This work was funded by the Medical Research Council grant G1000299.
The corresponding author would like to thank Gary Borthwick, University of Edinburgh, for assisting during the surgery.
The authors would like to acknowledge advice from Helen Douglas and Iain Mackay and allowing us to observe their Deep Inferior Epigastric (DIEP) flap procedure (Canniesburn Plastic Surgery Unit, Glasgow Royal Infirmary, 84 Castle Street, Glasgow G4 0SF, UK).
The authors would also like to thank Gary Blackie at the University of Edinburgh for his help in producing the video for this article.

All surgery is performed in accordance with guidelines set out by the United Kingdom's Home Office and the University of Edinburgh's Veterinary Services Department.
1. Surgical Procedure Set-up Notes
<ol>
 <li>Change into clean surgical scrubs, gown, scrub cap and mask. Clean all surfaces of the operating room including equipment with 2% chlorhexidine in 70% isopropyl alcohol. </li>
 <li>Prior to surgery, autoclave all surgical supplies and instruments that will be used in the procedure. Sterile packs per operation should include: 3 drapes, gauze, cotton tip applicators, silicone sheeting and the surgical instruments, see the Table of Specific Surgical Materials and Tools and Figure 1. Weigh the rat and measure out the appropriate volume of buprenorphine (0.04 mg/kg) to be administered subcutaneously 1 hr before completing the procedure. Lay out; 3 x 1 ml syringes for subcutaneous fluid administration during surgery, 2 x 6-0 Vicryl sutures, 1 x 5-0 Ethilon sutures, a sterile marker with ruler and a 10-blade disposable scalpel, 4 - 5 pairs of sterile gloves and a hand-held cautery unit. </li>
 <li>Place 4 x 10 ml sterile, 0.9% saline vials in a water bath warmed to 37 &deg;C. This will be used for subcutaneous fluid replacement (1 ml/kg/hr) and to rinse the surgical site. Set up the homeothermic blanket, rectal probe, heat lamp, operating microscope and anesthetic rig. Turn on the laser and its software, set-up another anesthetic rig and place a heat pad underneath the black mat on which the animal will lay during the scan. </li>
 <li>Use male, Lewis rats weighing 250-300 g. House rats for 7 days with food and water ad libitum with 12 hr light-dark cycles before any surgery is performed. </li>
</ol>
2. Anesthesia and Skin Preparation
<ol>
 <li>Place the rat in the anesthetic rig's induction chamber and administer 4% isoflurane with 1L/min O2 for 2-3 min to induce anesthesia. Remove the anesthetized rat from the chamber and place it supine on the clean, heated mat. Maintain isoflurane at 1.5% using a nose cone. Apply lacrilube or similar agent to prevent corneal abrasion during the procedure. Perform a foot-pad pinch test to ensure the animal is adequately anesthetized before proceeding. Repeat this last test before each major step in the procedure and adjust the inhalational anesthetic concentration accordingly. </li>
 <li>Closely shave the anterior abdominal using an electric shaver so that the entire abdominal surface is exposed. Apply depilating cream for the duration recommended by the supplier. Remove the cream and thoroughly rinse the skin with warmed sterile saline to remove all traces of the cream. Apply 2% chlorhexidine in 70% isopropyl alcohol to the skin and allow to dry before proceeding. This is the standard skin preparation in our unit based on current evidence for surgical site infection.34 Please discuss with your Veterinary Department what is standard procedure in your unit before electing a skin preparation protocol. </li>
 <li>Place 2 drapes either side of the rat and take care to keep them sterile. Put on sterile gloves and with the aid of an assistant open up the sterile packs. Place all the instruments on one drape and the sutures, gauze, cotton applicators, silicone sheeting and sterile marker pen with ruler on the other.</li>
<li> Identify the midline using xiphisternum and the tail as reference points. Mark the midline. Measure 0.8 cm below the xiphisternum and mark this point. Draw a line perpendicular to the midline from this point. Taking the midline as the center of the flap mark out 1 cm and 2 cm to the left and the right of the midline. Draw vertical lines parallel to the midline from the points. Measure 4 cm below the original horizontal line and draw another parallel to it. Following these instructions a 4 x 4 cm flap divided into 4 equal strips is delineated (see Figure 2). </li>
</ol>
3. Laser Doppler Imaging
<ol>
 <li>Carefully move the rat to the nose cone of the second anesthetic rig at the LDI scanner. Continue anesthesia at 1.5% isoflurane, 1L/min O2. Turn on the laser and follow the manufacturer's instructions to start scanning. After saving the scanned file return the rat back to the first rig and re-insert the rectal probe of the homeothermic blanket using soft white paraffin as a lubricant. </li>
</ol>
4. In situ TRAM flap - Myocutaneous Model of IRI
<ol>
 <li>Re-scrub hands and put on fresh sterile gloves. Cut out a 5 cm diameter circle in the center of the remaining sterile drape and place this over the exposed abdomen to create a draped sterile-field. </li>
 <li>Make an incision down the left lateral marked edge (Figure 3A and B). Achieve hemostasis. Make similar incisions down the horizontal lines to the left of the midline. Achieve hemostasis. </li>
 <li>The fat overlying the left inferior rectus sheath should be visible. Using forceps and fine iris scissors carefully get underneath this fat. Take care not to damage the perforators coming through the left, anterior rectus sheath. The plane opened up by such dissection is that immediately above the anterior abdominal wall fascia. Continue dissecting in this plane around the demarcated margins. In the left iliac fossa lies the large, superficial circumflex iliac vessels- these can be tied or cauterized. Then extend the dissection medially with caution and only as far as the lateral margin of the left rectus muscle. There is an obvious color change at this point from pink to near white (Figure 3C). Carefully irrigate the area with sterile saline and check that hemostasis has been achieved before placing moist gauze over the area. </li>
 <li>Repeat the procedure on the contralateral side but this time extend to the linea alba (mid line). Take care to identify and cauterize all the musculocutaneous perforators arising in the center of the right rectus abdominis muscle (Figure 3D). If this is not done properly it can result in a postoperative hematoma and spurious results. Similarly achieve hemostasis, irrigate and place moist gauze over the raised flap. </li>
 <li>Return to the inferior margin of the left anterior rectus (Figure 3E and F). Cauterize the most inferior perforator seen. Proceed to cut a small (approx. 0.6 cm x 0.6 cm) window in the anterior rectus sheath using microscissors and pointed curved Graeffe forceps. Blunt dissect slowly down the lateral margin of the muscle until the muscle thins out but before the posterior sheath is breached. Then rotate the forceps and blunt dissect medially until the belly of the muscle is atop the curved edge of the forceps and the tips are free at the medial margin. Feed approximately 6 cm of 5-0 Ethilon into the jaws of the forceps and tie off the inferior rectus sheath. Upon completion of this step the myocutaneous flap is isolated on one dominant vessel- the deep superior epigastric vessels. Cover with a moist gauze. </li>
 <li>Cut the silicone sheeting into ovals with smooth corners, (Figure 3G). These should be large enough to cover most of the area exposed under the fasciocutaneous portions of the flap. However, caution must be taken to ensure that the skin edge can be closed without any tension and the medial curve of the oval may have to be paired back to prevent it impairing flow through the remaining perforators. These are then sutured in place with 6-0 Vicryl (Figure 3H). Cover with moist gauze.</li>
 <li>Using simple interrupted 5-0 Ethilon sutures 'peg out' the flap to reduce heat and water loss (Figure 3I). Cover with a moist gauze. </li>
 <li>Extend the wound superiorly to the left of the xiphisternum (Figure 3I). Suture this to the upper left quadrant to improve the field-of-view.</li>
 <li>Cut away any overlying fat to reveal the superior, left, anterior rectus sheath. Cut a small (0.6 cm x 0.6 cm) window in this sheath (Figure 3J). Extend the wound medially until a change in muscle fiber trajectory from vertical to oblique and consistency from tightly packed to loose fibrils is seen. </li>
 <li>Insert the curved forceps carefully between these two muscles and open up a plane by blunt dissection. Carefully cut down only as far as the superior surface of these curved forceps cutting through the belly of the left rectus abdominis muscle to reveal the underlying deep, superior epigastric artery and vein (Figure 3K).</li>
 <li>Using micro-instruments and high power on the operating microscope, carefully separate the artery and vein and strip off any surrounding fat.</li>
 <li>Apply atraumatic Acland clamps to the artery and vein (B-1, "V" type) and start the timer to count down the 30 min ischemic period. Irrigate the clamped pedicle and cover with gauze. We did not employ vessel dilators such as verapamil or pabavarine but should vessel spasm be a problem, these drugs should be considered.</li>
 <li>Administer the buprenorphine (0.04 mg/kg) and warmed, sterile saline (1 ml/kg/hr). </li>
 <li>Starting at the top left corner, suture the flap in place with 6-0 Vicryl subcuticular sutures stopping and tying off at the xiphisternum. </li>
 <li>When the 30 min ischemic time is over, carefully remove the clamps and irrigate the pedicle with warmed saline. Check that flow has been re-established. Please note that this ischaemic time was stipulated by the UK Home Office authority. Researchers working in other authorities may be able to extend this time. Extending the ischemic time will likely lead to worse clinical outcome. </li>
 <li>Suture the cut edges of the rectus back in place with 6-0 Vicryl. Take care not to apply too much tension as this can lead to kinking of the vessels. </li>
 <li>Complete the subcutaneous suturing taking care to bury all knots below the skin (Figure 3K).</li>
 <li>Clean the wound area and allow it to dry. Re-draw the zones on the flap.</li>
 <li>Re-scan the animal to obtain a postoperative image. </li>
 <li>Re-apply lacrilube to the animal's eyes and place in a warmed incubator (37 &deg;C) for 1 hr to recover before returning to the husbandry unit.
 </li>
</ol>
Critical steps within the protocol
The crux of the procedure is in identifying the deep, superior epigastric vessels. This is shown clearly in the accompanying film. In brief, a window is cut in the anterior rectus sheath to expose the underlying muscle fibers which will be running longitudinally. By extending the superficial dissection of the anterior rectus sheath medially a change in muscle fiber trajectory is observed from longitudinal to oblique. Insert blunt ended, curved, Graeffe forceps (or similar) at the intersection of these two bundles of muscle fibers. Blunt dissect laterally. Cut down, using micro scissors, onto the upper surface of the curved forceps held in this plane between the muscle fiber bundles. On removal of the Graeffe forceps the deep, superior epigastric artery and vein will be observed at the midpoint of the rectus abdominis muscle body. Strip off the fat overlying the vessels using micro instruments and apply the clamps.
The fasciocutaneous portions of the rat TRAM flap are thin enough to permit the flap to take as a full thickness skin graft. To prevent this and to ensure that this is a true model of IRI a thin, flexible silicon sheet is placed underneath the fasciocutaneous portions of the flap.35 This step has been adopted by other researchers undertaking rat TRAM models.17,21,25 
Rats chew through knots so make sure all sutures are subcuticular and all knots are buried. In carrying out meticulous suturing autocannabilism of flaps as reported by other researchers can be avoided. 24 
Following administration of Buprenorphine reduce maintenance anesthesia to 1% Isoflurane (1L/min O2).

We have no disclosures.

Modifications and trouble shooting
The protocol presented here reproduces the IRI seen in free tissue transfer in an experimental system enabling further understanding of that process and provides a means to investigate means of ameliorating IRI and improving outcome. This could easily be modified to produce a more severe injury if it were based on the non-dominant, deep, inferior epigastric pedicle or if the ischemic time were increased. 
Limitations of technique
 <p...

The goal of this protocol is to demonstrate a reliable and reproducible model of the ischemia-reperfusion injury observed in free tissue transfer to enable interventional strategies to be investigated. 
Free tissue transfer is defined as the vascular detachment of an isolated block of tissue followed by autologous transplant of that tissue with anastomosis of the flap's transected vessels to native vessels at the recipient site. The procedure is known as FTT and the tissue being transferred referred to as the free flap.
Free tissue transfer is the gold standard approach for the correction of complex, composite defects where local options are unsuitable or unavailable.1-4 Ischemia reperfusion injury (IRI) is inevitable in free tissue transfer, contributes to flap failure5,6 and has no effective treatment. The elective nature of free flap surgeries permits administration of pharmacological agents to precondition against IRI. 
IRI results in impaired flow through the microcirculation by endothelial activation and metabolic dysfunction,7 increased capillary permeability and subsequent interstitial edema7, influx of inflammatory cells,8 release of inflammatory mediators, reactive oxygen species9 and complement deposition.10 This complex process of hypoxia and subsequent reperfusion injury ultimately leads to cell death. A model of myocutaneous IRI enables the effectiveness of preconditioning strategies on clinical outcomes to be assessed. Recent work has validated the use of animal models of IRI studies as a surrogate for human IRI by comparing the molecular changes observed in human subjects and existing animal data.10,11 
The rat transverse rectus abdominis myocutaneous (TRAM) flap was first described in 1987 in German12 and in 199313 in English. This model gained wide popularity13-25 as a cheap, robust model to investigate different strategies to reduce IRI associated with free tissue transfer.14,17-22 The majority of these studies were designed as unipedicled TRAM flaps based on the deep, inferior, epigastric vascular pedicle.15-18,20-22 Comparison of the data from these studies is complicated by the use of different sized cutaneous islands (10.5 - 30 cm2) and different lengths of postoperative follow-up (2 - 10 days). The average total percentage area flap necrosis in the control arm of these studies is 69 &plusmn; 6.2% (mean &plusmn; SEM). It should be noted that these six papers all employ the rectus abdominis muscle as a carrier for the vascular pedicle but do not expose, divide and microanastomose or clamp the vessels. Zhang et al.23 have described a true, free rat TRAM flap based on the superior epigastric vessels in which the flaps were raised, vessels divided and the myocutaneous flap transferred and microanastomosed to the groin vessels. This difficult technique required the microanastomosis of 0.45 - 0.5 mm caliber vessels. Only fifteen were performed and of these 67% survived.23 The model described by Zhang et al.23 is an excellent model for the human free TRAM flap as it truly mirrors the injury incurred during FTT. The other published models of a rat TRAM flap more accurately reflect the injuries incurred during a human pedicled TRAM but do not accurately reflect the IRI as these flap in do not undergo an ischemic period followed by reperfusion as the vascular pedicle is never clamped or divided and microanastomosis performed. This protocol and video describe a new model of free tissue transfer using the rat TRAM in which the IRI is replicated using microclamps. This more faithfully replicates IRI than the pedicle TRAM predecessors but is technically easier than performing the microanastomosis. Microclamps have been widely employed by transplant researchers to recreate IRI associated with solid organ transplant;26-33however, this is the first time it has been described in the rat TRAM flap.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Moor LD12 laser doppler imaging scanner</td>
 <td></td>
 <td></td>
 <td><a href="http://gb.moor.co.uk/product/moorldi2-laser-doppler-imager/8" target="_blank">http://gb.moor.co.uk/product/moorldi2-laser-doppler-imager/8</a></td>
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 <td>Complete homeothermic blanket system with flexible probe. Small. 230 VAC, 50 Hz</td>
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 <td>507221F</td>
 <td><a href="http://www.harvardapparatus.com" target="_blank">www.harvardapparatus.com</a></td>
 </tr>
 <tr>
 <td>Graeffe forceps 0.8 mm tips curved</td>
 <td></td>
 <td>11052-10</td>
 <td>2, <a href="http://www.finescience.de" target="_blank">http://www.finescience.de</a></td>
 </tr>
 <tr>
 <td>Acland clamps</td>
 <td></td>
 <td>00398 V</td>
 <td>B-1 'V' pattern clamps used on both artery and vein. <a href="http://www.merciansurgical.com/acland-clamps.pdf" target="_blank">http://www.merciansurgical.com/acland-clamps.pdf</a></td>
 </tr>
 <tr>
 <td>Clamp applicator</td>
 <td></td>
 <td>CAF-4</td>
 <td><a href="http://www.merciansurgical.com/acland-clamps.pdf" target="_blank">http://www.merciansurgical.com/acland-clamps.pdf</a></td>
 </tr>
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 <td>Gemini cautery unit</td>
 <td></td>
 <td>726067</td>
 <td><a href="www.harvardapparatus.com" target="_blank">www.harvardapparatus.com</a></td>
 </tr>
 <tr>
 <td>Micro-vessel dilators 11 cm 0.3 mm tips 00124</td>
 <td></td>
 <td>D-5a.2</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
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 <td>Micro Jewellers Forceps 11cm angulated 00109</td>
 <td></td>
 <td>JFA-5b</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
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 <td>Micro Jewellers Forceps 11 cm straight 00108</td>
 <td></td>
 <td>JF-5</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
 </tr>
 <tr>
 <td>Acland Single Clamps B-1V (Pair)</td>
 <td></td>
 <td>396</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
 </tr>
 <tr>
 <td>Micro Scissors Round Handles 15 cm Straight</td>
 <td></td>
 <td>67</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
 </tr>
 <tr>
 <td>Iris Scissors 11.5 cm Curves EASY-CUT</td>
 <td></td>
 <td>EA7613-11</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
 </tr>
 <tr>
 <td>Mayo Scissors 14 cm Straight Chamfered Blades EASY-CUT</td>
 <td></td>
 <td>EA7652-14</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
 </tr>
 <tr>
 <td>Derf Needle Holders 12 cm TC</td>
 <td></td>
 <td>703DE12</td>
 <td><a href="http://www.merciansurgical.com" target="_blank">http://www.merciansurgical.com</a></td>
 </tr>
 <tr>
 <td>Ethilon 5-0</td>
 <td></td>
 <td>W1618</td>
 <td><a href="http://www.farlamedical.co.uk/" target="_blank">http://www.farlamedical.co.uk/</a></td>
 </tr>
 <tr>
 <td>Vicryl rapide 6-0</td>
 <td></td>
 <td>W9913</td>
 <td><a href="http://www.millermedicalsupplies.com/" target="_blank">http://www.millermedicalsupplies.com/</a></td>
 </tr>
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 <td>Instrapac - Adson Toothed Forceps (Extra Fine)</td>
 <td></td>
 <td>7973</td>
 <td><a href="http://www.millermedicalsupplies.com/" target="_blank">http://www.millermedicalsupplies.com/</a></td>
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 <tr>
 <td>Castroviejo needle holders</td>
 <td></td>
 <td>12565-14</td>
 <td><a href="http://s-and-t.ne" target="_blank">http://s-and-t.ne</a></td>
 </tr>
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 <td>Heat Lamp</td>
 <td></td>
 <td></td>
 <td><a href="http://www.chicken-house.co.uk" target="_blank">http://www.chicken-house.co.uk</a></td>
 </tr>
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 <td>Silicone sheeting 0.3 mm translucent</td>
 <td></td>
 <td></td>
 <td><a href="http://www.silex.co.uk/" target="_blank">http://www.silex.co.uk/</a></td>
 </tr>
 <tr>
 <td>Image J software</td>
 <td></td>
 <td></td>
 <td><a href="http://rsbweb.nih.gov/ij/" target="_blank">http://rsbweb.nih.gov/ij/</a></td>
 </tr>
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 <td>Zeiss OPMI pico</td>
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 <td></td>
 <td><a href="http://www.zeiss.co.uk/" target="_blank">http://www.zeiss.co.uk/</a></td>
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 <td>Operating microscope</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
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 <td>Vet tech solution isofluorane rig</td>
 <td></td>
 <td></td>
 <td><a href="http://www.vet-tech.co.uk/" target="_blank">http://www.vet-tech.co.uk/</a></td>
 </tr>
 <tr>
 <td>Vet tech solution isofluorane rig</td>
 <td></td>
 <td></td>
 <td><a href="http://www.vet-tech.co.uk/" target="_blank">http://www.vet-tech.co.uk/</a></td>
 </tr>
 </tbody>
</table>

in situ transverse rectus abdominis myocutaneous flap: a rat model of myocutaneous ischemia reperfusion injury

Free tissue transfer is the gold standard of reconstructive surgery to repair complex defects not amenable to local options or those requiring composite tissue. Ischemia reperfusion injury (IRI) is a known cause of partial free flap failure and has no effective treatment. Establishing a laboratory model of this injury can prove costly both financially as larger mammals are conventionally used and in the expertise required by the technical difficulty of these procedures typically requires employing an experienced microsurgeon. This publication and video demonstrate the effective use of a model of IRI in rats which does not require microsurgical expertise. This procedure is an in situ model of a transverse abdominis myocutaneous (TRAM) flap where atraumatic clamps are utilized to reproduce the ischemia-reperfusion injury associated with this surgery. A laser Doppler Imaging (LDI) scanner is employed to assess flap perfusion and the image processing software, Image J to assess percentage area skin survival as a primary outcome measure of injury.

Rat models are more economical than larger animals models,36 are disease-resistant in nature and may be genetically manipulated. Loose skinned animals, such as rodents, were thought to have a different arrangement of cutaneous blood supply compared to fixed skinned animals such as humans and pigs. In loose skinned animals, skin is supplied primarily by direct cutaneous blood vessels passing through the subcutaneous fat to the overlying skin (Figure 4) By contrast, fixed skinned animals derive...

Watch this Scientific Journal Video about In situ Transverse Rectus Abdominis Myocutaneous Flap: A Rat Model of Myocutaneous Ischemia Reperfusion Injury at JoVE.com

In situ Transverse Rectus Abdominis Myocutaneous Flap: A Rat Model of Myocutaneous Ischemia Reperfusion Injury

Free tissue transfer is widely employed in reconstructive surgery to restore form and function following oncological resection and trauma. Preconditioning this tissue prior to surgery may improve outcome. This article describes an in situ transverse rectus abdominis myocutaneous flap (TRAM) in rats as a means for testing preconditioning strategies.

In situ Transverse Rectus Abdominis Myocutaneous Flap: A Rat Model of Myocutaneous Ischemia Reperfusion Injury

Free tissue transfer is the gold standard of reconstructive surgery to repair complex defects not amenable to local options or those requiring composite ...

in-situ-transverse-rectus-abdominis-myocutaneous-flap-rat-model

Research

JoVE Journal

Medicine

13.7K vistas.  Royal Infirmary of Edinburgh. Transferencia de tejido libre es ampliamente utilizado en cirugía reconstructiva para restaurar la forma y la función después de la resección oncológica y trauma. Preacondicionamiento este tejido antes de la cirugía puede mejorar el resultado. En este artículo se describe una In situ Músculo recto abdominal transverso colgajo miocutáneo (TRAM) en ratas como un medio para probar estrategias de preacondicionamiento.

Video: In situ Transverse Rectus Abdominis Myocutaneous Flap: A Rat Model of Myocutaneous Ischemia Reperfusion Injury

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 ...

Induction and Assessment of Ischemia-reperfusion Injury in Langendorff-perfused Rat Hearts

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for myocardial infarction and cardiac complications of thoracic surgery. The ability of certain drugs to precondition the myocardium against ischemia reperfusion injury has led to multiple clinical trials, with little success. The isolated heart model allows acute observation of the functional effects of ischemia reperfusion injury in real time, including the effects of various pharmacological interventions administered at any time-point before or within the ischemia-reperfusion injury window. Since brief periods of ischemia can precondition the heart against ischemic injury, in situ aortic cannulation is performed to allow for functional assessment of non-preconditioned myocardium. A saline filled balloon is placed into the left ventricle to allow for real-time measurement of pressure generation. Ischemic injury is simulated by the cessation of perfusion buffer flow, followed by reperfusion. The duration of both ischemia and reperfusion can be modulated to examine biochemical events at any given time-point. Although the Langendorff isolated heart model does not allow for the consideration of systemic events affecting ischemia and reperfusion, it is an excellent model for the examination of acute functional and biochemical events within the window of ischemia reperfusion injury as well as the effect of pharmacological intervention on cardiac pre- and postconditioning. The goal of this protocol is to demonstrate how to perform in situ aortic cannulation and heart excision followed by ischemia/reperfusion injury in the Langendorff model.

The isolated rat heart is an enduring model for ischemia reperfusion injury. Here, we describe the process of harvesting the beating heart from a rat via in situ aortic cannulation, Langendorff perfusion of the heart, simulated ischemia-reperfusion injury, and infarct staining to confirm the extent of ischemic insult.

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for ...

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, ...

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 ...

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand and face transplants and, more recently, abdominal wall, uterus, and urogenital transplants. Functional outcomes have exceeded initial expectations, and most recipients enjoy an improved quality of life. However, as clinical experience accumulates, chronic rejection and complications from the immunosuppression must be addressed. In many cases where grafts have failed, the causative pathology has been ischemic vasculopathy. <span style="font-size: 11pt; line-height: 115%; font-family: Arial, sans-serif; color: rgb(41, 43, 49); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">The biological mechanisms of the acute and chronic rejection associated with VCA, especially ischemic vasculopathy, are important areas of research. However, due to the very small number of VCA patients, the evaluation of proposed mechanisms is better addressed in an experimental model. Multiple groups have used animal models to address some of the relevant unsolved questions in VCA rejection and vasculopathy. Several model designs involving a variety of species are described in the literature. Here we present a reproducible model of VCA heterotopic hindlimb osteomyocutaneous flap in the rat that can be utilized for translational VCA research. This model allows for the serial evaluation of the graft, including biopsies and different imaging modalities, while maintaining a low level of morbidity.

Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand ...

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 ...

Delayed Intramyocardial Delivery of Stem Cells after Ischemia Reperfusion Injury in a Murine Model

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, cardiac stem cell therapy is studied by delivering SCs concurrently with the induction of myocardial injury. However, this approach presents two significant limitations: the early hostile pro-inflammatory ischemic environment may affect the survival of transplanted SCs, and it does not represent the subacute infarction scenario where SCs will likely be used. Here we describe a two-part series of surgical procedures for the induction of ischemia-reperfusion injury and delivery of mesenchymal stem cells (MSCs). This method of stem cell administration may allow for the longer viability and retention around damaged tissue by circumventing the initial immune response. A model of ischemia reperfusion injury was induced in mice accompanied by the delivery of mesenchymal stem cells (3.0 x 105), stably expressing the reporter gene firefly luciferase under the constitutively expressed CMV promoter, intramyocardially 7 days later. The animals were imaged via ultrasound and bioluminescent imaging for confirmation of injury and injection of cells, respectively. Importantly, there was no added complication rate when performing this two-procedure approach for SC delivery. This method of stem cell administration, collectively with the utilization of state-of-the-art reporter genes, may allow for the in vivo study of viability and retention of transplanted SCs in a situation of chronic ischemia commonly seen clinically, while also circumventing the initial pro-inflammatory response. In summary, we established a protocol for the delayed delivery of stem cells into the myocardium, which can be used as a potential new approach in promoting regeneration of the damaged tissue.

Stems cells are continuously investigated as potential treatments for individuals with myocardial damage, however, their decreased viability and retention within injured tissue can impact their long-term efficacy. In this manuscript we describe an alternative method for stem cell delivery in a murine model of ischemia reperfusion injury.

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, ...

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, ...