Name: implantation of human-sized coronary stents into rat abdominal aorta using a trans-femoral access
Uploaded: 2020-11-19T13:00:00.000+00:00
Duration: 5 min 4 s
Description: Watch this Scientific Journal Video about Implantation of Human-Sized Coronary Stents into Rat Abdominal Aorta Using a Trans-Femoral Access at JoVE.com

We would like to thank Mrs. Angela Freund for her invaluable technical assistance with embedding and slides production. We would also like to thank Mr. Tadeusz Stopinski at the Institute for Laboratory Animal Science &#38; Experimental Surgery for his insightful help with the veterinary work.

The experiments were performed in accordance with the German animal welfare law (TSchG) and Directive 2010/63/EU pertaining to the protection of animals used for scientific purposes. The official approval for this study was granted by the Governmental Animal Care and Use Committee (Protocol No.: AZ 87-51.04.2010.A065; Landesamt für Natur, Umwelt und Verbraucherschutz Nordrhein-Westfalen, Recklinghausen, Germany). The study protocol complied with the Guide for the Care and Use of Laboratory Animals. Postoperative pain treatment is based on the recommendations of the German Society for Laboratory Animal Science (GV-SOLAS) as well as Initiative Veterinary Pain Therapy.
1. Basic techniques and common procedures
<ol>
	<li>Use homozygous apoE-/- Sprague-Dawley rats. Identify the genotype of each animal by using standard methods17.</li>
	<li>Keep the animals under identical conditions (21 °C ± 2 °C, 60% ± 5% humidity, and a 12 h light/dark cycle) and ensure free access to water and food.</li>
	<li>Carry out all procedures under clean but nonsterile conditions.</li>
	<li>Once the rat is anesthetized, perform all procedures under a surgical microscope at a magnification of 16x.</li>
	<li>Use cotton swabs for compression hemostasis. Gauze swabs (5 cm x 5 cm) soaked with lactated Ringer solution are helpful to keep the groin moist.</li>
	<li>Follow waste disposal regulations to dispose used materials.</li>
</ol>
2. Preparations before surgery
<ol>
	<li>Prepare the veterinary drugs before starting the operation. Keep all solutions at room temperature, unless otherwise indicated.</li>
	<li>Thirty minutes before the procedure, administer 0.03-0.05 mg/kg buprenorphine subcutaneously.</li>
	<li>Anesthetize the rat with an intraperitoneal injection of 100 mg/kg body weight (BW) (S)-ketamine and 8 mg/kg BW xylazine.</li>
	<li>Assess the rat’s weight using a weighing scale.</li>
	<li>Place the rat on a heating pad and fix the upper and lower limbs using medical tape. Position the rat with its left hind limb fully extended and as much in line with its spine as possible so as to create a straight line between femoral artery and aorta. This will facilitate advancing the balloon-mounted stent through the aortic bifurcation.</li>
	<li>Maintain anesthesia with inhalation of 1.5 vol% isoflurane in 97.5% oxygen at a flow rate of 2 L/min. 
	NOTE: Allow the rat to breathe spontaneously, without intubation.</li>
	<li>Apply eye ointment to prevent eye damage during unconsciousness.</li>
	<li>Shave the fur from the groin and lower abdomen area of the rat and sterilize the corresponding skin with a povidone-iodine solution.</li>
	<li>Before starting the surgery, verify adequate depth of anesthesia by pinching the tail tip and the interdigital tissue. </li>
</ol>
3. Surgery
<ol>
	<li>Make a medial incision of ~0.5‒1 cm in the left groin to open the skin and the underlying fascia.</li>
	<li>Bluntly dissect and probe in the depths until the pulsating left femoral artery can be identified.</li>
	<li>Using very fine forceps, prepare the femoral artery by gently removing the surrounding connective tissue. Be careful to harm neither the femoral nerve nor the femoral vein, which is medial to the artery.</li>
	<li>Prepare about 1 cm of the femoral artery. Carefully put the tip of the forceps under the vessel to gently lift it.</li>
	<li>Thread pieces of 4-0 silk suture under the distal and proximal parts of the artery and form slings. Clamp the ends of each of the two thread slings between the branches of a surgical clamp. Use the surgical clamps to control the artery. Gently stretch and lift the slings in order to temporarily interrupt blood flow. 
	NOTE: Work fast to avoid a prolonged tourniquet which may lead to tissue damage.</li>
	<li>Using sharp micro scissors, perform an arteriotomy in the middle of the femoral artery.</li>
	<li>Introduce a guide wire through the arteriotomy. When reaching the proximal thread sling, release the tension of the thread by moving the surgical clamp and advance the guide wire further towards the abdominal aorta. 
	NOTE: Cut the guide wire using a wire cutter to facilitate handling.</li>
	<li>Place the proximal end of the guide wire between the diaphragm and the renal arteries. 
	NOTE: Advancing the guide wire too far bears the risk of aortic or cardiac injury. We recommend opening the abdomen to ensure adequate positioning of the guide wire and the stent at least for the first several animals.</li>
	<li>Introduce a crimped and balloon-mounted coronary stent measuring 2.25 mm x 8 mm (max. 2.5 mm x 8 mm) over the guide wire into the femoral artery and advance it to the abdominal aorta.</li>
	<li>Place the stent just above the aortic bifurcation but below the renal arteries. Deploy the stent by inflating the balloon catheter to 12 atm for 15 s by using an inflation syringe system.</li>
	<li>Deflate the balloon catheter and maintain negative pressure according to the manufacturer’s recommendations for the stent in use.</li>
	<li>Slowly withdraw the deflated catheter while leaving the stent in place.</li>
	<li>Just before taking out the catheter, create tension on the thread loop above the incision with the surgical clamp to interrupt blood flow again. Then remove the balloon catheter and directly ligate the vessel proximally.</li>
	<li>Tie the proximal and the distal thread loops to ligate the femoral artery and confirm adequate hemostasis of the arteriotomy. Collateral arteries will ensure further perfusion to the limb.</li>
	<li>Close the muscle overlying the artery, as well as the skin incision by using 10-0 non-resorbable sutures.</li>
</ol>
4. Animal care after stent implantation
<ol>
	<li>Immediately after the operation, allow the rat to recover for 60 min in a special intensive care unit cage with warmed air (30‒35 °C) and an oxygen supply.</li>
	<li>Watch the animals carefully until fully recovered. Afterwards, move the rats into a normal cage. Provide ad libitum access to water and food.</li>
	<li>Administer postoperative analgesia every 6-12 hours with 0.03-0.05 mg/kg buprenorphine (s.c., in 500µl NaCl) for a total of 72 hours under clinical assessment.</li>
	<li>Have the food mixed with clopidogrel (15 mg/kg) to avoid thrombosis of the implanted stent.</li>
	<li>To enhance hypercholesterolemic conditions and plaque formation, start western diet feeding at 6‒8 weeks after birth and continue until euthanasia. If desired, a cohort of animals fed normal rat chow can serve as control.</li>
</ol>
5. Tissue collection and processing
<ol>
	<li>Before starting the tissue explantation at the designated time point, euthanize the animal according to IACUC guidelines. Harvest the stented aorta for histological analysis at the end of the observation period.</li>
	<li>Open the abdomen by a midline incision and remove the stented segment of the aorta as well as adjacent non-stented parts of the aorta, measuring 0.5 cm each.</li>
	<li>Place the tissue into a solution of 4% buffered formalin for 24 h for fixation.</li>
	<li>Embed the stented arterial tissue in plastic and perform histological and immunohistochemical staining according to standard protocols18,19.</li>
</ol>
6. Histomorphometric analysis
<ol>
	<li>Perform histomorphometric analysis of sequential sections of the proximal, middle, and distal part of the stented aorta by means of a microscope linked to a computer with an appropriate image analysis software.</li>
	<li>Trace the contours of the external elastic lamina (EEL, between adventitia and media), internal elastic lamina (IEL, between media and neointima), and lumen with a graphic drawing tablet. From these values, calculate EEL area, IEL area, and lumen area with the software.</li>
	<li>Calculate the percent cross-sectional area in-stent restenosis (ISR): 
	<img alt="ISR formula: 100 × (1 - [Lumen area/IEL area]); mathematical equation; vascular study." src="/files/ftp_upload/61442/61442equ01.jpg" data-altupdatedat="1747911167000" data-alt="Equation 1"></li>
	<li>Calculate the total neointimal area (Ai): 
	<img alt="A&lt;sub&gt;i&lt;/sub&gt;=IEL/Lumen area; formula for area ratio; symbolic equation for research analysis." src="/files/ftp_upload/61442/61442equ02.jpg" data-altupdatedat="1747911167000" data-alt="Equation 2"></li>
	<li>Measure the neointimal thickness (NIT) over each stent strut as the distance between strut and lumen. Measure the NIT between the stent struts as the distance between IEL and lumen. 
	NOTE: Alternatively, calculate NIT as 
	<img alt="NIT formula: (2 × Ai)/(PL + PIEL); equation illustrating a research calculation method." src="/files/ftp_upload/61442/61442equ03.jpg" data-altupdatedat="1747911167000" data-alt="Equation 3"> 
	where PL and PIEL are the lumen and internal elastic lamina perimeter, respectively20.</li>
	<li>Perform additional analyses according to the requirements of the study.</li>
</ol>

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The authors have nothing to disclose.

This protocol describes the implantation of human-sized coronary stents into the abdominal aorta of apoE-/- rats. Several technical points are worth emphasizing. First, a mismatch between the stent size and the size of the aorta should be avoided. Placing too small a stent can lead to stent malapposition, whereas implantation of a stent that is too large for the aorta can cause overstretch, tearing, and injury of the vessel. Therefore, we recommend using stents between 2.0 and 2.5 mm in diameter, and to keep implantation pressure within the recommended range without overstretching the stent. The most suitable implantation pressure is usually given by the stent manufacturer. Excess injury of the femoral vein and subsequently the vena cava should be avoided because the vessel walls are extremely thin and very easy to injure, resulting in bleeding that is hard to stop. The femoral artery is distinguishable from the femoral vein by pulsation, which should be carefully observed. Another pitfall is the possibility of arterial injury and dissection when introducing the guidewire and / or the balloon catheter. Arterial dissection can be minimized by controlling and stretching the femoral artery distally with slings using silk ties while introducing the balloon catheter. It is imperative to immediately stop advancing the device when resistance is encountered. In this case, small movements between thumb and index finger will help change the direction of the device. In our experience, this is most frequently the case just below the inguinal ligament and further up, when the common iliac artery approaches the bifurcation, as it descends deeper into the retroperitoneal space here. There will certainly be a learning curve for the operator before survival rates are stable and with some experience, the average surgical time is about 20 min.
In humans, stents are usually implanted into severely narrowed atherosclerotic arteries. Although apoE deficiency in general renders animals more susceptible for the development of atherosclerotic lesions, we did not observe any plaque formation in our rats, most likely because western diet feeding was not started until stent implantation. If stent implantation in atherosclerotic lesions is desired, western diet should start at 6&#8210;8 weeks after birth and continue until sacrifice. Atherosclerotic lesions in susceptible strains will develop after 7&#8210;14 weeks on the high fat diet22. So far, only limited data on apoE-/- rats have been published. However, no study reported spontaneous lesion development before the age of 20 weeks23. Zhao et al. observed typical atherosclerosis in apoE-/- rats after at least 24 weeks with a continuous increase in plaque burden and lesion severity until sacrifice at 72 weeks15. Thus, according to the literature, it is improbable that rats would develop spontaneous atherosclerosis at 14&#8211;16 weeks of age. Therefore, we recommend using older rats and to start western diet as early as possible if stent implantation in pre-formed atherosclerotic lesions is desired for the study.
Six animals did not survive the surgery. Two animals died from stent thrombosis in spite of administration of clopidogrel. To reduce stent thrombosis, animals can be pre-treated for 48 h with aspirin or receive an intraperitoneal injection of enoxaparin post-operatively. Introducing clopidogrel one day before surgery might also reduce the risk of thrombosis, but any intensification of the anti-thrombotic therapy at the same time increases the risk of hemorrhage. Stent thrombosis is a common complication of PCI24,25,26 and can have several reasons. Potentially, in our study, stent thrombosis fatalities resulted from insufficient balloon inflation and concurrent stent malapposition. In contrast to stent implantation in humans, stent deployment in the rat abdominal aorta was not controlled by angiography. Therefore, ineffective balloon inflation cannot be detected and corrected during surgery. Similarly, stent deployment might lead to an unintentional occlusion of a branching vessel. Considering that it is not practicable to perform the surgery which requires the use of a surgical microscope under fluoroscopic control, we recommend opening the abdomen to confirm the precise deployment of the stent, at least for the first several procedures. Other potential causes for stent thrombosis might be inflammatory reactions, severe injury, or dissections of the vessel wall. The surgeon must be aware of any clinical signs indicating these complications, and the animals must be inspected each day throughout the observation period.
The rat abdominal aorta measures between 1.8 mm and 3.0 mm in diameter, depending on the animal&#8217;s weight27,28. Advancing a bulky stent through the even smaller femoral and iliac arteries may cause intimal tear and damage to the vessel wall. Therefore, this technique is limited to the implantation of smaller stents (between 2.0 and 2.5 mm in diameter) to avoid overstretching or injury of the vessel wall of the aorta.
Another limitation is the necessity of ligating the femoral artery in order to achieve hemostasis after the procedure, potentially bearing the risk of lower limb ischemia. However, previous studies showed that collateral arteries as well as adaptions of the microvasculature distal to the occlusion are able to maintain lower limb perfusion after ligation of the femoral artery in rats29, and none of our rats exhibited clinical signs of lower limb ischemia during the observation period. Still, the investigators should be aware of this potential risk, as limb ischemia not only represents a potential cause of post-operative death, but can also potentially induce a systemic inflammatory reaction, potentially biasing the results.
While rats in general are a cost-effective animal model, the use of genetically modified apoE-/- rats increases the cost. Another limitation is that it takes a comparatively long time until atherosclerotic plaques have developed in rats. Furthermore, there are some important hemodynamic differences between the aorta and the coronary arteries that deserve closer attention. Shear stress is higher in the aorta as compared to the coronaries, and bifurcations causing turbulent blood flow are absent. This diminishes the development of intimal hyperplasia and the extent of restenosis.
Restenosis is one of the major factors limiting the long-term success of coronary stents. A variety of animal models have been used to study the pathophysiology of restenosis, each featuring their own advantages and shortcomings. In comparison with other animal models, rats hold the advantage of a high throughput, an ease of handling and housing, reproducibility, as well as cost-effectiveness, while at the same time allowing for the implantation of human-sized coronary stents. The first protocol of abdominal aorta stenting in rats was reported by Langeveld et al.11. This model, however, requires a trans-abdominal access to introduce the stent, which is associated with a physical constriction of the aorta to achieve a temporary interruption of blood flow. The resulting manipulation and vessel injury might potentially cause inflammatory reactions, which might not only lead to complications, but also to pronounced ISR12. Later, Oyamada et al. modified the protocol by introducing the stent through the common iliac artery12. They compared the survival rate between the two different approaches (trans-aorta versus trans-iliac artery) and found a significantly higher mortality rate in animals with trans-abdominally deployed stents (57% versus 11%, p &#60; 0.05). Rats most commonly died from thrombosis at the incision / suture site, which is catastrophic when occurring in the abdominal aorta12. Further reducing trauma and mimicking the implantation technique in humans more closely, we used a trans-femoral access to introduce the stent and reported a mortality rate of 14%. Two rats each died from vessel closure failure, internal hemorrhage, and stent thrombosis. More recent studies, however, reported mortality rates as low as 6% after stent implantation in the rat abdominal aorta even with the trans-aortic access30,31. Still, the combined morbidity and mortality rate was 13.4%, in a study by Nevzati et al. after implantation of magnesium stents into the rat aorta30. While neither vessel closure failure nor internal hemorrhage were reported in their series, stent thrombosis was evident in 10.5% of rats30. On the other hand, Aquarius et al. did not report any stent thrombosis after treating sidewall aneurysms with flow diverters, however, this study used thinner stent strut devices, and dual antiplatelet therapy was administered to the rats31. We tried to strike a balance between the risk of stent thrombosis and bleeding risk and administered clopidogrel and heparin in our study. While this might have reduced the risk of stent thrombosis, which occurred in 4.76% of rats, it also might have been the reason for the comparatively higher risk of bleeding (9.52% of rats), either because of internal hemorrhage or vessel closure failure.
Here, we demonstrated the implantation of a drug-eluting stent into the rat abdominal aorta, but likewise this method can be used for the evaluation of other, similarly sized stent devices, for example bare metal stents or bioresorbable vascular scaffolds.
In summary, abdominal aorta stenting of apolipoprotein E-deficient rats is a reliable and reproducible model to investigate ISR after stent implantation. The model can be extended to the use of older rats, which are more likely to develop atherosclerotic lesions spontaneously, and by testing other devices used for human coronary intervention.

Percutaneous coronary intervention (PCI), combined with the deployment of a coronary stent, represents the gold standard in interventional treatment of coronary artery disease1. The long-term success of stents, however, can be limited by the occurrence of in-stent restenosis (ISR) that is determined by an excessive proliferation of neointimal tissue within the stent2,3. ISR may require a re-intervention either with coronary artery bypass or re-PCI. A variety of animal models have been suggested for the study of ISR, each of them featuring advantages and shortcomings. The major drawbacks of the most commonly used porcine coronary and rabbit iliac artery models, albeit developing lesions markedly similar to humans after stent implantation4,5, are large animal and housing costs which brings up logistical difficulties especially in long-term studies, as well as limitations in handling and equipment. Furthermore, availability of antibodies to cellular proteins of swine and rabbits is limited. On the other hand, murine models provide the major advantages of high throughput and reproducibility, as well as ease of handling, housing, and therefore cost-effectiveness. Furthermore, a higher number of antibodies are available. However, while apolipoprotein E-deficient (apoE-/-) mice have been broadly used for the study of atherosclerosis6,7,8, they are unsuitable for the study of ISR as stents have to be miniaturized to be implanted into mice, potentially changing the stents&#8217; mechanical properties. Moreover, the aortic wall of mice measures between 50 &#181;m in young mice and 85 &#181;m in old mice9, and stents have to be deployed using pressure levels as low as 2 atm, which might lead to malapposition of the stent10. Rats, however, allow for the implantation of commercially available human coronary stents, and demonstrate a vascular healing course similar to larger animals after aortic stent implantation, first reported by Langeveld et al.11. This technique originally required a trans-abdominal access, which necessitated a physical constriction of the aorta to achieve a temporary interruption of blood flow. To avoid the potentially associated vessel injury and inflammatory reactions, the technique was later refined by the introduction of a trans-iliac access, which additionally resulted in a higher survival rate of the animals12.
Because wildtype rats do not develop atherosclerotic lesions13, apoE-/- rats have been generated using nuclease techniques such as Transcription Activator-Like Effector Nuclease (TALEN)14, Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR/Cas9)15, and Zinc Finger (ZF)16. ApoE-/- rats have been commercially available since 2011. Providing an atherogenic background, apoE-/- rats allow for a more realistic evaluation of human-sized coronary stents, especially with regards to ISR.
Herein, we describe the method via the transfemoral access route and using a commercially available thin-strut cobalt-chromium drug-eluting stent (DES), however, it can also be applied for the study of other stent types, such as bare metal stents (BMS) or biodegradable stents.

<table><tbody><tr><td>&lt;strong&gt;Diet&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>SNIFF High Fat diet + Clopidogrel (15 mg/kg)</td><td>SNIFF Spezialdi&amp;auml;ten GmbH, Soest</td><td>custom prepared</td><td>Western Diet</td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Drugs and Anesthetics&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Buprenorphine</td><td>Essex Pharma</td><td>997.00.00</td><td></td></tr><tr><td>ISOFLO (Isoflurane Vapor) vaporiser</td><td>Eickemeyer</td><td>4802885</td><td></td></tr><tr><td>Isoflurane</td><td>Forene Abbott</td><td>B 506</td><td></td></tr><tr><td>Isotonic (0.9%) NaCl solution</td><td>DeltaSelect GmbH</td><td>PZN 00765145</td><td></td></tr><tr><td>Ringer&#039;s lactate solution</td><td>Baxter Deutschland GmbH</td><td>3775380</td><td></td></tr><tr><td>(S)-ketamine</td><td>CEVA Germany</td><td></td><td></td></tr><tr><td>Xylazine</td><td>Medistar Germany</td><td></td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Consumable supplies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>10 mL syringes</td><td>BD Plastipak</td><td>4606108V</td><td></td></tr><tr><td>2 mL syringes</td><td>BD Plastipak</td><td>4606027V</td><td></td></tr><tr><td>6-0 prolene suture</td><td>ETHICON</td><td>N-2719K</td><td></td></tr><tr><td>4-0 silk suture</td><td>Seraflex</td><td>IC 158000</td><td></td></tr><tr><td>Bepanthen Eye and Nose Ointment</td><td>Bayer Vital GmbH</td><td>6029009.00.00</td><td></td></tr><tr><td>Cotton Gauze swabs</td><td>Fuhrmann GmbH</td><td>32014</td><td></td></tr><tr><td>Durapore silk tape</td><td>3M</td><td>1538-1</td><td></td></tr><tr><td>Poly-Alcohol Skin Desinfection Solution</td><td>Antiseptica GmbH</td><td>72PAH200</td><td></td></tr><tr><td>Sterican needle 18 G</td><td>B. Braun</td><td>304622</td><td></td></tr><tr><td>Sterican needle 27 3/4 G</td><td>B.Braun</td><td>4657705</td><td></td></tr><tr><td>Tissue Paper</td><td>commercially available</td><td></td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Surgical instruments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Graefe forceps curved x1</td><td>Fine Science Tools Inc.</td><td>11151-10</td><td></td></tr><tr><td>Graefe forceps straight</td><td>Fine Science Tools Inc.</td><td>11050-10</td><td></td></tr><tr><td>Needle holder Mathieu</td><td>Fine Science Tools Inc.</td><td>12010-14</td><td></td></tr><tr><td>Scissors</td><td>Fine Science Tools Inc.</td><td>14074-11</td><td></td></tr><tr><td>Semken forceps</td><td>Fine Science Tools Inc.</td><td>11008-13</td><td></td></tr><tr><td>Small surgical scissors curved</td><td>Fine Science Tools Inc.</td><td>14029-10</td><td></td></tr><tr><td>Small surgical scissors straight</td><td>Fine Science Tools Inc.</td><td>14028-10</td><td></td></tr><tr><td>Standard pattern forceps</td><td>Fine Science Tools Inc.</td><td>11000-12</td><td></td></tr><tr><td>Vannas spring scissors</td><td>Fine Science Tools Inc.</td><td>15000-08</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Dissecting microscope</td><td>Leica MZ9</td><td></td><td></td></tr><tr><td>Temperature controlled heating pad</td><td>Sygonix</td><td>26857617</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Equipment for stent implantation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Drug-eluting stent Xience 2,25mm x 8mm</td><td>Abbott Vascular USA</td><td>1009544-18</td><td></td></tr><tr><td>Guide wire Fielder XT PTCA guide wire: 0.014&quot; x 300cm</td><td>ASAHI INTECC CO., LTD Japan</td><td>AGP140302</td><td></td></tr><tr><td>Inflation syringe system</td><td>Abbott 20/30 Priority Pack</td><td>1000186</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Tissue processing and analysis&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>30% H&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;2&lt;/sub&gt;</td><td>Roth</td><td>9681</td><td>Histology</td></tr><tr><td>Ethanol</td><td>Roth</td><td>K928.1</td><td>Histology</td></tr><tr><td>Giemsas Azur-Eosin-Methylenblau</td><td>Merck</td><td>109204</td><td>Histology</td></tr><tr><td>Graphic Drawing Tablet</td><td>WACOM Europe GmbH</td><td>CTL-6100WLK-S</td><td></td></tr><tr><td>Roti Histofix, Formaldehyd 4% buffered</td><td>Roth</td><td>P087</td><td>Histology</td></tr><tr><td>Technovit 9100</td><td>Morphisto</td><td>12225.K1000</td><td>Histology</td></tr></tbody></table>

implantation of human-sized coronary stents into rat abdominal aorta using a trans-femoral access

Percutaneous coronary intervention (PCI), combined with the deployment of a coronary stent, represents the gold standard in interventional treatment of coronary artery disease. In-stent restenosis (ISR) is determined by an excessive proliferation of neointimal tissue within the stent and limits the long-term success of stents. A variety of animal models have been used to elucidate pathophysiological processes underlying in-stent restenosis (ISR), with the porcine coronary and the rabbit iliac artery models being the most frequently used. Murine models provide the advantages of high throughput, ease of handling and housing, reproducibility, and a broad availability of molecular markers. The apolipoprotein E deficient (apoE-/- ) mouse model has been widely used to study cardiovascular diseases. However, stents must be miniaturized to be implanted into mice, involving important changes of their mechanical and (potentially) biological properties. The use of apoE-/- rats can overcome these shortcomings as apoE-/- rats allow for the evaluation of human-sized coronary stents while at the same time providing an atherogenic phenotype. This makes them an excellent and reliable model to investigate ISR after stent implantation. Here, we describe, in detail, the implantation of commercially available human coronary stents into the abdominal aorta of rats with an apoE-/- background using a trans-femoral access.

This protocol describes stent implantation in the abdominal aorta of rats using a trans-femoral access route (Figure 1). The first central point of this animal model is that it allows for the deployment of human-sized coronary stents. A commercially available crimped and balloon-mounted coronary stent can be placed into the abdominal aorta of rats. Thus, in addition, the same principle of stent deployment as in humans can be applied. Another advantage of the use of rats is the availability of genetically modified strains, such as apoE-/- rats, which are commercially available.
We recently employed this method to evaluate whether apolipoprotein E-deficient rats are more prone to develop ISR as compared to wildtype rats21. From a total of 42 male rats undergoing stent implantation, 36 rats completed the study protocol after 28 days (survival rate = 85.71%). Two rats each died from vessel closure failure, internal hemorrhage, and stent thrombosis. Stents from three animals could not be analyzed because the tissue was seriously damaged or disrupted due to processing failures. Most likely, this happened during the sawing procedure. We recommend training to perform this technique several times before the start of the study.
In the remaining 33 rats, human-sized coronary stents were successfully deployed with no sign of malapposition or vessel injury (Table 1). Body weight was similar in wildtype apoE+/+ and apoE-/- rats (530.1 ± 15.94 g versus 513.6 ± 16.45 g). Homozygous apoE-/- rats developed markedly elevated neointimal hyperplasia and ISR as compared to wildtype apoE+/+ rats (Figure 2). Although an apoE-/- background renders animals more susceptible for atherosclerosis, especially when fed western diet, we did not observe any antecedent atherosclerotic plaques in our rats, most likely because a western diet was not started until surgery and the subsequent observation period of four weeks was too short for atherosclerotic lesion development.
<img alt="Femoral artery stent placement diagram; surgical clamp, thread sling; coronary stent balloon setup." class="xfigimg" src="/files/ftp_upload/61442/61442fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Schema of the stent implantation into the abdominal aorta of rats using a trans-femoral access. 
(a) After interruption of the blood flow, a guide wire is introduced through a medial arteriotomy. (b) A crimped and balloon-mounted coronary stent is introduced over the guide wire into the femoral artery. (c) The balloon-mounted stent is advanced to the abdominal aorta, where it is deployed by balloon inflation. The stent should be placed above the bifurcation and below the renal arteries. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61442/61442fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Microscope image of plant stem cross-section, labeled cells, anatomical study, high magnification." class="xfigimg" src="/files/ftp_upload/61442/61442fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Representative photomicrographs of Giemsa-stained abdominal aorta at 28 days after stent implantation in western-diet-fed. 
(a) Wildtype apoE+/+ rats and (b) Homozygous apoE-/- rats. High power images: NI = neointima, St = stent strut, M = tunica media, L = lumen. Figure has been reproduced with modifications from Cornelissen, A. et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61442/61442fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>number of rats</td>
		</tr>
		<tr>
			<td>vessel closure failure</td>
			<td>2</td>
		</tr>
		<tr>
			<td>internal hemorrhage</td>
			<td>2</td>
		</tr>
		<tr>
			<td>stent thrombosis</td>
			<td>2</td>
		</tr>
		<tr>
			<td>tissue processing failure</td>
			<td>3</td>
		</tr>
		<tr>
			<td>successful completion of the protocol</td>
			<td>33</td>
		</tr>
	</tbody>
</table>
Table 1: Outcome of stent implantation in rat abdominal aorta using a trans-femoral access.

Watch this Scientific Journal Video about Implantation of Human-Sized Coronary Stents into Rat Abdominal Aorta Using a Trans-Femoral Access at JoVE.com

Implantation of Human-Sized Coronary Stents into Rat Abdominal Aorta Using a Trans-Femoral Access

This protocol describes the implantation of human coronary stents into the abdominal aorta of rats with an apoE-/- background using a trans-femoral access. Compared with other animal models, murine models carry the advantages of high throughput, reproducibility, ease of handling and housing, and a broad availability of molecular markers.

Percutaneous coronary intervention (PCI), combined with the deployment of a coronary stent, represents the gold standard in interventional treatment of ...

implantation-human-sized-coronary-stents-into-rat-abdominal-aorta

Nanyang Technological University

Research

JoVE Journal

Medicine

4.2K Views.  University Hospital RWTH Aachen. Dit protocol beschrijft de implantatie van menselijke coronaire stents in de abdominale aorta van ratten met een apoE-/- achtergrond met behulp van een trans-femurtoegang. In vergelijking met andere diermodellen hebben murinemodellen de voordelen van een hoge doorvoer, reproduceerbaarheid, gebruiksgemak en huisvesting en een brede beschikbaarheid van moleculaire markers.

Video: Implantation of Human-Sized Coronary Stents into Rat Abdominal Aorta Using a Trans-Femoral Access

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

Human Internal Mammary Artery (IMA) Transplantation and Stenting: A Human Model to Study the Development of In-Stent Restenosis

Preclinical in vivo research models to investigate pathobiological and pathophysiological processes in the development of intimal hyperplasia after vessel stenting are crucial for translational approaches1,2.
The commonly used animal models include mice, rats, rabbits, and pigs3-5. However, the translation of these models into clinical settings remains difficult, since those biological processes are already studied in animal vessels but never performed before in human research models6,7. In this video we demonstrate a new humanized model to overcome this translational gap. The shown procedure is reproducible, easy, and fast to perform and is suitable to study the development of intimal hyperplasia and the applicability of diverse stents.
This video shows how to perform the stent technique in human vessels followed by transplantation into immunodeficient rats, and identifies the origin of proliferating cells as human.

Human Internal Mammary Artery IMA Transplantation and Stenting: A Human Model to Study the Development of In-Stent Restenosis

This video shows a model to study the development of intimal hyperplasia after stent deployment using a human vessel (IMA) in an immunodeficient rat model.

Preclinical in vivo research models to investigate pathobiological and pathophysiological processes in the development of intimal hyperplasia after vessel ...

Implantation of Engineered Tissue in the Rat Heart

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in common laboratory rats or mice and these quite frequently serve as an intermediate step between bench-top experiments and large animal testing or human trials. Given that rodents provide an established, cost-effective, and physiologically-relevant model system in which to test novel combinations of scaffolding materials and cells, they are particularly well-suited for cardiovascular tissue engineering studies. Presently, we describe an open-heart surgical procedure to implant engineered tissue containing myogenic progenitor cells in the atrioventricular (AV) groove of a rat heart. These implants are intended to create an electrical conduit between the right atrium and right ventricle with the ultimate goal of providing an alternative treatment to conventional pacemaker implantation in pediatric patients with complete heart block[1]. The engineered tissue is implanted in the AV-groove by means of a thoracotomy. For our purposes, Lewis rats are anesthetized and invasively ventilated to maintain positive airway pressure during the sterile surgical procedure. The approach to the heart is performed by a right thoracotomy through an antero-lateral incision at the 5th intercostal space. The tissue construct is fixed in the AV groove using a single 7-0 Prolene suture and positioned between the right ventricle and atrium at the ventral portion of the heart. The epicardium is partially removed to allow direct contact between the recipient myocardial cells and those contained in the engineered tissue. Following implantation, the chest wall is closed in layers, any pneumothorax is evacuated, and the animal is extubated and treated with analgesic.

Here, we describe a cardiac surgical procedure to implant engineered tissue in the atrioventricular (AV)-groove of an adult Lewis rat.

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in ...

Biology

A Murine Model of Stent Implantation in the Carotid Artery for the Study of Restenosis

Despite the considerable progress made in the stent development in the last decades, cardiovascular diseases remain the main cause of death in western countries. Beside the benefits offered by the development of different drug-eluting stents, the coronary revascularization bears also the life-threatening risks of in-stent thrombosis and restenosis. Research on new therapeutic strategies is impaired by the lack of appropriate methods to study stent implantation and restenosis processes. Here, we describe a rapid and accessible procedure of stent implantation in mouse carotid artery, which offers the possibility to study in a convenient way the molecular mechanisms of vessel remodeling and the effects of different drug coatings.

A model of stent implantation in mouse carotid artery is described. Compared to other similar methods, this procedure is very rapid, simple and accessible, offering the possibility to study in a convenient way the vascular wall reaction to different drug-eluting stents and the molecular mechanisms of restenosis.

Despite  the considerable progress made in the stent development in the last decades,  cardiovascular diseases remain the main cause of death in western ...

A New Murine Model of Endovascular Aortic Aneurysm Repair

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of Abdominal aortic aneurysm (AAA) pathophysiology. There is no abdominal aortic aneurysm endograft exclusion model cheap and reproducible, which would allow deep investigations of AAA before and after treatment. We hereby describe how to induce, and then to exclude with a covered coronary stentgraft an abdominal aortic aneurysm in a rat. The well known elastase induced AAA model was first reported in 19901 in a rat, then described in mice2. Elastin degradation leads to dilation of the aorta with inflammatory infiltration of the abdominal wall and intra luminal thrombus, matching with human AAA. Endovascular exclusion with small covered stentgraft is then performed, excluding any interactions between circulating blood and the aneurysm thrombus. Appropriate exclusion and stentgraft patency is confirmed before euthanasia by an angiography thought the left carotid artery. Partial control of elastase diffusion makes aneurysm shape different for each animal. It is difficult to create an aneurysm, which will allow an appropriate length of aorta below the aneurysm for an easy stentgraft introduction, and with adequate proximal and distal neck to prevent endoleaks. Lots of failure can result to stentgraft introduction which sometimes lead to aorta tear with pain and troubles to stitch it, and endothelial damage with post op aorta thrombosis. Giving aspirin to rats before stentgraft implantation decreases failure rate without major hemorrhage. Clamping time activates neutrophils, endothelium and platelets, and may interfere with biological analysis.

Histological and biochemical modifications after aneurysm endograft exclusion are still unclear. We describe a new model of endograft implantation on a murine aneurysm. Through thrombus analysis with and without persistant circulating blood stream interface, and new endovascular biomaterials evaluation, this model will help to better understand endovascular aneurysm exclusion pathobiology.

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of ...

Imaging In-Stent Restenosis: An Inexpensive, Reliable, and Rapid Preclinical Model

Preclinical models of restenosis are essential to unravel the pathophysiological processes that lead to in-stent restenosis and to optimize existing and future drug-eluting stents.
A variety of antibodies and transgenic and knockout strains are available in rats. Consequently, a model for in-stent restenosis in the rat would be convenient for pathobiological and pathophysiological studies. 
In this video, we present the full procedure and pit-falls of a rat stent model suitable for high throughput stent research. We will show the surgical procedure of stent deployment, and the assessment of in-stent restenosis using the most elegant technique of OCT (Optical Coherence Tomography). This technique provides high accuracy in assessing plaque CSAs (cross section areas) and correlates well with histological sections, which require special and time consuming embedding and sectioning techniques. OCT imaging further allows longitudinal monitoring of the development of in-stent restenosis within the same animal compared to one-time snapshots using histology.

This video demonstrates how to use a preclinical inexpensive and reliable model to study pathobiological and pathophysiological processes of in-stent restenosis development. Longitudinal in vivo monitoring using OCT (Optical Coherence Tomography) and analysis of OCT images are also demonstrated.

Preclinical models of restenosis are essential to unravel the pathophysiological processes that lead to in-stent restenosis and to optimize existing and ...

Biodegradable Magnesium Stent Treatment of Saccular Aneurysms in a Rat Model - Introduction of the Surgical Technique

The steady progess in the armamentarium of techniques available for endovascular treatment of intracranial aneurysms requires affordable and reproducable experimental animal models to test novel embolization materials such as stents and flow diverters. The aim of the present project was to design a safe, fast, and standardized surgical technique for stent assisted embolization of saccular aneurysms in a rat animal model.
Saccular aneurysms were created from an arterial graft from the descending aorta.The aneurysms were microsurgically transplanted through end-to-side anastomosis to the infrarenal abdominal aorta of a syngenic male Wistar rat weighing &#62;500 g. Following aneurysm anastomosis, aneurysm embolization was performed using balloon expandable magnesium stents (2.5 mm x 6 mm). The stent system was retrograde introduced from the lower abdominal aorta using a modified Seldinger technique.
Following a pilot series of 6 animals, a total of 67 rats were operated according to established standard operating procedures. Mean surgery time, mean anastomosis time, and mean suturing time of the artery puncture site were 167 &#177; 22 min, 26 &#177; 6 min and 11 &#177; 5 min, respectively. The mortality rate was 6% (n=4). The morbidity rate was 7.5% (n=5), and in-stent thrombosis was found in 4 cases (n=2 early, n=2 late in stent thrombosis).
The results demonstrate the feasibility of standardized stent occlusion of saccular sidewall aneurysms in rats - with low rates of morbidity and mortality. This stent embolization procedure combines the opportunity to study novel concepts of stent or flow diverter based devices as well as the molecular aspects of healing.

Reproducable experimental animal models are needed for the testing of novel embolization materials, which have been designed to treat endovascular occlusion of intracranial aneurysms (IA). The present study aims to develop a safe and standardized surgical technique for stent assisted embolization of saccular aneurysms in a rat animal model.

The steady progess in the armamentarium of techniques available for endovascular treatment of intracranial aneurysms requires affordable and reproducable ...

Neuroscience

Novel Percutaneous Approach for Deployment of 3D Printed Coronary Stenosis Implants in Swine Models of Ischemic Heart Disease

Minimally invasive methods for creating models of focal coronary narrowing in large animals are challenging. Rapid prototyping using three-dimensionally (3D) printed coronary implants can be employed to percutaneously create a focal coronary stenosis. However, reliable delivery of the implants can be difficult without the use of ancillary equipment. We describe the use of a mother-and-child coronary guide catheter for stabilization of the implant and for effective delivery of the 3D printed implant to any desired location along the length of the coronary vessel. The focal coronary narrowing was confirmed under coronary cineangiography and the functional significance of the coronary stenosis was assessed using gadolinium-enhanced first-pass cardiac perfusion MRI. We showed that reliable delivery of 3D printed coronary implants in swine models (n = 11) of ischemic heart disease can be achieved through repurposing mother-and-child coronary guide catheters. Our technique simplifies the percutaneous delivery of coronary implants to create closed-chest swine models of focal coronary artery stenosis and can be performed expeditiously, with a low procedural failure rate.

We describe a novel, cost-effective, and efficient technique for percutaneous delivery of three-dimensionally printed coronary implants to create closed-chest swine models of ischemic heart disease. The implants were fixed in place using a mother-and-child extension catheter with high success rate.

Minimally invasive methods for creating models of focal coronary narrowing in large animals are challenging. Rapid prototyping using three-dimensionally ...

Using a Cell-Tracer Injection to Investigate the Origin of Neointima-Forming Cells in a Rat Saccular Side Wall Model

Microsurgical clipping creates a subsequent barrier of blood flow into intracranial aneurysms, whereas endovascular treatment relies on neointima and thrombus formation. The source of endothelial cells covering the endoluminal layer of the neointima remains unclear. Therefore, the aim of the present study was to investigate the origin of neointima-forming cells after cell-tracer injection in the already well-established Helsinki rat microsurgical sidewall aneurysm model.
Sidewall aneurysms were created by suturing decellularized or vital arterial pouches end-to-side to the aorta in male Lewis rats. Before arteriotomy with aneurysm suture, a cell-tracer injection containing CM-Dil dye was performed into the clamped aorta to label endothelial cells in the adjacent vessel and track their proliferation during follow-up (FU). Treatment followed by coiling (n = 16) or stenting (n = 15). At FU (7 days or 21 days), all rats underwent fluorescence angiography, followed by aneurysm harvesting and macroscopic and histological evaluation with immunohistological cell counts for specific regions of interest. 
None of the 31 aneurysms had ruptured upon follow-up. Four animals died prematurely. Macroscopically residual perfusion was observed in 75.0% coiled and 7.0% of stented rats. The amount of cell-tracer-positive cells was significantly elevated in decellularized stented compared to coiled aneurysms with respect to thrombus on day 7 (p = 0.01) and neointima on day 21 (p = 0.04). No significant differences were found in thrombus or neointima in vital aneurysms. 
These findings confirm worse healing patterns in coiled compared to stented aneurysms. Neointima formation seems particularly dependent on the parent artery in decellularized aneurysms, whereas it is supported by the recruitment from aneurysm wall cells in vital cell-rich walls. In terms of translation, stent treatment might be more appropriate for highly degenerated aneurysms, whereas coiling alone might be adequate for aneurysms with mostly healthy vessel walls.

We performed a one-point, lipophilic cell-tracer injection to track endothelial cells, followed by an arteriotomy and suturing of sidewall aneurysms on the abdominal rat aorta. Neointima formation seemed dependent on the parent artery in decellularized aneurysms and was promoted by the recruitment from aneurysm wall cells in vital cell-rich walls.

Microsurgical clipping creates a subsequent barrier of blood flow into intracranial aneurysms, whereas endovascular treatment relies on neointima and ...

Implantation of Electrospun Vascular Grafts with Optimized Structure in a Rat Model

Here, we present a protocol to fabricate macroporous PCL vascular graft and describe an evaluation protocol by using a rat model of abdominal aorta replacement. The electrospun vascular grafts often possess relatively small pores, which limit cell infiltration into the grafts and hinder the regeneration and remodeling of the neo-arteries. In this study, PCL vascular grafts with thicker fibers (5 - 6 &#181;m) and larger pores (~30 &#181;m) were fabricated by using a modified processing technique. The long-term performance of the graft was evaluated by implantation in a rat abdominal aorta model. Ultrasound analysis showed that the grafts remained patent without aneurysm or stenosis occurring even after 12 months of implantation. Macroporous structure improved the cell ingrowth and thus promoted tissue regenerated at 3 months. More importantly, there was no sign of adverse remodeling, such as calcification within the graft wall after 12 months. Therefore, electrospun PCL vascular grafts with modified macroporous processing hold potential to be an artery substitute for long-term implantation.

Here, we present a modified electrospinning method to fabricate PCL vascular grafts with thick fibers and large pores, and describe a protocol to evaluate the in vivo performance in a rat model of abdominal aorta replacement.

Here, we present a protocol to fabricate macroporous PCL vascular graft and describe an evaluation protocol by using a rat model of abdominal aorta ...

Implantation of Human-Sized Coronary Stents into Rat Abdominal Aorta Using a Trans-Femoral Access

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