Name: subcutaneous angiotensin ii infusion using osmotic pumps induces aortic aneurysms in mice
Uploaded: 2015-09-28T15:00:00
Duration: 7 min 21 s
Description: Watch this Scientific Journal Video about Subcutaneous Angiotensin II Infusion using Osmotic Pumps Induces Aortic Aneurysms in Mice at JoVE.com

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 border="0" cellpadding="0" cellspacing="0" width="711">
	<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 &#62; 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 ...

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Research

JoVE Journal

Medicine

Orthotopic Aortic Transplantation in Mice for the Study of Vascular Disease

Vascular procedures involving anastomoses in the mouse are generally thought to be difficult and highly dependent on the skill of the individual surgeon. This is largely true, but there are a number of important principles that can reduce the difficulty of these procedures and enhance reproducibility. Orthotopic aortic transplantation is an excellent procedure in which to learn these principles because it involves only two end-to-end anastomoses, but requires good suturing technique and handling of the vessels for consistent success. This procedure begins with the procurement of a length of abdominal aorta from a donor animal, followed by division of the native aorta in the recipient. The procured aorta is then placed between the divided ends of the recipient aorta and sutured into place using end-to-end anastomoses. To accomplish this objective successfully requires a high degree of concentration, good tools, a steady hand, and an appreciation of how easily the vasculature of a mouse can be damaged, resulting in thrombosis. Learning these important principles is what occupies most of the beginner's time when learning microsurgery in small rodents. Throughout this protocol, we refer to these important points. This model can be used to study vascular disease in a variety of different experimental systems1-8. In the context shown here, it is most often used for the study of post-transplant vascular disease, a common long-term complication of solid organ transplantation in which intimal hyperplasia occurs within the allograft. The primary advantage of the model is that it facilitates quantitative morphometric analyses and the transplanted vessel lies contiguous to the endogenous vessel, which can serve as an additional control9. The technique shown here is most often used for mice weighing 18-25 grams. We have accumulated most of our experience using the C57BL/6J, BALB/cJ, and C3H/HeJ strains.

We describe a technique in which a section of the abdominal aorta from a mouse is transplanted orthotopically to just below the renal arteries in an allogeneic or syngeneic recipient. This technique can be useful in studies in which transplantation of large arteries of uniform size is deemed advantageous.

Vascular procedures involving anastomoses in the mouse are generally thought to be difficult and highly dependent on the skill of the individual surgeon. ...

Generation of Subcutaneous and Intrahepatic Human Hepatocellular Carcinoma Xenografts in Immunodeficient Mice

In vivo experimental models of hepatocellular carcinoma (HCC) that recapitulate the human disease provide a valuable platform for research into disease pathophysiology and for the preclinical evaluation of novel therapies. We present a variety of methods to generate subcutaneous or orthotopic human HCC xenografts in immunodeficient mice that could be utilized in a variety of research applications. With a focus on the use of primary tumor tissue from patients undergoing surgical resection as a starting point, we describe the preparation of cell suspensions or tumor fragments for xenografting. We describe specific techniques to xenograft these tissues i) subcutaneously; or ii) intrahepatically, either by direct implantation of tumor cells or fragments into the liver, or indirectly by injection of cells into the mouse spleen. We also describe the use of partial resection of the native mouse liver at the time of xenografting as a strategy to induce a state of active liver regeneration in the recipient mouse that may facilitate the intrahepatic engraftment of primary human tumor cells. The expected results of these techniques are illustrated. The protocols described have been validated using primary human HCC samples and xenografts, which typically perform less robustly than the well-established human HCC cell lines that are widely used and frequently cited in the literature. In comparison with cell lines, we discuss factors which may contribute to the relatively low chance of primary HCC engraftment in xenotransplantation models and comment on technical issues that may influence the kinetics of xenograft growth. We also suggest methods that should be applied to ensure that xenografts obtained accurately resemble parent HCC tissues.

Human tumor xenografts in immunodeficient mice are valuable tools to study cancer biology. Specific protocols to generate subcutaneous and intrahepatic xenografts from human hepatocellular carcinoma cells or tumor fragments are described. Liver regeneration induced by partial hepatectomy in recipient mice is presented as a strategy to facilitate intrahepatic engraftment.

In  vivo experimental models of hepatocellular carcinoma (HCC) that recapitulate  the human disease provide a valuable platform for research into disease  ...

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

We present a protocol for measuring in vivo aortic stiffness in mice using high-resolution ultrasound imaging. Aortic diameter is measured by ultrasound and aortic blood pressure is measured invasively with a solid-state pressure catheter. Blood pressure is raised then lowered incrementally by intravenous infusion of vasoactive drugs phenylephrine and sodium nitroprusside. Aortic diameter is measured for each pressure step to characterize the pressure-diameter relationship of the ascending aorta. Stiffness indices derived from the pressure-diameter relationship can be calculated from the data collected. Calculation of arterial compliance is described in this protocol.
This technique can be used to investigate mechanisms underlying increased aortic stiffness associated with cardiovascular disease and aging. The technique produces a physiologically relevant measure of stiffness compared to ex vivo approaches because physiological influences on aortic stiffness are incorporated in the measurement. The primary limitation of this technique is the measurement error introduced from the movement of the aorta during the cardiac cycle. This motion can be compensated by adjusting the location of the probe with the aortic movement as well as making multiple measurements of the aortic pressure-diameter relationship and expanding the experimental group size.

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

We describe a technique for measuring aortic stiffness from its pressure-diameter relationship in vivo in mice. Aortic diameter is recorded by ultrasound and aortic pressure is measured invasively with a solid-state pressure catheter. Blood pressure is changed incrementally and the resulting diameter is measured.

We present a protocol for measuring in vivo aortic stiffness in mice using high-resolution ultrasound imaging. Aortic diameter is measured by ultrasound ...

Neurodegeneration in an Animal Model of Chronic Amyloid-beta Oligomer Infusion Is Counteracted by Antibody Treatment Infused with Osmotic Pumps

Decline in hippocampal-dependent explicit memory (memory for facts and events) is one of the earliest clinical symptom of Alzheimer&#39;s disease (AD). It is well established that synapse loss and ensuing neurodegeneration are the best predictors for memory impairments in AD. Latest studies have emphasized the neurotoxic role of soluble amyloid-beta oligomers (A&#946;o) that begin to accumulate in the human brain approximately 10 to 15 yr before the clinical symptoms become apparent. Many reports indicate that soluble A&#946;o correlate with memory deficits in AD models and humans. The A&#946;o-induced neurodegeneration observed in neuronal and brain slice cultures has been more challenging to reproduce in many animal models. The model of repeated A&#946;o infusions shown here overcome this issue and allow addressing two key domains for developing new disease modifying therapies: identify biological markers to diagnose early AD, and determine the molecular mechanisms underpinning A&#946;o-induced memory deficits at the onset of AD. Since soluble A&#946;o aggregate relatively fast into insoluble A&#946; fibrils that correlate poorly with the clinical state of patients, soluble A&#946;o are prepared freshly and injected once per day during six days to produce marked cell death in the hippocampus. We used cannula specially design for simultaneous infusions of A&#946;o and continuous infusion of A&#946;o antibody (6E10) in the hippocampus using osmotic pumps. This innovative in vivo method can now be used in preclinical studies to validate the efficiency of new AD therapies that might prevent the deposition and neurotoxicity of A&#946;o in pre-dementia patients.

To study the effects of A&#946;o in vivo, we developed a model based on repeated hippocampal infusions of soluble A&#946;o coupled with continuous infusion of A&#946;o antibody (6E10) in the hippocampus using osmotic pumps to counteract the neurotoxic effect of A&#946;o.

Decline in hippocampal-dependent explicit memory (memory for facts and events) is one of the earliest clinical symptom of Alzheimer&#39;s disease (AD). It ...

Minimally Invasive Transverse Aortic Constriction in Mice

Minimally invasive transverse aortic constriction (MTAC) is a more desirable method for the constriction of the transverse aorta in mice than standard open-chest transverse aortic constriction (TAC). Although transverse aortic constriction is a highly functional method for the induction of high pressure in the left ventricle, it is a more difficult and lengthy procedure due to its use of artificial ventilation with tracheal intubation. TAC is oftentimes also less survivable, as the newer method, MTAC, neither requires the cutting of the ribs and intercostal muscles nor tracheal intubation with a ventilation setup. In MTAC, as opposed to a thoracotomy to access to the chest cavity, the aortic arch is reached through a midline incision in the anterior neck. The thyroid is pulled back to reveal the sternal notch. The sternum is subsequently cut down to the second rib level, and the aortic arch is reached simply by separating the connective tissues and thymus. From there, a suture can be wrapped around the arch and tied with a spacer, and then the sternal cut and skin can be closed. MTAC is a much faster and less invasive way to induce left ventricular hypertension and enables the possibility for high-throughput studies. The success of the constriction can be verified using high-frequency trans-thoracic echocardiography, particularly color Doppler and pulsed-wave Doppler, to determine the flow velocities of the aortic arch and left and right carotid arteries, the dimension of the blood vessels, and the left ventricular function and morphology. A successful constriction will also trigger significant histopathological changes, such as cardiac muscle cell hypertrophy with interstitial and perivascular fibrosis. Here, the procedure of MTAC is described, demonstrating how the resulting flow changes in the carotid arteries can be examined with echocardiography, gross morphology, and histopathological changes in the heart.

Minimally invasive transverse aortic constriction (MTAC) conserves the essentials of regular transverse aortic constriction (TAC) while eliminating the use of a ventilator with tracheal intubation. It proves to be a highly desirable method for high-throughput studies on left ventricular overload, particularly in translational studies.

Minimally invasive transverse aortic constriction (MTAC) is a more desirable method for the constriction of the transverse aorta in mice than standard ...

A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to perform a partial thoracotomy to visualize the transverse aortic arch. However, the surgical trauma caused by the thoracotomy in open-chest models changes the respiratory physiology as the ribs are dissected and left unattached after chest closure. To prevent this, we established a minimally invasive, closed chest approach via lateral thoracotomy. Herein we approach the aortic arch via the 2nd intercostal space without entering the chest cavities, leaving the mouse with a less traumatic injury to recover from. We perform this operation using standard laboratory settings for open chest TAC procedures with equal survival rates. Apart from maintaining physiological breathing patterns due to the closed chest approach, the mice seem to benefit by showing rapid recovery, as the less invasive technique appears to facilitate a fast healing process and to reduce immune response after trauma.

Here, we present a protocol of Transverse Aortic constriction (TAC) via a lateral thoracotomy. This technique is a minimally invasive, closed chest surgical procedure aiming to simulate pressure overload and heart failure in mice utilizing standard TAC laboratory settings.

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to ...

Improved Registration of 3D CT Angiography with X-ray Fluoroscopy for Image Fusion During Transcatheter Aortic Valve Implantation

The fusion of 3D anatomical models derived from high-fidelity pre-interventional computed tomography angiography (CTA), and x-ray (XR) fluoroscopy to facilitate anatomical guidance is of huge interest for complex cardiac interventions like TAVI procedures with cerebral protection. Co-registration of CTA and XR has been introduced either based on additional intraoperative non-/contrast-enhanced cone-beam computed tomography (CBCT) or two separate aortograms. With the related increase of radiation exposure and/or contrast agent (CA) dose, a potential additional risk for the patient is introduced. Here, we propose a modified co-registration approach making use of arteriograms of the iliofemoral arteries, routinely performed during the femoral puncture and sheath introduction. On-the-fly refinement of the co-registration during the on-going procedure enables accurate co-registration without any additional angiograms, thus reducing CA, XR dose and procedure time, while simultaneously improving operator confidence and procedure safety.

The aim of this study was to improve the co-registration for image fusion (IF) of pre-interventional CT data with real-time x-ray (XR) fluoroscopy during transfemoral transcatheter aortic valve implantation (TAVI).

The fusion of 3D anatomical models derived from high-fidelity pre-interventional computed tomography angiography (CTA), and x-ray (XR) fluoroscopy to ...

Electrocardiogram Recordings in Anesthetized Mice using Lead II

The electrocardiogram is a valuable tool for evaluating the cardiac conduction system. Animal research has helped generate novel genetic and pharmacological information regarding the electrocardiogram. However, making electrocardiogram measurements in small animals in vivo, such as mice, has been challenging. To this end, we used an electrocardiogram recording method in anesthetized mice with many advantages: it is a technically simple procedure, is inexpensive, has short measuring time, and is affordable, even in young mice. Despite the limitations with using anesthesia, comparisons between control and experimental groups can be performed with enhanced sensitivity. We treated mice with agonists and antagonists of the autonomic nervous system to determine the validity of this protocol and compared our results with previous reports. Our ECG protocol detected increased heart rates and QTc intervals on treatment with atropine, decreased heart rates and QTc intervals after carbachol treatment, and higher heart rates and QTc intervals with isoprenaline but did not note any change in ECG parameters on administration of propranolol. These results are supported by previous reports, confirming the reliability of this ECG protocol. Thus, this method can be used as a screening approach to making ECG measurements that otherwise would not be attempted due to high cost and technical difficulties.

We present an ECG protocol that is technically easy, inexpensive, fast, and affordable in small mice, and can be performed with enhanced sensitivity. We suggest this method as a screening approach for studying pharmacological agents, genetic modifications, and disease models in mice.

The electrocardiogram is a valuable tool for evaluating the cardiac conduction system. Animal research has helped generate novel genetic and ...

The Intra-Aortic Balloon Pump

Cardiogenic shock remains one of the most challenging clinical syndromes in modern medicine. Mechanical support is being increasingly used in the management of cardiogenic shock. Intra-aortic balloon pump (IABP) is one of the earliest and most widely used types of mechanical circulatory support. The device acts by external counterpulsation and uses systolic unloading and diastolic augmentation of aortic pressure to improve hemodynamics. Although IABP provides less hemodynamic support when compared with newer mechanical circulatory support devices, it can still be the mechanical support device of choice in appropriate situations because of its relative simplicity of insertion and removal, need for smaller size vascular access and better safety profile. In this review, we discuss the equipment, procedural and technical aspects, hemodynamic effects, indications, evidence, current status and recent advances in the use of IABP in cardiogenic shock.

We describe the steps for the percutaneous implantation of the intra-aortic balloon pump (IABP), a mechanical circulatory support device. It acts by counterpulsation, inflating at the onset of diastole, augmenting diastolic aortic pressure and improving coronary blood flow and systemic perfusion, and deflating before systole, reducing left ventricular afterload.

Cardiogenic shock remains one of the most challenging clinical syndromes in modern medicine. Mechanical support is being increasingly used in the ...

Minimizing Post-Infusion Portal Vein Bleeding during Intrahepatic Islet Transplantation in Mice

Although the liver is currently accepted as the primary transplantation site for human islets in clinical settings, islets are transplanted under the kidney capsule in most rodent preclinical islet transplantation studies. This model is commonly used because murine intrahepatic islet transplantation is technically challenging, and a high percentage of mice could die from surgical complications, especially bleeding from the injection site post-transplantation. In this study, two procedures that can minimize the incidence of post-infusion portal vein bleeding are demonstrated. The first method applies an absorbable hemostatic gelatin sponge to the injection site, and the second method involves penetrating the islet injection needle through the fat tissue first and then into the portal vein by using the fat tissue as a physical barrier to stop bleeding. Both methods could effectively prevent bleeding-induced mouse death. The whole liver section showing islet distribution and evidence of islet thrombosis post-transplantation, a typical feature for intrahepatic islet transplantation, were presented. These improved protocols refine the intrahepatic islet transplantation procedures and may help laboratories set up the procedure to study islet survival and function in pre-clinical settings.

Here we present refined surgical procedures on successfully performing intraportal islet transplantation, a clinically relevant but technically challenging surgical procedure, in mice.

Although the liver is currently accepted as the primary transplantation site for human islets in clinical settings, islets are transplanted under the ...