This work was funded by the NINDS grant: K08NS105914

All research methods and animal care/maintenance were performed in compliance with the institutional guidelines at the University of California, Davis. The Institutional Animal Care and Use Committee (IACUC) of the University of California, Davis, approved all animal use protocols and experimental procedures (IACUC protocol #21874).
1. Animal housing
<ol>
	<li>Obtain Sprague-Dawley rats of age 8-10 months old. Prior to any experimental procedure, house the rats in a vivarium and allow at least 1 week for general adaptation in their cages following a 12-h light/dark cycle with food and water ad libitum.</li>
</ol>
2. Anesthesia and pre-operative procedures
<ol>
	<li>Anesthetize the rat with 4% isoflurane for 4 min. Hang the rat by its teeth in a supine position on an intubation platform, and intubate endotracheally using an endotracheal cannula and laryngoscope.</li>
	<li>Place the anesthetized and intubated rat on a ventilator (2% isoflurane and O2/N2 carrier gas). The rat is adequately anesthetized if no response to a painful stimulus such as a hindleg pinch is observed.</li>
	<li>Insert a rectal thermometer to continuously monitor the temperature.</li>
	<li>Perform all operative procedures using sterile technique. Clip the hair on the head and the femoral region and prep the skin with three alternating scrubs of Betadine and 70% alcohol prior to surgery.</li>
	<li>Aspirate any accumulated respiratory secretions by temporarily removing the rat from the ventilator and aspirating the secretions with PE-50 tubing connected to a 10 mL syringe.</li>
	<li>Protect the rat&#39;s eyes with sterile artificial tears eye ointment.</li>
	<li>Inject local bupivacaine (~0.1 mL of 0.25% solution) into the skin and subcutaneous tissues prior to the scalp incision.</li>
</ol>
3. Surgery protocol
<ol>
	<li>Placement of intraventricular needle and intracranial pressure (ICP) monitor
	<ol>
		<li>Place the rat in a prone position in a stereotactic frame and ear bar the rat.</li>
		<li>Make a 1.5-cm scalp incision along the midline with a 15-blade scalpel.</li>
		<li>Apply mild pressure with gauze for hemostasis.</li>
		<li>Using a sterile cotton tip applicator, separate the periosteum from the skull until the bregma landmark is visible.</li>
		<li>Locate and mark out bregma using stereotaxis and mark out the location of two bilateral burr holes, 1.4 mm lateral and 0.9 mm posterior to the bregma.</li>
		<li>Using a hand-held drill, create these two small (up to 2 mm) cranial burr holes in the right and left hemispheres. Irrigate out any excess bone chips with sterile Lactated Ringer&#39;s solution.</li>
		<li>In the right hemisphere, position a 22-G guide cannula at the level of the burr hole to insert the 28-G needle through the cannula to the depth of the right lateral ventricle (4.6 mm in relation to the bregma) in order to create IVH.</li>
		<li>Connect the fiberoptic pressure sensor to the read-out unit. Turn on the read-out unit and ensure the units selected are in mmHg. Then prime the sensor by submerging its tip into a small beaker with Lactated Ringer&#39;s solution until the read-out unit reads out zero. Once it is zeroed in the Lactated Ringers solution, it is all set to be inserted.</li>
		<li>In the left hemisphere, gently insert the pressure sensor to 2-3 mm depth into the cortex for real-time ICP monitoring.</li>
	</ol>
	</li>
	<li>Femoral artery cannulation and insertion of mean arterial pressure (MAP) monitor
	<ol>
		<li>After insertion of the ICP monitor, turn the lower trunk of the rat for easy access to the left thigh and groin area.</li>
		<li>After sterile preparation and local bupivacaine administration, make a 1.5 cm skin incision over the hindlimb with a 15-blade scalpel.</li>
		<li>Dissect out the left femoral artery first superficially with a hemostat and then deeper layers using forceps with fine tips under a microscope. Identify the deep blue femoral vein to help locate the adjacent artery.</li>
		<li>Tie off the distal femoral artery using 3-0 silk suture and place a temporary metal clip on the proximal portion of the femoral artery.</li>
		<li>Have a second fiberoptic pressure sensor connected to the read-out unit already primed. Insert the pressure sensor into the polyethylene (PE-50) tubing, which is inserted into a Tuohy Borst that is then closed. Connect the Tuohy Borst to a 3-way-stopcock connected to a 1 mL syringe at one end and a 22-G needle with PE-50 tubing at the other end.</li>
		<li>Under the microscope, make a 2 mm femoral arteriotomy with micro scissors and cannulate it with PE-50 tubing connected to the rest of the setup.</li>
	</ol>
	</li>
	<li>Intraventricular injection
	<ol>
		<li>Aspirate 500 &#181;L of blood using a 1 mL syringe and turn the 3-way-stopcock to have the pressure sensor read MAP.</li>
		<li>Prime the 28-G intraventricular needle connected to PE-50 tubing with the aspirated blood for IVH animals and Lactated Ringer&#39;s for the vehicle control animals. Then insert this needle into the guide cannula to the depth of the right lateral ventricle.</li>
		<li>Using 100 &#181;L/min rate, inject the blood or sterile Lactated Ringer&#39;s solution (200 &#181;L) into the right lateral ventricle by pumping the 1 mL syringe with the thumb. Prior to this and during intraventricular injection, monitor and record ICP, arterial blood pressure, and rectal temperature.</li>
		<li>Monitor and record the post-injection ICP and MAP values.</li>
	</ol>
	</li>
	<li>Closure
	<ol>
		<li>Following completion of the intraventricular injection, withdraw the PE-50 tubing containing the pressure sensor that was inserted into the femoral artery and apply the temporary clip to the femoral artery to prevent bleeding.</li>
		<li>Tie off the proximal portion of the femoral artery using the 3-0 silk suture.</li>
		<li>Close the femoral incision in an interrupted fashion using 3-0 silk.</li>
		<li>Remove the guide cannula with the intraventricular needle and the ICP monitor.</li>
		<li>Seal the burr holes with bone wax.</li>
		<li>Close the cranial incision with 3-0 silk suture in an interrupted fashion.</li>
		<li>Apply topical bupivacaine to the incision and inject 0.35 mL of carprofen (5 mg/kg) postoperatively. Do not leave animals unattended until they have regained sufficient consciousness to maintain sternal recumbency.</li>
		<li>Allow rats to fully recover after surgery under supervision and return them to their home cages with free access to food and water after recovery.</li>
	</ol>
	</li>
</ol>
4. Postoperative management
<ol>
	<li>Check all postoperative animals daily for seven postoperative days to monitor their recovery, neurological status, behavior, weight, and incisions.</li>
	<li>Administer 0.35 mL of carprofen (5 mg/kg) by subcutaneous injection at the time of surgery and on the 1st and 2nd postoperative days.</li>
	<li>Remove the sutures on the 7th postoperative day in a sterile fashion.</li>
</ol>

<ol>
	<li>Gates, P. C., Barnett, H. J. M., Vinters, H. V., Simonsen, R. L., Siu, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primary+intraventricular+hemorrhage+in+adults.">Primary intraventricular hemorrhage in adults.</a> Stroke. 17, 872-877 (1986).</li><li>Strajle, J., Garton, H. J. L., Maher, C. O., Muraszko, K., Keep, R. F., Xi, G. <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+hydrocephalus+after+neonatal+and+adult+intraventricular+hemorrhage.">Mechanisms of hydrocephalus after neonatal and adult intraventricular hemorrhage.</a> Translational Stroke Research. 3, 25-38 (2012).</li><li>Murao, K., Rossi, C., Cordonnier, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracerebral+hemorrhage+and+cognitive+decline.">Intracerebral hemorrhage and cognitive decline.</a> Revue Neurologique. 169, 772-778 (2013).</li><li>Al-Khindi, T., Macdonald, R. L., Schweizer, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cognitive+and+functional+outcome+after+aneurysmal+subarachnoid+hemorrhage.">Cognitive and functional outcome after aneurysmal subarachnoid hemorrhage.</a> Stroke. 41, 519-536 (2010).</li><li>Kreiter, K. 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=Predictors+of+cognitive+dysfunction+after+subarachnoid+hemorrhage.">Predictors of cognitive dysfunction after subarachnoid hemorrhage.</a> Stroke. 33, 200-208 (2002).</li><li>Zanaty, M., 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=Intraventricular+extension+of+an+aneurysmal+subarachnoid+hemorrhage+is+an+independent+predictor+of+a+worse+functional+outcome.">Intraventricular extension of an aneurysmal subarachnoid hemorrhage is an independent predictor of a worse functional outcome.</a> Clinical Neurology and Neurosurgery. 170, 67-72 (2018).</li><li>Gabrielian, L., Willshire, L. W., Helps, S. C., vanden Heuvel, C., Mathias, J., Vink, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracranial+pressure+changes+following+traumatic+brain+injury+in+rats:+lack+of+significant+change+in+the+absence+of+mass+lesions+or+hypoxia.">Intracranial pressure changes following traumatic brain injury in rats: lack of significant change in the absence of mass lesions or hypoxia.</a> Journal of Neurotrauma. 28, 2103-2111 (2011).</li><li>Kolar, M., Nohejlova, K., Duska, F., Mares, J., Pachl, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+of+cortical+perfusion+in+the+early+phase+of+subarachnoid+bleeding+in+a+rat+model+and+the+role+of+intracranial+hypertension.">Changes of cortical perfusion in the early phase of subarachnoid bleeding in a rat model and the role of intracranial hypertension.</a> Physiological Research. 66, 545-551 (2017).</li><li>Ariesen, M. J., Claus, S. P., Rinkel, G. J. E., Algra, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Risk+factors+for+intracerebral+hemorrhage+in+the+general+population.+A+systematic+review.">Risk factors for intracerebral hemorrhage in the general population. A systematic review.</a> Stroke. 34, 2060-2066 (2003).</li><li>MacLellan, C. L., Paquette, R., Colbourne, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+critical+appraisal+of+experimental+intracerebral+hemorrhage+research.">A critical appraisal of experimental intracerebral hemorrhage research.</a> Journal of Cerebral Blood Flow &amp; Metabolism. 32, 612-627 (2012).</li><li>Hartman, R., Lekic, T., Rojas, H., Tang, 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=Assessing+functional+outcomes+following+intracerebral+hemorrhage+in+rats.">Assessing functional outcomes following intracerebral hemorrhage in rats.</a> Brain Research. 1280, 148-157 (2009).</li></ol>

All authors report no conflict of interest.

This study investigated mechanisms to measure ICPs, MAPs, and CPPs in a nontraumatic IVH rat animal model. The results were recorded from the following groups: sham, VH 200 &#181;L, and vehicle control (artificial cerebrospinal fluid intraventricular injection) animals. This experimental design was chosen to investigate how ICPs can be monitored during IVH injection as we hypothesized that the spike in ICPs may contribute to the more significant secondary brain injury and thus memory deficit in IVH animal models. Therefo...

Intraventricular hemorrhage (IVH), a type of intracranial hemorrhage (ICH), is a devastating disease that carries significant mortality and morbidity. IVH is characterized as the accumulation of blood products inside the intracranial ventricles. Isolated IVH in uncommon and typically occurs in adults1. It may be associated with hypertensive hemorrhage, ruptured intracranial aneurysm or another vascular malformation, tumors, or trauma1. IVH leads to secondary brain injury as well as the development of hydrocephalus2. Survivors of IVH are often left with significant functional, memory, and cognitive impairments following their injury. These long-term cognitive and memory deficits are reported in as high as 44% of survivors of ICH3. In subarachnoid hemorrhage (SAH), another type of ICH, it is well known that approximately half of the survivors will have memory deficits, and for those who have IVH in addition to SAH, outcomes tend to be significantly worse4,5,6.
Underlying mechanisms of memory dysfunction following IVH remain to be elucidated. In vivo research utilizing nontraumatic IVH animal models with functional and memory dysfunction is essential in order to discover potential therapeutic targets for such patients. Animal models with more severe memory and functional dysfunction following IVH would be the best to study these changes. The senior author&#39;s lab has also been investigating specifically the role of high intracranial pressure (ICP) in the development of memory deficits in IVH rat models. Hence, methods to precisely measure ICPs during IVH were important to investigate. Herein, we report on methods of precisely measuring ICPs in an IVH rat model. Although ICP monitoring has previously been used in traumatic ICH as well as subarachnoid hemorrhage animal models, ICP monitoring in spontaneous IVH rodent&#160;models is not as commonly reported in the literature7,8. Hence, the experimental design presented herein included three groups of Sprague Dawley rats: sham, standard 200 &#181;l intraventricular hemorrhage, and vehicle control. For IVH group, an autologous intraventricular blood injection model was used. For vehicle control animals, intraventricular injection of sterile Lactated Ringer&#39;s solution was used. ICPs, mean arterial pressures (MAPs), and cerebral perfusion pressures (CPPs) were recorded intraoperatively, and results are reported herein.

<table><tbody><tr><td>0.25% bupivacaine</td><td>Hospira, Inc.</td><td>409115901</td><td></td></tr><tr><td>1 mL syringe</td><td>Covetrus</td><td>60734</td><td></td></tr><tr><td>10% providine iodine solution</td><td>Aplicare</td><td>MSD093947</td><td></td></tr><tr><td>20 mL syringe</td><td>Covidien</td><td>8881520657</td><td></td></tr><tr><td>22 G needles</td><td>Becton Dickinson</td><td>305155</td><td></td></tr><tr><td>28 G intraventricular needles</td><td>P technologies</td><td>8IC313ISPCXC</td><td>C313I/SPC 28-Gneedles to fit 22-G guide cannula with 6 mm projection</td></tr><tr><td>3-0 silk suture</td><td>Henry Schein, Inc.</td><td>SP116</td><td></td></tr><tr><td>3-way-stopcock</td><td>Merti Medical Systems</td><td>M3SNC</td><td></td></tr><tr><td>4% paraformaldehyde</td><td>Fisher Chemical</td><td>30525-89-4</td><td></td></tr><tr><td>AnyMaze software</td><td>Any-Maze behavioral tracking software</td><td></td><td>Stoelting CO, USA</td></tr><tr><td>Artificial ointment</td><td>Covetrus</td><td>48272</td><td></td></tr><tr><td>Blood collection vials with EDTA</td><td>Becton Dickinson</td><td>367856</td><td></td></tr><tr><td>Bone wax</td><td>CP Medical, Inc.</td><td>CPB31A</td><td></td></tr><tr><td>Carprofen</td><td>Zoetis, Inc.</td><td>54771-8507-1</td><td></td></tr><tr><td>Centrifuge</td><td>Beckman</td><td>BE-GS6R</td><td>Model GS-6R</td></tr><tr><td>Cotton tip applicators</td><td>Covetrus</td><td>71214</td><td></td></tr><tr><td>Drill</td><td>Dremel</td><td>1600A011JA</td><td></td></tr><tr><td>Fiberoptic pressure sensors with readout units</td><td>Opsens Medical</td><td>OPP-M200-X-80SC- 2.0PTFE-XN-100PIT-P1 and LIS-P1-N-62SC</td><td>Opp-M200 packaged pressure sensors with LifeSens system</td></tr><tr><td>Forceps</td><td></td><td>11923-13, 11064-07</td><td></td></tr><tr><td>Gauze</td><td>Covetrus</td><td>71043</td><td></td></tr><tr><td>Guillotine</td><td>World Precision Instruments</td><td>51330</td><td></td></tr><tr><td>Heating pad with rectal thermometer</td><td>CWE, Inc.</td><td>08-13000 ,08-13014</td><td>TC1000 Temperature controller</td></tr><tr><td>Hemostats</td><td></td><td>&amp;nbsp;13013-14,&amp;nbsp; 13008-12</td><td></td></tr><tr><td>Isoflurane</td><td>Covetrus</td><td>29405</td><td></td></tr><tr><td>Lactated ringers</td><td>Baxter Healthcare Corp.</td><td>Y345583</td><td></td></tr><tr><td>Laryngoscope</td><td>American Diagnostic Corporation</td><td>4080</td><td></td></tr><tr><td>Metal clip</td><td>Fine Scientic Tools</td><td>18056-14</td><td></td></tr><tr><td>Micro scissors</td><td>Fine Scientic Tools</td><td>15007-08</td><td></td></tr><tr><td>Microscope</td><td>Leica</td><td>model L2</td><td></td></tr><tr><td>Needle driver</td><td></td><td>12003-15</td><td></td></tr><tr><td>Polyethylene tubing</td><td>Thermo Fisher Scientific</td><td>14-170-12B</td><td>PE-50 tubing</td></tr><tr><td>Rats</td><td>Envigo</td><td></td><td>Sprague Dawley rats 8&amp;ndash;10 months old</td></tr><tr><td>Scalpel</td><td></td><td>&amp;nbsp;10010-00</td><td></td></tr><tr><td>Scissors</td><td></td><td>14090-11</td><td></td></tr><tr><td>Stereotaxic instrument</td><td>Kopf instruments</td><td></td><td>Model 940 with ear bars</td></tr><tr><td>Syringe pump</td><td>KD Scientific</td><td>780100</td><td>Model 100 series</td></tr><tr><td>Tuohy Borst</td><td>Abbott</td><td>23242</td><td></td></tr><tr><td>Ventilator</td><td>Harvard rodent ventilator</td><td>55-0000</td><td>Model 683</td></tr></tbody></table>

intracranial pressure monitoring in nontraumatic intraventricular hemorrhage rodent model

Survivors of intraventricular hemorrhage are often left with significant long-term memory impairment; thus, research utilizing intraventricular hemorrhage animal models is essential. In this study, we sought out ways to measure intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during nontraumatic intraventricular hemorrhage in rats. The experimental design included three Sprague Dawley groups: sham, standard 200 &#181;l intraventricular hemorrhage, and vehicle control groups. By introducing an intraparenchymal fiberoptic pressure sensor, precise intracranial pressure measurements were obtained in all groups. Cerebral perfusion pressures were calculated with the knowledge of intracranial pressure and mean arterial pressure values. As expected, the intraventricular hemorrhage and vehicle control groups both experienced a rise in the intracranial pressure and subsequent decline in cerebral perfusion pressure during intraventricular injection of autologous blood and artificial cerebrospinal fluid, respectively. The addition of an intraparenchymal fiberoptic pressure sensor is beneficial in monitoring precise intracranial pressure changes.

Intracranial, mean arterial, and cerebral perfusion pressures 
Both ICPs and MAPs were monitored intraoperatively in all animals (Figure 1). Rats were of 8-10 months old with a mean weight of 495 &#177; 17 g. Real-time ICP graphs were also collected (Figure 2). Excluding the sham group, ICPs increased significantly during intraventricular injection in IVH as well as vehicle control groups (Figure 3). I...

Watch this Scientific Journal Video about Intracranial Pressure Monitoring In Nontraumatic Intraventricular Hemorrhage Rodent Model at JoVE.com

Intracranial Pressure Monitoring In Nontraumatic Intraventricular Hemorrhage Rodent Model

Monitoring intracranial pressure in rodent&#160;models of nontraumatic intraventricular hemorrhage is not common in the current literature. Herein, we demonstrate a technique for measuring intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during intraventricular hemorrhage in a rat&#160;animal model.

Survivors of intraventricular hemorrhage are often left with significant long-term memory impairment; thus, research utilizing intraventricular hemorrhage ...

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Neuroscience

2.3K Views.  UC Davis Medical Center. Monitoring intracranial pressure in rodent models of nontraumatic intraventricular hemorrhage is not common in the current literature. Herein, we demonstrate a technique for measuring intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during intraventricular hemorrhage in a rat animal model.

Video: Intracranial Pressure Monitoring In Nontraumatic Intraventricular Hemorrhage Rodent Model

Epidural Intracranial Pressure Measurement in Rats Using a Fiber-optic Pressure Transducer

Elevated intracranial pressure (ICP) is a significant problem in several forms of ischemic brain injury including stroke, traumatic brain injury and cardiac arrest. This elevation may result in further neurological injury, in the form of transtentorial herniation1,2,3,4, midbrain compression, neurological deficit or increased cerebral infarct2,4. Current therapies are often inadequate to control elevated ICP in the clinical setting5,6,7 . Thus there is a need for accurate methods of ICP measurement in animal models to further our understanding of the basic mechanisms and to develop new treatments for elevated ICP.
In both the clinical and experimental setting ICP cannot be estimated without direct measurement. Several methods of ICP catheter insertion currently exist. Of these the intraventricular catheter has become the clinical 'gold standard' of ICP measurement in humans8. This method involves the partial removal of skull and the instrumentation of the catheter through brain tissue. Consequently, intraventricular catheters have an infection rate of 6-11%9. For this reason, subdural and epidural cannulations have become the preferred methods in animal models of ischemic injury. 
Various ICP measurement techniques have been adapted for animal models, and of these, fluid-filled telemetry catheters10 and solid state catheters are the most frequently used11,12,13,14,15. The fluid-filled systems are prone to developing air bubbles in the line, resulting in false ICP readings. Solid state probes avoid this problem (Figure 1). An additional problem is fitting catheters under the skull or into the ventricles without causing any brain injury that might alter the experimental outcomes. Therefore, we have developed a method that places an ICP catheter contiguous with the epidural space, but avoids the need to insert it between skull and brain. 
An optic fibre pressure catheter (420LP, SAMBA Sensors, Sweden) was used to measure ICP at the epidural location because the location of the pressure sensor (at the very tip of the catheter) was found to produce a high fidelity ICP signal in this model. There are other manufacturers of similar optic fibre technologies13 that may be used with our methodology. Alternative solid state catheters, which have the pressure sensor located at the side of the catheter tip, would not be appropriate for this model as the signal would be dampened by the presence of the monitoring screw. 
Here, we present a relatively simple and accurate method to measure ICP. This method can be used across a wide range of ICP related animal models.

A novel technique to record the pressures within the skull is described. The minimally invasive method uses a fibre-optic pressure sensing system to accurately measure intracranial pressure (ICP) in anaesthetized rats without causing significant brain trauma. The technique may be used in a wide range of experimental models.

Elevated intracranial pressure (ICP) is a significant problem in several forms of ischemic brain injury including stroke, traumatic brain injury and ...

Medicine

Optimized System for Cerebral Perfusion Monitoring in the Rat Stroke Model of Intraluminal Middle Cerebral Artery Occlusion

The translational potential of pre-clinical stroke research depends on the accuracy of experimental modeling. Cerebral perfusion monitoring in animal models of acute ischemic stroke allows to confirm successful arterial occlusion and exclude subarachnoid hemorrhage. Cerebral perfusion monitoring can also be used to study intracranial collateral circulation, which is emerging as a powerful determinant of stroke outcome and a possible therapeutic target. Despite a recognized role of Laser Doppler perfusion monitoring as part of the current guidelines for experimental cerebral ischemia, a number of technical difficulties exist that limit its widespread use. One of the major issues is obtaining a secure and prolonged attachment of a deep-penetration Laser Doppler probe to the animal skull. In this video, we show our optimized system for cerebral perfusion monitoring during transient middle cerebral artery occlusion by intraluminal filament in the rat. We developed in-house a simple method to obtain a custom made holder for twin-fibre (deep-penetration) Laser Doppler probes, which allow multi-site monitoring if needed. A continuous and prolonged monitoring of cerebral perfusion could easily be obtained over the intact skull.

Cerebral perfusion monitoring has been demonstrated to improve accuracy in ischemic stroke models. Technical difficulties often limit the use of this essential tool for cerebrovascular research. In this video, an optimized system is shown to obtain a single or multi-site hemodynamic monitoring during intraluminal middle cerebral artery occlusion in rats.

The translational potential of pre-clinical stroke research depends on the accuracy of experimental modeling. Cerebral perfusion monitoring in animal ...

A Murine Model of Subarachnoid Hemorrhage

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

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

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

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 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 Bedside, Single Burr Hole Approach to Multimodality Monitoring in Severe Brain Injury

Intracranial pressure (ICP) monitoring is a cornerstone of the intensive care management of patients with severe acute brain injuries, including traumatic brain injury. While elevations in ICP are common, data regarding the measurement and treatment of these ICP elevations are conflicting. There is increasing recognition that changes in the balance between supply and demand of brain tissue are critically important and therefore the measurement of multiple modalities is required. Approaches are not standard, and therefore this article provides a description of a bedside, single burr hole approach to multimodality monitoring that allows the passage of probes designed to measure not only ICP but brain tissue oxygen, blood flow, and intracranial electroencephalography. Patient selection criteria, operative procedures, and practical considerations for securing probes during critical care are described. This method is readily performed, safe, secure, and flexible for the adoption of a variety of multimodality monitoring approaches aimed at detecting or preventing secondary brain injuries.

A method of recording multimodality monitoring signals in patients with severe brain injuries using a bedside, single burr hole technique is described.

Intracranial pressure (ICP) monitoring is a cornerstone of the intensive care management of patients with severe acute brain injuries, including traumatic ...

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

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

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

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

Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

Erectile dysfunction (ED) is defined as the inability to attain or keep an erection of the penis, and this has become a prevalent male sexual disorder. Rodents are employed by many studies to research the physiology/pathology of erectile function. Erectile function in rodents can be evaluated by measuring the intracavernosal pressure (ICP). In practice, ICP can be monitored following electrical stimulation of the cavernous nerves (CNs). The arterial pressure of the carotid artery (the mean arterial pressure) is used as the reference for ICP. Using ICP recording protocols, many key parameters of erectile function can be measured from the ICP response curve. The ICP measurement provides more information than the apomorphine-induced penile erection test, and is cheaper than telemetric monitoring of the corpus spongiosum penis, making this method the most popular one to evaluate erectile function. However, compared to the easily-performed APO-induced erectile function test, successful ICP recordings require attention to detail, practice, and adherence to the operation method. In this work, an introduction to ICP recording in rats is provided to complement the procedure efficiently.

Intracavernosal pressure recording (ICP) is an important method to evaluate the erectile function of experimental animals. Here, a detailed protocol is demonstrated for the recording procedure of ICP by catheterizing the crura penis and then electrically stimulating the cavernous nerves in rats.

Erectile dysfunction (ED) is defined as the inability to attain or keep an erection of the penis, and this has become a prevalent male sexual disorder. ...

Biology

Modeling Posthemorrhagic Hydrocephalus of Prematurity in Rats

Posthemorrhagic hydrocephalus of prematurity (PHHP) is a serious sequela of severe intraventricular hemorrhage (IVH) in very preterm infants less than 32 weeks gestational age (GA). PHHP is defined by the accumulation of cerebrospinal fluid (CSF) associated with clinical symptoms of elevated intracranial pressure (ICP). Infants with PHHP suffer lifelong shunt dependence, with half requiring repeat surgery in the first year of life and many requiring multiple additional surgeries throughout the lifespan. Prenatal chorioamnionitis predisposes preterm infants to severe IVH and the need for surgical treatment of PHHP trends with neonatal sepsis. These clinical features suggest that systemic inflammation is an integral component of PHHP pathophysiology. 
Here, we define an animal model that recapitulates all clinical aspects and essential features of PHHP in rats. The goal of this protocol is to illustrate how in utero chorioamnionitis and postnatal IVH using lysed red blood cells can be combined to yield PHHP. This preclinical approach yields progressive macrocephaly and domed craniums, elevated intracranial pressure, and ventriculomegaly that can be detected via magnetic resonance imaging (MRI) or via microscopy. In addition to sustained disruption in CSF dynamics, rats also have cognitive delay and functional disability into adulthood. Accordingly, this preclinical platform facilitates unique and unparalleled translational studies of PHHP that can incorporate molecular, cellular, biochemical, histologic, imaging, and functional outcome measures. It can also be used for rigorous analysis of the choroid plexus, ependymal motile cilia, and glymphatic system in parallel. Last, it can also be an invaluable preclinical tool for the investigation of novel surgical intervention strategies and non-surgical therapeutic approaches for the treatment of hydrocephalus.

Posthemorrhagic hydrocephalus of prematurity (PHHP) can be modeled in neonatal rats by combining chorioamnionitis and intraventricular hemorrhage. The combination of these prenatal and postnatal events accurately recapitulates the clinical hallmarks of PHHP, including macrocephaly, ventriculomegaly, and elevated intracranial pressure, through the lifespan.

Posthemorrhagic hydrocephalus of prematurity (PHHP) is a serious sequela of severe intraventricular hemorrhage (IVH) in very preterm infants less than 32 ...

Establishing a Subarachnoid Hemorrhage Rat Model Induced by Autologous Blood Injection

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