Name: techniques of sleeve gastrectomy and modified roux-en-y gastric bypass in mice
Uploaded: 2017-03-20T12:30:00.000+00:00
Duration: 10 min 5 s
Description: Watch this Scientific Journal Video about Techniques of Sleeve Gastrectomy and Modified Roux-en-Y Gastric Bypass in Mice at JoVE.com

We thank Gilles Mithieux and Aude Barataud (INSERM U1213, Lyon, France) and Marie Liabeuf and Stephanie Lemarchand-Minde (Animal facility, l'Institut du Thorax, Nantes, France) for their help with the animal care protocol. This work was supported by grants from La r&#233;gion des Pays de la Loire, the Fondation d'Avenir, and the Casden Bank. We would like to thank Catherine Postic, Fadila Benhamed and Michelle Ca&#252;zac from l'institut Cochin for their hospitality and their help during the filming process.

Animal and housing: Obtain 8-week-old C57Bl/6 male mice. At 10 weeks old, give the C57Bl/6 mice free access to water and a high-fat diet (DIO diet: 35% kcal from fat, 25.8% caseine, 1.30% mineral AIN, 1.30% vitamins, 1.70% dicalcium phosphate, 0.7% calcium carbonate, 2.10% citrate potassium, 0.026% choline bitartrate, 8.9% sucrose, 0.384% cystine, 6.5% cellulose, 31.7% lard, 3.3% soybean oil, and 16.29% maltodextrine) for 8 (SG) or 14 (RYGB) weeks prior to the bariatric surgeries. To promote weight gain, give the mice dedicated to the RYGB surgery (sham &#38; RYGB) water containing 20% fructose (w/v) in addition to the high-fat diet.
Ethics Statement: All protocol steps described below follow the guidelines of the Ethics Committee for Animal Experimentation of Pays de la Loire under the approval number 01953.01.
1. General Preoperative Preparation
<ol>
	<li>Fast the mice for 6 h before the surgery. Gel diet food was given 3 days before surgery and exclusively one day before surgery. Induce anesthesia in a chamber with 5% isoflurane (0.4 L/min) and O2 (dioxygen; 0.4 L/min). Perform a toe-pinch test to confirm that the anesthetization is effective. Administer analgesics (0.1 mg/kg buprenorphine), antibiotics (10 mg/kg marbofloxacine), and pro-kinetics (1 mg/kg metoclopramide) via subcutaneous injections.</li>
	<li>Allocate a specific work area for surgical procedures (distant from laboratory traffic and ventilation fans). Clean the surgical area with dedicated hard surface disinfectants. Place a heating pad on the working space area to maintain mouse homeothermy during the surgery. Apply a clean, absorbent underpad over the heating pad.</li>
	<li>Shave the abdomen from sternum to pelvis using an electric razor. Wrap the mouse abdomen in sterilized plastic wrap (Figures 1A and 2A) and place the mouse in supine position on the heating pad.</li>
	<li>Apply vet ointment on the eyes to prevent dryness while under anesthesia. Place the mouse in the nose cone and maintain anesthesia with 2% isoflurane (0.4 L/min) and O2 (0.4 L/min).</li>
	<li>Before beginning the surgery, put on a sterile examination gown, a disposable scrubs cap, a facemask, and sterile gloves. Open an autoclaved package containing a sterilized surgical instrument set, which is required for all steps of the surgery. Cut a window in the sterilized plastic wrap and disinfect the skin with 2 successive povidone-iodine solutions.</li>
</ol>
2. Sleeve Gastrectomy: Surgical Procedure
<ol>
	<li>Median laparotomy (Figure 1B)
	<ol>
		<li>Under a binocular microscope and with a scalpel, perform a midline incision from the sternum to the middle of the abdomen to open the abdominal cavity (do not damage the abdominal muscles). Protect the skin with a sterile compress soaked with 37 &#176;C saline solution.</li>
	</ol>
	</li>
	<li>Stomach exposure (Figure 1C)
	<ol>
		<li>Gently mobilize the stomach using moistened-cotton swabs. Free the stomach from its lateral close connective tissue attachments using moistened cotton swabs or micro scissors.</li>
		<li>Gently, externalize the stomach fully and place a hemostatic collagen compress behind the stomach. Suture the pylorus and esophagus vessels along the stomach greater curvature with 8.0 non-absorbable sutures to avoid future bleeding (Figure 1D and E). Note:&#160;Hemostat use can induce tissue damage, make sure to only use it on tissue that aim to be resected.</li>
	</ol>
	</li>
	<li>Resection of the cardiac region of stomach (fundus)
	<ol>
		<li>Perform a gastrotomy on the anatomical line present between the pyloric region (corpus) and the cardiac region of stomach (fundus) (Figure 1F).</li>
		<li>Remove residual food with 2 moistened cotton swabs towards each side of the stomach. Using micro scissors, cut the stomach along the boundary between the fundus and the corpus of the stomach (Figure 1G).</li>
		<li>Press a hemostatic collagen compress for 2 min in case of major bleeding. Close the stomach opening with 8.0 non-absorbable sutures from the gastro-oesophagal junction to the end of the incision (Figure 1H).</li>
	</ol>
	</li>
	<li>Resection of the pyloric region of stomach (corpus)
	<ol>
		<li>Complete the stomach resection along the greater curvature so as to remove approximately 80% of the stomach (Figure 1I). Standardize the width of the SG to 3 mm. Press a hemostatic collagen compress for 2 min in case of major bleeding.</li>
		<li>Close the opening of the pyloric region of the stomach with 8.0 non-absorbable suture (Figure 1J). Gently roll 2 moistened cotton swabs towards each side of the stomach to ensure that the suture is leak-proof.</li>
		<li>Complete with 8.0 non-absorbable sutures in case of leakage. Remove the hemostatic collagen compresses and return the stomach to the abdominal cavity. Leave a hemostatic collagen compress against the suture.</li>
	</ol>
	</li>
	<li>Abdominal closure
	<ol>
		<li>Close the muscle layer of the abdominal wall with 5.0 non-absorbable sutures (Figure 1K). Reduce anesthesia by reducing the isoflurane concentration to 1%. Close the skin using 5.0 non-absorbable sutures (Figure 1L).</li>
	</ol>
	</li>
</ol>
3. Roux-en-Y Gastric Bypass: Surgical Procedure
<ol>
	<li>Median laparotomy
	<ol>
		<li>Under a binocular microscope and with a scalpel, perform a midline incision from the sternum to the middle of the abdomen to open the abdominal cavity (do not damage the abdominal muscles). Protect the skin with a sterile compress soaked with 37 &#176;C saline solution (Figure 2B).</li>
	</ol>
	</li>
	<li>Biliopancreatic limb and alimentary limb
	<ol>
		<li>Externalize the intestine (Figure 2C). Measure 8 cm from the pylorus and perform two ligatures of the intestine with 5.0 non-absorbable sutures (Figure 2D).</li>
		<li>Cut the intestine between the two ligatures (Figure 2E). Place the proximal limb of the two ends in the upper-left quadrant of the abdomen. Note: This will be used as the alimentary limb (Figure 2F).</li>
		<li>Place the distal limb of the two ends facing the alimentary limb 6 cm below the proximal limb (Figure 2F). 
		NOTE: This will be used as the biliary limb.</li>
	</ol>
	</li>
	<li>Jejuno-jejunostomy
	<ol>
		<li>Cut both the proximal limb and the intestine loop using micro scissors and perform two antimesenteric incisions of the same length (Figure 2G and 2H).</li>
		<li>Perform side-to-side anastomosis with two 8.0 non-absorbable sutures. Perform the dorsal side anastomosis first (Figure 2I), followed by the ventral side anastomosis (Figure 2J).</li>
		<li>Gently roll 2 moistened cotton swabs towards each side of the anastomosis to ensure that the suture is leak-proof. Complete with 8.0 non-absorbable sutures in case of leakage.</li>
	</ol>
	</li>
	<li>Gastro-jejunostomy
	<ol>
		<li>Gently mobilize the stomach using moistened cotton swabs. Free the stomach from its lateral close connective tissue attachments using moistened cotton swabs or micro scissors.</li>
		<li>Gently, externalize the stomach fully, placing a re-absorbable hemostatic collagen compress behind the stomach. Perform a ligature of the pylorus using a 5.0 non-absorbable suture passed through the omentum using curved micro forceps (Figure 2K and 2L).</li>
		<li>Cut both the ventral side of the stomach, 1.5 cm from the pylorus, and the distal limb using micro scissors, creating two incisions of the same length (Figure 2M).</li>
		<li>Perform side-to-side anastomosis with two 8.0 non-absorbable sutures. Start with the dorsal side anastomosis (Figure 2N), and then perform the ventral side anastomosis (Figure 2O). Gently roll 2 moistened-cotton swabs towards each side of the anastomosis to ensure that the suture is leak-proof. Complete with 8.0 non-absorbable sutures in case of leakage.</li>
	</ol>
	</li>
	<li>Abdominal closure
	<ol>
		<li>Close the muscle layer of the abdominal wall using 5.0 non-absorbable sutures (Figure 2P). Reduce the anesthesia by reducing the isoflurane concentration to 1%. Close the skin using 5.0 non-absorbable sutures (Figure 2Q).Administer 25 mL/kg of warm saline solution via subcutaneous injection.</li>
	</ol>
	</li>
</ol>
4. General Postoperative Care
<ol>
	<li>Stop isoflurane and continue with an O2 flow of 0.8 L/min until the mouse is fully awake. Do not leave the mouse unattended until it regains motor control, indicated by the animal starting to move around the cage and being able to stand and walk without falling.</li>
	<li>Place the mouse (only one mouse per cage) in an incubator under 30 &#176;C temperature condition for 5 days.</li>
	<li>Maintain iron (0.5 mg/kg/day; subcutaneous injection) and vitamin (800 mg/180 mL in water) supplementations in the RYGB mice until the end of the protocol.</li>
	<li>Return free access to gel diet food (high-fat gel diet: 10% lard, 10% liquid sugar, 57% water) for 5 days after surgery. Reintroduce a solid diet 3 days after surgery.</li>
	<li>Subcutaneously inject buprenorphine (0.1 mg/kg, twice daily, from day 0 to day 3 after surgery), meloxicam (1 mg/kg, from day 0 to day 3 after surgery), metoclopramide (1 mg/kg, from day 0 to day 5 after surgery), and marbofloxacine (10 mg/kg, from day 0 to day 3 after surgery).</li>
</ol>
5. Postoperative Metabolic Parameters Assessment
<ol>
	<li>Food intake measurement
	<ol>
		<li>Allow the mice to recover for 1 week after surgery. House the mice one per cage. Measure the amount of solid diet placed in the cage. Weight the remaining food 24 h later. Repeat this step as long as required.</li>
	</ol>
	</li>
	<li>Oral glucose tolerance test
	<ol>
		<li>Fast the mice for 6 h before the oral bolus. Harvest a blood drop from the tail tip and apply it on a glucose strip inserted in a glucometer to determine the time 0 blood glucose value.</li>
		<li>Administer an oral bolus of 20% D-glucose solution (2 g/kg). Measure the blood glucose levels at 15, 30, 60, and 120 min post gavage.</li>
	</ol>
	</li>
	<li>Hemoglobin level analysis
	<ol>
		<li>Using a hematology system, measure 20 &#181;L of fresh blood to sample the hemoglobin level.</li>
	</ol>
	</li>
</ol>
6. Euthanasia
<ol>
	<li>Anesthetize the mice by intraperitoneally injecting a xylazine/ketamine solution (10/80 mg/kg). Perform a cervical dislocation. Verify complete euthanasia by ensuring that the heart is not beating and the eyeball blink reflex is lost.</li>
</ol>

<ol>
	<li>Ng, M., Fleming, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global,+regional,+and+national+prevalence+of+overweight+and+obesity+in+children+and+adults+during+1980-2013:+a+systematic+analysis+for+the+Global+Burden+of+Disease+Study+2013.">Global, regional, and national prevalence of overweight and obesity in children and adults during 1980-2013: a systematic analysis for the Global Burden of Disease Study 2013.</a> The Lancet. 384 (9945), 766-781 (2014).</li><li>Calle, E. E., Thun, M. J., Petrelli, J. M., Rodriguez, C., Heath, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Body-mass+index+and+mortality+in+a+prospective+cohort+of+U.S.+adults.">Body-mass index and mortality in a prospective cohort of U.S. adults.</a> The New England Journal of Medicine. 341 (15), 1097-1105 (1999).</li><li>Sj&#246;str&#246;m, L., Lindroos, A. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lifestyle,+diabetes,+and+cardiovascular+risk+factors+10+years+after+bariatric+surgery.">Lifestyle, diabetes, and cardiovascular risk factors 10 years after bariatric surgery.</a> The New England Journal of Medicine. 351 (26), 2683-2693 (2004).</li><li>Buchwald, H., Avidor, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bariatric+surgery:+a+systematic+review+and+meta-analysis.">Bariatric surgery: a systematic review and meta-analysis.</a> JAMA. 292 (14), 1724-1737 (2004).</li><li>Yin, D. P., Gao, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+different+bariatric+surgeries+in+the+treatment+of+obesity+and+insulin+resistance+in+mice.">Assessment of different bariatric surgeries in the treatment of obesity and insulin resistance in mice.</a> Annals of surgery. 254 (1), 73-82 (2011).</li><li>Buchwald, H., Oien, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabolic/Bariatric+Surgery+Worldwide+2008.">Metabolic/Bariatric Surgery Worldwide 2008.</a> Obesity Surgery. 19 (12), 1605-1611 (2009).</li><li>Liu, W., Zassoko, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+duodenojejunal+bypass+surgery+in+mice:+A+model+designed+for+diabetic+research.">Establishment of duodenojejunal bypass surgery in mice: A model designed for diabetic research.</a> Microsurgery. 28 (3), 197-202 (2008).</li><li>Lan, Z., Zassoko, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+techniques+for+gastrojejunal+bypass+surgery+in+obese+mice.">Development of techniques for gastrojejunal bypass surgery in obese mice.</a> Microsurgery. , (2010).</li><li>Schlager, A., Khalaileh, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+mouse+model+for+sleeve+gastrectomy:+Applications+for+diabetes+research.">A mouse model for sleeve gastrectomy: Applications for diabetes research.</a> Microsurgery. 31 (1), 66-71 (2011).</li><li>Troy, S., Soty, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intestinal+Gluconeogenesis+Is+a+Key+Factor+for+Early+Metabolic+Changes+after+Gastric+Bypass+but+Not+after+Gastric+Lap-Band+in+Mice.">Intestinal Gluconeogenesis Is a Key Factor for Early Metabolic Changes after Gastric Bypass but Not after Gastric Lap-Band in Mice.</a> Cell Metabolism. 8 (3), 201-211 (2008).</li><li>Seyfried, F., Lannoo, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Roux-en-Y+gastric+bypass+in+mice--surgical+technique+and+characterisation.">Roux-en-Y gastric bypass in mice--surgical technique and characterisation.</a> Obesity surgery. 22 (7), 1117-1125 (2012).</li></ol>

The authors declare no conflicts of interest.

To overcome the growing epidemic of obesity, the first bariatric surgery procedures emerged in the 1960s in the United States. Since then, the number of procedures performed worldwide each year still increase, and today, these techniques represent the best therapeutic option for the management of morbid obesity6. Among the procedures developed, SG and RYGB are the two most popular methods used in clinical practice4. Animal models, notably rodents, have been used to decipher the mechanisms behind the general health improvement observed after bariatric surgery.
To decipher the molecular mechanisms sustaining the beneficial metabolic effects of bariatric surgeries, we developed two mouse models of bariatric surgery: the SG, which is often defined as a restrictive technique, and the RYGB, which is described as a restrictive and malabsorptive surgery. The realization of these mouse bariatric procedures requires experience in microsurgery. These models were developed using a technician specialized in animal surgery and a human visceral surgeon. From experience, surgical training was necessary to reach over 75% survival after RYGB. Mastering SG does not require as much time, and the learning curve is shorter, with mortality at about 95%. By contrast, RYGB is a more complex technique and requires 6 to 8 weeks of training to reach an acceptable postoperative survival rate.
In the literature, there are a variety of rodent bariatric procedures that have been previously described7,8,9. In this SG technique, we removed the cardiac region of the stomach (also called the fundus), not present in humans. The objective was to remove about 80% of the stomach. The pouch size of RYGB models in rats ranges from &#60;5% to &#62;20% of the initial stomach size. Several studies used mouse RYGB models, presenting either a complete preservation of the stomach10, a small gastric pouch5, or a total stomach diversion5. In a very comprehensive study, Deng Ping Yin and colleagues have evaluated the different mouse models of RYGB and reported that the murine stomach was not anatomically adapted to the creation of a gastric pouch5. For our RYGB procedure, we choose to perform a Roux-en-Y without cutting the stomach. As a limitation, our model did not induce a reduction in food intake, which contrasts what it observed in human models of RYGB.
The survival rate after RYGB operations in rats is rarely mentioned by the authors in the literature, but it seems to reach about 30 to 35% on average11. In our hands, mortality was mainly due to anastomotic leakage or stenosis or to late postoperative gastrojejunal anastomosis stenosis, leading to chronic obstruction with persistent excessive weight loss, which compromised animal welfare. RYGB requires specific perioperative management, including a gel diet during the first three postoperative days and a pain management regime. To improve long-term survival after surgery, we added vitamins postoperatively, particularly fat-soluble vitamins and iron, to prevent anemia. Conversely, mice do not need liposoluble vitamin supplementation after SG, since it was only a restrictive procedure. In the previous papers, the authors did not describe the health condition of operated animals during the postoperative course. Because it is a major surgery, we cannot assume the systematic well-being of the animals in these studies.
In conclusion, mouse models of either restrictive (SG) or restrictive-malabsorptive (RYGB) bariatric surgeries are feasible but require microsurgical training with a learning curve, especially for the RYGB procedure. To keep the animals in good health, perioperative management is very important in the RYGB procedure, particularly in terms of pain, but also in terms of vitamin supplementation. By achieving a faster, sterile surgery and by using microsurgical training, we can greatly improve the postoperative survival rate. These models are useful and essential tools to understand the metabolic effects of bariatric surgery.

The worldwide pandemic of obesity and diabetes is devastating in severity. Over two billion adults worldwide (30% of the population) are either overweight (BMI &#62;25 kg/m2) or obese (BMI &#62;30 kg/m2)1. This can come along with metabolic complications, such as type 2 diabetes, hypertension, and dyslipidemia, leading to increased morbidity and mortality. Obesity increases the overall mortality and the prevalence of cancer2. Due to the lack of any effective noninvasive treatments, bariatric surgery represents the only option that can lead to long-term, sustained weight loss3,4. A number of different surgical methods have been developed, but the sleeve gastrectomy (SG) and Roux-en-Y gastric bypass (RYGB) are the two procedures most commonly used in clinical practice. During the SG procedure, 80% of the initial volume of the stomach is removed; thus, this technique is one of the restrictive surgeries that improves satiety. The RYGB is one of the restrictive-malabsorptive techniques. During RYGB, a small gastric pouch (1-2% of the total gastric volume) is created and the intestine is rearranged into a Y-shape, which delays digestion and the absorption of nutrients. These two techniques lead to significant body weight reductions and general improvements in frequently-associated comorbidities (e.g., hypertension, type 2 diabetes, and dyslipidemia)3, with a higher efficiency seen in RYGB. However, the molecular mechanisms behind the pleiotropic beneficial effects of the bariatric surgeries are often not fully elucidated. Due to the numerous genetically-modified strains available, a mouse model is the most convenient animal model to explore these molecular mechanisms.
However, bariatric procedures are difficult to directly adapt to small animal models and require high surgical dexterity. While SG can be easily performed in rodents with a very good survival rate, RYGB is lethal in mice due to severe bowel obstructions5. Different modified RYGB techniques have been proposed to counteract this problem, notably the oesojejunostomy5. Here, we present another alternative: the gastrojejunostomy without a stomach excision. This modified RYGB reproduces most of the beneficial effects observed in humans (i.e., a significant body weight reduction and an improvement in glucose and lipid homeostasis).
This manuscript aims to summarize and discuss the technical and experimental details of SG and RYGB in mice and to facilitate these procedures with the help of videos. A specific highlight will be made regarding the optimization of the preoperative and postoperative healthcare protocols that allow the reduction of vitamin and iron deficiencies.

<table><tbody><tr><td>&lt;strong&gt;Drugs&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>High Fat diet</td><td>DIO diet</td><td>Safe</td><td></td></tr><tr><td>Isoflurane</td><td>Forane</td><td>Baxter</td><td></td></tr><tr><td>Buprenorphin</td><td>Buprecare</td><td>Animalcare</td><td></td></tr><tr><td>Marbofloxacine</td><td>Marbocyl</td><td>Vetoquinol</td><td></td></tr><tr><td>Ammonium iron citrate, vitamins PP-B12</td><td>Fercobsang</td><td>Vetoquinol</td><td></td></tr><tr><td>Vitamins A-D3-E-K-B</td><td>Vita Rongeur</td><td>Virbac</td><td></td></tr><tr><td>NaCl 0.9%</td><td>NaCl 0,9%</td><td></td><td></td></tr><tr><td>Povidone solution</td><td>Betadine Scrub</td><td>Betadine</td><td></td></tr><tr><td>Povidone solution</td><td>Betadine Solution</td><td>Betadine</td><td></td></tr><tr><td>Carboptol 980 NF</td><td>Ocrygel</td><td>TVM</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;References&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Sutures&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Prolene&amp;reg;</td><td>8.0, 6,5 mm</td><td>Ethicon</td><td></td></tr><tr><td>Prolene&amp;reg;</td><td>5.0, 13 mm</td><td>Ethicon</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;References&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Surgical equipments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Scissors</td><td></td><td>FST</td><td></td></tr><tr><td>Needle holder</td><td>Olsen-Hegar</td><td>FST</td><td></td></tr><tr><td>Micro scissors</td><td>Vannas</td><td>FST</td><td></td></tr><tr><td>Micro forceps&amp;nbsp;</td><td>Graefe</td><td>FST</td><td></td></tr><tr><td>Micro forceps curved</td><td>Graefe</td><td>FST</td><td></td></tr><tr><td>Curved micro needle holder</td><td>Castroviejo</td><td>FST</td><td></td></tr><tr><td>Hemostatic collagen compress</td><td>Pangen</td><td>Urgo</td><td></td></tr><tr><td>Absorbent underpads</td><td></td><td>VWR</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;References&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Specific equipments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Hematology system&amp;nbsp;</td><td>Hemavet 950FS</td><td>Hemavet</td><td></td></tr><tr><td>Glucose strips and glucometer</td><td>One touch Verio</td><td>Life scan</td><td></td></tr><tr><td>Stereo microscope</td><td>MZ6</td><td>Leica&amp;nbsp;</td><td></td></tr></tbody></table>

techniques of sleeve gastrectomy and modified roux-en-y gastric bypass in mice

Obesity is a major public health issue, with a prevalence of 4 to 28% for men and 6.2 to 36.5% for women in Europe (from 2003 to 2008). Morbid obesity is frequently associated with metabolic complications, such as type 2 diabetes, hypertension, and dyslipidemia, reducing life expectancy and quality. In the absence of any effective noninvasive treatments, bariatric surgery is a valuable therapeutic option for patients with morbid obesity (body mass index (BMI) &#62;40 kg/m2), leading to long-term, sustained weight loss and improvements in metabolic complications. However, the underlying cellular and molecular mechanisms sustaining the beneficial effects of bariatric surgery are not yet fully understood. Due to the numerous genetically-modified strains available, the mouse model is the most convenient animal model to explore the molecular mechanisms behind the pleiotropic beneficial effects of bariatric surgeries. Here, we detailed the optimized healthcare methods and surgical protocols in mice for the two most widely-used bariatric surgeries: the sleeve gastrectomy and the modified Roux-en-Y gastric bypass. Deciphering the molecular mechanisms underlying the therapeutic effects of bariatric surgeries offers the promise of identifying new therapeutics targets.

General conditions
The mean operative time for the SG procedure was 49.3 &#177; 1.5 min. We removed 62.8 &#177; 5.0 mg of stomach, which represents about 80% percent of the stomach. No mice died during the surgery or during the following seven days. One mouse (7.1%) died on the 11th postoperative day because of a gastric obstruction caused by a bezoar.
The mean operative time for the RYGB procedure was 89.1 &#177; 2.8 min. The mortality rate was significantly higher after RYGB (sham: 25% versus RYGB: 66.6%; p = 0.038). The main causes of postoperative deaths were anastomotic leaks (n = 8), anastomotic stenosis (n = 5), anesthesia complications (n = 4), and postoperative bleeding (n = 2). We also observed two long-term deaths due to gastrointestinal anastomosis stenosis with chronic obstruction and malnutrition. As shown in Figure 3, we noticed a progressive improvement of the post-operative survival rate throughout the protocol, indicating that intensive surgical training is required to master the RYGB surgical technique.
Body weight, food intake, and glucose homeostasis
The mice had similar body weights before surgery. SG demonstrated significant weight loss 14 days after surgery compared to sham control mice (sham: -6.6% versus SG: -16.0%; p &#60;0.01; Figure 4A). RYGB was associated with significant weight loss from postoperative days 14 (sham: -7.9% on day 14 post-surgery and -5.1% on day 28 post-surgery versus RYGB: -22.9% on day 14 post-surgery and -25.5% on day 28 post-surgery; Figure 4B). The daily food intake was reduced by 35% during the 14-day period following the SG (Figure 4C). In contrast, we did not observe such an anorexigenic effect between the sham and RYGB mice (Figure 4D). We also reported an improvement of glucose tolerance following SG and RYGB surgeries (Figure 4E, 4F).
Beneficial effect of iron supplementation on RYGB-induced anemia
We did not observe a significant decrease of hemoglobinemia 14 days after SG surgery (data not shown). In contrast, severe anemia was measured in RYGB-operated mice, but it did benefit from specific iron supplementation (Figure 5A). Daily iron supplementation was indeed crucial and prevented significant anemia 28 days after RYGB surgery (Figure 5B).
<img alt="Figure 1" src="/files/ftp_upload/54905/54905fig1.jpg" /> 
Figure 1: Sleeve Gastrectomy Procedure. (A) Preoperative preparation. (B) Median laparotomy. (C) Stomach exposure. (D) Pylorus vessel suture. (E) Esophagus vessel suture. (F) Initial gastrotomy. (G) Resection of the cardiac region of the stomach. (H) Closure of the stomach opening from the gastroesophageal junction to the end of the incision. (I) Resection of the pyloric region of the stomach. (J) Closure of the opening of the pyloric region. (K) Closure of the muscle layer of the abdominal wall. (L) Closure of the skin. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54905/54905fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54905/54905fig2.jpg" /> 
Figure 2: Roux-en-Y bypass Procedure. (A) Preoperative preparation. (B) Median laparotomy. (C) Externalization of the small intestine. (D) Biliopancreatic and alimentary limbs ligatures. (E) Small intestine section between the ligature. (F) Intestinal limb positioning for the jejuno-jejunostomy. (G-H) Antimesenteric incisions of the proximal limb and the intestine loop. (I) Dorsal side-to-side anastomosis. (J) Ventral side-to-side anastomosis. (K-L) Pylorus ligature. (M) Incision of the ventral side of the stomach. (N) Incision of the distal limb and dorsal side-to-side anastomosis. (O) Ventral side-to-side anastomosis. (P) Closure of the muscle layer of the abdominal wall. (Q) Closure of the skin. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54905/54905fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54905/54905fig3.jpg" /> 
Figure 3: Survival Rate of Mice 7 Days after the RYGB Surgery. The data show the benefit of surgical training on the survival rate of RYGB-operated mice. The values are shown as the mean value &#177; SEM. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54905/54905fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54905/54905fig4.jpg" /> 
Figure 4: Effect of SG (A, C, D) and RYGB (B, D, F) on Bodyweight, Daily Food Intake, and Glucose Tolerance. The data are shown as the mean value &#177; SEM. NS: not significant; * P &#60;0.05; ** P &#60;0.01 (Mann-Whitney test, ANOVA test). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54905/54905fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54905/54905fig5.jpg" /> 
Figure 5: Plasma Hemoglobin Levels after RYGB in Mice Without (A) or With (B) Iron Supplementation. The data are shown as the mean value &#177; SEM. NS: not significant; * P &#60;0.05 (Mann-Whitney test). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54905/54905fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Techniques of Sleeve Gastrectomy and Modified Roux-en-Y Gastric Bypass in Mice at JoVE.com

Techniques of Sleeve Gastrectomy and Modified Roux-en-Y Gastric Bypass in Mice

Bariatric surgery is the most efficient way to reduce body weight and the deadly metabolic complications (diabetes, obesity, and dyslipidemia) frequently associated with morbid obesity. Mouse models of bariatric surgery represent a unique asset for deciphering molecular mechanisms behind the beneficial effects of these surgeries on diabetes, hypertension, and dyslipidemia.

Obesity is a major public health issue, with a prevalence of 4 to 28% for men and 6.2 to 36.5% for women in Europe (from 2003 to 2008). Morbid obesity is ...

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11.9K Views.  INSERM, CNRS, UNIV Nantes. Bariatric surgery is the most efficient way to reduce body weight and the deadly metabolic complications (diabetes, obesity, and dyslipidemia) frequently associated with morbid obesity. Mouse models of bariatric surgery represent a unique asset for deciphering molecular mechanisms behind the beneficial effects of these surgeries on diabetes, hypertension, and dyslipidemia.

Video: Techniques of Sleeve Gastrectomy and Modified Roux-en-Y Gastric Bypass in Mice

Modified Technique for Coronary Artery Ligation in Mice

Myocardial infarction (MI) is one of the most important causes of mortality in humans1-3. In order to improve morbidity and mortality in patients with MI we need better knowledge about pathophysiology of myocardial ischemia. This knowledge may be valuable to define new therapeutic targets for innovative cardiovascular therapies4. Experimental MI model in mice is an increasingly popular small-animal model in preclinical research in which MI is induced by means of permanent or temporary ligation of left coronary artery (LCA)5. In this video, we describe the step-by-step method of how to induce experimental MI in mice.
The animal is first anesthetized with 2% isoflurane. The unconscious mouse is then intubated and connected to a ventilator for artificial ventilation. The left chest is shaved and 1.5 cm incision along mid-axillary line is made in the skin. The left pectoralis major muscle is bluntly dissociated until the ribs are exposed. The muscle layers are pulled aside and fixed with an eyelid-retractor. After these preparations, left thoracotomy is performed between the third and fourth ribs in order to visualize the anterior surface of the heart and left lung. The proximal segment of LCA artery is then ligated with a 7-0 ethilon suture which typically induces an infarct size ~40% of left ventricle. At the end, the chest is closed and the animals receive postoperative analgesia (Temgesic, 0.3 mg/50 ml, ip). The animals are kept in a warm cage until spontaneous recovery.

The surgical procedure used to induce experimental myocardial infarction in mice begins with left thoracotomy between the third and the fourth ribs in order to visualize the anterior surface of the heart and left lung. The left coronary artery is ligated, the chest is closed and the mouse is allowed to recover spontaneously.

Myocardial infarction (MI) is one of the most important causes of mortality in humans1-3. In order to improve morbidity and mortality in patients with MI ...

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.

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

Assessment of Gastric Emptying in Non-obese Diabetic Mice Using a [13C]-octanoic Acid Breath Test

Gastric emptying studies in mice have been limited by the inability to follow gastric emptying changes in the same animal since the most commonly used techniques require killing of the animals and postmortem recovery of the meal1,2. This approach prevents longitudinal studies to determine changes in gastric emptying with age and progression of disease. The commonly used [13C]-octanoic acid breath test for humans3 has been modified for use in mice4-6 and rats7 and we previously showed that this test is reliable and responsive to changes in gastric emptying in response to drugs and during diabetic disease progression8. In this video presentation the principle and practical implementation of this modified test is explained. As in the previous study, NOD LtJ mice are used, a model of type 1 diabetes9. A proportion of these mice develop the symptoms of gastroparesis, a complication of diabetes characterized by delayed gastric emptying without mechanical obstruction of the stomach10.
This paper demonstrates how to train the mice for testing, how to prepare the test meal and obtain 4 hr gastric emptying data and how to analyze the obtained data. The carbon isotope analyzer used in the present study is suitable for the automatic sampling of the air samples from up to 12 mice at the same time. This technique allows the longitudinal follow-up of gastric emptying from larger groups of mice with diabetes or other long-standing diseases.

Assessment of Gastric Emptying in Non-obese Diabetic Mice Using a [13C]-octanoic Acid Breath Test

Determination of gastric emptying with a non-invasive [13C]-octanoic acid breath test for tracking gastroparesis in female NOD LtJ mice.

Gastric emptying studies in mice have been limited by the inability to follow gastric emptying changes in the same animal since the most commonly used ...

Using the Sleeve Technique in a Mouse Model of Aortic Transplantation - An Instructional Video

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the aorta in mice using the sleeve method was short and easy averaging 20 min, permitting studies of iso/allo grafts. The following article describes the aortic transplantation procedure used in our laboratory. The mice were anesthetized with a mixture of 1.5% volume isoflurane and 100% oxygen through a face mask. At this point, the segment of the aorta between the renal arteries and its bifurcation was separated from the vena cava, freely prepared and clampedat the proximal and distal segments with a single silk suture. Prior to the removal of the aorta, a saline solution containing heparin was injected into the inferior vena cava. Then the aorta was cut between the clamps and a saline heparin solution was used to flush the lumen. The sleeve technique with monofilament sutures was used in order to transplant the abdominal aorta in the orthotopic position.

We present an orthotopic aortic transplantation model using the sleeve technique in mice. It is a very rapid anastomosis method, which can be employed in studies of vascular disease.

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the ...

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.

The number of people who are overweight and obese is continually increasing both in the adult and adolescent populations. This coincides with the ...

A Protocol for Roux-en-Y Gastric Bypass in Rats using Linear Staplers

Roux-en-Y gastric bypass (RYGB) is commonly performed for the treatment of severe obesity and type 2 diabetes. However, the mechanism of weight loss and metabolic changes are not well understood. Multiple factors are thought to play a role, including reduced caloric intake, decreased nutrient absorption, increased satiety, the release of satiety-promoting hormones, shifts in bile acid metabolism, and alterations in the gut microbiota.
The rat RYGB model presents an ideal framework to study these mechanisms. Prior work on mouse models have had high mortality rates, ranging from 17 to 52%, limiting their adoption. Rat models demonstrate more physiologic reserve to surgical stimulus and are technically easier to adopt as they allow for the use of surgical staplers. One challenge with surgical staplers, however, is that they often leave a large gastric pouch which is not representative of RYGB in humans. 
In this protocol, we present a RYGB protocol in rats that result in a small gastric pouch using surgical staplers. Utilizing two stapler fires which remove the forestomach of the rat, we obtain a smaller gastric pouch similar to that following a typical human RYGB. Surgical stapling also results in better hemostasis than sharp division. Additionally, the forestomach of the rat does not contain any glands and its removal should not alter the physiology of RYGB.
Weight loss and metabolic changes in the RYGB cohort were significant compared to the sham cohort, with significantly lower glucose tolerance at 14 weeks. Furthermore, this protocol has an excellent survival of 88.9% after RYGB. The skills described in this protocol can be acquired without previous microsurgical experience. Once mastered, this procedure will provide a reproducible tool for studying the mechanisms and effects of RYGB.

Roux-en-Y gastric bypass (RYGB) is performed to treat obesity and diabetes. However, the mechanisms underlying RYGB's efficacy are not fully understood, and studies are limited by technical difficulty leading to high mortality in animal models. This article provides instructions on how to perform RYGB in rats with high success rates.

Roux-en-Y gastric bypass (RYGB) is commonly performed for the treatment of severe obesity and type 2 diabetes. However, the mechanism of weight loss and ...

Single-Anastomosis Duodeno-Ileal Bypass with Sleeve Gastrectomy Model in Mice

Obesity is a major health issue worldwide. As a response, bariatric surgeries have emerged to treat obesity and its related comorbidities (e.g., diabetes mellitus, dyslipidemia, non-alcoholic steatohepatitis, cardiovascular events, and cancers) through restrictive and malabsorptive mechanisms. Understanding the mechanisms by which these procedures allow such improvements often require their transposition into animals, especially in mice, because of the ease of generating genetically modified animals. Recently, the single-anastomosis duodeno-ileal bypass with sleeve gastrectomy (SADI-S) has emerged as a procedure that uses both restrictive and malabsorptive effects, which is being used as an alternative to gastric bypass in case of major obesity. Thus far, this procedure has been associated with strong metabolic improvements, which has led to a marked increase in its use in daily clinical practice. However, the mechanisms underlying these metabolic effects have been poorly studied as a result of a lack of animal models. In this article, we present a reliable and reproducible model of SADI-S in mice, with a special focus on perioperative management. The description and use of this new rodent model will be helpful for the scientific community to better understand the molecular, metabolic, and structural changes induced by the SADI-S and to better define the surgical indications for clinical practice.

Single-anastomosis duodeno-ileal bypass (SADI-S) is an emerging bariatric procedure with important metabolic effects. In this article, we present a reliable and reproducible model of SADI-S in mice.

Obesity is a major health issue worldwide. As a response, bariatric surgeries have emerged to treat obesity and its related comorbidities (e.g., diabetes ...

A Murine Model of Vertical Sleeve Gastrectomy

Bariatric surgery, such as vertical sleeve gastrectomy (VSG), is a surgery of the gastrointestinal tract that is performed for the purpose of weight loss. Bariatric surgery is currently the most effective long-term treatment for obesity. In addition to weight loss, bariatric surgery produces additional health benefits such as remission of type 2 diabetes, remission of hypertension, and decreased risk of developing certain types of cancer. The mechanisms beyond weight loss for these benefits remain incompletely defined. Therefore, animal models of bariatric surgery are being developed and validated to identify the mechanisms leading to these benefits, with the goal of improving understanding of gastrointestinal physiology and identifying new therapeutic targets. VSG has become the most commonly performed bariatric procedure in the clinic in the United States because it is highly effective at producing weight loss and metabolic improvement, and is simpler to perform than other bariatric procedures. Therefore, we have developed and validated a murine model of VSG. This murine VSG model recapitulates many of the effects of VSG seen in humans, including improved glucose and blood pressure regulation. The method is based on isolation of the stomach, ligation of gastric vessels, and removal of 70% of the stomach by transecting along the greater curvature of the stomach. We have successfully applied this surgical protocol to various genetically modified mouse lines to define the mechanistic contributors to the benefits of VSG. Furthermore, this murine VSG model has been combined with other surgical techniques, to achieve deeper mechanistic insight. Therefore, this is a simple and versatile model for studying gastrointestinal physiology and the health benefits of bariatric surgery.

The following describes the performance of vertical sleeve gastrectomy in mice. This is a type of weight-loss surgery that involves removal of approximately 70% of the stomach.

Bariatric surgery, such as vertical sleeve gastrectomy (VSG), is a surgery of the gastrointestinal tract that is performed for the purpose of weight loss. ...

Biology

Roux-en-Y Gastric Bypass Procedure: A Bariatric Surgical Procedure for Gastric Resectioning in Murine Model

Source: Ayer, A., et al., <a href="https://www-jove-com.bdigitaluss.remotexs.co/v/54905">Techniques of Sleeve Gastrectomy and Modified Roux-en-Y Gastric Bypass in Mice.</a> J. Vis. Exp. (2017).
This video describes a bariatric surgery-Roux-en-Y procedure. This involves the cutting of the jejunum- the middle portion of the intestine- to generate two intestinal limbs. The biliary limb is attached to the existing intestinal loop and the alimentary limb is attached to the stomach, creating a gastric bypass. The gastric bypass created here helps to decrease the nutrient uptake and efficiently reduces body weight.

Source: Ayer, A., et al., Techniques of Sleeve Gastrectomy and Modified Roux-en-Y Gastric Bypass in Mice. J. Vis. Exp. (2017).
This video describes a ...

JoVE Encyclopedia of Experiments

Rodent Models

Adult

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

Before beginning the procedure, apply veterinary ointment to the eyes of an anesthetized mouse and place the mouse on a heating pad. Remove the hair from the umbilicus to the axilla, gently holding the skin taut to avoid skin lesions, and use cotton tip applicators to sequentially disinfect the exposed skin with povidone-iodine and ethanol.
Wearing sterile gloves and using sterile technique, place sterile surgical drapes on either side of the animal. Cut a small hole in the foil to allow access to the mouse&#39;s abdomen and use a piece of autoclaved aluminum foil as a surgical drape. Make a skin incision from the mid-abdomen to the xiphoid cartilage.
Under a dissecting microscope, use iris scissors to cut through the body wall along the linea alba and use new cotton tip applicators to gently elevate the stomach out of the abdomen. Bluntly dissect the greater omentum from the greater curvature of the stomach.
Then, use 7.0 monofilament absorbable sutures to place two ligatures around the short gastric artery that runs between the fundus of the stomach and the spleen. When both ligatures have been placed, use spring scissors to cut between the two sutures and fully exteriorize the stomach from the abdominal cavity.
Place gauze under the stomach and hydrate the organ with saline. For a vertical sleeve gastrectomy procedure, use a 7.0 monofilament absorbable suture with a taper needle to ligate the prominent branches of the gastric artery and vein, just below the intended line of transection. Place a simple continuous line of 6.0 monofilament absorbable suture through both gastric walls, just below the intended line of transection, to prevent spillage of the gastric contents.
Place thin-tipped hemostats above the suture line and use spring scissors to cut between the suture line and the hemostats. Remove the transected gastric tissue from the sterile surgical field and use fresh cotton tip applicators to clean the blood and digesta from the stomach.
Use a 6.0 monofilament absorbable suture with a taper needle to reinforce the apposition of the gastric walls in a simple discontinuous pattern. Using a 20G gavage needle attached to a 20-milliliter syringe, perform gastric lavage with fresh saline and securely close the stomach with a minimum of 20 knots.
Then, gently press the stomach with a cotton applicator to test for leaks. If no leaks are detected, lavage the stomach with at least 60 more milliliters of fresh saline to remove any infectious particulates.
After the procedure, use a cotton tip applicator to return the stomach under the liver within the abdominal cavity and place an applicator along the dorsal aspect of the abdominal cavity to absorb any excess fluid.
Use a blunt 18G needle to inject lactated Ringer&#39;s solution with or without antibiotics directly into the abdominal cavity, and use 6.0 monofilament absorbable sutures to close the abdominal muscle layer in a simple discontinuous pattern and the skin layer in a simple continuous pattern. Then, place tissue adhesive on the skin and fold the skin over the suture line to protect the suture from disruption by the animal.