This work was supported by a National Research Foundation of Korea (NRF) grant funded by the Korean government (MSIT) (No. 2020R1A4A3079570), the Ministry of Education (No. 2021R1I1A1A01051425), and the Industrial Strategic Technology Development Program (No. 20014873) funded by the Ministry of Trade, Industry &#38; Energy, Republic of Korea.

All animal research procedures were approved and performed in accordance with the Laboratory Animals Welfare Act, the Guide for the Care and Use of Laboratory Animals, and the Guidelines and Policies for Animal Experiments provided by the Institutional Animal Care and Use Committee (IACUC) at the College of Medicine of The Catholic University of Korea (approval number: CUMC-2021-0176-05). The present study utilized 3 month old male New Zealand white (NZW) rabbits weighing 3.5-4.0 kg, which were maintained under standard conditions in individual cages. The rabbits were fed either a normal diet or a 0.5% cholesterol-enriched diet supplemented with 50,000 U of vitamin D2 (see Table of Materials). The experimental design and analysis methods for the induction of the AVS rabbit model are depicted in Figure 1.
1. Preparation for the operation
<ol>
	<li>Ensure that all the medical and surgical instruments (see Table of Materials) are sterilized at the beginning of the operation.</li>
	<li>Prepare the dilation balloon catheter set following the steps below.
	<ol>
		<li>Connect the in-deflation device filled with a mixture of saline and commercially available contrast medium (1:1) to the luer lock part of the balloon catheter (see Table of Materials).</li>
		<li>Fill the balloon with inflation solution, and remove any air from the balloon catheter. 
		&#8203;NOTE: For the present study, the inflation solution consisted of 30% iodixanol with 0.9% saline (see Table of Materials).</li>
		<li>Verify proper balloon inflation by purging the balloon lumen with the inflation solution.</li>
	</ol>
	</li>
</ol>
2. Surgical procedure for the aortic valve injury
<ol>
	<li>Administer an intramuscular injection of tiletamine &#38; zolazepam (15 mg/kg) and xylazine (5 mg/kg) (see Table of Materials) to anesthetize the animal. 
	NOTE: Before administering the anesthesia, the rabbits were pre-treated with a subcutaneous glycopyrrolate injection (0.05 mg/kg) as a pre-anesthetic anticholinergic agent. The adequate anesthesia level was determined by a combination of criteria, including a lack of response to a toe pinch and a steady respiratory rate.</li>
	<li>Insert a 24 G intravenous (IV) catheter into the marginal auricular vein, and connect an infusion set with heparinized saline (100 U/kg heparin).</li>
	<li>Connect the rabbit with a multiparameter veterinary monitor (see Table of Materials) to continuously monitor the vital signs, such as the oxygen saturation signal (SpO2), temperature, and blood pressure. 
	NOTE: For the SpO2 monitoring, attach the SpO2 sensor to the rabbit&#39;s tongue. For the temperature monitoring, insert the probe into the rabbit&#39;s rectum. For the blood pressure monitoring, place the cuff on the forelimb.</li>
	<li>Place the rabbit in a supine position on an operating table equipped with a C-arm fluoroscopy (see Table of Materials), and remove the hair from the ventral neck area using animal hair clippers (Figure 2A).</li>
	<li>Sterilize the incision area with iodine, and cover the rabbit with surgical towels.</li>
	<li>Position the rabbit&#39;s heart at the center of the C-arm image. 
	NOTE: All researchers must wear protective gear with attached thermoluminescent dosimeters (TLDs) to reduce radiation exposure while performing the C-arm-guided surgery.
	<ol>
		<li>Turn on the C-arm, and select the fluoroscopic mode for cardiac imaging.</li>
		<li>Adjust the position of the rabbit to ensure that the heart is at the center of the imaging field.</li>
	</ol>
	</li>
	<li>Make a longitudinal incision of approximately 3 cm in the skin of the neck, and use surgical scissors to cut the fascia and fat tissue.</li>
	<li>Expose the left common carotid artery (LCCA) by carefully separating the muscles until approximately 3-3.5 cm of the LCCA is exposed (Figure 2B).</li>
	<li>Ligate the LCCA with a 3-0 silk suture (see Table of Materials) at the top and end of the exposed LCCA to stop the blood flow.</li>
	<li>Insert a 22 G IV catheter into the LCCA, and introduce a guide wire (0.016 in, see Table of Materials) into the left ventricle (LV) through the IV catheter, ensuring that the tip of the catheter is properly positioned in the imaging field of the C-arm. 
	NOTE: When inserting the IV catheter, carefully loosen the ligature suture on the way down to the aortic valve to allow for the advancement of the catheter.</li>
	<li>Withdraw the IV catheter, leaving the guide wire, and place a 4-F sheath (see Table of Materials) over the guide wire into the LCCA to introduce the balloon catheter (Figure 2C). 
	NOTE: After replacing the IV catheter with the sheath, any trapped air should be removed from the sheath device.</li>
	<li>Carefully insert the 8 mm balloon catheter over the guidewire into the aortic valve under C-arm fluoroscopic guidance (Figure 2D).</li>
	<li>Place the balloon catheter tip approximately 1-2 cm distal to the aortic valve, and inflate the balloon by purging the inflation solution with a pressure inflator at 6 atm.</li>
	<li>Advance the balloon into the LV apex, and pull it back into the LV outlet. Repeat this procedure five times, and then deflate the balloon (Figure 2E, F).</li>
	<li>Repeat steps 2.8-2.9 three times to ensure adequate valve injury.</li>
	<li>Withdraw the balloon catheter and guidewire. Slowly remove the sheath from the LCCA, and immediately tie the LCCA with the suture on the way down to the aortic valve.</li>
	<li>Clean the incision area with saline to remove the blood clots, and inspect the punctured site for arterial bleeding.</li>
	<li>Close the muscle and skin with a 3-0 non-absorbable suture, and sterilize all the sides of the wound with iodine.</li>
</ol>
3. Post-operative care
<ol>
	<li>Remove the monitoring patches and clips, and keep the rabbit in an intensive care incubator. 
	NOTE: After surgery, the rabbits were closely observed for 1 day in an intensive care incubator and then moved to a home cage.</li>
	<li>Manage post-operative pain with 5 mg/kg tramadol and 3 mg/kg ketoprofen and administer antibiotics (4 mg/kg gentamicin) twice daily for 3 days via subcutaneous injection (see Table of Materials). 
	NOTE: Post-operative pain management should adhere to veterinary and IACUC guidelines (e.g., opioid, NSAID, local anesthetic or combination).</li>
	<li>Feed a 0.5% cholesterol-enriched diet with 50,000 U of vitamin D2 (HC + VitD2) for 8 weeks.</li>
</ol>
4. Echocardiography
<ol>
	<li>After 8 weeks of balloon injury, anesthetize the rabbit using the same procedure described in step 2.1.</li>
	<li>Visualize the aortic valves using two-dimensional transthoracic views, and record M-mode images in short-axis and long-axis views.
	<ol>
		<li>Place the rabbit in a supine position on an echo table.</li>
		<li>Shave the chest area using clippers and hair removal cream.</li>
		<li>Apply ultrasound transducer gel (see Table of Materials) to the chest.</li>
		<li>Adjust the transducer to obtain the parasternal long-axis view and the parasternal short-axis view of the aortic valve.</li>
		<li>Use M-mode imaging to record images of the aortic valve in both long-axis and short-axis views, and save the images for later analysis.</li>
	</ol>
	</li>
</ol>
5. Histological analysis
<ol>
	<li>After the echocardiography, euthanize the rabbit by administering an intravenous injection of potassium chloride (KCl, 3 g/20 mL, 1 mL).</li>
	<li>Open the thoracic cavity, harvest the heart with the ascending aorta7, and place it on ice in phosphate-buffered saline (PBS).</li>
	<li>Immediately immerse the heart in a 4% paraformaldehyde (PFA) solution, and embed it in a paraffin block (see Table of Materials).</li>
	<li>Cut the paraffin-embedded heart block into 4 &#181;m thick sections using a microtome, and stain the sections with Masson&#39;s trichrome (MT), Alizarin Red, and von Kossa (see Table of Materials) to assess the collagen deposition and valve calcification, respectively8,9.</li>
</ol>

<ol>
	<li>Aliev, G., Burnstock, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Watanabe+rabbits+with+heritable+hypercholesterolaemia:+A+model+of+atherosclerosis.">Watanabe rabbits with heritable hypercholesterolaemia: A model of atherosclerosis.</a> Histology and Histopathology. 13 (3), 797-817 (1998).</li><li>Cimini, M., Boughner, D. R., Ronald, J. A., Aldington, L., Rogers, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+aortic+valve+sclerosis+in+a+rabbit+model+of+atherosclerosis:+An+immunohistochemical+and+histological+study.">Development of aortic valve sclerosis in a rabbit model of atherosclerosis: An immunohistochemical and histological study.</a> Journal of Heart Valve Disease. 14 (3), 365-375 (2005).</li><li>Drolet, M. C., Couet, J., Arsenault, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+aortic+valve+sclerosis+or+stenosis+in+rabbits:+role+of+cholesterol+and+calcium.">Development of aortic valve sclerosis or stenosis in rabbits: role of cholesterol and calcium.</a> Journal of Heart Valve Disease. 17 (4), 381-387 (2008).</li><li>Sider, K. L., Blaser, M. C., Simmons, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+calcific+aortic+valve+disease.">Animal models of calcific aortic valve disease.</a> International Journal of Inflammation. 2011, 364310 (2011).</li><li>Honda, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+mouse+model+of+aortic+valve+stenosis+induced+by+direct+wire+injury.">A novel mouse model of aortic valve stenosis induced by direct wire injury.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 34 (2), 270-278 (2014).</li><li>Niepmann, S. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Graded+murine+wire-induced+aortic+valve+stenosis+model+mimics+human+functional+and+morphological+disease+phenotype.">Graded murine wire-induced aortic valve stenosis model mimics human functional and morphological disease phenotype.</a> Clinical Research in Cardiology. 108 (8), 847-856 (2019).</li><li>Robbins, N., Thompson, A., Mann, A., Blomkalns, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+excision+of+murine+aorta;+A+versatile+technique+in+the+study+of+cardiovascular+disease.">Isolation and excision of murine aorta; A versatile technique in the study of cardiovascular disease.</a> Journal of Visualized Experiments. (93), e52172 (2014).</li><li>Wirrig, E. E., Gomez, M. V., Hinton, R. B., Yutzey, K. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=COX2+inhibition+reduces+aortic+valve+calcification+in+vivo.">COX2 inhibition reduces aortic valve calcification in vivo.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 35 (4), 938-947 (2015).</li><li>Jung, S. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiotemporal+dynamics+of+macrophage+heterogeneity+and+a+potential+function+of+Trem2(hi)+macrophages+in+infarcted+hearts.">Spatiotemporal dynamics of macrophage heterogeneity and a potential function of Trem2(hi) macrophages in infarcted hearts.</a> Nature Communications. 13 (1), 4580 (2022).</li><li>Freeman, R. V., Otto, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spectrum+of+calcific+aortic+valve+disease:+Pathogenesis,+disease+progression,+and+treatment+strategies.">Spectrum of calcific aortic valve disease: Pathogenesis, disease progression, and treatment strategies.</a> Circulation. 111 (24), 3316-3326 (2005).</li><li>Lindman, B. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcific+aortic+stenosis.">Calcific aortic stenosis.</a> Nature Reviews Disease Primers. 2, 16006 (2016).</li><li>Cuniberti, L. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+mild+aortic+valve+stenosis+in+a+rabbit+model+of+hypertension.">Development of mild aortic valve stenosis in a rabbit model of hypertension.</a> Journal of the American College of Cardiology. 47 (11), 2303-2309 (2006).</li><li>Marechaux, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+tissue+factor+in+experimental+aortic+valve+sclerosis.">Identification of tissue factor in experimental aortic valve sclerosis.</a> Cardiovascular Pathology. 18 (2), 67-76 (2009).</li><li>Hara, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progression+of+calcific+aortic+valve+sclerosis+in+WHHLMI+rabbits.">Progression of calcific aortic valve sclerosis in WHHLMI rabbits.</a> Atherosclerosis. 273, 8-14 (2018).</li></ol>

The authors have no conflicts of interest to declare with this work.

Animal AVS models are commonly used to study the pathological aspects of AVS, including the initiation and progression of AVS. This protocol introduces a new rabbit AVS model induced by a direct balloon injury to the aortic valve. In this study, the aortic valve injury model showed significant leaflet thickening and calcification. Compared to the mild AVS model induced by dietary supplementation, the aortic valve in the direct balloon injury model was selectively injured, leading to thickened cusps and restricted motion,...

It is increasingly recognized that the use of appropriate animal models can contribute to a better understanding of the pathological mechanisms underlying aortic valve stenosis (AVS) due to the lack of access to reliable sources of diseased human aortic valves associated with the progression of aortic stenosis (AS). Among the various animal models for studying AVS, rabbits are one of the most commonly used large-animal AVS models, and the AVS rabbit model is induced either through cholesterol/vitamin D2 supplementation or genetic manipulation1,2,3,4.
Although rabbit AVS models have provided significant insight into the development and progression of AVS, it still remains challenging to induce AVS consistently and reproducibly, as seen in our preliminary experiments.
In addition to diet-induced and genetically susceptible animal models, a new model of AVS has been established through direct mechanical injury in mice5,6. The mechanical injury model successfully induces aortic stenosis and represents a simple and reproducible AVS model in wild-type mice. To the best of our knowledge, there have been no prior studies examining the effects of a mechanical injury on the aortic valve in rabbit models. Thus, this study provides a new procedure for inducing AVS in male New Zealand white rabbits through a direct balloon injury to the aortic valve, which can accurately mimic the condition of valvular aortic stenosis. This protocol includes step-by-step descriptions of the preparation, the surgical procedure, and the post-operative care, which are useful for inducing reproducible AVS rabbit models.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3-0 Silk suture</td>
			<td>AILEE</td>
			<td>SK312</td>
			<td></td>
		</tr>
		<tr>
			<td>4% paraformaldehyde(PFA)</td>
			<td>Intron</td>
			<td>IBS-BP031-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Alizarin red Solution</td>
			<td>Millpore</td>
			<td>TMS-008-C</td>
			<td></td>
		</tr>
		<tr>
			<td>ASAHI SION BLUE&#160;</td>
			<td>ASAHI</td>
			<td></td>
			<td>Guide wire</td>
		</tr>
		<tr>
			<td>Back Table Cover</td>
			<td>Yuhan kimberly</td>
			<td>80101-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Balloon In-deflation Device</td>
			<td>Demax Medical</td>
			<td>DID30s</td>
			<td></td>
		</tr>
		<tr>
			<td>Bionet Veterinary monitor</td>
			<td>BIONET</td>
			<td>BM3 VET</td>
			<td></td>
		</tr>
		<tr>
			<td>C-Arm</td>
			<td>SIEMENS Healthcare GmbH</td>
			<td>Cios alpha</td>
			<td></td>
		</tr>
		<tr>
			<td>Certified Rabbit Diet</td>
			<td>Purina</td>
			<td>5322</td>
			<td>4.7% Hydrogenated Coconut Oil, 0.5% Cholesterol, &#38; 1% Molasse</td>
		</tr>
		<tr>
			<td>Curadle Smart Incubator</td>
			<td>Autoelex</td>
			<td>CS-CV206</td>
			<td>Intensive Care Unit (ICU)</td>
		</tr>
		<tr>
			<td>Ergocalciferol</td>
			<td>Sigma-aldrich&#160;</td>
			<td>E5750</td>
			<td>Vitamin D2</td>
		</tr>
		<tr>
			<td>Fechtner conjunctiva forceps titanium</td>
			<td>WORLD PRECISSION Instrument</td>
			<td>WP1820</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>HEBU</td>
			<td>HB203</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Shin Poong</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glycopyrrolate&#160;</td>
			<td>SamChunDang</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Greenflex NS</td>
			<td>DAI HAN PHARM</td>
			<td></td>
			<td>Normal saline 500 mL</td>
		</tr>
		<tr>
			<td>Hematoxylin solution</td>
			<td>Sigma-aldrich&#160;</td>
			<td>HT1079-1 SET</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>JW pharmaceutical</td>
			<td></td>
			<td>25,000 U</td>
		</tr>
		<tr>
			<td>Infusion set for single use</td>
			<td>SWOON MEDICAL</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Iodine</td>
			<td>Green pharmaceutical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Iodixanol</td>
			<td>GE Healthcare</td>
			<td>Visipaque</td>
			<td>Inflation solution (contrast agent)</td>
		</tr>
		<tr>
			<td>IV catheter 22 G</td>
			<td>BD&#160;</td>
			<td>382423</td>
			<td></td>
		</tr>
		<tr>
			<td>IV catheter 24 G</td>
			<td>BD</td>
			<td>382412</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketoprofen</td>
			<td>SamChunDang</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Luer-Lok syringe 10 mL</td>
			<td>Becton Dickinson Medical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Luer-Lok syringe 3 mL</td>
			<td>Becton Dickinson Medical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>OLYMPUS</td>
			<td>SZ61</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtome</td>
			<td>ThermoFisher Scientific</td>
			<td>HM 325</td>
			<td></td>
		</tr>
		<tr>
			<td>MT stain kit</td>
			<td>Sigma-aldrich</td>
			<td>HT15-1kt</td>
			<td></td>
		</tr>
		<tr>
			<td>Needel holder</td>
			<td>Solco</td>
			<td>009-1304</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle Holder with Lock and Suture</td>
			<td>JEUNGDO BIO &#38; PLANT</td>
			<td>H-1222-18</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin</td>
			<td>LK LABKOREA</td>
			<td>H06-660-107</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Gibco</td>
			<td>10010-023</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride 40</td>
			<td>Daihan Pharm</td>
			<td></td>
			<td>KCl</td>
		</tr>
		<tr>
			<td>Prelude Ideal Hydrophilic Sheath</td>
			<td>MERIT MEDICAL</td>
			<td>PID4F11018SS</td>
			<td>Sheath 4F</td>
		</tr>
		<tr>
			<td>PTA Balloon Dilatation catheter</td>
			<td>Boston Scientific</td>
			<td>H749-3903280208-0</td>
			<td>Balloon catheter 8.0 mm</td>
		</tr>
		<tr>
			<td>Rompun</td>
			<td>Elanco</td>
			<td></td>
			<td>Xylaxine</td>
		</tr>
		<tr>
			<td>sterile Gauze</td>
			<td>DAE HAN Medical</td>
			<td></td>
			<td>10 cm x 20 cm&#160;</td>
		</tr>
		<tr>
			<td>Surgical Gloves</td>
			<td>Ansell</td>
			<td>Ansell</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Gown</td>
			<td>Yuhan kimberly</td>
			<td>90002-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scissors</td>
			<td>Nopa, Germany</td>
			<td>AC020/16</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Tape</td>
			<td>3M micopore</td>
			<td>1530-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe 1 mL</td>
			<td>Shin Chang Medical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe 10 mL</td>
			<td>Shin Chang Medical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue cassette</td>
			<td>Scilav korea</td>
			<td>Cas3003</td>
			<td></td>
		</tr>
		<tr>
			<td>Transducer gel&#160;</td>
			<td>SUNGHEUNG</td>
			<td>SH102</td>
			<td></td>
		</tr>
		<tr>
			<td>Tridol</td>
			<td>Yuhan Corp.</td>
			<td></td>
			<td>Tramadol HCl</td>
		</tr>
		<tr>
			<td>Ultrasound system</td>
			<td>Philps</td>
			<td>Affiniti 50</td>
			<td></td>
		</tr>
		<tr>
			<td>Von Kossa stain kit</td>
			<td>Abcam</td>
			<td>ab105689</td>
			<td></td>
		</tr>
		<tr>
			<td>Zoletil 50</td>
			<td>Virbac korea</td>
			<td></td>
			<td>Tiletamine &#38; zolazepam</td>
		</tr>
	</tbody>
</table>

a rabbit aortic valve stenosis model induced by direct balloon injury

Animal models are emerging as an important tool to understand the pathologic mechanisms underlying aortic valve stenosis (AVS) because of the lack of access to reliable sources of diseased human aortic valves. Among the various animal models, AVS rabbit models are one of the most commonly used in large animal studies. However, traditional AVS rabbit models require a long-term period of dietary supplementation and genetic manipulation to induce significant stenosis in the aortic valve, limiting their use in experimental studies. To address these limitations, a new AVS rabbit model is proposed, in which stenosis is induced by a direct balloon injury to the aortic valve. The present protocol describes a successful technique for inducing AVS in New Zealand white (NZW) rabbits, with step-by-step procedures for the preparation, the surgical procedure, and the post-operative care. This simple and reproducible model offers a promising approach for studying the initiation and progression of AVS and provides a valuable tool for investigating the underlying pathological mechanisms of the disease.

Rabbit AVS model induced by aortic valve injury 
To induce the rabbit AVS model, male NZW rabbits weighing 3.5-4.0 kg were used for this study. According to the surgical procedures described in step 2 (Figure 2), the AVS model was established by aortic valve injury, which resulted in mechanical aortic valve degeneration and calcification. The control group included rabbits fed with a 0.5% cholesterol-enriched diet (high-cholesterol, HC) and 50,000 U of vitamin D2 (VitD2...

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A Rabbit Aortic Valve Stenosis Model Induced by Direct Balloon Injury

An appropriate animal model is needed to understand the pathologic mechanisms underlying aortic valve stenosis (AVS) and to evaluate the efficacy of therapeutic interventions. The present protocol describes a new procedure for developing the AVS rabbit model via a direct balloon injury in vivo.

Animal models are emerging as an important tool to understand the pathologic mechanisms underlying aortic valve stenosis (AVS) because of the lack of ...

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Research

JoVE Journal

Medicine

878 Views.  The Catholic University of Korea. An appropriate animal model is needed to understand the pathologic mechanisms underlying aortic valve stenosis (AVS) and to evaluate the efficacy of therapeutic interventions. The present protocol describes a new procedure for developing the AVS rabbit model via a direct balloon injury in vivo.

Video: A Rabbit Aortic Valve Stenosis Model Induced by Direct Balloon Injury

A Rat Carotid Balloon Injury Model to Test Anti-vascular Remodeling Therapeutics

The rat carotid balloon injury is a well-established surgical model that has been used to study arterial remodeling and vascular cell proliferation. It is also a valuable model system to test, and to evaluate therapeutics and drugs that negate maladaptive remodeling in the vessel. The injury, or barotrauma, in the vessel lumen caused by an inflated balloon via an inserted catheter induces subsequent neointimal growth, often leading to hyperplasia or thickening of the vessel wall that narrows, or obstructs the lumen. The method described here is sufficiently sensitive, and the results can be obtained in relatively short time (2 weeks after the surgery). The efficacy of the drug or therapeutic against the induced-remodeling can be evaluated either by the post-mortem pathological and histomorphological analysis, or by ultrasound sonography in live animals. In addition, this model system has also been used to determine the therapeutic window or the time course of the administered drug. These studies can leadto the development of a better administrative strategy and a better therapeutic outcome. The procedure described here provides a tool for translational studies that bring drug and therapeutic candidates from bench research to clinical applications.

The rat carotid balloon injury model described below allows researchers to evaluate drugs or therapeutics that negate injury-induced arterial hyperplasia. Detailed pre-surgical preparation, surgical procedure, and post-surgical cares of the animal are described.

The rat carotid balloon injury is a well-established surgical model that has been used to study arterial remodeling and vascular cell proliferation. It is ...

Reproducible Arterial Denudation Injury by Infrarenal Abdominal Aortic Clamping in a Murine Model

Percutaneous vascular interventions uniformly result in arterial denudation injuries that subsequently lead to thrombosis and restenosis. These complications can be attributed to impairments in re-endothelialization within the wound margins. Yet, the cellular and molecular mechanisms of re-endothelialization remain to be defined. While several animal models to study re-endothelialization after arterial denudation are available, few are performed in the mouse because of surgical limitations. This undermines the opportunity to exploit transgenic mouse lines and investigate the contribution of specific genes to the process of re-endothelialization. Here, we present a step-by-step protocol for creating a highly reproducible murine model of arterial denudation injury in the infrarenal abdominal aorta using external vascular clamping. Immunocytochemical staining of injured aortas for fibrinogen and &#946;-catenin demonstrate the exposure of a pro-thrombotic surface and the border of intact endothelium, respectively. The method presented here has the advantages of speed, excellent overall survival rate, and relative technical ease, creating a uniquely practical tool for imposing arterial denudation injury in transgenic mouse models. Using this method, investigators may elucidate the mechanisms of re-endothelialization under normal or pathological conditions.

Understanding the cellular and molecular mechanisms of re-endothelialization following arterial denudation injury is of paramount importance in preventing thrombosis and restenosis of arteries. Here we describe a protocol for reproducible arterial denudation injury of the infrarenal abdominal aorta. The procedure was developed to investigate the underlying mechanisms that regulate endothelial regeneration using mouse models.

Percutaneous vascular interventions uniformly result in arterial denudation injuries that subsequently lead to thrombosis and restenosis. These ...

The Rabbit Model of Accelerated Atherosclerosis: A Methodological Perspective of the Iliac Artery Balloon Injury

Acute coronary syndrome resulting from coronary occlusion following atherosclerotic plaque development and rupture is the leading cause of death in the industrialized world. New Zealand White (NZW) rabbits are widely used as an animal model for the study of atherosclerosis. They develop spontaneous lesions when fed with atherogenic diet; however, this requires long time of 4 - 8 months. To further enhance and accelerate atherogenesis, a combination of atherogenic diet and mechanical endothelial injury is often employed. The presented procedure for inducing atherosclerotic plaques in rabbits uses a balloon catheter to disrupt the endothelium in the left iliac artery of NZW rabbits fed with atherogenic diet. Such mechanical damage caused by the balloon catheter induces a chain of inflammatory reactions initiating neointimal lipid accumulation in a time dependent fashion. Atherosclerotic plaque following balloon injury show neointimal thickening with extensive lipid infiltration, high smooth muscle cell content and presence of macrophage derived foam cells. This technique is simple, reproducible and produces plaque of controlled length within the iliac artery. The whole procedure is completed within 20 - 30 min. The procedure is safe with low mortality and also offers high success in obtaining substantial intimal lesions. The procedure of balloon catheter induced arterial injury results in atherosclerosis within two weeks. This model can be used for investigating the disease pathology, diagnostic imaging and to evaluate new therapeutic strategies.

Animal models of atherosclerosis are essential to understand the mechanism and to investigate newer approaches to prevent plaque development or rupture, a leading cause of death in the industrialized world. This protocol uses a combination of balloon injury and cholesterol rich diet to induce atherosclerotic plaques in rabbit iliac artery.

Acute coronary syndrome resulting from coronary occlusion following atherosclerotic plaque development and rupture is the leading cause of death in the ...

Full-root Aortic Valve Replacement by Stentless Aortic Xenografts in Patients with Small Aortic Roots

In patients with small aortic roots who need an aortic valve replacement with biological valve substitutes, the implantation of the stented pericardial valve might not meet the functional needs. The implantation of a too-small stented pericardial valve, leading to an effective orifice area indexed to a body surface area less than 0.85 cm2/m2, is regarded as prosthesis-patient mismatch (PPM). A PPM negatively affects the regression of left ventricular hypertrophy and thus the normalization of left ventricular function and the alleviation of symptoms. Persistent left ventricular hypertrophy is associated with an increased risk of arrhythmias and sudden cardiac death. In the case of predictable PPM, there are three options: 1) accept the PPM resulting from the implantation of a stented pericardial valve when comorbidities of the patient forbid the more technically demanding operative technique of implanting a larger prosthesis, 2) enlarge the aortic root to accommodate a larger stented valve substitute, or 3) implant a stentless biological valve or a homograft. Compared to classical aortic valve replacement with stented pericardial valves, the full-root implantation of stentless aortic xenografts offers the possibility of implanting a 3-4 mm larger valve in a given patient, thus allowing significant reduction in transvalvular gradients. However, a number of cardiac surgeons are reluctant to transform a classical aortic valve replacement with stented pericardial valves into the more technically challenging full-root implantation of stentless aortic xenografts. Given the potential hemodynamic advantages of stentless aortic xenografts, we have adopted full-root implantation to avoid PPM in patients with small aortic roots necessitating an aortic valve replacement. Here, we describe in detail a technique for the full-root implantation of stentless aortic xenografts, with emphasis on the management of the proximal suture line and coronary anastomoses. Limitations of this technique and alternative options are discussed.

Full-root aortic valve replacement by stentless aortic xenograft is a viable option in patients with small aortic roots. We describe, a technique for the full-root implantation of stentless aortic xenografts, with emphasis on the management of the proximal suture line and coronary anastomoses, and discuss its limitations and alternative options.

In patients with small aortic roots who need an aortic valve replacement with biological valve substitutes, the implantation of the stented pericardial ...

Standardized Technique of Aortic Valve Re-implantation for Valve-sparing Aortic Root Replacement

Despite the obvious advantages of the preservation of a normal aortic valve during aortic root replacement, the complexity of valve sparing procedures prevents a number of cardiac surgeons from incorporating them into their practice. The aim of this protocol is to describe a simplified and user-friendly technique of an aortic valve-sparing root replacement (VSRR) procedure by re-implantation of the aortic valve. Proper selection of patients and limitations of the technique are discussed.
In 54 consecutive patients, normal appearing aortic valves were re-implanted in a commercially available polyester prosthesis with pre-shaped sinuses by a simplified and standardized technique. Placement of the first row of the proximal suture line, choice of the prosthesis size, and adjustment of the height of the commissures of the patient to the fixed height of the sinus portion of the prosthesis were slightly modified from the reference techniques with the aim of increasing its feasibility for use by other cardiac surgeons. Early mortality and morbidity as well as 5-year survival, freedom from aortic valve reoperation, and freedom from recurrent moderate regurgitation were collected in all patients.
Thirty-day mortality, re-sternotomy for bleeding, re-sternotomy for mediastinitis, and the incidence of stroke were very low, 1.8% for each (1 of 54). No patient required permanent pace-maker implantation. At 5 years, survival, freedom from aortic valve reoperation, and freedom from recurrent moderate regurgitation were 97.5%, 95.2%, and 91.6%, respectively.
Mid-term results of our standardized technique of re-implantation of the aortic valve for valve-sparing aortic root replacement are very good and compare with more complex techniques reported by experienced surgeons. By following the present protocol of the standardized re-implantation technique, a greater number of cardiac surgeons can perform this procedure with comparable good results.

Valve-sparing aortic root replacement has the advantage of preserving the patient&#39;s own aortic valve. The complexity of the reported techniques to date restricts their use to a limited number of cardiac surgeons. This protocol describes step-by-step a standardized technique reproducible by a greater number of cardiac surgeons.

Despite the obvious advantages of the preservation of a normal aortic valve during aortic root replacement, the complexity of valve sparing procedures ...

A Rabbit Model of Aqueous-Deficient Dry Eye Disease Induced by Concanavalin A Injection into the Lacrimal Glands: Application to Drug Efficacy Studies

Dry eye disease (DED), a multifactorial inflammatory disease of the ocular surface, affects 1 in 6 humans worldwide with staggering implications for quality of life and health care costs. The lack of informative animal models that recapitulate its key features impedes the search for new therapeutic agents for DED. Available DED animal models have limited reproducibility and efficacy. A model is presented here in which DED is induced by injecting the mitogen concanavalin A (Con A) into the orbital lacrimal glands of rabbits. Innovative aspects of this model are the use of ultrasound (US) guidance to ensure optimal and reproducible injection of Con A into the inferior lacrimal gland; injection of Con A into all orbital lacrimal glands that limits compensatory production of tears; and use of periodic repeat injections of Con A that prolong the state of DED at will. DED and its response to test agents are monitored with a panel of parameters that assess tear production, the stability of the tear film, and the status of the corneal and conjunctival mucosa. They include tear osmolarity, tear break-up time, Schirmer&#39;s tear test, rose bengal staining, and tear lactoferrin levels. The induction of DED and the monitoring of its parameters are described in detail. This model is simple, robust, reproducible, and informative. This animal model is suitable for the study of tear physiology and of the pathophysiology of DED as well as for the assessment of the efficacy and safety of candidate agents for the treatment of DED.

This article describes the development of a method to induce acute or chronic dry eye disease in rabbits by injecting concanavalin A to all portions of the orbital lacrimal gland system. This method, superior to those already reported, generates a reproducible, stable model of dry eye suitable for the study of pharmacological agents.

Dry eye disease (DED), a multifactorial inflammatory disease of the ocular surface, affects 1 in 6 humans worldwide with staggering implications for ...

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

Acute Kidney Injury Model Induced by Cisplatin in Adult Zebrafish

Cisplatin is commonly used as chemotherapy. Although it has positive effects in cancer-treated individuals, cisplatin can easily accumulate in the kidney due to its low molecular weight. Such accumulation causes the death of tubular cells and can induce the development of Acute Kidney Injury (AKI), which is characterized by a quick decrease in kidney function, tissue damage, and immune cells infiltration. If administered in specific doses cisplatin can be a useful tool as an AKI inducer in animal models. The zebrafish has appeared as an interesting model to study renal function, kidney regeneration, and injury, as renal structures conserve functional similarities with mammals. Adult zebrafish injected with cisplatin shows decreased survival, kidney cell death, and increased inflammation markers after 24 h post-injection (hpi). In this model, immune cells infiltration and cell death can be assessed by flow cytometry and TUNEL assay. This protocol describes the procedures to induce AKI in adult zebrafish by intraperitoneal cisplatin injection and subsequently demonstrates how to collect the renal tissue for flow cytometry processing and cell death TUNEL assay. These techniques will be useful to understand the effects of cisplatin as a nephrotoxic agent and will contribute to the expansion of AKI models in adult zebrafish. This model can also be used to study kidney regeneration, in the search for compounds that treat or prevent kidney damage and to study inflammation in AKI. Moreover, the methods used in this protocol will improve the characterization of tissue damage and inflammation, which are therapeutic targets in kidney-associated comorbidities.

This protocol describes the procedures to induce Acute Kidney Injury (AKI) in adult zebrafish using cisplatin as a nephrotoxic agent. We detailed the steps to evaluate the reproducibility of the technique and two techniques to analyze inflammation and cell death in the renal tissue, flow cytometry and TUNEL, respectively.

Cisplatin is commonly used as chemotherapy. Although it has positive effects in cancer-treated individuals, cisplatin can easily accumulate in the kidney ...

A Murine Model of Ischemic Retinal Injury Induced by Transient Bilateral Common Carotid Artery Occlusion

Diverse vascular diseases such as diabetic retinopathy, occlusion of retinal veins or arteries and ocular ischemic syndrome can lead to retinal ischemia. To investigate pathological mechanisms of retinal ischemia, relevant experimental models need to be developed. Anatomically, a main retinal blood supplying vessel is the ophthalmic artery (OpA) and OpA originates from the internal carotid artery of the common carotid artery (CCA). Thus, disruption of CCA could effectively cause retinal ischemia. Here, we established a mouse model of retinal ischemia by transient bilateral common carotid artery occlusion (tBCCAO) to tie the right CCA with 6-0 silk sutures and to occlude the left CCA transiently for 2 seconds via a clamp, and showed that tBCCAO could induce acute retinal ischemia leading to retinal dysfunction. The current method reduces reliance on surgical instruments by only using surgical needles and a clamp, shortens occlusion time to minimize unexpected animal death, which is often seen in mouse models of middle cerebral artery occlusion, and maintains reproducibility of common retinal ischemic findings. The model can be utilized to investigate the pathophysiology of ischemic retinopathies in mice and further can be used for in vivo drug screening.

Here, we describe a mouse model of retinal ischemia by transient bilateral common carotid artery occlusion using simple sutures and a clamp. This model can be useful for understanding the pathological mechanisms of retinal ischemia caused by cardiovascular abnormalities.

Diverse vascular diseases such as diabetic retinopathy, occlusion of retinal veins or arteries and ocular ischemic syndrome can lead to retinal ischemia. ...

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.