The research work presented in this manuscript was supported by a grant (HL107319 to Alan Daugherty and HL107326 to Lisa A. Cassis) from the National Institutes of Health of the United States of America. The content in this manuscript is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. The publication of this manuscript was sponsored by DURECT Corporation.

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.
<ol>
	<li>Weigh study mice before calculating the amount of AngII needed for infusion.</li>
	<li>Use the template (Table 1) to calculate the AngII mass needed for the experiment. Use the &#8220;Mean Pumping Rate&#8221; indicated in the Instruction of pumps as the &#8220;Pumping rate&#8221; in Step 4 of the template. In the template, record Steps 1 - 5 manually, and Steps 6 - 10 are calculated automatically.
	<ol>
		<li>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 &#8220;0&#34; or &#8220;1 g&#8221; based on our own experience from previous studies.</li>
		<li>Calculate a 300 &#181;l total volume of AngII solution for each mouse since each pump requires approximately 250 &#181;l.</li>
	</ol>
	</li>
</ol>

2. Dissolution of AngII
<ol>
	<li>Store lyophilized AngII vials at -20 &#176;C. Equilibrate AngII vials to RT before opening.</li>
	<li>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.</li>
	<li>Add the calculated volume of sterile saline (1,200 &#181;l) into the plastic tube containing the lyophilized AngII, cap, and mix thoroughly by inversion until the solution is clear.</li>
	<li>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.
	<ol>
		<li>For example, pipette 3.6 &#181;l sterile saline into tube #1, then 296.4 &#181;l AngII solution, and mix thoroughly by pipetting up and down gently.</li>
	</ol>
	</li>
	<li>Label mouse numbers on plastic tubes with caps (4 ml; sterile). These will be used for incubating pumps as described in Step 3.13.</li>
</ol>
3. Osmotic Pump Filling
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#8220;Pump Weight empty&#8221; in the template (Table 1), will be used to calculate the filled ratio.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#181;l, per Instructions.</li>
	<li>Stop filling the pump and carefully remove the needle as soon as a bead of fluid rises out of the pump.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Weigh filled pump. Record the weight under &#8220;Pump Weight Filled&#8221; in the template.</li>
	<li>Calculate Filling Ratio (%) = (Pump Weight &#8220;filled&#8221; - &#8220;empty&#8221;) x 1,000/mean fill volume x 100.
	<ol>
		<li>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 &#60; 95% (implicating that air bubbles may be present in pump).</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Place tubes in a 37 &#186;C 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.</li>
</ol>
4. Preparation for Pump Implantation
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
5. Surgical Procedure of Pump Implantation
<ol>
	<li>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.</li>
	<li>Place mouse in a laminar hood, with its nose flush with a cone connected to isoflurane outflow (Figure 2A). Place the mouse&#8217;s head toward the surgeon&#8217;s dominant hand. Use vet ointment on mouse&#8217;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.</li>
	<li>Swab and wipe shaved area with betadine followed by three wipes with 70% ethanol.&#160; Don or change sterile gloves.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
6. Harvesting, Fixing, Cleaning, and Imaging of Aortas
<ol>
	<li>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.</li>
	<li>Place harvested aortas in plastic tubes containing at least 3 ml of 4% paraformaldehyde or 10% neutrally buffered formalin for 24 - 48 hr 27.</li>
	<li>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.</li>
</ol>
7. En face Imaging of Aortas
<ol>
	<li>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.</li>
	<li>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.</li>
</ol>

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Publication of this article is sponsored by Alzet.

Osmotic pumps delivering AngII subcutaneously is a routine approach to inducing aortic aneurysms in mice. Based on data from many laboratories, there have been consistent findings that this is a reliable and reproducible method to study both AAAs3,4 and TAAs22-26 in mice. Therefore, this mouse model is considered a model that recapitulates several features of human aortic aneurysms and provides mechanistic insights into these devastating diseases.
While aging is a risk factor for AAAs in humans, it has not been systematically studied for AngII-induced AAAs in mice. However, it appears incidence and severity of AngII-induced AAAs are similar in mice at the age of 8 - 48 weeks4,5,7.&#160; Currently, there are only a few studies reporting AngII-induced TAAs in mice at the age of 8 - 24 weeks22-26, which did not show apparent age-related differences on TAA formation.
Female mice have a much lower incidence of AAAs than male mice infused with AngII4,28. It is also worth noting that the incidence of AngII-induced AAAs is much higher in hyper- than normo-cholesterolemic mice, which is more than 50% versus less than 30%, respectively. Additionally, aortic rupture is frequent (approximately 10 - 30%) in both normo- and hypercholesterolemic mice during AngII infusion. Infusion of AngII at a rate of 1,000 ng/kg/min into hypercholesterolemic mice, such as LDL receptor -/- mice fed a Western diet or apolipoprotein (apoE) -/- mice fed a normal or Western diet, has maximal effects on AAA development3,4,29. This infusion rate is optimal for a study in which manipulating a gene of interest in hypercholesterolemic mice is expected to reduce AAAs. If a manipulation in hypercholesterolemic mice is expected to augment AAAs, it is recommended to infuse AngII at a rate of 500 ng/kg/min or lower30. In contrast to AAAs, there is no demonstrated association between male gender or hypercholesterolemia and AngII-induced TAAs25. However, similarly to AAAs, if manipulation of a gene of interest is expected to augment TAAs, we recommend a lower infusion rate than 1,000 ng/kg/min for AngII infusion.
It is also important to know that incidence and severity of AngII-induced aortic aneurysms vary among individual mice and between studies. If mice do not develop aortic aneurysms, one potential possibility is that AngII might have not been successfully delivered into mice. For validation of a high infusion rate of AngII, such as 1,000 ng/kg/min, measurement of blood pressure is recommended prior to, and during, AngII infusion using a non-invasive tail-cuff method31. AngII infusion at a rate of 1,000 ng/kg/min increases systolic blood pressure in mice. Also, plasma renin concentrations may be measured during AngII infusion or at termination since AngII has a negative feedback on renin secretion. Therefore, AngII infusion leads to reductions in plasma renin concentrations. If a mouse infused with AngII has no apparent aortic pathologies, no increase of blood pressure, and no decrease of plasma renin concentration, it would indicate that AngII has not been delivered efficiently through the implanted osmotic mini pump. We would recommend removing this mouse from the study. It is also important to note that some mice do not develop aortic aneurysms despite increased systolic blood pressures and decreased plasma renin concentrations. These mice should remain in the study.
In summary, AngII infusion is achieved by subcutaneous implantation using osmotic pumps to induce aortic aneurysms in mice. This method delivers AngII constantly at a defined rate for designated durations that are used to study both AAAs and TAAs.

Aortic aneurysms exhibit permanent luminal expansion of the aorta that portends rupture and usually leads to death. This disease occurs in both abdominal and thoracic aortic regions, which are termed as abdominal aortic aneurysms (AAAs) and thoracic aortic aneurysms (TAAs), respectively. Due to an incomplete understanding of molecular mechanisms and pathophysiologic processes, there is no proven medical therapy that can prevent expansion or rupture of either type of aortic aneurysms. Since it is difficult to acquire patient samples and perform experiments in humans directly, research focusing on defining mechanisms of AAAs has been frequently extrapolated from animal models. A commonly used animal model is subcutaneous infusion of angiotensin II (AngII) into mice. Compared to other surgical approaches for inducing AAAs in mice, such as intra-aortic elastase perfusion or peri-aortic application of calcium chloride that require laparotomy1,2, this method does not require entry into the body cavity and requires minimal surgical expertise3,4.
Subcutaneous infusion of AngII through osmotic pumps to induce AAAs was initially reported in low density-lipoprotein (LDL) receptor -/- mice fed a saturated fat-enriched diet3, and subsequently in apoE -/- mice fed a normal laboratory diet4. Many recent studies have also demonstrated that AngII induces AAAs in normolipidemic mice5-7. The approach of infusing AngII has been applied to induce AAAs and explore molecular mechanisms as well as development of potential therapeutic strategies (e.g.,&#160;5-15) since this model recapitulates many features observed in human AAAs. For example, risk factors of human AAAs such as smoking, aging, and male gender also augment AngII-induced AAAs in mice16,17. The association of hypercholesterolemia with AAAs in humans requires clarification. However, it has been consistent that hypercholesterolemia augments AngII-induced AAAs in mice18. Pathologies of AngII-induced AAAs in mice are highly heterogenous and are characterized by profound macrophage infiltration, collagen degradation, thrombotic formation and resolution, and neovascularization19-21. In contrast to the most common infrarenal aortic location of AAAs in humans, AngII-induced AAAs in mice occur in the suprarenal aortic region. Another ubiquitous feature of AngII-induced AAAs is the transmural medial break, leading to transmural thrombosis. It is unclear whether transmural elastin rupture occurs in humans since pathological development of AAAs in humans has not been exclusively studied due to lack of aneurysmal tissues from earlier stages.
AngII infusion into mice also leads to profound expansion of the thoracic aortic region, that is predominantly restricted to the ascending aorta which is the most common region for TAAs in humans19,22-26. Similar to AngII-induced AAAs, TAAs induced during AngII infusion also recapitulate many features of human TAAs25. However, in contrast to AngII-induced AAAs, AngII-induced TAAs are not associated with hypercholesterolemia and do not have gender differences.
The overall goal of subcutaneous AngII infusion into mice is to study pathological features and molecular mechanisms of AAAs and TAAs.

<table><tbody><tr><td>Angiotensin II</td><td>Bachem</td><td>H-1705</td><td>compound used to induce aortic aneurysms</td></tr><tr><td>Alzet Osmotic Pumps</td><td>DURECT Corporation</td><td>Alzet Model 2004</td><td>feasible for 28-day infusion in mice weighed &gt; 20 g</td></tr><tr><td>Saturated fat-enriched diet</td><td>Harlan Teklad</td><td>TD.88137</td><td>42% calories/calories to stimulate hypercholesterolemia in LDL receptor -/- mice</td></tr></tbody></table>

subcutaneous angiotensin ii infusion using osmotic pumps induces aortic aneurysms in mice

Osmotic pumps continuously deliver compounds at a constant rate into small animals. This article introduces a standard protocol used to induce aortic aneurysms via subcutaneous infusion of angiotensin II (AngII) from implanted osmotic pumps. This protocol includes calculation of AngII amount and dissolution, osmotic pump filling, implantation of osmotic pumps subcutaneously, observation after pump implantation, and harvest of aortas to visualize aortic aneurysms in mice. Subcutaneous infusion of AngII through osmotic pumps following this protocol is a reliable and reproducible technique to induce both abdominal and thoracic aortic aneurysms in mice. Infusion durations range from a few days to several months based on the purpose of the study. AngII 1,000 ng/kg/min is sufficient to provide maximal effects on abdominal aortic aneurysmal formation in male hypercholesterolemic mouse models such as apolipoprotein E deficient or low-density lipoprotein receptor deficient mice. Incidence of abdominal aortic aneurysms induced by AngII infusion via osmotic pumps is 5 - 10 times lower in female hypercholesterolemic mice and also lower in both genders of normocholesterolemic mice. In contrast, AngII-induced thoracic aortic aneurysms in mice are not hypercholesterolemia or gender-dependent. Importantly, multiple features of this mouse model recapitulate those of human aortic aneurysms.

The 4 male LDL receptor -/- mice described in the Protocol section were euthanized after 4 weeks of AngII infusion. Aortas were harvested, cleaned, and imaged to visualize aortic dilations. As shown in Figure 3, aortas have several different characteristics including expansion of the suprarenal region (AAAs; Figure 3A), expansion of the ascending region (TAAs; Figure 3B), or expansion of both regions (presence of both AAAs and TAAs; Figure 3C), whereas morphology in one mouse was grossly normal (Figure 3D). Dilation of the abdominal aorta is quantified by measuring the ex vivo maximal width of the suprarenal region, as illustrated by the red line in Figure 3A. To measure ascending aortic dilation, aortas were cut open and pinned as shown in Figure 4. Intimal surface area was measured in the ascending aortic region (area surrounded by the red lines in Figure 4A) to quantitate TAAs. A ruler was included in each image to standardize measurements, as shown in both Figures 3 and 4.
<img alt="Figure 1" src="/files/ftp_upload/53191/53191fig1.jpg" /> 
Figure 1. Representative image of filled osmotic pump. Each pump contains two separate parts: a main body and a flow moderator. After filling the pump body with AngII, the flow moderator is inserted to seal the pump.
<img alt="Figure 2" src="/files/ftp_upload/53191/53191fig2.jpg" /> 
Figure 2. Process of pump implantation surgery. (A) Mouse is placed in a laminar hood with a nose cone that is continuously releasing isoflurane and oxygen; (B) A straight hemostat is inserted into the skin incision to make a subcutaneous tunnel; (C) Pump is inserted through the skin incision gently; (D) The skin incision is stapled after pump insertion.
<img alt="Figure 3" src="/files/ftp_upload/53191/53191fig3.jpg" /> 
Figure 3. Aortic images (ex vivo) from mice infused with AngII. AngII 1,000 ng/kg/min was infused in male LDL receptor -/- mice for 28 days. (A) AAAs accompanied by thrombosis; Red line (2.05 mm) shows the measurement of maximal aortic width in the suprarenal region. (B) Ascending aortic dilation (TAA) with grossly normal abdominal aorta; (C) Profound dilations in both the ascending and suprarenal aortic regions (TAAs and AAAs); (D) grossly normal aorta with no apparent dilation of either ascending or suprarenal aortic region.
<img alt="Figure 4" src="/files/ftp_upload/53191/53191fig4.jpg" /> 
Figure 4. En face images of thoracic aortic regions from mice infused with AngII. AngII 1,000 ng/kg/min was infused in male LDL receptor -/- mice for 28 days. Surface area outlined by a red line represents the ascending aortic region including part of the aortic arch.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>1</td>
			<td colspan="2">Dose required</td>
			<td></td>
			<td>1,000</td>
			<td colspan="2">ng/kg/min</td>
			<td></td>
		</tr>
		<tr>
			<td>2</td>
			<td colspan="3">Start body weight (largest mouse)</td>
			<td>24.8</td>
			<td>g</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3</td>
			<td colspan="3">Total estimated body weight gain</td>
			<td>1</td>
			<td>g</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4</td>
			<td>Pumping rate</td>
			<td></td>
			<td></td>
			<td>0.25</td>
			<td>&#181;l/hr</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5</td>
			<td colspan="2">Number of mice</td>
			<td></td>
			<td>4</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>6</td>
			<td colspan="2">Dose per hour for animal</td>
			<td></td>
			<td>1518</td>
			<td>ng</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>7</td>
			<td>Conc needed</td>
			<td></td>
			<td></td>
			<td>6072</td>
			<td>ng/&#956;l</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>8</td>
			<td colspan="2">For 300 ul solution</td>
			<td></td>
			<td>1.82</td>
			<td colspan="2">mg/300 &#956;l</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td colspan="2">SOLUTION NEEDED</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>9</td>
			<td colspan="2">Total AngII (mg)</td>
			<td></td>
			<td>7.3</td>
			<td>mg&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10</td>
			<td colspan="2">Dissolved in saline&#160;</td>
			<td></td>
			<td>1,200</td>
			<td>&#956;l&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td colspan="2"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse</td>
			<td>Body Weight</td>
			<td>Dilution factor</td>
			<td colspan="2">Volume (&#956;l)</td>
			<td colspan="2">Pump Weight (g)</td>
			<td>Filled Ratio</td>
		</tr>
		<tr>
			<td>#</td>
			<td>(g)</td>
			<td></td>
			<td>AngII</td>
			<td>Saline</td>
			<td>Empty</td>
			<td>Filled</td>
			<td>(%)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>24.5</td>
			<td>1.0</td>
			<td>296.4</td>
			<td>3.6</td>
			<td>1.1443</td>
			<td>1.3877</td>
			<td>99</td>
		</tr>
		<tr>
			<td>2</td>
			<td>23.0</td>
			<td>0.9</td>
			<td>278.2</td>
			<td>21.8</td>
			<td>1.1677</td>
			<td>1.4145</td>
			<td>100</td>
		</tr>
		<tr>
			<td>3</td>
			<td>24.8</td>
			<td>1.0</td>
			<td>300.0</td>
			<td>0.0</td>
			<td>1.1438</td>
			<td>1.3904</td>
			<td>100</td>
		</tr>
		<tr>
			<td>4</td>
			<td>21.8</td>
			<td>0.9</td>
			<td>263.7</td>
			<td>36.3</td>
			<td>1.1438</td>
			<td>1.3904</td>
			<td>100</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="7">Dilution factor = body weight of the mouse/body weight of the largest mouse</td>
			<td></td>
		</tr>
	</tbody>
</table>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Mouse</td>
			<td>Body Weight</td>
			<td>Dilution factor</td>
			<td colspan="2">Volume (&#956;l)</td>
			<td colspan="2">Pump Weight (g)</td>
			<td>Filled Ratio</td>
		</tr>
		<tr>
			<td>#</td>
			<td>(g)</td>
			<td></td>
			<td>AngII</td>
			<td>Saline</td>
			<td>Empty</td>
			<td>Filled</td>
			<td>(%)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>24.5</td>
			<td>1.0</td>
			<td>296.4</td>
			<td>3.6</td>
			<td>1.1443</td>
			<td>1.3877</td>
			<td>99</td>
		</tr>
		<tr>
			<td>2</td>
			<td>23.0</td>
			<td>0.9</td>
			<td>278.2</td>
			<td>21.8</td>
			<td>1.1677</td>
			<td>1.4145</td>
			<td>100</td>
		</tr>
		<tr>
			<td>3</td>
			<td>24.8</td>
			<td>1.0</td>
			<td>300.0</td>
			<td>0.0</td>
			<td>1.1438</td>
			<td>1.3904</td>
			<td>100</td>
		</tr>
		<tr>
			<td>4</td>
			<td>21.8</td>
			<td>0.9</td>
			<td>263.7</td>
			<td>36.3</td>
			<td>1.1438</td>
			<td>1.3904</td>
			<td>100</td>
		</tr>
	</tbody>
</table>
Table 1: Calculation for 28-day infusion via osmotic pumps.

Watch this Scientific Journal Video about Subcutaneous Angiotensin II Infusion using Osmotic Pumps Induces Aortic Aneurysms in Mice at JoVE.com

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

Subcutaneous implantation of osmotic pumps provides a convenient approach for prolonged and consistent delivery of compounds. This approach has been used extensively to study both abdominal and thoracic aortic aneurysms in mice.

Osmotic pumps continuously deliver compounds at a constant rate into small animals.  This article introduces a standard protocol used to induce aortic ...

subcutaneous-angiotensin-ii-infusion-using-osmotic-pumps-induces

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

37.0K Views.  University of Kentucky. Subcutaneous implantation of osmotic pumps provides a convenient approach for prolonged and consistent delivery of compounds. This approach has been used extensively to study both abdominal and thoracic aortic aneurysms in mice.

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

Visualizing Leukocyte Rolling and Adhesion in Angiotensin II-Infused Mice: Techniques and Pitfalls

Epifluorescence intravital video microscopy (IVM) of blood vessels is an established method to evaluate the activation of immune cells and their ability to role and adhere to the endothelial layer. Visualization of circulating cells by injection of fluorescent dyes or fluorophore-coupled antibodies is commonly used. Alternatively, fluorescent reporter mice can be used. Interactions of leukocytes, in particular lysozyme M+ (LysM+) monocytes, with the vessel wall play pivotal roles in promoting vascular dysfunction and arterial hypertension. We here present the technique to visualize and quantify leukocyte rolling and adhesion in carotid arteries in angiotensin II (AngII)-induced hypertension in mice by IVM.
The implantation of a catheter damages the vascular wall and leads to altered blood cell responses. We compared different injection techniques and administration routes to visualize leukocytes in a LysMCre+IRG+ mouse with widespread expression of red fluorescent protein and conditional expression of green fluorescent protein in LysM+ cells. To study LysM+ cell activation, we used AngII infused mice in which rolling and adhesion of leukocytes to the endothelium is increased. We either injected acridine orange using a jugular catheter or directly though the tail vein and compared the amount of rolling and adhering cells. We found that jugular catheter implantation per se increased the number of rolling and adhering LysM+ cells in sham-infused LysMCre+IRG+ mice compared to controls. This activation was augmented in AngII-infused mice. Interestingly, injecting acridine orange directly through the tail vein did not increase LysM+ cell adhesion or rolling in sham-infused mice. We thereby demonstrated the importance of transgenic reporter mice expressing fluorescent proteins to not interfere with in vivo processes during experimentation. Furthermore, tail vein injection of fluorescent tracers might be a possible alternative to jugular catheter injections.

This manuscript describes the use of transgenic reporter mice and different administration routes of fluorescent dyes in angiotensin II-induced hypertension using intravital video microscopy of blood vessels to evaluate the activation of immune cells and their ability to roll and adhere to the endothelium.

Epifluorescence intravital video microscopy (IVM) of blood vessels is an established method to evaluate the activation of immune cells and their ability ...

Immunology and Infection

Induction of Atherosclerotic Plaques Through Activation of Mineralocorticoid Receptors in Apolipoprotein E-deficient Mice

Atherosclerosis is due to a chronic inflammatory response affecting vascular endothelium and is promoted by several factors such as hypertension, dyslipidemia, and diabetes. To date, there is evidence to support a role for circulating aldosterone as a risk factor for the development of cardiovascular disease. Transgenic mouse models have been generated to study cellular and molecular processes leading to atherosclerosis. In this manuscript, we describe a protocol that takes advantage of continuous infusion of aldosterone in ApoE-/- mice and generates atherosclerotic plaques in the aortic root after 4 weeks of treatment. We, therefore, illustrate a method for quantification and characterization of atherosclerotic lesions at the aortic root level. The added value of aldosterone infusion is represented by the generation of atherosclerotic lesions rich in lipid and inflammatory cells after 4 weeks of treatment. We describe in detail the staining procedures to quantify lipid and macrophage content within the plaque. Notably, in this protocol, we perform heart tissue-embedding in OCT in order to preserve the antigenicity of cardiac tissue and facilitate detectability of antigens of interest. Analysis of the plaque phenotype represents a valid approach to study the pathophysiology of atherosclerosis development and to identify novel pharmacological targets for the development of anti-atherogenic drugs.

We describe a protocol to induce atherosclerosis in the aortic root of ApoE-/- mice fed with an atherogenic diet, through a continuous release of aldosterone. Methods to characterize plaque composition are also described.

Atherosclerosis is due to a chronic inflammatory response affecting vascular endothelium and is promoted by several factors such as hypertension, ...

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 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.

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. ...

Improved Renal Denervation Mitigated Hypertension Induced by Angiotensin II Infusion

The benefits of renal sympathetic denervation (RDN) on blood pressure have been proved in a large number of clinical trials in recent years. However, the regulatory mechanism of RDN on hypertension remains elusive. Thus, it&#39;s essential to establish a simpler RDN model in mice. In this study, osmotic mini pumps filled with Angiotensin II were implanted in 14-week-old C57BL/6 mice. One week after the implantation of the mini-osmotic pump, a modified RDN procedure was performed on bilateral renal arteries of the mice using phenol. Age-sex-matched mice were given saline and served as sham group. Blood pressure was measured at baseline and every week subsequently for 21 days. Then, renal artery, abdominal aorta and heart were collected for histological examination using H&#38;E and Masson staining. In this study, we present a simple, practical, repeatable, and standardized RDN model, which can control hypertension and alleviate cardiac hypertrophy. The technique can denervate peripheral renal sympathetic nerves without renal artery damage. Compared to previous models, the modified RDN facilitates the study of the pathobiology and pathophysiology of hypertension.

Here, we present a protocol for renal sympathetic denervation (RDN) in mice with hypertension induced by Angiotensin II infusion. The procedure is repeatable, convenient and allows to study the regulatory mechanisms of RDN on hypertension and cardiac hypertrophy.

The benefits of renal sympathetic denervation (RDN) on blood pressure have been proved in a large number of clinical trials in recent years. However, the ...

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 ...

Investigating the Molecular Mechanisms Underlying Murine Abdominal Aortic Aneurysm

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

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

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

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

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

Surgically Induced Cardiac Volume Overload by Aortic Regurgitation in Mouse

Aortic regurgitation (AR) is a common valvular heart disease that exerts volume overload on the heart and represents a global public health problem. Although mice are widely applied to shed light on the mechanisms of cardiovascular disease, mouse models of AR, especially those induced by surgery, are still paucity. Here, a mouse model of AR was described in detail which is surgically induced by disruption of the aortic valves under high-resolution echocardiography. In accordance with regurgitated blood flow, AR mouse hearts present a distinctive and clinically relevant volume overload phenotype, which is characterized by eccentric hypertrophy and cardiac dysfunction, as evidenced by echocardiographic and invasive hemodynamic evaluation. Our proposal, in a reliable and reproducible manner, provides a practical guide on the establishment and assessment of a mouse model of AR for future studies on molecular mechanisms and therapeutic targets of volume overload cardiomyopathy.

This protocol presents a practical guide on the surgery for creation of aortic regurgitation (AR) in the mouse. Assessment of the AR mouse by echocardiography and invasive hemodynamic measurement recapitulates its clinically relevant characteristics of volume overload-induced eccentric hypertrophy, suggesting its promising application in the study of cardiac hypertrophy.

Aortic regurgitation (AR) is a common valvular heart disease that exerts volume overload on the heart and represents a global public health problem. ...

Biology

Mouse Abdominal Aortic Aneurysm Model Induced by Perivascular Application of Elastase

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

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

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

A Calcium Phosphate-Induced Mouse Abdominal Aortic Aneurysm Model

An abdominal aortic aneurysm (AAA) is a life-threatening cardiovascular disease that occurs worldwide and is characterized by irreversible dilation of the abdominal aorta. Currently, several chemically induced murine AAA models are used, each simulating a different aspect of the pathogenesis of AAA. The calcium phosphate-induced AAA model is a rapid and cost-effective model compared to the angiotensin II- and elastase-induced AAA models. The application of CaPO4 crystals to the mouse aorta results in elastic fiber degradation, loss of smooth muscle cells, inflammation, and calcium deposition associated with aortic dilation. This article introduces a standard protocol for the CaPO4-induced AAA model. The protocol includes material preparation, the surgical application of the CaPO4&#160;to the adventitia of the infrarenal abdominal aorta, the harvesting of aortas to visualize aortic aneurysms, and histological analyses in mice.

This protocol describes a calcium phosphate-induced abdominal aortic aneurysm (AAA) mouse model to study the pathological features and molecular mechanisms of AAAs.

An abdominal aortic aneurysm (AAA) is a life-threatening cardiovascular disease that occurs worldwide and is characterized by irreversible dilation of the ...