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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>&#160;13013-14,&#160; 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&#8211;10 months old</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td></td>
			<td>&#160;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>Touhy 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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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

Intracranial Injection of Adeno-associated Viral Vectors

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific subsets of cells in different brain regions both in vivo and in brain sections. Unlike the injection of fluorescent dyes, viral labeling offers targeting of individual cell types and is less expensive and time consuming than establishing transgenic mouse lines. In this technique, an adeno-associated viral (AAV) vector is injected intracranially using stereotaxic coordinates, a micropipette and an automated pump for precise delivery of AAV to the desired area with minimal damage to the surrounding tissue. Injection parameters can be tailored to individual experiments by adjusting the animal age at injection, injection location, volume of injection, rate of injection, AAV serotype and the promoter driving gene expression. Depending on the conditions chosen, virally-induced transgene expression can allow visualization of groups of cells, individual cells or fine cellular processes, down to the level of dendritic spines. The experiment shown here depicts an injection of double-stranded AAV expressing green fluorescent protein for the labeling of neurons and glia in the mouse primary visual cortex.

Here we present the intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex.

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific ...

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% ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Long-term Continuous EEG Monitoring in Small Rodent Models of Human Disease Using the Epoch Wireless Transmitter System

Many progressive neurologic diseases in humans, such as epilepsy, require pre-clinical animal models that slowly develop the disease in order to test interventions at various stages of the disease process. These animal models are particularly difficult to implement in immature rodents, a classic model organism for laboratory study of these disorders. Recording continuous EEG in young animal models of seizures and other neurological disorders presents a technical challenge due to the small physical size of young rodents and their dependence on the dam prior to weaning. Therefore, there is not only a clear need for improving pre-clinical research that will better identify those therapies suitable for translation to the clinic but also a need for new devices capable of recording continuous EEG in immature rodents. Here, we describe the technology behind and demonstrate the use of a novel miniature telemetry system, specifically engineered for use in immature rats or mice, which is also effective for use in adult animals.

Here we demonstrate the use of a wireless enabling technology for electroencephalogram (EEG) in neonatal rodent models of human disease. With telemetry, there are no encumbering connections, thus allowing natural behaviors.

Many progressive neurologic diseases in humans, such as epilepsy, require pre-clinical animal models that slowly develop the disease in order to test ...

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 ...

Intracranial Pharmacotherapy and Pain Assays in Rodents

Pain is a salient sensory experience with affective and cognitive dimensions. However, central mechanisms for pain remain poorly understood, hindering the development of effective therapeutics. Intracranial pharmacology presents an important tool for understanding the molecular and cellular mechanisms of pain in the brain, as well as for novel treatments. Here we present a protocol that integrates intracranial pharmacology with pain behavior testing. Specifically, we show how to infuse analgesic drugs into a select brain region, which may be responsible for pain modulation. Furthermore, to determine the effect of the candidate drug in the central nerve system, pain assays are performed after intracranial treatment. Our results demonstrate that intracranial administration of analgesic drugs in a targeted region can provide relief of pain in rodents. Thus, our protocol successfully demonstrates that intracranial pharmacology, combined with pain behavior testing, can be a powerful tool for the study of pain mechanisms in the brain.

Here we present a protocol to perform intracranial pharmacological experiments followed by pain behavior assays in rodents. This protocol allows researchers to deliver molecular and cellular targets in the brain, for pharmacologic agents in the treatment of pain.

Pain is a salient sensory experience with affective and cognitive dimensions. However, central mechanisms for pain remain poorly understood, hindering the ...

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

Translaminar Autonomous System Model for the Modulation of Intraocular and Intracranial Pressure in Human Donor Posterior Segments

There is a current unmet need for a new preclinical human model that can target disease etiology ex vivo using intracranial pressure (ICP) and intraocular pressure (IOP) which&#160;can identify various pathogenic paradigms related to the glaucoma pathogenesis. Ex vivo human anterior segment perfusion organ culture models have previously been successfully utilized and applied as effective technologies for the discovery of glaucoma pathogenesis and testing of therapeutics. Preclinical drug screening and research performed on ex vivo human organ systems can be more translatable to clinical research. This article describes in detail the generation and operation of a novel ex vivo human translaminar pressure model called the translaminar autonomous system (TAS). The TAS model can independently regulate ICP and IOP using human donor posterior segments. The model allows for studying pathogenesis in a preclinical manner. It can reduce the use of living animals in ophthalmic research. In contrast to in vitro experimental models, optic nerve head (ONH) tissue structure, complexity, and integrity can also be maintained within the ex vivo TAS model.

We describe and detail the use of the translaminar autonomous system. This system utilizes the human posterior segment to independently regulate the pressure inside the segment (intraocular) and surrounding the optic nerve (intracranial) to generate a translaminar pressure gradient that mimics features of glaucomatous optic neuropathy.

There is a current unmet need for a new preclinical human model that can target disease etiology ex vivo using intracranial pressure (ICP) and intraocular ...

Double Direct Injection of Blood into the Cisterna Magna as a Model of Subarachnoid Hemorrhage

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% of the total stroke-related mortality with an important morbidity in terms of neurological outcome. A delayed cerebral vasospasm (CVS) may occur most often in association with a delayed cerebral ischemia. Different animal models of SAH are now being used including endovascular perforation and direct injection of blood into the cisterna magna or even the prechiasmatic cistern, each exhibiting distinct advantages and disadvantages. In this article, a standardized mouse model of SAH by double direct injection of determined volumes of autologous whole blood into the cisterna magna is presented. Briefly, mice were weighed and then anesthetized by isoflurane inhalation. Then, the animal was placed in a reclining position on a heated blanket maintaining a rectal temperature of 37 &#176;C and positioned in a stereotactic frame with a cervical bend of about 30&#176;. Once in place, the tip of an elongated glass micropipette filled with the homologous arterial blood taken from carotid artery of another mouse of the same age and gender (C57Bl/6J) was positioned at a right angle in contact with the atlanto-occipital membrane by means of a micromanipulator. Then 60 &#181;L of blood was injected in the cisterna magna followed by a 30&#176; downward tilt of the animal for 2 minutes. The second infusion of 30 &#181;L of blood into the cisterna magna was performed 24 h after the first one. The individual follow-up of each animal is carried out daily (careful evaluation of weight and well-being). This procedure allows a predictable and highly reproducible distribution of blood, likely accompanied by intracranial pressure elevation that can be mimicked by an equivalent injection of an artificial cerebral spinal fluid (CSF), and represents an acute to mild-model of SAH inducing low mortality.

We described in this protocol a standardized subarachnoid hemorrhage (SAH) mouse model by a double injection of autologous whole-blood into the cisterna magna. The high degree of standardization of the double-injection procedure represents a middle-to-acute model of SAH with relative safety regarding mortality.

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% ...

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

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

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

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