Name: the rabbit blood-shunt model for the study of acute and late sequelae of subarachnoid hemorrhage: technical aspects
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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><tbody><tr><td>Equipment</td><td></td><td></td><td></td></tr><tr><td>Operation microscope</td><td>Zeiss, Jena, Germany</td><td></td><td>Zeiss, OPMI-MD surgical microscope</td></tr><tr><td>Surgical equipment</td><td>B. Braun, Germany</td><td></td><td>Forceps medical no. 5; vessel scissors 8 cm; microclip 4 mm</td></tr><tr><td>Respirator</td><td>Hugo Sachs</td><td></td><td></td></tr><tr><td>Hair clipper</td><td>3M Surgical Clipper&amp;nbsp;</td><td></td><td>Starter Kit 9667A</td></tr><tr><td>Body warm plate</td><td>FHC</td><td></td><td></td></tr><tr><td>Blood gas analyzer</td><td>Radiometer, Copenhagen, Denmark</td><td>ABL 725</td><td></td></tr><tr><td>Cardiac monitoring</td><td>Camino Multi-Parameter Monitor, Integra, Plainsboro, NJ, US</td><td>AP-05</td><td></td></tr><tr><td>Software analysis</td><td>BIOPAC Systems, Inc., Goleta, CA, USA</td><td></td><td>Biopac MP100 and acqKnowledge software,version 3.8.1</td></tr><tr><td>Software analysis</td><td>ImagePro Discovery, MediaCybernetics, Silver Spring, MD, USA</td><td></td><td>Image-Pro Plus version&amp;nbsp;</td></tr><tr><td>Angiography apparatus</td><td>DFP 2000 A-Toshiba</td><td>MIIXR0001EAA</td><td></td></tr><tr><td>ICP monitor</td><td>Camino Laboratories, San Diego, CA, USA</td><td></td><td>ICP monitor, Model 110-4B</td></tr><tr><td>Blood flow monitor</td><td>Oxford Optronix Ltd., Oxford, UK</td><td></td><td>CAL KIT microsphere solution</td></tr><tr><td>Laser-Doppler flowmetry fine needle probes</td><td>Oxford Optronix Ltd., Oxford, UK</td><td></td><td>MNP110XP, 0.48 mm diameter</td></tr><tr><td>Pressure tube</td><td>B. Braun, Germay</td><td></td><td>PE 1.0 mm &amp;times; 2.0 mm</td></tr><tr><td>Anesthesia monitor</td><td>GE Medical Systems, Switzerland&amp;nbsp;</td><td></td><td>Datex S5 Monitor</td></tr><tr><td>Material</td><td></td><td></td><td></td></tr><tr><td>20 G vascular catheter</td><td>Smiths Medical</td><td></td><td>Jelco i.v. catheter, REF 4057</td></tr><tr><td>5.5 F three-lumen central venous catheter&amp;nbsp;</td><td>Connectors, Tagelswangen, Switzerland</td><td></td><td>Silicone catheter STH-C040</td></tr><tr><td>22 G x 40 mm needle&amp;nbsp;</td><td>Emergo Group Inc., Netherlands</td><td></td><td></td></tr><tr><td>High-speed microdrill</td><td>Stryker, Solothurn, Switzerland</td><td>5400-15&amp;nbsp;</td><td></td></tr><tr><td>Bone wax</td><td>Ethicon, Johnson &amp;amp; Johnson,NJ, USA</td><td>ETHW31G</td><td></td></tr><tr><td>Bipolar forceps</td><td>Aesculap, Inc., PA, US</td><td>US349SP&amp;nbsp;</td><td></td></tr><tr><td>Ketamin</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Xylazine</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Buprenorphine</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Fentanyl</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Transdermal fentanyl matrix patches&amp;nbsp;</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Lidocaine 1%&amp;nbsp;</td><td></td><td></td><td>Any generic product</td></tr><tr><td>4% papaverin HCl&amp;nbsp;</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Neomycin sulfate&amp;nbsp;</td><td>Research Organics Inc., OH, USA</td><td></td><td>Any generic product</td></tr><tr><td>Povidone-iodine&amp;nbsp;</td><td></td><td></td><td>Any generic product</td></tr><tr><td>0.9% sodium chloride</td><td></td><td></td><td>Any generic product</td></tr><tr><td>Iopamidol&amp;nbsp;</td><td>Abott Laboratories, IL, USA</td><td></td><td>Any generic product</td></tr><tr><td>3-0 resorbable suture</td><td>Ethicon Inc., USA</td><td>VCP824G</td><td></td></tr><tr><td>5-0 non absorbable suture</td><td>Ethicon Inc., USA</td><td>8618G</td><td></td></tr><tr><td>4-0 polyfilament sutures</td><td>Ethicon Inc., USA</td><td>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 ...

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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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Medicine

13.6K Views.  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

Autologous Blood Injection to Model Spontaneous Intracerebral Hemorrhage in Mice

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model. While ICH
accounts for 10-15% of all strokes, there remains no specific
effective therapy. The autologous blood injection model in mice
involves the stereotaxic injection of arterial blood into the basal
ganglia mimicking a spontaneous hypertensive hemorrhage in man. The
response to hemorrhage can then be studied in vivo and the
neurobehavioral deficits quantified, allowing for description of the
ensuing pathology and the testing of potential therapeutic agents. The
procedure described in this protocol uses the double injection
technique to minimize risk of blood reflux up the needle track, no
anticoagulants in the pumping system, and eliminates all dead space
and expandable tubing in the system.

The autologous blood injection model of intracerebral hemorrhage in mice described in this protocol uses the double injection technique to minimize risk of blood reflux up the needle track, no anticoagulants in the pumping system, and eliminates all dead space and expandable tubing in the system.

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model.  While ICH
accounts for 10-15% ...

Neuroscience

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) ...

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 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 ...

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% ...

Analysis of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage with High Frequency Transcranial Duplex Ultrasound

Cerebral vasospasm that occurs in the weeks after subarachnoid hemorrhage, a type of hemorrhagic stroke, contributes to delayed cerebral ischemia. A problem encountered in experimental studies using murine models of SAH is that methods for in vivo monitoring of cerebral vasospasm in mice are lacking. Here, we demonstrate the application of high frequency ultrasound to perform transcranial Duplex sonography examinations on mice. Using the method, the internal carotid arteries (ICA) could be identified. The blood flow velocities in the intracranial ICAs were accelerated significantly after induction of SAH, while blood flow velocities in the extracranial ICAs remained low, indicating cerebral vasospasm. In conclusion, the method demonstrated here allows functional, noninvasive in vivo monitoring of cerebral vasospasm in a murine SAH model.

The aim of this manuscript is to present a sonography-based method that allows in vivo imaging of blood flow in cerebral arteries in mice. We demonstrate its application to determine changes in blood flow velocities associated with vasospasm in murine models of subarachnoid hemorrhage (SAH).

Cerebral vasospasm that occurs in the weeks after subarachnoid hemorrhage, a type of hemorrhagic stroke, contributes to delayed cerebral ischemia. A ...

Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model

Despite advances in treatment over the last decades, subarachnoid hemorrhage (SAH) continues to carry a high burden of morbidity and mortality, largely afflicting a fairly young population. Several animal models of SAH have been developed to investigate the pathophysiological mechanisms behind SAH and to test pharmacological interventions. The pre-chiasmatic, single injection model in the rat presented in this article is an experimental model of SAH with a predetermined blood volume. Briefly, the animal is anesthetized, intubated, and kept under mechanical ventilation. Temperature is regulated with a heating pad. A catheter is placed in the tail artery, enabling continuous blood pressure measurement as well as blood sampling. The atlantooccipital membrane is incised and a catheter for pressure recording is placed in the cisterna magna to enable intracerebral pressure measurement. This catheter can also be used for intrathecal therapeutic interventions. The rat is placed in a stereotaxic frame, a burr hole is drilled anteriorly to the bregma, and a catheter is inserted through the burr hole and placed just anterior to the optic chiasm. Autologous blood (0.3 mL) is withdrawn from the tail catheter and manually injected. This results in a rise of intracerebral pressure and a decrease of cerebral blood flow. The animal is kept sedated for 30 min and given subcutaneous saline and analgesics. The animal is extubated and returned to its cage. The pre-chiasmatic model has a high reproducibility rate and limited variation between animals due to the pre-determined blood volume. It mimics SAH in humans making it a relevant model for SAH research.

Subarachnoid hemorrhage continues to carry a high burden of mortality and morbidity in man. To facilitate further research into the condition and its pathophysiology, a pre-chiasmatic, single injection model is presented.

Despite advances in treatment over the last decades, subarachnoid hemorrhage (SAH) continues to carry a high burden of morbidity and mortality, largely ...

Endovascular Perforation Model for Subarachnoid Hemorrhage Combined with Magnetic Resonance Imaging (MRI)

The endovascular filament perforation model to mimic subarachnoid hemorrhage (SAH) is a commonly used model - however, the technique can cause a high mortality rate as well as an uncontrollable volume of SAH and other intracranial complications such as stroke or intracranial hemorrhage. In this protocol, a standardized SAH mouse model is presented, induced by endovascular filament perforation, combined with magnetic resonance imaging (MRI) 24 h after operation to ensure the correct bleeding site and exclude other relevant intracranial pathologies. Briefly, C57BL/6J mice are anesthetized with an intraperitoneal ketamine/xylazine (70 mg/16 mg/kg body weight) injection and placed in a supine position. After midline neck incision, the common carotid artery (CCA) and carotid bifurcation are exposed, and a 5-0 non-absorbable monofilament polypropylene suture is inserted in a retrograde fashion into the external carotid artery (ECA) and advanced into the common carotid artery. Then, the filament is invaginated into the internal carotid artery (ICA) and pushed forward to perforate the anterior cerebral artery (ACA). After recovery from surgery, mice undergo a 7.0 T MRI 24 h later. The volume of bleeding can be quantified and graded via postoperative MRI, enabling a robust experimental SAH group with the option to perform further subgroup analyses based on blood quantity.

Endovascular Perforation Model for Subarachnoid Hemorrhage Combined with Magnetic Resonance Imaging MRI

Here we present a standardized SAH mouse model, induced by endovascular filament perforation, combined with magnetic resonance imaging (MRI) 24 h after operation to ensure the correct bleeding site and exclude other relevant intracranial pathologies.

The endovascular filament perforation model to mimic subarachnoid hemorrhage (SAH) is a commonly used model - however, the technique can cause a high ...

Establishing a Subarachnoid Hemorrhage Rat Model Induced by Autologous Blood Injection

To place the laser Doppler probe, make a 15-millimeter incision caudally in the midline, starting just anterior to the eyes. After removing the connective tissue and the muscles with forceps, use the end of a sterile cotton swab as a regime to identify the bregma and the coronal sutures.
After placing the armed retractor, place a 25-gauge spinal needle in the stereotaxic frame exactly on the bregma and note the position. Next, remove the needle from the bregma. Move the frame anteriorly by 6.5 millimeters, then replace the needle in the midline to mark the side of drilling.
Drill until the dura mater is identified below the bone. Using straight forceps, gently remove the bone fragments, then fill the cavity with bone wax. Next, 3 to 4 millimeters lateral to the right of the bregma and just anterior to the coronal suture for the laser Doppler, drill another hole taking care not to penetrate the dura mater.
Look for the vessels where the laser Doppler can measure the blood flow. Place the laser Doppler and check the values. A minimum value of 100 flux units is required. If the values remain acceptable after removing the microscope, add 1 drop of glue to fix the probe. Recheck to confirm whether the value is above 80 flux units.
To induce subarachnoid hemorrhage, insert the needle gently through the skull in the midline between the hemispheres until resistance is felt due to the base of the skull. Retract the needle by 1 millimeter to ensure correct placement just anteriorly to the optic chiasm. To ensure the most homogeneous result when injecting the blood, turn the needle clockwise by 90 degrees such that the needle tip points to the right. Then, remove the stiletto.
After a 15-minute equilibration, adjust the level of anesthesia to obtain a mean arterial blood pressure in the range 80 to 100 millimeters of mercury. Then, perform a blood gas analysis to confirm that the pH, partial pressure of carbon dioxide and partial pressure of oxygen are within physiological parameters.
Next, using a 1-milliliter syringe with a blunt 23-gauge needle, withdraw 500 microliters of blood from the tail catheter. To avoid injection of air, fill the dead space of the spinal needle chamber with blood. After adjusting the volume of the blood in the syringe to 300 microliters, connect the syringe to the spinal needle. Then, grasping firmly around the syringe, inject the blood manually to surpass mean arterial pressure. Observe a steep rise in the intracerebral pressure and a concurrent steep fall in the cerebral blood flow.

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

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