Name: 비디오: 쥐 수직 소매 위 절제술: 실험용 마우스 모델에서 위를 축소하는 수술 절차 - 개념
Uploaded: 2023-04-30T13:32:37.000+00:00
Duration: 1 min 39 s
Description: 1.1K 조회수. 출처: Garibay, D., Cummings, B. P. 수직 소매 위절제술의 쥐 모델. J. Vis. 특급 (2017) 이 비디오에서는 실험용 마우스 모델에서 수직 소매 위 절제술 절차가 수행됩니다. 이 절차는 더 큰 곡률을 따라 위의 상당 부분을 제거하는 데 도움이 됩니다.

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Vertical Sleeve Gastrectomy and Sham Procedures
<ol>
	<li>On the day of surgery, weigh mice to obtain a baseline weight measurement.</li>
	<li>Equip the surgical room with an anesthesia system, heated water bath, and heating pad. Set the water bath to 37 &#176;C and use it to warm 0.9% saline solution for irrigation. Clean the surgical field with 70% ethanol and use sterile technique to open autoclaved surgery pack and tools. 
	NOTE: The heating pad is used to keep the animal warm during surgery and recovery from anesthesia.</li>
	<li>Using an induction chamber, anesthetize the mouse with 5% isoflurane and O2&#160;flow rate of 1 L/min. Confirm that the mouse is in the appropriate plane of anesthesia by performing a toe pinch. Maintain the mouse with 1-3% isoflurane and an O2&#160;flow rate of 0.6 L/min once a consistent plane of anesthesia has been achieved.</li>
	<li>Once the mouse is anesthetized, place eye ointment (see Table of Materials) on its eyes to keep them moist during surgery. At this time, also administer an analgesic, such as meloxicam (2 mg/kg subcutaneously). Clip the hair from the umbilicus to the axilla holding the skin down gently to provide tension.
	<ol>
		<li>As the mouse&#39;s skin is thin and delicate, take care to not cause skin lesions when clipping the hair. Clean the skin with povidone-iodine and alcohol.</li>
		<li>Don sterile gloves and prepare the sterile surgical field and instruments. Perform the rest of the procedure using sterile technique. Prepare sterile surgical field by placing sterile surgical drapes on either side of the mouse. Use autoclaved tin foil to create a surgical drape for the mouse.</li>
		<li>Cut a small hole in the tin foil to allow access to the mouse&#39;s abdomen.</li>
	</ol>
	</li>
	<li>Make an incision of the skin from the mid-abdomen (umbilicus) to the level of the xiphoid cartilage using an iris scissor. Identify the linea alba and use iris scissors to cut through the body wall along the linea alba. Use CTAs to gently elevate the stomach out of the abdomen and then bluntly dissect the greater omentum off of the greater curvature of the stomach.</li>
	<li>Ligate the short gastric artery that runs between the fundus of the stomach and the spleen by placing two ligatures using 7-0 monofilament absorbable suture. Use spring scissors to cut between the two ligatures. Once the artery is transected, fully exteriorize the stomach from the abdominal cavity. Place gauze under the stomach and wet with saline to keep the tissue moist.</li>
	<li>For the VSG procedure, ligate the prominent branches of the gastric artery and vein with 7-0 monofilament absorbable suture with a taper needle using 3 throws for each knot. Place the ligatures just below (i.e., towards the lesser curvature of the stomach) the intended line of transection. 
	NOTE: The intended line of transection starts at ~2 mm above (meaning towards the greater curvature of the stomach) the cardiac notch of the stomach and at least 2 mm below the margo pilcatus and extends to the proximal end of the right lobe of the pancreas. Typically, there are 4 vessels per gastric wall that need to be ligated; however, this may vary between mice.
	<ol>
		<li>To prevent spillage of gastric contents during the gastrectomy, place a simple continuous line of suture passing through both gastric walls just below the intended line of transection using a 6-0 monofilament absorbable suture with a taper needle. Begin the suture line to the surgeon&#39;s right of the esophagus and below the margo pilcatus and end just above the pancreas.</li>
		<li>Place thin-tipped hemostats above the suture line and use spring scissors to cut between the suture line and hemostats. Remove the transected gastric tissue from the sterile surgical field. Use CTAs to clean blood and digesta off of the stomach.</li>
		<li>Reinforce apposition of the gastric walls using 6-0 monofilament absorbable suture with a taper needle using a simple discontinuous pattern. 
		NOTE: A simple discontinuous suture pattern is recommended since this provides more secure closure than the simple continuous suture pattern.</li>
		<li>Flush the gastric remnant with saline throughout the procedure in order to keep the tissue moist and clean. Use a 20G gavage needle attached to a 20 mL syringe to perform gastric lavage with saline. Use this size to ensure adequate pressure is provided without risking gastric tissue damage from excessive pressure or accidental gastric tissue damage from the use of a regular pointed needle.</li>
		<li>Use a minimum of 20 knots to securely close the stomach; pay attention to closure along the esophageal side, as this side is more difficult to access and therefore is often a site of dehiscence.</li>
		<li>Ensure that there are no leaks by gently pressing on the stomach with CTAs. If leaks are identified, place additional simple discontinuous knots on the areas of leakage and then leak test again. Leak test until no leakage is detected.</li>
		<li>Do a final thorough lavage of the stomach using at least 60 mL of saline to ensure that no infectious particulate matter has been left behind.</li>
	</ol>
	</li>
	<li>Place the stomach back into the abdominal cavity under the liver using a CTA. Place CTAs along the dorsal aspect of the abdominal cavity to absorb all excess fluid. Using a blunt 18G needle, inject lactated Ringer&#39;s solution (LRS) with or without antibiotics (0.5 mL LRS +/- 20 mg/kg Enrofloxacin) directly into the abdominal cavity, just prior to closure. 
	NOTE: This replaces fluid loss experienced during surgery and provides a method for direct application of antibiotics to the surgical site to assist with recovery.</li>
	<li>Close the abdominal muscle layer using a 6-0 monofilament absorbable suture with a taper needle in a simple discontinuous pattern. Then, close the skin layer with 6-0 monofilament absorbable suture in a simple continuous pattern.</li>
	<li>Place tissue adhesive on the skin and fold the skin over the suture line to bury the suture so that the mouse cannot disrupt the wound closure post-operatively.</li>
	<li>Turn the isoflurane off and let the mouse recover on the heating pad for 10-15 min before returning it to its home cage. Do not leave the animal unattended until it has regained consciousness and is able to move.</li>
	<li>Keep animals housed singly until fully recovered from surgery. 
	NOTE: If accurate food intake measurements are to be obtained throughout the study, mice should continue to be singly housed throughout the study. There is a risk of foreign body obstruction from eating home cage bedding. Animals should be housed in cages free of bedding except for nesting material to provide enrichment (see&#160;Table of Materials).</li>
</ol>

<table><tbody><tr><td>6-0 Suture</td><td>Ethicon</td><td>Z432</td><td>Monofilamentabsorbable/taper</td></tr><tr><td>7-0 Suture&amp;nbsp;</td><td>Covidien</td><td>8866127-01</td><td>Monofilamentabsorbable/taper</td></tr><tr><td>Cotton swabs</td><td>Fisherbrand</td><td>23-400-118</td><td>Small</td></tr><tr><td>Cotton swabs</td><td>Fisherbrand</td><td>233-400-101</td><td>Large</td></tr><tr><td>Foil</td><td>Various</td><td></td><td></td></tr><tr><td>Surgery drape</td><td>Various</td><td></td><td></td></tr><tr><td>0.9% Saline solution</td><td>Various</td><td></td><td></td></tr><tr><td>Microneedle driver</td><td>Fine Science Tools</td><td>12075-14</td><td></td></tr><tr><td>Spring scissors</td><td>Fine Science Tools</td><td>15396-00</td><td></td></tr><tr><td>Alcohol</td><td>Various</td><td></td><td></td></tr><tr><td>Eye Ointment</td><td>Paralube&amp;Acirc;&amp;reg;Vet Ointment</td><td>17033-211-38</td><td></td></tr><tr><td>Meloxicam (Metacam)</td><td>Boehringer Ingelheim</td><td>141-213</td><td>5 mg/ml</td></tr><tr><td>Thin tipped hemostats</td><td>Fine Science Tools</td><td>13021-12</td><td></td></tr><tr><td>20 mL syringe</td><td>Various</td><td></td><td></td></tr><tr><td>Enrofloxacin</td><td>Baytril</td><td>08713254-186599</td><td>22.7 mg/ml</td></tr></tbody></table>

murine vertical sleeve gastrectomy: a surgical procedure to downsize stomach in experimental mouse models

Source: Garibay, D., Cummings, B. P. <a href="https://www.jove.com/video/56534" target="_blank">A Murine Model of Vertical Sleeve Gastrectomy.</a> J. Vis. Exp. (2017)
In this video, a vertical sleeve gastrectomy procedure is performed on an experimental mouse model. This procedure helps remove a significant portion of the stomach along its greater curvature.

Murine Vertical Sleeve Gastrectomy: A Surgical Procedure to Downsize Stomach in Experimental Mouse Models

Source: Garibay, D., Cummings, B. P. A Murine Model of Vertical Sleeve Gastrectomy. J. Vis. Exp. (2017)
In this video, a vertical sleeve gastrectomy ...

Vertical sleeve gastrectomy is a surgical procedure that involves removing a significant portion of the stomach along its greater curvature. To begin, prep an anesthetized mouse in the supine position.
Sterilize the abdominal skin with a suitable disinfectant. Incise the body wall from the mid-abdomen to the beginning of the breast bone to expose the stomach. Elevate the stomach from the abdominal cavity.
Dissect the greater omentum, or the layer of peritoneal tissue attached to the greater curvature of the stomach, to free up the organ.
Next, locate the short gastric artery extending from the spleen and supplying blood to the greater curvature of the stomach. Place two ligatures along the artery and cut between them to sever the vascular connection.
Identify the intended transection line. Locate the blood vessels extending towards the region of the stomach to be transected and ligate them just below the transection line to prevent blood loss.
Next, suture the opposing gastric walls below the transection line to seal them and prevent gastric content spillage. Now, clamp a hemostat above the sutured region to appropriately stage the transection line. Cut along the transection line to remove the unwanted gastric tissue.
Return the downsized stomach into the abdominal cavity. Suture the incisions and allow the mouse to recover.

murine-vertical-sleeve-gastrectomy-surgical-procedure-to-downsize

Research

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Animal model studies

1.1K 조회수. 출처: Garibay, D., Cummings, B. P. 수직 소매 위절제술의 쥐 모델. J. Vis. 특급 (2017) 이 비디오에서는 실험용 마우스 모델에서 수직 소매 위 절제술 절차가 수행됩니다. 이 절차는 더 큰 곡률을 따라 위의 상당 부분을 제거하는 데 도움이 됩니다. 

비디오: 쥐 수직 소매 위 절제술: 실험용 마우스 모델에서 위를 축소하는 수술 절차 - 개념

Roux-en-Y Gastric Bypass Operation in Rats

Currently, the most effective therapy for the treatment of morbid obesity to induce significant and maintained body weight loss with a proven mortality benefit is bariatric surgery1,2. Consequently, there has been a steady rise in the number of bariatric operations done worldwide in recent years with the Roux-en-Y gastric bypass (gastric bypass) being the most commonly performed operation3. Against this background, it is important to understand the physiological mechanisms by which gastric bypass induces and maintains body weight loss. These mechanisms are yet not fully understood, but may include reduced hunger and increased satiation4,5, increased energy expenditure6,7, altered preference for food high in fat and sugar8,9, altered salt and water handling of the kidney10 as well as alterations in gut microbiota11. Such changes seen after gastric bypass may at least partly stem from how the surgery alters the hormonal milieu because gastric bypass increases the postprandial release of peptide-YY (PYY) and glucagon-like-peptide-1 (GLP-1), hormones that are released by the gut in the presence of nutrients and that reduce eating12.
During the last two decades numerous studies using rats have been carried out to further investigate physiological changes after gastric bypass. The gastric bypass rat model has proven to be a valuable experimental tool not least as it closely mimics the time profile and magnitude of human weight loss, but also allows researchers to control and manipulate critical anatomic and physiologic factors including the use of appropriate controls. Consequently, there is a wide array of rat gastric bypass models available in the literature reviewed elsewhere in more detail 13-15. The description of the exact surgical technique of these models varies widely and differs e.g. in terms of pouch size, limb lengths, and the preservation of the vagal nerve. If reported, mortality rates seem to range from 0 to 35%15. Furthermore, surgery has been carried out almost exclusively in male rats of different strains and ages. Pre- and postoperative diets also varied significantly.
Technical and experimental variations in published gastric bypass rat models complicate the comparison and identification of potential physiological mechanisms involved in gastric bypass. There is no clear evidence that any of these models is superior, but there is an emerging need for standardization of the procedure to achieve consistent and comparable data. This article therefore aims to summarize and discuss technical and experimental details of our previously validated and published gastric bypass rat model.

Numerous studies using gastric bypass rat models have been recently conducted to uncover the underlying physiological mechanisms of Roux-en-Y gastric bypass operations. This article aims to demonstrate and discuss the technical and experimental details of our published gastric bypass rat model to understand advantages and limitations of this experimental tool.

쥐에서 Roux-EN-Y 위장 바이패스 수술

Currently, the most effective therapy for the treatment of morbid obesity to induce significant and maintained body weight loss with a proven mortality ...

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JoVE Journal

의학

Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood flow triggers a signaling cascade resulting in vasodilation thereby alleviating mechanical stress on the vascular wall. The pattern of shear stress is also well known to be a critical factor in the development of atherosclerosis with laminar shear stress being atheroprotective and disturbed shear stress being pro-atherogenic. While we have a detailed understanding of the various intermediate cell signaling pathways, the receptors that first translate the mechanical stimulus into chemical mediators are not completely understood. Mechanosensitive ion channels are critical to the response to shear and regulate shear-induced cell signaling thereby controlling the production of vasoactive mediators. These channels are among the earliest activated signaling components to shear and have been linked to shear-induced vasodilation through promoting nitric oxide production (e.g., inwardly rectifying K+ [Kir] and transient receptor potential [TRP] channels) and endothelium hyperpolarizing factor (e.g., Kir and calcium-activated K+ [KCa] channels) and shear-induced vasoconstriction through an undetermined mechanism that involves piezo channels. Understanding the biophysical mechanism by which these channels are activated by shear forces (i.e., directly or through a primary mechano-receptor) could provide potential new targets to resolve the pathophysiology associated with endothelial dysfunction and atherogenesis. It is still a major challenge to record flow-induced activation of ion channels in real time using electrophysiology. The standard methods to expose cells to well-defined shear stress, such as the cone and plate rheometer and closed parallel plate flow chamber do not allow real time study of ion channel activation. The goal of this protocol is to describe a modified parallel plate flow chamber that allows real time electrophysiological recording of mechanosensitive ion channels under well-defined shear stress.

The goal of this protocol is to describe a modified parallel plate flow chamber for use in investigating real time activation of mechanosensitive ion channels by shear stress.

잘 정의된 전단 응력 하에서 단세포 이온 전류의 전기 생리학적 기록

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood ...

면역학 및 감염병학

Sleeve Gastrectomy in Mice using Surgical Clips

The number of people who are overweight and obese is continually increasing both in the adult and adolescent populations. This coincides with the increased universal phenomenon of type 2 diabetes (T2D) and other metabolic problems. Bariatric surgery, such as SG, is currently one of the most effective and commonly used long-term treatment for obesity and T2D, but the association between them is not completely explored yet. The mechanisms underlying the outcomes seen after bariatric surgery in humans can be investigated based on preclinical animal studies. The SG reduces body weight, glucose levels and many metabolic parameters, and is easy to perform with a low incidence of complications. The goal of this work is to provide a simple method and an uncomplicated preclinical model of bariatric surgery in animals for researchers.

The prevalence of diabetes and obesity is continuously increasing worldwide. The mechanisms between diabetes, obesity and their associated mortality and co-morbidities need to be further investigated. Here, we present a protocol for sleeve gastrectomy (SG) in animals as an uncomplicated preclinical model of bariatric surgery.

Murine Vertical Sleeve Gastrectomy: A Surgical Procedure to Downsize Stomach in Experimental Mouse Models

내레이션 대본

Murine Vertical Sleeve Gastrectomy: A Surgical Procedure to Downsize Stomach in Experimental Mouse Models