Reliable Creation of a Murine Abdominal Aortic Aneurysm Model Using Elastin Sensitization and Perivascular Elastase

Summary

This protocol provides a repeatable and reliable method for inducing abdominal aortic aneurysms (AAAs) in C57BL/6 mice using elastin sensitization and perivascular elastase application, as an AAA model that incorporates an immune-sensitization component.

Abstract

Degradation of the elastic lamellar matrix in the abdominal aortic wall is a histopathologic feature of abdominal aortic aneurysms (AAA). Growing evidence implicates elastin degradation and related host immune responses in the pathogenesis of AAA. Effective modeling of these processes is important for improving the understanding and treatment of this common disease. While previous AAA models have used elastin-disrupting techniques for aneurysm induction, the single use of elastase in many of these models does not account for the ongoing immune response against elastin and its degradation products (EDP). To model this ongoing immune response, we developed a model that sensitizes C57BL/6 mice to EDP by subcutaneously injecting EDP 7 days prior to aneurysm induction surgery. Aneurysm induction surgery proceeds with exposure of the abdominal aorta and application of topical elastase in the periaortic compartment, with the intention to expose elastin and EDP on the mouse aorta. With these moieties exposed, the animal should mount an immune response against these antigens to which it was previously sensitized. Repeat “booster” EDP injection 7 days later serves to propagate this immune response. Herein, we describe the protocol for the reliable creation of AAA in C57BL/6 mice via multiple sensitizations with EDP, followed by AAA induction surgery with perivascular elastase application. This model can be created quickly and at moderate cost, while inducing >30% aortic dilation in 82% of mice by video micrometry and 73% by ultrasound measurement, and >50% dilation in 64% by video micrometry and 27% by ultrasound, as measured at day 21 post-induction surgery. This technique is also associated with a 95% intraoperative survival rate.

Introduction

Abdominal aortic aneurysm (AAA) affects >35 million people worldwide, and 4–8% of men aged 65-80^1,2. Rupture, the most devastating complication of AAA, typically occurs when the mechanical wall stress of the aneurysm exceeds the strength of the aortic tissue, leading to massive hemorrhage and an overall mortality rate of 80−90%^3,4. Current management is aimed at prevention, with ultrasound screening of high-risk individuals and surgical repair when the risk of rupture exceeds the risk of repair, typically at 5.5 cm in diameter⁵. However, most AAAs are asymptomatic, and many are diagnosed incidentally⁶. As such, efforts should be made to identify high-risk patients and develop non-surgical treatments to reduce AAA growth and, therefore, the risk of rupture.

To develop medical treatments for AAA, the pathogenesis must be better understood. Several experimental models are currently used, including small-animal models utilizing angiotensin II infusion⁷, xenografting⁸, calcium chloride application⁹, and infusion¹⁰ and/or topical application of elastase¹¹. Each of these models uses a different understanding of the pathologic basis of AAA, and most have been modified, improved, or combined with other methods since inception.

Importantly, AAA histology demonstrates loss of the normal lamellar elastin matrix, which is also seen in lung tissue in chronic obstructive pulmonary disease^12,13. Elastin degradation via matrix metalloproteinase activity provides an important linkage between AAA pathogenesis and smoking, the most important modifiable risk factor for AAA formation^12,14. Notably, reduced aortic elastin, elastin fiber disruption, and increased circulating elastin peptides are observed in human AAA and are associated with increased AAA wall distensibility^15,16. Therefore, stimulating elastin degradation is crucial in creating a translatable AAA model. Existing AAA models that rely on elastin degradation, while effective, can be complicated by high animal mortality rates, especially in models that use aortic cannulation¹⁷, and by inconsistent aneurysm formation, particularly in models that rely solely on topical elastase application¹⁸. Furthermore, all models relying on induction surgeries are subject to operator learning curves and technical variation.

To address these issues, we leverage the host immune response to sensitize animals to elastin by injecting elastin degradation products (EDP). One week later, aneurysm induction surgery occurs in a standardized stepwise fashion, using periaortic application of elastase to create EDP moieties for the sensitized host to mount an immune response against. Subsequent “booster” EDP sensitization, a week after induction surgery, ensures the host immune response continues. Repetitive immune sensitization enables reliable aortic dilation. We utilize a murine AAA model due to lower cost, ease of maintenance, rapid disease development, and technical consistency.

This model does not reliably result in aortic rupture and is therefore not intended for the study of factors associated with AAA rupture. However, this model does reliably produce progressive aortic dilation to 21 days post-induction surgery. As such, the model is best suited to studying therapeutic modalities that may reduce or reverse aortic dilation as the aneurysm is forming.

Protocol

1. Ultrasound imaging

1. Prepare the mouse
   1. Place the mouse into the anesthesia induction chamber.
   2. Administer inhaled isoflurane at 1.8−2% and oxygen until the mouse is anesthetized. Ensure that anesthetic gas canisters are connected to the anesthesia apparatus to reduce operator exposure to volatile anesthetics when isoflurane is used.
   3. Place the mouse in supine position on a stage heated to 37 °C with the mouse’s nose placed in the anesthesia nose cone.
   4. Tape the hindlimbs to the stage with labeling tape.
   5. Apply a hair removal agent using a cotton-tipped applicator to the abdomen and wait 30 seconds.
   6. Remove the abdominal hair using a 5 cm x 5 cm cotton gauze pad.
   7. Place warmed ultrasound transmission gel on the mouse’s abdomen.
2. Place a 38 MHz ultrasound probe on the abdomen and identify the pelvic girdle.
3. Identify the aorta as a tubular midline structure that is incompressible with moderate posteriorly directed pressure applied through the ultrasound probe.
   NOTE: The inferior vena cava (IVC) will appear as a luminal structure to the mouse’s right of the aorta and is compressible with moderate posteriorly directed pressure applied through the ultrasound probe.
4. Measure the anterior-posterior diameter of the abdominal aorta just cephalad to the point of bifurcation.
5. Move the probe proximally along the aorta and identify the left renal vein crossing anterior to the aorta and entering the IVC.
6. Measure the anterior-posterior diameter of the abdominal aorta just caudad to the point where the left renal vein crosses anteriorly to the abdominal aorta.
7. Move the probe proximally and identify the diaphragm.
   NOTE: The diaphragm can be identified as a hyperechoic band just cephalad to the liver.
8. Measure the anterior-posterior diameter of the abdominal aorta just caudad to the point where it enters the abdomen.
9. Place the mouse in its cage or in a recovery basket that is lined with towels and placed on a stage heated to 37 °C.
10. Monitor the mouse until it emerges from anesthesia.

2. Initial elastin sensitization

1. Prepare the elastin degradation products (EDP) while wearing personal protective equipment, including gloves, eye protection, and facemasks.
   1. Obtain a formulation of murine pancreatic elastase in PBS suspension at 0.475 units/mL (4.75 units/mg protein).
   2. Take 168 μL of the elastase and add it to 5 mg of murine elastin. Add another 200 μL of 1X phosphate-buffered saline (PBS) to this mixture, then pipette to mix.
   3. Incubate the elastase and elastin mixture at 37 °C for 20 minutes with intermittent gentle agitation.
   4. Add 632 μL cold 1X PBS to the elastase and elastin mixture.
   5. Add 1 mL of complete Freund’s adjuvant.
2. Place the mouse into the anesthesia induction chamber.
3. Administer inhaled isoflurane at 1.8−2% and oxygen until the mouse is anesthetized.
4. Place the mouse in a prone position on a stage heated to 37 °C with the mouse’s nose placed in the anesthesia nose cone.
5. Inject 50 μL of EDP subcutaneously into each flank for a total of 100 μL EDP in each mouse.
6. Place the mouse in a recovery basket that is lined with towels and placed on a stage heated to 37 °C.
7. Monitor the mouse until it emerges from anesthesia.

3. Preparation for surgery

1. Prepare the surgical tools.
   1. Assemble a surgical kit with 2 toothed forceps, 1 sharp straight forceps, 1 sharp angled forceps, 1 needle driver, scissors, at least 3 self-retaining magnetic retractors, 1 skin stapler, and 1 polypropylene bottle cap.
   2. Add 4-0 polyglycolic acid braided suture to the kit in a sterile fashion immediately prior to surgery.
   3. NOTE: 3-0 or 5-0 suture sizes will also suffice.
   4. Fill the polypropylene bottle cap with sterile warmed 1X PBS.
      NOTE: A bottle of sterile 1X PBS is kept in a warmer at 37 °C for refills.
2. Prepare the elastase.
   1. Dilute 27 μL of 5.1 mg/mL (≥4.0 units/mg protein) porcine pancreatic elastase into 73 μL of 1X PBS for a final volume of 100 μL for surgery on 10 mice (10 μL/mouse).
   2. Prepare the surgical suite.
   3. Autoclave a beaker of cotton-tipped applicators, tapered-tip cotton-tipped applicators, and 5 cm x 5 cm gauze, and place them next to the surgical trays prior to surgery.
   4. Place a magnetic surgical stage on a heating pad set to 37 °C on a microscope stage. Tape the heating pad into place on the microscope stage, but leave the magnetic component untethered to allow for repositioning during surgery.
   5. Place an absorbent pad or paper towel on the heating pad. Fold a 5 cm x 5 cm gauze in half and place it under the anesthesia nose cone.
   6. Place a 30 mL cup filled with hair removal agent, a 30 mL cup filled with 5% povidone-iodine, a tube of eye lubricant, surgical tape, non-sterile 5 cm x 5 cm gauze, non-sterile cotton tipped applicators, sealed 5 cm x 5 cm alcohol swabs, a roll of adhesive food service film, and a spray bottle of 70% ethanol near the surgical area.
   7. Draw extended release 1.3 mg/mL buprenorphine into a 1 mL syringe at a volume of 75 μL per mouse, for a dose of 3.25 mg/kg in a 30 g mouse.
3. Prepare the mouse for surgery.
   1. Measure and record the mouse’s weight in grams.
   2. Place the mouse into the anesthesia induction chamber.
   3. Administer inhaled isoflurane at 1.8−2% and oxygen until the mouse is anesthetized.
   4. Place the mouse in supine position on a stage heated to 37 °C with the mouse’s nose placed in the anesthesia nose cone.
   5. Inject 75 μL of buprenorphine subcutaneously.
   6. Tape the hindlimbs to the stage with labeling tape.
   7. Apply a hair removal agent using a cotton-tipped applicator to the abdomen and wait 30 seconds.
   8. Remove the abdominal hair using a 5 cm x 5 cm cotton gauze pad.
   9. Apply eye lubricant to the mouse’s eyes.
   10. Apply povidone-iodine to the mouse’s abdomen using a 5 cm x 5 cm gauze pad and allow it to dry.
   11. Wipe the abdomen using the 5 cm x 5 cm alcohol swabs. Reapply povidone-iodine and allow it to dry.
   12. Test the anesthesia by pinching the mouse’s toe and ensuring no response.
   13. Place a sheet of adhesive food service film over the surgical stage.
   14. Spray the surgical stage with 70% ethanol and allow it to dry.
   15. Wash your hands and don sterile surgical gloves.

4. Surgery procedure (7 days after initial elastin sensitization)

1. Expose the abdominal aorta.
   1. Make a skin incision using scissors or a scalpel along the abdominal midline for ~3 cm, starting 0.5−1 cm cephalad to the penis and extending to 0.5 cm caudad to the xiphoid.
   2. Grasp the skin using toothed forceps and gently pull up to identify the linea alba, which will appear as an avascular plane between the rectus muscles.
   3. Using scissors, incise the linea alba while continuing to pull up on the skin. Take care to only cut the linea alba, avoiding intra-abdominal structures.
   4. Place magnetic self-retaining retractors, with one retracting the right side of the abdominal wall, one retracting the left side of the abdominal wall, and one retracting the structures in the caudal aspect of the incision, such as the bladder and seminal vesicles.
   5. Using moistened cotton-tipped applicators, eviscerate the bowel away from yourself and cover it with a moist 5 cm x 5 cm gauze. Alternatively, pack the bowel using a rolled-up moistened 5 cm x 5 cm gauze into the left side of the mouse’s abdomen.
      NOTE: Depending on the mouse's anatomy and the placement of the retractors, the bowel may be eviscerated or packed in the opposite direction for optimal visualization.
   6. Identify the IVC and abdominal aorta in the midline posteriorly.
   7. Using cotton-tipped applicators, enter the retroperitoneum by gently opening the tissue plane covering the posterior vascular structures.
      NOTE: Posteriorly directed pressure on the tissue of interest, followed by pulling the tips of the cotton applicators apart from each other, applies greater degrees of traction. For gentler traction, rotate the tips of the cotton applicators in opposite directions.
   8. Place the lateral retractors deeper to obtain better traction and visualization of the aorta.
   9. Continue using cotton-tipped applicators to gently separate the IVC and infrarenal abdominal aorta, avoiding release of the lateral attachments of the aorta at this stage.
   10. Continue developing the plane between the IVC and aorta until a triangular space becomes visible between the aorta and IVC (Figure 1).
       NOTE: This step is typically easiest to accomplish just caudad to the left renal vein.
   11. Using blunt dissection or sharp forceps, release the lateral aortic attachments by pulling the aorta apart from the connective tissue and lumbar musculature on the mouse’s left.
   12. Place the angled sharp forceps in the space developed between the aorta and the IVC, and gently open and close the forceps in a repeated fashion to open the posterior aortic attachments.
   13. Using the same opening and closing technique, develop the posterior plane along the aorta, working from cephalad to caudal.
   14. Obtain a video micrometry image of the entire aorta and save it for analysis.
2. Apply perivascular elastase.
   1. Using a pipette, drop 10 μL of elastase solution along the length of the exposed aorta.
      NOTE: By only dissecting the tissues medial and lateral to the aorta, a gutter will emerge in which the elastase pools.
   2. Place moistened 5 cm x 5 cm gauze pads over the open abdomen and allow the elastase solution to incubate for 10 minutes.
   3. Remove the gauze pad covering and irrigate the abdomen with sterile 1X PBS three times to remove residual elastase solution.
3. Close the abdomen.
   1. Return abdominal organs to their original positions. Observe for and remove any twisting or kinking of bowel.
   2. Using toothed forceps, peel the rectus muscle and fascia away from the skin for a 0.25 cm skin flap.
   3. Close the abdominal fascia using a 4-0 braided polyglycolic acid (or similar) suture in a simple running fashion, taking care to incorporate the corners of the incision and traveling no more than 2 mm between bites.
   4. Staple the overlying skin together using 3−4 staples.

5. Post-operative care

1. Remove the mouse from the surgery stage and place it in a recovery basket that is lined with towels and placed on a stage heated to 37 °C.
2. Monitor the mouse until it emerges from anesthesia.
3. Monitor the mouse daily postoperatively for 4 days, noting general health and the status of the abdominal incision.
4. Postoperatively, perform a repeat ultrasound as described above to monitor the growth of the aneurysm.
5. Remove the staples one week after surgery or at the time of the week 1 ultrasound.

6. Repeat elastin sensitization (7 days after aneurysm induction surgery)

1. Perform the same steps as described above for the initial elastin sensitization by preparing the EDP solution in the same fashion, with a single substitution of incomplete Freund’s adjuvant instead of complete Freund’s adjuvant.

7. Tissue harvest procedures

1. Harvest tissues on postoperative day 21 for various analyses.
2. Prepare the mouse in the same fashion as was done for the induction surgery.
3. Expose the aorta again in the same fashion as was done during the induction surgery, taking care not to damage any specimens to be collected for histological analyses.
   NOTE: At this point, the mouse will have developed adhesive scarring, so more use of sharp dissection with forceps rather than blunt dissection may be necessary.
4. Obtain video micrometry images.
5. Perform a cardiac puncture to collect blood for analysis and euthanize the mouse.
6. Perfuse the mouse for analysis while wearing appropriate personal protective equipment, including gloves, eye protection, and facemasks.
   1. If the tissues are designated for flow cytometry (or similar) analysis, perfuse the left ventricle with 1X PBS without a vascular dilator at 100−120 mm Hg for a volume of 8 mL.
   2. If the tissues are designated for PCR analyses, perfuse the left ventricle with 1X PBS with vascular dilator (0.1 mM adenosine and 0.01 mM sodium nitroprusside) followed by RNA stabilization solution.
   3. If the tissues are designated for histological analyses, perfuse the left ventricle with 1X PBS with a vascular dilator, followed by 10% buffered formalin phosphate.
7. Harvest the desired tissues, taking care not to structurally damage any tissues intended for histological analysis. Mark orientation with sutures or marking agents.

Results

A total of 28 male C57BL/6J mice have undergone this surgical protocol, with one perioperative mortality from an IVC injury (3.6%). There were three additional delayed mortalities, one of which was due to an incisional hernia and the other two of which were for unknown reasons, for a total 21-day survival of 85.7%. The average time from induction of anesthesia to the end of anesthetic administration was 46.4 ± 8.0 minutes. Sham-surgery mice underwent a nearly identical protocol to the aneurysm group, including initial EDP sensitization, laparotomy, exposure of the aorta, and repeat EDP sensitization. The key difference was that the sham group did not receive topical perivascular elastase at the time of induction surgery.

Aortic diameters were measured using ultrasound and video micrometry. Representative ultrasound and intraoperative images are shown in Figure 2. An aortic aneurysm was defined as an aortic diameter ratio at day 21 of ≥1.5 compared to baseline. The ultrasound mean aortic diameter ratio versus baseline at day 21 in the aneurysm group was 1.43 ± 0.17, versus 1.04 ± 0.11 in sham surgery controls (Figure 3, p=0.00046). Based on ultrasound measurements at day 21, 8 of 11 aneurysm mice (72.7%) demonstrated >30% aortic diameter increase, versus 0 of 5 sham surgery controls (p=0.026). However, 3 of 11 (27.3%) aneurysm mice reached criteria for aneurysm, with >50% diameter increase (p=0.51).

Video micrometry measurements noted a mean aortic diameter ratio at day 21 of 1.96 ± 0.82, versus 1.06 ± 0.07 in sham surgery controls (Figure 3, p=0.0032). Video micrometry at day 21 demonstrated that 9 of 11 aneurysm mice (81.8%) had >30% increase in aortic diameter, compared with 0 of 5 sham surgery controls (p=0.0048). A >50% diameter increase was seen in 7 of 11 (63.6%) aneurysm mice (p=0.034). Differences between ultrasound and video micrometry measurements likely reflect differences between anterior-posterior ultrasound measurements and medial-lateral video micrometry measurements, as well as differences in surrounding tissue inflammation and scarring.

When comparing ultrasound data from sham-surgery controls to “normal” C57BL/6 mice that had not undergone EDP sensitization or surgery, neither group showed any aortic diameter increases >25% from baseline. The mean aortic diameter ratio at day 21 for “normal” mice was 1.08 ± 0.13 versus 1.04 ± 0.11 (p=0.69).

Representative histological images after tissue harvest on day 21 are shown in Figure 4. These histological images demonstrate the major changes observed in this model. Importantly, cellular infiltration is seen, with a thickened intima (Figure 4A). Substantial collagen deposition is seen surrounding an aneurysmal aorta (Figure 4B). Furthermore, the extent of elastin degradation in the aortic wall is shown, with thinned, fragmented, or absent lamellar elastin in the aneurysm model, compared to thick and well-defined lamellar elastin layers in a non-aneurysmal section (Figure 4C).

Representative gross aortic specimens after tissue harvest on day 21 are shown in Figure 5. These gross images show the location, extent, and degree of aortic dilation in the infrarenal segment. The aneurysmal aorta (Figure 5B) is grossly larger than the normal aorta (Figure 5A) in the entire infrarenal segment, with an area of greater dilation seen near the aortic bifurcation. The gross integrity of the aortic wall is preserved in both specimens.

Figure 1: Representative image of typical induction surgery dissection with the mouse head to the left. The three structures forming the boundaries of the “triangular space” are highlighted. This space is the location through which the dissection of the posterior aspect of the aorta is typically begun. LRV = left renal vein. IVC = inferior vena cava. Please click here to view a larger version of this figure.

Figure 2: Representative intraoperative and ultrasound images of a single mouse. (A and C) baseline images. (B and D) Day 21 images. (A and B) Intraoperative images are taken at the time of induction or harvest surgery, with the mouse head to the left. (C and D) Ultrasound images showing the aortic diameter measured in the anterior-posterior dimension. Please click here to view a larger version of this figure.

Figure 3: Aortic size ratios at day 21 versus baseline. (A) Ultrasound-based. (B) Video micrometry-based measurements. Aortic size ratios are calculated by dividing the aortic diameter at day 21 by the baseline aortic diameter. Values are displayed as mean, with error bars showing standard error of the mean (SEM). Mann-Whitney U tests were used for statistical analyses. The mean ultrasound ratio in the sham group was 1.04 (SEM = 0.05). The mean ultrasound ratio in the aneurysm group was 1.43 (SEM = 0.05). The mean video micrometry ratio in the sham group was 1.06 (SEM = 0.03). The mean video micrometry ratio in the aneurysm group was 1.96 (SEM = 0.25). There were 5 mice in the sham surgery group and 11 mice in the aneurysm group. Please click here to view a larger version of this figure.

Figure 4: Representative histological aortic tissue sections. Aortic tissue sections collected on tissue harvest after 21 days post-induction surgery are shown in the same animal. (A) Standard 5.6X magnification hematoxylin and eosin staining, demonstrating intimal thickening and cellular infiltration throughout the aortic wall. (B) 5.6X magnification picrosirius red staining, highlighting collagen fibrosis surrounding the aorta in red. (C) 5.6X magnification of Verhoeff-Van Gieson staining, highlighting elastin in black. Notably, fragmented elastin is shown throughout the aortic wall. The scale bars for panels A-C represent a distance of 400 μm. The top panel of the inset in Panel C shows 20X magnification staining of normal aortic wall lamellar elastin, with the bottom panel showing a 20X magnified area of the tissue section shown in panel C, highlighting the loss and fragmentation of elastin in the aortic wall. The scale bars in the inset panels represent a distance of 199.7 μm. Please click here to view a larger version of this figure.

Figure 5: Representative gross aortic specimens. Infrarenal abdominal aortic specimens collected on day 21 tissue harvest are shown at 1.25X magnification. (A) A gross specimen from the sham surgery group. (B) A gross specimen from the aneurysm group. In the images, the left-most extent of the specimens is at the aorta, immediately caudal to the renal arteries. The right-most extent of the specimens is at the aortic bifurcation. Scale bar = 0.812 mm. Please click here to view a larger version of this figure.

Discussion

Since it was first described in rats in 1990, the elastase-based AAA model has been modified several times and used in various animals^{10,19,20,21,22,23}. Some of the modifications include aortic cannulation, which is likely to increase mortality and introduce a traumatic component to the aortic pathology¹⁷. Less-invasive modifications include 3-aminopropionitrile fumarate salt (β-Aminopropionitrile (BAPN)), a lysyl oxidase inhibitor, dissolved in animal drinking water¹⁸. The rationale for most modifications to the model is to improve consistency in aneurysm formation, ease of technical procedures, and animal survival. Variations in surgical technique between models and between operators using the same model may lead to variable results and difficulty in interpreting the study. In particular, the method of aortic exposure, the elastin preparation used, and the incubation time of perivascular elastase are likely to affect aneurysm formation and animal survival. However, this iteration of the model is notable for its operational speed, which enables a larger number of animals and thereby improves statistical power, and for its high survival, which enables more efficient use of time and resources. Furthermore, this model focuses on elastin degradation in the aortic wall and does not address other confounding mechanisms that can increase nonspecific inflammation or tissue damage, thereby inducing aneurysms.

The most crucial step in this protocol is separating the abdominal aorta from its connections. Separating the aorta from its connections to the adjacent IVC is the highest-risk portion of the aneurysm induction surgery, as the IVC wall is fragile and IVC injuries are difficult to manage, frequently leading to hemorrhagic mortality. Small arterial or venous injuries may be treated with direct pressure through a cotton-tipped applicator. To optimize dissection of the connective tissues surrounding the aorta while avoiding IVC injuries, it is important to rely on blunt dissection for as much of the exposure as possible. Observationally, sharp dissection involves applying point forces with inconsistent intensities that are difficult to measure, which can lead to inadvertent tearing of the IVC wall. If the cotton tip of the applicator becomes too moist, it can be difficult to obtain the requisite traction required to complete this dissection. In this case, either use a fresh cotton-tipped applicator or use a sterile 5 cm x 5 cm gauze to squeeze the excess fluid out of the cotton tip. To maximize the success of blunt dissection, the lateral aortic attachments must be left in place while the medial attachments, bordering the IVC, are dissected. If the lateral aortic attachments are released first, the traction to pull the aorta apart from the IVC is lost. Once both the medial and lateral aortic attachments are released, the posterior attachments are easily accessed and dissected. Exposure in this fashion also helps form a gutter for the topical elastase to fill, obviating the need to wrap the aorta with elastase-soaked fabric and reducing the risk of aortic injury and heterogeneity in elastase application. Importantly, the operator must understand when the risk of further dissection endangers the animal's survival, as there is a marginal additional benefit from aneurysm formation with extensive dissection. It is not necessary for AAA formation to have an abdominal aorta completely free of all attachments.

Other key procedural considerations in this protocol include the judicious monitoring of the volatile anesthetic. Despite repeated isoflurane exposures, the animals do not appear to develop substantial tolerance to the anesthetic. A sufficient concentration of isoflurane for 25−35g C57BL/6J mice at 14 weeks old is 1.8−2%. Lower concentrations risk inadequate anesthesia, and higher concentrations risk avoidable anesthetic-related mortality. The use of a volatile anesthetic is advantageous for rapid induction and emergence from anesthesia while allowing variable operating times, as the anesthetic agent is administered continuously. Entry into the abdominal cavity must also be approached with caution to avoid injury to intra-abdominal organs. The placement of retractors must also be carefully considered to avoid retraction injuries, particularly to the bowel. If retractors must be placed on bowel or solid organs, padding them with moistened cotton gauze reduces the risk of clinically significant injury. After completion of the elastase incubation and PBS irrigation of the abdomen, abdominal closure is important to avoid delayed mortality from adhesive bowel obstruction, volvulus, or incisional hernia. Taking care to return the bowel to its original configuration, without twisting or kinking, can minimize bowel complications. For the closure of the abdominal wall specifically, the distance between suture bites must be minimal to avoid abdominal contents herniating between suture gaps. Postoperatively, it is important to monitor the animals for appropriate recovery. The mice should appear healthy, alert, and active within minutes of emergence from volatile anesthesia.

Serial ultrasound was performed at multiple time points for several reasons. We perform a baseline ultrasound, prior to any EDP sensitization or induction surgery, to rule out any pre-existing anatomic anomalies or disease in the animal and to establish an aortic diameter baseline. Subsequent measurements allow for the monitoring of aortic dilation over time, which helps us understand the typical trends and timing of aortic size changes. Ultrasound also provides a second measurement to compare against video micrometry results, as both methods of measurement have limitations. The primary limitations of ultrasound measurements are the operator dependence and difficulty of measuring the same aortic locations at different timepoints. The standardization of ultrasound protocol, which begins with identifying key anatomic landmarks, helps to reduce between-measurement variability. Video micrometry is limited by the quality of the repeat exposure of the aorta, as well as the presence of inflammation and scar tissue that can make identification of the exact borders of the aortic wall challenging. We have previously used polymer-based vascular contrast agents with subsequent micro-computed tomography to determine aortic morphology. However, this technique is limited as it requires sacrifice of the animal at the time of imaging, with the inability to perform other assays on the same specimen. One advantage of ultrasound and video micrometry is that they do not preclude the use of any specimens for further assays. To reduce potential bias, all ultrasound and video micrometry measurements are performed without knowledge of the animal’s experimental group. However, blinding is imperfect, as the degree of inflammatory changes and scarring are observationally greater in the aneurysm group at the time of day 21 tissue harvest and video micrometry, which may lead to unconscious bias.

In prior iterations of this protocol, our group used only a single EDP sensitization step prior to induction surgery and did not consistently induce AAA. However, with the current protocol that includes a second EDP injection, if AAA are not consistently forming, the main factor to evaluate is the surgical technique for adequate exposure of the abdominal aorta. When evaluating mortality, early mortalities are typically related to blood loss and subsequent hypovolemic shock. In cases where the animal appears to have lost substantial blood volume, the operator may consider leaving a small amount of sterile PBS irrigation in the abdomen prior to closure to aid hydration. Alternatively, subcutaneous PBS can be administered. However, these methods are infrequently required. Late mortalities are often associated with bowel or infectious complications, including small bowel obstructions, hernias, and abscesses. Taking care to return the bowel to its original configuration, avoiding organ injury during aortic exposure, and using a meticulous closure technique can substantially minimize the risk of these complications.

There are several limitations to this method. One major limitation is the operator learning curve for the induction surgery. Additionally, this protocol has only been used in male mice. As such, the consistency across female mice or other animal models is unknown. Interestingly, when obtaining blood samples via terminal left ventricular cardiac cannulation, blood volumes are higher with a closed ventricular puncture method. However, if completion dissection images are desired, an open heart stick is necessary to avoid post-mortem tissue damage during dissection and tissue collection. If completion dissection images are not desired, the repeat exposure of the abdominal aorta can be skipped. Furthermore, the exact timing of maximal aortic dilation is somewhat variable. Prior groups have shown success with an endpoint at 14 days¹⁹. While many of the mice in this protocol demonstrate aortic dilation and aneurysms at 14 days, the degree of consistent aneurysmal dilation is better at 21 days. Additionally, because tissue harvest timing is based on aneurysm formation, capturing dynamic cytokine expression profiles is difficult, with prior studies showing that most cytokines return to baseline levels by day 7 after induction surgery¹⁸.

A major limitation that may reduce reproducibility is the likely heterogeneity of the EDP solution made in-house. While the protocol is standardized, the composition of the EDP mixture is not routinely verified using analytic techniques. As such, the EDP preparation may require further optimization of pH balance, enzyme or elastin amounts, or digestion conditions. Given the mixture is not verified using analytic techniques, the EDP preparation may also vary somewhat from cohort to cohort. It is likely that some residual elastase activity remains in the injected mixture for a time, as the reaction is simply quenched with cold PBS solution, but the elastase is not inactivated by thermal or chemical methods. The residual elastase is also not bound or precipitated, so it remains in solution. As such, it is possible that, upon injection into the animal and subsequent return to physiologic temperature, the enzyme resumes activity for a period before host clearance. The improvement of the EDP mixture is an ongoing area of study, with experiments planned to test concentrations, buffers, and incubation conditions, with subsequent verification and purification using mass spectrometry and high-performance liquid chromatography. While commercial EDP preparations were previously available, purchasing them can incur substantial costs, and the additional benefit of using them is unclear. In prior iterations of this protocol, the animal's sensitivity to EDP has been quantified using ear volume assays akin to a tuberculin skin test²⁴.

Overall, this protocol is effective for modeling aortic dilation in a widely available mouse strain. Given increasing interest in the role of immunologic and inflammatory mechanisms in AAA formation, this model provides a robust and reproducible method for translational study. In particular, this model was used to examine the effects of novel therapeutics in mitigating aortic diameter growth over time. Future experiments are also planned to elucidate the immunologic factors at play in this model, including immunohistochemistry and surface marker expression studies. This iteration of the murine elastase-induced AAA model is safe, with high perioperative and overall survival rates, and fast, with relatively short operative and follow-up times. Despite the limitations, we believe this protocol can consistently produce aortic dilation in mice with an immune-sensitization component, thereby supporting a better understanding of AAA and its treatments.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1 mL Tuberculin Syringes	Becton Dickinson	309628
10% Buffered Formalin Phosphate	Fisher Chemical	SF100-4
70% Isopropyl Alcohol Swabs	Cardinal Health	S-16183
Angled sharp forceps	Dumont	I I25 I-35
AutoClip 9mm (Skin stapler)	MikRon	427630
Betadine	Atlantis Consumer Healthcare	67618-155-16
Cotton gauze pad	Dynarex Corporation	3223
Cotton tipped applicator	Puritan Medical Products	25-806 2WC
Elastase from porcine pancreas (≥4.0 units/mg protein)	Sigma-Aldrich	E1250	Stored in aqueous solution at 4 °C
Elastase from mouse pancreas (4.75 units/mg protein)	Elastin Products Co., Inc.	MS838	Stored in aqueous solution (0.1 mg in 1 mL PBS)at 4 °C
Elastin	MP Biomedicals	101636	Stored in aqueous solution at 4 °C
Enersight Desktop Software	Leica	https://www.leica-microsystems.com/products/microscope-software/p/enersight/
Ethiqa XR (extended release buprenorphine)	Fidelis	NDC 86084-100-30
Eye lubricant	OptixCare	Opx-4242
Flexacam i5 (Stereo)	Leica	https://www.leica-microsystems.com/products/microscope-cameras/p/12730536-leica/
Freund’s complete adjuvant	MP Biomedicals	642851	Stored in aqueous solution at 4 °C
Freund’s incomplete adjuvant	MP Biomedicals	642861	Stored in aqueous solution at 4 °C
Gloves (Sterile)	Encore	7824PF
Hypodermic Needles (23 G)	Becton Dickinson	305145
Hypodermic Needles (27 G)	Becton Dickinson	305109
Isospire (Isoflurane)	Dechra Veterinary	07-894-8943
Labeling Tape	Fisher Scientific	15959
Masks	Dukal Corp.	1540
Needle driver	Scanlan	6006-36
Omnicon F/air Anesthetic Gas Canister	A.M. Bickford Inc.	80120
Press'N Seal	Glad	12587704417
RC2 Rodent Circuit Controller	VetEquip Incorporated	922100
RNAlater Stabilization solution	ThermoFisher Scientific	AM7021
Scissors	Fine Science Tools	14005-12
Skin staples (clips)	MikRon	205016
Small Rodent Warmer Heater	Stoelting	53851
Small Rodent Warmer Stage	Stoelting	53812
Stereo Microscope M60	Leica	https://www.leica-microsystems.com/products/light-microscopes/stereo-microscopes/p/leica-m80/
Straight sharp forceps	Dumont	I I 25 I-30
SurgiSuite self-retaining magnetic retractor set and stage	Kent Scientific	13-005-180
Tahoe Portable Oxygen Concentrator	Vetland Medical	595-2300A
Toothed forceps	Fine Science Tools	11043-08, 11042-08
Vevo 2100 Ultrasound	FUJIFILM Visualsonics	EDU00496
Visorb polyglycolic acid suture	CP Medical	397A
VWR Absorbents	VWR	95057-860