JMS received funding from NIH/NINDS R01 NS110793 and K12 (Neurosurgeon Research Career Development Program). BAM received funding from NIH/NINDS K08 NS112580-01A1, University of Kentucky Neuroscience Research Priority Area Award, and a Hydrocephalus Association Innovator Award.

NOTE: All animal protocols were approved by the institutions&#39; Animal Care and Use Committee. See the Table of Materials for details about all materials, reagents, equipment, and software used in this protocol.
1. Preparation of hemoglobin and CSF solutions
<ol>
	<li>Prepare a sterile&#160;artificial CSF (aCSF) solution by adding 500 &#181;L of the aCSF solution to a 1.5 mL microtube and store on ice.</li>
	<li>Prepare a sterile 150 mg/mL hemoglobin solution by adding 75 mg of hemoglobin to 500 &#181;L of aCSF in a 1.5 mL microtube and store on ice.</li>
</ol>
2. Preparation of the animal for injection
<ol>
	<li>Turn the heating pad to the medium setting to maintain the body temperature of the rat.</li>
	<li>Anesthetize postnatal day 4 (P4) rats in an induction chamber filled with 3% isoflurane. 
	NOTE: Confirm sufficient anesthesia using the toe/tail-pinch response every 15 min. Monitor anesthesia with visual observation of tissue color, body temperature, and respiratory rate.</li>
	<li>Administer pain relief with a 5 mg/kg subcutaneous carprofen injection to the anesthetized rat.</li>
	<li>Place the anesthetized rat prone in the stereotactic apparatus with the nose positioned in the anesthesia adaptor with a constant flow of 1.5% isoflurane.</li>
	<li>Tighten non-rupture ear bars on the external auditory meatus to secure the head. 
	NOTE: Apply vet ointment to keep the eyes moisturized if the eyes are open at the age of injection.</li>
	<li>Clean the head, alternating with sterile cotton-tipped applicators soaked in betadine and 70% ethanol.
	<ol>
		<li>Touch the betadine-soaked applicator to the center of the scalp and spread the betadine in circles, moving outward.</li>
		<li>Repeat step 2.6.1.1 with the ethanol-soaked applicator.</li>
		<li>Repeat step 2.6.1.1 and step 2.6.1.2 3x.</li>
	</ol>
	</li>
	<li>Apply a sterile surgical drape to protect the surgical field.&#160;</li>
	<li>Using a sterile scalpel, make a 0.3 cm incision vertically down the center of the head to expose the bregma of the skull. 
	NOTE: If injecting from the lambda, expose the lambda of the skull instead of the bregma.</li>
	<li>Use a sterile cotton-tipped applicator to dry the area.</li>
</ol>
3. Setting up the stereotactic injector
<ol>
	<li>Draw the hemoglobin solution prepared in step 1.2 into a 0.3 mL sterile syringe with a 30G needle and place the syringe into the stereotactic injector system. 
	&#8203;NOTE: If generating control conditions, draw aCSF solution prepared in step 1.1 into a 0.3 mL sterile syringe and proceed with the protocol.</li>
	<li>Turn on the stereotactic injector interface and click on the Configuration button to input the injection volume and rate settings.
	<ol>
		<li>Click on Volume and set the volume at 20,000 nL (20 &#181;L).</li>
		<li>Click on Infusion rate and set the rate at 8,000 nL/min (8 &#181;L/min).</li>
	</ol>
	</li>
	<li>Exit Configuration by clicking on the Reset Pos&#160;button.</li>
	<li>Flush the needle tip by clicking on the Infuse button until a small bead of hemoglobin solution emerges at the needle tip.</li>
	<li>Gently wick the hemoglobin solution from the needle tip with a sterile cotton-tipped applicator.</li>
</ol>
4. Animal injection
<ol>
	<li>Set the bregma as zero on the stereotactic injector system by adjusting the mediolateral and anteroposterior positions of the syringe before lowering the tip of the flushed syringe needle to gently touch the skull at the bregma. 
	NOTE: If injecting from the lambda, set lambda as zero.</li>
	<li>Identify the coordinates of choice.
	<ol>
		<li>If injecting from the bregma, in P4 rats described here, use 1.5 mm lateral, 0.4 mm anterior, and 2.0 mm deep from bregma.</li>
		<li>If injecting from the lambda, use the following coordinates for P4 rats: 1.1 mm lateral, 4.6 mm anterior, and 3.3 mm deep from lambda.</li>
	</ol>
	</li>
	<li>Raise the syringe needle 1 cm above the skull to clear the scalp. When the syringe is raised, proceed to set the mediolateral and anteroposterior coordinates.</li>
	<li>Lower the syringe needle to gently touch the skull. Check that the needle is touching the skull.</li>
	<li>Set the dorsoventral coordinate over a 30 s period. 
	NOTE: While setting the dorsoventral coordinate, the needle will puncture the skull. Care must be taken to assure that the syringe passes through the skull without deforming the skull. Skull deformation is avoided by slowly withdrawing the needle along the dorsoventral coordinate if deformation occurs, then placing the needle back along the same trajectory. This allows the needle to pass through the hole in the skull with less force and no deformation.</li>
	<li>On the stereotactic injector interface, click on the Run button to begin injection.</li>
	<li>After the injection is finished, leave the syringe needle in place for 2 min to minimize backflow of the solution.</li>
	<li>Withdraw the syringe slowly along the dorsoventral coordinate over 2&#160;min until the needle tip is 2 cm above the scalp.</li>
	<li>Rotate the stereotactic injector arm away from the operative field.</li>
</ol>
5. Postoperative care
<ol>
	<li>Close the scalp with a 6-0 monofilament suture. Make one simple interrupted suture at the center of the 0.3 cm incision.</li>
	<li>Remove the pup from anesthesia and place it onto a secure area on the heating pad.</li>
	<li>Return the rodent to the home cage to recover from the anesthesia under the care of its dam. 
	NOTE: Timely return to the care of the dam reduces early postoperative mortality.</li>
	<li>Monitor the animals for anesthesia by loss of the righting reflex hourly post surgery for 3 h.</li>
	<li>Monitor the animals daily for 7 days for normal activity, food intake, and weight gain. Monitor the incision site for wound healing, closure, and reappearance of fur at the site of surgery. 
	NOTE: In the rare case that neurological changes such as seizures, central depression, or decreased appetite are observed during monitoring, euthanize the animal using intravascular perfusion or cervical dislocation under anesthesia.</li>
	<li>To prevent infection once the suture is closed and the wound is healing, apply topical triple-antibiotic at the incision site.</li>
</ol>
6. MRI acquisition and quantification
<ol>
	<li>Perform MRI on a 4.7T or 9.4T small-animal scanner.</li>
	<li>Turn the heating pad to the medium setting to maintain the body temperature of the rat.</li>
	<li>Induce anesthesia in a chamber using 3% isoflurane. 
	NOTE: Confirm sufficient anesthesia using the toe/tail-pinch response every 15 min. Monitor anesthesia with visual observation of tissue color, body temperature, and respiratory rate.</li>
	<li>Place the anesthetized rat prone in the MRI with the nose positioned in the anesthesia adaptor with a constant flow of 1.5% isoflurane.</li>
	<li>Perform T2-weighted imaging by selecting a T2-weighted fast spin echo sequence.
	<ol>
		<li>If using a 4.7T MRI scanner, enter the following parameters into the MRI software: repetition&#8239;time = 3,000 ms, echo time = 27.50 ms, number of averages = 3, field of view = 18.0 mm x 18.0 mm, matrix = 128 x 128, number of axial slices = 24, thickness = 0.50 mm.</li>
		<li>If using a 9.4T MRI scanner, enter the following parameters into the MRI software: repetition time = 5,000 ms, echo time = 66.00 ms, echo spacing = 16.50 ms, number of averages = 2, repetitions = 1, rare factor = 8, field of view = 16.0 mm x 16.0 mm, matrix = 256 x 256, number of axial slices = 32, thickness = 0.50 mm.</li>
	</ol>
	</li>
	<li>Click on the Continue button to start the sequence.</li>
</ol>
7. Image processing and analysis
<ol>
	<li>Use native T2-weighted data to analyze brain volume. Use segmentation software to manually delineate the lateral ventricles6. Click on Paintbrush Mode and select square brush style. Adjust the brush size to 1. Click Layout inspector and select axial view. Click zoom to fit. Place the cursor on the image; trace and fill the lateral ventricle space.</li>
	<li>Click on Segmentation in the toolbar | Volume and Statistics to view the segmented volumes.</li>
</ol>

<ol>
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L., Maher, C. O., Muraszko, K. M., Keep, R. 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S., Love, S., Silver, I. A., Porter, H. J., Whitelaw, A. G. L., Thoresen, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Posthemorrhagic+ventricular+dilation+in+the+neonate:+Development+and+characterization+of+a+rat+model.">Posthemorrhagic ventricular dilation in the neonate: Development and characterization of a rat model.</a> Journal of Neuropathology and Experimental Neurology. 62 (3), (2003).</li><li>Balasubramaniam, J., Xue, M., Buist, R. J., Ivanco, T. L., Natuik, S., del Bigio, ., R, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistent+motor+deficit+following+infusion+of+autologous+blood+into+the+periventricular+region+of+neonatal+rats.">Persistent motor deficit following infusion of autologous blood into the periventricular region of neonatal rats.</a> Experimental Neurology. 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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=Strain-specific+differences+in+perinatal+rodent+oligodendrocyte+lineage+progression+and+its+correlation+with+human.">Strain-specific differences in perinatal rodent oligodendrocyte lineage progression and its correlation with human.</a> Developmental Neuroscience. 33 (34), (2011).</li><li>Engelhardt, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+blood-brain+barrier.">Development of the blood-brain barrier.</a> Cell and Tissue Research. 314 (1), (2003).</li><li>Daneman, R., Zhou, L., Kebede, A. A., Barres, B. 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The authors declare that they have no conflicts of interest.

This IVH model utilizing hemoglobin injection allows for the study of the pathology of IVH specifically mediated by hemoglobin. For complementary studies, hemoglobin can also be easily delivered in vitro and does not confound biochemical assays for proteins made by microglia/macrophages that are present in whole blood.
The leading theories of IVH-PHH include the mechanical obstruction of CSF circulation, the disruption of cilia lining the ependymal walls, inflammation, fibrosis, and i...

Neonatal IVH originates from the germinal matrix, a site of rapid cell division that is adjacent to the lateral ventricles of the developing brain. This highly vascular structure is vulnerable to hemodynamic instability related to premature birth. Blood is released into the lateral ventricles in germinal matrix hemorrhage (GMH)-IVH when fragile blood vessels within the germinal matrix rupture. In the case of grade IV IVH, periventricular hemorrhagic infarction may also contribute to the release of blood products within the brain.1 The combination of GMH-IVH may cause PHH, particularly after high-grade hemorrhage (grades III and IV)1. PHH can be treated with the placement of a ventriculoperitoneal shunt, but shunt placement does not reverse the brain injury that may occur from IVH. Although modern neonatal intensive care has lowered rates of IVH2, 3, there are no specific treatments for the brain injury or hydrocephalus caused by IVH once it has occurred. A significant limitation in developing preventative treatments for IVH-induced brain injury and PHH is the incomplete understanding of IVH pathophysiology.
Recently, early CSF levels of key blood breakdown product hemoglobin have been shown to be associated with the later development of PHH in neonates with high-grade IVH4. Furthermore, CSF levels of iron-handling pathway proteins&#8212;hemoglobin, ferritin, and bilirubin&#8212;are associated with ventricle size in neonatal IVH. This was also shown in a multicenter cohort of infants with preterm PHH, where higher ventricular CSF levels of ferritin were associated with larger ventricle size5.
In this study, we developed a clinically relevant model of IVH-induced brain injury and hydrocephalus utilizing hemoglobin injection into the brain ventricles, which allows for quantification of brain injury and PHH and the testing of new therapeutic strategies (Figure 1)6, 7. This IVH model utilizes neonatal rat pups, which are placed under general anesthesia for the duration of the procedure. A midline incision is made on the scalp, and coordinates derived from skull landmarks&#8212;the bregma or lambda&#8212;are used to target the lateral ventricles for injection. Slow injection using an infusion pump delivers hemoglobin into the ventricle. This protocol is easy to use, versatile, and can model different components of IVH that result in PHH.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >0.3 mL insulin syringe</td>
			<td >BD Microfine + Insulin Syringe</td>
			<td >230-4533</td>
			<td >0.3-0.5 mL synringes will work</td>
		</tr>
		<tr >
			<td >1.5 mL microtube</td>
			<td >USA Scientific</td>
			<td >1615-5500</td>
			<td>Lot No. K194642H -3 511</td>
		</tr>
		<tr >
			<td >4.7T MRI</td>
			<td >Agilent/Varian</td>
			<td >4.7T/33 cm</td>
			<td>Agilent/Varian DirectDrive 4.7-T (200-MHz) MRI system</td>
		</tr>
		<tr >
			<td >6-0 monofilament suture</td>
			<td >ETHICON</td>
			<td >667G</td>
			<td></td>
		</tr>
		<tr >
			<td >9.4T MRI</td>
			<td >Bruker</td>
			<td >BioSpec 94/20</td>
			<td>Used in this protocol without the cryoprobe</td>
		</tr>
		<tr >
			<td >Analytical balance</td>
			<td >CCURIS Instruments</td>
			<td >W3200-320</td>
			<td></td>
		</tr>
		<tr >
			<td >Artificial CSF (aCSF)</td>
			<td >Tocris Bioscience</td>
			<td >3525</td>
			<td>Batch No: 72A</td>
		</tr>
		<tr >
			<td >Betadine</td>
			<td >Purdue Products L.P.</td>
			<td >301005-00</td>
			<td >NDC 67618-150-09</td>
		</tr>
		<tr >
			<td >Carprofen (injectable)</td>
			<td >Zoetis Inc.&#160;</td>
			<td>PI 4019448</td>
			<td>Rimadyl</td>
		</tr>
		<tr >
			<td >Ethanol</td>
			<td >Decon Laboratories</td>
			<td >2701</td>
			<td></td>
		</tr>
		<tr >
			<td >Heating pad</td>
			<td >Sunbeam</td>
			<td >E12107-819</td>
			<td>UL 612A, Z-1228-001</td>
		</tr>
		<tr >
			<td >Hemoglobin</td>
			<td >MP Biomedicals</td>
			<td >100714</td>
			<td>LOT NO. SR02321</td>
		</tr>
		<tr >
			<td >Isoflurane</td>
			<td >Piramal Critical Care</td>
			<td >NDC 66794-017-25</td>
			<td></td>
		</tr>
		<tr >
			<td >Isoflurane vaporizer</td>
			<td >VETEQUIP</td>
			<td >911103</td>
			<td></td>
		</tr>
		<tr >
			<td >Light for stereotactic insturment</td>
			<td >Dolan-Jenner industries</td>
			<td >Fiber-Lite MI-150</td>
			<td></td>
		</tr>
		<tr >
			<td >Microinjection syringe pump</td>
			<td >World Precision Instruments</td>
			<td >MICRO21</td>
			<td>Serial 184034 T08K</td>
		</tr>
		<tr >
			<td >MRI software</td>
			<td >Bruker BioSpin</td>
			<td >Paravision 360 3.2</td>
			<td></td>
		</tr>
		<tr >
			<td >Oxygen</td>
			<td >Airgas Healthcare</td>
			<td >UN1072</td>
			<td>LOT NUMBER S1432080XA02</td>
		</tr>
		<tr >
			<td >Sprague Dawley rats</td>
			<td >Charles River Laboratories</td>
			<td >Strain code: 001</td>
			<td></td>
		</tr>
		<tr >
			<td >Stereotactic instrument</td>
			<td >KOPF Instuments</td>
			<td >Model 900LS Lazy Susan</td>
			<td></td>
		</tr>
		<tr >
			<td >Sterile cotton tipped applicator</td>
			<td >Fischerbrand</td>
			<td >23-400-118</td>
			<td></td>
		</tr>
		<tr >
			<td >Surgical blade</td>
			<td >covetrus</td>
			<td >#10</td>
			<td></td>
		</tr>
		<tr >
			<td >Topical triple antibiotic</td>
			<td >Triple Antibiotic Ointment</td>
			<td >NDC 51672-2120-1</td>
			<td></td>
		</tr>
		<tr >
			<td >Ventricle volume quantification software</td>
			<td >ITK-SNAP</td>
			<td >ITK-SNAP 4.0.0 beta</td>
			<td></td>
		</tr>
	</tbody>
</table>

modeling neonatal intraventricular hemorrhage through intraventricular injection of hemoglobin

Neonatal intraventricular hemorrhage (IVH) is a common consequence of premature birth and leads to brain injury, posthemorrhagic hydrocephalus (PHH), and lifelong neurological deficits. While PHH can be treated by temporary and permanent cerebrospinal fluid (CSF) diversion procedures (ventricular reservoir and ventriculoperitoneal shunt, respectively), there are no pharmacological strategies to prevent or treat IVH-induced brain injury and hydrocephalus. Animal models are needed to better understand the pathophysiology of IVH and test pharmacological treatments. While there are existing models of neonatal IVH, those that reliably result in hydrocephalus are often limited by the necessity for large-volume injections, which may complicate modeling of the pathology or introduce variability in the clinical phenotype observed.
Recent clinical studies have implicated hemoglobin and ferritin in causing ventricular enlargement after IVH. Here, we develop a straightforward animal model that mimics the clinical phenotype of PHH utilizing small-volume intraventricular injections of the blood breakdown product hemoglobin. In addition to reliably inducing ventricular enlargement and hydrocephalus, this model results in white matter injury, inflammation, and immune cell infiltration in periventricular and white matter regions. This paper describes this clinically relevant, simple method for modeling IVH-PHH in neonatal rats using intraventricular injection and presents methods for quantifying ventricle size post injection.

The success of injection was confirmed by radiologic&#160;and immunohistochemical means. Animals that underwent hemoglobin injection developed moderate acute ventriculomegaly when assessed via MRI (Figure 2A), with significantly larger lateral ventricles at 24 h and 72 h post hemoglobin injection compared to aCSF-injected animals (Figure 2B,C). While there was no significant difference in lateral ventricle volume between hemoglobin-injected and ...

Watch this Scientific Journal Video about Modeling Neonatal Intraventricular Hemorrhage Through Intraventricular Injection of Hemoglobin at JoVE.com

Modeling Neonatal Intraventricular Hemorrhage Through Intraventricular Injection of Hemoglobin

We present a model of neonatal intraventricular hemorrhage using rat pups that mimics the pathology seen in humans.

Neonatal intraventricular hemorrhage (IVH) is a common consequence of premature birth and leads to brain injury, posthemorrhagic hydrocephalus (PHH), and ...

modeling-neonatal-intraventricular-hemorrhage-through

Research

JoVE Journal

Neuroscience

2.7K Views.  University of Kentucky. We present a model of neonatal intraventricular hemorrhage using rat pups that mimics the pathology seen in humans.

Video: Modeling Neonatal Intraventricular Hemorrhage Through Intraventricular Injection of Hemoglobin

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% ...

Neonatal Subventricular Zone Electroporation

Neural stem cells (NSCs) line the postnatal lateral ventricles and give rise to multiple cell types which include neurons, astrocytes, and ependymal cells1. Understanding the molecular pathways responsible for NSC self-renewal, commitment, and differentiation is critical for harnessing their unique potential to repair the brain and better understand central nervous system disorders. Previous methods for the manipulation of mammalian systems required the time consuming and expensive endeavor of genetic engineering at the whole animal level2. Thus, the vast majority of studies have explored the functions of NSC molecules in vitro or in invertebrates.
Here, we demonstrate the simple and rapid technique to manipulate neonatal NPCs that is referred to as neonatal subventricular zone (SVZ) electroporation. Similar techniques were developed a decade ago to study embryonic NSCs and have aided studies on cortical development3,4 . More recently this was applied to study the postnatal rodent forebrain5-7. This technique results in robust labeling of SVZ NSCs and their progeny. Thus, postnatal SVZ electroporation provides a cost and time effective alternative for mammalian NSC genetic engineering.

We demonstrate a minimally invasive technique referred to as neonatal subventricular zone electroporation. The technique consists of injecting plasmid DNA into the lateral ventricles of neonatal pups and applying electrical current to deliver and genetically manipulate neural stem cells

Neural stem cells (NSCs) line the postnatal lateral ventricles and give rise to multiple cell types which include neurons, astrocytes, and ependymal ...

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 ...

Neonatal Pial Surface Electroporation

Over the past several years the pial surface has been identified as a germinal niche of importance during embryonic, perinatal and adult neuro- and gliogenesis, including after injury. However, methods for genetically interrogating these progenitor populations and tracking their lineages had been limited owing to a lack of specificity or time consuming production of viruses. Thus, progress in this region has been relatively slow with only a handful of investigations of this location. Electroporation has been used for over a decade to study neural stem cell properties in the embryo, and more recently in the postnatal brain. Here we describe an efficient, rapid, and simple technique for the genetic manipulation of pial surface progenitors based on an adapted electroporation approach. Pial surface electroporation allows for facile genetic labeling and manipulation of these progenitors, thus representing a time-saving and economical approach for studying these cells.

The pial surface is a unique progenitor zone in the CNS that is receiving increasing attention. Herein, we detail a method for rapid genetic manipulation of this progenitor zone using a modified electroporation method. This procedure can be used for cellular and molecular investigations of cell lineages and signaling pathways involved in cell differentiation and to elucidate the fate and properties of daughter cells.

Over the past several years the pial surface has been identified as a germinal niche of importance during embryonic, perinatal and adult neuro- and ...

Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene expression in the mouse brain. Here we describe a technique first introduced by Passini and Wolfe for direct intracranial delivery of virally-encoded transgenes into the neonatal mouse brain. In its most basic form, the procedure requires only an ice bucket and a microliter syringe. However, the protocol can also be adapted for use with stereotaxic frames to improve consistency for researchers new to the technique. The method relies on the ability of adeno-associated virus (AAV) to move freely from the cerebral ventricles into the brain parenchyma while the ependymal lining is still immature during the first 12-24 hr after birth. Intraventricular injection of AAV at this age results in widespread transduction of neurons throughout the brain. Expression begins within days of injection and persists for the lifetime of the animal. Viral titer can be adjusted to control the density of transduced neurons, while co-expression of a fluorescent protein provides a vital label of transduced cells. With the rising availability of viral core facilities to provide both off-the-shelf, pre-packaged reagents and custom viral preparation, this approach offers a timely method for manipulating gene expression in the mouse brain that is fast, easy, and far less expensive than traditional germline engineering.

Here we demonstrate a technique for widespread neuronal transduction by intraventricular injection of adeno-associated virus into the neonatal mouse brain. This method provides a rapid and easy way to attain lifelong expression of virally-delivered transgenes.

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene ...

Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the hematogenous route, which involves infection of the endothelial cells and pericytes of the brain. The second is the intracerebroventricular (ICV) route. Once within the central nervous system (CNS), viruses may spread to the subarachnoid space, meninges, and choroid plexus via the cerebrospinal fluid. In experimental models, the earliest stages of CNS viral distribution are not well characterized, and it is unclear whether only certain cells are initially infected. Here, we have analyzed the distribution of cytomegalovirus (CMV) particles during the acute phase of infection, termed primary viremia, following ICV or intravascular (IV) injection into the neonatal mouse brain. In the ICV injection model, 5 &#181;l of murine CMV (MCMV) or fluorescent microbeads were injected into the lateral ventricle at the midpoint between the ear and eye using a 10-&#181;l syringe with a 27 G needle. In the IV injection model, a 1-ml syringe with a 35 G needle was used. A transilluminator was used to visualize the superficial temporal (facial) vein of the neonatal mouse. We infused 50 &#181;l of MCMV or fluorescent microbeads into the superficial temporal vein. Brains were harvested at different time points post-injection. MCMV genomes were detected using the in situ hybridization method. Fluorescent microbeads or green fluorescent protein expressing recombinant MCMV particles were observed by fluorescent microscopy. These techniques can be applied to many other pathogens to investigate the pathogenesis of encephalitis.

Here, we describe a simple method of intracerebroventricular and intravascular injection of viral particles or fluorescent microbeads into the neonatal mouse brain. The localization pattern of the virus and nanoparticles could be detected by microscopic evaluation or by in situ hybridization.

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the ...

Establishing Mouse Models for Zika Virus-induced Neurological Disorders Using Intracerebral Injection Strategies: Embryonic, Neonatal, and Adult

The Zika virus (ZIKV) is a flavivirus currently endemic in North, Central, and South America. It is now established that the ZIKV can cause microcephaly and additional brain abnormalities. However, the mechanism underlying the pathogenesis of ZIKV in the developing brain remains unclear. Intracerebral surgical methods are frequently used in neuroscience research to address questions about both normal and abnormal brain development and brain function. This protocol utilizes classical surgical techniques and describes methods that allow one to model ZIKV-associated human neurological disease in the mouse nervous system. While direct brain inoculation does not model the normal mode of virus transmission, the method allows investigators to ask targeted questions concerning the consequence after ZIKV infection of the developing brain. This protocol describes embryonic, neonatal, and adult stages of intraventricular inoculation of ZIKV. Once mastered, this method can become a straightforward and reproducible technique that only takes a few hours to perform.

Here we describe a method for establishing a model of Zika virus-induced microcephaly in mouse. This protocol includes methods for embryonic, neonatal, and adult-stage intracerebral inoculation of the Zika virus.

The Zika virus (ZIKV) is a flavivirus currently endemic in North, Central, and South America. It is now established that the ZIKV can cause microcephaly ...

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 ...

Minimally Invasive Endoscopic Intracerebral Hemorrhage Evacuation

Intracerebral hemorrhage (ICH) is a subtype of stroke with high mortality and poor functional outcomes, largely because there are no evidence-based treatment options for this devastating disease process. In the past decade, a number of minimally invasive surgeries have emerged to address this issue, one of which is endoscopic evacuation. Stereotactic ICH Underwater Blood Aspiration (SCUBA) is a novel endoscopic evacuation technique performed in a fluid-filled cavity using an aspiration system to provide an additional degree of freedom during the procedure. The SCUBA procedure utilizes a suction device, endoscope, and sheath and is divided into two phases. The first phase involves maximal aspiration and minimal irrigation to decrease clot burden. The second phase involves increasing irrigation for visibility, decreasing aspiration strength for targeted aspiration without disturbing the cavity wall, and cauterizing any bleeding vessels. Using the endoscope and aspiration wand, this technique aims to maximize hematoma evacuation while minimizing collateral damage to the surrounding brain. Advantages of the SCUBA technique include the use of a low-profile endoscopic sheath minimizing brain disruption and improved visualization with a fluid-filled cavity rather than an air-filled one.

This paper details the surgical protocol for minimally invasive endoscopic intracerebral hemorrhage evacuation using the SCUBA technique.

Intracerebral hemorrhage (ICH) is a subtype of stroke with high mortality and poor functional outcomes, largely because there are no evidence-based ...

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% ...