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 de...

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 th...

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

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8.1K Views.  UT Southwestern Medical Center. This manuscript describes a consistent way to quickly perform survival rodent orchiectomies and ovariectomies. 

Video: Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

Murine Echocardiography and Ultrasound Imaging

Rodent models of cardiac pathophysiology represent a valuable research tool to investigate mechanism of disease as well as test new therapeutics.1 Echocardiography provides a powerful, non-invasive tool to serially assess cardiac morphometry and function in a living animal.2 However, using this technique on mice poses unique challenges owing to the small size and rapid heart rate of these animals.3 Until recently, few ultrasound systems were capable of performing quality echocardiography on mice, and those generally lacked the image resolution and frame rate necessary to obtain truly quantitative measurements. Newly released systems such as the VisualSonics Vevo2100 provide new tools for researchers to carefully and non-invasively investigate cardiac function in mice. This system generates high resolution images and provides analysis capabilities similar to those used with human patients. Although color Doppler has been available for over 30 years in humans, this valuable technology has only recently been possible in rodent ultrasound.4,5 Color Doppler has broad applications for echocardiography, including the ability to quickly assess flow directionality in vessels and through valves, and to rapidly identify valve regurgitation. Strain analysis is a critical advance that is utilized to quantitatively measure regional myocardial function.6 This technique has the potential to detect changes in pathology, or resolution of pathology, earlier than conventional techniques. Coupled with the addition of three-dimensional image reconstruction, volumetric assessment of whole-organs is possible, including visualization and assessment of cardiac and vascular structures. Murine-compatible contrast imaging can also allow for volumetric measurements and tissue perfusion assessment.

This video demonstrates use of a rail-mounted high-frequency ultrasound probe to perform echocardiography on an anesthetized mouse. The methods describe both conventional two-dimensional and M-mode measurements of cardiac function in addition to newer, more powerful tools such as color Doppler, strain analysis, as well as general and targeted contrast imaging.

Rodent models of cardiac pathophysiology represent a valuable research tool to investigate mechanism of disease as well as test new therapeutics.1  ...

The Measurement and Treatment of Suppression in Amblyopia

Amblyopia, a developmental disorder of the visual cortex, is one of the leading causes of visual dysfunction in the working age population. Current estimates put the prevalence of amblyopia at approximately 1-3%1-3, the majority of cases being monocular2. Amblyopia is most frequently caused by ocular misalignment (strabismus), blur induced by unequal refractive error (anisometropia), and in some cases by form deprivation.
Although amblyopia is initially caused by abnormal visual input in infancy, once established, the visual deficit often remains when normal visual input has been restored using surgery and/or refractive correction. This is because amblyopia is the result of abnormal visual cortex development rather than a problem with the amblyopic eye itself4,5 . Amblyopia is characterized by both monocular and binocular deficits6,7 which include impaired visual acuity and poor or absent stereopsis respectively. The visual dysfunction in amblyopia is often associated with a strong suppression of the inputs from the amblyopic eye under binocular viewing conditions8. Recent work has indicated that suppression may play a central role in both the monocular and binocular deficits associated with amblyopia9,10 .
Current clinical tests for suppression tend to verify the presence or absence of suppression rather than giving a quantitative measurement of the degree of suppression. Here we describe a technique for measuring amblyopic suppression with a compact, portable device11,12 . The device consists of a laptop computer connected to a pair of virtual reality goggles. The novelty of the technique lies in the way we present visual stimuli to measure suppression. Stimuli are shown to the amblyopic eye at high contrast while the contrast of the stimuli shown to the non-amblyopic eye are varied. Patients perform a simple signal/noise task that allows for a precise measurement of the strength of excitatory binocular interactions. The contrast offset at which neither eye has a performance advantage is a measure of the "balance point" and is a direct measure of suppression. This technique has been validated psychophysically both in control13,14 and patient6,9,11 populations.
In addition to measuring suppression this technique also forms the basis of a novel form of treatment to decrease suppression over time and improve binocular and often monocular function in adult patients with amblyopia12,15,16 . This new treatment approach can be deployed either on the goggle system described above or on a specially modified iPod touch device15.

Amblyopia is a developmental disorder of the visual cortex that is often accompanied by strong suppression of one eye. We present a new technique for measuring and treating interocular suppression in patients with amblyopia that can be deployed using virtual reality goggles or a portable iPod Touch device.

Amblyopia, a developmental disorder of the visual cortex, is one of the leading causes of visual dysfunction in the working age population. Current ...

Isolating and Using Sections of Bovine Mesenteric Artery and Vein as a Bioassay to Test for Vasoactivity in the Small Intestine

Mammalian gastrointestinal systems are constantly exposed to compounds (desirable and undesirable) that can have an effect on blood flow to and from that system. Changes in blood flow to the small intestine can result in effects on the absorptive functions of the organ. Particular interest in toxins liberated from feedstuffs through fermentative and digestive processes has developed in ruminants as an area where productive efficiencies could be improved. The video associated with this article describes an in vitro bioassay developed to screen compounds for vasoactivity in isolated cross-sections of bovine mesenteric artery and vein using a multimyograph. Once the blood vessels are mounted and equilibrated in the myograph, the bioassay itself can be used: as a screening tool to evaluate the contractile response or vasoactivity of compounds of interest; determine the presence of receptor types by pharmacologically targeting receptors with specific agonists; determine the role of a receptor with the presence of one or more antagonists; or determine potential interactions of compounds of interest with antagonists. Through all of this, data are collected real-time, tissue collected from a single animal can be exposed to a large number of different experimental treatments (an in vitro advantage), and represents vasculature on either side of the capillary bed to provide an accurate picture of what could be happening in the afferent and efferent blood supply supporting the small intestine.

The small intestine is frequently exposed to toxins that can influence blood flow and negatively impact nutrient absorption. Using a multimyograph and mesenteric artery and vein isolates, compounds or toxins of interest can be screened for vasoactivity.

Mammalian gastrointestinal systems are constantly exposed to compounds (desirable and undesirable) that can have an effect on blood flow to and from that ...

An Ex vivo Model to Study Hormone Action in the Human Breast

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial cells tend to lose hormone receptor expression. Widely used hormone receptor positive breast cancer cell lines are of limited relevance to the in vivo situation. Here, we describe an ex vivo model to study hormone action in the human breast. Fresh human breast tissue specimens from surgical discard material such as reduction mammoplasties or mammectomies are mechanically and enzymatically digested to obtain tissue fragments containing ducts and lobules and multiple stromal cell types. These tissue microstructures kept in basal medium without growth factors preserve their intercellular contacts, the tissue architecture, and remain hormone responsive for several days. They are readily processed for RNA and protein extraction, histological analysis or stored in freezing medium. Fluorescence activated cell sorting (FACS) can be used to enrich for specific cell populations. This protocol provides a straightforward, standard approach for translational studies with highly complex, varied human specimens.

An Ex vivo Model to Study Hormone Action in the Human Breast

We have developed a novel ex vivo model to study hormone action in the human breast. It is based on tissue microstructures isolated from surgical breast tissue specimens which preserve tissue architecture, intercellular interactions, and paracrine signaling.

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial ...

Povidone Iodine Rectal Preparation at Time of Prostate Needle Biopsy is a Simple and Reproducible Means to Reduce Risk of Procedural Infection

Single institution and population-based studies highlight that infectious complications following transrectal ultrasound guided prostate needle biopsy (TRUS PNB) are increasing. Such infections are largely attributable to quinolone resistant microorganisms which colonize the rectal vault and are translocated into the bloodstream during the biopsy procedure. A povidone iodine rectal preparation (PIRP) at time of biopsy is a simple, reproducible method to reduce rectal microorganism colony counts and therefore resultant infections following TRUS PNB.
All patients are administered three days of oral antibiotic therapy prior to biopsy. The PIRP technique involves initially positioning the patient in the standard manner for a TRUS PNB. Following digital rectal examination, 15 ml&#160;of a 10% solution of commercially available povidone iodine is mixed with 5 ml&#160;of 1% lidocaine jelly to create slurry. A 4 cm x 4 cm sterile gauze is soaked in this slurry and then inserted into the rectal vault for 2 min after which it is removed. Thereafter, a disposable cotton gynecologic swab is used to paint both the perianal area and the rectal vault to a distance of 3 cm from the anus. The povidone iodine solution is then allowed to dry for 2 - 3 min prior to proceeding with standard transrectal ultrasonography and subsequent biopsy.
This PIRP technique has been in practice at our institution since March of 2012 with an associated reduction of post-biopsy infections from 4.3% to 0.6% (p = 0.02). The principal advantage of this prophylaxis regimen is its simplicity and reproducibility with use of an easily available, inexpensive agent to reduce infections. Furthermore, the technique avoids exposing patients to additional systemic antibiotics with potential further propagation of multi-drug resistant organisms. Usage of PIRP at TRUS PNB, however, is not applicable for patients with iodine or shellfish allergies.

Here, we present a protocol for use at the time of transrectal ultrasound guided prostate needle biopsy (TRUS PNB) that is a simple and cost-effective means to reduce infections following the procedure.

Single institution and population-based studies highlight that infectious complications following transrectal ultrasound guided prostate needle biopsy ...

Transplantation of Adipose Tissue-Derived Stem Cell Sheet to Reduce Leakage After Partial Colectomy in A Rat Model

Anastomotic leakage is a disastrous complication after colorectal surgery. Although current methods for leakage prevention have different levels of clinical efficacy, they are until now imperfect solutions. Stem cell therapy using ASC sheets could provide a solution to this problem. ASCs are considered as promising candidates for promoting tissue healing because of their trophic and immunomodulatory properties. Here, we provide methods to produce high-density ASC sheets, that are transplanted onto a colorectal anastomosis in a rat model to reduce the leakage. ASCs formed cell sheets in thermo-responsive culture dishes that could be easily detached. On the day of the transplantation, a partial colectomy with a 5-suture colorectal anastomosis was performed. Animals were immediately transplanted with 1 ASC sheet per rat. ASC sheets adhered spontaneously to the anastomosis without any glue, suture, or any biomaterial. Animal groups were sacrificed 3 and 7 days postoperatively. Compared to transplanted animals, the incidence of anastomotic abscesses and leakage was higher in control animals. In our model, the transplantation of ASC sheets after colorectal anastomosis was successful and associated with a lower leakage rate.

The preparation and transplantation of adipose tissue derived stem cell (ASC) sheets onto insufficiently sutured colorectal anastomoses in a rat model is presented. This novel application shows that ASC sheets can reduce colorectal anastomosis leakage.

Anastomotic leakage is a disastrous complication after colorectal surgery. Although current methods for leakage prevention have different levels of ...

A Clinical Trial Assessing the Safety, Efficacy, and Delivery of Olive-Oil-Based Three-Chamber Bags for Parenteral Nutrition

Limited evidence exists to precisely estimate efficacy and safety differences between parenteral nutrition (PN) prepared using olive-oil-based three-chamber bags (3CBs) and soybean-oil-based compounded bags (CoBs) in hospitalized adult patients. We designed a multicenter, randomized, prospective, open-label, noninferiority protocol to compare the efficacy, safety, and distribution of olive-oil-based 3CBs and soybean-oil-based CoB formulations in adult Chinese patients requiring PN during surgical intervention. Subjects were randomized to receive either one of the study treatments using an interactive voice or web-based recognition system in accordance with the randomization code. Randomization was further stratified based on the study site and surgical category. Both treatment groups received similar amounts of calories and protein. In addition, the two study treatments contained a similar composition of the amino-acid component. The only difference between the two PN formulations was the lipid constitution. The duration of administration of study treatments was a minimum of 5 days up to a maximum of 14 days after the surgical procedure. The primary efficacy endpoint was serum prealbumin levels on day 5 of the study. Noninferiority was proved if the anti-log of the lower bound of the 95% confidence interval (CI) of the treatment difference was at least 0.80. Other efficacy measures included treatment preparation time; duration to achieve tolerability of oral nutrition; associated infectious complications; length of hospitalization; and laboratory assessment of markers of nutrition, inflammation, metabolism, and oxidative stress. A total of 458 patients were enrolled in the study. The results showed that olive-oil-based 3CBs were non-inferior to soybean-based CoBs, besides being well tolerated. The infection rate was found to be significantly lower in the olive-oil-based 3CB group. Thus, this study may be used as a reference for future research on lipid emulsion and 3CBs.

Here, we present a protocol for a study comparing the efficacy, safety, and delivery of olive-oil-based 3CB and soybean-oil-based CoB formulations in adults requiring parenteral nutrition. The results revealed that olive-oil-based 3CBs is non-inferior and well tolerated compared to soybean formulations.

Limited evidence exists to precisely estimate efficacy and safety differences between parenteral nutrition (PN) prepared using olive-oil-based ...

Mechanisms Underlying Gut Hormone Secretion Using the Isolated Perfused Rat Small Intestine

The gut is the largest endocrine organ of the body, producing more than 15 different peptide hormones that regulate appetite and food intake, digestion, nutrient absorption and distribution, and post-prandial glucose excursions. Understanding the molecular mechanisms that regulate gut hormone secretion is fundamental for understanding and translating gut hormone physiology. Traditionally, the mechanisms underlying gut hormone secretion are either studied in vivo (in experimental animals or humans) or using gut hormone-secreting primary mucosal cell cultures or cell lines. Here, we introduce an isolated perfused rat small intestine as an alternative method for studying gut hormone secretion. The virtues of this model are that it relies on the intact gut, meaning that it recapitulates most of the physiologically important parameters responsible for the secretion in in vivo studies, including mucosal polarization, paracrine relationships and routes of perfusion/stimulus exposure. In addition, and unlike in vivo studies, the isolated perfused rat small intestine allows for almost complete experimental control and direct assessment of secretion. In contrast to in vitro studies, it is possible to study both the magnitude and the dynamics of secretion and to address important questions, such as what stimuli cause secretion of different gut hormones, from which side of the gut (luminal or vascular) is secretion stimulated, and to analyze in detail molecular sensors underlying the secretory response. In addition, the preparation is a powerful model for the study of intestinal absorption and details regarding the dynamics of intestinal absorption including the responsible transporters.

Here, we present a powerful and physiological model to study the molecular mechanisms underlying gut hormone secretion and intestinal absorption &#8212; the isolated perfused rat small intestine.

The gut is the largest endocrine organ of the body, producing more than 15 different peptide hormones that regulate appetite and food intake, digestion, ...

Cooling or Warming the Esophagus to Reduce Esophageal Injury During Left Atrial Ablation in the Treatment of Atrial Fibrillation

Ablation of the left atrium using either radiofrequency (RF) or cryothermal energy is an effective treatment for atrial fibrillation (AF) and is the most frequent type of cardiac ablation procedure performed. Although generally safe, collateral injury to surrounding structures, particularly the esophagus, remains a concern. Cooling or warming the esophagus to counteract the heat from RF ablation, or the cold from cryoablation, is a method that is used to reduce thermal esophageal injury, and there are increasing data to support this approach. This protocol describes the use of a commercially available esophageal temperature management device to cool or warm the esophagus to reduce esophageal injury during left atrial ablation. The temperature management device is powered by standard water-blanket heat exchangers, and is shaped like a standard orogastric tube placed for gastric suctioning and decompression. Water circulates through the device in a closed-loop circuit, transferring heat across the silicone walls of the device, through the esophageal wall. Placement of the device is analogous to the placement of a typical orogastric tube, and temperature is adjusted via the external heat-exchanger console.

The goal of this protocol is to describe the use of esophageal temperature modulation to counteract esophageal thermal injury from left atrial ablation for the treatment of atrial fibrillation.

Ablation of the left atrium using either radiofrequency (RF) or cryothermal energy is an effective treatment for atrial fibrillation (AF) and is the most ...

Network Pharmacology Prediction and Experimental Validation of Trichosanthes-Fritillaria thunbergii Action Mechanism Against Lung Adenocarcinoma

We aimed to study the mechanism of Trichosanthes-Fritillaria thunbergii in treating lung adenocarcinoma (LUAD) based on network pharmacology and experimental verification. The effective components and potential targets of Trichosanthis and Fritillaria thunbergii were collected by high-throughput experiment and reference-guided (HERB) database of traditional Chinese medicine and a similarity ensemble approach (SEA) database, and the LUAD-related targets were queried by the GeneCards and Online Mendelian Inheritance in Man (OMIM) databases. A drug-component-disease-target network was constructed by Cytoscape software. Protein-protein interaction (PPI) network, gene ontology (GO) function, and Kyoto encyclopedia of genes and genomes (KEGG) pathway enrichment analyses were conducted to obtain core targets and key pathways. An aqueous extract of Trichosanthes-Fritillaria thunbergii and A549 cells were used for the subsequent experimental validation. Through the HERB database and literature search, 31 effective compounds and 157 potential target genes of Trichosanthes-Fritillaria thunbergii were screened, of which 144 were regulatory targets of Trichosanthes-Fritillaria thunbergii in the treatment of lung adenocarcinoma. The GO functional enrichment analysis showed that the mechanism of action of Trichosanthes-Fritillaria thunbergii against lung adenocarcinoma is mainly protein phosphorylation. The KEGG pathway enrichment analysis suggested that the treatment of lung adenocarcinoma by Trichosanthes-Fritillaria thunbergii mainly involves the PI3K/AKT signaling pathway. The experimental validation showed that an aqueous extract of Trichosanthes-Fritillaria thunbergii could inhibit the proliferation of A549 cells and the phosphorylation of AKT. Through network pharmacology and experimental validation, it was verified that the PI3K/AKT signaling pathway plays a vital role in the action of Trichosanthes-Fritillaria thunbergii in treating lung adenocarcinoma.

Network Pharmacology Prediction and Experimental Validation of Trichosanthes-Fritillaria thunbergii Action Mechanism Against Lung Adenocarcinoma

This study reveals the mechanism of Trichosanthes-Fritillaria thunbergii in treating lung adenocarcinoma based on network pharmacology and experimental verification. The study also demonstrates that the PI3K/AKT signaling pathway plays a vital role in the action of Trichosanthes-Fritillaria thunbergii in treating lung adenocarcinoma.

Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

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Murine Echocardiography and Ultrasound Imaging

The Measurement and Treatment of Suppression in Amblyopia

Isolating and Using Sections of Bovine Mesenteric Artery and Vein as a Bioassay to Test for Vasoactivity in the Small Intestine

An Ex vivo Model to Study Hormone Action in the Human Breast

Povidone Iodine Rectal Preparation at Time of Prostate Needle Biopsy is a Simple and Reproducible Means to Reduce Risk of Procedural Infection

Transplantation of Adipose Tissue-Derived Stem Cell Sheet to Reduce Leakage After Partial Colectomy in A Rat Model

A Clinical Trial Assessing the Safety, Efficacy, and Delivery of Olive-Oil-Based Three-Chamber Bags for Parenteral Nutrition

Mechanisms Underlying Gut Hormone Secretion Using the Isolated Perfused Rat Small Intestine

Cooling or Warming the Esophagus to Reduce Esophageal Injury During Left Atrial Ablation in the Treatment of Atrial Fibrillation

Network Pharmacology Prediction and Experimental Validation of Trichosanthes-Fritillaria thunbergii Action Mechanism Against Lung Adenocarcinoma