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&#228;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. 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.

Video: Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit

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.

Detection of Neuritic Plaques in Alzheimer&#039;s Disease Mouse Model

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 ...

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These neurotropic cancer cells have a unique ability to migrate unidirectionally along nerves towards the central nervous system (CNS). The dorsal root ganglia (DRG)/cancer cell model is a three dimensional (3D) in vitro model frequently used for studying the interaction between neural stroma and cancer cells. In this model, mouse or human cancer cell lines are grown in ECM adjacent to preparations of freshly dissociated cultured DRG. In this article, the DRG isolation protocol from mice, and implantation in petri dishes for co-culturing with pancreatic cancer cells are demonstrated. Five days after implantation, the cancer cells made contact with the DRG neurites. Later, these cells formed bridgeheads to facilitate more extensive polarized, neurotropic migration of cancer cells.

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

This video article shows the use of the dorsal root ganglia (DRG)/cancer cell model in pancreatic ductal adenocarcinoma.

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These ...

The Use of Trace Eyeblink Classical Conditioning to Assess Hippocampal Dysfunction in a Rat Model of Fetal Alcohol Spectrum Disorders

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain undergoes rapid organizational change and is similar to accelerated brain changes that occur during the third trimester in humans. This model of fetal alcohol spectrum disorders (FASDs) produces severe brain damage, mimicking the amount and pattern of binge-drinking that occurs in some pregnant alcoholic mothers. We describe the use of trace eyeblink classical conditioning (ECC), a higher-order variant of associative learning, to assess long-term hippocampal dysfunction that is typically seen in alcohol-exposed adult offspring. At 90 days of age, rodents were surgically prepared with recording and stimulating electrodes, which measured electromyographic (EMG) blink activity from the left eyelid muscle and delivered mild shock posterior to the left eye, respectively. After a 5 day recovery period, they underwent 6 sessions of trace ECC to determine associative learning differences between alcohol-exposed and control rats. Trace ECC is one of many possible ECC procedures that can be easily modified using the same equipment and software, so that different neural systems can be assessed. ECC procedures in general, can be used as diagnostic tools for detecting neural pathology in different brain systems and different conditions that insult the brain.

Trace eyeblink classical conditioning (ECC) was used to assess hippocampal-dependent associative learning in adult rats that were administered a high concentration (11.9% v/v) of alcohol during early neonatal brain development. In general, ECC procedures are sound diagnostic tools for detecting brain dysfunction across many psychological and biomedical settings.

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain ...

A Volumetric Method for Quantification of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor determining patient outcome and is therefore frequently taken as a study endpoint. However, in small animal studies on SAH, quantification of cerebral vasospasm is a major challenge. Here, an ex vivo method is presented that allows quantification of volumes of entire vessel segments, which can be used as an objective measure to quantify cerebral vasospasm. In a first step, endovascular casting of the cerebral vasculature is performed using a radiopaque casting agent. Then, cross-sectional imaging data are acquired by micro computed tomography. The final step involves 3-dimensional reconstruction of the virtual vascular tree, followed by an algorithm to calculate center lines and volumes of the selected vessel segments. The method resulted in a highly accurate virtual reconstruction of the cerebrovascular tree shown by a diameter-based comparison of anatomical samples with their virtual reconstructions. Compared with vessel diameters alone, the vessel volumes highlight the differences between vasospastic and non-vasospastic vessels shown in a series of SAH and sham-operated mice.

The goal of this article is to present a method that allows a 3-dimensional reconstruction of the cerebrovascular tree in mice after micro computed tomography and determination of volumes of entire vessel segments that can be used to quantify cerebral vasospasm in murine models of subarachnoid hemorrhage.

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor ...

Fluorescence Angiography for Evaluation of Aneurysm Perfusion and Parent Artery Patency in Rat and Rabbit Aneurysm Models

Brain aneurysm treatment focuses on achieving complete occlusion, as well as preserving blood flow in the parent artery. Fluorescein sodium and indocyanine green are used to enable the observation of blood flow and vessel perfusion status, respectively. The aim of this study is to apply FVA to verify real-time blood flow, vessel perfusion status and occlusion of aneurysms after induction of sidewall aneurysms in rabbits and rats, as well as to validate the procedure in these species.
Twenty sidewall aneurysms were created in 10 rabbits by suturing a decellularized arterial vessel pouch on the carotid artery of a donor rabbit. In addition, 48 microsurgical sidewall aneurysms were created in 48 rats. During follow-up at one month after creation, the parent artery/aneurysm complex was dissected and FVA was performed using an intravenous fluorescein (10%, 1 mL) injection via an ear vein catheterization in rabbits and a femoral vein catherization in rats. Aneurysms were then harvested, and patency was evaluated macroscopically.
Macroscopically, 14 out of 16 aneurysms in rabbits indicated no residual parent artery perfusion with totally occluded luminae, however 11 (79%) were detected by FVA. Four aneurysms were excluded due to technical problems. In rats, residual aneurysm perfusion was macroscopically observed in 25 out of 48 cases. Of the 23 without macroscopic evidence of perfusion, FVA confirmed the incidence of 22 aneurysms (96%). There were no adverse events associated with FVA. Fluorescein is easily applicable and no special equipment is needed. It is a safe and extremely effective method for evaluating parent artery integrity and aneurysm patency/residual perfusion in an experimental setting with rabbits and rats. FVA using fluorescein as a contrast agent appears to be effective in controlling patency of aneurysms and the underlying vessel and can even be adapted to bypass surgery.

We present a protocol to efficiently evaluate aneurysm perfusion and vessel patency of sidewall aneurysm in rats and rabbits, using fluorescein-based fluorescence video angiography (FVA). With a positive predictive value of 92.6%, it is a simple but very effective and economical method with no special equipment required.

Brain aneurysm treatment focuses on achieving complete occlusion, as well as preserving blood flow in the parent artery. Fluorescein sodium and ...

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

Double Direct Injection of Blood into the Cisterna Magna as a Model of Subarachnoid Hemorrhage

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% of the total stroke-related mortality with an important morbidity in terms of neurological outcome. A delayed cerebral vasospasm (CVS) may occur most often in association with a delayed cerebral ischemia. Different animal models of SAH are now being used including endovascular perforation and direct injection of blood into the cisterna magna or even the prechiasmatic cistern, each exhibiting distinct advantages and disadvantages. In this article, a standardized mouse model of SAH by double direct injection of determined volumes of autologous whole blood into the cisterna magna is presented. Briefly, mice were weighed and then anesthetized by isoflurane inhalation. Then, the animal was placed in a reclining position on a heated blanket maintaining a rectal temperature of 37 &#176;C and positioned in a stereotactic frame with a cervical bend of about 30&#176;. Once in place, the tip of an elongated glass micropipette filled with the homologous arterial blood taken from carotid artery of another mouse of the same age and gender (C57Bl/6J) was positioned at a right angle in contact with the atlanto-occipital membrane by means of a micromanipulator. Then 60 &#181;L of blood was injected in the cisterna magna followed by a 30&#176; downward tilt of the animal for 2 minutes. The second infusion of 30 &#181;L of blood into the cisterna magna was performed 24 h after the first one. The individual follow-up of each animal is carried out daily (careful evaluation of weight and well-being). This procedure allows a predictable and highly reproducible distribution of blood, likely accompanied by intracranial pressure elevation that can be mimicked by an equivalent injection of an artificial cerebral spinal fluid (CSF), and represents an acute to mild-model of SAH inducing low mortality.

We described in this protocol a standardized subarachnoid hemorrhage (SAH) mouse model by a double injection of autologous whole-blood into the cisterna magna. The high degree of standardization of the double-injection procedure represents a middle-to-acute model of SAH with relative safety regarding mortality.

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% ...

A Rat Model of EcoHIV Brain Infection

It has been well studied that the EcoHIV infected mouse model is of significant utility in investigating HIV associated neurological complications. Establishment of the EcoHIV infected rat model for studies of drug abuse and neurocognitive disorders, would be beneficial in the study of neuroHIV and HIV-1 associated neurocognitive disorders (HAND). In the present study, we demonstrate the successful creation of a rat model of active HIV infection using chimeric HIV (EcoHIV). First, the lentiviral construct of EcoHIV was packaged in cultured 293 FT cells for 48 hours. Then, the conditional medium was concentrated and titered. Next, we performed bilateral stereotaxic injections of the EcoHIV-EGFP into F344/N rat brain tissue. One week after infection, EGFP fluorescence signals were detected in the infected brain tissue, indicating that EcoHIV successfully induces an active HIV infection in rats. In addition, immunostaining for the microglial cell marker, Iba1, was performed. The results indicated that microglia were the predominant cell type harboring EcoHIV. Furthermore, EcoHIV rats exhibited alterations in temporal processing, a potential underlying neurobehavioral mechanism of HAND as well as synaptic dysfunction eight weeks after infection. Collectively, the present study extends the EcoHIV model of HIV-1 infection to the rat offering a valuable biological system to study HIV-1 viral reservoirs in the brain as well as HAND and associated comorbidities such as drug abuse.

Here, we present a protocol to establish a new rat model of active HIV infection using chimeric HIV (EcoHIV), which is critical for enhancing our understanding of HIV-1 viral reservoirs in the brain and offering a system to study HIV-associated neurocognitive disorders and associated comorbidities (i.e., drug abuse).

It has been well studied that the EcoHIV infected mouse model is of significant utility in investigating HIV associated neurological complications. ...