The authors thank Laurie von Melchner, Bern University Hospital, Department of Neurosurgery, Bern, Switzerland, for proofreading and editing the manuscript and Paskus Jeremiah, Boston Children&#8217;s Hospital, Boston, MA for proofreading the initial draft. We appreciate the skillful management of animal care, anesthesia, and operative assistance from Daniel Mettler, DVM, Max M&#252;ller, DVM, Daniel Zalokar, and Olgica Beslac, Experimental Surgical Institute, Department of Clinical Research, University of Bern, Bern, Switzerland. We thank Michael Lensch, Head Research Nurse, Department of Intensive Care Medicine, Bern University Hospital and University of Bern, Bern, Switzerland, for real-time data monitoring and post-processing of the physiological parameters. We thank Edin Nevzati, Carl Muroi, and Salome Erhardt, for their excellent laboratory technical and operative assistance.
This work was supported by the Department of Intensive Care Medicine, Bern University Hospital and University of Bern, Bern, Switzerland, the Department of Clinical Research, University of Bern, Bern, Switzerland, and the Research Fund from the Kantonsspital Aarau, Aarau, Switzerland. We thank Elsevier, for reprint permission for Figures 1 and 2.

Three month old female New Zealand rabbits weighing 2.5 &#8211; 3.5 kg were used for this procedure. The study was performed in accordance with the National Institutes of Health guidelines for the care and use of experimental animals and with the approval of the Animal Care Committee of the Canton of Bern, Switzerland (approval #105/13). All surgical procedures were performed under sterile conditions at the Experimental Surgical Institute of the Department of Clinical Research at Bern University Hospital in Bern, Switzerland. A veterinary anesthesiologist monitored the animals during surgery and throughout recovery.
1. Animal Preparation, Positioning and Subclavian Artery Cannulation
<ol>
	<li>Induce general anesthesia in the rabbit with intramuscular injection of ketamine (30 mg/kg; Ketalar, 50 mg/ml) and xylazine (6 mg/kg; Xylapan 20 mg/ml) and control depth of anesthesia by checking the rabbit&#8217;s response to noxious stimulation (e.g., toe pinch). See 1.7 in case of positive response.</li>
	<li>Pull down the lower lids of both eyes and apply a small amount of ointment to the eyelids to prevent dryness and further irritation.</li>
	<li>Catheterize the lateral ear vein with a 20 Gbutterfly venflon (20 G vascular catheter), fix with adhesive tape, and connect to a gravity bag containing 0.9% sodium chloride (500 ml/24 hr) and ketamine (40 mg/kg/hr) / xylazine (4 mg/kg/hr) for continuous intravenous (i.v.) anesthesia. Administer additional analgesics every 15 min i.v. (Fentanyl, 1 mcg/kg). Note: Avoid volatile gas anesthetics, which is associated with decreased CPP, increasing CBF, and decreasing cerebral metabolic rate for oxygen.11 Intravenous anesthetics provide more ideal characteristics for neuroanesthesia by preserving CBF and cerebral vasoconstriction,12 which is of uppermost importance when studying cerebral vasospasm. In addition, although mortality is increased in spontaneously breathing animals, it may better mimic the human situation of acute SAH.</li>
	<li>Provide oxygen (1 - 2 L/min) via a respiratory mask that allows end-tidal carbon dioxide (EtCO2) monitoring.</li>
	<li>Install a 3 channel electrocardiogram (ECG).Place three subcutaneous electrodes in a triangular arrangement on the ventral side of the rabbit; specifically, place one electrode over the right midthoracic region (with distance to the sterile shaved field for the subclavian artery cannulation) and two electrodes in the lower abdomen distributed over both limbs.</li>
	<li>Monitor the depth of anesthesia every 15 min during surgery by following respiratory rate, heart rate (HR) monitored from the ECG signal, and reaction to noxious stimulation.</li>
	<li>In case of positive response to noxious stimulation (toe pinch), adapt the depth of anesthesia by ketamine bolus (6 mg/kg) i.v. and xylazine bolus (0.05 mg/kg) i.v. and/or an additional analgesia bolus with Fentanyl (1 mcg/kg) i.v.</li>
	<li>Fix the rabbit in a supine position on a body warming plate, tilting the head 20&#176; down and turning it slightly contralaterally to the side on which the subclavian artery will be exposed.</li>
	<li>apply eye ointment and prepare the area for surgery by shaving the hair over the right pectoral muscle around the middle third of the clavicle, and over the frontal-, parietal- and occipital skull, the neck, and over the right common femoral artery.</li>
	<li>Disinfect the skin for 3 min with a broad spectrum antiseptic, e.g., Povidone-iodine.</li>
	<li>Cover the rabbit with sterile sheets. Perform all further procedures under sterile conditions and frequently apply 4% papaverin HCl and antibiotic solution (neomycin sulfate 5 mg/ml) topically to prevent arterial vasospasm by vessel manipulation and local infections.</li>
	<li>Infiltrate the pectoral muscle with local anesthetics (Lidocaine 1% maximal 6 mg/kg). Make a parasternal skin incision and prepare the pectoral muscle. Using the microscope, dissect the subclavian artery and secure with a proximal and distal ligature (4-0 polyfilament sutures) around the exposed end. Keep one ligature close to the proximal control in place to secure the catheter and ligate the vessel distally.</li>
	<li>Perform an arteriotomy in the wall of the subclavian artery by incising the artery with a curved microscissor and cannulate the subclavian artery retrogradely with a small intravasal 3-way stopcock. Secure the catheter by double knot ligature towards the distal ligature in order to prevent arterial twisting or bending of the proximal part of the artery and to avoid slippage or massive bleeding.</li>
</ol>
2. Blood Pressure and Arterial Blood Gas Monitoring
<ol>
	<li>Connect the 3-way stop cock to i) the intravasal cannula for arterial blood gas (ABG) analyses, pH, PaCO2, PaO2, bicarbonate, base excess, and SO2, ii) the invasive arterial blood pressure measurement device, and iii) the shunt device.</li>
	<li>Collect blood samples for ABG status (PaCO2, PaO2) and continuously monitor standard cardiovascular and respiratory parameters (blood pressure, HR, ECG, respiratory rate and end-tidal CO2) and transfer data via the analog output interface to an analog&#8211;digital converter/data logger and store. 
	NOTE: The pressures will be zeroed at heart levels before and after each session, and pressure calibration of the analog/digital-converter and data-logging system will be done once before the series starts.</li>
</ol>
3. Baseline Digital Subtraction Angiography
<ol>
	<li>Place an external sizing device (small sphere) over both mandible angles in order to calibrate the angiogram. 
	NOTE: This allows exact comparison of post hoc measurements of baseline and follow up vessel diameter.</li>
	<li>Perform digital subtraction angiography (DSA) by retrograde intra-arterial bolus injection of non-ionic Iopamidol (0.6 ml/kg, 5 ml/sec for 2 sec) through the cannulated artery and flush the cannula immediately after bolus injection with saline in order to prevent occlusion of the latter.</li>
	<li>Obtain images (7 images in 14 sec) of the vertebrobasilar system using a rapid sequential angiography recording in a 5&#176; left-anterior oblique position.</li>
	<li>Infiltrate the area around the right common femoral artery with local anesthetics (Lidocaine 1%, maximal 6 mg/kg). Make a small inguinal skin incision. Using the microscope for visualization, dissect the common femoral artery and secure with a proximal and distal ligature (4-0 polyfilament sutures) around the exposed end.</li>
	<li>After arteriotomy, cannulate the femoral artery with a 5-F sheath. Flush the side port of the sheath with saline.</li>
	<li>Advance a 5-F catheter into the brachiocephalic artery through the sheath under fluoroscopy. Create a road map, then advance a guide wire to the vertebrobasilar system. Inject a bolus of non-ionic Iopamidol (0.6 ml/kg, 5 ml/sec in 2 sec) for DSA as described in step 3.2.</li>
</ol>
4. Rotation to Prone Position
<ol>
	<li>Following baseline DSA, reposition the rabbit from supine to prone position. Be careful not to manipulate or relocate the position of the intra-arterial catheters.</li>
	<li>Position the head in a head-holder at a 30&#176; angle, oriented head down.</li>
</ol>
5. Cisterna Magna Puncture
<ol>
	<li>Disinfect the skin over the head and neck with Povidone-iodine 3 times for 1 min each, and cover the surgical area with sterile sheets.</li>
	<li>Insert a 22 G x 40 mm pediatric spinal access needle transcutaneously into the cisterna magna without any prior skin incision or muscle displacement.</li>
	<li>Confirm that the animal is fully anesthetized by ensuring a lack of toe-pinch response before sliding the needle down along the bony external occipital protuberance until a gap is detected; do not push the needle any further.</li>
	<li>Confirm the correct positioning of the needle by observing spontaneous dripping of cerebrospinal fluid with the rabbit&#8217;s head tilted down at a 20&#176; angle for a few min.</li>
</ol>
6. Installation of Intracranial Pressure and Cerebral Blood Flow Monitoring
<ol>
	<li>After midline skin and galea incision, insert a small surgical retractor.</li>
	<li>Make three round osteotomies (2 mm diameter) using a high-speed microdrill in the frontal part of the skull according to the outer skull landmarks (Figure 1)9, i.e., over the olfactory bulb and bilateral frontal for placement of a neuromonitoring device if necessary. Use a millimeter scale ruler to determine the coordinates for burr hole placement as follows: Intracranial pressure (ICP) monitoring in the midpupillary line, one to two mm from the midsagittal line; intraparenchymal laser-Doppler probes four to five mm anterior and lateral to the bregma (Figure 1).</li>
	<li>Visualize the dura, and perform meticulous hemostasis: use bone wax for bone hemostasis by virtue of its tamponade action and perform local hemostasis using bipolar coagulation of the dura.</li>
	<li>Place the intraparenchymal intracranial pressure (ICP) monitor tip into the right olfactory bulb to a depth of 2 mm and then calibrate.</li>
	<li>Place the two laser-Doppler flowmetry fine needle probes using an external clamp system and insert them into the corresponding burr holes in both the right and left frontal hemispheres lateral to the ventricular system, i.e., in the midline to avoid interference with cerebrospinal fluid. Place needle probes to a depth of 2.5 mm.</li>
	<li>After placement of the neuromonitoring probes, seal all burr holes with a thick plug of bone wax in order to keep the cranium fluid-tight.</li>
	<li>Measure baseline parameters of mean arterial blood pressure (MAP), ICP and cerebral blood flow (CBF)using a multiparameter monitor and four channel laser-Doppler tissue blood perfusion monitor.</li>
</ol>
7. Shunt Induction
<ol>
	<li>Connect the spinal access needle in the cisterna magna to the previously catheterized subclavian artery via blood-filled pressure monitoring tubing. Use the 3-way stopcock for blood pressure measurement and as blood sampling port. 
	NOTE: The severity of SAH depends on the amount of blood, and can be roughly estimated by the extensiveness of subarachnoid clots at the time of brain harvest5,11.</li>
	<li>Continuously monitor MAP, HR, ECG, respiratory rate and end-tidal CO2 at a sampling rate of 1 Hz from 6 min before until at least 20 min after SAH.</li>
	<li>Confirm that the animal is fully anesthetized by ensuring a lack of toe pinch response before opening the shunt connection between the subclavian artery and the cisterna magna to induce SAH by the pressure gradient. 
	NOTE: A controlled SAH can be achieved by closing the shunt at any time point (e.g., at a desired level of ICP).</li>
	<li>Record steady state values during a time period of approximately 15 min.</li>
	<li>After ICP reaches its peak, keep the spinal access needle in place until ICP returns to a steady state close to baseline values. If the ICP plateau is maintained for more than 10 sec or if ICP decreases spontaneously, close the shunt.</li>
	<li>Remove CBF fine-needle probes and ICP probe, plug burr holes with bone wax, remove all catheters (including subclavian catheter, since catheter manipulation with consecutive bleeding is associated with high morbidity and mortality, and increases infection rate), perform rigorous wound irrigation with Neomycin sulfate and suture the skin.</li>
</ol>
8. Postoperative Management
<ol>
	<li>The procedure lasts for about 2 hr. Due to the half-lives of ketamine and xylazine, the recovery time of the animal is quite short -- approximately 1 hr. The animals are kept under a heating lamp during recovery. Additional fluids are not provided. During this initial postoperative recovery phase, apply buprenorphine 0.02 mg/kg s.c. every 8 hr for 24 hr.</li>
	<li>Apply transdermal fentanyl matrix patches releasing 12.5 &#956;g/hr in the shaved neck region of the animals for effective analgesia over the next 72 hr.</li>
	<li>Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
	<li>Do not return an animal that has undergone surgery in the company of other animals until fully recovered.</li>
</ol>
9. Follow up Digital Subtraction Angiography to Assess DCVS at Day 3
<ol>
	<li>Perform Steps 1.1 to 3.6 as described above.</li>
	<li>Euthanize the animals by intraarterial bolus injection of sodium thiopenthal (40 mg/kg) (Pentothal, Ospedalia AG, H&#252;nenberg, Switzerland). In cases where histology and immunohistochemistry is needed, perform an intracardiac perfusion-fixation at RT at a perfusion pressure of 100 cm H2O.</li>
</ol>

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None. The authors have no financial or commercial interest in any of the drugs, materials, or equipment used. No specific funding was received for this work. The authors are solely responsible for the design and conduct of the presented study and report no conflict of interest concerning the materials and methods used in this study or the findings specified in the paper. They confirm the adherence of ethical standards. The study was performed in accordance with the National Institutes of Health guidelines for the care and use of experimental animals and with the approval of the Animal Care Committee of the Canton of Bern, Switzerland (approval #109/07 and #107/09).

The shunt model produces pathology similar to that observed in humans after acute SAH3,8,10. It has been suggested that EBI may exacerbate, maintain and even trigger DCVS12, and as such this model may aid in investigating both the early and late DCVS phases, including EBI and DCVS interactions following SAH. In particular, repeatable in vivo DCVS monitoring techniques including DSA13, computed tomography angiography14, and transcranial Doppler15 are more rea...

Aneurysmal subarachnoid hemorrhage (SAH) is one of the most life threatening neuropathological conditions, frequently leading to permanent neurological damage or death1. Past research has focused on delayed cerebral vasospasm (DCVS) as the primary etiology of neurological deficits associated with SAH2. However, the generally poor clinical outcomes of patients suffering from SAH after treatment of vasospasm has led to an expansion of the research focus to include the effects of early brain injury (EBI) after SAH3. Greater understanding of the significance of both EBI and DCVS in contributing to poor clinical outcomes after SAH is essential for the development of more effective therapeutic strategies.
Up to now, single and double autologous blood injection into the cisterna magna has been the standard method for SAH induction for the study of DCVS2-6. Though commonly used in previous studies, this model most likely does not reproduce the neuropathological key changes associated with SAH induced EBI7. In contrast, endovascular perforation is known to produce severe acute pathophysiological changes that partially mimic the symptoms of EBI7.
This report describes a novel rabbit model of SAH designed to enable investigation of both EBI and DCVS, thereby allowing more accurate characterization of SAH-induced pathology8-10. With the described technique, the standard cisterna magna model is adapted by connecting the arterial system of the subclavian artery and the cisterna magna via an extracorporeal shunt. The blood flow is thereby linked to the rabbit&#8217;s physiology and driven by the pressure gradient between the arterial blood and intracranial pressure. The bleeding stops when intracerebral pressure (ICP) equals diastolic blood pressure and the blood in the shunt system coagulates. Utilizing the host&#8217;s physiology reduces examiner-dependent SAH induction, leading to a more consistent model of SAH that reliably produces both EBI and DCVS phenotypes3,8-10.

<table border="0" cellpadding="0" cellspacing="0" width="905">
	<tbody>
		<tr height="34">
			<td height="34" style="height:35px;width:279px;">Name</td>
			<td style="width:236px;">Company</td>
			<td style="width:223px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:279px;">Equipment</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">operation microscope</td>
			<td style="width:236px;">Zeiss, Jena, Germany</td>
			<td colspan="2">Zeiss, OPMI-MD surgical microscope</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:279px;">surgical equipment</td>
			<td style="width:236px;">B. Braun, Germany</td>
			<td style="width:223px;">forceps medical n&#176;5, vessel sciccors 8cm, microclip 4mm</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">respirator</td>
			<td style="width:236px;">Hugo Sachs</td>
			<td style="width:223px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">hair clipper</td>
			<td style="width:236px;">3M Surgical Clipper&#160;</td>
			<td style="width:223px;">&#160;Starter Kit 9667A</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">body warm plate</td>
			<td style="width:236px;">FHC</td>
			<td style="width:223px;"></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:279px;">blood gas analyzer</td>
			<td style="width:236px;">Radiometer, Copenhagen, Denmark</td>
			<td style="width:223px;">ABL 725</td>
			<td></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:279px;">cardiac monitoring</td>
			<td style="width:236px;">Camino Multi-Parameter Monitor, Integra, Plainsboro, NJ, US</td>
			<td style="width:223px;">AP-05</td>
			<td></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;">software analysis</td>
			<td style="width:236px;">BIOPAC Systems, Inc., Goleta, CA, USA</td>
			<td style="width:223px;">Biopac MP100 and acqKnowledge software,version 3.8.1</td>
			<td></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;">software analysis</td>
			<td style="width:236px;">ImagePro Discovery, MediaCybernetics, Silver Spring, MD, USA</td>
			<td style="width:223px;">image-Pro Plus version&#160;</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">angiography apparatus</td>
			<td style="width:236px;">DFP 2000 A-Toshiba</td>
			<td style="width:223px;">MIIXR0001EAA</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">ICP monitor</td>
			<td style="width:236px;">Camino Laboratories, San Diego, CA, USA</td>
			<td style="width:223px;">ICP monitor, Model 110-4B</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">blood flow monitor</td>
			<td style="width:236px;">Oxford Optronix Ltd., Oxford, UK</td>
			<td style="width:223px;">CAL KIT microsphere solution</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">laser-Doppler flowmetry fine needle probes&#160;</td>
			<td style="width:236px;">Oxford Optronix Ltd., Oxford, UK</td>
			<td style="width:223px;">MNP110XP, 0.48 mm diameter</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">pressure tube</td>
			<td>B. Braun, Germay</td>
			<td style="width:223px;">PE 1.0 mm &#215; 2.0 mm</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">anesthesia monitor</td>
			<td>GE Medical Systems, Switzerland&#160;</td>
			<td style="width:223px;">Datex S5 Monitor</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:279px;">Material</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">20 G vascular catheter</td>
			<td style="width:236px;">Smiths Medical</td>
			<td style="width:223px;">Jelco i.v. catheter, REF 4057</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">5.5F three-lumen central venous catheter&#160;</td>
			<td style="width:236px;">Connectors, Tagelswangen, Switzerland</td>
			<td>silicone catheter STH-C040</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">22Gx40mm needle&#160;</td>
			<td style="width:236px;">Emergo Group Inc., Netherlands</td>
			<td style="width:223px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">high-speed microdrill</td>
			<td style="width:236px;">Stryker, Solothurn, Switzerland</td>
			<td>5400-15&#160;</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">bone wax</td>
			<td style="width:236px;">Ethicon, Johnson &#38; Johnson,NJ, USA</td>
			<td style="width:223px;">ETHW31G</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">bipolar forceps</td>
			<td style="width:236px;">Aesculap, Inc., PA, US</td>
			<td>US349SP&#160;</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">Ketamin</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Xylazine</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">Buprenorphine</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fentanyl</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">transdermal fentanyl matrix patches&#160;</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Lidocaine 1%&#160;</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">4% papaverin HCl&#160;</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Neomycin sulfate&#160;</td>
			<td style="width:236px;">Research Organics Inc., OH, USA</td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Povidone-iodine&#160;</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">0.9% sodium chloride</td>
			<td style="width:236px;"></td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">Iopamidol&#160;</td>
			<td style="width:236px;">Abott Laboratories, IL, USA</td>
			<td style="width:223px;"></td>
			<td>Any generic product</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">3-0 resorbable suture</td>
			<td style="width:236px;">Ethicon Inc., USA</td>
			<td>VCP824G</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">5-0 non absorbable suture</td>
			<td style="width:236px;">Ethicon Inc., USA</td>
			<td>8618G</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:279px;">4-0 polyfilament sutures</td>
			<td style="width:236px;">Ethicon Inc., USA</td>
			<td style="width:223px;">VCP284G</td>
			<td></td>
		</tr>
	</tbody>
</table>

the rabbit blood-shunt model for the study of acute and late sequelae of subarachnoid hemorrhage: technical aspects

Early brain injury and delayed cerebral vasospasm both contribute to unfavorable outcomes after subarachnoid hemorrhage (SAH). Reproducible and controllable animal models that simulate both conditions are presently uncommon. Therefore, new models are needed in order to mimic human pathophysiological conditions resulting from SAH.
This report describes the technical nuances of a rabbit blood-shunt SAH model that enables control of intracerebral pressure (ICP). An extracorporeal shunt is placed between the arterial system and the subarachnoid space, which enables examiner-independent SAH in a closed cranium. Step-by-step procedural instructions and necessary equipment are described, as well as technical considerations to produce the model with minimal mortality and morbidity. Important details required for successful surgical creation of this robust, simple and consistent ICP-controlled SAH rabbit model are described.

The rabbit blood shunt model of SAH described in this report produces EBI in the hippocampus (Figure 2A, B), basal cortex (Figure 2A, B), and cerebral vasculature (Figure 2C) as early as 24 hr after injury and shows a characteristic blood distribution (Figure 2D)8. In addition, the model triggers moderate to severe degrees of DCVS on day three after SAH induction (Figure 3)10. The mortality rate is ...

Watch this Scientific Journal Video about The Rabbit Blood-shunt Model for the Study of Acute and Late Sequelae of Subarachnoid Hemorrhage: Technical Aspects at JoVE.com

The Rabbit Blood-shunt Model for the Study of Acute and Late Sequelae of Subarachnoid Hemorrhage: Technical Aspects

The experimental intracranial pressure-controlled blood shunt subarachnoid hemorrhage (SAH) model in the rabbit combines the standard procedures &#8212; subclavian artery cannulation and transcutaneous cisterna magna puncture, which enables close mimicking of human pathophysiological conditions after SAH. We present step-by-step instructions and discuss key surgical points for successful experimental SAH creation.

Early brain injury and delayed cerebral vasospasm both contribute to unfavorable outcomes after subarachnoid hemorrhage (SAH). Reproducible and ...

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13.5K vistas.  University and Bern University Hospital (Inselspital). The experimental intracranial pressure-controlled blood shunt subarachnoid hemorrhage (SAH) model in the rabbit combines the standard procedures — subclavian artery cannulation and transcutaneous cisterna magna puncture, which enables close mimicking of human pathophysiological conditions after SAH. We present step-by-step instructions and discuss key surgical points for successful experimental SAH creation.

Video: The Rabbit Blood-shunt Model for the Study of Acute and Late Sequelae of Subarachnoid Hemorrhage: Technical Aspects

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 ...

Biomarkers in an Animal Model for Revealing Neural, Hematologic, and Behavioral Correlates of PTSD

Identification of biomarkers representing the evolution of the pathophysiology of Post Traumatic Stress Disorder (PTSD) is vitally important, not only for objective diagnosis but also for the evaluation of therapeutic efficacy and resilience to trauma. Ongoing research is directed at identifying molecular biomarkers for PTSD, including traumatic stress induced proteins, transcriptomes, genomic variances and genetic modulators, using biologic samples from subjects' blood, saliva, urine, and postmortem brain tissues. However, the correlation of these biomarker molecules in peripheral or postmortem samples to altered brain functions associated with psychiatric symptoms in PTSD remains unresolved. Here, we present an animal model of PTSD in which both peripheral blood and central brain biomarkers, as well as behavioral phenotype, can be collected and measured, thus providing the needed correlation of the central biomarkers of PTSD, which are mechanistic and pathognomonic but cannot be collected from people, with the peripheral biomarkers and behavioral phenotypes, which can.
Our animal model of PTSD employs restraint and tail shocks repeated for three continuous days - the inescapable tail-shock model (ITS) in rats. This ITS model mimics the pathophysiology of PTSD 17, 7, 4, 10. We and others have verified that the ITS model induces behavioral and neurobiological alterations similar to those found in PTSD subjects 17, 7, 10, 9. Specifically, these stressed rats exhibit (1) a delayed and exaggerated startle response appearing several days after stressor cessation, which given the compressed time scale of the rat's life compared to a humans, corresponds to the one to three months delay of symptoms in PTSD patients (DSM-IV-TR PTSD Criterian D/E 13), (2) enhanced plasma corticosterone (CORT) for several days, indicating compromise of the hypothalamopituitary axis (HPA), and (3) retarded body weight gain after stressor cessation, indicating dysfunction of metabolic regulation.
The experimental paradigms employed for this model are: (1) a learned helplessness paradigm in the rat assayed by measurement of acoustic startle response (ASR) and a charting of body mass; (2) microdissection of the rat brain into regions and nuclei; (3) enzyme-linked immunosorbent assay (ELISA) for blood levels of CORT; (4) a gene expression microarray plus related bioinformatics tools 18. This microarray, dubbed rMNChip, focuses on mitochondrial and mitochondria-related nuclear genes in the rat so as to specifically address the neuronal bioenergetics hypothesized to be involved in PTSD.

We describe a rat model of post traumatic stress disorder (PTSD) that reveals the persistent alterations in neuroendocrine function and the delayed long-term, exaggerated fear response, characteristic of PTSD patients. The animal model and methods described here are useful for correlating biomarkers in brain nuclei, which are mechanistic but cannot be measured in patients, with biomarkers in peripheral white blood cells, which can.

Identification of biomarkers representing the evolution of the pathophysiology of Post Traumatic Stress Disorder (PTSD) is vitally important, not only for ...

NADH Fluorescence Imaging of Isolated Biventricular Working Rabbit Hearts

Since its inception by Langendorff1, the isolated perfused heart remains a prominent tool for studying cardiac physiology2. However, it is not well-suited for studies of cardiac metabolism, which require the heart to perform work within the context of physiologic preload and afterload pressures. Neely introduced modifications to the Langendorff technique to establish appropriate left ventricular (LV) preload and afterload pressures3. The model is known as the isolated LV working heart model and has been used extensively to study LV performance and metabolism4-6. This model, however, does not provide a properly loaded right ventricle (RV). Demmy et al. first reported a biventricular model as a modification of the LV working heart model7, 8. They found that stroke volume, cardiac output, and pressure development improved in hearts converted from working LV mode to biventricular working mode8. A properly loaded RV also diminishes abnormal pressure gradients across the septum to improve septal function. Biventricular working hearts have been shown to maintain aortic output, pulmonary flow, mean aortic pressure, heart rate, and myocardial ATP levels for up to 3 hours8.
When studying the metabolic effects of myocardial injury, such as ischemia, it is often necessary to identify the location of the affected tissue. This can be done by imaging the fluorescence of NADH (the reduced form of nicotinamide adenine dinucleotide)9-11, a coenzyme found in large quantities in the mitochondria. NADH fluorescence (fNADH) displays a near linearly inverse relationship with local oxygen concentration12 and provides a measure of mitochondrial redox state13. fNADH imaging during hypoxic and ischemic conditions has been used as a dye-free method to identify hypoxic regions14, 15 and to monitor the progression of hypoxic conditions over time10.
The objective of the method is to monitor the mitochondrial redox state of biventricular working hearts during protocols that alter the rate of myocyte metabolism or induce hypoxia or create a combination of the two. Hearts from New Zealand white rabbits were connected to a biventricular working heart system (Hugo Sachs Elektronik) and perfused with modified Krebs-Henseleit solution16 at 37 &deg;C. Aortic, LV, pulmonary artery, and left &amp; right atrial pressures were recorded. Electrical activity was measured using a monophasic action potential electrode. To image fNADH, light from a mercury lamp was filtered (350&plusmn;25 nm) and used to illuminate the epicardium. Emitted light was filtered (460&plusmn;20 nm) and imaged using a CCD camera. Changes in the epicardial fNADH of biventricular working hearts during different pacing rates are presented. The combination of the heart model and fNADH imaging provides a new and valuable experimental tool for studying acute cardiac pathologies within the context of realistic physiological conditions.

The objective is to monitor the mitochondrial redox state of isolated hearts within the context of physiologic preload and afterload pressures. A biventricular working rabbit heart model is presented. High spatiotemporal resolution fluorescence imaging of NADH is used to monitor the mitochondrial redox state of epicardial tissue.

Since its inception by Langendorff1, the isolated perfused heart remains a prominent tool for studying cardiac physiology2. However, it is not well-suited ...

A Low Mortality Rat Model to Assess Delayed Cerebral Vasospasm After Experimental Subarachnoid Hemorrhage

Objective: To characterize and establish a reproducible model that demonstrates delayed cerebral vasospasm after aneurysmal subarachnoid hemorrhage (SAH) in rats, in order to identify the initiating events, pathophysiological changes and potential targets for treatment.
Methods: Twenty-eight male Sprague-Dawley rats (250 - 300 g) were arbitrarily assigned to one of two groups - SAH or saline control. Rat subarachnoid hemorrhage in the SAH group (n=15) was induced by double injection of autologous blood, 48 hr apart, into the cisterna magna. Similarly, normal saline (n=13) was injected into the cisterna magna of the saline control group. Rats were sacrificed on day five after the second blood injection and the brains were preserved for histological analysis. The degree of vasospasm was measured using sections of the basilar artery, by measuring the internal luminal cross sectional area using NIH Image-J software. The significance was tested using Tukey/Kramer's statistical analysis.
Results: After analysis of histological sections, basilar artery luminal cross sectional area were smaller in the SAH than in the saline group, consistent with cerebral vasospasm in the former group. In the SAH group, basilar artery internal area (.056 &mu;m &plusmn; 3) were significantly smaller from vasospasm five days after the second blood injection (seven days after the initial blood injection), compared to the saline control group with internal area (.069 &plusmn; 3; p=0.004). There were no mortalities from cerebral vasospasm.
Conclusion: The rat double SAH model induces a mild, survivable, basilar artery vasospasm that can be used to study the pathophysiological mechanisms of cerebral vasospasm in a small animal model. A low and acceptable mortality rate is a significant criterion to be satisfied for an ideal SAH animal model so that the mechanisms of vasospasm can be elucidated 7, 8. Further modifications of the model can be made to adjust for increased severity of vasospasm and neurological exams.

Aneurysmal subarachnoid hemorrhage (SAH) is bleeding that occurs into the subarachnoid space when an aneurysm ruptures. While the morbidity and mortality from this event has been on a decline due to improved treatment approaches, the risk of vasospasm after subarachnoid hemorrhage continues to be the same as it was several years ago. The importance of establishing a comprehensive and reproducible animal model to identify initiating events of cerebral vasospasm has been the focus of research since the first use of rats in an experimental vasospasm model in 1979 by Barry et al. Early work in rats demonstrated that a single injection of autologous blood into the cisterna magna led to acute (within minutes) but not delayed cerebral vasospasm 3, 6, 14. Here we characterize a low mortality SAH rat model that results in reproducible delayed vasospasm.

Objective: To characterize and establish a reproducible model that demonstrates delayed cerebral vasospasm after aneurysmal subarachnoid hemorrhage (SAH) ...

Human Neuroendocrine Tumor Cell Lines as a Three-Dimensional Model for the Study of Human Neuroendocrine Tumor Therapy

Neuroendocrine tumors (NETs) are rare tumors, with an incidence of two per 100,&#x200A;000 individuals per year, and they account for 0.5% of all human malignancies.1 Other than surgery for the minority of patients who present with localized disease, there is little or no survival benefit of systemic therapy. Therefore, there is a great need to better understand the biology of NETs, and in particular define new therapeutic targets for patients with nonresectable or metastatic neuroendocrine tumors. 3D cell culture is becoming a popular method for drug screening due to its relevance in modeling the in vivo tumor tissue organization and microenvironment.2,3 The 3D multicellular spheroids could provide valuable information in a more timely and less expensive manner than directly proceeding from 2D cell culture experiments to animal (murine) models.
To facilitate the discovery of new therapeutics for NET patients, we have developed an in vitro 3D multicellular spheroids model using the human NET cell lines. The NET cells are plated in a non-adhesive agarose-coated 24-well plate and incubated under physiological conditions (5% CO2, 37 &deg;C) with a very slow agitation for 16-24 hr after plating. The cells form multicellular spheroids starting on the 3rd or 4th day. The spheroids become more spherical by the 6th day, at which point the drug treatments are initiated. The efficacy of the drug treatments on the NET spheroids is monitored based on the morphology, shape and size of the spheroids with a phase-contrast light microscope. The size of the spheroids is estimated automatically using a custom-developed MATLAB program based on an active contour algorithm. Further, we demonstrate a simple method to process the HistoGel embedding on these 3D spheroids, allowing the use of standard histological and immunohistochemical techniques. 
This is the first report on generating 3D spheroids using NET cell lines to examine the effect of therapeutic drugs. We have also performed histology on these 3D spheroids, and displayed an example of a single drug's effect on growth and proliferation of the NET spheroids. Our results support that the NET spheroids are valuable for further studies of NET biology and drug development.

We present a simple agarose overlay platform to grow 3D multicellular spheroids using neuroendocrine cancer cell lines. This method provides a very convenient way to examine the effect of therapeutic drugs on the neuroendocrine tumor cells. It could also help us establish human neuroendocrine tumor spheroids for cancer therapy.

Neuroendocrine tumors (NETs) are rare tumors, with an incidence of two per 100,&#x200A;000 individuals per year, and they account for 0.5% of all human ...

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

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

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

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

Technical Aspects of the Mouse Aortocaval Fistula

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta and inferior vena cava (IVC) are exposed. The proximal infrarenal aorta and the distal aorta are dissected for clamp placement and needle puncture, respectively. Special attention is paid to avoid dissection between the aorta and the IVC. After clamping the aorta, a 25 G needle is used to puncture both walls of the aorta into the IVC. The surrounding connective tissue is used for hemostatic compression. Successful creation of the AVF will show pulsatile arterial blood flow in the IVC. Further confirmation of successful AVF can be achieved by post-operative Doppler ultrasound.

The goal is to produce an arteriovenous fistula that is simple and reproducible. This method does not use sutures or glue adhesive. Therefore the samples can be used with the least amount of foreign materials for analysis.

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta ...

A Murine Model of Subarachnoid Hemorrhage

In this video publication a standardized mouse model of subarachnoid hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of Willis perforation (CWp) and proven by intracranial pressure (ICP) monitoring. Thereby a homogenous blood distribution in subarachnoid spaces surrounding the arterial circulation and cerebellar fissures is achieved. Animal physiology is maintained by intubation, mechanical ventilation, and continuous on-line monitoring of various physiological and cardiovascular parameters: body temperature, systemic blood pressure, heart rate, and hemoglobin saturation. Thereby the cerebral perfusion pressure can be tightly monitored resulting in a less variable volume of extravasated blood. This allows a better standardization of endovascular filament perforation in mice and makes the whole model highly reproducible. Thus it is readily available for pharmacological and pathophysiological studies in wild type and genetically altered mice.

A standardized mouse model of subarachnoid hemorrhage by intraluminal Circle of Willis perforation is described. Vessel perforation and subarachnoid bleeding are monitored by intracranial pressure monitoring. In addition various vital parameters are recorded and controlled to maintain physiologic conditions.

In this video publication a standardized mouse model of subarachnoid  hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of  Willis ...

A Murine Model of Arterial Restenosis: Technical Aspects of Femoral Wire Injury

Cardiovascular disease caused by atherosclerosis is the leading cause of death in the developed world. Narrowing of the vessel lumen, due to atherosclerotic plaque development or the rupturing of established plaques, interrupts normal blood flow leading to various morbidities such as myocardial infarction and stroke. In the clinic endovascular procedures such as angioplasty are commonly performed to reopen the lumen. However, these treatments inevitably damage the vessel wall as well as the vascular endothelium, triggering an excessive healing response and the development of a neointimal plaque that extends into the lumen causing vessel restenosis (re-narrowing). Restenosis remains a major cause of failure of endovascular treatments for atherosclerosis. Thus, preclinical animal models of restenosis are vitally important for investigating the pathophysiological mechanisms as well as translational approaches to vascular interventions. Among several murine experimental models, femoral artery wire injury is widely accepted as the most suitable for studies of post-angioplasty restenosis because it closely resembles the angioplasty procedure that injures both endothelium and vessel wall. However, many researchers have difficulty utilizing this model due to its high degree of technical difficulty. This is primarily because a metal wire needs to be inserted into the femoral artery, which is approximately three times thinner than the wire, to generate sufficient injury to induce prominent neointima. Here, we describe the essential surgical details to effectively overcome the major technical difficulties of this model. By following the presented procedures, performing the mouse femoral artery wire injury becomes easier. Once familiarized, the whole procedure can be completed within 20 min.

The mouse femoral artery wire injury model of restenosis is technically challenging. In this protocol we show the key technical details essential for successfully performing wire injury to induce consistent&#160;neointima for studies of restenosis.

Cardiovascular disease caused by atherosclerosis is the leading cause of death in the developed world. Narrowing of the vessel lumen, due to ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...