The current research is funded by the Solorz-Zak Research Foundation.

All surgical procedures were subjected to ethical review and approved by the government of Upper Bavaria (reference number: 55.2-1-54-2532.3-13-13 and -2532-136-11). Animals are male C57BL/6 mice with a body weight of approximately 25 g.
1. Animal Preparation
<ol>
	<li>Induce anesthesia by putting the mouse into a chamber. Flush the chamber with 5% isoflurane until the animal loses consciousness.</li>
	<li>Inject premixed anesthetics intraperitoneally: fentanyl (0.05 mg/kg), midazolam (5 mg/kg) and medetomidine (0.5 mg/kg). Check reflexes before and regularly during the procedure. Reinject one third of the initial amount hourly to maintain anesthesia.</li>
	<li>Intubate orotracheally with a tube made from a 20 G venous catheter11. For intubation fix the animal on a slanted platform (30&#176;), retract the tongue with bent forceps, visualize the vocal cords under an operation microscope and insert the tube into the trachea during inspiration.</li>
	<li>Place the mouse in a prone position and check the correct placement of the tube with a piece of cotton or a microcapnograph.</li>
	<li>Connect the intubation tube to the respirator. Ventilate the mouse with room air supplemented with 25% oxygen with a frequency of 180-220 breaths/min and a stroke volume of 200-250 &#956;l.</li>
	<li>Connect the intubation tube to the microcapnograph. Maintain the end-expiratory pCO2 at 30 mmHg by adjusting the ventilation frequency.</li>
	<li>Insert a rectal temperature probe and place the animal on a heating pad in order to maintain 37 &#176;C core body temperature.</li>
	<li>Apply the annular pulsoximeter sensor on the right hind paw.</li>
	<li>Open the skin above the skull with a pair of scissors. The incision should be approximately 0.5 cm long and between ear and eye.</li>
	<li>Dissect the left temporal muscle with a scalpel from the temporal bone.</li>
	<li>Glue the laser Doppler flowmeter (LDF) probe on the left temporal bone. Hold the probe in a fixed position until the glue hardens.</li>
	<li>Drill a hole of approximately 1.5 mm diameter with a dental drill into the left temporal bone. Cool the bone with saline to prevent heat damage.</li>
	<li>Insert the ICP probe into the cranial cavity. Push forward as dorsally as possible to prevent brain tissue damage and bleeding.</li>
	<li>If the probe is at the right position, fix, and seal it with cement. Let the cement dry for 5 min.</li>
	<li>Turn the mouse carefully to a supine position.</li>
	<li>For continuous blood pressure monitoring, catheterize the left femoral artery.</li>
	<li>Connect the femoral catheter to the blood pressure monitoring device.</li>
</ol>
2. SAH Induction
<ol>
	<li>Open the skin with a pair of scissors from sternum to chin (2 cm). Dissect the connective tissue bluntly and push the salivary glands aside.</li>
	<li>Expose the left common carotid artery (CCA) and mobilize it. Preserve the vagal nerve, which runs in the same connective tissue sheath as the CCA. Move cranial and expose and mobilize the ICA and the ECA using the same technique.</li>
	<li>Ligate the ECA as far cranially as possible.</li>
	<li>Prearrange two more ligations for the filament around the ECA.</li>
	<li>Occlude the CCA and the ICA temporarily with microclips. Position the microclips with a microclip applicator. Make sure the clips are applied correctly by gently pulling them back.</li>
	<li>Cut a hole for the filament insertion into the ECA with a vessel scissors.</li>
	<li>Insert a Prolene 5-0 filament with 12 mm length into the ECA.</li>
	<li>Close the insertion site with one prearranged ligation.</li>
	<li>Remove the microclips with a microclip applicator from the CCA and the ICA.</li>
	<li>Advance the filament with a forceps into the ICA until the ICP rises. A sudden rise of the ICP indicates bleeding induction.</li>
	<li>Withdraw the filament immediately and ligate the ECA by closing both prearranged ligations consecutively. This prevents bleeding out of the insertion site.</li>
	<li>Suture the skin wound.</li>
	<li>Monitor the physiologic parameters of the animal for another 20 min.</li>
	<li>Remove ICP and LDF probes and suture the skin wound.</li>
</ol>
3. End of Experiment
<ol>
	<li>Perfuse the animal transcardially with 20 ml of saline (room temperature) followed by 20 ml of 4% PFA in PBS (4 &#176;C).</li>
	<li>Dissect the brain out of the skull. Cut the skull in the midline and between the orbital cavities. Then peel the bone from the brain.</li>
	<li>Evaluate the blood distribution in the subarachnoid space.</li>
</ol>
4.&#160;Considerations in Case of Survival Surgery (not shown in the video)
<ol>
	<li>Inject Carprofen (4 mg/kg subcutaneously) for postoperative analgesia directly after anesthesia induction. During the postoperative observation period Carprofen (4 mg/kg subcutaneously) is injected every 24 hr.</li>
	<li>Instead of invasive blood pressure measurement make use of a noninvasive blood pressure monitoring system.</li>
	<li>For termination of anesthesia inject antagonizing agents subcutaneously: naloxone (1.2 mg/kg), flumazenil (0.5 mg/kg) and atipamezole (2.5 mg/kg).</li>
	<li>Extubate the animal.</li>
	<li>After regaining of reflexes put the animal into a preheated chamber. Keep the animal at approximately 32 &#176;C for 24 hr.</li>
	<li>Animals are checked regularly for spontaneous breathing and their general condition during the first hours after surgery. If the brain stem is affected animals have breathing problems and should be euthanized.&#160;</li>
	<li>Animals are checked daily for their general condition and body weight as well as for neurological and sensory deficits15.</li>
</ol>

<ol>
	<li>Cahill, J., Zhang, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subarachnoid+hemorrhage:+is+it+time+for+a+new+direction.">Subarachnoid hemorrhage: is it time for a new direction.</a> Stroke. 40, 86-87 (2009).</li><li>van Gijn, J., Kerr, R. S., Rinkel, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subarachnoid+haemorrhage.">Subarachnoid haemorrhage.</a> Lancet. 369, 306-318 (2007).</li><li>Lin, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+murine+model+of+subarachnoid+hemorrhage-induced+cerebral+vasospasm.">A murine model of subarachnoid hemorrhage-induced cerebral vasospasm.</a> J. Neurosci. Methods. 123, 89-97 (2003).</li><li>Altay, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+for+subarachnoid+hemorrhage+to+induce+vasospasm+in+mice.">A novel method for subarachnoid hemorrhage to induce vasospasm in mice.</a> J. Neurosci. Methods. 183, 136-140 (2009).</li><li>Feiler, S., Friedrich, B., Scholler, K., Thal, S. C., Plesnila, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standardized+induction+of+subarachnoid+hemorrhage+in+mice+by+intracranial+pressure+monitoring.">Standardized induction of subarachnoid hemorrhage in mice by intracranial pressure monitoring.</a> J. Neurosci. Methods. 190, 164-170 (2010).</li><li>Bederson, J. B., Germano, I. M., Guarino, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cortical+blood+flow+and+cerebral+perfusion+pressure+in+a+new+noncraniotomy+model+of+subarachnoid+hemorrhage+in+the+rat.">Cortical blood flow and cerebral perfusion pressure in a new noncraniotomy model of subarachnoid hemorrhage in the rat.</a> Stroke. 26, 1086-1091 (1995).</li><li>Veelken, J. A., Laing, R. J., Jakubowski, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Sheffield+model+of+subarachnoid+hemorrhage+in+rats.">The Sheffield model of subarachnoid hemorrhage in rats.</a> Stroke. 26, 1279-1283 (1995).</li><li>Kamii, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amelioration+of+vasospasm+after+subarachnoid+hemorrhage+in+transgenic+mice+overexpressing+CuZn-superoxide+dismutase.">Amelioration of vasospasm after subarachnoid hemorrhage in transgenic mice overexpressing CuZn-superoxide dismutase.</a> Stroke. 30, 867-871 (1999).</li><li>Parra, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+model+of+subarachnoid+hemorrhage+associated+cerebral+vasospasm:+methodological+analysis.">Mouse model of subarachnoid hemorrhage associated cerebral vasospasm: methodological analysis.</a> Neurol. Res. 24, 510-516 (2002).</li><li>Broderick, J. P., Brott, T. G., Duldner, J. E., Tomsick, T., Leach, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Initial+and+recurrent+bleeding+are+the+major+causes+of+death+following+subarachnoid+hemorrhage.">Initial and recurrent bleeding are the major causes of death following subarachnoid hemorrhage.</a> Stroke. 25, 1342-1347 (1994).</li><li>Thal, S. C., Plesnila, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-invasive+intraoperative+monitoring+of+blood+pressure+and+arterial+pCO2+during+surgical+anesthesia+in+mice.">Non-invasive intraoperative monitoring of blood pressure and arterial pCO2 during surgical anesthesia in mice.</a> J. Neurosci. Methods. 159, 261-267 (2007).</li><li>Hockel, K., Trabold, R., Scholler, K., Torok, E., Plesnila, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impact+of+anesthesia+on+pathophysiology+and+mortality+following+subarachnoid+hemorrhage+in+rats.">Impact of anesthesia on pathophysiology and mortality following subarachnoid hemorrhage in rats.</a> Exp. Transl. Stroke Med. 4, 5 (2012).</li><li>Wang, Z., Schuler, B., Vogel, O., Arras, M., Vogel, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+is+the+optimal+anesthetic+protocol+for+measurements+of+cerebral+autoregulation+in+spontaneously+breathing+mice?.">What is the optimal anesthetic protocol for measurements of cerebral autoregulation in spontaneously breathing mice?.</a> Exp. Brain Res. 207, 249-258 (2010).</li><li>Schwartz, A. Y., Masago, A., Sehba, F. A., Bederson, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+models+of+subarachnoid+hemorrhage+in+the+rat:+a+refinement+of+the+endovascular+filament+model.">Experimental models of subarachnoid hemorrhage in the rat: a refinement of the endovascular filament model.</a> J. Neurosci. Methods. 96, 161-167 (2000).</li><li>Feiler, S., Plesnila, N., Thal, S. C., Zausinger, S., Scholler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contribution+of+matrix+metalloproteinase-9+to+cerebral+edema+and+functional+outcome+following+experimental+subarachnoid+hemorrhage.">Contribution of matrix metalloproteinase-9 to cerebral edema and functional outcome following experimental subarachnoid hemorrhage.</a> Cerebrovasc. Dis. 32, 289-295 (2011).</li></ol>

Authors have nothing to disclose.

Treatment options after SAH are scarce and mostly inefficacious. Therefore the pathophysiology of post-hemorrhagic brain damage needs to be further understood in order to identify new therapeutic targets and develop novel therapeutic approaches. Standardized and well reproducible animal models in genetically modified animals, i.e. mice, are crucial for such investigations. The CWp model has become a widely used model for SAH as it resembles the pathophysiology in humans closely; however, its use in mice is hampe...

SAH is the stroke subtype with the least beneficial outcome for patients: 40% of the patients die within a month after the bleeding1 and survivors rarely have a clinically favorable outcome.
The large majority of spontaneous SAHs (80%) are caused by rupture of intracranial aneurysms which are mostly located along the anterior and posterior communicating artery, the basilar artery, and middle cerebral artery (MCA)2.
Such aneurysms are difficult to model in animals and therefore animal models of SAH are either performed by injection of blood into the subarachnoid space/cerebral ventricles or by endovascular perforation of a subarachnoid vessel.
Autologous blood injection into the cisterna magna is easy to perform and reproducible as the blood volume can be directly controlled3. Unfortunately some aspects of the SAH pathophysiology, e.g. the vessel injury, cannot be modeled by this procedure. Another technical approach for induction of SAH is the opening of an intracisternal vein4.
However, the intraluminal CWp at the MCA branch appears to be the procedure that models the pathophysiology in humans most closely5. The method was developed and first described in rats by Bederson and colleagues and at the same time by Veelken and colleagues6,7. Later the intraluminal perforation model was adapted to mice8,9. A filament is inserted into the external carotid artery (ECA) and advanced to the skull base via the internal carotid artery (ICA). At the branching point of the MCA the filament perforates the vessel and induces a bleeding into the subarachnoid space at the skull base. The blood then distributes into the remaining subarachnoid space along fissures and blood vessels. Bleeding is stopped by clot formation at the site of perforation, but rebleedings, which are often detrimental in patients10, can occur. Accordingly, the endovascular filament model became a widely used SAH model during the past few years. The most frequently mentioned drawback of the filament perforation model is that bleeding volume cannot be directly controlled and may therefore be variable. This variability can significantly be reduced by tight control of animal physiology and post-hemorrhagic ICP.
Mice have the great advantage that a large number of genetically modified strains are available. However, due to their small size surgical procedures tend to be more complex than in larger species, e.g. rats or rabbits. Therefore the downscaling of techniques developed for rats to mice often does not lead to the desired results, e.g. as mice have a very limited body weight and blood volume noninvasive techniques for blood pressure and blood gas analysis as well as for hemoglobin saturation and heart rate monitoring have to be applied whenever possible. Accordingly, the aim of the current publication is to describe the filament perforation model for SAH in mice and to demonstrate how this model can be performed in a standardized and highly reproducible manner.

<table border="1">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>operation microscope</td>
			<td>Leica</td>
			<td>KL2500</td>
			<td></td>
		</tr>
		<tr>
			<td>isoflurane vaporizer</td>
			<td>Harvard Instruments</td>
			<td>Continuous Flow Vaporizer</td>
			<td></td>
		</tr>
		<tr>
			<td>respirator</td>
			<td>Hugo Sachs</td>
			<td>Minivent 845</td>
			<td></td>
		</tr>
		<tr>
			<td>microcapnograph</td>
			<td>Hugo Sachs</td>
			<td>Type 340</td>
			<td></td>
		</tr>
		<tr>
			<td>temperature controller</td>
			<td>FHC</td>
			<td>DC Temperature Controller</td>
			<td></td>
		</tr>
		<tr>
			<td>dental drill</td>
			<td>Paggen</td>
			<td>Labset- N</td>
			<td></td>
		</tr>
		<tr>
			<td>ICP monitor</td>
			<td>Codman</td>
			<td>ICP monitor</td>
			<td></td>
		</tr>
		<tr>
			<td>blood pressure monitor</td>
			<td>AD Instruments</td>
			<td>Bridge Amp FE221</td>
			<td></td>
		</tr>
		<tr>
			<td>syringe pump</td>
			<td>World Precision Instruments</td>
			<td>SP101IZ</td>
			<td></td>
		</tr>
		<tr>
			<td>pulsoximeter</td>
			<td>Kent Scientific</td>
			<td>MouseSTAT</td>
			<td></td>
		</tr>
		<tr>
			<td>LDF</td>
			<td>Perimed</td>
			<td>Periflux 5000</td>
			<td></td>
		</tr>
		<tr>
			<td>analog data monitor</td>
			<td>AD Instruments</td>
			<td>Power Lab 16/35</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Material</td>
		</tr>
		<tr>
			<td>cement for ICP probe fixation</td>
			<td>Speiko</td>
			<td>Carboxylate cement</td>
			<td></td>
		</tr>
		<tr>
			<td>glue for LDF probe fixation</td>
			<td>Bob Smith Industries</td>
			<td>Cyanoacrylate glue (Maxi Cure and Insta Set)</td>
			<td></td>
		</tr>
		<tr>
			<td>venous catheter</td>
			<td>Johnson &#38; Johnson</td>
			<td>Jelco winged i.v. catheter; REF 4076</td>
			<td>modified intubation tube</td>
		</tr>
		<tr>
			<td>tubing for femoral catheter</td>
			<td>Smiths Medical</td>
			<td>Fine Bore Polythene Tubing; ID 0.28 mm OD 0.61 mm; REF 800/100/100</td>
			<td>cut to 30 cm length</td>
		</tr>
		<tr>
			<td>filament for vessel perforation</td>
			<td>Ethicon</td>
			<td>Prolene 5-0</td>
			<td>cut to 12 mm length</td>
		</tr>
		<tr>
			<td>surgical equipment</td>
			<td>Fine Scientific Instruments</td>
			<td>forceps medical #5, vessel scissors 8 cm, microclip 4 mm jaw</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Mortality
Once the surgery technique is mastered the procedure does not elicit any intraoperative mortality. Also bleeding can be achieved in virtually all animals. Postoperative mortality is 30-40% with most animals dying on day 1 after surgery (Figure 5).
ICP values after SAH
The ICP before bleeding is around 4 mmHg. Bleeding results in a sharp increase of the ICP up to 120 mmHg. ICP values then stabilize within 5 min at appr...

Watch this Scientific Journal Video about A Murine Model of Subarachnoid Hemorrhage at JoVE.com

A Murine Model of Subarachnoid Hemorrhage

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-subarachnoid-hemorrhage

Research

JoVE Journal

Medicine

19.5K Views.  University of Munich Medical Center.  Стандартная модель мыши из субарахноидального кровоизлияния по внутрипросветной Круга Уиллиса перфорации описывается. Судно перфорация и субарахноидальное кровотечение контролируются мониторинга внутричерепного давления. Кроме того различные жизненны?...

Video: A Murine Model of Subarachnoid Hemorrhage

A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional capacity. Transcutaneous surface stimulation of skeletal muscle involves a current flow between a cathode and an anode, thereby inducing excitement of the motor unit and the surrounding muscle fibers. 
NMES is an attractive modality to evaluate skeletal muscle adaptive responses for several reasons. First, it provides a reproducible experimental model in which physiological adaptations, such as myofiber hypertophy and muscle strengthening6, angiogenesis7-9, growth factor secretion9-11, and muscle precursor cell activation12 are well documented. Such physiological responses may be carefully titrated using different parameters of stimulation (for Cochrane review, see 13). In addition, NMES recruits motor units non-selectively, and in a spatially fixed and temporally synchronous manner14, offering the advantage of exerting a treatment effect on all fibers, regardless of fiber type. Although there are specified contraindications to NMES in clinical populations, including peripheral venous disorders or malignancy, for example, NMES is safe and feasible, even for those who are ill and/or bedridden and for populations in which rigorous exercise may be challenging. 
Here, we demonstrate the protocol for adapting commercially available electrodes and performing a NMES protocol using a murine model. This animal model has the advantage of utilizing a clinically available device and providing instant feedback regarding positioning of the electrode to elicit the desired muscle contractile effect. For the purpose of this manuscript, we will describe the protocol for muscle stimulation of the anterior compartment muscles of a mouse hindlimb.

A murine model of neuromuscular electrical stimulation (NMES), a safe and inexpensive clinical modality, to the anterior compartment muscles is described. This model has the advantage of modifying a readily available clinical device for the purpose of eliciting targeted and specific muscle contractions in mice.

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional ...

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) ...

Murine Model of Wound Healing

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation. There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of humans.

A murine model of cutaneous wound healing that can be used to assess therapeutic compounds in physiological and pathophysiological settings.

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. ...

A New Murine Model of Endovascular Aortic Aneurysm Repair

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

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

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

Intrastriatal Injection of Autologous Blood or Clostridial Collagenase as Murine Models of Intracerebral Hemorrhage

Intracerebral hemorrhage (ICH) is a common form of cerebrovascular disease and is associated with significant morbidity and mortality. Lack of effective treatment and failure of large clinical trials aimed at hemostasis and clot removal demonstrate the need for further mechanism-driven investigation of ICH. This research may be performed through the framework provided by preclinical models. Two murine models in popular use include intrastriatal (basal ganglia) injection of either autologous whole blood or clostridial collagenase. Since, each model represents distinctly different pathophysiological features related to ICH, use of a particular model may be selected based on what aspect of the disease is to be studied. For example, autologous blood injection most accurately represents the brain&#39;s response to the presence of intraparenchymal blood, and may most closely replicate lobar hemorrhage. Clostridial collagenase injection most accurately represents the small vessel rupture and hematoma evolution characteristic of deep hemorrhages. Thus, each model results in different hematoma formation, neuroinflammatory response, cerebral edema development, and neurobehavioral outcomes. Robustness of a purported therapeutic intervention can be best assessed using both models. In this protocol, induction of ICH using both models, immediate post-operative demonstration of injury, and early post-operative care techniques are demonstrated. Both models result in reproducible injuries, hematoma volumes, and neurobehavioral deficits. Because of the heterogeneity of human ICH, multiple preclinical models are needed to thoroughly explore pathophysiologic mechanisms and test potential therapeutic strategies.

Preclinical models of intracerebral hemorrhage are utilized to mimic certain aspects of clinical disease. Thus, mechanisms of injury and potential therapeutic strategies may be explored. In this protocol, two models of intracerebral hemorrhage are described, intrastriatal (basal ganglia) injections of autologous blood or collagenase.

Intracerebral hemorrhage (ICH) is a common form of cerebrovascular disease and is associated with significant morbidity and mortality. Lack of effective ...

A Preclinical Murine Model of Hepatic Metastases

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, murine pancreatic tumor model of hepatic metastases via a hemispleen injection of syngeneic murine pancreatic tumor cells is described. This model mimics many of the clinical conditions in patients with metastatic disease to the liver. Mice consistently develop metastases in the liver allowing for investigation of the metastatic process, experimental therapy testing, and tumor immunology research.

A preclinical, murine model of hepatic metastases performed via a hemispleen injection technique.

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, ...

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 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 ...

A Murine Model of Irreversible and Reversible Unilateral Ureteric Obstruction

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. Presented here is a model of obstructive nephropathy induced by unilateral ureteric obstruction (UUO), which can either be irreversible (UUO) or reversible (R-UUO). In the irreversible UUO model, the ureter may be obstructed for variable periods of time in order to induce increasingly severe renal inflammation and interstitial fibrotic scarring. In the reversible R-UUO model the ureter is obstructed to induce hydronephrosis, tubular dilation and inflammation. After a suitable period of time the ureteric obstruction is then surgically reversed by anastomosis of the severed previously obstructed ureter to the bladder in order to allow complete decompression of the kidney and restoration of urinary flow to the bladder. The irreversible UUO model has been used to investigate various aspects of renal inflammation and scarring including the pathogenesis of disease and the testing of potential anti-inflammatory or anti-fibrotic therapies. The more challenging model of R-UUO has been used by some investigators and does offer significant research potential as it allows the study of inflammatory and immune processes and tissue remodeling in an injured and scarred kidney following the removal of the injurious stimulus. As a result, the R-UUO model offers investigators the opportunity to explore the resolution of kidney inflammation together with key aspects of tissue repair. These experimental models are of relevance to human disease as patients often present with obstruction of the renal tract that requires decompression and are commonly left with significant residual kidney impairment that has no current treatment options and may lead to eventual end stage kidney failure.

The murine model of irreversible unilateral ureteric obstruction (UUO) is presented together with the model of reversible UUO in which the ureteric obstruction is relieved by anastomosis of the severed ureter into the bladder. These models enable the study of renal inflammation and scarring as well as tissue remodeling.

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. Presented here is a model of obstructive ...

Pre-clinical Orthotopic Murine Model of Human Prostate Cancer

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information about this disease. The human orthotopic prostate cancer xenograft mouse model provides a useful alternative approach for understanding the specific interactions between genetically and molecularly altered tumor cells, their organ microenvironment, and for evaluation of efficacy of therapeutic regimens. This is a well characterized model designed to study the molecular events of primary tumor development and it recapitulates the early events in the metastatic cascade prior to embolism and entry of tumor cells into the circulation. Thus it allows elucidation of molecular mechanisms underlying the initial phase of metastatic disease. In addition, this model can annotate drug targets of clinical relevance and is a valuable tool to study prostate cancer progression. In this manuscript we describe a detailed procedure to establish a human orthotopic prostate cancer xenograft mouse model.

Prostate cancer is the second most common cause of cancer-related deaths in the United States. An orthotopic cancer model provides a useful approach to understand the biology of prostate cancer and to evaluate the efficacy of therapeutic regimens. This protocol describes detailed steps necessary to establish an orthotopic prostate cancer mouse model.

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information ...

Integrated Compensatory Responses in a Human Model of Hemorrhage

Hemorrhage is the leading cause of trauma-related deaths, partly because early diagnosis of the severity of blood loss is difficult. Assessment of hemorrhaging patients is difficult because current clinical tools provide measures of vital signs that remain stable during the early stages of bleeding due to compensatory mechanisms. Consequently, there is a need to understand and measure the total integration of mechanisms that compensate for reduced circulating blood volume and how they change during ongoing progressive hemorrhage. The body&#39;s reserve to compensate for reduced circulating blood volume is called the &#39;compensatory reserve&#39;. The compensatory reserve can be accurately evaluated with real-time measurements of changes in the features of the arterial waveform measured with the use of a high-powered computer. Lower Body Negative Pressure (LBNP) has been shown to simulate many of the physiological responses in humans associated with hemorrhage, and is used to study the compensatory response to hemorrhage. The purpose of this study is to demonstrate how compensatory reserve is assessed during progressive reductions in central blood volume with LBNP as a simulation of hemorrhage.

The purpose of this protocol is to demonstrate the techniques for measuring compensatory responses to reduced central blood volume using lower body negative pressure as a noninvasive experimental model of human hemorrhage which can be used to quantify the total integration of compensatory mechanisms to blood volume deficit in humans.