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>

<ol>
	<li>Coady, M. 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=What+is+the+appropriate+size+criterion+for+resection+of+thoracic+aortic+aneurysms.">What is the appropriate size criterion for resection of thoracic aortic aneurysms.</a> Journal of Thoracic and Cardiovascular Surgery. 113 (3), 489-491 (1997).</li><li>Aggarwal, S., Qamar, A., Sharma, V., Sharma, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abdominal+aortic+aneurysm:+A+comprehensive+review.">Abdominal aortic aneurysm: A comprehensive review.</a> Experimental and Clinical Cardiology. 16 (1), 11-15 (2011).</li><li>Bickerstaff, L. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thoracic+aortic+aneurysms:+a+population-based+study.">Thoracic aortic aneurysms: a population-based study.</a> Surgery. 92 (6), 1103-1108 (1982).</li><li>Cheng, D., 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+aortic+repair+versus+open+surgical+repair+for+descending+thoracic+aortic+disease+a+systematic+review+and+meta-analysis+of+comparative+studies.">Endovascular aortic repair versus open surgical repair for descending thoracic aortic disease a systematic review and meta-analysis of comparative studies.</a> Journal of the American College of Cardiology. 55 (10), 986-1001 (2010).</li><li>Walsh, S. 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. S., 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=Altered+patterns+of+gene+expression+distinguishing+ascending+aortic+aneurysms+from+abdominal+aortic+aneurysms:+complementary+DNA+expression+profiling+in+the+molecular+characterization+of+aortic+disease.">Altered patterns of gene expression distinguishing ascending aortic aneurysms from abdominal aortic aneurysms: complementary DNA expression profiling in the molecular characterization of aortic disease.</a> The Journal of Thoracic and Cardiovascular Surgery. 126 (2), 344-357 (2003).</li><li>Ailawadi, G., Eliason, J. L., Upchurch, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+concepts+in+the+pathogenesis+of+abdominal+aortic+aneurysm.">Current concepts in the pathogenesis of abdominal aortic aneurysm.</a> Journal of Vascular Surgery. 38 (3), 584-588 (2003).</li><li>Ikonomidis, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+murine+model+of+thoracic+aortic+aneurysms.">A murine model of thoracic aortic aneurysms.</a> Journal of Surgical Research. 115 (1), 157-163 (2003).</li><li>Daugherty, A., Manning, M. W., Cassis, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Angiotensin+II+promotes+atherosclerotic+lesions+and+aneurysms+in+apolipoprotein+E-deficient+mice.">Angiotensin II promotes atherosclerotic lesions and aneurysms in apolipoprotein E-deficient mice.</a> Journal of Clinical Investigation. 105 (11), 1605-1612 (2000).</li><li>Trachet, B., 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=Ascending+Aortic+Aneurysm+in+Angiotensin+II-Infused+Mice:+Formation,+Progression,+and+the+Role+of+Focal+Dissections.">Ascending Aortic Aneurysm in Angiotensin II-Infused Mice: Formation, Progression, and the Role of Focal Dissections.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 36 (4), 673-681 (2016).</li><li>Johnston, W. F., 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=Inhibition+of+interleukin-1beta+decreases+aneurysm+formation+and+progression+in+a+novel+model+of+thoracic+aortic+aneurysms.">Inhibition of interleukin-1beta decreases aneurysm formation and progression in a novel model of thoracic aortic aneurysms.</a> Circulation. 130 (11), 51-59 (2014).</li><li>Pope, N. H., 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=Interleukin-6+Receptor+Inhibition+Prevents+Descending+Thoracic+Aortic+Aneurysm+Formation.">Interleukin-6 Receptor Inhibition Prevents Descending Thoracic Aortic Aneurysm Formation.</a> Annals of Thoracic Surgery. 100 (5), 1620-1626 (2015).</li><li>Vandivort, T. C., An, D., Parks, W. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+Improved+Method+for+Rapid+Intubation+of+the+Trachea+in+Mice.">An Improved Method for Rapid Intubation of the Trachea in Mice.</a> Journal of Visualized Experiments. (108), e53771 (2016).</li><li>Ailawadi, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+nonintrinsic+regional+basis+for+increased+infrarenal+aortic+MMP-9+expression+and+activity.">A nonintrinsic regional basis for increased infrarenal aortic MMP-9 expression and activity.</a> Journal of Vascular Surgery. 37 (5), 1059-1066 (2003).</li><li>Ruddy, J. M., Jones, J. A., Spinale, F. G., Ikonomidis, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regional+heterogeneity+within+the+aorta:+relevance+to+aneurysm+disease.">Regional heterogeneity within the aorta: relevance to aneurysm disease.</a> Journal of Thoracic and Cardiovascular Surgery. 136 (5), 1123-1130 (2008).</li><li>Trigueros-Motos, L., 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=Embryological-origin-dependent+differences+in+homeobox+expression+in+adult+aorta:+role+in+regional+phenotypic+variability+and+regulation+of+NF-kappaB+activity.">Embryological-origin-dependent differences in homeobox expression in adult aorta: role in regional phenotypic variability and regulation of NF-kappaB activity.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 33 (6), 1248-1256 (2013).</li><li>Lu, G. S. G., Davis, J. P., Schaheen, B., Downs, E., Roy, R. J., Ailawadi, G., Upchurch, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+chronic+advanced+staged+abdominal+aortic+aneurysm+murine+model.">A novel chronic advanced staged abdominal aortic aneurysm murine model.</a> Journal of Vascular Surgery. 66 (1), 232-242 (2017).</li></ol>

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 border="0" cellpadding="0" cellspacing="0" style="width:761px;" width="760">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Angiocatheter (22G)</td>
			<td style="width:191px;"></td>
			<td style="width:161px;"></td>
			<td style="width:167px;">Used for ET Tube</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Dumont Tweezers; Pattern #7 x2</td>
			<td style="width:191px;">Roboz</td>
			<td style="width:161px;">RS-4982</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Graefe Tissue Forceps</td>
			<td style="width:191px;">Roboz</td>
			<td style="width:161px;">RS-5158</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Harms Forceps x2</td>
			<td style="width:191px;">Roboz</td>
			<td style="width:161px;">RS-5097</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:243px;">Intracardiac Needle Holder; Extra Delicate; Carbide Jaws; 7&#34; Length</td>
			<td style="width:191px;">Roboz</td>
			<td style="width:161px;">RS-7800</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">&#160;KL 1500 LED Light Source</td>
			<td style="width:191px;">Leica</td>
			<td style="width:161px;">150-400</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">&#160;M205A Dissction Microscope</td>
			<td style="width:191px;">Leica</td>
			<td style="width:161px;">CH 94-35</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:243px;">Iris Scissors, 11cm, Tungsten Carbide</td>
			<td style="width:191px;">World Precision Instruments</td>
			<td style="width:161px;">500216-G</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:243px;">Metal Clip board</td>
			<td style="width:191px;"></td>
			<td style="width:161px;"></td>
			<td style="width:167px;">Use with the Mouse Retractor Set&#160;</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:243px;">Mouse Retractor Set</td>
			<td style="width:191px;">Kent</td>
			<td style="width:161px;">SURGI-5001</td>
			<td style="width:167px;">Need 2 short and 1 tall fixators</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:243px;">Mouse Ventilator MiniVent Type 845, 115 V, Power Supply with US Connector</td>
			<td style="width:191px;">Harvard Apparatus</td>
			<td style="width:161px;">73-0043</td>
			<td style="width:167px;">MiniVent Ventilator for Mice (Model 845), Single Animal, Volume Controlled</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:243px;">Sigma Aldrich</td>
			<td style="width:191px;">Elastase from porcine pancreas</td>
			<td style="width:161px;">E0258-50MG</td>
			<td style="width:167px;">Can be purchased in various size bottles</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:243px;">Small Vessel Cauterizer Kit</td>
			<td style="width:191px;">Fine Science Tools</td>
			<td style="width:161px;">18000-00</td>
			<td style="width:167px;">Recommend using rechargable AA batteries</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:243px;">Spring Scissors, 10.5cm</td>
			<td style="width:191px;">World Precision Instruments</td>
			<td align="right" style="width:161px;">14127</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Steril Swabs (Sponges)</td>
			<td style="width:191px;">Sugi</td>
			<td align="right" style="width:161px;">31603</td>
			<td style="width:167px;">Can be cut to size</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Surgi Suite Surgical Platform</td>
			<td style="width:191px;">Kent</td>
			<td style="width:161px;"></td>
			<td style="width:167px;">Attach to clip board&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Tech IV Isoflurane Vap</td>
			<td style="width:191px;">Jorgensen&#160;Laboratories&#160;</td>
			<td style="width:161px;">J0561A</td>
			<td style="width:167px;">Anesthesia vaporizer&#160;</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 ...

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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. ...

murine-surgical-model-topical-elastase-induced-descending-thoracic

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JoVE Journal

Medicine

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

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 ...

Murine Left Anterior Descending (LAD) Coronary Artery Ligation: An Improved and Simplified Model for Myocardial Infarction

Ischemic heart disease (IHD), or acute coronary syndrome (ACS), is one of the leading causes of death in the United States. IHD is characterized by reduced blood supply to the heart, resulting in the loss of oxygen to and the ensuing necrosis of the heart muscle. The MI model has gained popularity for its use as a short-term ischemia-reperfusion model and a long-term permanent ligation model. Below, we describe a reliable method for the permanent ligation of the LAD. With mouse genetic engineering technology becoming more advanced, and with an increasing availability of quality murine surgical instruments, the mouse has become a popular model for MI surgeries. Our surgical model incorporates the use of an easily reversible anesthetic for the rapid recovery of the mouse; a minimally invasive endotracheal intubation without involving a tracheotomy; and a thoracentesis through the original thoracotomy site without creating an additional incision in the chest, as is done in some other methods, to effectively remove excess blood and air from the chest cavity. This method is comparatively less invasive than other methods, which dramatically reduces surgical and post-surgical complications and mortality and improves reproducibility.

Murine Left Anterior Descending LAD Coronary Artery Ligation: An Improved and Simplified Model for Myocardial Infarction

We provide a reliable method for left anterior descending artery (LAD) ligation in a mouse model. This method is comparatively less invasive than other methods, involving endotracheal intubation, a left-sided thoracotomy approach, and thoracentesis. This method can be used as a model for both acute and chronic myocardial infarction (MI).

Ischemic heart disease (IHD), or acute coronary syndrome (ACS), is one of the leading causes of death in the United States. IHD is characterized by ...

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 ...

Quantitative Micro-CT Analysis of Aortopathy in a Mouse Model of &#946;-aminopropionitrile-induced Aortic Aneurysm and Dissection

Aortic aneurysm and dissection is associated with significant morbidity and mortality in the population and can be highly lethal. While animal models of aortic disease exist, in vivo imaging of the vasculature has been limited. In recent years, micro-computerized tomography (micro-CT) has emerged as a preferred modality for imaging both large and small vessels both in vivo and ex vivo. In conjunction with a method of vascular casting, we have successfully used micro-CT to characterize the frequency and distribution of aortic pathology in &#946;-aminopropionitrile-treated C57/Bl6 mice. Technical limitations of this method include variations in the quality of the perfusion introduced by poor animal preparation, the application of proper methodologies for vessel size quantification, and the non-survivability of this procedure. This article details a methodology for the intravascular perfusion of a lead-based radiopaque silicone rubber for the quantitative characterization of aortopathy in a mouse model of aneurysm and dissection. In addition to visualizing aortic pathology, this method may be used for examining other vascular beds in vivo or vascular beds removed post-mortem.

Quantitative Micro-CT Analysis of Aortopathy in a Mouse Model of &amp;#946;-aminopropionitrile-induced Aortic Aneurysm and Dissection

This article describes a detailed methodology of using a radiopaque lead-based silicone rubber to perfuse the murine vasculature for aortic diameter quantification in a mouse model of aortic aneurysm and dissection.

Aortic aneurysm and dissection is associated with significant morbidity and mortality in the population and can be highly lethal. While animal models of ...

Ultrasound Imaging of the Thoracic and Abdominal Aorta in Mice to Determine Aneurysm Dimensions

Contemporary high-resolution ultrasound instruments have sufficient resolution to facilitate the measurement of mouse aortas. These instruments have been widely used to measure aortic dimensions in mouse models of aortic aneurysms. Aortic aneurysms are defined as permanent dilations of the aorta, which occur most frequently in the ascending and abdominal regions. Sequential measurements of aortic dimensions by ultrasound are the principal approach for assessing the development and progression of aortic aneurysms in vivo. Although many reported studies used ultrasound imaging to measure aortic diameters as a primary endpoint, there are confounding factors, such as probe position and cardiac cycle, that may impact the accuracy of data acquisition, analysis, and interpretation. The purpose of this protocol is to provide a practical guide on the use of ultrasound to measure the aortic diameter in a reliable and reproducible manner. This protocol introduces the preparation of mice and instruments, the acquisition of appropriate ultrasound images, and data analysis.

Ultrasound imaging has become a common modality to determine the luminal dimensions of thoracic and abdominal aortic aneurysms in mice. This protocol describes the procedure to acquire reliable and reproducible two-dimensional ultrasound images of the ascending and abdominal aorta in mice.

Contemporary high-resolution ultrasound instruments have sufficient resolution to facilitate the measurement of mouse aortas. These instruments have been ...

Murine Myocardial Infarction Model using Permanent Ligation of Left Anterior Descending Coronary Artery

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine models of MI with permanent ligation of left-anterior descending (LAD) coronary artery closely mimics MI in humans. Murine models benefit from the extensive genetic engineering available nowadays. Here we propose a reproducible murine surgical model of myocardial infarction by permanent LAD coronary ligation. Our technique comprises anesthesia with ketamine/xylazine that can be rapidly reversed by administration of an antagonist, intubation without tracheotomy for mechanical-assisted ventilation, ventilation with application of extrinsic positive end-expiratory pressure (PEEP) to avoid alveolar collapse, a thoracotomy method limiting to the minimum surgical lesions made to skeletal muscles, and lung inflation without thoracentesis. This method is sparsely invasive, reproducible and reduces post-surgery mortality and complications.

Herein we describe a surgical procedure showing how to achieve permanent ligation of the left-anterior descending coronary artery in mice. This model is of high relevance to investigate the pathophysiology of myocardial infarction and the concomitant biological processes.

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine ...

Porcine Model of Infrarenal Abdominal Aortic Aneurysm

Large animal models to study abdominal aortic aneurysms are sparse. The purpose of this model is to create reproducible, clinically significant infrarenal abdominal aortic aneurysms (AAA) in swine. To achieve this, we use a combination of balloon angioplasty, elastase and collagenase, and a lysyl oxidase inhibitor, called &#946;-aminopropionitrile (BAPN), to create clinically significant infrarenal aortic aneurysms, analogous to human disease.
Noncastrated male swine are fed BAPN for 7 days prior to surgery to achieve a steady state in the blood. A midline laparotomy is performed and the infrarenal aorta is circumferentially dissected. An initial measurement is recorded prior to aneurysm induction with a combination of balloon angioplasty, elastase (500 units)/collagenase (8000 units) perfusion, and topical elastase application. Swine are fed BAPN daily until terminal procedure on either postoperative day 7, 14, or 28, at which time the aneurysm is measured, and tissue procured. BAPN + surgery pigs are compared to pigs that underwent surgery alone.
Swine treated with BAPN and surgery had a mean aortic dilation of 89.9% &#177; 47.4% at day 7, 105.4% &#177; 58.1% at day 14, and 113.5% &#177; 30.2% at day 28. Pigs treated with surgery alone had significantly smaller aneurysms compared to BAPN + surgery animals at day 28 (p &#60; 0.0003). The BAPN + surgery group had macroscopic and immunohistochemical evidence of end stage aneurysmal disease.
Clinically significant infrarenal AAA can be induced using balloon angioplasty, elastase/collagenase perfusion and topical application, supplemented with oral BAPN. This model creates large, clinically significant AAA with hallmarks of human disease. This has important implications for the elucidation of AAA pathogenesis and testing of novel therapies and devices for the treatment of AAA. Limitations of the model include variation in BAPN ingested by swine, quality of elastase perfusion, and cost of BAPN.&#160;

This novel model creates robust infrarenal abdominal aortic aneurysms in swine using a combination of balloon angioplasty, elastase/collagenase perfusion, topical elastase application, and oral compound &#946;-aminopropionitrile administration, which interferes with collagen cross-linking.

Large animal models to study abdominal aortic aneurysms are sparse. The purpose of this model is to create reproducible, clinically significant infrarenal ...

Drug Treatment by Central Venous Catheter in a Mouse Model of Angiotensin II Induced Abdominal Aortic Aneurysm and Monitoring by 3D Ultrasound

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse models, are applied to advance the understanding of the disease pathogenesis and to identify potential therapeutic targets. Testing novel drug candidates to block AAA growth in these models generally requires repeated drug administration during the time course of the experiment. Here, we describe a compiled protocol for AAA induction, insertion of an intravenous catheter to facilitate prolonged therapy, and serial AAA monitoring by 3D ultrasound. Aneurysms are induced in apolipoprotein E (ApoE) deficient mice by angiotensin II release over 28 days from osmotic mini-pumps implanted subcutaneously into the mouse back. Subsequently, the surgical procedure for external jugular vein catheterization is conducted to allow for daily intravenous drug treatment or repeated blood sampling via a subcutaneous vascular access button. Despite the two dorsal implants, the monitoring of AAA development is readily facilitated by sequential semi-automated 3D ultrasound analysis, which yields comprehensive information on the expansion of aortic diameter and volume and on aneurysm morphology, as illustrated by experimental examples.

This protocol describes the consecutive implantation of an osmotic pump to induce abdominal aortic aneurysm by angiotensin II release in apolipoprotein E (ApoE) deficient mice and of a vascular access port with a jugular vein catheter for repeated drug treatment. Monitoring of aneurysm development by 3D ultrasound is effectively conducted despite dorsal implants.

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse ...