transplantation will include changes in the health of the recipient that reflect the age of the transplanted ovary, rather than the chronological age of the recipient.

Described here are orthotopic ovarian transplantation protocols for modifying ovarian influence on the physiology of the recipient mouse. This surgical procedure is used to expose female mice to various types of ovarian influence and may include heterochronic transplantations and transplantation of ovaries that have undergone various ex vivo treatments.

Ovarian transplantation was first conducted at Utah State University in 1963. In more recent work, heterochronic transplantation of mammalian ovaries is ...

Developmental Biology

Assessment of Sexual Behavior of Male Mice

Sexual behavior is highly species-specific. Although rodents have slightly different sexual behaviors, mice and rats have a similar sexual behavioral pattern. The purpose of this article is to describe the hormone-induced estrus ovariectomized female model and the experimental procedure for the assessment of sexual behavior of male mice. The most important sexual behavioral elements are demonstrated in the video and illustrations. The critical steps, advantages, and limitations of the sexual behavior test are explained as well. Finally, the behavior parameters are presented, and mounting, intromission, and ejaculation processes in mating are distinguished. Behavioral parameters are assessed in terms of the occurred duration and counts during the test period.

This article describes how to perform sexual behavior tests in male mice.

Sexual behavior is highly species-specific. Although rodents have slightly different sexual behaviors, mice and rats have a similar sexual behavioral ...

Behavior

A Hyperandrogenic Mouse Model to Study Polycystic Ovary Syndrome

Hyperandrogenemia plays a critical role in reproductive and metabolic function in females and is the hallmark of polycystic ovary syndrome. Developing a lean PCOS-like mouse model that mimics women with PCOS is clinically meaningful. In this protocol, we describe such a model. By inserting a 4 mm length of DHT (dihydrotestosterone) crystal powder pellet (total length of pellet is 8 mm), and replacing it monthly, we are able to produce a PCOS-like mouse model with serum DHT levels 2 fold higher than mice not implanted with DHT (no-DHT). We observed reproductive and metabolic dysfunction without changing body weight and body composition. While exhibiting a high degree of infertility, a small subset of these PCOS-like female mice can get pregnant and their offspring show delayed puberty and increased testosterone as adults. This PCOS-like lean mouse model is a useful tool to study the pathophysiology of PCOS and the offspring from these PCOS-like dams.

We describe the development of a lean PCOS-like mouse model with&#160;dihydrotestosterone pellet to study the pathophysiology of PCOS and the offspring from these PCOS-like dams.

Hyperandrogenemia plays a critical role in reproductive and metabolic function in females and is the hallmark of polycystic ovary syndrome. Developing a ...

Generation of a Mouse Artificial Decidualization Model with Ovariectomy for Endometrial Decidualization Research

Endometrial decidualization is a unique differentiation process of the endometrium, closely related to menstruation and pregnancy. Impairment of decidualization leads to various endometrial disorders, such as infertility, recurrent miscarriage, and preterm birth. The development and use of the endometrial decidualization model in reproductive studies have been a highlight for reproductive researchers for a long time. The mouse has been extensively used in studying reproduction and decidualization. There are three well-established mouse models regarding decidualization, namely natural pregnancy decidualization (NPD), artificial decidualization (AD), and in vitro decidualization (IVD). Among them, AD is considered a reliable model for mouse decidualization, which is easy to implement and close to NPD. This paper focuses on a modified method of the generation and application process of the mouse artificial decidualization model with ovariectomy to avoid ovarian effects, which can obtain highly reproducible results with small within group variances. This method provides a good and reliable animal model for the study of endometrial decidualization.

Here, we describe the method of generating an artificial decidualization model using the ovariectomized mouse, a classic endometrial decidualization experiment in the research field of endometrial decidualization.

Endometrial decidualization is a unique differentiation process of the endometrium, closely related to menstruation and pregnancy. Impairment of ...

Methods for Studying Uterine Contributions to Pregnancy Establishment in an Ovariectomized Mouse Model

For pregnancy to be established, a viable blastocyst must successfully interact with a receptive uterine lining (endometrium) to facilitate implantation and placenta formation and enable ongoing pregnancy. The limitations to pregnancy success caused by embryonic defects are well known and have been largely overcome in recent decades with the rise of in vitro fertilization (IVF) and assisted reproductive technologies. As yet, however, the field has not overcome the limitations caused by an inadequately receptive endometrium, thus resulting in stagnating IVF success rates. Ovarian and endometrial functions are closely intertwined, as hormones produced by the ovary are responsible for the endometrium's menstrual cyclicity. As such, when using rodent models of pregnancy, it can be difficult to ascertain whether an observed result is due to an ovarian or uterine deficit. To overcome this, an ovariectomized mouse model was developed with embryo transfer or artificial decidualization to allow the study of uterine-specific contributions to pregnancy. This article will provide instructions on how to perform ovariectomy and offer insights into various techniques for supplying exogenous hormones to support successful artificial decidualization or pregnancy following embryo transfer from healthy donors. These techniques include subcutaneous injection, slow-release pellets, and osmotic mini pumps. The key advantages and disadvantages of each method will be discussed, enabling researchers to choose the best study design for their specific research question.

Pregnancy establishment is a dynamic process involving complex embryo and uterine crosstalk. The precise contributions of the maternal uterine environment to these processes remain an active area of investigation. Here, detailed protocols are provided to aid in designing in vivo animal models to address these research questions.

For pregnancy to be established, a viable blastocyst must successfully interact with a receptive uterine lining (endometrium) to facilitate implantation ...

Murine Prostate Micro-dissection and Surgical Castration

Mouse models are used extensively to study prostate cancer and other diseases. The mouse is an excellent model with which to study the prostate and has been used as a surrogate for discoveries in human prostate development and disease. Prostate micro-dissection allows consistent study of lobe-specific prostate anatomy, histology, and cellular characteristics in the absence of contamination of other tissues. Testosterone affects prostate development and disease. Androgen deprivation therapy is a common treatment for prostate cancer patients, but many prostate tumors become castration-resistant. Surgical castration of mouse models allows for the study of castration resistance and other facets of hormonal biology on the prostate. This procedure can be coupled with testosterone reintroduction, or hormonal regeneration of the prostate, a powerful method to study stem cell lineages in the prostate. Together, prostate micro-dissection and surgical castration opens up a multitude of opportunities for robust and consistent research of prostate development and disease. This manuscript describes the protocols for prostate micro-dissection and surgical castration in the laboratory mouse.

This manuscript describes the protocols for prostate micro-dissection and surgical castration in the laboratory mouse. We also depict representative results produced by these protocols. Finally, we discuss the advantages and utilization of these protocols.

Mouse models are used extensively to study prostate cancer and other diseases. The mouse is an excellent model with which to study the prostate and has ...

Ovariectomy and 17&beta;-estradiol Replacement in Rats and Mice: A Visual Demonstration

Estrogens are a family of female sexual hormones with an exceptionally wide spectrum of effects. When rats and mice are used in estrogen research they are commonly ovariectomized in order to ablate the rapidly cycling hormone production, replacing the 17&beta;-estradiol exogenously. There is, however, lack of consensus regarding how the hormone should be administered to obtain physiological serum concentrations. This is crucial since the 17&beta;-estradiol level/administration method profoundly influences the experimental results1-3. We have in a series of studies characterized the different modes of 17&beta;-estradiol administration, finding that subcutaneous silastic capsules and per-oral nut-cream Nutella are superior to commercially available slow-release pellets (produced by the company Innovative Research of America) and daily injections in terms of producing physiological serum concentrations of 17&beta;-estradiol4-6. Amongst the advantages of the nut-cream method, that previously has been used for buprenorphine administration7, is that when used for estrogen administration it resembles peroral hormone replacement therapy and is non-invasive. The subcutaneous silastic capsules are convenient and produce the most stable serum concentrations. This video article contains step-by-step demonstrations of ovariectomy and 17&beta;-estradiol hormone replacement by silastic capsules and peroral Nutella in rats and mice, followed by a discussion of important aspects of the administration procedures.

Ovariectomy and 17&#946;-estradiol Replacement in Rats and Mice: A Visual Demonstration

Ovariectomy and subsequent replacement of 17&beta;-estradiol by means of silastic capsules and peroral Nutella are demonstrated in rats and mice.

Estrogens are a family of female sexual hormones with an exceptionally wide spectrum of effects. When rats and mice are used in estrogen research they are ...

Biology

Noninvasive Monitoring of Lesion Size in a Heterologous Mouse Model of Endometriosis

Here, we describe a protocol for the implementation of a heterologous mouse model in which progression of endometriosis can be assessed in real time through noninvasive monitoring of fluorescence emitted by implanted ectopic human endometrial tissue. For this purpose, biopsies of human endometrium are obtained from donor women ongoing oocyte donation. Human endometrial fragments are cultured in the presence of adenoviruses engineered to express cDNA for the reporter fluorescent protein mCherry. Upon visualization, labeled tissues with an optimal rate of fluorescence after infection are subsequently chosen for the implantation in recipient mice. One week prior to the implantation surgery, recipient mice are oophorectomized, and estradiol pellets are placed subcutaneously to sustain the survival and growth of lesions. On the day of surgery mice are anesthetized, and peritoneal cavity accessed through a small (1.5 cm) incision by the linea-alba. Fluorescently labeled implants are tweezed, briefly soaked in glue and attached to the peritoneal layer. Incisions are sutured, and animals left to recover for a couple of days. Fluorescence emitted by endometriotic implants is usually non-invasively monitored every 3 days for 4 weeks with an in vivo imaging system. Variations in the size of endometriotic implants can be estimated in real time by quantification of the mCherry signal and normalization against the initial time-point showing maximal fluorescence intensity.
Traditional preclinical rodents of models of endometriosis do not allow non-invasive monitoring of lesion in real time but rather allow evaluation of the effects of drugs assayed at the end point. This protocol allows one to track lesions in real time and is more useful to explore the therapeutic potential of drugs in preclinical models of endometriosis. The main limitation of the model thus generated is that non-invasive monitoring is not possible over long periods of time due to the episomal expression of Ad-virus.

Here, we present a protocol for live-imaging of fluorescently labeled human endometrial fragments grafted in mice. The method allows studying the effects of drugs of choice on endometriotic lesion size through monitoring and quantification of fluorescence emitted by the fluorescent reporter on real time

Here, we describe a protocol for the implementation of a heterologous mouse model in which progression of endometriosis can be assessed in real time ...

Developing an Ovariectomized Mouse Model with Elevated FSH Level

Exploring Independent Effects of Follicle-Stimulating Hormone In Vivo in a Mouse Model

During the transition from a reproductive to a nonreproductive phase (menopause), many women experience significant physiological and pathological changes, including decreased bone mass, increased blood lipids, and increased visceral adiposity. Levels of follicle-stimulating hormone (FSH) rise during the menopausal transition. Many studies have shown that FSH in various extragonadal tissues and organs is associated with the pathogenesis of multiple diseases. Thus, building an animal model that can help study the independent effects of FSH in vivo is particularly important. In this study, C57BL/6 female mice were ovariectomized and supplemented with estradiol valerate (OVX + E2) to eliminate the effect of the hypothalamic-pituitary-gonadal axis. The OVX + E2 mice received solvent (N.S.) or different doses of recombinant FSH via intraperitoneal injection to create a mouse model (OVF) characterized by relatively stable estrogen and rising FSH levels. Thus, we successfully generated an experimental mouse model to mimic the early stage of menopause transition, characterized by elevated serum FSH levels. The OVF model has the advantages of being stable, low cost, and easy to operate, which is suitable for studies to explore the extragonadal actions of FSH. Here, we describe detailed protocols for the mouse OVF model.

Author Spotlight: Investigating the Relationship Between FSH and Pathophysiological Changes in Perimenopausal Women - Insights from a Mouse Model 

Follicle-stimulating hormone (FSH) in various extragonadal tissues and organs is associated with the pathogenesis of multiple diseases. The ovariectomized and FSH-treated mouse model (OVF) can be used to explore the extragonadal actions of FSH.

Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

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