Funding for this project was provided by the Division of Intramural Research of the National Heart, Lung, and Blood Institute, National Institutes of Health.
We would like to thank Dr. James Hawkins for his mentorship and support. We also thank Drs. Mihai Oltean and Robert Linford for their assistance in locating the superior mesenteric artery. We would like to extend our thanks to Drs. Patricia Carvalho Obeid Ellrich, Claudio Correa Natalini, and George Howell III for providing their expertise during the development of this protocol. Finally, we would like to thank Stephen Wincovitch for his assistance in acquiring the beautiful photomicrographs featured in this paper and Dr. Alicia Olivier for her help labeling and rendering the final figures.

The procedures described here were approved by the National Heart, Lung, and Blood Institute Animal Care and Use Committee at the National Institutes of Health and conform to the policies outlined in The Public Health Service Policy on Humane Care and Use of Laboratory Animals, The Animal Welfare Act, and the Guide for the Care and Use of Laboratory Animals.
1. Surgical setup
<ol>
	<li>Follow aseptic procedures. Don a mask, hair cover, and clean jumpsuit/lab coat/surgical scrubs.</li>
	<li>Prepare the following sterilized materials: surgical instruments (see Table of Materials), warm saline, cotton swabs, gauze, surgical staples, surgical drapes, and gloves. Also obtain surgical tape, which does not need to be sterilized. Sterilize the materials with either autoclave or ethylene oxide sterilization techniques.</li>
	<li>Place a heated circulating water blanket in the surgery area and cover it with a sterile towel or drape.</li>
	<li>Use a precision isoflurane vaporizer, pressurized medical air (21% O2), and a Bain non-rebreathing circuit with a nose cone designed for mice to provide surgical anesthesia.</li>
</ol>
2. Animal preparation
<ol>
	<li>Anesthetize the mouse in an induction chamber by delivering 2%-4% isoflurane with 21% O2 medical air at a rate of 0.5 L/min for each liter of chamber volume. 
	NOTE: It is preferable to use 21% O2 medical air over 100% O2 for this particular model, as saturating the blood with O2 may interfere with the IRI.</li>
	<li>Remove the mouse from the chamber and move it to a clean surface separated from the surgery area. Fit it with a nose cone delivering 1.5% isoflurane with 21% O2 medical air.</li>
	<li>Inject 1 mg/kg buprenorphine subcutaneously into the dorsal cervicothoracic area.</li>
	<li>Inject 200-600 IU/kg heparin intraperitoneally to prevent thrombus formation during the period of occlusion.</li>
	<li>Apply ophthalmic ointment to the eyes to prevent corneal damage.</li>
	<li>Remove hair from the ventral abdomen using clippers.</li>
	<li>Move the mouse onto the heated water blanket in the surgery area. Again, fit it with a nose cone delivering 1.5% isoflurane with 21% O2 medical air to achieve a surgical plane of anesthesia.</li>
	<li>Position the mouse in dorsal recumbency and secure the limbs to the table with surgical tape.</li>
	<li>Monitor the animal&#39;s body temperature rectally using a rodent-specific thermometer. Keep the body temperature at 36.5 &#177; 0.5 &#176;C for the entire duration of the surgery.</li>
	<li>Disinfect the ventral abdomen using sterile gauze soaked in either chlorhexidine scrub or povidone-iodine scrub, followed by 70% alcohol. Repeat this sequence three times, alternating between the scrub and the alcohol. A new set of scrubs and alcohol gauzes should be used each time.
	<ol>
		<li>Apply the scrub and alcohol in a circular motion, starting with small circles at the center of the surgical site and gradually working toward the edges by increasing the size of the circles. Discard the gauze once the edge of the surgical site has been reached. Do not scrub backward from edge to center.</li>
	</ol>
	</li>
	<li>Perform a toe pinch test (pedal reflex) to ensure the animal is fully anesthetized.</li>
	<li>Don sterile gloves. Aseptically drape the surgical site.</li>
</ol>
3. Surgery and ischemia
<ol>
	<li>Make a 3-5 cm ventral midline abdominal incision in the skin using a #15 scalpel blade, dissect it free from the underlying muscle fascia, and reflect it laterally. Continue the incision through the abdominal wall along the linea alba using either micro dissecting scissors or spring-loaded micro scissors and place a retractor in position.</li>
	<li>Place sterile gauze pads moistened with warm sterile saline around the operating area.</li>
	<li>Remove the small intestine from the abdominal cavity, flip it cranially and to the animal&#39;s left, and place it on the moistened pads. Place another moistened gauze pad over the tissues to prevent desiccation. Periodically drip warm sterile saline on the gauze to keep the tissues moist.</li>
	<li>Isolate the SMA, which is located ventral to the inferior vena cava, caudal to the celiac artery, and cranial to the renal artery. 
	NOTE: Figure 1 shows the location of the SMA where it is isolated during surgery. The SMA normally lies ventral to the inferior vena cava and extends toward the right. When the intestines are exteriorized and flipped to the left during the surgery, the SMA lies to the left of the inferior vena cava.</li>
	<li>Place an atraumatic microvascular clip across the base of the SMA where it branches off the abdominal aorta, ensuring the clip does not occlude the superior mesenteric vein.</li>
	<li>Verify ischemia of the small intestine by noting the color change from pink to pale-white and the loss of mesenteric pulsation.</li>
	<li>Return the viscera to its original position inside the abdominal cavity for the duration of the ischemic period. Remove the retractor and cover the incision with moist gauze. Periodically add warm sterile saline to the gauze to prevent desiccation and maintain body temperature.</li>
	<li>After a 45 min period of ischemia (the beginning of which is marked by the initial application of the clip), remove the occluding clip. Verify the restoration of blood flow by observing a mesenteric pulsation and flushed color.</li>
	<li>Apply warm sterile saline intraperitoneally just prior to final closure to maintain appropriate hydration.</li>
	<li>Close the abdominal muscles with a 6-0 polyglactin 910 suture. Administer bupivacaine (up to 2 mg/kg) along the muscle incision line for pain relief. Close the skin with surgical staples or wound clips.</li>
</ol>
4. Recovery and reperfusion
<ol>
	<li>Return the mouse to a warm chamber or cage on a circulating water blanket, hand warmer, or other appropriate heat source. Deliver 21% O2 at a flow rate of 0.5 L/min for each liter of chamber volume. Let the mouse recover here for 90 min. Monitor the mouse every 5-10 min for signs of pain or distress, such as hunched posture, squinting, and reluctance to move.</li>
</ol>
5. Euthanasia and blood collection
<ol>
	<li>At the end of the 90 min recovery period, return the mouse to the induction chamber and deliver 2%-4% isoflurane with 21% O2 at a rate of 0.5 L/min of chamber volume to re-induce full anesthesia.</li>
	<li>Transfer the animal back to the surgery area and fit it with a nose cone delivering 2%-4% isoflurane with 21% O2 to achieve deep anesthesia. 
	NOTE: CO2 is not an appropriate method of euthanasia for this procedure, as it induces physiologic changes which may interfere with ischemic injury or tissue analytes13.</li>
	<li>Re-open the ventral midline incision and perform a terminal bleed by collecting as much blood as possible from the abdominal vena cava using a 23 G needle and syringe. Expect to collect between 0.3-0.5 mL of blood (less in mice who have undergone IRI, more in those who received sham laparotomy). 
	NOTE: The purpose of the terminal bleed is to aid in humane euthanasia and to collect and preserve blood for future testing (i.e., serum chemistry, PCR, ELISA).</li>
	<li>Following blood collection, the abdominal aorta is severed to allow for complete exsanguination.</li>
	<li>Perform either cervical dislocation or thoracotomy as a secondary measure to ensure successful euthanasia.</li>
</ol>
6. Tissue processing for histology
<ol>
	<li>After euthanasia, collect the desired tissues. Ensure that the tissue processing is done promptly, as autolysis begins immediately after death14,15.
	<ol>
		<li>Intestines: Collect the entire length of the small intestine and large intestine. Discard the cecum.</li>
		<li>Liver: Collect the left lateral, left median, and right median&#160;lobes.</li>
		<li>Kidneys: Collect both kidneys. By convention, the left kidney is cut longitudinally, and the right is cut as a cross section at the time of necropsy. 
		NOTE: The colon, liver, and kidneys may be used for assessing multi-organ failure or other systemic effects of IRI. The small intestines are used to assess the primary injury. It is not necessary to keep track of individual sections of the liver lobe and kidneys, as each organ will be analyzed and scored as one unit. The intestinal segments, however, should be kept separate and then labeled and scored individually.</li>
	</ol>
	</li>
	<li>Divide the intestine into four sections: duodenum, jejunum, ileum, and colon. Ensure that the three small intestinal segments are equal in length. Do this by folding the small intestines into a &#34;Z&#34; shape, where the top line is the duodenum, the middle line is the jejunum, and the bottom line is the ileum. It is important to keep track of the proximal versus the distal end.</li>
	<li>Flush the lumen of the intestinal segments with saline using a 10 mL syringe affixed with a 20 G angio-catheter.</li>
	<li>Prior to making sections, lay each intestinal segment flat with the luminal side facing up. Using a 3 mL syringe affixed with a 27 G needle and generously apply 10% buffered formalin dropwise to coat the entire length of the mucosa. Then, roll each intestinal segment individually and place in separate, labeled tissue cassettes.
	<ol>
		<li>To roll, lay each segment flat with the luminal side facing up, then roll circumferentially around a toothpick. The proximal portion should form the inner part of the roll. The lumen should face the inside/center. Try to roll as gently as possible to avoid compressing the villi.</li>
		<li>When rolled, the intestine should look like a Swiss roll. Place the Swiss roll spiral face-up inside the cassette.</li>
	</ol>
	</li>
	<li>Place the tissues in labeled vials filled with 10% buffered formalin to fix them at room temperature. Over-fixing is better than under-fixing. The vials should be large with plenty of formalin-at least 20x more fixative than tissue.
	<ol>
		<li>Intestines: Place the four cassettes together in a specimen cup. Fix for 24-48 h.</li>
		<li>Liver: Place the liver lobes together in a 50 mL conical tube. Fix for 24-48 h.</li>
		<li>Kidneys: Place the kidneys together in a 50 mL conical tube. Fix for 48-72 h. 
		NOTE: Untrimmed kidneys take longer to fix than trimmed kidneys. To shorten the fixation time to 24-48 h, the kidneys may be cut along the median plane, longitudinally (left kidney) and transversely (right kidney), and placed in cassettes before being deposited in the formalin.</li>
	</ol>
	</li>
	<li>After the tissues have been fixed in formalin for the designated time, rinse with phosphate-buffered saline (PBS) or distilled water and transfer to labeled vials filled with 70% EtOH. The tissue can be stored in the EtOH indefinitely at room temperature while awaiting histology.
	<ol>
		<li>Intestines: Place the four cassettes together in a specimen cup.</li>
		<li>Liver: Place the liver lobes together in a 50 mL conical tube.</li>
		<li>Kidneys: Place the kidneys together in a 50 mL conical tube.</li>
	</ol>
	</li>
	<li>When ready, have the tissues processed onto glass slides using hematoxylin and eosin (H&#38;E) staining. Trim the formalin-fixed tissues and then embed them in paraffin. Mount five-micron sections on the slides and stain with H&#38;E.</li>
</ol>
7. Tissue scoring
<ol>
	<li>Tissue scoring should preferably be performed by experienced personnel who are blinded to the sample groups.</li>
	<li>Intestinal ischemia is scored using the Chiu/Park scoring system17.</li>
	<li>Kidney damage is scored using the Jablonski scoring system18,19.</li>
	<li>Liver damage is scored using the Suzuki scoring system20,21. 
	NOTE: There are many scoring systems currently in use for assessing tissue damage in rodent models of intestinal IRI. The scoring systems used in this study were selected to minimize arbitrary estimation and to maximize intentional qualitative assessment (Table 1).</li>
</ol>

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Renal Physiology. 300 (1), 167-176 (2011).</li><li>Megison, S. M., Horton, J. W., Chao, H., Walker, P. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+model+for+intestinal+ischemia+in+the+rat.">A new model for intestinal ischemia in the rat.</a> The Journal of Surgical Research. 49 (2), 168-173 (1990).</li><li>Gubernatorova, E. O., Perez-Chanona, E., Koroleva, E. P., Jobin, C., Tumanov, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+model+of+intestinal+ischemia-reperfusion+injury.">Murine model of intestinal ischemia-reperfusion injury.</a> Journal of Visualized Experiments. (111), 53881 (2016).</li><li>Trompeter, M., Brazda, T., Remy, C. T., Vestring, T., Reimer, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-occlusive+mesenteric+ischemia:+Etiology,+diagnosis,+and+interventional+therapy.">Non-occlusive mesenteric ischemia: Etiology, diagnosis, and interventional therapy.</a> European Radiology. 12 (5), 1179-1187 (2002).</li><li>Kozar, R. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Superior+mesenteric+artery+occlusion+models+shock-induced+gut+ischemia-reperfusion.">Superior mesenteric artery occlusion models shock-induced gut ischemia-reperfusion.</a> The Journal of Surgical Research. 116 (1), 145-150 (2004).</li></ol>

The authors of this paper have no conflicts of interest to disclose.

Despite the widespread use of this intestinal IRI model, it is not without its limitations. For instance, sole occlusion of just the base of the SMA does not completely obstruct blood flow to the intestine. This is likely due to extensive collateral circulation in the mesentery, which can draw blood from neighboring branches of the abdominal aorta. In one study in cats, SMA occlusion decreased blood flow by 35% in the proximal duodenum, 61% in the distal duodenum, 71% in the jejunum and ileum, and 63% in the proximal colon. Blood flow was not reduced in the mid and distal colon, which receive much of their circulation from the inferior mesenteric artery23. In rodents, the jejunum and ileum are most often cited as the intestinal segments which incur the most significant tissue injury following SMA occlusion9.
A wide range of ischemia times after SMA occlusion have been cited in the literature, from 1 to 90 min or more. Different ischemic times result in different levels of reperfusion injury; Park et al. observed reperfusion injury when the ischemic interval was between 40 and 60 min, but not when the ischemic interval was shorter or longer24. Such results suggest that shorter times do not produce enough ischemia to incite reperfusion injury, while longer times damage the tissue so severely that it is impossible to demonstrate the reperfusion injury that follows. In addition, longer ischemic times carry the risk of increased mortality. As seen in our study, 50% (3/6) of the initial mice that underwent 60 min ischemia died after only 90 min of reperfusion. Shortening the ischemia time to 45 min lowered the mortality to 20% (1/5) without changing the tissue injury scores. Based on our study, it appears that the ideal window of ischemic damage can be attained by SMA occlusion for about 45 min.
Another variable is the reperfusion time before tissue collection. As with ischemia times, reperfusion times vary widely across studies, from 60 min to over 24 h. Several papers have reported that the intestinal mucosa incurs maximal morphologic damage at 2 to 3 h of reperfusion, with complete repair achieved at 24 h25,26,27. Collecting tissue before this 2 to 3 h window risks not capturing the full extent of the reperfusion injury, while tissues harvested closer to 24 h will have already started the repair process. We initially opted for a reperfusion time of 120 min, but then changed to 90 min in an effort to lower mortality. This change did not change tissue injury results, suggesting that a 30 min deviation from the 2 to 3 h window is acceptable.
Oxygen concentration is also an important variable in the development of IRI. Wilding et al. found that, compared to mice receiving 21% O2, those anesthetized with isoflurane delivered with 100% O2 experienced ventilation-perfusion mismatch due to atelectasis. In the same study, rats receiving 100% O2 developed acute respiratory acidosis and elevated mean arterial pressure28. Such physiologic changes are best avoided when inducing an injury such as IRI, in which a number of systemic factors are involved. Thus, 21% O2 seems to be more appropriate than 100% O2 as the carrier gas for isoflurane delivery.
The use of heparin in this protocol is open to debate. Heparin is known to have anti-coagulative and anti-inflammatory effects29. We found that changing from 60 min ischemia and 120 min reperfusion to 45 min ischemia and 90 min reperfusion with 400 IU/kg heparin did not change microscopic intestinal injury but did lower mortality. One possible explanation is that heparin prevented fatal thromboembolism to distant organs such as the lungs and brain, however we did not find evidence of this on necropsy by gross or microscopic examination of the initial two mice that died. Using shorter ischemia and reperfusion times without heparin may be just as effective at reducing mortality. If that were the case, it would be prudent to forego the use of heparin to minimize interference with IRI. However, including heparin in the protocol may be appropriate for those wishing to model surgical causes of IRI, as surgical patients often receive heparin peri-operatively.
Isoflurane has been shown to have tissue protective effects in cases of intestinal inflammation and ischemia, and its use may interfere with a clinically relevant IRI model30,31,32. However, organofluorine inhalants (i.e., isoflurane, sevoflurane) are commonly used anesthetics in both veterinary and human medicine. In addition, the length of anesthesia required for this protocol exceeds 120 min, and thus an inhalant is more appropriate than a shorter-acting injectable which would need to be re-dosed.
No microscopic lesions were present in the proximal colon, liver, or kidney. The lack of microscopic changes was perhaps due to the relatively short 90 to 120 min reperfusion time. In addition, the proximal colon has a blood supply from the inferior mesenteric artery. However, a lack of visible damage does not rule out systemic injury. Reverse transcription-quantitative polymerase chain reaction (RT-qPCR) is likely a better methodology to demonstrate systemic injury by measuring inflammatory cytokines such as TNF-&#945;.
Several variations of this intestinal IRI model have been developed over the years. In 1990, Megison et al. demonstrated that occluding collateral vessels in addition to the SMA produced a more consistent reduction of mesenteric blood flow but an increase in the mortality rate33. A more recent study showed that, in lieu of occluding the SMA at its base, ligating its peripheral and collateral branches to induce ischemia in the distal ileum yielded reproducible injury without mortality34. Occlusion of the local arterial branches ensures maximal ischemia and may address the issue of multifocal, segmental reductions of blood flow seen with ligating the SMA just at its base. While this alternative method of modeling intestinal IRI has application for research into the local tissue effects of intestinal IRI, it is unknown whether it can accurately model the systemic inflammation and multi-organ failure which can be associated with intestinal injury.
SMA occlusion is not an appropriate model for all types of intestinal IRI. Non-occlusive mesenteric ischemia, for instance, is characterized by splanchnic hypoperfusion stemming from decreased cardiac output. Therefore, this technique would not be optimal to study intestinal IRI caused by myocardial infarction, congestive heart failure, aortic insufficiency, or renal or hepatic disease35. Kozar et al. reported that SMA occlusion is, however, a clinically relevant model for gut IRI induced by shock36. Although less economical, the use of other species such as pigs may have benefits over rodents for modeling certain intestinal injury conditions. A comprehensive review by Gonzalez et al. in 2014 describes animal models currently in use for investigating intestinal IRI9.
Despite its limitations, the technique of occluding the SMA at its base remains one of the most commonly utilized rodent models of intestinal ischemia9. As it only requires one vascular clamp and a basic setup, the surgery itself is quite simple. It also yields reproducible injury, as evidenced by the data presented here. SMA occlusion in rodents can reliably model occlusive causes of intestinal IRI and can have practical application in both veterinary and human medicine. As such, it is important that the procedures we have outlined here be carried out with consistency.

Ischemia-reperfusion injury (IRI) can occur in any organ and involves two sequential components. An initial cessation of blood flow causes affected tissues to become ischemic and then subsequent reperfusion induces further cell injury. Damage from the reperfusion often exceeds that caused by ischemia1. The pathophysiology of IRI involves a complex cascade of events, the most notable of which is free radical formation upon the reintroduction of oxygen, which occurs during reperfusion2. Activation of the inflammatory cells and cytokines also plays a role2. In cases of intestinal IRI, bacterial translocation into the bloodstream following endothelial damage can lead to systemic inflammatory response syndrome2. If the damage due to IRI is severe enough, resultant systemic effects can lead to multi-organ failure3.
Cases of intestinal IRI are associated with high morbidity and mortality4,5,6.&#160;Intestinal IRI is associated with many pathologic conditions and surgical procedures in both veterinary and human medicine. In veterinary medicine, animals are especially prone to intestinal IRI conditions, such as gastric dilatation volvulus (GDV), mesenteric torsion, and colic7,8. In humans, IRI is a major and frequently occurring problem in abdominal aortic aneurysm surgery, strangulated hernias, acute mesenteric ischemia, volvulus, trauma, shock, neonatal necrotizing enterocolitis, and small bowel resection or transplantation9.
Most in vivo rodent studies of intestinal IRI involve occlusion of the base of the superior mesenteric artery (SMA), the branch of the abdominal aorta that supplies blood to the majority of the small intestines and the proximal portion of the large intestines10,11,12. Despite this model&#39;s widespread use and relative simplicity, a detailed protocol using inhalant anesthesia in 21% O2 medical air has not been published. The lack of a standard protocol poses difficulty for researchers who are unfamiliar with the procedure and prevents consistency across studies. We demonstrate the steps necessary to conduct the surgical model of intestinal IRI in 8-14-week-old male and female Swiss Webster mice. This model of intestinal IRI yields reproducible injury, as demonstrated by consistent histopathology.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Adson forceps</td>
			<td>Roboz</td>
			<td>RS-5236</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Alm retractor</td>
			<td>Roboz</td>
			<td>RS-6510</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Anesthesia machine</td>
			<td>Datex-Ohmeda</td>
			<td>Aestiva 5</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia: isoflurane</td>
			<td>Baxter Healthcare Corporation</td>
			<td>NDC 10019-360-40</td>
			<td>Dose: 1-4%, INH</td>
		</tr>
		<tr>
			<td>Angiocath 20 g&#160; x 2</td>
			<td>Smiths Medical</td>
			<td>5057</td>
			<td>Flushing intestines with saline and formalin</td>
		</tr>
		<tr>
			<td>Atraumatic microvascular clip</td>
			<td>Teleflex</td>
			<td>065100</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Buffered formalin 10%</td>
			<td>Fisher Scientific</td>
			<td>23-245684</td>
			<td>Tissue fixation</td>
		</tr>
		<tr>
			<td>Bupivicaine 0.25%</td>
			<td>Hospira, Inc.</td>
			<td>NDC 0409-1160-18</td>
			<td>Dose: up to 2 mg/kg drop-wise</td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Par Pharmaceutical</td>
			<td>NDC 42023-179-05</td>
			<td>Dose: 1 mg/kg, SQ</td>
		</tr>
		<tr>
			<td>Chlorhexidine scrub 2%</td>
			<td>Vet One</td>
			<td>510083</td>
			<td>Surgical site prep</td>
		</tr>
		<tr>
			<td>Circulating water blanket</td>
			<td>Cincinnati Sub Zero</td>
			<td>Blanketrol 2</td>
			<td>Body temp maintenance</td>
		</tr>
		<tr>
			<td>Clippers - Wahl BravMini, Purple Hair clippers</td>
			<td>Lambert Vet Supply</td>
			<td>008WA-41590-0438</td>
			<td>Surgical site prep</td>
		</tr>
		<tr>
			<td>Conical tubes 50 ml</td>
			<td>Fisher Scientific</td>
			<td>14-432-22</td>
			<td>Tissue fixation and storage</td>
		</tr>
		<tr>
			<td>Dry ice</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>PCR tissue samples</td>
		</tr>
		<tr>
			<td>EtOH 200 proof</td>
			<td>The Warner-Graham Company</td>
			<td>64-17-5</td>
			<td>Tissue storage</td>
		</tr>
		<tr>
			<td>Heparin (optional)</td>
			<td>Meitheal Pharmaceuticals</td>
			<td>NDC 71288-402-11</td>
			<td>Dose: 200-600 IU/kg</td>
		</tr>
		<tr>
			<td>Induction chamber</td>
			<td>VetEquip</td>
			<td>941456</td>
			<td></td>
		</tr>
		<tr>
			<td>Indus Instruments THM100 Rodent Monitor</td>
			<td>Indus Instruments</td>
			<td>N/A</td>
			<td>For monitoring rodent body temperature during surgery</td>
		</tr>
		<tr>
			<td>Isopropyl Alcohol 70%</td>
			<td>Humco</td>
			<td>NDC 0395-4202-28</td>
			<td>For scrubbing surgical site</td>
		</tr>
		<tr>
			<td>Microcentrifuge Tubes: 0.6mL</td>
			<td>Fisher Scientific</td>
			<td>05-408-121</td>
			<td>PCR tissue samples. 8 per mouse, Terminal bleed collection, serum storage</td>
		</tr>
		<tr>
			<td>Microsoft Excel</td>
			<td>Microsoft</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Nose cone</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Can be homemade with syringe tube or bubble tubing</td>
		</tr>
		<tr>
			<td>O2 medical air 21%</td>
			<td>Roberts Oxygen</td>
			<td>N/A</td>
			<td>Rate: 0.5 L/min for each L chamber volume</td>
		</tr>
		<tr>
			<td>Ophthalmic ointment</td>
			<td>Akorn, Inc.</td>
			<td>NDC 17478-062-35</td>
			<td>Surgical prep</td>
		</tr>
		<tr>
			<td>PBS pH 7.4 (1x)</td>
			<td>ThermoFisher Scientific</td>
			<td>10010-031</td>
			<td>For tissue rinsing and making 70% EtOH</td>
		</tr>
		<tr>
			<td>Specimen cups</td>
			<td>Cardinal Healthcare</td>
			<td>C13005</td>
			<td>For holding tissue cassettes in formalin</td>
		</tr>
		<tr>
			<td>Sterile Castroviejo Needle Holder</td>
			<td>Roboz</td>
			<td>RS-6412</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile cotton swabs</td>
			<td>Medline</td>
			<td>BXTA50002Z</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile gauze</td>
			<td>Medline</td>
			<td>PRM21423Z</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile Micro Dissecting Scissors</td>
			<td>Roboz</td>
			<td>RS-5980</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile micro dissecting spring scissors</td>
			<td>Roboz</td>
			<td>RS-5693</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile micro forceps</td>
			<td>Roboz</td>
			<td>RS-5264</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile saline (0.9%)</td>
			<td>Braun</td>
			<td>R5201-01</td>
			<td>Must be warmed</td>
		</tr>
		<tr>
			<td>Sterile scalpel blade #15</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>32295-015</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile scalpel handle</td>
			<td>Roboz</td>
			<td>RS-9843</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile surgical drape</td>
			<td>Medline</td>
			<td>DYNJSD1092</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile surgical gloves</td>
			<td>Medline</td>
			<td>MSG2270</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile surgical stapler</td>
			<td>Roboz</td>
			<td>RS-9260</td>
			<td>Surgical instrument</td>
		</tr>
		<tr>
			<td>Sterile surgical staples</td>
			<td>Roboz</td>
			<td>RS-9262</td>
			<td>Abdominal skin closure</td>
		</tr>
		<tr>
			<td>Sterile suture: Vicryl (polyglactin 910) 6-0, 27&#34; Taper RB-1 Needle</td>
			<td>Ethicon</td>
			<td>J212G</td>
			<td>Closing abdominal muscle</td>
		</tr>
		<tr>
			<td>Surgical tape</td>
			<td>Medline</td>
			<td>MMM15271Z</td>
			<td>Securing mouse in dorsal recumbancy</td>
		</tr>
		<tr>
			<td>Syringe 10 ml x 2</td>
			<td>Medline</td>
			<td>SYR110010</td>
			<td>Flushing intestines with saline and formalin</td>
		</tr>
		<tr>
			<td>Tissue cassettes</td>
			<td>Fisher Scientific</td>
			<td>22-038-665</td>
			<td>Rolled intestinal segments. 4 per mouse.</td>
		</tr>
		<tr>
			<td>Towel or drape</td>
			<td>Medline</td>
			<td>GEM2140</td>
			<td>Water blanket cover</td>
		</tr>
	</tbody>
</table>

rodent model of intestinal ischemia-reperfusion injury via occlusion of the superior mesenteric artery

Intestinal ischemia-reperfusion injury (IRI) is associated with a myriad of conditions in both veterinary and human medicine. Intestinal IRI conditions, such as gastric dilatation volvulus (GDV), mesenteric torsion, and colic, are observed in animals such as dogs and horses. An initial interruption of blood flow causes tissues to become ischemic. Although necessary to salvage viable tissue, subsequent reperfusion can induce further injury. The main mechanism responsible for IRI is free radical formation upon reperfusion and reintroduction of oxygen into damaged tissue, but there are many other components involved. The resulting local and systemic effects often impart a poor prognosis. 
Intestinal IRI has been the subject of extensive research over the past 50 years. An in vivo rodent model in which the base of the superior mesenteric artery (SMA) is temporarily ligated is currently the most common method used to study intestinal IRI. Here, we describe a model of intestinal IRI utilizing isoflurane anesthesia in 21% O2 medical air that yields reproducible injury, as demonstrated by consistent histopathology of the small intestines. Tissue injury was also assessed in the colon, liver, and kidneys.

We demonstrated a model of intestinal IRI in mice that yielded consistent and reproducible results. The small intestine, proximal colon, kidneys, and liver were sectioned and stained with H&#38;E. A veterinary pathologist graded tissue injury using the scoring systems previously mentioned (Table 1). Statistical analysis was performed using single factor analysis of variance (ANOVA) followed by Tukey&#39;s post hoc with pairwise comparisons, which determined whether or not there was a significant difference in the data within and across groups. A p-value less than or equal to 0.05 was considered the cutoff for establishing statistical significance. All statistical tests and graphing were carried out in a spreadsheet software (e.g., Microsoft Excel) with the Real Statistics Resource Pack add-on. Data are presented as the mean &#177; standard error of the mean (SEM).
Microscopic lesion scores of the three small intestinal segments (duodenum, jejunum, and ileum) were significantly increased for animals undergoing intestinal ischemia-reperfusion injury (IRI; N = 7) versus those that underwent sham laparotomy (Sham; N = 6) (Figure 2 and Figure 3). The standard error for these data was narrow, demonstrating consistency of the results within and across groups. Each intestinal segment in the Sham group yielded the exact same average Park/Chiu score of 0.83. The SEM for the duodenum, jejunum, and ileum in the Sham group was 0.31, 0.40, and 0.31, respectively. The average Park/Chiu scores for the duodenum, jejunum, and ileum in the IRI group were 4.07 &#177; 0.44, 4.14 &#177; 0.46, and 5.14 &#177; 0.40, respectively.
In this study, 50% (3/6) of the initial mice that underwent 60 min ischemia and 120 min reperfusion (60/120 group) died. Two of the three mice were submitted for necropsy. Both mice had epithelial necrosis, congestion, and hemorrhage of the small intestine. In addition, the mice had lymphocytolysis, a nonspecific change associated with physiologic stress. Neither mouse had lesions in the heart, lung, liver, or kidneys. Shortening the times to 45 min ischemia and 90 min reperfusion and adding in 400 IU/kg heparin (45/90/H group) lowered the mortality to 20% (1/5) without changing the intestinal injury scores (Figure 4). The mean Park/Chiu score for the 60/120 group was 4.56 &#177; 0.38 (N = 3), and the mean score for the 45/90/H group was 4.375 &#177; 0.38 (N = 4).
Microscopic findings indicative of injury in the proximal colon, liver, and kidney were not seen in either the 60/120 mice or the 45/90/H mice.
Table 1: Scoring systems for the intestines, kidneys, and liver. Intestinal damage was graded using the Chiu/Park system17. Kidney damage was graded using the Jablonski scoring system18,19. Liver damage was graded using the Suzuki scoring system20,21. This table is adapted with permissions from scoring systems presented in Quaedackers et al.17, Du et al.19, and Behrends et al.21. <a href="https://www.jove.com/files/ftp_upload/64314/Table 1 - Scoring Systems.zip" target="_blank">Please click here to download this Table.</a>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64314/64314fig01.jpg" /> 
Figure 1: Location and isolation of the superior mesenteric artery (SMA). (A) Normally, the SMA lies ventral to the inferior vena cava and extends toward the animal&#39;s right. It is situated between the celiac artery and the renal artery. This figure is adapted with permission from The Anatomy of the Laboratory Mouse&#160;by Margaret Cook (1965)22. (B) In this procedure, the intestines are exteriorized and flipped to the left (covered with moistened gauze in this picture), so the SMA (yellow arrow) lies to the left of the inferior vena cava (blue arrow). Abbreviations: RK = right kidney; D = duodenum. <a href="https://www.jove.com/files/ftp_upload/64314/64314fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64314/64314fig02.jpg" /> 
Figure 2: Small intestinal segments stained with hematoxylin and eosin. Sections of jejunum (A) and ileum (B) from mice in the Sham group featured villi that were long and thin without distortion. Sections of jejunum (C) and ileum (D) from mice in the IRI group featured areas of necrosis (asterisks) and hemorrhage with blunting and distortion of the remaining villi (arrows). The photos are from mice that underwent 45 min ischemia and 90 min reperfusion and received 400 IU/kg heparin. The photos were taken at 20x magnification with 10% zoom. Scale bar = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/64314/64314fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64314/64314fig03.jpg" /> 
Figure 3: Park/Chiu scores for small intestinal segments. Microscopic damage to all three intestinal segments (duodenum, jejunum, and ileum) for animals undergoing intestinal ischemia-reperfusion injury (IRI) was significantly increased compared to those that underwent sham laparotomy (Sham). * p&#160;&#60; 0.05 for IRI versus Sham. <a href="https://www.jove.com/files/ftp_upload/64314/64314fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64314/64314fig04.jpg" /> 
Figure 4: Park/Chiu scores for small intestines undergoing 60 min ischemia and 120 reperfusion versus 45 min ischemia and 90 min reperfusion with 400 IU/kg heparin. Decreasing the times from 60 min ischemia and 120 min reperfusion (60/120) to 45 min ischemia and 90 min reperfusion with 400 IU/kg heparin (45/90/H) did not create a statistically significant difference in Park/Chiu injury scores of the small intestines of mice in the IRI group. It did, however, reduce mortality from 50% to 20%. <a href="https://www.jove.com/files/ftp_upload/64314/64314fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Rodent Model of Intestinal Ischemia-Reperfusion Injury via Occlusion of the Superior Mesenteric Artery at JoVE.com

Rodent Model of Intestinal Ischemia-Reperfusion Injury via Occlusion of the Superior Mesenteric Artery

We describe how to generate a widely used surgical model of intestinal ischemia-reperfusion injury (IRI) in rodents. The procedure involves occlusion of the superior mesenteric artery followed by the restoration of blood flow. This model is useful for studies investigating occlusive causes of intestinal IRI in both veterinary and human medicine.

Intestinal ischemia-reperfusion injury (IRI) is associated with a myriad of conditions in both veterinary and human medicine. Intestinal IRI conditions, ...

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1.1K Views.  National Institutes of Health. We describe how to generate a widely used surgical model of intestinal ischemia-reperfusion injury (IRI) in rodents. The procedure involves occlusion of the superior mesenteric artery followed by the restoration of blood flow. This model is useful for studies investigating occlusive causes of intestinal IRI in both veterinary and human medicine.

Video: Rodent Model of Intestinal Ischemia-Reperfusion Injury via Occlusion of the Superior Mesenteric Artery

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

A powerful model for perioperative and critical care related acute kidney injury is presented. Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

A Murine Model of Myocardial Ischemia-reperfusion Injury through Ligation of the Left Anterior Descending Artery

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms at work during MI and developing effective therapeutic approaches requires methodology to reproducibly simulate the clinical incidence and reflect the pathophysiological changes associated with MI. Here, we describe a surgical method to induce MI in mouse models that can be used for short-term ischemia-reperfusion (I/R) injury as well as permanent ligation. The major advantage of this method is to facilitate location of the left anterior descending artery (LAD) to allow for accurate ligation of this artery to induce ischemia in the left ventricle of the mouse heart. Accurate positioning of the ligature on the LAD increases reproducibility of infarct size and thus produces more reliable results. Greater precision in placement of the ligature will improve the standard surgical approaches to simulate MI in mice, thus reducing the number of experimental animals necessary for statistically relevant studies and improving our understanding of the mechanisms producing cardiac dysfunction following MI. This mouse model of MI is also useful for the preclinical testing of treatments targeting myocardial damage following MI.

We introduce a surgical method to induce experimental ischemia/reperfusion (I/R) injury to simulate myocardial infarction (MI) in mouse models that allows for more clarity in positioning of the ligation on the left anterior descending artery (LAD) to increase the reproducibility of MI experiments in mice.

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms ...

Murine Model of Intestinal Ischemia-reperfusion Injury

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, hypotension, necrotizing enterocolitis, bowel transplantation, trauma and chronic inflammation. Intestinal ischemia-reperfusion (IR) injury is a consequence of acute mesenteric ischemia, caused by inadequate blood flow through the mesenteric vessels, resulting in intestinal damage. Reperfusion following ischemia can further exacerbate damage of the intestine. The mechanisms of IR injury are complex and poorly understood. Therefore, experimental small animal models are critical for understanding the pathophysiology of IR injury and the development of novel therapies.
Here we describe a mouse model of acute intestinal IR injury that provides reproducible injury of the small intestine without mortality. This is achieved by inducing ischemia in the region of the distal ileum by temporally occluding the peripheral and terminal collateral branches of the superior mesenteric artery for 60 min using microvascular clips. Reperfusion for 1 hr, or 2 hr after injury results in reproducible injury of the intestine examined by histological analysis. Proper position of the microvascular clips is critical for the procedure. Therefore the video clip provides a detailed visual step-by-step description of this technique. This model of intestinal IR injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Here we describe the detailed procedure of intestinal ischemia-reperfusion in mice which results in reproducible injury without mortality to encourage the standardization of this technique across the field. This model of intestinal ischemia-reperfusion injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, ...

A Mouse Model of Retinal Ischemia-Reperfusion Injury Through Elevation of Intraocular Pressure

Retinal ischemia-reperfusion (I/R) is a pathophysiological process contributing to cellular damage in multiple ocular conditions, including glaucoma, diabetic retinopathy, and retinal vascular occlusions. Rodent models of I/R injury are providing significant insights into mechanisms and treatment strategies for human I/R injury, especially with regard to neurodegenerative damage in the retinal neurovascular unit. Presented here is a protocol for inducing retinal I/R injury in mice through elevation of intraocular pressure (IOP). In this protocol, the ocular anterior chamber is cannulated with a needle, through which flows the drip of an elevated saline reservoir. Using this drip to raise IOP above systolic arterial blood pressure, a practitioner temporarily halts inner retinal blood flow (ischemia). When circulation is reinstated (reperfusion) by removal of the cannula, severe cellular damage ensues, resulting ultimately in retinal neurodegeneration. Recent studies demonstrate inflammation, vascular permeability, and capillary degeneration as additional elements of this model. Compared to alternative retinal I/R methodologies, such as retinal arterial ligation, retinal I/R injury by elevated IOP offers advantages in its&#160;anatomical specificity, experimental tractability, and technical accessibility, presenting itself as a valuable tool for examining neuronal pathogenesis and therapy in the retinal neurovascular unit.

This article describes a procedure for inducing retinal ischemia-reperfusion injury by elevated intraocular pressure in mice. Retinal ischemia-reperfusion injury by elevated intraocular pressure serves to model human pathologies characterized by compromised oxygen and nutrient delivery in the retina, enabling researchers to examine potential cellular mechanisms and treatments for human diseases of the retinal neurovascular unit.

Retinal ischemia-reperfusion (I/R) is a pathophysiological process contributing to cellular damage in multiple ocular conditions, including glaucoma, ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

Two-vessel Occlusion Mouse Model of Cerebral Ischemia-reperfusion

In this study, a middle cerebral artery (MCA) occlusion mouse model is employed to study cerebral ischemia-reperfusion. A reproducible and reliable mouse model is useful for investigating the pathophysiology of cerebral ischemia-reperfusion and determining potential therapeutic strategies for patients with stroke. Variations in the anatomy of the circle of Willis of C57BL/6 mice affects their infarct volume after cerebral-ischemia-induced injury. Studies have indicated that distal MCA occlusion (MCAO) can overcome this problem and result in a stable infarct size. In this study, we establish a two-vessel occlusion mouse model of cerebral ischemia-reperfusion through the interruption of the blood flow to the right MCA. We distally ligate the right MCA and right common carotid artery (CCA) and restore blood flow after a certain period of ischemia. This ischemia-reperfusion injury induces an infarct of stable size and a behavioral deficit. Peripheral immune cells infiltrate the ischemic brain within the 24 h infiltration period. Additionally, the neuronal loss in the cortical area is less for a longer reperfusion duration. Therefore, this two-vessel occlusion model is suitable for investigating the immune response and neuronal recovery during the reperfusion period after cerebral ischemia.

A mouse model of cerebral ischemia-reperfusion is established to investigate the pathophysiology of stroke. We distally ligate the right middle cerebral artery and right common carotid artery and restore blood flow after 10 or 40 min of ischemia.

In this study, a middle cerebral artery (MCA) occlusion mouse model is employed to study cerebral ischemia-reperfusion. A reproducible and reliable mouse ...

Evaluation of Coronary Flow Reserve After Myocardial Ischemia Reperfusion in Rats

Coronary artery disease is the leading cause of death worldwide. After an acute myocardial infarction, early and successful myocardial intervention via recanalization of the coronary artery is the most effective strategy for reducing the size of ischemic myocardium. The coronary microvasculature cannot be visualized and imaged&#160;in vivo, but there are several invasive and noninvasive techniques that can be used to assess parameters which depend directly on coronary microvascular function. The endothelial function after ischemia reperfusion can be assessed also at the level of the coronary circulation via the coronary flow reserve (CFR).&#160;In this study, peak velocity of left anterior descending (LAD) coronary arteries was measured in rats in vivo via Transthoracic Doppler Echocardiography during resting and stress challenge (induced by Dobutamine). A normal heart can increase its coronary blood flow up to four times above the resting values during stress induction. Following ischemia reperfusion, we found a significantly diminished CFR, which can be used as a marker of coronary microvascular dysfunction. CFR has opened a window on the importance of microvascular dysfunction and has been shown to predict cardiovascular risk independent of whether the severe obstructive disease is present.

Coronary flow reserve (CFR), is defined as the ratio of maximal coronary blood flow to the resting coronary blood flow. We present a protocol for evaluating CFR in rats via ultrasound, which offers the opportunity to predict cardiovascular risk factors in the absence of obstructive coronary disease.

Coronary artery disease is the leading cause of death worldwide. After an acute myocardial infarction, early and successful myocardial intervention via ...

A Pre-clinical Rat Model for the Study of Ischemia-reperfusion Injury in Reconstructive Microsurgery

Ischemia-reperfusion injury is the main cause of flap failure in reconstructive microsurgery. The rat is the preferred preclinical animal model in many areas of biomedical research due to its cost-effectiveness and its translation to humans. This protocol describes a method to create a preclinical free skin flap model in rats with ischemia-reperfusion injury. The described 3 cm x 6 cm rat free skin flap model is easily obtained after the placement of several vascular ligatures and the section of the vascular pedicle. Then, 8 h after the ischemic insult and completion of the microsurgical anastomosis, the free skin flap develops the tissue damage. These ischemia-reperfusion injury-related damages can be studied in this model, making it a suitable model for evaluating therapeutic agents to address this pathophysiological process. Furthermore, two main monitoring techniques are described in the protocol for the assessment of this animal model: transit-time ultrasound technology and laser speckle contrast analysis.

Here, we describe a pre-clinical animal model for studying the pathophysiology of ischemia-reperfusion injury in reconstructive microsurgery. This free skin flap model based on the superficial caudal epigastric vessels in the rat may also allow for the evaluation of different therapies and compounds to counteract ischemia-reperfusion injury-related damage.

Ischemia-reperfusion injury is the main cause of flap failure in reconstructive microsurgery. The rat is the preferred preclinical animal model in many ...

Organ Ischemia-Reperfusion Injury by Simulating Hemodynamic Changes in Rat Liver Transplant Model

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, including ischemia-reperfusion injury (IRI) of extrahepatic organs. This model requires numerous experiments and devices. The duration of anhepatic phase is closely related to the time to develop IRI after transplantation. In this experiment, we used hemodynamic changes to induce extrahepatic organ damage in rats and determined the maximum tolerance time. The time until the most severe organ injury varied for different organs. This method can easily be replicated and can also be used to study IRI of the extrahepatic organs after liver transplantation.

This paper provides a detailed description of how to build an animal model of the anhepatic phase (liver ischemia) in rats to facilitate basic research into ischemia-reperfusion injury after liver transplantation.

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, ...