<p class="jove_content">This study was funded by the American Society for Metabolic and Bariatric Surgery Research Award. Ethicon graciously supplied sutures, staplers, and clips. The lead author&#39;s doctoral research was funded in by the University of Alberta Clinician Investigator Program and the Alberta Innovates Clinician Fellowship. We would also like to thank Michelle Tran for her medical illustration of the RYGB anatomy.</p>

<p class="jove_content">Animal use protocols were approved by the Health Science Animal Care and Use Committee at the University of Alberta (AUP00003000). See <strong class="xfig">Figure 1</strong> for a diagram demonstrating the RYGB anatomy.</p>
<p class="jove_title">1. Roux-en-Y gastric bypass</p>
<ol>
	<li>Preparation of animals and operative setup
	<ol>
		<li>One week prior to the surgery, provide the rats with oral rehydration therapy and liquid diet in addition to their solid diet and water to acclimatize them to this new diet.</li>
		<li>Fast rats with only access to water for 12-18 h prior to the surgery.
		<ol>
			<li>Ensure rats are fasted on a raised wire platform so that they cannot consume bedding material.</li>
		</ol>
		</li>
		<li>Inject rats with subcutaneous long buprenorphine sustained release (SR) at a dose of 1 mg/kg immediately before surgery.</li>
		<li>Autoclave all surgical instruments, towels, and drapes.</li>
		<li>Clean the operating surface, heating pad, and anesthetic nose cone with 70% ethanol.</li>
		<li>Set up the operating surface with an operating microscope, anesthetic machine, and supplies in a manner that is ergonomic for the operating surgeon.</li>
		<li>Use a temperature-regulated heating pad and set to 37 &#176;C.</li>
		<li>Place a sterile drape or towel over the heating pad.</li>
		<li>Fill a 50 mL sterile conical tube with 0.9% saline.</li>
	</ol>
	</li>
	<li>Anesthetic induction and preparation
	<ol>
		<li>Induce anesthesia using 4% isoflurane as per previously established protocols<sup class="xref">15</sup>.</li>
		<li>Apply pressure to the hindfoot of all four limbs to ensure there is no pain response.</li>
		<li>Check for adequate anesthesia and respiratory rate after every 5 min.</li>
		<li>Apply lubricant to both eyes to prevent drying.</li>
		<li>Shave hair from the abdomen.</li>
		<li>Clean the abdomen with a povidone-iodine solution. Allow the solution to dry, and change into sterile gloves.</li>
		<li>Drape the rat with an opening in the drape to expose the abdomen.</li>
		<li>Place instruments, sutures, cotton swabs, and a 10 mL syringe in a location that permits easy access during the procedure.</li>
	</ol>
	</li>
	<li>Median laparotomy
	<ol>
		<li>Make a 3 cm incision in the upper midline of the abdomen using a scalpel, just below the xyphoid process as a landmark.</li>
		<li>Using scissors, divide the fascia and peritoneum, with care to stay midline on the linea alba to reduce bleeding from the rectus abdominus. If there is bleeding, control it with thermal or electrocautery.</li>
	</ol>
	</li>
	<li>Mobilizing the stomach
	<ol>
		<li>Using two wet cotton swabs, bluntly dissect gastric attachments.</li>
		<li>When encountering dense adhesions, use thermal cautery to divide gastric attachments with care to avoid cauterizing the stomach. Sharply divide the ligament between the stomach and the accessory liver lobe to reduce the risk of liver tearing with stomach mobilization.</li>
		<li>For larger blood vessels, especially at the short gastric arteries, ligate using 6-0 polypropylene suture.</li>
		<li>Create a window on the right distal side of the esophagus but proximal to the left gastric artery. Ensure that a cotton swab can reach into this area posteriorly. The stomach is adequately mobilized when it can be exteriorized outside of the abdomen.</li>
	</ol>
	</li>
	<li>Identify and divide the jejunum
	<ol>
		<li>Identify the ligament of Treitz by following the jejunum proximally until observing it is attachment to the transverse mesocolon.</li>
		<li>Measure 7 cm distally, identify a location between mesenteric vessels, and divide the bowel with micro scissors. Avoid Peyer&#39;s patches when dividing the bowel. Take care to only divide the bowel and not the mesentery.</li>
		<li>Place a clean, saline soaked sponge prior to dividing the bowel to minimize contamination.</li>
		<li>Check for the presence of a small crossing vessel in the mesentery at the border of the small bowel and divide this with cautery to avoid bleeding.</li>
		<li>Continue to divide the mesentery 1 cm towards the mesenteric base.</li>
		<li>Identify the proximal and distal jejunum. Place the proximal jejunum under a wet gauze on the rat&#39;s right and the distal jejunum on the rat&#39;s left.</li>
	</ol>
	</li>
	<li>Stapling the stomach
	<ol>
		<li>Insert a 45 mm linear cutting stapler with 3.5 mm staple height across the white line of the forestomach to create a smaller pouch. Wait for 10 s before firing the stapler.</li>
		<li>Place pressure using gauze on the staple lines for 1 min to ensure hemostasis. If hemostasis is not achieved with pressure alone, bleeding along the staple line is oversewn using 6-0 polypropylene figure of eight sutures.</li>
		<li>Perform a second staple fire across the stomach into the window created previously. Wait for 10 s before firing the stapler. Pressure is held along the staple line to ensure hemostasis, and oversewing may be needed.</li>
	</ol>
	</li>
	<li>Gastrojejunostomy
	<ol>
		<li>Make a gastrotomy immediately after stapling the stomach. Delays in this can cause gastric distension and aspiration as the stomach is discontinuous after the second gastric staple.</li>
		<li>Using an 11-blade scalpel, create a gastrotomy at the distal pouch. Express gastric contents through the gastrotomy. This is important to prevent gastric distension and aspiration. Lengthen this gastrotomy using micro scissors to approximately 5 mm. The gastrotomy is made large enough for the cotton swab tip just to fit through.</li>
		<li>Mobilize the distal end of the jejunum adjacent to the gastrotomy and place such that the mesentery is not twisted.</li>
		<li>While suturing the anastomosis, ensure that the bowel is kept moist by covering it with saline-soaked gauze and reapplying saline regularly.</li>
		<li>Using 6-0 polydioxanone or polypropylene suture, place a stay suture at the inferior margin of the anastomosis and gently retract using a snap. Tie with three knots.</li>
		<li>Place a stay suture at the superior margin of the anastomosis and gently retract using a snap. Tie with six knots.</li>
		<li>Suture the anterior side of the anastomosis in a continuous fashion, taking bites 1 mm wide and 1 mm apart with care to avoid taking the backside.</li>
		<li>Once the suture has reached the inferior stay suture, tie these together with an additional six knots.</li>
		<li>Once the anterior side is complete, flip the bowel and stomach over and pass the inferior stay suture through the mesenteric defect. Reapply the snap and retract inferiorly.</li>
		<li>For the posterior side of the anastomosis, place full thickness interrupted 6-0 sutures, 1 mm wide, and spaced 1 mm apart, with care to avoid taking the backside. These are tied with six knots each.<br />
		NOTE: The anterior side of the anastomosis is sutured in a continuous fashion while the posterior side is done in an interrupted fashion. This prevents potential stricture or stenosis associated with a continuous circumferential closure.</li>
		<li>Check for the leakage by gently pushing luminal contents across the anastomosis. If there are areas with leakage, carefully reinforce them with interrupted sutures. Take care to avoid taking the backwall when reinforcing with extra sutures.</li>
	</ol>
	</li>
	<li>Jejunojejunostomy
	<ol>
		<li>From the gastrojejunostomy, measure 20 cm distally.</li>
		<li>Create a jejunotomy on the antimesenteric side using the 11-blade scalpel. Avoid making the jejunotomy over Peyer&#39;s patches.</li>
		<li>Extend this jejunotomy using micro scissors, such that it is the same size as the biliopancreatic limb. Ensure that a cotton swab just fits inside.</li>
		<li>Place the biliopancreatic limb such that there is no twisting of the mesentery.</li>
		<li>Perform the anastomosis similarly to the gastrojejunostomy with 6-0 stay sutures on the superior and inferior sides. The anterior side is performed with continuous sutures while the posterior side is performed with interrupted sutures.</li>
		<li>Ensure that the bowel is kept moist with saline during this anastomosis.</li>
		<li>Check for leakage by gently pushing luminal contents through the anastomosis. If there are areas with leakage, reinforce them with interrupted sutures.</li>
	</ol>
	</li>
	<li>Reposition the bowel and stomach
	<ol>
		<li>Ensure that there is no twisting of the pouch, remnant stomach, or liver. Ensure that the left lobe of the liver is anterior to the stomach and not trapped behind the pouch as this can cause compressive liver ischemia.</li>
		<li>Position the bowel in the abdomen in its natural position such that there is no twisting.</li>
	</ol>
	</li>
	<li>Abdominal closure
	<ol>
		<li>Close the fascia with 3-0 polyglactin in a continuous fashion. 3-0 polydioxanone may also be used.</li>
		<li>Close the skin with 2-0 silk in a continuous fashion.</li>
	</ol>
	</li>
	<li>Anesthetic emergence
	<ol>
		<li>Decrease isoflurane to zero but continue supplemental oxygen.</li>
		<li>Administer a local anesthetic as a splash block to the incision.</li>
		<li>Administer 10 mL of subcutaneous 5% dextrose in normal saline (D5NS) in the subcutaneous tissue behind the neck.</li>
		<li>Place an Elizabethan rat collar before the rat is fully awake. Take care to fit it snugly but not too tight to cause discomfort.<br />
		&#8203;NOTE: The collar is kept on until day 5 to prevent wound dehiscence.</li>
	</ol>
	</li>
</ol>
<p class="jove_title">2. Sham surgery</p>
<p class="jove_content">NOTE: Sham surgery is performed similar to RYGB, however, no anastomoses are performed.</p>
<ol>
	<li>A gastrotomy is created and then closed with 6-0 polydioxanone or polypropylene sutures.</li>
	<li>A jejunotomy is created 7 cm distal to the ligament of Treitz and then closed with 6-0 polydioxanone or polypropylene sutures.</li>
</ol>
<p class="jove_title">3. Postoperative care</p>
<ol>
	<li>Postoperative care
	<ol>
		<li>House rats individually and keep them on raised wire platforms until solid food is reintroduced to prevent consumption of bedding and luminal obstruction.<br />
		NOTE: Postoperative diet is resumed gradually as edema at the gastrojejunostomy can cause obstruction with the early resumption of solid diet.</li>
		<li>Inspect the feet daily while rats are on raised wire platforms for any skin changes.</li>
		<li>Keep rats on water and oral rehydration therapy diet for the first 72 h.</li>
		<li>Administer 10 mL of D5NS every 12 h for the first 72 h.</li>
		<li>Administer subcutaneous short-acting buprenorphine at 0.01 mg/kg if rats appear to be in pain. The Rat Grimace Scale is used to assess for pain<sup class="xref">16</sup>.</li>
		<li>On postoperative day 3, add rodent liquid diet. Continue to provide water and oral rehydration therapy.</li>
		<li>On postoperative day 5, restart high-fat diet. Continue to provide water and liquid diet. Remove the Elizabethan collar.</li>
		<li>On postoperative day 7, discontinue liquid diet.</li>
		<li>Remove skin sutures on postoperative day 10-14.</li>
	</ol>
	</li>
</ol>

<ol><li>World Health Organization. <em>Obesity and Overweight</em>. , Available from: 
     <a target="_blank" href="http://www.who.int/news-room/fact-sheets/detail/obesity-and-overweight">http://www.who.int/news-room/fact-sheets/detail/obesity-and-overweight</a> (2018).</li>
<li>Courcoulas, A. P., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Three-year+outcomes+of+bariatric+surgery+vs+lifestyle+intervention+for+type+2+diabetes+mellitus+treatment%5BTitle%5D)+AND+%22JAMA+Surgery%22%5BJournal%5D)">Three-year outcomes of bariatric surgery vs lifestyle intervention for type 2 diabetes mellitus treatment.</a> <em>JAMA Surgery</em>. <strong>15213</strong> (10), 1-9 (2015).</li>
<li>Ardestani, A., Rhoads, D., Tavakkoli, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Insulin+cessation+and+diabetes+remission+after+bariatric+surgery+in+adults+with+insulin-treated+type+2+diabetes%5BTitle%5D)+AND+%22Diabetes+Care%22%5BJournal%5D)">Insulin cessation and diabetes remission after bariatric surgery in adults with insulin-treated type 2 diabetes.</a> <em>Diabetes Care</em>. <strong>38</strong> (4), 659-664 (2015).</li>
<li>Casella, G., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ten-year+duration+of+type+2+diabetes+as+prognostic+factor+for+remission+after+sleeve+gastrectomy%5BTitle%5D)+AND+%22Surgery+for+Obesity+and+Related+Diseases%22%5BJournal%5D)">Ten-year duration of type 2 diabetes as prognostic factor for remission after sleeve gastrectomy.</a> <em>Surgery for Obesity and Related Diseases</em>. <strong>7</strong> (6), 697-702 (2011).</li>
<li>Panunzi, S., De Gaetano, A., Carnicelli, A., Mingrone, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Predictors+of+remission+of+diabetes+mellitus+in+severely+obese+individuals+undergoing+bariatric+surgery%5BTitle%5D)+AND+%22Annals+of+Surgery%22%5BJournal%5D)">Predictors of remission of diabetes mellitus in severely obese individuals undergoing bariatric surgery.</a> <em>Annals of Surgery</em>. <strong>261</strong> (3), 459-467 (2015).</li>
<li>Edelman, S., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Control+of+type+2+diabetes+after+1+year+of+laparoscopic+adjustable+gastric+banding+in+the+helping+evaluate+reduction+in+obesity+%28HERO%29+study%5BTitle%5D)+AND+%22Diabetes%2C+Obesity+and+Metabolism%22%5BJournal%5D)">Control of type 2 diabetes after 1 year of laparoscopic adjustable gastric banding in the helping evaluate reduction in obesity (HERO) study.</a> <em>Diabetes, Obesity and Metabolism</em>. <strong>16</strong> (10), 1009-1015 (2014).</li>
<li>Mingrone, G., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Metabolic+surgery+versus+conventional+medical+therapy+in+patients+with+type+2+diabetes%3A+10-year+follow-up+of+an+open-label%2C+single-centre%2C+randomised+controlled+trial%5BTitle%5D)+AND+%22The+Lancet%22%5BJournal%5D)">Metabolic surgery versus conventional medical therapy in patients with type 2 diabetes: 10-year follow-up of an open-label, single-centre, randomised controlled trial.</a> <em>The Lancet</em>. <strong>397</strong> (10271), 293-304 (2021).</li>
<li>Thaler, J. P., Cummings, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Minireview%3A+Hormonal+and+metabolic+mechanisms+of+diabetes+remission+after+gastrointestinal+surgery%5BTitle%5D)+AND+%22Endocrinology%22%5BJournal%5D)">Minireview: Hormonal and metabolic mechanisms of diabetes remission after gastrointestinal surgery.</a> <em>Endocrinology</em>. <strong>150</strong> (6), 2518-2525 (2009).</li>
<li>Mingrone, G., Castagneto-Gissey, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mechanisms+of+early+improvement+%2F+resolution+of+type+2+diabetes+after+bariatric+surgery%5BTitle%5D)+AND+%22Diabetes+and+Metabolism%22%5BJournal%5D)">Mechanisms of early improvement / resolution of type 2 diabetes after bariatric surgery.</a> <em>Diabetes and Metabolism</em>. <strong>35</strong> (6), 518-523 (2009).</li>
<li>Arapis, K., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Remodeling+of+the+residual+gastric+mucosa+after+Roux-en-Y+gastric+bypass+or+vertical+sleeve+gastrectomy+in+diet-induced+obese+rats%5BTitle%5D)+AND+%22PloS+One%22%5BJournal%5D)">Remodeling of the residual gastric mucosa after Roux-en-Y gastric bypass or vertical sleeve gastrectomy in diet-induced obese rats.</a> <em>PloS One</em>. <strong>10</strong> (3), 012414(2015).</li>
<li>Bruinsma, B. G., Uygun, K., Yarmush, M. L., Saeidi, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Surgical+models+of+Roux-en-Y+gastric+bypass+surgery+and+sleeve+gastrectomy+in+rats+and+mice%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">Surgical models of Roux-en-Y gastric bypass surgery and sleeve gastrectomy in rats and mice.</a> <em>Nature Protocols</em>. <strong>10</strong> (3), 495-507 (2015).</li>
<li>Bueter, M., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Roux-en-Y+gastric+bypass+operation+in+rats+protocol%5BTitle%5D)+AND+%22Journal+of+visualized+experiments+%3A+JoVE%22%5BJournal%5D)">Roux-en-Y gastric bypass operation in rats protocol.</a> <em>Journal of visualized experiments : JoVE</em>.  (64), (2012).</li>
<li>Stevenson, M., Lee, J., Lau, R. G., Brathwaite, C. E. M., Ragolia, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Surgical+mouse+models+of+vertical+sleeve+gastrectomy+and+Roux-en+Y+gastric+bypass%3A+a+Review%5BTitle%5D)+AND+%22Obesity+Surgery%22%5BJournal%5D)">Surgical mouse models of vertical sleeve gastrectomy and Roux-en Y gastric bypass: a Review.</a> <em>Obesity Surgery</em>. , (2019).</li>
<li>Hao, Z., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Reprogramming+of+defended+body+weight+after+Roux-En-Y+gastric+bypass+surgery+in+diet-induced+obese+mice%5BTitle%5D)+AND+%22Obesity%22%5BJournal%5D)">Reprogramming of defended body weight after Roux-En-Y gastric bypass surgery in diet-induced obese mice.</a> <em>Obesity</em>. <strong>24</strong> (3), 654-660 (2016).</li>
<li>McErlane, S. Adult rodent anesthesia SOP. <em>UBC Animal Care Guidelines</em>. , Available from: 
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<li>Sotocinal, S. G., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+Rat+Grimace+Scale%3A+a+partially+automated+method+for+quantifying+pain+in+the+laboratory+rat+via+facial+expressions%5BTitle%5D)+AND+%22Molecular+pain%22%5BJournal%5D)">The Rat Grimace Scale: a partially automated method for quantifying pain in the laboratory rat via facial expressions.</a> <em>Molecular pain</em>. , 1-10 (2011).</li>
<li>Elder, K. A., Wolfe, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bariatric+surgery%3A+A+review+of+procedures+and+outcomes%5BTitle%5D)+AND+%22Gastroenterology%22%5BJournal%5D)">Bariatric surgery: A review of procedures and outcomes.</a> <em>Gastroenterology</em>. <strong>132</strong> (6), 2253-2271 (2007).</li>
<li>Lim, R. B. Bariatric procedures for the management of severe obesity: Descriptions. <em>UpToDate</em>. , Available from: 
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</ol>

<p class="jove_content">Ethicon supplied two 45 mm linear cutting staplers, multiple 3.5 mm stapler reloads, and 6-0 polypropylene sutures. Authors have no other conflicts of interest to declare.</p>

<p class="jove_content">RYGB involves the creation of a small gastric pouch (less than 30 mL), and the creation of a biliopancreatic limb and a Roux limb (<strong class='xfig'>Figure 1</strong>). In humans, the biliopancreatic limb is typically 30 to 50 cm and transports secretions from the gastric remnant, liver, and pancreas. The Roux limb is typically 75 to 150 cm in length and is the primary channel for ingested food. The common channel is the remaining small bowel distal to where the two limbs join and is where the majority of digestion and absorption occur, as pancreatic enzymes and bile mix with ingested food<sup class='xref'>17</sup>.</p>
<p class="jove_content">The mechanism of weight loss in RYGB is multimodal. The small gastric pouch reduces food intake through mechanical restriction. The bypass results in a malabsorptive component as a significant portion of the small intestine is not absorbing calories and nutrients. More recently, studies have demonstrated that gut hormones play a significant role in weight loss after RYGB as well. These are primarily through ghrelin, peptide-YY, cholecystokinin (CCK), and GLP-1 hormone pathways<sup class='xref'>18</sup>.</p>
<p class="jove_content">Rat models provide a powerful method to study the mechanisms behind both the weight and metabolic effects of RYGB. In this paper, we present a RYGB protocol that has low mortality with significant weight loss and metabolic effects. Once the operator became familiar with the technique, the procedure took approximately 90 minutes to perform. The protocol can also be modified with longer biliopancreatic and Roux limb lengths to potentially increase weight loss and metabolic effect. Furthermore, it is technically more feasible than other models as it allows the use of surgical staplers to achieve hemostasis and minimize operative time. Models that rely on sharp division of the stomach without stapling often result in higher mortality due to significant blood loss. The technical skills required to perform the procedure were relatively easy to acquire, and learners were able to comfortably perform the procedure after approximately five to ten non-recovery procedures.</p>
<p class="jove_content">One of the critical steps of this protocol is to limit blood loss during mobilization of the stomach. Careful use of thermal cautery combined with suture ligation of vessels is important. It is also important to perform at least half the circumference of the anastomoses in an interrupted manner. This prevents excessive stricturing at the anastomoses. Furthermore, checking for leaks is crucial as these can lead to sepsis and death. Prior to closing the abdomen, it is essential that the left lobe of the liver is placed in its natural, anterior position and that there is no rotation in the bowel or the stomach as this can lead to visceral ischemia.</p>
<p class="jove_content">Postoperative care is vital to this protocol. Raised wire platforms are required during both fasting and postoperative periods as the consumption of solid material leads to anastomotic obstructions. It is vitally important to provide subcutaneous fluid as the rats may not tolerate oral fluids in the immediate postoperative period. The rats should be acclimatized to oral rehydration therapy and liquid diet as rats may avoid new diets due to associations with postoperative pain. This dietary protocol contributes to significant weight loss in the immediate postoperative period in both the RYGB and sham cohorts, and weight recovery in the sham group took about five weeks. However, strict adherence to this postoperative protocol is vital to reduce morbidity and mortality after RYGB. Additionally, frequent examination of the rats using the Rat Grimace Scale is important to detect for morbidity. In our study, one rat developed a late anastomotic leak which was rapidly detected using this scale and allowed for early euthanasia to reduce suffering.</p>
<p class="jove_content">One of the advantages of this method is that it results in a smaller pouch through the use of surgical staplers to reduce gastric bleeding. When we attempted to sharply divide the stomach without staplers, it leads to excessive bleeding and a much higher mortality rate. However, this also leads to the removal of the forestomach, and this may lead to physiologic changes that are different from that of human RYGB. However, the forestomach is unique to rodents and contains no glands, and should not cause any changes to gut hormones.</p>
<p class="jove_content">The most important limitation of this method is that it requires two surgical stapler reloads per rat, which can be costly. However, excellent survival outcomes potentially reduce cost by requiring less rats for a study, resulting in better utilization of husbandry facilities, surgical equipment and research personnel.</p>

<p class="jove_content">Obesity and type 2 diabetes have become worldwide epidemics<sup class="xref">1</sup>. Although medical weight loss can improve diabetes in patients, those with severe diabetes benefit most from bariatric surgery. Bariatric surgery has proven to be safe and effective at weight loss and improving or curing type 2 diabetes<sup class="xref">2</sup><sup>,</sup><sup class="xref">3</sup>, even in those with long-standing disease<sup class="xref">4</sup>. Metabolic bariatric procedures, such as the current gold-standard Roux-en-Y gastric bypass (RYGB) surgery, induce rapid and sustained improvements in glucose homeostasis while also reducing the need for diabetic medications<sup class="xref">5</sup><sup>,</sup><sup class="xref">6</sup><sup>,</sup><sup class="xref">7</sup>.</p>
<p class="jove_content">After RYGB, glucose homeostasis improvement occurs rapidly and is independent of the weight loss<sup class="xref">8</sup>. Two major theories have been proposed to explain the metabolic changes associated with diabetes remission that occur following metabolic surgery. First, the hindgut hypothesis postulates that, after bypass, higher concentrations of undigested nutrients reach the distal intestine enhancing the release of hormones such as GLP-1. On the other hand, the foregut hypothesis suggests that bypassing the proximal intestine reduces the secretion of anti-incretin hormones. Both of these effects could lead to early improvement of glucose metabolism<sup class="xref">9</sup>.</p>
<p class="jove_content">Animal models have the potential to be a powerful tool to study these mechanisms. However, a major barrier in utilizing mouse or rat models is the technical difficulty in performing these procedures. Most studies have relied on mouse or rat models<sup class="xref">10</sup><sup>,</sup><sup class="xref">11</sup><sup>,</sup><sup class="xref">12</sup>. Mouse models have been difficult as the mouse stomach is too small to use stapler devices<sup class="xref">11</sup>, and mortality rates are unacceptably high, ranging from 17 to 52%<sup class="xref">13</sup>. In rats, some protocols remain technically difficult to perform due to complex ligation of gastric vessels prior to dividing the stomach<sup class="xref">12</sup><sup>,</sup><sup class="xref">14</sup>. Other models divide the stomach using a stapler but leave a large pouch not consistent with the post RYGB human anatomy<sup class="xref">11</sup>. In this model, we provide detailed instructions on how to perform RYGB using linear staplers in a rat model resulting in a gastric pouch more in keeping with that of human anatomy. Overall, this procedure was associated with excellent survival rates and metabolic outcomes.</p>

<table><tbody><tr><td>2-0 Silk Sutures</td><td>Ethicon</td><td>K533</td><td></td></tr><tr><td>3-0 Vicryl Sutures</td><td>Ethicon</td><td>J219H</td><td></td></tr><tr><td>4% Isoflurane</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>5% Dextrose and 0.9% Sodium Chloride Solution - 1000 mL</td><td>Baxter</td><td>2B1064</td><td></td></tr><tr><td>50 mL Conical Centrifuge Tubes</td><td>Fisher Scientific</td><td>14-432-22</td><td></td></tr><tr><td>6-0 Prolene Sutures</td><td>Ethicon</td><td>8805H</td><td></td></tr><tr><td>Anesthetic Machine</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Animal Hair Shaver</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Betadine Solution</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Castrojievo Needle Holder with lock 14 cm (smooth curved)</td><td>World Precision Instruments</td><td>503258</td><td></td></tr><tr><td>ECHELON FLEX Articulating Endoscopic Linear Cutter</td><td>Ethicon</td><td>EC45A</td><td></td></tr><tr><td>Economy Tweezers #4</td><td>World Precision Instruments</td><td>501978</td><td></td></tr><tr><td>ENDOPATH ETS Articulating Linear Cutter 45mm Reloads</td><td>Ethicon</td><td>6R45B</td><td></td></tr><tr><td>Far Infrared Warming Pad Controller with warming pad (15.2 cm W x 20.3 cm L), pad temperature probe, and 10 disposable, non-sterile sleeve protectors</td><td>Kent Scientific</td><td>RT-0515</td><td></td></tr><tr><td>Large Rat Elizabethan Collar</td><td>Kent Scientific</td><td>EC404VL-10</td><td></td></tr><tr><td>Liquid Diet Feeding Tube (150 mL)</td><td>Bio-Serv</td><td>9007</td><td></td></tr><tr><td>Liquid Diet Feeding Tube Holder (short adjustable)</td><td>Bio-Serv</td><td>9015</td><td></td></tr><tr><td>Micro Mosquito Forceps</td><td>World Precision Instruments</td><td>500452</td><td></td></tr><tr><td>Micro Scissors</td><td>World Precision Instruments</td><td>503365</td><td></td></tr><tr><td>Mouse Diet, High Fat Fat Calories (60%), Soft Pellets</td><td>Bio-Serv</td><td>S3282</td><td></td></tr><tr><td>No. 11 Blade and Scalpel Handle</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>OPMI Vario Surgical Microscope</td><td>ZEISS</td><td>S88</td><td></td></tr><tr><td>Raised Floor Grid</td><td>Tecniplast</td><td>GM500150 Raised Floor Grid</td><td></td></tr><tr><td>Rodent Liquid Diet, Lieber-DeCarli &#039;82, Control, 4 Liters/Bag</td><td>Bio-Serv</td><td>F1259</td><td></td></tr><tr><td>Sodium Chloride Irrigation 0.9% Solution - 500 mL</td><td>Baxter</td><td>JF7633</td><td></td></tr><tr><td>Sterile Cotton Swabs</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Sterile Drape</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Sterile Towel</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Thermal Cautery Unit</td><td>World Precision Instruments</td><td>501293</td><td></td></tr></tbody></table>

a protocol for roux-en-y gastric bypass in rats using linear staplers

<p class="jove_content">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.</p>
<p class="jove_content">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. </p>
<p class="jove_content">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.</p>
<p class="jove_content">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.</p>

<p class="jove_content"><strong>Animals and housing</strong><br>
36 male Wistar rats were housed in pairs and were fed 60% sterile rodent high-fat diet starting from six weeks of age (<strong class="xfig">Figure 2</strong>). At 16 weeks of age, they underwent RYGB or sham surgery. After the first postoperative week, rats were resumed on a high fat diet. Half of the rats were euthanized at 2 weeks post-operative and the other half were euthanized at 14 weeks postoperative.</p>
<p class="jove_content"><strong>Mortality</strong><br>
Overall, 33 (91.7%) rats survived to the planned study endpoint. All rats undergoing early euthanasia underwent necropsy by a veterinarian. Two rats were euthanized within 24 hours. One RYGB had aspiration pneumonitis and one sham rat had fascial dehiscence with unsalvageable bowel. Another RYGB rat was euthanized at two weeks due to anastomotic leak from the gastrojejunostomy. Overall, 88.9% of RYGB rats survived to study endpoint.</p>
<p class="jove_content"><strong>Body Weight</strong><br>
Rats undergoing RYGB had a lower postoperative weight than sham rats. <strong class="xfig">Figure 3</strong> demonstrates absolute weights for rats postoperatively while <strong class="xfig">Figure 4</strong> demonstrates postoperative percentage weight change which was statistically significant at all timepoints postoperatively. At 14 weeks, rats who had RYGB had a mean percentage weight change of 6.4% while rats with sham surgery had 23.7% (p = 0.0001).</p>
<p class="jove_content"><strong>Intraperitoneal glucose tolerance testing</strong><br>
Fasting blood glucose was not significantly different between any of the cohorts. However, the area under the curve was significantly lower in RYGB compared with sham at 13 weeks (18.1 vs 23.8 mmol-h/L, p=0.046, <strong class="xfig">Figure 5</strong>) but was the same for RYGB vs sham at 1 week (20.8 vs 23.3 mmol-h/L, p=0.68).</p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Digestive system diagram; pre vs. post-Roux-en-Y gastric bypass surgery anatomy." class="xfigimg" src="/files/ftp_upload/62575/62575fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"><br>
<strong class="xfig">Figure 1</strong><strong>: Roux-en-Y gastric bypass anatomy</strong> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62575/62575fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Weight progression graph over preoperative weeks, displaying data curves and error bars for analysis." class="xfigimg" src="/files/ftp_upload/62575/62575fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"><br>
<strong class="xfig">Figure 2</strong><strong>: Preoperative absolute weight on high fat diet; RYGB, Roux-en-Y gastric bypass</strong> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62575/62575fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Weight change graph over weeks for RYGB vs sham, with error bars, tracking study results." class="xfigimg" src="/files/ftp_upload/62575/62575fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"><br>
<strong class="xfig">Figure 3</strong><strong>: Postoperative absolute weight on high-fat diet; RYGB, Roux-en-Y gastric bypass</strong> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62575/62575fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Weight change graph comparing RYGB and Sham groups over 14 weeks; percentage change analysis." class="xfigimg" src="/files/ftp_upload/62575/62575fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"><br>
<strong class="xfig">Figure 4</strong><strong>: Postoperative percentage weight change on high-fat diet; RYGB, Roux-en-Y gastric bypass</strong> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62575/62575fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Blood glucose levels over time graph; RYGB vs. Sham, indicating post-surgical effects." class="xfigimg" src="/files/ftp_upload/62575/62575fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"><br>
<strong class="xfig">Figure 5</strong><strong>: Intraperitoneal glucose tolerance testing in gastric bypass vs sham at 13 weeks. RYGB, Roux-en-Y gastric bypass</strong> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62575/62575fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>

Watch this Scientific Journal Video about A Protocol for Roux-en-Y Gastric Bypass in Rats using Linear Staplers at JoVE.com

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

<p class="jove_content">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.</p>

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 ...

a-protocol-for-roux-en-y-gastric-bypass-in-rats-using-linear-staplers

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5.4K Views.  University of Alberta. 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.

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

Extraction

<p>Source: Lara Al Hariri at the University of Massachusetts Amherst, MA, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Preparation of the Laboratory<button class="expand">Expand</button>
	<p><em>Here, we show the laboratory preparation for 10 students working in pairs, with some excess. Please adjust quantities as needed.</em></p>
	<ul class="lab-items">
		<li data-sub-note-item="1-2">
		<div class="lab-step">Put on a lab coat, splash-proof safety glasses, and nitrile gloves.</div>
		</li>
		<li data-sub-note-item="1-3">
		<div class="lab-step">Place labeled containers for halogenated waste and solid reagent waste in a satellite waste hood. Place a glass waste container nearby.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Label a 20-mL vial as &#8216;unknown mixture&#8217;.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Measure out 0.5 g of cellulose and add it to the labeled vial. Measure out 4.75 g each of benzoic acid and caffeine and add them to the vial. Stir the solids well with a glass rod.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Tightly cap the vial and place it by the balances. Set a box of 90-mm circular filter paper by the balances. Make sure that the area has enough weighing boats, laboratory wipes, and clean spatulas.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Set out the following glassware and equipment at each student lab station (we suggest that students work in pairs):
		<table align="center" style="border-collapse: collapse; font-size: 17px; line-height: 24px;">
			<tbody>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;Lab stand with ring fixture</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;Hotplate</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;Vacuum pump (or house vacuum)</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;Medium 3-prong clamp</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;125-mL separatory funnel with a stopper</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;125-mL Erlenmeyer flask</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;3&#160;&#160;&#160;&#160;10-mL glass graduated cylinders</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;50-mL glass graduated cylinders</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;50-mL beakers</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;3&#160;&#160;&#160;&#160;100-mL beakers</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;600-mL beaker</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;250-mL filter flask</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;B&#252;chner funnel</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;60-mL glass funnels</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;100-mm watch glasses</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;Glass stirring rods</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;24/40 rubber adapters</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;Pasteur pipette bulbs</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;2&#160;&#160;&#160;&#160;Small spatulas</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;Package of pH paper</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px; ">&#160;1&#160;&#160;&#160;&#160;Roll of lab tape and pen</td>
				</tr>
			</tbody>
		</table>
		</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Distribute boxes of lab wipes and Pasteur pipettes around the lab so that each workstation has them nearby.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Set out anhydrous magnesium sulfate, sodium bicarbonate, vacuum grease, and several spatulas on a common bench.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Shortly before the lab, place labeled bottles of dichloromethane and 1 M NaOH in a fume hood.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Put a bottle of 3 M HCl in a separate fume hood to avoid inadvertent contact between acids and bases.</div>
		</li>
	</ul>
	</li>
</ol>

Extraction: Liquid-Liquid and Acid-Base Extraction - Prep

Wydobywanie

Source: Lara Al Hariri at the University of Massachusetts Amherst, MA, USA

	Preparation of the LaboratoryExpand
	Here, we show the laboratory preparation ...

Education

JoVE Lab Manual

Chemistry

Lab Manual Sub Articles

Lab Techniques

<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Preparation of Laboratory<button class="expand">Expand</button>
	<p><em>Here we show the laboratory preparation for 10 students working individually, with some excess. Please adjust quantities as needed.&#160;</em></p>
	<ul class="lab-items">
		<li data-sub-note-item="1-2">
		<div class="lab-step">To set up for this lab experiment, wear the appropriate personal protective equipment, including a lab coat, chemical splash goggles, and gloves.</div>
		</li>
		<li data-sub-note-item="1-3">
		<div class="lab-step">Obtain enough small plastic bottles with caps for the number of students and label them &#8216;NaCl&#8217;.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Use a top-loading balance to weigh 50 g of NaCl directly into each bottle. Place the bottles of NaCl at the student workstations.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Fill enough 250-mL wash bottles with deionized water for each student to have one.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Set out the following glassware and equipment at each student lab station:
		<table align="center" height="224" style="border-collapse: collapse; font-size: 17px; line-height: 24px;">
			<tbody>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;Stirring rod</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;20-mL volumetric flask</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;10-mL volumetric pipette</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;Pipetter</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;600-mL beaker</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;50-mL graduated cylinder</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;12-in ruler</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;Egg</td>
				</tr>
				<tr>
					<td style="border: 1px solid; padding: 0px;">&#160;1&#160;&#160;&#160;&#160;Eyedropper</td>
				</tr>
			</tbody>
		</table>
		</div>
		</li>
	</ul>
	</li>
</ol>

Accuracy, Precision, Uncertainty, and Significant Figures - Prep

Techniki laboratoryjne

Source: Smaa Koraym at Johns Hopkins University, MD, USA

	Preparation of LaboratoryExpand
	Here we show the laboratory preparation for 10 students ...

Buffers

<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse=""><strong>Preparing 50 mM NaH<sub>2</sub>PO<sub>4</sub> Buffer, pH 7</strong><button class="expand">Expand</button>
	<p>In the first part of this experiment, you will prepare a sodium phosphate solution buffered at pH 7.0. Monosodium phosphate is a weak acid with the conjugate base, disodium phosphate. Unadjusted monosodium phosphate solutions usually have a pH of about 4 - 6.</p>
	<p>Buffers are most effective close to their pKa, which is 6.8 to 7.2 for monosodium phosphate. So, you will use NaOH to push the equilibrium towards the conjugate base without altering the overall composition of the buffer equilibrium.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Before starting the experiment, calculate the mass of monosodium phosphate that you will need to make 200 mL of a 50 mM solution.
		<h4><strong>Table 1: Preparation of 50 mM NaH<sub>2</sub>PO<sub>4</sub> Buffer, pH 7.0</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always" height="247" width="358">
			<tbody>
				<tr>
					<td colspan="2"><strong>Molar mass of NaH<sub>2</sub>PO<sub>4</sub> = 119.98 g/mol</strong></td>
				</tr>
				<tr>
					<td>Solution volume (mL)</td>
					<td style="text-align: center;">200</td>
				</tr>
				<tr>
					<td>Molarity (mM)</td>
					<td style="text-align: center;">50</td>
				</tr>
				<tr>
					<td>Moles of NaH<sub>2</sub>PO<sub>4</sub> (mol)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Mass needed (mg)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Initial volume of 1 M NaOH (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Final volume of 1 M NaOH (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Volume of NaOH used (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Volume of DI water needed (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/45/Table_1._Preparation_of_50_mM_NaH2PO4_Buffer,_pH_7.0.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">To begin, put on the necessary personal protective equipment, including a lab coat, splash-resistant safety glasses, and gloves. <strong>Note: </strong>This section of the lab uses NaOH, which is corrosive and toxic. Use caution when pouring and transporting NaOH.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Fill a 250-mL plastic wash bottle with deionized water.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Label two 100-mL beakers, one for neutral aqueous waste and one for basic aqueous waste.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Calibrate your pH meter using the provided buffers. Store the probe in its storage solution when you are done.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Tare weighing paper, and use a clean spatula to measure out the required amount of monosodium phosphate according to your calculations. Record the exact amount of monosodium phosphate in your lab notebook. <strong>Note: </strong>Monosodium phosphate absorbs moisture from the air, so work quickly to obtain an accurate measurement and close the container tightly when you are done with it.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Place the monosodium phosphate in a 400-mL beaker and clean the spatula with a laboratory wipe. Throw out the weighing paper and the laboratory wipe before returning to your fume hood.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Use a graduated cylinder to measure out 175 mL of deionized water. Pour the water into the beaker of monosodium phosphate and add a magnetic stir bar.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Stir the solution until the salt has dissolved completely and the solution appears homogeneous. This usually takes 2 - 3 min.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Measure 15 mL of deionized water and pour it into the beaker. Continue stirring the solution until it appears homogeneous again (about 1 &#8211; 2 min). Turn off the stir motor.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Rinse the pH probe with deionized water, and then clamp it in the solution with the sensor above the stir bar.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Bring a 10-mL graduated cylinder and watch glass to the dispensing hood and measure out 10 mL of 1 M NaOH. Note the exact volume in the graduated cylinder.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Cover your NaOH with the watch glass, and carefully bring it to your fume hood.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Resume stirring the monosodium phosphate solution. While monitoring the pH reading, use a disposable pipette to slowly add NaOH to the stirring solution in a drop-wise manner. Once the buffer pH reaches 7.0, return any NaOH still in the pipette to the graduated cylinder.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Calculate the volume of NaOH that you added to the buffer. Subtract that volume from 10 mL to determine how much deionized water you must add to your buffer to reach a total solution volume of 200 mL.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">Measure the deionized water with another 10-mL graduated cylinder and add it to the stirring solution to finish making the buffer.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Rinse the pH sensor with deionized water, cover it with the storage solution filled cap, and unplug or turn off the probe.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Label a 250-mL polyethylene bottle as &#39;50 mM monosodium phosphate buffer, pH 7.0&#39;. Retrieve the magnetic stir bar from the solution.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Use a funnel to pour the buffer solution into the labeled bottle and cap it tightly.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse=""><strong>Absorption Spectroscopy of Free and Bound Neutral Red</strong><button class="expand">Expand</button>
	<p>Buffers can be used to evaluate compounds at specific pH values. In this section, you will record the absorbance spectrum of the indicator neutral red in various buffers. The protonated form of neutral red is red, and the deprotonated form is yellow-orange, meaning that they absorb green and blue-violet light, respectively. Thus, the acidic and basic forms have distinct absorption wavelengths. Protein binding alters the properties of neutral red, changing both its absorbance and its pKa.</p>
	<p>After measuring the absorbance spectrum of free neutral red at several pH values, you will add riboflavin-binding protein, or RP, to the solutions and measure the absorbances again. Absorbance intensity is related to concentration, so you will use the spectra to determine the pKa of free and bound neutral red after the lab.</p>
	<ul class="lab-items">
		<li data-sub-note-item="2-3">
		<div class="lab-step">Draw the following table in your lab notebook. Number the cuvettes 1 through 10 and list the nine buffer pH values that you will use. The 10th cuvette will be a deionized water blank. <strong> Note: </strong>Always hold cuvettes by the textured sides and wipe the transparent sides just before putting the cuvette in the spectrophotometer. Remember to align the transparent sides with the beam of light in the spectrophotometer.
		<h4><strong>Table 2: Absorbance of Free and Bound Neutral Red</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td style="text-align: center;"><strong>Cuvette #</strong></td>
					<td style="text-align: center;"><strong>Buffer pH </strong></td>
					<td style="text-align: center;"><strong>Abs. at &#955;<sub>max </sub>(Free NRH<sup>+</sup>)</strong></td>
					<td style="text-align: center;"><strong>Abs. at &#955;<sub>max</sub> (Bound NRH)</strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">1</td>
					<td style="text-align: center;">5.0</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">2</td>
					<td style="text-align: center;">5.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">3</td>
					<td style="text-align: center;">6.0</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">4</td>
					<td style="text-align: center;">6.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">5</td>
					<td style="text-align: center;">7.0</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">6</td>
					<td style="text-align: center;">7.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">7</td>
					<td style="text-align: center;">8.0</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">8</td>
					<td style="text-align: center;">8.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">9</td>
					<td style="text-align: center;">11</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">10</td>
					<td style="text-align: center;">blank</td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/45/Table_2._Absorbance_of_Free_and_Bound_Neutral_Red.xlsx" target="_blank">Click Here to download Table 2</a></div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Obtain ten 1.5-mL cuvettes and caps and label the caps 1 through 10 to match the table in your lab notebook.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Label a 25-mL beaker as &#8216;DIH2O&#8217; and fill it with deionized water.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Flush your neutral aqueous waste down the drain and relabel the beaker as &#39;aqueous buffer waste&#39;.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Label a 400-mL beaker for used micropipette tips.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Attach a tip to a 1-mL micropipette and use it to dispense 1000 &#181;L of deionized water into cuvette 10. This is your solvent blank.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">Place 925 &#181;L of your pH 7.0 monosodium phosphate buffer in cuvette 5.</div>
		</li>
		<li data-sub-note-item="2-10">
		<div class="lab-step">Bring the remaining empty cuvettes to the buffer table. Guided by the table in your lab notebook, dispense 925 &#181;L of each buffer into the appropriate cuvette using the labeled micropipettes. Be careful not to use the same pipette tip for different buffers.</div>
		</li>
		<li data-sub-note-item="2-11">
		<div class="lab-step">Once you have finished, bring the buffers back to your workbench. There, set a 200-&#181;L micropipette to dispense 75 &#181;L, and attach a tip to the micropipette.</div>
		</li>
		<li data-sub-note-item="2-12">
		<div class="lab-step">Dispense 75 &#181;L of neutral red into each of the nine buffer cuvettes. Invert each cuvette several times to thoroughly mix the solutions. <strong>Note: </strong>Replace the pipette tip if it touches a buffer.</div>
		</li>
		<li data-sub-note-item="2-13">
		<div class="lab-step">Once you have mixed neutral red with every buffer, take a picture, or write down the colors of the solutions.</div>
		</li>
		<li data-sub-note-item="2-14">
		<div class="lab-step">Turn on a hand-held spectrophotometer and allow the light source to warm up. Once it is ready, create a new experiment to measure absorbance versus wavelength for the free neutral red.</div>
		</li>
		<li data-sub-note-item="2-15">
		<div class="lab-step">Insert the cuvette of deionized water and acquire a spectrum of the deionized water. Set it as a solvent background or blank.</div>
		</li>
		<li data-sub-note-item="2-16">
		<div class="lab-step">Then, remove the blank and insert the cuvette of neutral red in the lowest pH buffer, which will have the highest concentration of protonated neutral red.</div>
		</li>
		<li data-sub-note-item="2-17">
		<div class="lab-step">Acquire a spectrum, which should show one strong peak. Identify the wavelength corresponding to the highest point of this peak, or the maximum. Record this wavelength in your lab notebook as lambda max for protonated free neutral red.</div>
		</li>
		<li data-sub-note-item="2-18">
		<div class="lab-step">Save the data and remove the cuvette. Record the absorbance in your notebook, then collect spectra for cuvettes 2 - 9 using the same procedure.</div>
		</li>
		<li data-sub-note-item="2-19">
		<div class="lab-step">Record the wavelength of the isosbestic point in your lab notebook. That is where all the spectra cross at a single wavelength. If a spectrum does not cross at that point, empty and clean the cuvette, prepare a fresh sample, and try again.</div>
		</li>
		<li data-sub-note-item="2-20">
		<div class="lab-step">Once you finish collecting spectra for all nine cuvettes, save and export the data.</div>
		</li>
		<li data-sub-note-item="2-21">
		<div class="lab-step">Create a new experiment to measure absorbance versus wavelength for the bound neutral red.</div>
		</li>
		<li data-sub-note-item="2-22">
		<div class="lab-step">Fit a new tip to the 200-&#181;L micropipette and add 75 &#181;L of riboflavin-binding protein solution to each cuvette, including the blank. Eject the tip, secure the cuvette caps, and invert the cuvette several times to mix the solutions.</div>
		</li>
		<li data-sub-note-item="2-23">
		<div class="lab-step">Insert cuvette 10 into the spectrophotometer and set it as the solvent blank.</div>
		</li>
		<li data-sub-note-item="2-24">
		<div class="lab-step">Acquire a spectrum of the lowest pH sample, and identify the wavelength corresponding to the maximum of the most intense peak. Record it in your lab notebook as the lambda max of protonated-bound neutral red.</div>
		</li>
		<li data-sub-note-item="2-25">
		<div class="lab-step">Acquire spectra for cuvettes 2 - 9 in the same way as you did for the free neutral red samples. Record the intensities at lambda max for protonated-bound neutral red in your table, along with the wavelength at the isosbestic point.</div>
		</li>
		<li data-sub-note-item="2-26">
		<div class="lab-step">When you are done, save your data, export it for later analysis, and shut down the spectrophotometer.</div>
		</li>
		<li data-sub-note-item="2-27">
		<div class="lab-step">Put away all equipment and dispose of your used pipette tips in an approved waste container or trash bin.</div>
		</li>
		<li data-sub-note-item="2-28">
		<div class="lab-step">Empty the cuvettes into the aqueous buffer waste beaker and rinse the cuvettes into the beaker with deionized water.</div>
		</li>
		<li data-sub-note-item="2-29">
		<div class="lab-step">Empty the excess NaOH into the basic waste and rinse the graduated cylinder with water.</div>
		</li>
		<li data-sub-note-item="2-30">
		<div class="lab-step">Flush the basic aqueous waste down the drain with copious tap water, along with the other aqueous waste.</div>
		</li>
		<li data-sub-note-item="2-31">
		<div class="lab-step">Wash your glassware, cuvette caps, and stir bar per your lab standard procedures. Clean your work surfaces with a damp paper towel and throw out used paper towels and lab wipes in the lab trash.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="3" data-parent-collapse=""><strong>Results</strong><button class="expand">Expand</button>
	<p>Now, let&#39;s analyze our absorbance data to determine the pKa&#39;s of neutral red.</p>
	<h4><strong>Table 3: Determination of Neutral Red pKa&#39;s</strong></h4>
	<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
		<tbody>
			<tr>
				<td></td>
				<td>Free NRH<sup>+</sup></td>
				<td>Bound NRH<sup>+</sup></td>
			</tr>
			<tr>
				<td>&#160;&#955;<sub>max</sub> (nm)</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td>&#160;&#916;A (nm)</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td>&#160;Midpoint (nm)</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td>&#160;pKa</td>
				<td></td>
				<td></td>
			</tr>
		</tbody>
	</table>
	<p><a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/45/Table_3._Determination_of_Neutral_Red_pKa.xlsx" target="_blank">Click Here to download Table 3</a></p>
	<ul class="lab-items">
		<li data-sub-note-item="3-2">
		<div class="lab-step">Plot both sets of intensity data with absorbance intensity at lambda max on the y-axis and pH on the x-axis, with the points joined by smooth lines.</div>
		</li>
		<li data-sub-note-item="3-3">
		<div class="lab-step">Calculate the difference between the starting and ending absorbance intensities for each data series.</div>
		</li>
		<li data-sub-note-item="3-4">
		<div class="lab-step">Determine the absorbance midway between the starting and ending absorbances for each series, or the absorbance midpoints.</div>
		</li>
		<li data-sub-note-item="3-5">
		<div class="lab-step">Find where the midpoint occurs on each line and identify the pH value at that point. These pH values are the pKa&#39;s of free and bound neutral red.</div>
		</li>
		<li data-sub-note-item="3-6">
		<div class="lab-step">Here, we see an increase in pKa of about one between free and bound neutral red, indicating that bound-protonated neutral red is a weaker acid than free protonated neutral red by an order of magnitude.</div>
		</li>
	</ul>
	</li>
</ol>

Buffers: Common Ion Effect, Henderson-Hasselbalch Equation, and Buffer Capacity - Procedure

Source: Smaa Koraym at Johns Hopkins University, MD, USA

	Preparing 50 mM NaH2PO4 Buffer, pH 7Expand
	In the first part of this experiment, you will ...

Optimal Foraging

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Simulating Foraging with Artificial Habitats and Prey
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: The foragers will hunt for prey represented by pinto beans in four buckets of rice with varying prey densities. Without knowing what these densities are, foragers must obtain as many prey items as possible in as little time as possible. 
</div></li><li data-sub-note-item="1-2"><div class="lab-step">HYPOTHESES: In this experiment, the experimental hypothesis could be that foragers will catch the most prey in the higher prey density bucket and also spend the most time foraging there.  The null hypothesis would then be that foragers will catch an equal number of prey in each bucket and that foragers will spend an equal amount of time foraging in each patch. 
</div></li><li data-sub-note-item="1-3"><div class="lab-step">To begin the foraging activity, record the names and role of each participant at the top of the student data collection sheet.  NOTE: Before starting the experiment, the forager is allowed to become familiar with the foraging process &#8211; the following steps are a practice round. 
</div></li><li data-sub-note-item="1-4"><div class="lab-step">The forager should kneel or sit in front of the average density bucket. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Once the timer says start, the forager may begin looking for the pinto beans representing the prey. The forager can spend as much time as they like foraging in this bucket. 
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Once a prey item is located, the forager must signal that one has been found and place the prey in the cup next to the bucket.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Once they believe they have acquired most of the prey in the patch, they should tell the timer to stop the time and the practice round is over. 
</div></li><li data-sub-note-item="1-8"><div class="lab-step">To begin the simulation proper, the forager should sit or kneel in front of bucket A. 
</div></li><li data-sub-note-item="1-9"><div class="lab-step">The timer must be ready to start the stopwatch. 
</div></li><li data-sub-note-item="1-10"><div class="lab-step">The recorder should be ready to record the time each prey item is located, in addition to the time that the forager arrived at and left each bucket. 
</div></li><li data-sub-note-item="1-11"><div class="lab-step">When all parties are ready, the timer should start the stopwatch and say &#8220;start&#8221;,  and the forager should begin foraging. NOTE: The stopwatch will run through the rest of the round for buckets B and C.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">As the forager signals finding pinto beans and places them in the cup next to the bucket, the timer should tell the recorder the time.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">The recorder should then write the time in the student data collection sheet.  <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/34/Table_1_Optimal_foraging.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Once the forager has decided to move on to the next bucket, they should tell the timer &#8220;done&#8221;, and begin moving to the next patch. 
</div></li><li data-sub-note-item="1-15"><div class="lab-step">The recorder should record the time the forager left and the time they arrived at bucket B in the student data collection sheet.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Repeat the same procedure for buckets B and C. Once the forager has completed the activity, it is time for the next forager to begin using the same procedure. 
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">Before performing any data analysis, review the data collection sheet and compile your individual data for captured prey, foraging time, and travel time for each bucket in your student calculations table. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/34/Table_2_Optimal_foraging.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="2-2"><div class="lab-step">After this, calculate your individual capture rate for each bucket by dividing the number of prey captured by the sum of the foraging time and travel time and record these values in your table. Individual capture rate =  Number of prey captured / (foraging time + travel time)
</div></li><li data-sub-note-item="2-3"><div class="lab-step">To calculate your Giving-Up-Time or a GUT for each bucket, subtract the time that the last prey item was acquired from the time you left the bucket.  GUT = time forager left bucket  -  time last prey item acquired
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Next, calculate the mean GUT for your foraging group and record the values in your table.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Finally, calculate your scaled GUT for each bucket by dividing the bucket GUT by your group&#39;s mean GUT. Scaled GUT =  Bucket GUT / Group mean GUT NOTE: The scaled GUT will allow class data to be pooled later on without differences in individual foragers skewing the data. 
</div></li><li data-sub-note-item="2-6"><div class="lab-step">The instructor should now compile the data into the classroom spreadsheet.  <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/34/Table_3_Optimal_foraging.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Plot a bar graph of the average number of prey collected in each bucket. On the x-axis, A should represent the low prey density bucket, B should represent the medium prey density bucket, and C the high prey density bucket. Plot the y-axis as the number of prey items caught. 
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Next, plot the graph of the class average scaled GUT, where the x-axis represents the buckets with varying levels of prey density and the y-axis shows the values for the average scaled GUT. 
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Plot a third graph of the average time spent foraging in each bucket. 
</div></li><li data-sub-note-item="2-10"><div class="lab-step">Now review the Marginal Value Theorem (MVT) from the concepts video and examine the three graphs that you generated from your class data. Take note of whether the graphs conform to the predictions of MVT. 
</div></li></ul></li></ol>

Optimal Foraging: Foraging with Artificial Habitats and Prey - Procedure

Optymalne żerowanie

Simulating Foraging with Artificial Habitats and Prey
ExpandNOTE: The foragers will hunt for prey represented by pinto beans in four buckets of rice with ...

Biology

Ecology

Physiology of the Circulatory System

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Measuring Blood Pressure
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">To prepare for the blood pressure exercise, simply place the appropriate number of alcohol swabs, sphygmomanometers, and stethoscopes at the front of the classroom.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Be sure to check over each of the component parts of the sphygmomanometers, including the tubing, cuff, manometer, and bulb to ensure they are undamaged.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Respiration in Fish
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To prepare for the fish respiration experiment, one week before the goldfish arrive, set up two 10-gallon tanks in the room near a window.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Add around 9 gallons of water to the tanks to a depth of about 1 inch below the rim.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Then, mix in dechlorinator according the directions on the bottle.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Add in liquid beneficial bacteria according to the manufacturer&#39;s instructions.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Now, hang the filter on the side of the tank, fill up the trough with water, and then plug the filter in to begin cycling the water.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Once you plug in the filter, check to see if it is running properly.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Leave the tank to cycle for one week before adding fish. This will establish the beneficial bacteria, creating a healthy environment for the animals.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">The fish should arrive in a plastic bag filled with water. Float this bag in the 10-gallon tank for approximately 20 minutes, so that the water in the bag and tank acclimate to the same temperature.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">When the water has acclimated, carefully open the bag and add some of the tank water, and then gently pour the fish into the tank. IMPORTANT: The fish should not be fed the day they are introduced into the tank.
</div></li><li data-sub-note-item="2-10"><div class="lab-step">Feed the fish the following day with just a pinch of crushed food.
</div></li><li data-sub-note-item="2-11"><div class="lab-step">Change 10% of the water everyday, making sure the fresh water is at the same temperature as the tank water. Don&#39;t forget to add fresh dechlorinator when you change the water.
</div></li><li data-sub-note-item="2-12"><div class="lab-step">One hour before class, set up five water baths in the front of the classroom. Label each water bath with one of the five temperature conditions.
</div></li><li data-sub-note-item="2-13"><div class="lab-step">Next, fill the water baths up halfway with either aquarium water or dechlorinated tap water.
</div></li><li data-sub-note-item="2-14"><div class="lab-step">Then, set the temperatures in accordance with the labels, and place a temperature probe into each water bath.
</div></li><li data-sub-note-item="2-15"><div class="lab-step">If the temperature is warmer than 20 - 22 &#176;C, use dechlorinated ice cubes to cool the water down before the lab.
</div></li><li data-sub-note-item="2-16"><div class="lab-step">Finally, place enough clear containers with screw caps next to the tanks, so that each pair of students will have two.
</div></li></ul></li></ol>

Fizjologia układu krążenia

Measuring Blood Pressure
ExpandTo prepare for the blood pressure exercise, simply place the appropriate number of alcohol swabs, sphygmomanometers, and ...

Fundamentals

Animal Behavior

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Making Choice Chambers
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">Before beginning the experiments, use one of the Sharpies from the lab table to trace a line around the circumference of the bottom of both bottles. One line should be 1 cm from the base of the bottle and the other should be 1.5 cm from the base of the bottle.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Use a razor blade to make a small cut in each bottle at the mark. 
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, insert the scissors and carefully cut along the line until the bottoms of the bottles are removed completely.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Fit one bottle bottom into the other.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Tape the bottles together to make a secure chamber.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Use the razor blade and scissors to poke a hole into the center of the chamber. 
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Take a piece of tape and fold the edge onto itself to make a tab and place the tape over the hole.  Note: The tab helps with easy removal and reattachment of the tape to the chamber.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Use the Sharpie to label one side of the chamber A and the other side B.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Upon receiving the colony of <em>Drosophila</em>, perform a count to make sure there are 40 living individuals in the vial.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Geotaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To perform a geotaxis assay, first attach a clean, empty bottle cap to each end of the choice chamber and turn the fly vial upside down onto its lid.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Open the tape flap in the center of the choice chamber.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Insert the funnel into the hole and tap all of the flies into the chamber. HYPOTHESES: The alternate hypothesis is that more flies will be moving against gravity, or up, regardless of the orientation of the choice chamber. Alternatively, if the flies are positively geotactic, then more flies will move down, or with gravity. The null hypothesis is that the flies will have no preference with regard to gravity and will move randomly within the chamber. 
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Once the <em>Drosophila</em> are in the chamber, close the tape flap, rotate the chamber so that it stands vertically with side A on the bottom, and start the timer.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Immediately, count the number of <em>Drosophila</em> that are moving up and the number that are moving down. Record these data for time point 0 in the table. 
</div></li><li data-sub-note-item="2-6"><div class="lab-step">After 30 seconds, rotate the chamber so that side B is on the bottom and count the number of <em>Drosophila</em> that are moving up or down. Record these data for time point 30 in the table.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">After 60 seconds, rotate the chamber so that side A is on the bottom again and record the number of moving flies for time point 60.  <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/15/Table_1_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Hold the chamber over the fly vial and carefully open the cap to return all of the <em>Drosophila</em> back to their empty vial. 
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Phototaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">To perform a phototaxis analysis, first cover side A of the choice chamber with tin foil, taking care that the foil reaches to the exact center between the two sides. 
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Ensure the caps are secured on both ends of the chamber.  HYPOTHESES: In this exercise, the alternative hypothesis is that more flies will be found in the light side of the choice chamber, if the flues are attracted to light, or on the dark side if they are repelled by light. The null hypothesis is that <em>Drosophila</em> will have no preference with regard to either stimulus. Therefore, equal numbers of flies will be found on either side of the choice chamber.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Open the tape flap in the center of the choice chamber, insert the funnel into the hole, and tap all 40 flies into the chamber as previously described.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Set the timer to sound an alarm at 10 minutes.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">When the alarm sounds, stop the timer, and count the number of flies in the open side B of the chamber.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Subtract that number from the total number of flies added to the choice chamber to calculate the number of flies in the covered side A.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Record these data in the table for time point 10.  <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/15/Table_2_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Carefully, return all of the flies back to their empty vial.
</div></li></ul></li><li data-note-item="4" data-parent-collapse="">Chemotaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="4-1"><div class="lab-step">HYPOTHESES: In this exercise, the alternative hypothesis is that more flies will be found on the side of the chamber with the apple cider vinegar if the flies are attracted to the odor. Or, if the flies are repelled by the vinegar, then more of them will be found on the side of the chamber presenting the odorless water. The null hypothesis is that <em>Drosophila</em> will have no preference with regard to either stimulus and will move randomly in the chamber.  Use an individual disposable plastic pipette to dispense two drops of the apple cider vinegar onto one cotton ball for side A of the chamber. 
</div></li><li data-sub-note-item="4-2"><div class="lab-step">Use an individual disposable plastic pipette to dispense two drops of the distilled water onto a cotton ball for side B of the chamber. 
</div></li><li data-sub-note-item="4-3"><div class="lab-step">Fold two pieces of tape over themselves to create individual pieces of double-sided tape and use the tape to attach each cotton ball to the inside of the single bottle cap.
</div></li><li data-sub-note-item="4-4"><div class="lab-step">Screw the bottle caps with the cotton balls onto their corresponding sides of the chamber.
</div></li><li data-sub-note-item="4-5"><div class="lab-step">Transfer all of the flies into the chamber and set the timer to sound an alarm after 10 minutes. 
</div></li><li data-sub-note-item="4-6"><div class="lab-step">When the alarm sounds, stop the timer and record the number of flies in each side of the chamber. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/15/Table_3_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="4-7"><div class="lab-step">At the end of the final experiment, funnel the flies into a morgue tube containing 70% ethanol. 
</div></li><li data-sub-note-item="4-8"><div class="lab-step">Dismantle the choice chambers for recycling.  NOTE: Since all of the chemicals used in these experiments are lab safe, the liquids can be disposed of down the sink, the chambers rinsed, and the solids can be thrown away in the trash. 
</div></li></ul></li><li data-note-item="5" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="5-1"><div class="lab-step">Gather all of the data collected for the geotaxis, phototaxis, and chemotaxis experiments.
</div></li><li data-sub-note-item="5-2"><div class="lab-step">Plot these values as bar graphs for different conditions and different time points.
</div></li><li data-sub-note-item="5-3"><div class="lab-step">Use these data to draw inferences about the behavior of <em>drosophila melanogaster</em>. 
</div></li></ul></li></ol>

Animal Behavior: Studying Taxis Behavior of Drosophila - Procedure

Zachowanie zwierząt

Making Choice Chambers
ExpandBefore beginning the experiments, use one of the Sharpies from the lab table to trace a line around the circumference of the ...

Artificial Selection

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Observing Artificial Selection in Plants 
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">To begin the experiment, collect five two-week old grown plants from the generation one population that appear healthy. Note: Plants should appear green, not wilted, and not have damaged leaves. Experimental Hypothesis: The experimental hypothesis for this lab might be that the number of lead trichomes varies naturally, but the average number will increase over generations if only plants with the highest number of trichomes are interbred or decrease if those with the least are interbred. Null Hypothesis: The null hypothesis might be that the average number of trichomes will not change over subsequent generations after artificial selection.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Label each plant with a letter from A to E and with both partners&#8217; names.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Record any natural differences you see between your plant samples. Note: All of the plants are the same age, so any observed variation is due to expressed phenotype, not development.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Use a hand lens to count the number of trichomes found on the petiole of the lowest leaf, not including the round, first-grown cotyledons for each plant. Note: It can be helpful to shine a light onto the plant and to hold it against a dark background, such as a lab bench, during the trichome analysis. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Both partners will conduct counts on each plant independently and compare answers. If the counts are different, enlist a third counter or take the average of the two results.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">After this, determine the median number of trichomes for the entire class.  <a target="__blank" style="display:block;padding:20px;" href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/2/Lab_2_Table_A.xlsx">Click Here to download Table A</a>
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Determine the top and bottom 10% of the plants from the class trichome count. These groups will make up the two artificially selected lines: high hairiness and low hairiness, respectively.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">If more than 10% of the plants have no trichomes, select 10% of those plants randomly for the low hairiness plant line.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Separate the two groups of selected plants and label them clearly.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Move the plants to a well-lit area and keep them well watered until they develop flowers. Note: Flowers typically develop around 14 days.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">When the flowers are fully developed, use a pollination wand or paintbrush to rub the stigma and anther of one plant&#8217;s flowers for a few seconds.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Using the same wand or brush, rub the inside of each flower of every other plant in the same selection line.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Using a different wand or brush, repeat steps 11 through 12 on the other line. Note: Take care not to cross the lines. Spread the pollen only among the appropriately selected plants.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Return the generation one plants to the growing areas. When the plants reach approximately 1 month old, they should produce mature seed pods.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Allow the plants to dry out once the seed pods turn yellow.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Break the seed pods open and collect the seeds.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Obtain new pots for the generation two seeds.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Fill the pots with moist potting soil about halfway and add a fertilizer pellet to each pot.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Fill the pots with more soil to fill just below the lip.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Make a shallow depression in the soil of each pot with a probe.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Touch a clean toothpick to a gluestick and use the toothpick to transplant the seeds into the soil one by one.
</div></li><li data-sub-note-item="1-22"><div class="lab-step">Cover the seeds with a thin layer of soil.
</div></li><li data-sub-note-item="1-23"><div class="lab-step">Label the pots with their hairiness line and place them in the designated growing area.
</div></li><li data-sub-note-item="1-24"><div class="lab-step">Use a plastic dropper to water the plants as needed to ensure they do not dry out.
</div></li><li data-sub-note-item="1-25"><div class="lab-step">When the plants are two weeks old, collect five plants from each line and label them A to E.
</div></li><li data-sub-note-item="1-26"><div class="lab-step">Then, count the trichomes on these plants as previously described (steps 4- 5).
<a target="__blank" style="display:block;padding:20px;" href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/2/Lab_2_Table_B.xlsx">Click Here to download Table B</a>
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Data Analysis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">Create a frequency graph based on the generation one class data with the trichome count on the x axis and the number of plants on the y axis. Label the mean. 
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Then, create frequency graphs for both of the selected hairiness lines.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Compare the frequency graphs for generation one and both hairiness lines.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Take note of phenotypic shifts observed in the selected hairiness lines.
</div></li></ul></li></ol>

Artificial Selection: Visualization in Plants and Data Analysis - Procedure

Sztuczna selekcja

Observing Artificial Selection in Plants 
ExpandTo begin the experiment, collect five two-week old grown plants from the generation one population that ...

Evolution

Genetics of Organisms

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">REF and WEF Fly Genetics
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">To prepare for the experiment, be sure to obtain cultures of <em>Drosophila melanogaster</em> at least two weeks ahead of the planned experiment day. NOTE: Because adult females store sperm after mating, you will need this time to generate virgin females by breeding the flies.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">You should get two stock cultures: wildtype, that is homozygous dominant red eyed flies, and white chromosome one mutant, which are homozygous recessive white eyed flies.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Purchase or create <em>Drosophila </em>habitats consisting of a clean vial, 3 cm of prepared food media, and a stopper to close the open end of the vial.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">When not being moved or counted, keep the flies in an incubator or room kept at 23 - 25 &#176;C, where the lifecycle will take around 12 days. NOTE: From the egg, <em>Drosophila </em>progress through four main stages of development: embryo, larva, pupa, and adult. The complete lifecycle takes about 10 days at 25 &#176;C, but it can be affected by temperature. For example, at 18 &#176;C, the lifecycle is about 19 days and at 29 &#176;C, the lifecycle is only seven days.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">As an overview, to set up for the student experiment, you will generate vials for each student group containing white eyed virgin female flies only, white eyed males only, red eyed virgin females only, and red eyed males only. These will be referred to as the experimental vials.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">When your vials from the supplier arrive, referred to as the stock vials, these will contain a mix of male and female flies, as well as larvae, pupae, and eggs.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Beginning with your stock vials, you will take around 3 - 5 males and the same number of females, keeping white eyed and red eyed flies separate at all times, and place these together into a new nursery vial containing food. NOTE: You can sex the flies using a magnifying glass. Look at the posterior of the fly. Males are slightly smaller and have darker solid pigment on the end of their abdomen. Females are larger and have stripes rather than a solid spot.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">You will leave the flies in these nursery vials for around 3 - 5 days to mate and lay eggs.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">After this, you will move them back to their stock vials.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Next, you will observe as the eggs and larvae in the nursery vials mature, checking the vials every 6 - 8 hours. As it takes females around 8 - 10 hours to become sexually mature, this should ensure that any females found in the nursery vials are virgin.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">As the new adult flies appear, you should transfer them from the nursery to the appropriate experimental tube. For example, if it is a male, transfer it to the experimental male tube for the appropriate eye color. If it is a female, it should be placed in an experimental vial labeled with eye color and virgin female.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Keep adding flies to the appropriate vials until each experimental vial has at least five flies. NOTE: You will need to make enough experimental vials so that each student group has one of each eye color type and sex, with a few class spares. You should create enough nursery vials to account for this. A good rule of thumb is that it generally takes about two weeks for 25 healthy adults to produce about 50 new adults.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">To begin the process, first label empty habitats to make the nursery vials to raise larvae and pupae for each strain of flies: red eyed and white eyed.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">To move the adult flies into the nursery vial to lay eggs, use an anesthesia wand according to the manufacturer&#39;s instructions, placing it into the stock vial of mature flies. Wait until the animals become unconscious. Now they can be gently tapped onto a piece of white paper and sorted into piles of the same sex using a brush.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Transfer a few male and female flies to the appropriate vial labeled with their eye color phenotype and sex.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Place the nursery vials to incubate at 23 to 25 &#176;C and the flies will lay eggs.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Remove the adults after 3 - 4 days using anesthesia and put them back into the stock vial.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Remove any adult flies that have eclosed in the 6 - 8 hour period using an anesthesia wand.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Then, tip out the adults onto paper and sort them into groups based on their sex before placing each fly into an appropriate experimental vial marked male or virgin female and red or white for the phenotype. NOTE: Each experimental vial should contain five flies. Once this number is reached, then a new vial should be started. If you miss the 6 - 8 hour window, females may not be virgins, so these should be discarded into a morgue tube containing 70% ethanol or returned to the stock vial. Males do not need to be virgins, but do need to be kept in male only containers.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Each lab group will need one tube of five virgin red eyed females, one of five virgin white eyed females, one tube of red eyed males, and one of five white eyed males. In addition, they will need a morgue vial containing ethanol, two anesthesia wands, fly anesthetic, a paintbrush, a magnifying glass, some plain paper, and two empty fly vials for crosses.
</div></li></ul></li></ol>

Genetics of Organisms: REF and WEF Fly Genetics - Prep

Genetyka organizmów

REF and WEF Fly Genetics
ExpandTo prepare for the experiment, be sure to obtain cultures of Drosophila melanogaster at least two weeks ahead of the ...

Genetics

Bacterial Transformation

<ol class="note-items"><li data-note-item="1" data-parent-collapse=""><strong>Bacterial Transformation</strong>
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">First, ensure that each student or student group has a filled ice bucket.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Then, set up a tube rack for each work station.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Into the tube racks, place two empty, sterile 1.5 mL tubes, and then two one mL aliquots of SOC media.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Then, place the plasmid tube into the ice bucket and add two tubes of competent E. coli cells, allowing them to thaw if frozen.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Set out two LB agar plates per group and two LB agar ampicillin plates per group.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Ensure the students have the appropriate pipettes.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">And finally, set a water bath to 42 &#176;C and a temperature-controlled shaker to 37 &#176;C.
</div></li></ul></li></ol>

Bacterial Transformation using Plasmids - Prep

Transformacja bakteryjna

Bacterial Transformation
ExpandFirst, ensure that each student or student group has a filled ice bucket.
Then, set up a tube rack for each work station.
...

Animal Diversity

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Form and Function of Crickets and Crayfish
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: To observe the relationship between form and function in animals, you will be examining and dissecting a cricket and a crayfish. Each pair of students will receive one of each of these organisms. When dissecting, be sure to wear gloves, closed-toed shoes, and safety goggles. Take care when handling dissecting tools, as they may be sharp. HYPOTHESES: In this experiment, the experimental hypothesis could be that the cricket and crayfish have similar body structures because they are both arthropods, but that they will have some specific adaptations for their different habitats and lifestyles. The null hypothesis might be that the cricket and crayfish have identical adaptations and body plans.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">To begin, place the cricket on its back and observe its appendages.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">In Table 1, make detailed notes about your observations and try to predict the potential function of each of the forms you observe. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/27/Table_1_Animal_Diversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Do the same with the wings of the cricket if they are present.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Consider how the structure of the wings benefits the cricket. 
</div></li><li data-sub-note-item="1-6"><div class="lab-step">If the cricket does not have wings, consider the reasons behind their absence. 
</div></li><li data-sub-note-item="1-7"><div class="lab-step">If the cricket is female, it will have an ovipositor. This can be observed as a long dark tube protruding from the posterior end and it is used to deposit eggs.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Consider why the ovipositor is useful. 
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Next, flip the cricket over onto its back and observe the mouth parts under a microscope. NOTE: Use a probe to move structures when necessary.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Make a hypothesis about the size and type of prey the cricket consumes based upon your observations.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Continue to examine the external parts of the cricket, making hypotheses about their function until you have filled at least five rows in the table with your observations.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Using tweezers to hold the cricket down, carefully remove the wings and legs from the body with the scissors.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Next, position the body of the cricket so that its underside is facing down.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Again with the scissors, cut the thin outer exoskeleton from the anus to the head trying not to damage any of the organs underneath.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Use the tweezers to open the incision and dissecting pins to hold open the exoskeleton.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Next, use the water pitcher to carefully add water to the Petri dish until it reaches just below the top edge. This should make the exposed organs of the cricket easier to see.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Move the Petri dish to the stage of the dissecting microscope and turn on the light.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Use the cricket figure as a reference and observe the organs under the dissecting microscope. Where necessary, use the probe to move organs that may be blocking others.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Identify as many internal structures as possible and fill out the form and predicted function for at least three internal structures.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Next, you&#39;ll observe the outer and inner structures of the crayfish. Begin with the crayfish on its back and observe the appendages.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">In Table 1, take notes about the structures you notice.
</div></li><li data-sub-note-item="1-22"><div class="lab-step">Observe the tail. Devise a hypothesis for its jointed structure. Consider how a crayfish moves in its environment.
</div></li><li data-sub-note-item="1-23"><div class="lab-step">Next, with the crayfish still on its back, look for the swimmerets under its tail.  NOTE: If the crayfish is male, the swimmerets nearest the legs are long extending between the legs. These are specialized reproductive structures that transfer sperm to the female. Females do not have these long structures, but instead have swimmerets that store developing embryos.
</div></li><li data-sub-note-item="1-24"><div class="lab-step">Consider the benefits of this strategy. Be sure to observe both a male and female crayfish, checking with neighboring students to identify these structures.
</div></li><li data-sub-note-item="1-25"><div class="lab-step">Next, observe the mouth parts of your crayfish using a probe.
</div></li><li data-sub-note-item="1-26"><div class="lab-step">Based on these structures, make a hypothesis about the size and type of prey you think crayfish consume.
</div></li><li data-sub-note-item="1-27"><div class="lab-step">Before beginning the dissection, be sure to fill in Table 1 with at least five external structures you observed in the crayfish along with their predicted function. With the organism lying legs down, use scissors to cut an incision in the exoskeleton from the tail to between the eyes. Take care to not disturb the organs underneath.
</div></li><li data-sub-note-item="1-28"><div class="lab-step">Next, remove the carapace as demonstrated in the video.
</div></li><li data-sub-note-item="1-29"><div class="lab-step">Open the tail and use the pins to secure the exoskeleton down to the dissecting tray exposing the organs.
</div></li><li data-sub-note-item="1-30"><div class="lab-step">Observe the organs first with the naked eye. Use the figure to identify the different body parts of the animal.
</div></li><li data-sub-note-item="1-31"><div class="lab-step">Then, if there is enough space on the stage, place the dissecting tray with the crayfish under the dissecting microscope to view these structures more closely.
</div></li><li data-sub-note-item="1-32"><div class="lab-step">Fill out at least three rows of Table 1 with these observations and predicted functions of internal structures in the crayfish.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To analyze the data, first separate into groups. Within your groups, compare observations from your tables.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Using these observations, create class predictions that relate the observed forms to the hypothesized functions for both the cricket and the crayfish. As a group, fill out Table 2 with at least five of these predictions and discuss how you would go about testing these hypotheses. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/27/Table_2_Animal_Diversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Compare the internal and external structures you observed in the cricket and crayfish. Fill out Table 3 with similarities and differences you noticed in the structures of these organisms. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/27/Table_3_Animal_Diversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li></ul></li></ol>