Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm

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Summary

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.

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Tyerman, Z., Dahl, J., Shannon, A., Johnston, W. F., Pope, N. H., Lu, G., Upchurch Jr., G. R., Ailawadi, G., Salmon, M. Murine Surgical Model of Topical Elastase Induced Descending Thoracic Aortic Aneurysm. J. Vis. Exp. (150), e60105, doi:10.3791/60105 (2019).

Abstract

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.

Introduction

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 >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. 

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Protocol

Animal protocols were approved by the University of Virginia Institutional Animal Care and Use Committee (No. 3634).

1. Induction of anesthesia and intubation

  1. 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.
  2. 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.

2. Securing the mouse to the surgical board

  1. Connect the endotracheal (ET) tube, confirm chest rise, and lay the mouse in the right lateral decubitus position. 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 µL of tidal volume.
    NOTE: Toxicity is rapid if isoflurane is left at 5%. 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.
  2. 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.
  3. 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.
  4. Tape the left leg in its natural position. Draw the left arm ventrally over rolled gauze and tape down.

3. Preparation for surgery

  1. Shave the left flank of the mouse with electric clippers from left shoulder to left abdomen.
  2. 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.
  3. CAUTION: Ensure that all ethanol is completely dried before proceeding as electrocautery is used and can ignite the ethanol.

4. Entry into thorax

  1. 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.
  2. 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.
  3. 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.
  4. Use scissors to cut along the top aspect of the cauterized rib, until diaphragm is visible.

5. Aortic exposure

  1. Place rib retractors and use them to open the thoracotomy incision. Carefully remove sponge from the surface of the lung.
  2. 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.
  3. 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).

6. Elastase exposure

  1. Saturate 0.5 mm x 1 mm sponge with 12 μL of porcine pancreatic elastase and place it upon the exposed surface of the aorta.
    NOTE: Do not let the sponge touch the contralateral lung.
  2. 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.
  3. Use rolled 2" x 2" gauze to absorb the remaining saline irrigation. Turn isoflurane down to 2%.

7. Closure of chest

  1. Remove the lung sponge. Remove the caudal lung retractor. Remove the rostral lung retractor.
  2. 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.
  3. 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.
  4. Tie non-absorbable sutures. Reapproximate the muscle layers with a running 5-0 absorbable multifilament suture. Turn isoflurane down to 1%.
  5. Close skin with 7-10 interrupted 5-0 non-absorbable sutures. Turn isoflurane to 0%.

8. Recovery

  1. Administer 6 µg (0.02 mL of a 0.3 mg/mL suspension)  of buprenorphine subcutaneously. Remove the tape from the right foot, tail, and left arm followed by the right arm tape.
  2. 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. 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.

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.

  1. 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.
  2. Incise skin from dorsal left flank rostrally to the level of the left shoulder. Then incise at a 90° angle through the axilla to the sternum.
    NOTE: This incision should completely encircle the original skin incision.
  3. 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.
  4. 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.
  5. 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.
  6. 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.
  7. 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.

    Equation 1              Equation 1
     
  8. 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.
  9. Using a tuberculin syringe and needle, wash the aortic lumen with saline and process tissue as desired. 

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Representative Results

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 et al.11 describing the model showed significant increase in aortic dilation in wild type elastase treated mice (WT TAA) at 3, 7, 14, and 21 days compared to controls in wild type mice (WT controls). Maximal aortic dilation occurred at day 14 (WT TAA: 99.6 ± 24.7% versus WT control: 14.4 ± 8.2%; p < 0.0001) (Figure 3)11. A separate experiment by Pope et al.12 shows similar dilation at 14 days.

When examining histology, WT TAAs have elastin fibers that are thinned and fragmented. There is also less smooth muscle cell (SMαA) staining, while macrophages (Mac2) and interleukin-1β (IL-1β) expression are increased (Figure 4).

Figure 1
Figure 1: Sample photographs of the thoracic aortic aneurysm (TAA) model and equipment setup. From Left to Right, (1) initial exposure of the thoracic aorta through a left thoracotomy, (2) dissection of the pleura, (3) application of an elastase-soaked sponge, and (4) sponge removal. For aortic harvest (5), the thoracotomy was reopened, and thoracic aorta was dissected free and measured. (6) Illustration to the right demonstrates relative association of aorta with esophagus, lung, and hemiazygos vein. This figure has been adapted from Johnston et al.11. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Sample video micrometry measurement of TAA. C = unexposed control segment, A = elastase treated aneurysmal segment, RL = right lung, E = esophagus, T = tail, H = head. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Aortic diameter over time during TAA formation. Mean aortic diameter (mm ± SD) in WT mice exposed to elastase (WT TAA, gray) or exposed to saline (WT control, green) over time. Significant differences occurred between WT TAA and WT control at all time points marked with asterisk (p < 0.05). Maximal aortic dilation occurs at day 14 (WT TAA: 99.6 ± 24.7% versus WT control: 14.4 ± 8.2%; p < 0.0001). Error bars represent standard deviation. This figure has been adapted from Johnston et al.11. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Histological examination of WT elastase TAA and WT saline control tissue. Sample aortic cross-sections of WT control (saline sponge) and WT TAAs (elastase sponge) on day 14. Staining for elastin, smooth muscle cells (SMαA), macrophages (Mac2), and interleukin-1β (IL-1β). Scale bar = 50 μm. This figure has been adapted from Johnston et al.11. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Dose-response examination of individual elastase bottle. Each group of points represents a pre-specified time when the elastase-soaked sponge is allowed to dwell. These data are used to estimate the ideal digestion time for 100-130% dilation. Please click here to view a larger version of this figure.

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Discussion

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 relatively quick to execute and results in maximal dilation at 14 days as opposed to 16 weeks in the CaCl2 model8. This efficiency has facilitated the study of specific pharmacologic interventions which could potentially be translatable to human trials11,12. However, this protocol has several challenging aspects and limitations.

Our model utilizes purified porcine pancreatic elastase, which has been used in several previous AAA models. The mechanism of action is believed to be breakdown of elastin in the media of the aorta and creation of a robust inflammatory response, leading to a less compliant and eventually weaker wall. The digestive effect of elastase varies from bottle to bottle. To ensure a consistent digestive effect, a dose-response curve should be undertaken for each new bottle of elastase. The dose corresponds to the time during which the elastase-soaked gauze is allowed to dwell. An example of such a curve is shown in Figure 5. Specific elastase sponge exposure times are examined with the goal to find an exposure duration which creates aneurysms of 100-130% dilation at 14 days. In this example curve, a 4-minute digestion time is designated for this bottle of elastase (of note, the smallest quantity of elastase available is in 10 mL, which is enough for approximately 830 individual mice). Excessive elastase digestion can cause intraoperative bleeding or premature rupture and insufficient exposure can fail to cause sufficient digestion and aneurysmal dilation.

Because this model involves opening the thoracic cavity, the use of positive pressure ventilation is mandatory. Initially we utilized an anterior neck dissection and orotracheal intubation under direct vision, but this method was time consuming as it required an additional incision and closure. We now utilize a direct orotracheal intubation as detailed by Vandivort et al.13. When training new surgeons on this model, we recommend practicing the intubation technique extensively to master it before moving on to the actual surgery. Once the mouse is successfully intubated, great care should be taken not to dislodge the tube as edema increases with each intubation attempt, successively diminishing the chances of success and increasing the chances of a fatal airway injury. Great care should also be taken to minimize the risk of extubation during the surgery since the time required for reintubation is almost invariably fatal. As detailed in our protocol, securing the right arm and tail under a small amount of tension will prevent rostral or caudal shifting during surgery and therefore minimize the risk of extubation.

Control of anesthesia is vital in this model. As Vandivort et al.13 discussed in their study on intubation, a deep level of sedation is necessary. If the mouse is conscious during the intubation procedure, resistance to ET tube insertion can cause tracheal injury and esophageal intubation. To reduce the risk of isoflurane toxicity, once tracheal placement of the ET tube has been confirmed, the percent mixture of isoflurane should be reduced by approximately half that of used for induction. To achieve the ideal percentage of isoflurane, the mouse should have a spontaneous diaphragmatic contraction every 10-20 seconds. As the surgery progresses, the isoflurane should be progressively lowered and completely turned off after the last skin closure stitch is placed. The tape restraints should be removed while the mouse is still recovering from anesthesia as removal while the mouse is awake can be traumatic. To assist with recovery and dissipation of residual isoflurane we recently modified the procedure such that the mouse is recovered in a high oxygen warming environment immediately after extubation.

The mouse lung is extremely delicate and injury from puncture, inappropriate handling, or elastase exposure almost always results in death. The most common point of lung injury is the initial entry into the pleura. When using cautery on the ribs (step 4.2), we advise paying close attention to the spontaneous breathing frequency and only activating the cautery for a few seconds immediately after a breath. This helps to prevent an unexpected expansion of the ribcage which can drive the cautery tip through the chest wall and into the lung. The use of the wetted cotton-tipped applicator in the next step to gently break the tissues and the pleural membrane also helped minimize our rate of lung injury since sharp instruments are not meeting the lung surface. Finally, if the elastase-soaked sponge contacts either lung directly, severe damage can occur. This can be avoided by not entering the contralateral pleural space during aortic dissection and readying the sponge so that it can be placed directly on the aorta with little repositioning. At the end of the surgery, an attempt to re-inflate the left lung should be made by occluding the outflow tube of the ventilator. If that is unsuccessful, direct injection of 0.5 to 1.0 mL of air through the 3-way stopcock can re-expand the lung. If both of those methods do not work, closure should proceed as normal. We have found that if no major lung injury is present, mice tolerate a partially inflated lung relatively well and the lung is almost always inflated at tissue harvest.

This model has several limitations. Experimental models do not fully model human aortic disease. dTAA occurs in 14 days after one application of elastase in our model while human TAA formation is multifactorial and occurs over years if not decades. However, a mouse model that takes months to years to develop aneurysms is not useful in an investigative capacity. Unlike human disease, elastase induced aneurysms in our model begin to decrease in size when they reach maximal dilatation at two weeks post-operatively. This regression can be overcome with the use of β-aminopropionitrile (BAPN) supplemented in the mouse’s diet postoperatively. When combined with the elastase exposure model, BAPN supplementation allows continual growth and eventually rupture, better mimicking the chronicity of human disease. Our lab has previously studied BAPN in AAA17 and we are currently using it in the dTAA model to further refine our protocol. Primary experiments with the dTAA model have demonstrated rupture rates of approximately 30% at 28 days.

In the future, we plan to use this model to evaluate other genetic and pharmacological treatments that may be translatable to aneurysm treatment in humans. Because this protocol describes an easy and safe way to access the thoracic aorta, it can be used by other labs that wish to test interventions on the thoracic aorta.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

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.

Materials

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

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References

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  3. Bickerstaff, L. K., et al. Thoracic aortic aneurysms: a population-based study. Surgery. 92, (6), 1103-1108 (1982).
  4. Cheng, D., et al. Endovascular aortic repair versus open surgical repair for descending thoracic aortic disease a systematic review and meta-analysis of comparative studies. Journal of the American College of Cardiology. 55, (10), 986-1001 (2010).
  5. Walsh, S. R., et al. Endovascular stenting versus open surgery for thoracic aortic disease: systematic review and meta-analysis of perioperative results. Journal of Vascular Surgery. 47, (5), 1094-1098 (2008).
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  10. Trachet, B., et al. Ascending Aortic Aneurysm in Angiotensin II-Infused Mice: Formation, Progression, and the Role of Focal Dissections. Arteriosclerosis, Thrombosis, and Vascular Biology. 36, (4), 673-681 (2016).
  11. Johnston, W. F., et al. Inhibition of interleukin-1beta decreases aneurysm formation and progression in a novel model of thoracic aortic aneurysms. Circulation. 130, (11), Suppl 1 51-59 (2014).
  12. Pope, N. H., et al. Interleukin-6 Receptor Inhibition Prevents Descending Thoracic Aortic Aneurysm Formation. Annals of Thoracic Surgery. 100, (5), 1620-1626 (2015).
  13. Vandivort, T. C., An, D., Parks, W. C. An Improved Method for Rapid Intubation of the Trachea in Mice. Journal of Visualized Experiments. (108), e53771 (2016).
  14. Ailawadi, G., et al. A nonintrinsic regional basis for increased infrarenal aortic MMP-9 expression and activity. Journal of Vascular Surgery. 37, (5), 1059-1066 (2003).
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  16. Trigueros-Motos, L., et al. Embryological-origin-dependent differences in homeobox expression in adult aorta: role in regional phenotypic variability and regulation of NF-kappaB activity. Arteriosclerosis, Thrombosis, and Vascular Biology. 33, (6), 1248-1256 (2013).
  17. Lu, G. S. G., Davis, J. P., Schaheen, B., Downs, E., Roy, R. J., Ailawadi, G., Upchurch, G. R. A novel chronic advanced staged abdominal aortic aneurysm murine model. Journal of Vascular Surgery. 66, (1), 232-242 (2017).

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