Subcutaneous Angiotensin II Infusion using Osmotic Pumps Induces Aortic Aneurysms in Mice

Ethics Statement: Mouse studies are performed with approval of the University of Kentucky Institutional Animal Care and Use Committee (IACUC protocol number: 2006-0009). Mice are euthanized at termination using an overdose cocktail of ketamine (~ 210 mg/kg) and xylazine (~ 30 mg/kg).

1. Calculation of AngII Amount

NOTE: This protocol uses the example of infusion of AngII (1,000 ng/kg/min) for 4 weeks in 4 male LDL receptor -/- mice fed a saturated fat-enriched diet.

1. Weigh study mice before calculating the amount of AngII needed for infusion.
2. Use the template (Table 1) to calculate the AngII mass needed for the experiment. Use the “Mean Pumping Rate” indicated in the Instruction of pumps as the “Pumping rate” in Step 4 of the template. In the template, record Steps 1 – 5 manually, and Steps 6 – 10 are calculated automatically.
   1. In the template, assume that mice will gain 1 g of body weight during infusion of AngII 1,000 ng/kg/min for 4 weeks.
      NOTE: Each mouse may have very different body weight gain that will depend on many variables, such as mouse strain and diet. We routinely use “0" or “1 g” based on our own experience from previous studies.
   2. Calculate a 300 µl total volume of AngII solution for each mouse since each pump requires approximately 250 µl.

2. Dissolution of AngII

1. Store lyophilized AngII vials at -20 °C. Equilibrate AngII vials to RT before opening.
2. Weigh the calculated AngII mass (7.3 mg as shown in Table 1) into a sterile plastic tube.
   NOTE: Per Merck Index, do not use glass tubes for the dissolution since an aqueous solution of AngII has a strong affinity for binding to glass.
3. Add the calculated volume of sterile saline (1,200 µl) into the plastic tube containing the lyophilized AngII, cap, and mix thoroughly by inversion until the solution is clear.
4. Label mouse numbers #1, #2, #3, and #4 on individual sterile plastic tubes with caps (0.5 – 1.5 ml). Prepare AngII solution under a laminar hood for each mouse based on body weight as calculated in Step 1.2 and Table 1.
   1. For example, pipette 3.6 µl sterile saline into tube #1, then 296.4 µl AngII solution, and mix thoroughly by pipetting up and down gently.
5. Label mouse numbers on plastic tubes with caps (4 ml; sterile). These will be used for incubating pumps as described in Step 3.13.

3. Osmotic Pump Filling

1. Obtain pumps in two separate parts: the main body of the pump and the flow moderator (Figure 1). Each box has 10 pump bodies and flow moderators that are wrapped individually. Record the lot number.
   NOTE: Always wear gloves because oils transferred from hands to the exterior casing of pumps will adversely affect pumping function. Use sterile gloves, gauze, tubes, filling needle, and weigh boats to prepare the pumps, to avoid risk of infection from the implant.
2. Open only the number of pump bodies and flow moderators needed for the study, as these cannot be stored once opened. If more than 10 pumps are needed, ensure that the lot numbers of the pumps are the same for one study, since pumps from different lot numbers have different Mean Fill Volume and Pump Rate.
3. Weigh each pump (including both the main body and flow moderator), and note the weight to 4 decimal places (e.g., 1.1443 g of mouse #1). This weight, termed “Pump Weight empty” in the template (Table 1), will be used to calculate the filled ratio.
4. Attach the pump filling needle to a 1 cc sterile syringe and carefully fill the syringe with AngII solution from the appropriately numbered plastic tube. It is important to avoid drawing air into the syringe.
5. Remove all air bubbles carefully from the syringe while the needle is positioned downwards. Keep the needle/syringe in this position to prevent the introduction of bubbles into the pump.
6. Gently insert the filling needle into the pump body. Advance the tip of the needle into the pump. Do not rest the tip of the needle tightly on the bottom of the pump.
7. Push the syringe plunger slowly to fill the pump with AngII solution. A dark shadow inside the pump indicates the filling level. The filling volume is approximately 246 µl, per Instructions.
8. Stop filling the pump and carefully remove the needle as soon as a bead of fluid rises out of the pump.
9. Insert flow moderator into pump through the hole on top of pump body until no gap is seen between the head of the flow moderator and the top of the pump body (Figure 1).
10. Insertion of moderator into the pump body leads to some fluid leaking from the opening of the flow moderator. Carefully blot all extra fluid that might have leaked during placement of moderators.
11. Weigh filled pump. Record the weight under “Pump Weight Filled” in the template.
12. Calculate Filling Ratio (%) = (Pump Weight “filled” – “empty”) x 1,000/mean fill volume x 100.
    1. Calculate Filling Ratio as indicated in Table 1. Ideally, filling ratio should be equal or greater than 100%. Refill pump if the filling ratio is < 95% (implicating that air bubbles may be present in pump).
13. Place filled pump into the labeled 4 ml tube (Step 2.5) with the moderator head facing upward. Add sufficient volume of sterile saline to cover the pump. Keep pump in tube of saline until implantation.
14. Place tubes in a 37 ºC incubator. Incubate pumps O/N (at least 12 hr) to allow partial priming, and then implant into mice. Pumping of AngII starts approximately 24 hr after implantation, which allows mice to recover from surgery prior to any potential stress arising during AngII infusion.

4. Preparation for Pump Implantation

1. Autoclave (gravity mode, dry cycle, 15 min) gauze, cotton swabs and surgical tools including scissors, hemostat, forceps, staples, and stapler at least 1 day prior to surgery.
2. In a procedure room, prepare vaporizer for anesthesia using isoflurane. Open up sterile drapes in a laminar flow hood, and place nose cone for isoflurane anesthesia. Place betadine, 70% ethanol, sterile water, bead sterilizer, antiseptic handrub, swabs, gauze, and filled pumps in a laminar hood.
3. Don a mask and gown, then open an outer drape in a laminar hood with clean hands. Put on sterile gloves and open the sterile inside pack.

5. Surgical Procedure of Pump Implantation

1. Place mouse in an induction chamber with inflow of isoflurane at a flow rate of 1.5 – 2%. Monitor the mouse for an additional 2 – 3 min after recumbency. Shave an area about the size of a quarter, over the left or right shoulder.
2. Place mouse in a laminar hood, with its nose flush with a cone connected to isoflurane outflow (Figure 2A). Place the mouse’s head toward the surgeon’s dominant hand. Use vet ointment on mouse’s eyes to prevent dryness while under anesthesia. Ensure that the mouse has no response to stimulation of pain prior to the surgery. For example, pedal response is a good indicator for pain.
3. Swab and wipe shaved area with betadine followed by three wipes with 70% ethanol.  Don or change sterile gloves.
4. Use a surgical scalpel to make an ~ 1 cm incision behind the ear over the shoulder blade of the front leg. This incision should be perpendicular to the tail. Use care to cut only the skin and not underlying tissues.
5. Hold forceps in one hand to open incision, and use the other hand to make a subcutaneous tunnel under the skin using a hemostat (Figure 2B).
6. Advance hemostat tip toward the tail, and create a pocket for pump. This is accomplished by carefully opening the jaws of the hemostat under the skin to open up a pouch. Withdraw the hemostat from incision.
7. Insert pump into the incision with the moderator head positioned to the rear of the mouse (Figure 2C). Gently push the pump completely into the pocket. There should be enough free space to close the wound with no tension or stretching of the skin.
8. Once a pump has been inserted, firmly pinch both sides of the incision, straightening so the edges meet, and place 1 or 2 wound clips to close (Figure 2D). Inspect the incision site to ensure that there is complete closure of the wound, and that the pump is not pressing directly on the site.
9. Apply topical lidocaine cream (4% wt/wt) with a clean cotton swab.Remove mouse from the nose cone, and place it on a heating pad until it regains consciousness. After recovering, the mouse is returned to its cage.
10. Place surgical instruments into a bead sterilizer for 10 seconds between mice. Allow instruments to cool before use. Clean gloves with antiseptic handrub between mice. Monitor all mice until full recovery is achieved.
11. Monitor mice closely after the surgery. Inject a bolus of sterile saline (0.2 – 0.3 ml) subcutaneously if a mouse shows signs of distress, dehydration or apparent weight loss. Observe mice at least twice a day during the first 10 days, and at least once every day subsequently. Perform a necropsy immediately if any mice die during AngII infusion. Remove wound clips between 7 – 14 days after the surgery.

6. Harvesting, Fixing, Cleaning, and Imaging of Aortas

1. Cut open the mouse thoracic and abdominal cavities ventrally, cut open right atrium, perfuse with saline through left ventricle of the heart to remove blood in the aorta, and then harvest the aorta ²⁷.
2. Place harvested aortas in plastic tubes containing at least 3 ml of 4% paraformaldehyde or 10% neutrally buffered formalin for 24 – 48 hr ²⁷.
3. Remove adventitial tissues carefully. Pin aorta on black wax with pins. Acquire aortic images with same magnification. Include a ruler in each image for calibration, as shown in Figure 3.

7. En face Imaging of Aortas

1. Cut open aorta longitudinally through the outer and inner curvature of aortic arch, and cut open major branches including innominate, left carotid artery, and left subclavian artery. Pin aorta flat with outer adventitial laying adjacent to the black wax.
2. Acquire en face picture of the intimal surface of aorta at the same magnification. Include a ruler in each image for calibration, as shown in Figure 4.