The authors thank Olgica Beslac and Kay Nettelbeck for their excellent support and technical assistance during the peri-operative phase and Alessandra Bergadano, DVM, PhD, for the dedicated supervision of the long-term animal health.

All veterinary care was performed in accordance with the institutional guidelines (all experiments were approved by the Local Committee for Animal Care of the Canton Bern, Switzerland (BE 108/16)) and conducted under supervision of a board-certified veterinarian anesthesiologist. The ARRIVE guidelines and the 3R principles were strictly followed17,18.
NOTE: House all animals at a room temperature of 22&#8210;24 Celsius (&#176;C)&#160;and maintain a 12 hours (h) light/dark cycle. Provide free access to water, pellet and ad libitum hay diet every time. Statistical analyses were performed using the non-parametric Wilcoxon-Mann-Whitney-U test. A probability value (p) of &#8804; 0.05 was considered significant.
1. Presurgical phase
<ol>
	<li>Perform a detailed preoperative clinical examination of all rabbits planned for surgery immediately next to a quiet, aseptic operating room maintaining a temperature of 23 &#177; 3 &#176;C.
	<ol>
		<li>Record the weight of each animal, macroscopically evaluate the mucous membranes, capillary refill time and pulse quality.</li>
		<li>Further on perform cardiac auscultation with a stethoscope and abdominal palpation.</li>
		<li>Based on the clinical findings, attribute an American Society of Anesthesiologists (ASA) classification to each rabbit19. Include only animals with an ASA I score in the study.</li>
		<li>Shave both outer ears with an electric shaver and apply prilocaine-lidocaine cream on both auricular arteries and veins.</li>
	</ol>
	</li>
	<li>Sedate the rabbit with a combination of 20 milligram (mg)/killogram (kg) of ketamine, 100 mg/kg of dexmedetomidine and 0.3 mg/kg of methadone injected subcutaneously (SC) via a syringe.</li>
	<li>Leave each animal undisturbed for at least 15 min.</li>
	<li>Thereafter, under supplementary oxygenation with 3 liter (l) /minute (min) through a loose face mask and steady monitoring through a pulse oximeter, place a 22 G cannula in the left auricular central artery and another 22 G cannula in the auricular vein of the contralateral ear.</li>
	<li>Shave the surgical field (neck) and inject 0.75% peri-incisional ropivacaine intradermally. Next shave the forehead and prepare to place pediatric electroencephalographic (EEG) sensors.</li>
	<li>Induce general anesthesia with propofol 1-2 mg/kg intravenously (IV) to effect. Then immediately intubate the trachea of all rabbits with a silicone tube (3 millimeter (mm) internal diameter) under capnographic control. Afterwards, transport all rabbits to the operating room, place them in dorsal recumbency and connect the tube to a pediatric circle system.</li>
	<li>Achieve anesthesia deepening and maintenance through isoflurane in oxygen, targeting a maximal end tidal isoflurane concentration of 1.3%.</li>
	<li>Ensure clinical and instrumental monitoring (pulse oximetry, doppler and invasive blood pressure, 3-lead electrocardiogram, EEG, rectal temperature monitoring and inhaled and exhaled gases) until tracheal extubation.</li>
	<li>To maintain hydration, provide Ringer&#8217;s lactate at a continuous rate infusion (CRI) of 5 ml/kg/h through the venous access. Always confirm proper anesthesia using toe pinches at an interval of 10 min.</li>
	<li>Disinfect the surgical field using povidone iodine from the manubrium sterni to both jaw angles. Now, perform sterile draping of the surgical field.</li>
	<li>During surgery, provide analgesia with lidocaine at a CRI of 50 microgram (&#181;g)/kg/min and fentanyl at 3&#8210;10 &#181;g/kg/h. Apply spontaneous or assisted ventilation as well as permissive hypercapnia. Perform arterial blood gas analysis at least one time during surgery.</li>
	<li>Treat relevant hypotension (mean arterial pressure &#60; 60 mmHg) with noradrenaline. Prevent hypothermia (rectal temperature &#8804; 38 &#176;C) using a heating pad or a heating forced-air warming system.</li>
</ol>
2. Surgical phase &#8211; Step I
<ol>
	<li>Start the surgery with a median skin incision from the manubrium sterni to the level of the jaw angles/larynx. Sharply dissect the skin and soft tissue with a scalpel, surgical scissors and forceps. Separate the subcutis and the fat pad medially by blunt dissection.</li>
	<li>Enter the anterior upper ridge of the sternocleidomastoid muscle medially on the left side by blunt dissection, using micro forceps and surgical scissors.</li>
	<li>Macroscopically, perform blunt preparation and carefully separate the left CCA from the vagal nerve distally to avoid laryngeal paresis by further using micro forceps and surgical scissors (Figure 2). Note that the bifurcation of the left CCA serves as intraoperative landmark (Figure 3 and Figure 4A). For all the following steps, use a soft tissue spreader to improve surgical visualization.</li>
	<li>After successful preparation and liberation of the left distal CCA from the vagal nerve, administer papaverine (40 mg/ml, 1:1 diluted in 0.9% isotonic sodium chloride solution) locally. Continuously protect all vessel segments with micro swabs followed by further papaverine administration externally. Place the papaverine-soaked left CCA below the autologous muscle tissue to protect the vessel from drying under the light of the operation microscope.</li>
	<li>Switch sides while maximizing the surgeon&#8217;s comfort during the operative procedure. Repeat the same surgical procedure on the right side. Dissect the CCA distally and proximally up to the predefined landmarks (carotid bifurcation at the level of the jaw angles/larynx and internal jugular vein; Figure 4A,B). Reinsert a spreader and administer micro swabs and papaverine as described previously.</li>
	<li>Before the ligation of the right proximal CCA, inject heparin (500 international units (IU)/kg) systemically via a venous ear catheter.</li>
	<li>Use a surgical microscope from now on. First, ligate the right proximal CCA with a 4-0 non-absorbable suture directly at the end of the macroscopically visible proximal landmark to avoid any tension on the arterial vessel.
	<ol>
		<li>Secondly, apply a 6-0 non-absorbable ligature exactly 4&#8210;5 mm distally by using a vessel clip for measurement, considering that after cutting distally from the first 4-0 ligature, the resulting arterial pouch&#160;will be of standardized length of about 3&#8210;4 mm in every animal (Figure 5A,C).</li>
	</ol>
	</li>
	<li>After tightening the 6-0 ligature, clamp the right CCA as far distally as possible with a temporary vessel clip (as normally used in cerebral aneurysm surgery) to avoid any endothelial damage and to create a long vessel segment for irrigation in order to prevent thrombogenesis (Figure 5B).</li>
	<li>Now perform a cut distally to the 4-0 non-absorbable ligature. To harvest the arterial pouch (Figure 5C), perform a second cut distally to the 6-0 non-absorbable ligature.</li>
	<li>Clean the arterial pouch meticulously from all soft tissue and measure its length, width and depth (Figure 5C) with a vessel clip. If no further modification is needed, keep the autologous arterial graft in a heparinized solution (500 IU/100 ml&#160;in 0.9% isotonic sodium chloride) at room temperature until further use.</li>
</ol>
3. Arterial pouch degradation
<ol>
	<li>If an arterial pouch degradation is needed, clean it meticulously of soft tissue and preincubate it with 100 IU of porcine elastase dissolved in 5 ml&#160;of Tris-buffer at room temperature on the day of experiment for 20 min. Do not use a brush technique. Incubate the arterial pouch intra- and extra-luminally by using a shaker.</li>
	<li>Before putting the pouch in a heparinized solution of 0.9% isotonic sodium chloride, gently swipe it three times for 3 min with anatomical forceps in 0.9% isotonic sodium chloride solution to wash out the remaining porcine elastase.</li>
	<li>If needed, keep the lumen of the arterial pouch opened up with a microtube made of silicone; meticulously protect the left and right CCA during the whole surgical procedure with wet micro paddings.</li>
</ol>
4. Surgical phase &#8211; Step II
<ol>
	<li>For further preparation of the CCA, place two round micro swabs directly beneath it to move the artery more superficially. Now, put one micro swab with a purple padding under the left CCA at the distal third for better visualization of the artery.</li>
	<li>Flush the right proximal CCA with a solution of 0.9% isotonic sodium chloride combined with 500 IU of heparin dissolved in 100 ml&#160;of 0.9% isotonic sodium chloride. In order to create a tension free anastomosis, place the right CCA under the fat pad/peritracheal musculature by using surgical scissors for tunneling it to the left side. Remove the soft tissue of the artery.
	<ol>
		<li>Now perform a 2 mm fish mouth incision on the proximal side of the right CCA using a micro scissor and forceps.</li>
	</ol>
	</li>
	<li>Change the side on the operating table. Clip the left distal CCA with another temporary vessel clip followed by the proximal left CCA with two temporary vessel clips. Protect all exposed vessel segments from drying out under the surgical light&#160;using wet micro swabs.</li>
	<li>Liberate the distal third of the left CCA completely from soft tissue and perform an arteriotomy. Use surgical micro forceps and gently grab some soft tissue. Now elevate the artery and incise the left distal CCA slowly with a surgical micro scissor. Flush the vessel segments with heparin (500 IU dissolved in 100 ml of 0.9% isotonic sodium chloride solution).</li>
	<li>After performing the arteriotomy with curved micro forceps and micro scissors, enlarge the arteriotomy located at the distal third of the left CCA distally, measuring about 2-fold of the diameter of the right blunt of the carotid artery and the autologous graft. This allows sufficient blood flow into the arterial pouch.</li>
	<li>Take the arterial pouch out of the heparinized saline solution. Place the pouch in the surgical field, where the bifurcation is planned. Start suturing the rear of the right carotid blunt caudally-located with a non-absorbable 9-0 suture, followed by a suture on the cranially-located rear side at the level of the fish mouth incision. Finish sewing the rear from distal to proximal by single stitches.</li>
	<li>While suturing, keep all elastase preincubated pouches moist with continuous irrigation. While suturing the vessel wall of the pouch, use curved surgical micro forceps to gently open up the lumen with its tip. Whenever suturing parts of the left or proximal right CCA, use straight surgical micro forceps. Afterwards, suture the horizontal back side.</li>
	<li>Next suture the horizontal front side, starting at the dome of the aneurysm moving to its base. Afterwards, start with single stitches distally on the front side moving caudally.
	<ol>
		<li>For all steps 4.5&#8210;4.8 while suturing the anastomosis pay attention just to grab the part of the vessel close to the arteriotomy to avoid iatrogenic stenosis. Also, continuously moisten all vessel segments during the whole surgical procedure extraluminally with a syringe filled with heparinized sodium chloride solution (500 IU dissolved in 100 ml&#160;of 0.9% isotonic sodium chloride) and protect them with wet micro swabs.</li>
		<li>Before finishing the anastomosis, irrigate the whole complex with heparinized 0.9% isotonic sodium chloride solution intraluminal (500 IU dissolved in 100 ml&#160;of 0.9% isotonic sodium chloride). Beware that elastase modified arterial pouches have to be sewn on as quickly as possible because of their strong tendency to dry out and to thrombose. Because of the aggressive behavior of the residual elastase concentration in the pouch regarding digesting circumferential vessels, proceed fast with the surgery to re-perfuse the vessel complex quickly.</li>
	</ol>
	</li>
	<li>Remove all temporary vascular clamps stepwise.
	<ol>
		<li>Remove the distal clamp from the left CCA. Accept minor bleeding and staunch it by gently imprinting micro swabs on the anastomosis. Afterwards, remove the clamp of the right CCA, press gently with micro swab and forceps to avoid thrombus formation.</li>
		<li>If needed, replace the temporary vascular clips to provide enough coagulation. Afterwards, relieve both vessel clips from the left side proximally. If needed in any step, replace clips to allow coagulation or to perform re-stitching.</li>
	</ol>
	</li>
	<li>At this stage perform fluorescence angiography of the vessel complex (Figure 6 and Figure 7). 
	NOTE: Fluorescence angiography is performed by administering 1 ml of fluorescein IV, using 2 bandpass filters, a smartphone with video camera, and a bicycle spotlight. This procedure has been already described elsewhere20,21,22.</li>
	<li>Lastly, close the operative situs. Readapt and gently suture the fat pad with a 3-0 resorbable suture with single nodes to protect the anastomosis. Close subcutis and skin in the same fashion.</li>
</ol>
5. Postsurgical phase
<ol>
	<li>Discontinue the isoflurane and systemic analgesia administration at the end of the surgery and provide tracheal extubation as soon as the swallowing reflex has returned.</li>
	<li>Administer 0.5 mg/kg of meloxicam IV, 10 mg/kg aspirin (ASS) IV, 100 &#181;g of vitamin B12&#160;SC&#160;and 20 mg/kg of clamoxyl IV.</li>
	<li>Provide supplementary oxygenation and active warming until the rabbits have spontaneously regained sternal recumbency.</li>
	<li>Perform postoperative follow-up and animal care four times a day for the first three days, in accordance to the guidelines for the assessment and management of pain in rodents and rabbits23,24.</li>
	<li>Administer post-operative analgesia via a fentanyl patch (12 &#181;g/h) applied on the outer ear, meloxicam once a day SC for three days and methadone as rescue therapy SC, according with the score sheet for pain evaluation. Administer 250 IU/kg low-molecular heparin (LMH) subcutaneously for three days in all rabbits.</li>
</ol>

This work was supported by the research funds of the Research Council, Kantonsspital Aarau, Aarau, Switzerland and the Swiss national science foundation SNF (310030_182450). The authors are solely responsible for the design and conduct of the presented study and declare no competing interests.

Our study demonstrates the feasibility of creating a true bifurcation aneurysm model with different wall conditions in rabbits. Overall, 14 female New Zealand White rabbits with a mean weight of 3.7 &#177; 0.09 kg and mean age of 112 &#177; 3 days were included in the study. 85.72% of all aneurysms remained patent during a follow-up at 28 days. Two animals died prematurely (12.5% mortality).
Previous studies suggested a variety of extracranial aneurysm models to analyze the management of endov...

Subarachnoid hemorrhage resulting from intracranial aneurysm (IA) rupture&#160;can effectively be controlled by either endovascular or microsurgical occlusion techniques1,2,3,4. Different endovascular therapies, to overcome the main limitation of IA recurrence after coiling, gained importance over the past decades generating an increased need of testing endovascular devices. To test these novel treatment approaches, appropriate animal models that respect rheological properties, hemodynamics and aneurysm wall conditions are highly warranted5,6,7. In this context, clinical as well as preclinical studies have already revealed the important role of aneurysm wall conditions regarding aneurysm rupture and recurrence after occlusion, especially focusing on the loss of mural cells7,8,9.
So far, experimental aneurysms in rabbits have most often been created either by elastase incubated common carotid artery (CCA) stumps or venous pouches sutured into an artificial CCA bifurcation.10,11,12,13,14,15,16 Thus, a true arterial pouch bifurcation model has never been described.
The aim of this study was to design a safe, fast, and standardized technique for microsurgical creation of bifurcation aneurysms with different wall conditions in a rabbit model (Figure 1). This was achieved by suturing non-modified and modified arterial pouches into an artificial created bifurcation of both CCAs.

<table><tbody><tr><td>3-0 resorbable suture</td><td>Ethicon Inc., USA</td><td>VCP428G</td><td></td></tr><tr><td>4-0 non-absorbable suture</td><td>B. Braun, Germany</td><td>G0762563</td><td></td></tr><tr><td>6-0 non-absorbable suture</td><td>B. Braun, Germany</td><td>C0766070</td><td></td></tr><tr><td>9-0 non-absorbable suture</td><td>B. Braun, Germany</td><td>G1111140</td><td></td></tr><tr><td>Adrenaline</td><td>Amino AG</td><td>1445419</td><td>any generic</td></tr><tr><td>Amiodarone</td><td>Helvepharm AG</td><td>5078567</td><td>any generic</td></tr><tr><td>Anesthesia machine</td><td>Dr&amp;auml;ger</td><td></td><td>any other</td></tr><tr><td>Aspirin</td><td>Sanofi-Aventis (Suisse) SA</td><td>622693</td><td>any generic</td></tr><tr><td>Atropine</td><td>Labatec Pharma SA</td><td>6577083</td><td>any generic</td></tr><tr><td>Bandpass filter blue</td><td>Thorlabs</td><td>FD1B</td><td>any other</td></tr><tr><td>Bandpass filter green</td><td>Thorlabs</td><td>FGV9</td><td>any other</td></tr><tr><td>Bipolar forceps</td><td></td><td></td><td>any other</td></tr><tr><td>Bicycle spotlight</td><td></td><td></td><td>any other</td></tr><tr><td>Biemer vessel clip (2 x)</td><td>B. Braun Medical AG, Aesculap, Switzerland</td><td>FD560R</td><td>temporary</td></tr><tr><td>Bispectral index (neonatal)</td><td></td><td></td><td>any other</td></tr><tr><td>Blood pressure cuff (neonatal)</td><td></td><td></td><td>any other</td></tr><tr><td>Clamoxyl</td><td>GlaxoSmithKline AG</td><td>758808</td><td>any generic</td></tr><tr><td>Dexmedetomidine</td><td>Ever Pharma</td><td>136740-1</td><td>any generic</td></tr><tr><td>Electrocardiogram electrodes</td><td></td><td></td><td>any other</td></tr><tr><td>Elastase</td><td>Sigma Aldrich</td><td>45125</td><td>any generic</td></tr><tr><td>Ephedrine</td><td>Amino AG</td><td>1435734</td><td>any generic</td></tr><tr><td>Esmolol</td><td>OrPha Swiss GmbH</td><td>3284044</td><td>any generic</td></tr><tr><td>Fentanyl (intravenous use)</td><td>Janssen-Cilag AG</td><td>98683</td><td>any generic</td></tr><tr><td>Fentanyl (transdermal)</td><td>Mepha Pharma AG</td><td>4008286</td><td>any generic</td></tr><tr><td>Fluoresceine</td><td>Curatis AG</td><td>5030376</td><td>any generic</td></tr><tr><td>Fragmin</td><td>Pfizer PFE Switzerland GmbH</td><td>1906725</td><td>any generic</td></tr><tr><td>Glyco</td><td></td><td></td><td>any generic</td></tr><tr><td>Heating pad</td><td></td><td></td><td>any other</td></tr><tr><td>Isotonic sodium chloride solution (0.9%)</td><td>Fresenius KABI</td><td>336769</td><td>any generic</td></tr><tr><td>Ketamine</td><td>Pfizer</td><td>342261</td><td>any generic</td></tr><tr><td>Laboratory shaker</td><td>Stuart</td><td>SRT6</td><td>any other</td></tr><tr><td>Lidocaine</td><td>Streuli Pharma AG</td><td>747466</td><td>any generic</td></tr><tr><td>Longuettes</td><td></td><td></td><td>any other</td></tr><tr><td>Metacam</td><td>Boehringer Ingelheim</td><td>P7626406</td><td>any generic</td></tr><tr><td>Methadone</td><td>Streuli Pharma AG</td><td>1084546</td><td>any generic</td></tr><tr><td>Microtubes</td><td></td><td></td><td>any other</td></tr><tr><td>Micro needle holder</td><td></td><td></td><td>any other</td></tr><tr><td>Midazolam</td><td>Accord Healthcare AG</td><td>7752484</td><td>any generic</td></tr><tr><td>Needle holder</td><td></td><td></td><td>any other</td></tr><tr><td>O2-Face mask</td><td></td><td></td><td>any other</td></tr><tr><td>Operation microscope</td><td>Wild Heerbrugg</td><td></td><td>any other</td></tr><tr><td>Papaverine</td><td>Bichsel</td><td></td><td>any generic</td></tr><tr><td>Prilocaine-lidocaine creme</td><td>Emla</td><td></td><td>any generic</td></tr><tr><td>Propofol</td><td>B. Braun Medical AG, Switzerland</td><td></td><td>any generic</td></tr><tr><td>Pulse oxymeter</td><td></td><td></td><td>any generic</td></tr><tr><td>Rectal temperature probe (neonatal)</td><td></td><td></td><td>any other</td></tr><tr><td>Ropivacaine</td><td>Aspen Pharma Schweiz GmbH</td><td>1882249</td><td>any generic</td></tr><tr><td>Scalpell</td><td>Swann-Morton</td><td>210</td><td>any other</td></tr><tr><td>Small animal shaver</td><td></td><td></td><td>any other</td></tr><tr><td>Smartphone</td><td></td><td></td><td>any other</td></tr><tr><td>Soft tissue forceps</td><td></td><td></td><td>any other</td></tr><tr><td>Soft tissue spreader</td><td></td><td></td><td>any other</td></tr><tr><td>Stainless steel sponge bowls</td><td></td><td></td><td>any other</td></tr><tr><td>Sterile micro swabs</td><td></td><td></td><td>any other</td></tr><tr><td>Stethoscope</td><td></td><td></td><td>any other</td></tr><tr><td>Straight and curved micro-forceps</td><td></td><td></td><td>any other</td></tr><tr><td>Straight and curved micro-scissors</td><td></td><td></td><td>any other</td></tr><tr><td>Straight and curved forceps</td><td></td><td></td><td>any other</td></tr><tr><td>Surgery drape</td><td></td><td></td><td>any other</td></tr><tr><td>Surgical scissors</td><td></td><td></td><td>any other</td></tr><tr><td>Syringes 1 ml, 2ml and 5 ml</td><td></td><td></td><td>any other</td></tr><tr><td>Tris-Buffer</td><td>Sigma Aldrich</td><td>93302</td><td>any generic</td></tr><tr><td>Vascular clip applicator</td><td>B. Braun, Germany</td><td>FT495T</td><td></td></tr><tr><td>Vein and arterial catheter 22 G</td><td></td><td></td><td>any generic</td></tr><tr><td>Vitarubin</td><td>Streuli Pharma AG</td><td>6847559</td><td>any generic</td></tr><tr><td>Yasargil titan standard clip (2 x)</td><td>B. Braun Medical AG, Aesculap, Switzerland</td><td>FT242T</td><td>temporary</td></tr></tbody></table>

arterial pouch microsurgical bifurcation aneurysm model in the rabbit

Endovascular treatment for intracranial aneurysms gained importance over the past decades, consequently there is an increased need of testing endovascular devices. Animal models respecting rheological, hemodynamic and aneurysm wall conditions are highly warranted. Therefore, the aim of the present study was to design a novel standardized and reproducible surgical technique to create autologous arterial pouch bifurcation aneurysms with non-modified and modified wall conditions in rabbits.
Bifurcation aneurysms were created by end-to-side anastomosis of the right on the left common carotid artery, both serving as parent arteries for the arterial pouch, which was microsurgically sewn on. Grafts were taken from the proximal right common carotid artery, either for the control (n = 7, immediate autologous re-implantation) or modified (n = 7, incubated with 100 international units elastase for 20 minutes before autologous re-implantation) group. Pouch and parent artery patency were controlled by fluorescence angiography immediately after creation. At follow-up (28 days), all rabbits underwent contrast enhanced magnetic resonance angiography and fluorescence angiography followed by aneurysm harvesting, macroscopic and histological evaluation.
A total of 16 female New Zealand White rabbits were operated upon. Two animals died prematurely. At follow-up, 85.72% of all aneurysms remained patent. Both groups revealed an increase in aneurysm size over time; this was more pronounced in the control group (6.48 &#177; 1.81 mm3 at time of creation vs. 19.85 &#177; 6.40 mm3 at follow-up, p = 0.037) than in the modified group (8.03 &#177; 1.08 mm3 at time of creation vs. 20.29 &#177; 6.16 mm3 at follow-up, p = 0.054).
Our findings demonstrate the adequacy of this new rabbit model which allows for the creation of bifurcation aneurysms with different wall conditions in a microsurgical approach. Given the excellent long-term patency and the property of aneurysm growth over time, this model may serve as an important tool for preclinical evaluation of novel endovascular therapies.

Following a pilot series of seven animals, totally 16 animals were included in the experimental protocol. Two animals died prematurely and were therefore excluded from the final analysis (12.5% mortality). Calculated on 14 animals, immediate aneurysm patency rate during fluorescence angiography was 71.43% in both, the control and modified group. Four aneurysms had to be reopened with consecutive thrombus evacuation and after a repeated fluorescence angiography there was a documented patency in all cases (100%). Aneurysm ...

Watch this Scientific Journal Video about Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit at JoVE.com

Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit

Developing and testing endovascular devices for intracranial aneurysm treatment is still of great importance. Most aneurysm models used today miss either the important characteristics of an arterial degenerated wall or the hemodynamics of a true bifurcation. Therefore, we aimed to design a novel arterial pouch bifurcation model in rabbits.

Endovascular treatment for intracranial aneurysms gained importance over the past decades, consequently there is an increased need of testing endovascular ...

arterial-pouch-microsurgical-bifurcation-aneurysm-model-in-the-rabbit

Research

JoVE Journal

Neuroscience

2.3K Views.  Kantonsspital Aarau. Het ontwikkelen en testen van endovasculaire apparaten voor intracraniale aneurysmabehandeling is nog steeds van groot belang. De meeste aneurysmamodellen die tegenwoordig worden gebruikt, missen de belangrijke kenmerken van een arteriële gedegenereerde wand of de hemodynamiek van een echte bifurcatie. Daarom wilden we een nieuw arteriële buidelbifurcatiemodel bij konijnen ontwerpen.

Video: Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit

Microsurgical Venous Pouch Arterial-Bifurcation Aneurysms in the Rabbit Model: Technical Aspects

For ruptured human cerebral aneurysms endovascular embolization has become an equivalent alternative to aneurysm clipping.1 However, large clinical trials have shown disappointing long-term results with unacceptable high rates of aneurysm recanalization and delayed aneurysm rupture.2 To overcome these problems, animal experimental studies are crucial for the development of better endovascular devices.3-5
Several animal models in rats, rabbits, canines and swine are available.6-8 Comparisons of the different animal models showed the superiority of the rabbit model with regard to hemodynamics and comparability of the coagulation system and cost-effectiveness.9-11 
The venous pouch arterial bifurcation model in rabbits is formed by a venous pouch sutured into an artificially created true bifurcation of both common carotid arteries (CCA). The main advantage of this model are true bifurcational hemodynamics.12 The major drawbacks are the sofar high microsurgical technical demands and high morbidity and mortality rates of up to 50%.13 These limitations have resulted in less frequent use of this aneurysm model in the recent years. These shortcomings could be overcome with improved surgical procedures and modified peri- and postoperative analgetic management and anticoagulation.14-16 Our techniques reported in this paper demonstrate this optimized technique for microsurgical creation of arterial bifurcation aneurysms.

An optimized technique for the microsurgical creation of arterial bifurcation aneurysms mimicking bifurcation human cerebral aneurysms is described. A venous pouch is sutured into an artificially created true bifurcation of both common carotid arteries. Facilitated microsurgical techniques and aggressive postoperative anticoagulation and analgesia lead to minimized morbidity rates and high aneurysm patency rates.

For ruptured human cerebral aneurysms endovascular embolization has become an equivalent alternative to aneurysm clipping.1 However, large clinical trials ...

Medicine

Detection of Neuritic Plaques in Alzheimer's Disease Mouse Model

Alzheimer's disease (AD) is the most common neurodegenerative disorder leading to dementia. Neuritic plaque formation is one of the pathological hallmarks of Alzheimer's disease. The central component of neuritic plaques is a small filamentous protein called amyloid &beta; protein (A&beta;)1, which is derived from sequential proteolytic cleavage of the beta-amyloid precursor protein (APP) by &beta;-secretase and &gamma;-secretase. The amyloid hypothesis entails that A&gamma;-containing plaques as the underlying toxic mechanism in AD pathology2. The postmortem analysis of the presence of neuritic plaque confirms the diagnosis of AD. To further our understanding of A&gamma; neurobiology in AD pathogenesis, various mouse strains expressing AD-related mutations in the human APP genes were generated. Depending on the severity of the disease, these mice will develop neuritic plaques at different ages. These mice serve as invaluable tools for studying the pathogenesis and drug development that could affect the APP processing pathway and neuritic plaque formation. In this protocol, we employ an immunohistochemical method for specific detection of neuritic plaques in AD model mice. We will specifically discuss the preparation from extracting the half brain, paraformaldehyde fixation, cryosectioning, and two methods to detect neurotic plaques in AD transgenic mice: immunohistochemical detection using the ABC and DAB method and fluorescent detection using thiofalvin S staining method.

One of the pathological characteristics of AD is the formation of Amyloid &#x03B2; protein positive neuritic plaques. In this protocol we describe two methods to detect neuritic plaques in transgenic AD model mice: immunohistochemical detection using the ABC and DAB method and fluorescent detection using thioflavin S staining method.

Alzheimer's disease (AD) is the most common neurodegenerative disorder leading to dementia. Neuritic plaque formation is one of the pathological hallmarks ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

The Helsinki Rat Microsurgical Sidewall Aneurysm Model

Experimental saccular aneurysm models are necessary for testing novel surgical and endovascular treatment options and devices before they are introduced into clinical practice. Furthermore, experimental models are needed to elucidate the complex aneurysm biology leading to rupture of saccular aneurysms.
Several different kinds of experimental models for saccular aneurysms have been established in different species. Many of them, however, require special skills, expensive equipment, or special environments, which limits their widespread use. A simple, robust, and inexpensive experimental model is needed as a standardized tool that can be used in a standardized manner in various institutions.
The microsurgical rat abdominal aortic sidewall aneurysm model combines the possibility to study both novel endovascular treatment strategies and the molecular basis of aneurysm biology in a standardized and inexpensive manner. Standardized grafts by means of shape, size, and geometry are harvested from a donor rat&#39;s descending thoracic aorta and then transplanted to a syngenic recipient rat. The aneurysms are sutured end-to-side with continuous or interrupted 9-0 nylon sutures to the infrarenal abdominal aorta.
We present step-by-step procedural instructions, information on necessary equipment, and discuss important anatomical and surgical details for successful microsurgical creation of an abdominal aortic sidewall aneurysm in the rat.

Microsurgical sidewall aneurysms in rats are created by end-to-side anastomosis of an aortic graft to the abdominal aorta. We present step-by-step instructions and discuss anatomical and surgical details for successful experimental saccular aneurysm creation.

Experimental saccular aneurysm models are necessary for testing novel surgical and endovascular treatment options and devices before they are introduced ...

A Rabbit Venous Interposition Model Mimicking Revascularization Surgery using Vein Grafts to Assess Intimal Hyperplasia under Arterial Blood Pressure

Although vein grafts have been commonly used as autologous grafts in revascularization surgeries for ischemic diseases, the long-term patency remains poor because of the acceleration of intimal hyperplasia due to the exposure to arterial blood pressure. The present protocol is designed for the establishment of experimental venous intimal hyperplasia by interposing rabbit jugular veins to the ipsilateral carotid arteries. The protocol does not require surgical procedures deep in the body trunk and the extent of the incision is limited, which is less invasive for the animals, allowing long-term observation after implantation. This simple procedure enables researchers to investigate strategies to attenuate the progression of intimal hyperplasia of the implanted vein grafts. Using this protocol, we reported the effects transduction of microRNA-145 (miR-145), which is known to control the phenotype of vascular smooth muscle cells (VSMCs) from the proliferative to the contractile state, into harvested vein grafts. We confirmed the attenuation of intimal hyperplasia of vein grafts by transducing miR-145 before implantation surgery through the phenotype change of the VSMCs. Here we report a less invasive experimental platform to investigate the strategies that can be used to attenuate intimal hyperplasia of vein grafts in revascularization surgeries.

The present protocol aims to experimentally create venous intimal hyperplasia by subjecting veins to arterial blood pressure for developing strategies to attenuate venous intimal hyperplasia following revascularization surgery using vein grafts.

Although vein grafts have been commonly used as autologous grafts in revascularization surgeries for ischemic diseases, the long-term patency remains poor ...

Creation of Two Saccular Elastase-Digested Aneurysms with Different Hemodynamics in One Rabbit

Preclinical animal models with hemodynamic, morphologic, and histologic characteristics close to human intracranial aneurysms play a key role in the understanding of the pathophysiological processes and the development and testing of new therapeutic strategies. This study aims to describe a new rabbit aneurysm model that allows the creation of two elastase-digested saccular aneurysms with different hemodynamic conditions within the same animal.
Five female New Zealand white rabbits with a mean weight of 4.0 (&#177; 0.3) kg and mean age of 25 (&#177;5) weeks underwent microsurgical stump and bifurcation aneurysm creation. One aneurysm (stump) was created by right common carotid artery (CCA) exposure at its origin at the brachiocephalic trunk. A temporary clip was applied at the CCA origin and another, 2 cm above. This segment was treated with a local injection of 100 U of elastase for 20 min. A second aneurysm (bifurcation) was created by suturing an elastase-treated arterial pouch into the end-to-side anastomosis of the right CCA to left CCA. Patency was controlled by fluorescence angiography immediately after creation.
The average duration of surgery was 221 min. The creation of two aneurysms in the same animal was successful in all rabbits without complication. All aneurysms were patent immediately after surgery except for one bifurcation aneurysm, which showed an extreme tissue reaction due to elastase incubation and an immediate intraluminal thrombosis. No mortality was observed during surgery and up to one-month follow-up. Morbidity was limited to a transient vestibular syndrome (one rabbit), which recovered spontaneously within one day.
Demonstrated here for the first time is the feasibility of creating a two-aneurysm rabbit model with stump and bifurcation hemodynamic characteristics and highly degenerated wall conditions. This model allows the study of the natural course and potential treatment strategies on the basis of aneurysm biology under different flow conditions.

This protocol describes the steps for the creation of a rabbit model with two elastase-digested aneurysms with different hemodynamics (stump and bifurcation constellation). This allows the testing of novel endovascular devices in aneurysms with different angioarchitecture and hemodynamic conditions within a single animal.

Preclinical animal models with hemodynamic, morphologic, and histologic characteristics close to human intracranial aneurysms play a key role in the ...

Microsurgical Creation of Giant Bifurcation Aneurysms in Rabbits for the Evaluation of Endovascular Devices

Giant aneurysms are dangerous lesions requiring endovascular treatment, with high rates of aneurysm recanalization and re-rupture. Reliable in vivo models are rare but are required for testing new endovascular devices.&#160;We demonstrate the technical aspects of the creation of giant bifurcation aneurysms in New Zealand white rabbits (2.5-5.5 kg). A 25-30 mm long venous pouch is taken from the external jugular vein, and a bifurcation between both carotid arteries is created microsurgically. The pouch is sutured in the bifurcation to mimic a giant aneurysm. This protocol summarizes our previously published standard technique for venous pouch true arterial bifurcation aneurysms and highlights its essential modification steps for giant aneurysms. Using this modified technique, we were able to create an animal model for giant aneurysms with high comparability to humans regarding the hemodynamics and coagulation systems. Furthermore, low morbidity and high aneurysm patency rates were achieved. The proposed giant aneurysm model offers an excellent possibility for testing new endovascular devices.

Here, we describe the technique for the microsurgical creation of giant bifurcation aneurysms in rabbits for the evaluation of endovascular devices.

Giant aneurysms are dangerous lesions requiring endovascular treatment, with high rates of aneurysm recanalization and re-rupture. Reliable in vivo models ...

Swine Models of Aneurysmal Diseases for Training and Research

Large animal models, specifically swine, are widely used to research cardiovascular diseases and therapies, as well as for training purposes. This paper describes two different aneurysmal swine models that may help researchers to study new therapies for aneurysmal diseases. These aneurysmal models are created by surgically adding a pouch of tissue to carotid arteries in swine. When the model is used for research, the pouch must be autologous; for training purposes, a synthetic pouch suffices.
First, the right external jugular vein (EJV) and right common carotid artery (CCA) must be surgically exposed. The EJV is ligated and a vein pouch fashioned from a short segment. This pouch is then sutured to an elliptical arteriotomy performed in the CCA. Animals must be kept heparinized during model creation, and local vasodilators may be used to decrease vasospasms. Once the suture is completed, correct blood flow should be inspected, checking for bleeding from the suture line and vessel patency. Finally, the surgical incision is closed by layers and an angiography performed to image the aneurysmal model.
A simplification of this aneurysmal carotid model that decreases invasiveness and surgical time is the use of a synthetic, rather than venous, pouch. For this purpose, a pouch is tailored in advance with a segment of a polytetrafluoroethylene (PTFE) prosthesis, one end of which is sutured close using polypropylene vascular suture and sterilized prior to surgery. This "sac" is then attached to an arteriotomy performed in the CCA as described. 
Although these models do not reproduce many of the physiopathological events related to aneurysm formation, they are hemodynamically similar to the situation found in the clinical setting. Therefore, they can be used for research or training purposes, allowing physicians to learn and practice different endovascular techniques in animal models that are close to the human system.

Description of two different aneurysmal swine models for neuroradiology training courses and research studies. This study provides evidence of the feasibility of these aneurysm porcine model creations and the reproducible methods that are close to the clinical setting.

Large animal models, specifically swine, are widely used to research cardiovascular diseases and therapies, as well as for training purposes. This paper ...

A Modified Technique for Arteriovenous Fistula Construction in Rabbits

Juxta-anastomotic stenosis is a challenging problem that often causes non-maturation and decreases the patency of an arteriovenous fistula (AVF). Injury to the veins and arteries during the operation and hemodynamic changes can lead to intimal hyperplasia, leading to juxta-anastomotic stenosis. To reduce injury to the veins and arteries during the operation, this study proposes a new modified no-touch technique (MNTT) for AVF construction that can decrease the rate of juxta-anastomotic stenosis and improve the AVF patency. To unravel the hemodynamic changes and mechanisms of the MNTT, this study presented an AVF procedure using this technique. Although this procedure is technically challenging, 94.4% procedural success was achieved after adequate training. Ultimately, 13 out of 34 rabbits had a functional AVF 4 weeks after the surgery, leading to a 38.2% AVF patency rate. However, at 4 weeks, the survival rate was 86.1%. Ultrasonography showed active blood flow through AVF anastomosis. Furthermore, the spiral laminar flow was observed in the vein and artery near the anastomosis, suggesting that this technique may improve the hemodynamics of the AVF. On histological observation, significant venous intimal hyperplasia was observed at the AVF anastomosis, whereas no significant intimal hyperplasia was observed at the proximal external jugular vein (EJV) of the anastomosis. This technique will improve the understanding of the mechanisms underlying the use of MNTT for AVF construction and provide technical support for the further optimization of the surgical approach in AVF construction.

The present protocol proposes the creation of an arteriovenous fistula in rabbits using a modified no-touch technique. The technique involves the side-to-side anastomosis of the common carotid artery and external jugular vein without the dissection of the perivenous tissues or cutting off the artery.

Juxta-anastomotic stenosis is a challenging problem that often causes non-maturation and decreases the patency of an arteriovenous fistula (AVF). Injury ...

Developing an Experimental Aneurysm Model in a Rabbit Using an Elastase-Treated Vessel Pouch

<a href='https://app.jove.com/v/61157/arterial-pouch-microsurgical-bifurcation-aneurysm-model-in-the-rabbit'>Source: Wanderer, S., et al. Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit. J. Vis. Exp. (2020)</a>This video demonstrates the creation of an experimental aneurysm model in a rabbit using an elastase-treated vessel pouch. After excising a segment of the right CCA, the pouch is treated with elastase, then sutured to the left CCA to restore blood flow. The resulting stress on the weakened pouch walls induces aneurysm formation.

Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit

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