Name: murine surgical model of topical elastase induced descending thoracic aortic aneurysm
Uploaded: 2019-08-24T15:00:00.000+00:00
Duration: 8 s
Description: Watch this Scientific Journal Video about Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm at JoVE.com

This work was supported by AHA Scientist Development Grant 14SDG18730000 (M.S.), NIH K08 HL098560 (G.A.) and RO1 HL081629 (G.R.U.) grants. This project was supported by the Thoracic Surgery Foundation for Research and Education (TSFRE) Research Grant (PI: G. Ailawadi). The content is solely the responsibility of the authors and does not necessarily represent the views of the NHLBI or the TSFRE. We thank Anthony Herring and Cindy Dodson for their knowledge and technical expertise.

Animal protocols were approved by the University of Virginia Institutional Animal Care and Use Committee (No. 3634).
1. Induction of anesthesia and intubation
<ol>
	<li>Place an 8-10-week-old male C57BL/6 mouse in a closed chamber with continuous flow of 5% isoflurane and oxygen mixture for 5 min, until respirations are visibly slowed. 
	NOTE: Different strains, genders and ages of mice can be used depending on the experimental protocol. Female mice may be more difficult to intubate because of smaller size and thus smaller airway.</li>
	<li>Intubate the mouse as described by Vandivort et al.13. 
	NOTE: The intubation step is the most difficult portion of this model to both learn and perform. The above referenced authors do an excellent job explaining the steps in their video.</li>
</ol>
2. Securing the mouse to the surgical board
<ol>
	<li>Connect the endotracheal (ET) tube, confirm chest rise, and lay the mouse in the right lateral decubitus position.&#160;Turn isoflurane to approximately 3% and apply lubrication to both eyes. The ventilator should be set to provide approximately 200 breaths per minute and 225 &#181;L of tidal volume. 
	NOTE: Toxicity is rapid if isoflurane is left at 5%.&#160;However, response to isoflurane is variable so anesthetic flow rate should be titrated such that spontaneous diaphragmatic contractions occur approximately every 10-20 s and oxygenation appears adequate on inspection of mucous membranes and exposed tissue.</li>
	<li>Secure the ET tube to the board with tape. Extend the right arm rostrally so the front paw is in line with the nose and secure it with tape.</li>
	<li>Pull the tail caudally in order to create a line of tension between the right arm and tail, producing extension of the spine. 
	NOTE: Securing both the tail and the right paw in line prevents over insertion or dislodgement of the ET tube.</li>
	<li>Tape the left leg in its natural position. Draw the left arm ventrally over rolled gauze and tape down.</li>
</ol>
3. Preparation for surgery
<ol>
	<li>Shave the left flank of the mouse with electric clippers from left shoulder to left abdomen.</li>
	<li>Use a cotton tipped applicator to brush betadine solution over the surgical site. Use a new cotton tipped applicator to brush 70% ethanol solution over the surgical site. Allow to dry. Place a sterile drape.</li>
	<li>CAUTION: Ensure that all ethanol is completely dried before proceeding as electrocautery is used and can ignite the ethanol.</li>
</ol>
4. Entry into thorax
<ol>
	<li>Make a 1 cm lateral incision at the midpoint of the hemithorax using surgical scissors. Use handheld electrocautery to divide muscle layers until the ribs are visible.</li>
	<li>Directly cauterize a 2 mm portion of the rib. 
	NOTE: When using electrocautery on the rib, observe the spontaneous breathing frequency. If the mouse has a diaphragmatic contraction during cautery, the tip may enter the thorax and puncture the lung, which is usually fatal in this model.</li>
	<li>Using a wetted, fine cotton-tipped applicator in the rib space superior to the cauterized portion, rotate the tip upon the tissue to break into the pleural space. Place wetted 3 mm x 2 mm sponge into the thorax to help collapse the lung. 
	NOTE: Only let soft, wet, sponges contact the lung as it is extremely delicate.</li>
	<li>Use scissors to cut along the top aspect of the cauterized rib, until diaphragm is visible.</li>
</ol>
5. Aortic exposure
<ol>
	<li>Place rib retractors and use them to open the thoracotomy incision. Carefully remove sponge from the surface of the lung.</li>
	<li>Place wetted 6 mm x 4 mm sponge so that it lays covering the lung, with ends pointing rostrally and caudally. Place the (wide flat) lung retractor on the sponge and gently slide the retractor ventrally until the descending thoracic aorta is exposed.</li>
	<li>Use #7 forceps to dissect the connective tissue and fat off the aorta for an approximately 5 mm section. 
	NOTE: Small veins may be running transversely across the aorta; avoid tearing them during dissection (using at least 14x magnification can help to avoid this complication).</li>
</ol>
6. Elastase exposure
<ol>
	<li>Saturate 0.5 mm x 1 mm sponge with 12 &#956;L of porcine pancreatic elastase and place it upon the exposed surface of the aorta. 
	NOTE: Do not let the sponge touch the contralateral lung.</li>
	<li>After the predetermined time (usually 3-5 min), remove the elastase sponge with #7 forceps. Remove the lung retractor. Irrigate the chest cavity with 1 mL of sterile saline. 
	NOTE: Remove the lung retractor before irrigation with saline as it will allow the lung sponge to become saturated and soft, making it easier to remove from the surface of the lung.</li>
	<li>Use rolled 2&#34;&#160;x 2&#34;&#160;gauze to absorb the remaining saline irrigation. Turn isoflurane down to 2%.</li>
</ol>
7. Closure of chest
<ol>
	<li>Remove the lung sponge. Remove the caudal lung retractor. Remove the rostral lung retractor.</li>
	<li>Place 3 interrupted 6-0 non-absorbable sutures to oppose the ribs, tie a loose knot in each but do not tie down. Take great care to not touch the lung with the suture needle.</li>
	<li>Re-inflate the lung by occluding the outflow tube on ventilator or by rapidly blowing 0.5-1.0 mL of air through the ET tube by utilizing the 3 way stop cock. 
	NOTE: To avoid barotrauma, avoid excessive use of the air-filled syringe, and use no more than 1.0 mL of air.</li>
	<li>Tie non-absorbable sutures. Reapproximate the muscle layers with a running 5-0 absorbable multifilament suture. Turn isoflurane down to 1%.</li>
	<li>Close skin with 7-10 interrupted 5-0 non-absorbable sutures. Turn isoflurane to 0%.</li>
</ol>
8. Recovery
<ol>
	<li>Administer 6 &#181;g (0.02 mL of a 0.3 mg/mL suspension)&#160;&#160;of buprenorphine subcutaneously. Remove the tape from the right foot, tail, and left arm followed by the right arm tape.</li>
	<li>When the mouse moves extremities, extubate by pulling it by its tail to slide off the ET tube. Place in the high oxygen content warmer chamber in the supine position. 
	NOTE: It is safe to move mouse from oxygen chamber into cage when it can turn itself over to normal standing position.&#160;Furthermore, mice should be monitored for signs of pain, distress, or failure to thrive frequently for the first 24-48 hours after surgery and provided additional analgesia or soft food as indicated.</li>
</ol>
9. Exposure of aortic aneurysm (terminal harvest procedure)
NOTE: In general, tissue harvest is carried out at 14 days, as this represents the period of maximal dilatation. However, depending on the experiment, the harvest procedure timing can be carried out at any time between 3 days and 4+ weeks, depending on the experiment.
<ol>
	<li>Intubate and secure mouse to operative field as described above (sections 1-3). Use scissors to incise skin medially from left flank to central abdomen taking care to not enter the peritoneum.</li>
	<li>Incise skin from dorsal left flank rostrally to the level of the left shoulder. Then incise at a 90&#176; angle through the axilla to the sternum. 
	NOTE: This incision should completely encircle the original skin incision.</li>
	<li>Using cautery, dissect skin flap toward the ventral aspect of the mouse, exposing the left hemithorax. Use scissors to enter the abdomen and incise along the left costal margin from ventral to dorsal to expose the underside of the left diaphragm. The lateral most edge of the diaphragm may open which is desired.</li>
	<li>If not created with the prior step, incise the diaphragm at its most lateral edge. Place tip of cautery in this hole and cauterize the diaphragm off the costal margin to the xyphoid process. Using a damp fine tipped cotton-tipped applicator, gently free the lung from adhesions to the chest wall and push lung medially. 
	NOTE: If adhesions do not easily come off the chest wall, use cautery to remove them; doing so helps avoid tearing the lung which can cause heavy bleeding.</li>
	<li>Cauterize the inside of the chest wall from rib one to the costal margin, dorsal to the mid axillary line but at least 2 mm from the aorta. Cut chest wall along the cauterized line. 
	NOTE: This technique avoids bleeding from the intercostal arteries.</li>
	<li>Cut along the superior margin of rib one and then caudally along the lateral edge of the sternum, removing the left rib cage. Place retractors on the lung and pull medially. Place retractor on diaphragm and draw caudally to expose as much aorta as possible.</li>
	<li>Use a dry cotton-tipped applicator to remove adhesions from aortic aneurysm and an unaffected distal segment. Measure the diameter of the unaffected control segment and the widest portion of the elastase treated aneurysm using video micrometry. 
	NOTE: The video micrometry measurements are used to calculate the percent dilation of the aneurysmal segment compared to a control segment with Equation 1. A control segment that is 0.5 mm distal to the aneurysmal segment 1 is selected. 
	 
	<img alt="Equation 1" src="/files/ftp_upload/60105/60105eq1.jpg" /> &#160; &#160; &#160; &#160; &#160; &#160; &#160;Equation 1 
	&#160;</li>
	<li>Grasp aorta with Harms forceps, just distal to the treated segment. Use scissors to cut distal to forceps, then dissect the aorta off spinal column. Cut aorta proximal to treated segment and remove aneurysmal aorta.</li>
	<li>Using a tuberculin syringe and needle, wash the aortic lumen with saline and process tissue as desired.&#160;</li>
</ol>

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R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endovascular+stenting+versus+open+surgery+for+thoracic+aortic+disease:+systematic+review+and+meta-analysis+of+perioperative+results.">Endovascular stenting versus open surgery for thoracic aortic disease: systematic review and meta-analysis of perioperative results.</a> Journal of Vascular Surgery. 47 (5), 1094-1098 (2008).</li><li>Absi, T. 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The authors have nothing to disclose.

The thoracic and abdominal aorta are cellularly and embryologically distinct, which is relevant to aneurysmal disease14,15,16. Therefore, a specific animal model to study TAA is needed. Though other murine dTAA models have been published8, ours is the only model to create descending thoracic aortic dilatation which can be considered truly aneurysmal (over 50% dilation). Furthermore, our model is relativel...

The purpose of this method is to study the development, pathophysiology, and structural changes in the murine descending thoracic aorta during aortic aneurysm formation. Our model offers a reproducible and consistent method to induce thoracic aortic aneurysms (dTAA) in mice thereby allowing for the testing of various genetic and pharmacologic inhibitors. This work can help identify drugs and gene-therapies which could be translated to a viable treatment strategy for humans with dTAA disease.
dTAAs form when the wall of the thoracic aorta becomes weakened and dilates over time until reaching a critical diameter when tearing or rupture can then occur. Clinically, dTAA can progress in silence, increasing in size until the structure of the aortic wall is so distorted that it eventually fails, with catastrophic consequences. Concerningly, symptoms usually develop only when the aneurysm has reached a perilous size (100-150% dilation) and is at high risk for dissection or rupture1,2. dTAA rupture is almost universally lethal3, and elective surgical repair carries significant morbidity4,5. Furthermore, most patients carry the diagnosis of an aortic aneurysm for approximately 5 years before surgical repair6,7. This window represents an opportune time to intervene non-surgically. Thus, medical therapies to treat or slow progression of dTAA are needed and would represent a significant advancement to the field of aneurysm research. There are currently no medical treatments for dTAA available, mostly because of an incomplete understanding of dTAA pathogenesis.
Over the last 20 years, several dTAA animal models have been developed, but each of these models were distinct from our own and did not produce robust aneurysms. A murine dTAA model most similar to ours was developed by Ikonomidis et al.8, which includes direct application of CaCl2 to the adventitia of the aorta. Though our model was adapted from many of the techniques set forth by Ikonomidis, our model is unique in three separate ways. First, in our model the aorta is exposed to topical elastase for 3-5 minutes, compared to 15 minutes of CaCl2 exposure. Second, aortic dilation occurs in 2 weeks, compared to 16 weeks in the CaCl2 model. Last, our model consistently produces aneurysms of approximately 100% dilatation, compared to the aortic dilatations of 20-30% produced by CaCl2 application (which cannot be truly considered aneurysms as they are defined as an increase in aortic diameter &#62;50%). There are other non-surgical murine models of aneurysm formation, such as the Apo E knockout mouse, which form robust aneurysms with infusion of angiotensin II. However, these mice develop supra-renal or ascending thoracic aortic aneurysms rather than aneurysms specifically in the descending thoracic aorta9,10.
The rational for this protocol is to have a simple, inexpensive, and time suitable way to study dTAA in a murine model. The mouse model provides a unique opportunity to utilize many genetic and cell-specific knockouts that have been found to be impactful in other vascular diseases. The use of our specific TAA model has been well received and experiments utilizing it have been published in high impact journals11,12. To this point, the model has been used to investigate possible genetic and pharmacologic treatments that had a significant effect in the abdominal aortic aneurysm (AAA) murine models; however, as our lab has expanded use of the dTAA model, we are finding targets unique to dTAA formation which could be used as targeted therapies in humans.
This model is most appropriate for labs that have murine micro-surgical capabilities. Though it is technically challenging, it can be executed consistently even by researchers with no prior surgical experience. For a researcher with no murine surgical experience the model can be mastered in approximately 20 operative sessions (or approximately 50 mice). For the researcher with prior surgical experience, the model can be mastered in 5 operative sessions (approximately 20 mice). We believe with a high-quality video, the time to mastery can be further reduced. After proficiency is achieved, the procedure can be completed in 35 minutes for the surgery, and 20 minutes for the terminal harvest. The surgeons in our lab can complete 10-12 full surgeries per day, with an operative mortality rate of 5-10%. The most common cause of mortality is lung injury upon entry to the chest, anesthetic toxicity, or tear of the aorta during dissection. In addition to dTAA research, this model also serves as a guide for safe and easy access to the thoracic aorta and lung hilum for researchers studying other interventions in the chest.&#160;

<table><tbody><tr><td>Angiocatheter (22G)</td><td></td><td></td><td>Used for ET Tube</td></tr><tr><td>Dumont Tweezers; Pattern #7 x2</td><td>Roboz</td><td>RS-4982</td><td></td></tr><tr><td>Graefe Tissue Forceps</td><td>Roboz</td><td>RS-5158</td><td></td></tr><tr><td>Harms Forceps x2</td><td>Roboz</td><td>RS-5097</td><td></td></tr><tr><td>Intracardiac Needle Holder; Extra Delicate; Carbide Jaws; 7&quot; Length</td><td>Roboz</td><td>RS-7800</td><td></td></tr><tr><td>&amp;nbsp;KL 1500 LED Light Source</td><td>Leica</td><td>150-400</td><td></td></tr><tr><td>&amp;nbsp;M205A Dissction Microscope</td><td>Leica</td><td>CH 94-35</td><td></td></tr><tr><td>Iris Scissors, 11cm, Tungsten Carbide</td><td>World Precision Instruments</td><td>500216-G</td><td></td></tr><tr><td>Metal Clip board</td><td></td><td></td><td>Use with the Mouse Retractor Set&amp;nbsp;</td></tr><tr><td>Mouse Retractor Set</td><td>Kent</td><td>SURGI-5001</td><td>Need 2 short and 1 tall fixators</td></tr><tr><td>Mouse Ventilator MiniVent Type 845, 115 V, Power Supply with US Connector</td><td>Harvard Apparatus</td><td>73-0043</td><td>MiniVent Ventilator for Mice (Model 845), Single Animal, Volume Controlled</td></tr><tr><td>Sigma Aldrich</td><td>Elastase from porcine pancreas</td><td>E0258-50MG</td><td>Can be purchased in various size bottles</td></tr><tr><td>Small Vessel Cauterizer Kit</td><td>Fine Science Tools</td><td>18000-00</td><td>Recommend using rechargable AA batteries</td></tr><tr><td>Spring Scissors, 10.5cm</td><td>World Precision Instruments</td><td>14127</td><td></td></tr><tr><td>Steril Swabs (Sponges)</td><td>Sugi</td><td>31603</td><td>Can be cut to size</td></tr><tr><td>Surgi Suite Surgical Platform</td><td>Kent</td><td></td><td>Attach to clip board&amp;nbsp;</td></tr><tr><td>Tech IV Isoflurane Vap</td><td>Jorgensen&amp;nbsp;Laboratories&amp;nbsp;</td><td>J0561A</td><td>Anesthesia vaporizer&amp;nbsp;</td></tr></tbody></table>

murine surgical model of topical elastase induced descending thoracic aortic aneurysm

According to the Center for Disease Control, aortic aneurysms (AAs) were considered a leading cause of death in all races and both sexes from 1999-2016. An aneurysm forms as a result of progressive weakening and eventual dilation of the aorta, which can rupture or tear once it reaches a critical diameter. Aneurysms of the descending aorta in the chest, called descending thoracic aortic aneurysms (dTAA), make up a large proportion of aneurysm cases in the United States. Uncontained dTAA rupture is almost universally lethal, and elective repair has a high rate of morbidity and mortality. The purpose of our model is to study dTAA specifically, to elucidate the pathophysiology of dTAA and to search for molecular targets to halt the growth or reduce the size of dTAA. By having a murine model to study thoracic pathology precisely, targeted therapies can be developed to specifically test dTAA. The method is based on the placement of porcine pancreatic elastase (PPE) directly on the outer murine aortic wall after surgical exposure. This creates a destructive and inflammatory reaction, which weakens the aortic wall and allows for aneurysm formation over weeks to months. Though murine models possess limitations, our dTAA model produces robust aneurysms of predictable size. Furthermore, this model can be used to test genetic and pharmaceutical targets which may arrest dTAA growth or prevent rupture. In human patients, interventions such as these could help avoid aneurysm rupture, and difficult surgical intervention.

The application of our protocol results in robust dTAA in mice compared to saline controls. The TAAs developed are fusiform in shape and occur only in the treated portion of the aorta (Figure 1 and Figure 2)11. Figure 2 shows an example of a video micrometry measurement at tissue harvest. Using Equation 1, the aortic dilation is 130% in this example.
The original study by Johnston ...

Watch this Scientific Journal Video about Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm at JoVE.com

Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm

We describe a surgical protocol to consistently induce robust descending thoracic aortic aneurysms in mice. The procedure involves left thoracotomy, thoracic aorta exposure, and placement of a sponge soaked in porcine pancreatic elastase on the aortic wall.

According to the Center for Disease Control, aortic aneurysms (AAs) were considered a leading cause of death in all races and both sexes from 1999-2016. ...

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7.5K Views.  University of Virginia School of Medicine. We describe a surgical protocol to consistently induce robust descending thoracic aortic aneurysms in mice. The procedure involves left thoracotomy, thoracic aorta exposure, and placement of a sponge soaked in porcine pancreatic elastase on the aortic wall.

Video: Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm

Creation of Murine Experimental Abdominal Aortic Aneurysms with Elastase 

Transient intraluminal infusion of porcine pancreatic elastase into the infrarenal segment of the abdominal aorta is the most widely used animal model of abdominal aortic aneurysm (AAA) ever since it was first described in rats by Anidjar and colleagues.1 The rationale for its development was based on the disrupted nature of elastin observed in AAAs. This rat model has been modified to produce AAAs in the infrarenal aortic region of mice.2 The model has the ability to add broad insight into the pathobiology of AAA due to the emergence of numerous transgenic and gene knockout mice. Moreover, it is a viable platform to test potential therapeutic agents for AAA. In this video, we demonstrate the elastase infusion AAA procedure used in our laboratory. 
Mice are anesthetized using 2.5% isoflurane, and a laparotomy is performed under sterile conditions. The abdominal aortais isolated with the assistance of an operating stereomicroscope (Leica). After placing temporary ligatures around the proximal and distal aorta, an aortotomy is created at the bifurcation with the tip of a 30-gauge needle. A heat-tapered segment of PE-10 polyethylene tubing is introduced through the aortotomy and secured. The aortic lumen is subsequently perfused for 5-15 minutes at 100 mm Hg with saline containing type I porcine pancreatic elastase (4.5 U/mL; Sigma Chemical Co.). After removing the perfusion catheter, the aortotomy is repaired without constriction of the lumen.

Creation of Murine Experimental Abdominal Aortic Aneurysms with Elastase

This video shows how to induce abdominal aortic aneurysms (AAA) in mice via transient intraluminal infusion of porcine pancreatic elastase into the infrarenal segment of the abdominal aorta. The model has the ability to add broad insight into the pathobiology of AAA due to the emergence of numerous transgenic and gene knockout mice.

Transient intraluminal infusion of porcine pancreatic elastase into the infrarenal segment of the abdominal aorta is the most widely used animal model of ...

Biology

A New Murine Model of Endovascular Aortic Aneurysm Repair

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of Abdominal aortic aneurysm (AAA) pathophysiology. There is no abdominal aortic aneurysm endograft exclusion model cheap and reproducible, which would allow deep investigations of AAA before and after treatment. We hereby describe how to induce, and then to exclude with a covered coronary stentgraft an abdominal aortic aneurysm in a rat. The well known elastase induced AAA model was first reported in 19901 in a rat, then described in mice2. Elastin degradation leads to dilation of the aorta with inflammatory infiltration of the abdominal wall and intra luminal thrombus, matching with human AAA. Endovascular exclusion with small covered stentgraft is then performed, excluding any interactions between circulating blood and the aneurysm thrombus. Appropriate exclusion and stentgraft patency is confirmed before euthanasia by an angiography thought the left carotid artery. Partial control of elastase diffusion makes aneurysm shape different for each animal. It is difficult to create an aneurysm, which will allow an appropriate length of aorta below the aneurysm for an easy stentgraft introduction, and with adequate proximal and distal neck to prevent endoleaks. Lots of failure can result to stentgraft introduction which sometimes lead to aorta tear with pain and troubles to stitch it, and endothelial damage with post op aorta thrombosis. Giving aspirin to rats before stentgraft implantation decreases failure rate without major hemorrhage. Clamping time activates neutrophils, endothelium and platelets, and may interfere with biological analysis.

Histological and biochemical modifications after aneurysm endograft exclusion are still unclear. We describe a new model of endograft implantation on a murine aneurysm. Through thrombus analysis with and without persistant circulating blood stream interface, and new endovascular biomaterials evaluation, this model will help to better understand endovascular aneurysm exclusion pathobiology.

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of ...

Reproducable Paraplegia by Thoracic Aortic Occlusion in a Murine Model of Spinal Cord Ischemia-reperfusion

Background 
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered advancements to decrease the occurrence this injury.&#160;The current model demonstrates reproducible lower&#160;extremity paralysis following thoracic aortic occlusion.
Methods 
Adult male C57BL6 mice were anesthetized with isoflurane. Through a cervicosternal incision the aorta was exposed. The descending thoracic aorta and left subclavian arteries were identified without entrance into pleural space. Skeletonization of these arteries was followed by immediate closure (Sham) or occlusion for 4 min (moderate ischemia) or 8 min (prolonged ischemia). The sternotomy and skin were closed and the mouse was transferred to warming bed for recovery.&#160; Following recovery, functional analysis was obtained at 12 hr intervals until 48 hr.
Results 
Mice that underwent sham surgery showed no observable hind&#160;limb deficit. Mice subjected to moderate ischemia for 4 min had minimal functional deficit at 12 hr followed by progression to complete paralysis at 48 hr. Mice subjected to prolonged ischemia had an immediate paralysis with no observable hind-limb movement at any point in the postoperative period. There was no observed intraoperative or post&#160;operative mortality.
Conclusion 
Reproducible lower extremity paralysis whether immediate or delayed can be achieved in a murine model. Additionally, by using a median sternotomy and careful dissection, high survival rates, and reproducibility can be achieved.

The lack of mechanistic understanding of spinal cord ischemia-reperfusion injury has hindered further adjuncts to prevent paraplegia following high risk aortic operations. Thus, the development of animal models is imperative. This manuscript demonstrates reproducible lower extremity paralysis following thoracic aortic occlusion in a murine model.

Background
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered ...

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 ...

Surgical Models of Gastroesophageal Reflux with Mice

Multiple surgical procedures have been reported to induce gastroesophageal reflux in animals. Herein, we report three surgical models with mice aiming to induce reflux of gastric contents, duodenal contents or mixed contents. Surgical procedures and general principles have been described in detail. A researcher with surgical experience should be able to grasp the technique after a short period of practice. After surgery, most mice can survive and develop reflux esophagitis similar to that in humans. However, it should be noted that histological differences between mouse and human esophagus are the inherent limitations of these surgical models. If used for research on Barrett&#8217;s esophagus and adenocarcinoma, these procedures may need to be combined with genetic modifications.

This article demonstrates surgical procedures of gastroesophageal reflux with mice. These models are useful tools for research on mechanisms and treatment of gastroesophageal reflux disease and potentially Barrett&#8217;s esophagus and esophageal adenocarcinoma.

Multiple surgical procedures have been reported to induce gastroesophageal reflux in animals. Herein, we report three surgical models with mice aiming to ...

Creation of a Rodent Model of Abdominal Aortic Aneurysm by Blocking Adventitial Vasa Vasorum Perfusion

The adventitial vasa vasorum (VV) provides oxygen and nourishment to the aortic wall. Hypoxia in the aortic wall can cause enlarged abdominal aortic aneurysms (AAAs). This article introduces and describes a standard protocol used to induce AAAs through adventitial VV hypoperfusion created with a combination of polyurethane catheter insertion into the aortic lumen and suture ligation of the infrarenal abdominal aorta.
The protocol involves the use of male rats weighing 300-400 g, which are provided food and water ad libitum. After laparotomy with a ventral midline abdominal incision, exfoliation of the aorta is performed, which blocks blood flow from the perivascular tissue. Aortotomy involving a small incision adjacent to the renal artery branches is performed, and a polyurethane catheter is inserted using an 18-gauge indwelling needle. After repairing the incision, tight ligation of the aorta over the catheter blocks VV blood flow from the proximal direction through the aortic wall without disturbing the aortic blood flow. This technique can induce an AAA with progressive aortic dilatation.
The greatest benefit of this model is that VV hypoperfusion causes tissue hypoxia and the development of an infrarenal AAA, which has morphological and pathological characteristics similar to those of a human AAA.

Polyurethane catheter insertion into the aortic lumen and suture ligation of the aorta induce chronic hypoxia due to hypoperfusion of the adventitial vasa vasorum. This article describes a novel animal model of abdominal aortic aneurysm (AAA) with characteristics similar to those of AAA in humans.

The adventitial vasa vasorum (VV) provides oxygen and nourishment to the aortic wall. Hypoxia in the aortic wall can cause enlarged abdominal aortic ...

Investigating the Molecular Mechanisms Underlying Murine Abdominal Aortic Aneurysm

A Mouse Abdominal Aortic Aneurysm Model by Periadventitial Calcium Chloride and Elastase Infiltration

Abdominal aortic aneurysm (AAA) is a life-threatening disease associated with high mortality rates. It is characterized by the permanent dilation of the abdominal aorta with at least a 50% increase in arterial diameter. Various animal models of AAA have been introduced to mimic the pathophysiological changes and study the underlying mechanisms of AAA. Among these models, the calcium chloride (CaCl2)- and elastase-induced AAA models are commonly used in mice. However, these methods have certain limitations. Traditional intraluminal porcine pancreatic elastase (PPE) perfusion is associated with high technical difficulty and a high rupture rate, while periadventitial administration of PPE yields inconsistent results. In addition, the CaCl2-induced AAA model lacks human AAA features, such as atherothrombosis and aneurysm rupture. Therefore, the combined application of CaCl2 and PPE has been proposed as an approach to enhance success rates and induce greater diameter increases in AAA animal models. This manuscript presents a comprehensive protocol for establishing a mouse AAA model through periaortic infiltration of PPE and CaCl2 in the infrarenal segment of the abdominal aorta. By following this protocol, we can achieve an AAA formation rate of approximately 90% with technical simplicity and reproducibility. Further ultrasound and histological experiments confirm that this model effectively replicates the morphological and pathological changes observed in human AAA.

Author Spotlight: Development and Characterization of a Mouse Model for Abdominal Aortic Aneurysm

This manuscript presents a protocol for establishing a mouse abdominal aortic aneurysm model using calcium chloride and elastase, combining the advantages of previous modeling methods. This model can be utilized to investigate the pathophysiological mechanisms underlying abdominal aortic aneurysms.

Abdominal aortic aneurysm (AAA) is a life-threatening disease associated with high mortality rates. It is characterized by the permanent dilation of the ...

Mouse Abdominal Aortic Aneurysm Model Induced by Perivascular Application of Elastase

Abdominal aortic aneurysm (AAA), although primarily asymptomatic, is potentially life-threatening as the rupture of AAA usually has a devastating outcome. Currently, there are several distinct experimental models of AAA, each emphasizing a different aspect in the pathogenesis of AAA. The elastase-induced AAA model is the second most used rodent AAA model. This model involves direct infusion or application of porcine pancreatic elastase (PPE) to the infrarenal segment of the aorta. Due to technical challenges, most elastase-induced AAA model nowadays is performed with the external application rather than an intraluminal infusion of PPE. The infiltration of elastase will cause degradation of elastic lamellae in the medial layers, resulting in the loss of aortic wall integrity and subsequent dilation of the abdominal aorta. However, one disadvantage of the elastase-induced AAA model is the inevitable variation of how the surgery is performed. Specifically, the surgical technique of isolating the infrarenal segment of the aorta, the material used for aorta wrapping and PPE incubation, the enzymatic activity of PPE, and the time duration of PPE application can all be important determinants that affect the eventual AAA formation rate and aneurysm diameter. Notably, the difference in these factors from different studies on AAA can lead to reproducibility issues. This article describes a detailed surgical process of the elastase-induced AAA model through direct application of PPE to the adventitia of the infrarenal abdominal aorta in the mouse. Following this procedure, a stable AAA formation rate of around 80% in male and female mice is achievable. The consistency and reproducibility of AAA studies using an elastase-induced AAA model can be significantly enhanced by establishing a standard surgical procedure.

The present protocol describes a standardized surgical method for the elastase-induced AAA model through the direct application of elastase to the adventitia of infrarenal abdominal aorta in mice.

Abdominal aortic aneurysm (AAA), although primarily asymptomatic, is potentially life-threatening as the rupture of AAA usually has a devastating outcome. ...

Abdominal Aortic Aneurysm Development in Mice Using the Elastase and BAPN Method

Advanced Abdominal Aortic Aneurysm Modeling in Mice by Combination of Topical Elastase and Oral &#223;-aminopropionitrile

The topical elastase murine model of abdominal aortic aneurysm (AAA) is enhanced when combined with &#223;-aminopropionitrile (BAPN)-supplemented drinking water to reliably produce true infrarenal aneurysms with behaviors that mimic human AAAs. Topically applying elastase to the adventitia of the infrarenal aorta causes structural damage to the elastic layers of the aortic wall and initiates aneurysmal dilation. Co-administering BAPN, a lysyl oxidase inhibitor, promotes sustained wall degeneration by reducing collagen and elastin crosslinking. This combination results in large AAAs that progressively expand, form intraluminal thrombus, and are capable of rupture. Refining surgical techniques, such as circumferentially isolating the entire infrarenal aortic segment, can help standardize the procedure for a consistent and thorough application of porcine pancreatic elastase despite different operators and anatomic variations between mice. Therefore, the elastase/BAPN model is a refined approach to surgically inducing AAA in mice, which may better recapitulate human aneurysms and provide additional opportunities to study aneurysm growth and rupture risk.

Author Spotlight: Enhanced Murine AAA Model Using Elastase to Mimic Human Aneurysms

This protocol describes a methodical surgical approach to modeling advanced abdominal aortic aneurysms in mice by a combination of directly applying elastase to the infrarenal aorta and administering &#223;-aminopropionitrile through drinking water.

The topical elastase murine model of abdominal aortic aneurysm (AAA) is enhanced when combined with &#223;-aminopropionitrile (BAPN)-supplemented drinking ...

Developing an Experimental Aneurysm Model in a Rabbit Using an Elastase-Treated Vessel Pouch

<a href='https://app.jove.com/v/61157/arterial-pouch-microsurgical-bifurcation-aneurysm-model-in-the-rabbit'>Source: Wanderer, S., et al. Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit. J. Vis. Exp. (2020)</a>This video demonstrates the creation of an experimental aneurysm model in a rabbit using an elastase-treated vessel pouch. After excising a segment of the right CCA, the pouch is treated with elastase, then sutured to the left CCA to restore blood flow. The resulting stress on the weakened pouch walls induces aneurysm formation.

Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm

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Chapters in this video

Creation of Murine Experimental Abdominal Aortic Aneurysms with Elastase

A New Murine Model of Endovascular Aortic Aneurysm Repair

Reproducable Paraplegia by Thoracic Aortic Occlusion in a Murine Model of Spinal Cord Ischemia-reperfusion

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

Surgical Models of Gastroesophageal Reflux with Mice

Creation of a Rodent Model of Abdominal Aortic Aneurysm by Blocking Adventitial Vasa Vasorum Perfusion

A Mouse Abdominal Aortic Aneurysm Model by Periadventitial Calcium Chloride and Elastase Infiltration

Mouse Abdominal Aortic Aneurysm Model Induced by Perivascular Application of Elastase

Advanced Abdominal Aortic Aneurysm Modeling in Mice by Combination of Topical Elastase and Oral ß-aminopropionitrile

Developing an Experimental Aneurysm Model in a Rabbit Using an Elastase-Treated Vessel Pouch