The authors thank Mr. Masaaki Kaneta, Ms. Hiromi Suzuki, and Ms. Mitsue Kawashima (Department of Regenerative and Infectious Pathology, Hamamatsu University School of Medicine) for their excellent technical assistance. This work was supported by the Japan Society for the Promotion of Science, KAKENHI Grant Number 23590445.

All the experimental protocols were approved by the Animal Care Committee of Hamamatsu University of School of Medicine.
1. Preparation of MCMV (Smith strain) and Recombinant M32-enhanced Green Fluorescent Protein (EGFP)-MCMV
<ol>
	<li>Generate recombinant M32-EGFP-MCMV according to the method as follows (1.2 - 1.9) and as previously described8.</li>
	<li>Use recombinant viruses derived from the Smith strain of wild-type MCMV (accession number: U68299). Insert EGFP (4,361 base pairs; bp) between 37,089 and 41,450 bp (M32 - M31 locus) in the MCMV genome by homologous recombination.</li>
	<li>Amplify by polymerase chain reaction the MCMV M32 (41,450 - 39,286 bp, left flanking sequence and M31 (37,089 - 39,246 bp, right flanking sequence gene loci using the following primers: M32, 5&#8242;-CTACTAGCTAGCCTTCCGCGAGTCGCTGTATT-3&#8242; (forward primer), 5&#8242;-CTACTAGCTAGCCTTCCGCGAGTCGCTGTATT-3&#8242; (reverse primer); M31, 5&#8242;-CTAAATTAACTTAAGTCGTCCTCTTCGTCTCACAA-3&#8242; (forward primer), 5&#8242;-AACTAGTCTAGATCGCTCCTGGTTGGTTTTTA-3&#8242; (reverse primer).</li>
	<li>Insert the M32 locus into the EGFP-expressing vector using NheI and BamHI restriction sites. Insert the M31 locus into the M32-EGFP-recombinant plasmid using AflII and XbaI sites. Cleave the M32-EGFP-M31 sequence with NheI and AflII from the M32-EGFP-M31 recombination plasmid and dissolve in H2O.</li>
	<li>Transfect the cleaved construct into the nuclei of MCMV (Smith strain)-infected NIH3T3 cells 24 hr post-infection (hpi) using an electroporation system to induce homologous recombination.</li>
	<li>At 3 days post-infection, co-culture the cells with uninfected mouse embryonic fibroblasts (MEFs) at a ratio of 1/1,000 in six-well plates and screen for EGFP-expressing foci of infected cells.</li>
	<li>Harvest the virus-containing supernatants from the wells containing green fluorescent foci and dilute tenfold. For purification, choose wells that display single green fluorescent foci.</li>
	<li>When all foci resulting from the limiting-dilution infection display EGFP expression, the virus preparation is pure, indicating that no wild-type virus remains. Quantify the virus by the plaque-assay method as previously described17.</li>
	<li>Treat MCMV in certified biosafety cabinets at biosafety level 2 wearing gloves and mask.</li>
	<li>Passage MCMV (Smith strain) and recombinant MCMV in MEFs prepared from 12-day-old ICR mouse embryos as previously described17.</li>
	<li>Remove cells from the supernatants of infected MEF cultures by centrifugation at 3,000 &#215; g for 20 min at 16 &#176;C.</li>
	<li>Ultracentrifuge the supernatants for 40 min at 70,000 &#215; g. Resuspend the pellets containing virions in 1 ml of Tris-buffered saline and transfer onto a preformed linear sorbitol gradient (25% - 70%). Ultracentrifuge again at 70,000 x g for 60 min18.</li>
	<li>Harvest the virion-containing band with a syringe. Pellet the harvested virions by an additional ultracentrifugation step at 70,000 &#215; g for 40 min.</li>
	<li>Resuspend the pellet in 1 ml of phosphate-buffered saline (PBS) and store at -80 &#176;C until the infection experiments.</li>
	<li>Quantify the virus titer by the plaque-assay method as previously described19.</li>
	<li>Visualize the EGFP expression of the recombinant MCMV particles (excitation at 489 nm, emission at 508 nm) by fluorescent microscopy (Figure 1A) and the particle structure by transmission electron microscopy (TEM)8 (Figure 1B).</li>
</ol>
2. Preparation of Nile Red Fluorescent Microbeads
<ol>
	<li>Purchase carboxyl fluorescent Nile red microbeads and place 500 &#181;l of microbeads into tubes according to diameter (0.04 - 0.06 &#181;m, approximately 3.63 &#215; 1012 particles; 0.1 - 0.3 &#181;m, approximately 5.75 &#215; 1010 particles; and 1.7 - 2.2 &#181;m, approximately 6.85 &#215; 107 particles).</li>
	<li>Treat beads with 500 &#181;l of 0.1 M NaOH for approximately 1 day to remove any endotoxins and resuspend the beads in sterile water at RT.</li>
	<li>Adsorb the beads O/N with 10% mouse serum obtained from C57BL/6 mice at RT prior to use.</li>
	<li>To separate aggregates, vortex the beads and sonicate thoroughly before use.</li>
</ol>
3. ICV Injection of MCMV and Fluorescent Microbeads into Neonatal Mice
<ol>
	<li>Maintain normal pregnant ICR mice in a temperature-controlled facility under a 12 hr light/dark cycle. The neonates are designated as P 0.5 on the day of birth.</li>
	<li>Sterilize a 10-&#181;l syringe and a 27 G&#160;needle with 70% alcohol.</li>
	<li>Load the injection solution (5 &#181;l) containing MCMV or fluorescent microbeads into the needle by carefully pulling the plunger of the syringe.</li>
	<li>Restrain neonatal mouse (P 0.5) by putting the mouse on ice for 3 - 4 min. Once the animal is under anesthesia, use the toe-pinch response method to determine the depth of anesthesia.</li>
	<li>Mark the injection site with a non-toxic laboratory pen at a location approximately 0.7 - 1.0 mm lateral to the sagittal suture and 0.7 - 1.0 mm caudal from the neonatal bregma (Figure 2A).</li>
	<li>Insert the needle 2 mm deep, perpendicular to the skull surface at the marked injection site. For reference, mark 2 mm from the tip of needle with a non-toxic maker.</li>
	<li>Slowly inject 5 &#181;l of MCMV (approximately 5 &#215; 105 PFU) into the lateral ventricle without opening the scalp (Figure 2B).</li>
	<li>In another group of mice, inject a 5 &#181;l solution containing fluorescent microbeads (0.1 - 0.3 &#181;m, approximately 5.75 &#215; 108 particles) by the same method.</li>
	<li>Slowly remove the needle 10 - 20 sec after discontinuing the plunger movement to prevent backflow.</li>
	<li>To recover, keep the mice for 5 - 10 min in a warm container until movement and general responsiveness are restored.</li>
	<li>Harvest the brains as described in section 5 at a range of time points (3, 12, 24, 48, and 72 hr) post-injection.</li>
</ol>
4. IV Injection of MCMV or Fluorescent Microbeads into Neonatal Mice
<ol>
	<li>Restrain neonatal mouse (P 0.5) by putting the mouse on ice for 3 - 4 min.</li>
	<li>Use a 1-ml syringe with a 35 G needle to perform the intravenous injection of MCMV in P 0.5 neonates.</li>
	<li>Use a transilluminator (vein finder) to visualize the superficial temporal (facial) vein. Before injection, secure the neonate to the transilluminator using surgical tape (Figure 2C).</li>
	<li>While wearing magnifying glasses (1.5X), slowly infuse 50 &#181;l of MCMV (approximately 5.45 &#215; 109 particles) or fluorescent microbeads (0.04 - 0.06 &#181;m, approximately 3.63 &#215; 1011 particles; 0.1 - 0.3 &#181;m, approximately 5.75 &#215; 109 particles; and 1.7 - 2.2 &#181;m, approximately 6.85 &#215; 106 particles) into the superficial temporal vein (Figure 2D).</li>
	<li>After removing the needle, use gauze containing 70% alcohol to apply pressure to the injection site until the bleeding ceases.</li>
	<li>Give the neonate approximately 5 min in a warm container to recover before returning it to the cage.</li>
	<li>Harvest the brains as described in section 5 at a range of time points (3, 12, 24, and 72 hr) post-injection.</li>
</ol>
5. Brain Tissue Sample Preparation for Paraffin Sections
<ol>
	<li>Place the injected neonates in a small plastic dish&#160;on crushed ice.</li>
	<li>Once the animal is under anesthesia, use the toe-pinch response method to determine the depth of anesthesia.</li>
	<li>Make one central or two end horizontal end cuts through the rib cage to open up the thoracic cavity.</li>
	<li>Make a cut in the atrium with sharp scissors. Infuse 4% paraformaldehyde (PFA) solution into the atrium. Stop perfusion when spontaneous movement (PFA dance) and lightened-color of the liver are observed. Do not exsanguinate.</li>
	<li>Dissect the neonate (as previously described20) by first removing the head using a pair of scissors.</li>
	<li>Make a midline incision along the integument from the neck to the nose to expose the skull.</li>
	<li>Place the sharp end of a pair of iris scissors into the foramen magnum on one side, carefully sliding along the inner surface of the skull.</li>
	<li>Make a cut extending to the distal edge of the posterior skull surface, and make an identical cut on the contralateral side. Clear away the skull around the cerebellum.</li>
	<li>Carefully peel back the skull on one side to prevent damage to the brain. Repeat this procedure on the other side of the brain.</li>
	<li>Using a spatula, sever the olfactory bulbs and nervous connections along the ventral surface of the brain.</li>
	<li>Gently separate the brain from the head, trimming any dura that still connect the brain to the skull using scissors, and remove the brain.</li>
	<li>Place the brain in a vial of fixative containing 4% PFA at least 10 times the volume of the brain for 24 hr at 4&#176;C.</li>
	<li>Dehydrate the tissue through a series of graded ethanol baths to displace the water, and then infiltrate with paraffin wax, forming a block. Cut the paraffin block with a microtome into slices 4 &#181;m thick21.</li>
	<li>Deparaffinize and rehydrate the slides of the infected brains22. Proceed to in situ hybridization for paraffin embedded sections.</li>
</ol>
6. Brain Tissue Sample Preparation for Frozen Sections
<ol>
	<li>Remove the brain following the steps outlined in 5.1 - 5.11.</li>
	<li>To observe the fluorescent microbeads and M32-EGFP-MCMV, place the resected brain on flat bottom cryomolds. Add embedding medium to completely cover the brain sample. Snap freeze the cryomolds in precooled n-hexane (-80 &#186;C), using forceps to hold the edge of the cryomolds prior to attaching it to the chuck.</li>
	<li>For sectioning, attach the frozen tissue block on the precooled cryostat chuck. Transfer the frozen tissue with the chuck into a cryostat chamber and lower the temperature to between -10 and -20 &#176;C, and cut the slices approximately 8 - 10 &#181;m thick23.</li>
	<li>Fix the sections with cytofixative (mixture of isopropyl alcohol and polyethylene glycol) by spraying. Air dry the sections for 30 min at RT immediately after spraying, and store them at - 80 &#186;C until further use.</li>
	<li>Equilibrate the sections back to RT and wash three times with PBS.</li>
	<li>Stain the sections with fluorescein isothiocyanate (FITC)-conjugated Griffonia simplicifolia isolectin B4 at a concentration of 1:100 for 10 min in PBS at RT (Figure 5) or with PE-conjugated CD31 antibody at a concentration of 1:100 for 30 min in PBS at RT (Figure 6).</li>
	<li>Wash the sections with PBS three times.</li>
	<li>After the washing, stain the sections with 4&#8242;,6-diamidino-2-phenylindole (DAPI) to visualize cell nuclei. Mount the sections in anti-fade reagent and image DAPI (excitation at 345 nm, emission at 455 nm), EGFP (excitation at 489 nm, emission at 508 nm), and Nile red (excitation at 553 nm, emission at 637 nm) with a fluorescent microscope (Figures 3, 5, 6).</li>
</ol>
7. Fluorescent in situ Hybridization (FISH) for Paraffin Embedded Sections
<ol>
	<li>Prepare the FISH probe labeled directly with fluorophore for DNA in situ hybridization by nick translation using a bacterial artificial chromosome containing the whole MCMV DNA genome (pSM3fr)19. The concentration of the FISH probe is 0.1 &#181;g/&#181;l.</li>
	<li>Deparaffinize and rehydrate the slides of the infected brains22.</li>
	<li>Treat the tissue sections with RNase (100 &#181;g/ml in PBS) to detect viral DNA.</li>
	<li>Perform antigen retrieval with a 0.05% NP40, 0.01 M citrate buffer (pH 6.0) at 95 - 98&#176;C for 20 min. Cool the slides down to RT for 20 min.</li>
	<li>In a glass Coplin staining jar, wash the slides in pure water three times for 2 min each time. Perform an additional antigen retrieval step with a 0.06% pepsin, 0.01 N HCl solution for 5 min at 37 &#176;C.</li>
	<li>Rinse slides three times in pure water for 2 min each time in a glass Coplin staining jar.</li>
	<li>Dehydrate the tissue sections again by transferring the slides from 70% ethanol, to 85% ethanol, and then to 100% ethanol.</li>
	<li>For preparing 10 ml of the hybridization buffer, mix 1.25 ml in situ hybridization salts (3 M NaCl, 100 mM Tris-HCl pH 8.0, 100 mM sodium phosphate pH 6.8, 50 mM EDTA) with 5 ml deionized formamide, 2.5 ml 50% Dextran sulfate, 250 &#181;l 50x Denhardt&#39;s solution, 125 &#181;l 100 mg/ml tRNA, and 875 &#181;l H2O.</li>
	<li>Dilute the DNA probe directly labeled with fluorophores that recognize the whole MCMV genome randomly in the hybridization buffer. The final volume should be 10 &#181;l (7 &#181;l hybridization buffer, 1 &#181;l probe (0.1 &#181;g/&#181;l), and 2 &#181;l distilled water).</li>
	<li>Add the probe mix (10 &#181;l) to each slide and cover with a coverslip (15 &#215; 15 mm). Seal the coverslip with rubber cement. Denature the probe mix for 5 min at 85 &#176;C, and complete the hybridization step O/N at 42 &#176;C.</li>
	<li>Wash the slides with 0.3% NP40, 0.4x SSC at 73&#176;C for 2 min; with 0.1% NP40, 0.4x SSC at 73 &#176;C for 1 min; and with 2x SSC twice.</li>
	<li>Counterstain the nuclei with DAPI (10 ng/ml) and cover the slide with a coverslip.</li>
	<li>Mount the sections in anti-fade reagent and image DAPI (excitation at 345 nm, emission at 455 nm) and EGFP (excitation at 489 nm, emission at 508 nm) with a fluorescent microscope (Figure 4).</li>
</ol>

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</ol>

The authors have nothing to disclose.

In animal models, ICV, intraperitoneal, direct placental, and IV infections have been used to study the pathogenesis of viral encephalitis. We focused on the ICV and IV injection models of neonatal mice for the simplicity of the procedures and the benefit of direct injection of particles into the target region. Although intraperitoneal infection is an easy method, viral particles spread systemically via an indirect process5,24. Direct placental infection is a good method to study embryonic systemic infection. ...

When studying viral encephalitis, the initial distribution of viral particles is very important to understand disease pathogenesis and to identify viral targets in the brain. Most viruses range in size from 20 to 300 nm, although the Pandoravirus is more than 700 nm in size1. The distribution of the viral particles in the acute phase of infection may depend on the size of the particles, the distribution of cellular receptors, or the affinity of the cellular receptors for viruses. In animal models, intracerebroventricular (ICV), intraperitoneal, direct placental, and intravenous (IV) infections have been used to study the pathogenesis of viral encephalitis. ICV inoculation with virus is often used to establish central nervous system (CNS) infections in mice. Studies using this technique report widespread infection, particularly of cells in the periventricular zones and in regions of the brain in direct contact with the cerebrospinal fluid (CSF), similar to the effects of viral ventriculoencephalitis. The small size of adeno-associated virus (AAV) particles (20 - 25 nm in diameter) facilitates their dissemination throughout the brain in ICV infections2-4. Intraperitoneal5, direct placental6, and IV injections7 represent hematogenic systemic administration. The penetration of viral particles through the blood-brain barrier (BBB) allows them to reach the parenchyma of the neonatal brain, representing diffuse microglial nodules8,9.
Cytomegalovirus (CMV) is a common virus that belongs to the herpes virus family. In the United States, 50% - 80% of the people have had CMV infection by age 40. CMV infections are rarely harmful but can cause diseases in immunocompromised patients and fetuses. Of all deliveries, 0.2% - 2% are born with CMV10, resulting in severe symptoms such as microcephaly, periventricular calcification, cerebellar hypoplasia, microphthalmia, and optic nerve atrophy11,12. Furthermore, mental retardation, sensorineural hearing loss, visual defects, seizure, and epilepsy occur in about 10% of non-fatally CMV-infected infants13,14. CNS dysfunction is the most common characteristic symptom of CMV congenital anomaly. More children are permanently disabled each year by congenital CMV than by Down syndrome, fetal alcohol syndrome, or spina bifida15. There are no vaccinations against CMV available at the present, calling for a need of a safe and effective vaccine. Studying the interaction of CMV particles with their receptors in the earliest phase of infection is important to understand the effect of vaccination.
Ventriculoencephalitis and diffuse microglial nodules are the two main pathological characteristics of CMV encephalitis16. It has been uncertain how the CMV particles (150 - 300 nm) spread through the brain in the acute phase of infection and how the distribution of cellular receptors and their affinity for viruses contribute to the viral spread. Kawasaki et al. have evaluated ICV and IV infections from the perspective of the distribution of particles and their receptors (&#946;1 integrin) in the earliest phase of infection. We have found that the dissemination of CMV particles and the expression of &#946;1 integrin are well correlated in the earliest phase of infection in both ICV and IV infections8. ICV infection is a model of ventriculoencephalitis and IV infection is a model of diffuse microglial nodules. Studying the dynamics of viral or fluorescent particles would give useful information on the effect of particle size, viral interactions with cellular receptors, and the mechanism of BBB penetration in the brain. The following protocol could be used to investigate any viral infection and viral vector in the CNS.

<table><tbody><tr><td>Tris; tris(hydroxymethyl)- aminomethane</td><td>Sigma-Aldrich</td><td>T-6791</td><td></td></tr><tr><td>HCl</td><td>Sigma-Aldrich</td><td>H-1758</td><td></td></tr><tr><td>pEGFP-N1 vector&amp;nbsp;</td><td>Clontech</td><td>#6085-1</td><td></td></tr><tr><td>D-sorbitol</td><td>Sigma-Aldrich</td><td>S-1876</td><td></td></tr><tr><td>SPHERO TM Fluorescent Polystyrene Nile Red 0.04 - 0.06</td><td>Spherotech, Inc.</td><td>FP-00556-2</td><td></td></tr><tr><td>SPHERO TM Fluorescent Polystyrene Nile Red 0.1 - 0.3</td><td>Spherotech, Inc.</td><td>FP-0256-2</td><td></td></tr><tr><td>SPHERO TM Fluorescent Polystyrene Nile Red 1.7 - 2.2</td><td>Spherotech, inc.&amp;nbsp;</td><td>FP-2056-2</td><td></td></tr><tr><td>10% mouse serum</td><td>DAKO&amp;nbsp;</td><td>X0910</td><td></td></tr><tr><td>C57BL/6 mouse</td><td>SLC, Inc.</td><td></td><td></td></tr><tr><td>ICR mouse</td><td>SLC, Inc.</td><td></td><td></td></tr><tr><td>Modified Microliter Syringes (7000 Series)</td><td>Hamilton company</td><td></td><td></td></tr><tr><td>35-gauge needle</td><td>Saito Medical</td><td></td><td></td></tr><tr><td>A Wee Sight Transilluminator</td><td>Phillips Healthcare</td><td>1017920</td><td></td></tr><tr><td>O.C.T.Compound</td><td>Sakura Finetek</td><td>4583</td><td></td></tr><tr><td>RNase A</td><td>Sigma-Aldrich</td><td>R4642</td><td></td></tr><tr><td>Nonidet(R) P-40</td><td>Nacalai</td><td>25223-04</td><td></td></tr><tr><td>citrate buffer (pH 6) x 10</td><td>Sigma-Aldrich</td><td>C9999-100ml</td><td></td></tr><tr><td>pepsin</td><td>Sigma-Aldrich</td><td>P6887</td><td></td></tr><tr><td>EDTA</td><td>dojindo</td><td>N001</td><td></td></tr><tr><td>Formamide</td><td>TCI</td><td>F0045</td><td></td></tr><tr><td>Dextran sulfate sodium salt</td><td>Sigma-Aldrich</td><td>42867-5G</td><td></td></tr><tr><td>Denhardt&#039;s Solution (50x)</td><td>ThermoFishcer sceintific</td><td>750018</td><td></td></tr><tr><td>Yeast tRNA (10 mg/ml)</td><td>ThermoFishcer sceintific</td><td>AM7119</td><td></td></tr><tr><td>SSC 20x</td><td>Sigma-Aldrich</td><td>S6639</td><td></td></tr><tr><td>DAPI</td><td>ThermoFishcer sceintific</td><td>D1306</td><td></td></tr><tr><td>n-Hexane</td><td>Sigma-Aldrich</td><td>296090</td><td></td></tr><tr><td>superfrost plus glass</td><td>ThermoFishcer sceintific</td><td>12-55-18</td><td></td></tr><tr><td>Cytokeep II</td><td>Nippon Shoji Co.</td><td></td><td></td></tr><tr><td>FITC-conjugated Griffonia simplicifolia isolectin B4</td><td>Vector laboratories, Inc.</td><td>L1104</td><td></td></tr><tr><td>Anti-Mouse CD31 (PECAM-1) PE</td><td>ebioscience</td><td>12-0311</td><td></td></tr><tr><td>ProLong&amp;nbsp; Gold</td><td>ThermoFishcer sceintific</td><td>P36934</td><td></td></tr><tr><td>BIOREVO</td><td>KEYENCE</td><td>BZ-9000E</td><td></td></tr></tbody></table>

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.

In studies on the pathogenesis of viral encephalitis, the infection method is important. The hematogenous route represents an acute infection of the endothelial cells and pericytes of the brain, while the ICV route represents an acute infection spreading via the CSF through the subarachnoid space, reaching to the meninges and choroid plexus. To analyze the first distribution of particles in acute encephalitis, in situ hybridization detecting the MCMV genomes and direct observatio...

Watch this Scientific Journal Video about Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain at JoVE.com

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

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 ...

intracerebroventricular-intravascular-injection-viral-particles

Research

JoVE Journal

Neuroscience

19.4K Views.  Hamamatsu University School of Medicine. 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. 

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

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 ...

Delivery of Therapeutic Agents Through Intracerebroventricular (ICV) and Intravenous (IV) Injection in Mice

Despite the protective role that blood brain barrier plays in shielding the brain, it limits the access to the central nervous system (CNS) which most often results in failure of potential therapeutics designed for neurodegenerative disorders 1,2. Neurodegenerative diseases such as Spinal Muscular Atrophy (SMA), in which the lower motor neurons are affected, can benefit greatly from introducing the therapeutic agents into the CNS. The purpose of this video is to demonstrate two different injection paradigms to deliver therapeutic materials into neonatal mice soon after birth. One of these methods is injecting directly into cerebral lateral ventricles (Intracerebroventricular) which results in delivery of materials into the CNS through the cerebrospinal fluid 3,4. The second method is a temporal vein injection (intravenous) that can introduce different therapeutics into the circulatory system, leading to systemic delivery including the CNS 5. Widespread transduction of the CNS is achievable if an appropriate viral vector and viral serotype is utilized. Visualization and utilization of the temporal vein for injection is feasible up to postnatal day 6. However, if the delivered material is intended to reach the CNS, these injections should take place while the blood brain barrier is more permeable due to its immature status, preferably prior to postnatal day 2. The fully developed blood brain barrier greatly limits the effectiveness of intravenous delivery. Both delivery systems are simple and effective once the surgical aptitude is achieved. They do not require any extensive surgical devices and can be performed by a single person. However, these techniques are not without challenges. The small size of postnatal day 2 pups and the subsequent small target areas can make the injections difficult to perform and initially challenging to replicate.

Delivery of Therapeutic Agents Through Intracerebroventricular ICV and Intravenous IV Injection in Mice

This article demonstrates two very different methods of injection: 1) into the brain (intracerebroventricular) and 2) systemic (intravenous) to introduce the therapeutic agents into the central nervous system of neonatal mice.

Despite the protective role that blood brain barrier plays in shielding the brain, it limits the access to the central nervous system (CNS) which most ...

Medicine

Intraventricular Transplantation of Engineered Neuronal Precursors for In Vivo Neuroarchitecture Studies

Gene control of neuronal cytoarchitecture is currently the subject of intensive investigation. Described here is a simple method developed to study in vivo gene control of neocortical projection neuron morphology. This method is based on (1) in vitro lentiviral engineering of neuronal precursors as "test" and "control" cells, (2) their co-transplantation into wild-type brains, and (3) paired morphometric evaluation of their neuronal derivatives. Specifically, E12.5 pallial precursors from panneuronal, genetically labeled donors, are employed for this purpose. They are engineered to take advantage of selected promoters and tetON/OFF technology, and they are free-hand transplanted into neonatal lateral ventricles. Later, upon immunofluorescence profiling of recipient brains, silhouettes of transplanted neurons are fed into NeurphologyJ open source software, their morphometric parameters are extracted, and average length and branching index are calculated. Compared to other methods, this one offers three main advantages: it permits achieving of fine control of transgene expression at affordable costs, it only requires basic surgical skills, and it provides statistically reliable results upon analysis of a limited number of animals. Because of its design, however, it is not adequate to address non cell-autonomous control of neuroarchitecture. Moreover, it should be preferably used to investigate neurite morphology control after completion of neuronal migration. In its present formulation, this method is exquisitely tuned to investigate gene control of glutamatergic neocortical neuron architecture. Taking advantage of transgenic lines expressing EGFP in other specific neural cell types, it can be re-purposed to address gene control of their architecture.

Based on in vitro lentiviral engineering of neuronal precursors, their co-transplantation into wild-type brains and paired morphometric evaluation of "test" and "control" derivatives, this method allows accurate modeling of in vivo gene control of neocortical neuron morphology in a simple and affordable way.

Gene control of neuronal cytoarchitecture is currently the subject of intensive investigation. Described here is a simple method developed to study in ...

Genetics

Use of In vivo Imaging to Monitor the Progression of Experimental Mouse Cytomegalovirus Infection in Neonates

Human Cytomegalovirus (HCMV or HHV-5) is a life-threatening pathogen in immune-compromised individuals. Upon congenital or neonatal infection, the virus can infect and replicate in the developing brain, which may induce severe neurological damage, including deafness and mental retardation. Despite the potential severity of the symptoms, the therapeutic options are limited by the unavailability of a vaccine and the absence of a specific antiviral therapy. Furthermore, a precise description of the molecular events occurring during infection of the central nervous system (CNS) is still lacking since observations mostly derive from the autopsy of infected children. Several animal models, such as rhesus macaque CMV, have been developed and provided important insights into CMV pathogenesis in the CNS. However, despite its evolutionary proximity with humans, this model was limited by the intracranial inoculation procedure used to infect the animals and consistently induce CNS infection. Furthermore, ethical considerations have promoted the development of alternative models, among which neonatal infection of newborn mice with mouse cytomegalovirus (MCMV) has recently led to significant advances. For instance, it was reported that intraperitoneal injection of MCMV to Balb/c neonates leads to infection of neurons and glial cells in specific areas of the brain. These findings suggested that experimental inoculation of mice might recapitulate the deficits induced by HCMV infection in children. Nevertheless, a dynamic analysis of MCMV infection of neonates is difficult to perform because classical methodology requires the sacrifice of a significant number of animals at different time points to analyze the viral burden and/or immune-related parameters. To circumvent this bottleneck and to enable future investigations of rare mutant animals, we applied in vivo imaging technology to perform a time-course analysis of the viral dissemination in the brain upon peripheral injection of a recombinant MCMV expressing luciferase to C57Bl/6 neonates.

Use of In vivo Imaging to Monitor the Progression of Experimental Mouse Cytomegalovirus Infection in Neonates

Human Cytomegalovirus (HCMV) infection of neonates represents an important cause of mental retardation, yet the molecular events leading to virus-induced pathogenesis are still poorly understood. To investigate the dynamics of brain infection, we adapted whole-animal in vivo imaging to perform time-course analysis of neonates infected with a luciferase-recombinant virus.

Human Cytomegalovirus (HCMV or HHV-5) is a life-threatening  pathogen in immune-compromised individuals. Upon congenital or neonatal  infection, the virus ...

Immunology and Infection

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for tissue formation during embryonic development and tissue homeostasis during adulthood 1. Currently, great efforts are invested towards controlling the switch of somatic stem cells from expansion to differentiation because this is thought to be fundamental for developing novel strategies for regenerative medicine 1,2. However, a major challenge in the study and use of somatic stem cell is that their expansion has been proven very difficult to control.
Here we describe a system that allows the control of neural stem/progenitor cell (altogether referred to as NSC) expansion in the mouse embryonic cortex or the adult hippocampus by manipulating the expression of the cdk4/cyclinD1 complex, a major regulator of the G1 phase of the cell cycle and somatic stem cell differentiation 3,4. Specifically, two different approaches are described by which the cdk4/cyclinD1 complex is overexpressed in NSC in vivo. By the first approach, overexpression of the cell cycle regulators is obtained by injecting plasmids encoding for cdk4/cyclinD1 in the lumen of the mouse telencephalon followed by in utero electroporation to deliver them to NSC of the lateral cortex, thus, triggering episomal expression of the transgenes 5-8. By the second approach, highly concentrated HIV-derived viruses are stereotaxically injected in the dentate gyrus of the adult mouse hippocampus, thus, triggering constitutive expression of the cell cycle regulators after integration of the viral construct in the genome of infected cells 9. Both approaches, whose basic principles were already described by other video protocols 10-14, were here optimized to i) reduce tissue damage, ii) target wide portions of very specific brain regions, iii) obtain high numbers of manipulated cells within each region, and iv) trigger high expression levels of the transgenes within each cell. Transient overexpression of the transgenes using the two approaches is obtained by different means i.e. by natural dilution of the electroporated plasmids due to cell division or tamoxifen administration in Cre-expressing NSC infected with viruses carrying cdk4/cyclinD1 flanked by loxP sites, respectively 9,15.
These methods provide a very powerful platform to acutely and tissue-specifically manipulate the expression of any gene in the mouse brain. In particular, by manipulating the expression of the cdk4/cyclinD1 complex, our system allows the temporal control of NSC expansion and their switch to differentiation, thus, ultimately increasing the number of neurons generated in the mammalian brain. Our approach may be critically important for basic research and using somatic stem cells for therapy of the mammalian central nervous system while providing a better understanding of i) stem cell contribution to tissue formation during development, ii) tissue homeostasis during adulthood, iii) the role of adult neurogenesis in cognitive functions, and perhaps, iv) better using somatic stem cells in models of neurodegenerative diseases.

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Controlling the expansion of somatic stem cells is a major factor hampering their study and use in therapy. Here we describe a system to temporally control neural stem cells expansion during development and adulthood, which can be used to increase the number of neurons generated in the mouse brain.

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for ...

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 ...

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 ...

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are ...

Biology

A Comprehensive Protocol for Delivering Intrathecal Injections in Neonatal Mice

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and treatments. By directly delivering medications and treatments into the cerebrospinal fluid of the central nervous system, this method achieves higher localized drug concentrations while reducing systemic side-effects compared to other routes such as intravenous, subcutaneous, or intramuscular injections. Its importance extends beyond clinical settings, as intrathecal injection plays a vital role in preclinical studies focused on treating neurogenetic disorders in rodents and other large animals, including non-human primates. However, despite its widespread application, intrathecal injection in young, particularly neonatal pups, poses significant technical challenges due to their small size and fragile nature. Successful and reliable administration of intrathecal injections in newborn mice requires meticulous attention to detail and careful consideration of various factors. Thus, there is a crucial need for a standardized protocol that not only provides instructions but also highlights key technical considerations and good laboratory practices to ensure procedural consistency, as well as the safety and welfare of the animals.
To address this unmet need, we present a detailed and comprehensive protocol for performing intrathecal injections specifically in newborn pups on postnatal day 1 (P1). By following the step-by-step instructions, researchers can confidently perform intrathecal injections in neonatal pups, enabling the accurate delivery of drugs, antisense oligos, and viruses for gene replacement or genome editing-based treatments. Furthermore, the importance of adhering to good laboratory practices is emphasized to maintain the well-being of animals and ensure reliable experimental outcomes. This protocol aims to address the technical challenges associated with intrathecal injections in neonatal mice, ultimately facilitating advances in the field of neurogenetic research that aims to develop potential therapeutic interventions.

Author Spotlight: Intrathecal Injection &#8211; An Efficient and Reliable Delivery Method to Test the Efficacy of Gene Editing in Neonatal Mouse Brains

The present protocol outlines step-by-step instructions for performing intrathecal injections in neonatal mice for gene editing and drug delivery.

Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and ...

An Intracerebroventricular Injection to Analyze Murine Cytomegalovirus Distribution in a Neonatal Mouse Brain

Source: Kawasaki, H., et al. <a href="https://app.jove.com/v/54164">Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain.</a> J. Vis. Exp. (2016).
This video demonstrates the intracerebroventricular injection process for studying murine cytomegalovirus (MCMV) distribution in the neonatal mouse brain. MCMV, engineered to express an enhanced green fluorescent protein (EGFP) for visibility, is injected into the lateral ventricle of a mouse pup&#39;s brain. The injected virus circulates through the cerebrospinal fluid, infecting cells in the periventricular region, including the marginal zone, choroid plexus, and subventricular zone.

1. ICV Injection of MCMV and Fluorescent Microbeads into Neonatal Mice

	Maintain normal pregnant ICR mice in a temperature-controlled facility under a 12 ...

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

Summary

Explore More Videos

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Intraventricular Transplantation of Engineered Neuronal Precursors for In Vivo Neuroarchitecture Studies

Use of In vivo Imaging to Monitor the Progression of Experimental Mouse Cytomegalovirus Infection in Neonates

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

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

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

Intravenous Injections in Neonatal Mice

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

An Intracerebroventricular Injection to Analyze Murine Cytomegalovirus Distribution in a Neonatal Mouse Brain