The work was supported by the Lundbeck Foundation and the Lundbeck Grant of Excellence (no. R59-A5404). Funders had no role in any part of the manuscript.

This procedure is done in accordance with the European Union&#39;s Directive 2010/63/EU regarding the protection of animals used for scientific purposes and approved by the Danish Animal Experiments Inspectorate (license no. 2016-15-0201-00940). Surgery is performed using aseptic technique to the widest extent possible, including sterile instruments, gloves, catheters, and sutures. The study used male and female Sprague-Dawley rats weighing 230-350 g, group housed in 12-h light/dark cycle, with constant temperature of 22 &#176;C (&#177; 2&#160;&#176;C), and humidity of 55% (&#177; 10%). The animals are provided with standard chow and water ad libitum. The animals are housed in single cages following surgery but can be returned to group caging when the ICP-probe has been removed. The anesthetic in this protocol is isoflurane gas but a 1.5 mL/kg of 3:2 intraperitoneal mixture of ketamine (100 mg/mL) and xylazine (20 mg/mL) is also effectful21.
1. Preparations
<ol>
	<li>Modify a 16 G peripheral vein catheter for intubation. To modify, shorten the needle by 1 cm and bend the remaining distal 1 cm by 30&#176; toward the injection valve. Remove the catheter wings (multiple use).</li>
	<li>To make an ICP probe, cut a 20 mm piece of polythene tubing (inner diameter (ID): 0.58 mm, outer diameter (OD): 0.96 mm) and burn one end to make a circular plate, keeping an open lumen. Circumvent the polythene tubing with 1 mm of silicone tubing (ID: 1.0 mm, OD: 3.0 mm) before connecting 10 mm of silicone tubing (ID: 0.76 mm, OD: 2.4 mm) to the end of the polythene tubing.</li>
	<li>Power on the laptop and open the data acquisition software. Calibrate the blood pressure (BP) and intra cerebral pressure (ICP) transducers, and the Laser-Doppler according to the manufacturer&#39;s instructions.</li>
	<li>Prepare the blood gas analyzer apparatus. 
	CAUTION: Make sure there is enough isoflurane in the vaporizer.</li>
	<li>Turn on the O2 and atmospheric air&#160;flow. Set the flow of O2 at 30% and atmospheric air&#160;at 70%.</li>
	<li>Place the heating pad and set the temperature to 37 &#176;C.</li>
</ol>
2. Anesthesia
<ol>
	<li>Place the rat in the anesthesia chamber with a flow of 30% of O2 and 70% of atmospheric air. Administer 5% of isoflurane gas into the chamber. Adequate anesthesia will take around 4 min. Control the breathing carefully.</li>
	<li>When anesthetized, place the rat in supine position on a heavy plate circumvented by a rubber band. Place the front teeth of the rat below the rubber band.</li>
	<li>Draw the tongue out carefully with curved forceps. Clean the larynx with a cotton tip. Place an external light in the midline of the throat to visualize the vocal cords.</li>
	<li>Intubate during inspiration using the modified 16 G peripheral vein catheter. When correctly inserted, remove the stiletto. Connect the catheter to the ventilator. 
	NOTE: Correct placement of the tube is confirmed by chest movements in sync with respiration rate. If movements of the abdomen are seen, extubate and reintroduce the rat into the anesthesia bell. Do not repeat the procedure more than three times due to the risk of damaging the airways.</li>
	<li>When intubated, keep the animal on artificial respiration with 30% of O2 and 70% of atmospheric air. Maintain the anesthesia at 1.5%-3% of isoflurane. Adjust the isoflurane to keep the blood pressure between 80-100 mmHg.</li>
	<li>Keep the inspiratory volume of the respirator at 3 mL and the frequency at 40-45 inspirations/min. Adjust the inspiratory volume according to the blood gas analysis.</li>
	<li>Make a stitch through the inner soft tissue of the cheek with a 2-0 suture. Tie the suture around the injection tube and the injection valve of the peripheral vein catheter to fasten the catheter.</li>
	<li>Move the rat to the operating field and place it in supine position with the tail facing toward the surgeon.</li>
	<li>Apply the eye gel when needed to counter dry eyes.</li>
	<li>Perform a toe pinch to confirm an adequate depth of anesthesia.&#160;Assess and maintain&#160;anesthesia depth during surgery.</li>
</ol>
3. Tail catheter
<ol>
	<li>Disinfect the proximal 3-4 cm of the tail with 0.5% of chlorhexidine ethanol. 
	NOTE: From now on, use the surgical microscope upon the surgeon&#39;s discretion.</li>
	<li>Make a 15-20 mm skin incision in the proximal end of the tail on the ventral side. Be careful not to incise the artery.</li>
	<li>Loosen the skin from the underlying connective tissue using a curved forceps.</li>
	<li>Carefully penetrate the fascia exposing the artery.</li>
	<li>Carefully release the tail artery from the underlying tissue using a curved forceps.</li>
	<li>Slip three black silk threads under the vessel. Place one thread as distally as possible and tie a surgical knot tightly around the artery. Hold the loose ends of the thread with a hemostat.</li>
	<li>Tie the two remaining threads loosely around the artery.</li>
	<li>Push the proximal thread as proximally as possible. Apply a hemostat to hold the ends of the proximal thread. Pull the hemostat lightly, but enough to restrict and block the blood flow. Place the hemostat on the abdomen.</li>
	<li>Cut the tip of the catheter at a 45&#176; angle. Cut the sharp point to prevent arterial wall penetration.</li>
	<li>Using a Vannas scissor, make an artery incision 1/3 of the artery&#39;s diameter at a 30&#176; angle, 3-5 mm from the distal knot.</li>
	<li>Insert the catheter into the artery using two straight forceps. Use one forceps to hold the catheter and the other to carefully pull the artery over the catheter.</li>
	<li>Insert the catheter up the vessel to the proximal knot and loosen the knot from the hemostat. Visualize the blood flow in the catheter. Fasten the middle thread loosely to the catheter.</li>
	<li>Continue insertion to, and if possible, just beyond, the point where the artery is covered again by fascia.</li>
	<li>Control the catheter placement and possible leak by flushing with saline.</li>
	<li>Fasten the two proximal threads using surgical knots. 
	NOTE: The blood pressure measurement needs to be pulsatile; if not, the catheter is not properly placed.</li>
	<li>Fasten the catheter at the end of the incision by tying a surgical knot using the distal thread.</li>
	<li>Stitch the skin incision loosely together with two non-resorbable monofilament 4-0 suture. Be careful not to penetrate the catheter. 
	NOTE: Throughout the surgery be aware of the amplitude of pulsation. If this is low, flush the catheter with saline.</li>
	<li>Loosen the arterial catheter from the pressure transducer to allow blood flow for blood gas sampling. Place a micro capillary tube at the end of the catheter. Let the blood flow into the tube. Re-attach the catheter to the transducer after blood collection and flush the catheter.</li>
	<li>Insert the capillary tube in the blood gas analyzer. Measure the pH, pCO2, and pO2 and note them down 
	&#8203;NOTE: Depending on the blood gas and blood pressure values, change the ventilation rate. If the mean arterial pressure (MAP) is too low, try to turn down the flowrate of isoflurane. Test the reflexes to ensure proper depth of anesthesia.</li>
</ol>
4. ICP probe
<ol>
	<li>Place the rat in the stereotaxic frame. It is important to position the rat symmetrically.</li>
	<li>Place a cylindrical pillow under the stereotaxic frame to create anterior flexion of the neck.</li>
	<li>Shave the rat&#39;s scalp, neck, and the area behind the ears. Remove the superfluous hair.</li>
	<li>Disinfect the area with 0.5% of chlorhexidine ethanol.</li>
	<li>Anesthetize locally with 0.7 mL of 10 mg/5 &#181;g/mL lidocaine with adrenaline, insert the needle at the caudal end of the skull in the midline. Inject into the musculature of the neck with 0.3-0.4 mL. Inject the rest subcutaneously around and anterior to the bregma.</li>
	<li>Make a skin incision from the needle puncture ~8 mm caudally in the midline.</li>
	<li>Dissect all the muscles bluntly in layers to identify the atlantooccipital membrane (marble colored triangle caudally to the skull in the midline).</li>
	<li>Use the Alm retractor to restrain the neck musculature. Place the pronged retractor caudally if needed.</li>
	<li>Check whether the sterile ICP-probe is connected to the ICP transducer. Flush the ICP probe with saline. Ensure no air bubbles are present in the ICP probe.</li>
	<li>Incise the atlantooccipital membrane using a 23 G needle. Make a hole to coax the ICP probe through the membrane.</li>
	<li>Coax the probe through the atlantooccipital membrane gently. Pull the probe lightly and ensure that it shows a pulsating curve ranging between 0-5 mmHg. If not, remove the probe, check the connection to the transducer, and confirm the flow through the lumen.</li>
	<li>Apply two drops of the tissue glue. Move the 1 mm silicone tubing forward to the membrane and apply additional glue to minimize the risk of ICP-probe displacement.</li>
	<li>Remove the retractor(s).</li>
	<li>Make one horizontal mattress suture to the cephalic end of the incision and one simple interrupted suture to the caudal end using a non-resorbable monofilament 4-0 suture.</li>
</ol>
5. Placement of the needle and the Laser-Doppler probe
<ol>
	<li>Make an incision in the midline just anterior to the eyes, 15 mm caudally.</li>
	<li>Remove the connective tissue and the muscles with forceps. Use the end of a sterile cotton swab as a rougine making it possible to identify the bregma and the coronal sutures.</li>
	<li>Place the Alm retractor.</li>
	<li>Place a 25 G spinal needle in the stereotaxic frame. Place the needle exactly on the bregma and note the position. 
	NOTE: Place the midline joint of the stereotaxic frame at 30&#176; toward the animal in the vertical plane.</li>
	<li>Remove the needle from the bregma, move the frame&#160;65 mm anteriorly and then replace the needle in the midline to mark the site of drilling.</li>
	<li>Drill until the dura mater is identified below the bone. Gently remove the bone fragments using straight forceps and fill the cavity with bone wax.</li>
	<li>Drill another hole 3-4 mm lateral to the right of the bregma and just anterior to the coronal suture for the Laser-Doppler. It is not necessary to drill all the way through the bone. Be careful not to penetrate the dura mater.</li>
	<li>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 FU is required. Remove the microscope (artificial light).</li>
	<li>If the values are still acceptable, add one drop of glue to fix the probe.</li>
	<li>Recheck to confirm whether the value is above 80 FU. If the value is below 80 FU, remove and reposition the probe to reach a value above 80 FU. 
	&#8203;NOTE: The value, FU, is an arbitrary unit showing cerebral blood flow (CBF).</li>
</ol>
6. Induction of SAH
<ol>
	<li>Insert the needle gently through the skull in the midline between the hemispheres until resistance of the base of the skull is felt. Retract the needle by 1 mm to ensure correct placement just anteriorly to the optic chiasm.</li>
	<li>Turn the needle 90&#176; clockwise so that the needle tip points to the right to ensure the most homogenous result when injecting the blood. Remove the stiletto (Figure 3).</li>
	<li>Equilibrate for 15 min and adjust the level of anesthesia to obtain a mean arterial blood pressure in the range 80-100 mmHg.</li>
	<li>Perform a blood gas analysis. Adjust the level of anesthesia accordingly.</li>
	<li>Withdraw 500 &#181;L of blood from the tail catheter using a 1 mL syringe with a blunt 23 G needle.</li>
	<li>Fill the dead space of spinal needle chamber with blood to avoid injection of air. Remove the 23 G needle from the blood-filled syringe and confirm that the syringe contains 300 &#181;L of blood.</li>
	<li>Connect the syringe to the spinal needle. Grasp firmly and inject the blood manually to surpass MAP.</li>
	<li>Observe a steep rise in ICP and a steep fall in CBF on the laptop. 
	&#8203;NOTE: CBF should be 50% or lower compared to the baseline score for at least 5 min for the surgery to be successful, see Figure 4.&#160;Sham rats do not undergo the steps 6.1-6.7, thereby omitting the introduction of the spinal needle into the cerebrum, minimizing possible spontaneous hemorrhage, and iatrogenic brain damage.</li>
</ol>
7. Recovery and awakening
<ol>
	<li>Administer 0.1 mL/100 g of animal weight of 5.0 mg/mL of carprofen and 1 mL/100 g of animal weight of isotonic saline subcutaneously. Make sure the liquids are at least at room temperature before administering.</li>
	<li>Subsequently keep the rat under anesthesia for 30 min following the SAH.</li>
	<li>Remove the needle, the laser doppler probe, and then fill the cavities with bone wax. Close the incision using two horizontal mattress sutures with non-resorbable monofilament 4-0 suture.</li>
	<li>To use the ICP probe for injections into the cisterna magna, remove the silicone tubing and insert a pinpoint adapter to the polythene tubing.</li>
	<li>If no intervention is planned, cut the simple, interrupted suture. Shorten the ICP probe as much as possible using a scissor and then glue the end to prevent leak of cerebrospinal fluid (CSF). Close the incision with a non-absorbable monofilament 4-0 suture.</li>
	<li>Remove the rat from the stereotaxic frame and place in a supine position. Remove the loose sutures from the tail incision.</li>
	<li>Place a single suture proximal and deep to the arterial catheter. Remove the catheter and tie the suture to prevent bleeding. Suture the tail-incision with a non-absorbable monofilament 4-0 suture.</li>
	<li>Turn off the isoflurane.</li>
	<li>Clean the rat and its fur as much as possible.</li>
	<li>When the pedal withdrawal reflex is regained and the rat has spontaneous respiration when decoupled from the ventilator, extubate it.</li>
	<li>Place the rat in a single cage with food and water ad libitum. Place one half of the cage under a heating plate and place the rat in this area of the cage.</li>
	<li>Perform intrathecal administration by adapting the pinport injector to a precision syringe and administer the treatment through the pinport adapter. This intervention is feasible in animals that are awake. See Figure 5.</li>
</ol>
8. ICP-probe removal (if not removed during surgery)
NOTE: Use a surgical microscope upon the surgeon&#39;s discretion.
<ol>
	<li>Place the rat in the anesthesia chamber as described earlier.</li>
	<li>When anesthetized, place the rat in supine position in the operation field with heating pad.</li>
	<li>Place the nose in the anesthesia mask. Set the levels of O2 to 30%, atmospheric air&#160;to 70%, and isoflurane to 2%.</li>
	<li>Continuously apply the eye gel to counter dry eyes.</li>
	<li>Cut the caudal simple interrupted suture. Open the incision and remove the possible necrotic tissue or blood clots.</li>
	<li>Shorten the ICP probe as much as possible using a scissor and glue the end to prevent the leak of cerebrospinal fluid (CSF). Close the incision with a non-absorbable monofilament 4-0 suture.</li>
	<li>Turn off the isoflurane.</li>
	<li>When the rat starts to move, place it in a single cage with food and water ad libitum. Place one half of the cage over a heating plate and place the rat in this area.</li>
	<li>When returned to habitual state, reintroduce the animals to each other in a joint cage under supervision for the first 15 min.</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subacute+phase+of+subarachnoid+haemorrhage+in+female+rats:+increased+intracranial+pressure,+vascular+changes+and+impaired+sensorimotor+function.">Subacute phase of subarachnoid haemorrhage in female rats: increased intracranial pressure, vascular changes and impaired sensorimotor function.</a> Microvascular Research. 135, 104127 (2020).</li><li>Ansar, S., Vikman, P., Nielsen, M., Edvinsson, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cerebrovascular+ETB,+5-HT1B,+and+AT1+receptor+upregulation+correlates+with+reduction+in+regional+CBF+after+subarachnoid+hemorrhage.">Cerebrovascular ETB, 5-HT1B, and AT1 receptor upregulation correlates with reduction in regional CBF after subarachnoid hemorrhage.</a> American Journal of Physiology - Heart and Circulatory Physiology. 293 (6), 3750-3758 (2007).</li><li>Hansen-Schwartz, J., 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=Subarachnoid+hemorrhage+enhances+endothelin+receptor+expression+and+function+in+rat+cerebral+arteries.">Subarachnoid hemorrhage enhances endothelin receptor expression and function in rat cerebral arteries.</a> Neurosurgery. 52 (5), 1188-1194 (2003).</li><li>Hayman, E. G., Wessell, A., Gerzanich, V., Sheth, K. N., Simard, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+global+cerebral+edema+formation+in+aneurysmal+subarachnoid+hemorrhage.">Mechanisms of global cerebral edema formation in aneurysmal subarachnoid hemorrhage.</a> Neurocritical Care. 26 (2), 301-310 (2017).</li><li>Miyata, 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=Vasa+vasorum+formation+is+associated+with+rupture+of+intracranial+aneurysms.">Vasa vasorum formation is associated with rupture of intracranial aneurysms.</a> Journal of Neurosurgery. , 1-11 (2019).</li><li>Tada, Y., 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=Roles+of+hypertension+in+the+rupture+of+intracranial+aneurysms.">Roles of hypertension in the rupture of intracranial aneurysms.</a> Stroke. 45 (2), 579-586 (2014).</li><li>Nuki, Y., 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=Elastase-induced+intracranial+aneurysms+in+hypertensive+mice.">Elastase-induced intracranial aneurysms in hypertensive mice.</a> Hypertension. 54 (6), 1337-1344 (1979).</li><li>Marbacher, S., Wanderer, S., Strange, F., Gr&#252;ter, B. E., Fandino, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Saccular+aneurysm+models+featuring+growth+and+rupture:+A+systematic+review.">Saccular aneurysm models featuring growth and rupture: A systematic review.</a> Brain Sciences. 10 (2), 101 (2020).</li><li>Altay, O., 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=Isoflurane+on+brain+inflammation.">Isoflurane on brain inflammation.</a> Neurobiology of Disease. 62, 365-371 (2014).</li><li>Hockel, K., Trabold, R., Sch&#246;ller, K., T&#246;r&#246;k, 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> Experimental and Translational Stroke Medicine. 4 (1), 5 (2012).</li><li>Kamp, M. 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=A+Systematic+and+meta-analysis+of+mortality+in+experimental+mouse+models+analyzing+delayed+cerebral+ischemia+after+subarachnoid+hemorrhage.">A Systematic and meta-analysis of mortality in experimental mouse models analyzing delayed cerebral ischemia after subarachnoid hemorrhage.</a> Translational Stroke Research. 8 (3), 206-219 (2017).</li><li>Povlsen, G. K., Johansson, S. E., Larsen, C. C., Samraj, A. K., Edvinsson, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+events+triggering+delayed+vasoconstrictor+receptor+upregulation+and+cerebral+ischemia+after+subarachnoid+hemorrhage.">Early events triggering delayed vasoconstrictor receptor upregulation and cerebral ischemia after subarachnoid hemorrhage.</a> BMC Neuroscience. 14, 34 (2013).</li></ol>

The authors have no conflicting interests to declare.

The pre-chiasmatic single injection model of SAH mimics several important elements of human SAH, including the spike in ICP, reduction of CBF, transient global ischemia, upregulation of neuroinflammatory markers, and CVS14,15,16,18,19,20. The ICP-probe was also used as a port for intrathecal administration (<strong class="xfi...

Non-traumatic subarachnoid hemorrhage (SAH) is a form of stroke, representing around 5% of all cases. The most common cause of non-traumatic SAH is the sudden rupture of an aneurysm (aSAH), which accounts for 85% of SAHs. Other causes include the rupture of an arterio-venous malformation, coagulopathies, and rupture of veins in perimesencephalic hemorrhage1. The incidence rate is 9 per 100,000 person-years with mortality around one in three and another third requiring the support of daily living following SAH2,3.
Following initial stabilization and diagnosis confirmation, treatment depends on the severity of the hemorrhage. The most severely afflicted patients will have an extra-ventricular drain inserted into the ventricles to reduce the intracerebral pressure (ICP) and be admitted to the neurointensive care unit, where they are monitored closely. Patients will undergo an angiography to identify the (probable) aneurysm and afterward have the aneurysm coiled or clipped to prevent rebleeding4. Despite numerous trials of pharmacological therapies, only nimodipine, a calcium-channel antagonist, has shown to improve outcomes5. Multiple clinical trials are currently underway. Please see the review by Daou and colleagues for an extensive list6.
The rupture of an aneurysm has been described as the sudden onset of the worst headache ever experienced or a thunderclap headache. The rupture results in a steep rise in the ICP followed by a reduction in the cerebral blood flow (CBF). This reduction results in global ischemia of the brain, which can result in a loss of consciousness. This more mechanistic pathway, along with the initiated breakdown of the extravasated elements of blood, gives rise to cytokine release and activation of the innate immune system resulting in sterile neuroinflammation. Furthermore, breakdown of the blood-brain barrier, resulting in cerebral edema and disturbance in the ion homeostasis, is often observed. All these changes and more, coined early brain injury (EBI), occur within the first couple of days and results in neuronal loss and apoptosis7.
Approximately 1/3 of patients afflicted with aSAH will develop delayed cerebral ischemia (DCI) between day 4-148. DCI is defined as either the debut of a focal, neurological impairment or a drop of minimum two points on the Glasgow coma scale lasting for a minimum of 1 h, when other causes, including seizures and re-bleeding is excluded. DCI is associated with an increased risk of death and decreased functional outcome following aSAH9. Cerebral vasospasm (CVS), the narrowing of the cerebral arteries, has been known to be associated with DCI for decades and was formerly thought to be the sole reason for DCI. It has since been shown that CVS can occur without the development of DCI and more factors, including microvascular thrombosis and constriction, cortical spreading depression, and an inflammatory response of EBI have since been identified10,11,12.
Due to the large influence of EBI and DCI on the course of the disease and the outcome of the patients afflicted, animal models need to mimic these to the largest degree possible, while still being reproducible. Researchers have employed a wide range of different models in a variety of animals from mice to non-human primates to try and simulate aSAH. Sprague-Dawley and Wistar wildtype rats are currently the most commonly used laboratory animals, and the most common models are the endovascular perforation model, the cisterna-magna double injection model, and lastly the pre-chiasmatic single injection model, which will be described in this article13.
The pre-chiasmatic, single injection model was originally developed by Prunell and colleagues to counter some of the shortcomings of the other experimental models14. The surgery, when mastered, is highly reproducible and minimizes variation between animals. The model mimics SAH in humans on multiple points, including the sudden rise in ICP following the injection of blood, resulting in transient global ischemia due to a fall in the CBF15,16. It affects the anterior circulation, which is where most aSAH in humans occur17. The mortality ranges from 10%-33% depending on the study and amount of blood injected14,18. Delayed cell death and neuroinflammation can be detected on day 2 and 7 thereby providing variables to study the consequences of EBI and DCI19,20.
The study presents an updated description of the pre-chiasmatic single injection model in the rat along with a description of how to utilize the ICP-probe as a port for intrathecal administration of pharmaceutics.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>16 G peripheral vein catheter</td>
			<td>BD Venflon</td>
			<td>393229</td>
			<td>Needle shortened, distal 1 cm curved. Wings removed</td>
		</tr>
		<tr>
			<td>Anesthesia bell/ chamber</td>
			<td>Unknown</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Blood gas analyzer</td>
			<td>Radiometer</td>
			<td>ABL80</td>
			<td></td>
		</tr>
		<tr>
			<td>Blood pressure (BP) monitor</td>
			<td>Adinstruments</td>
			<td>ML117</td>
			<td>Connects to Powerlab</td>
		</tr>
		<tr>
			<td>Curved forceps, 12 cm x 3</td>
			<td>F.S.T</td>
			<td>11001-12</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Cylindrical pillow, 28 cm x 4 cm</td>
			<td>Homemade</td>
			<td></td>
			<td>Made from surgical towels</td>
		</tr>
		<tr>
			<td>Data acquisition hardware</td>
			<td>Adinstruments</td>
			<td>ML870 Powerlab</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquistion software</td>
			<td>Adinstruments</td>
			<td>LabChart 6.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>KMD</td>
			<td>1189</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill controller</td>
			<td>Silfradent</td>
			<td>300 IN</td>
			<td></td>
		</tr>
		<tr>
			<td>Flexible light</td>
			<td>Schott</td>
			<td>KL200</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Minco</td>
			<td>1135</td>
			<td></td>
		</tr>
		<tr>
			<td>Hypodermic needle, 20 G</td>
			<td>KD Medical</td>
			<td>301300</td>
			<td>Connects to stereotaxic frame</td>
		</tr>
		<tr>
			<td>ICP monitor</td>
			<td>Adinstruments</td>
			<td>ML117</td>
			<td>Connects to Powerlab</td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer</td>
			<td>Ohmeda</td>
			<td>TEC3</td>
			<td></td>
		</tr>
		<tr>
			<td>Laptop</td>
			<td>Lenovo</td>
			<td>T410</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser doppler monitor</td>
			<td>Adinstruments</td>
			<td>ML191</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser doppler probe</td>
			<td>Oxford Optronics</td>
			<td>MSF100XP</td>
			<td>Connects to laser doppler monitor</td>
		</tr>
		<tr>
			<td>Needle holder, 13 cm</td>
			<td>F.S.T</td>
			<td>12001-13</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Precision syringe, 0.025 mL</td>
			<td>Hamilton</td>
			<td>547407</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic frame</td>
			<td>Kopf Instruments</td>
			<td>M900</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td>Carl Zeiss</td>
			<td>F170</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture needle</td>
			<td>Allgaier</td>
			<td>1245</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Temperaure controller</td>
			<td>CWE,INC.</td>
			<td>TC-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Transducer x 2</td>
			<td>Adinstruments</td>
			<td>MLT0699</td>
			<td>Connects to BP and ICP monitor</td>
		</tr>
		<tr>
			<td>Ventilator</td>
			<td>Ugo Basile</td>
			<td>7025</td>
			<td></td>
		</tr>
		<tr>
			<td>Veterinary clipper</td>
			<td>Aesculap</td>
			<td>GT421</td>
			<td></td>
		</tr>
		<tr>
			<td>3-pronged Blair retractor, 13.5 cm</td>
			<td>Agnthos</td>
			<td>17022--13</td>
			<td></td>
		</tr>
		<tr>
			<td>Blunt Alm retractor</td>
			<td>F.S.T</td>
			<td>17008-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved forceps, 12 cm x 2</td>
			<td>F.S.T</td>
			<td>11001-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder, 13 cm</td>
			<td>F.S.T</td>
			<td>12001-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Dumont forceps, 11 cm</td>
			<td>F.S.T</td>
			<td>11252-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Halsted-Mosquito hemostat x 2</td>
			<td>F.S.T</td>
			<td>13008-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Iris scissor, 9 cm</td>
			<td>F.S.T</td>
			<td>14090-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Vannas scissor, 10.5 cm</td>
			<td>F.S.T</td>
			<td>15018-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorpable swabs</td>
			<td>Kettenbach</td>
			<td>31603</td>
			<td></td>
		</tr>
		<tr>
			<td>Black silk thread, 4-0, 5 x 15 cm</td>
			<td>V&#246;mel</td>
			<td>14757</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone wax</td>
			<td>Aesculap</td>
			<td>1029754</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbomer eye gel 2 mg/g</td>
			<td>Paranova</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton swab</td>
			<td>Heinz Herenz</td>
			<td>WA-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton tipped applicator x 4</td>
			<td>Selefa</td>
			<td>120788</td>
			<td></td>
		</tr>
		<tr>
			<td>Hypodermic needle, 23 G x2</td>
			<td>KD Medical</td>
			<td>900284</td>
			<td>Connects to stopcock. Remove distal end</td>
		</tr>
		<tr>
			<td>Hypodermic needle, 23 G x3</td>
			<td>KD Medical</td>
			<td>900284</td>
			<td>Remove distal end. 2 connects to stopcock, 1 to syringe</td>
		</tr>
		<tr>
			<td>ICP probe:</td>
			<td>Homemade</td>
			<td></td>
			<td>Made of the following:</td>
		</tr>
		<tr>
			<td>Polythene tubing, 20 mm</td>
			<td>Smiths medical</td>
			<td>800/100/200</td>
			<td>Inner diameter (ID): 0.58 mm, Outer diameter (OD): 0.96 mm.</td>
		</tr>
		<tr>
			<td>Silicone tubing, 10 mm</td>
			<td>Fisher</td>
			<td>15202710</td>
			<td>ID: 0.76 mm, OD: 2.4 mm.</td>
		</tr>
		<tr>
			<td>Silicone tubing, 2 mm</td>
			<td>Fisher</td>
			<td>11716513</td>
			<td>ID: 1.0 mm, OD: 3.0 mm.</td>
		</tr>
		<tr>
			<td>Micro hematocrit tubes</td>
			<td>Brand</td>
			<td>7493 11</td>
			<td></td>
		</tr>
		<tr>
			<td>OP-towel, 45 cm x75 cm</td>
			<td>M&#246;lnlycke</td>
			<td>800430</td>
			<td></td>
		</tr>
		<tr>
			<td>PinPort adapter, 22 G</td>
			<td>Instech</td>
			<td>PNP3F22</td>
			<td></td>
		</tr>
		<tr>
			<td>PinPort injector</td>
			<td>Instech</td>
			<td>PNP3M</td>
			<td></td>
		</tr>
		<tr>
			<td>Polythene tubing, 2 x 20 cm</td>
			<td>Smiths medical</td>
			<td>800/100/200</td>
			<td>Connects to syringe. ID: 0.58 mm, OD: 0.96 mm.</td>
		</tr>
		<tr>
			<td>Rubberband</td>
			<td>Unknown</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel, 10 blade</td>
			<td>Kiato</td>
			<td>23110</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinalneedle, 25 G x 3.5&#39;&#39;</td>
			<td>Braun</td>
			<td>5405905-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Stopcock system, Discofix x 2</td>
			<td>Braun</td>
			<td>16494C</td>
			<td>Connects to transducer</td>
		</tr>
		<tr>
			<td>Suture, 4-0, monofil, non-resorbable x 3</td>
			<td>Ethicon</td>
			<td>EH7145H</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, 1 mL</td>
			<td>BD Plastipak</td>
			<td>1710023</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, luer-lock, 10 mL x 4</td>
			<td>BD Plastipak</td>
			<td>305959</td>
			<td>Connects to transducer</td>
		</tr>
		<tr>
			<td>Tissue adhesive glue</td>
			<td>3M</td>
			<td>1469SB</td>
			<td></td>
		</tr>
		<tr>
			<td>0.5% Chlorhexidine spirit</td>
			<td>Faaborg Pharma</td>
			<td>210918</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen 50 mg/mL</td>
			<td>ScanVet</td>
			<td>43715</td>
			<td>Diluted 1:10</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Baxter</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isotonic saline</td>
			<td>Amgros</td>
			<td>16404</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine-Adrenaline 10 mg/5 &#181;g/mL</td>
			<td>Amgros</td>
			<td>16318</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Women have an increased risk of aSAH compared to men. Despite this, male rodents are primarily used in experiments due to possible bias from heterogeneity of estrus cycle in females. The representative results presented here are from a recent publication comparing female and male rats, confirming that the model produces similar results in female animals compared to male21. The study included 34 female Sprague-Dawley rats (18 SAHs and 16 shams). Shams did not have the spinal needle descended to the...

Watch this Scientific Journal Video about Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model at JoVE.com

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

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 ...

pre-chiasmatic-single-injection-autologous-blood-to-induce

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

2.2K Views.  Rigshospitalet. 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.

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

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals are not viable in those cases where the modified gene is expressed or deleted in the whole organism. Moreover, a variety of compensatory mechanisms often make it difficult to interpret the results. The compensatory effects can be alleviated by either timing the gene expression or limiting the amount of transfected cells.
The method of postnatal non-ventricular microinjection and in vivo electroporation allows targeted delivery of genes, siRNA or dye molecules directly to a small region of interest in the newborn rodent brain. In contrast to conventional ventricular injection technique, this method allows transfection of non-migratory cell types. Animals transfected by means of the method described here can be used, for example, for two-photon in vivo imaging or in electrophysiological experiments on acute brain slices.

This protocol describes a non-viral method of delivery of genetic constructs to a certain area of living rodent brain. The method consists of plasmid preparation, micropipette fabrication, neonatal rat pup surgery, microinjection of the construct, and in vivo electroporation.

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals ...

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% ...

Visual Evoked Potential Recording in a Rat Model of Experimental Optic Nerve Demyelination

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to monitor the disease course in the follow-up period. Changes in the VEP parameters closely correlate with pathological damage in the optic nerve. This protocol provides a detailed description about the rodent model of optic nerve microinjection, in which a partial demyelination lesion is produced in the optic nerve. VEP recording techniques are also discussed. Using skull implanted electrodes, we are able to acquire reproducible intra-session and between-session VEP traces. VEPs can be recorded on individual animals over a period of time to assess the functional changes in the optic nerve longitudinally. The optic nerve demyelination model, in conjunction with the VEP recording protocol, provides a tool to investigate the disease processes associated with demyelination and remyelination, and can potentially be employed to evaluate the effects of new remyelinating drugs or neuroprotective therapies.

Focal demyelination is induced in the optic nerve using lysolecithin microinjection. Visual evoked potentials are recorded via skull electrodes implanted over the visual cortex to examine the signal conduction along the visual pathway in vivo. This protocol details the surgical procedures underlying electrode implantation and optic nerve microinjection.

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to ...

Rat Model of Widespread Cerebral Cortical Demyelination Induced by an Intracerebral Injection of Pro-Inflammatory Cytokines

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and death, caused by white matter lesions in the spinal cord and cerebellum, as well as by demyelination in grey matter. Whilst conventional models of experimental allergic encephalomyelitis are suitable for the investigation of the cell-mediated inflammation in the spinal and cerebellar white matter, they fail to address grey matter pathologies. Here, we present the experimental protocol for a novel rat model of cortical demyelination allowing the investigation of the pathological and molecular mechanisms leading to cortical lesions. The demyelination is induced by an immunization with low-dose myelin oligodendrocyte glycoprotein (MOG) in an incomplete Freund&#39;s adjuvant followed by a catheter-mediated intracerebral delivery of pro-inflammatory cytokines. The catheter, moreover, enables multiple rounds of demyelination without causing injection-induced trauma, as well as the intracerebral delivery of potential therapeutic drugs undergoing a preclinical investigation. The method is also ethically favorable as animal pain and distress or disability are controlled and relatively minimal. The expected timeframe for the implementation of the entire protocol is around 8 - 10 weeks.

The protocol presented here allows the reproduction of a widespread grey matter demyelination of both cortical hemispheres in adult male Dark Agouti rats. The method comprises of intracerebral implantation of a catheter, subclinical immunization against myelin oligodendrocyte glycoprotein, and intracerebral injection of a pro-inflammatory cytokine mixture through the implanted catheter.

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and ...

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 ...

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a&#160;good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) ...

A Benchtop Approach to the Location Specific Blood Brain Barrier Opening using Focused Ultrasound in a Rat Model

Stereotaxic surgery is the gold standard for localized drug and gene delivery to the rodent brain. This technique has many advantages over systemic delivery including precise localization to a target brain region and reduction of off target side effects. However, stereotaxic surgery is highly invasive which limits its translational efficacy, requires long recovery times, and provides challenges when targeting multiple brain regions. Focused ultrasound (FUS) can be used in combination with circulating microbubbles to transiently open the blood brain barrier (BBB) in millimeter sized regions. This allows intracranial localization of systemically delivered agents that cannot normally cross the BBB. This technique provides a noninvasive alternative to stereotaxic surgery. However, to date this technique has yet to be widely adopted in neuroscience laboratories due to the limited access to equipment and standardized methods. The overall goal of this protocol is to provide a benchtop approach to FUS BBB opening (BBBO) that is affordable and reproducible and can therefore be easily adopted by any laboratory.

Focused ultrasound with microbubble agents can open the blood brain barrier focally and transiently. This technique has been used to deliver a wide range of agents across the blood brain barrier. This article provides a detailed protocol for the localized delivery to the rodent brain with or without MRI guidance.

Stereotaxic surgery is the gold standard for localized drug and gene delivery to the rodent brain. This technique has many advantages over systemic ...

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% ...

Massive Pontine Hemorrhage by Dual Injection of Autologous Blood

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred microliters of autologous blood was taken from the tail vein and stereotaxically injected into the pons. The injection process was divided into 2 steps: First, 10 &#181;L of blood was injected into a specific location, anteroposterior position (AP) -9.0 mm; lateral (Lat) 0 mm; vertical (Vert) -9.2 mm, followed by a second injection of the residual blood located at AP -9.0 mm; Lat 0 mm; Vert -9.0 mm with a 20-minute interval. The balance beam test, limb placement test, and the modified Voestch neuroscore were used to evaluate neurological function. Magnetic Resonance Imaging (MRI) was used to assess the volume of hemorrhage in vivo. The symptoms of this model were in line with patients with massive pontine hemorrhage.

We present a protocol to establish a massive pontine hemorrhage model in a rat via dual injection of autologous blood.

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred ...

A Rat Model of EcoHIV Brain Infection

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

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

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