The authors would like to acknowledge Dr. Abdellatif Benraiss at the University of Rochester for his mentorship and discussions relevant to learning the adult surgical and neonate techniques. The authors would also like to acknowledge Dr. James Lauderdale at UGA for the use of his stereotaxic equipment and discussions related to setting up the methodology for this technique, and the Advancement for Research College Scientists (ARCS) foundation for their support and our NIH support (NINDS grants R01NS096176-02, R01NS097231-01, &#38; F99NS105187-01).

All animal use protocols follow the animal care guidelines of the University of Southern California and the University of Georgia.&#160;Euthanasia methods for pregnant dams and adults are performed according to approved protocols: carbon dioxide asphyxiation, followed by cervical dislocation as a secondary method to ensure euthanasia. Neonatal pups are euthanized by decapitation.
Caution: The following protocol involves handling a pathogenic virus. Proper precaution should be taken while handl...

<ol>
	<li>Dreher, A. 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=Spectrum+of+Disease+and+Outcome+in+Children+with+Symptomatic+Congenital+Cytomegalovirus+Infection.">Spectrum of Disease and Outcome in Children with Symptomatic Congenital Cytomegalovirus Infection.</a> J of Pediatr. 164 (4), 855-859 (2014).</li><li>Lanzieri, T. 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=Long-term+outcomes+of+children+with+symptomatic+congenital+cytomegalovirus+disease.">Long-term outcomes of children with symptomatic congenital cytomegalovirus disease.</a> J of Perinatol. 37 (7), 875-880 (2017).</li><li>Naseer, M. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+WDR62+mutation+causes+primary+microcephaly+in+a+large+consanguineous+Saudi+family.">A novel WDR62 mutation causes primary microcephaly in a large consanguineous Saudi family.</a> Ann Saudi Med. 37 (2), 148-153 (2017).</li><li>Abuelo, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microcephaly+Syndromes.">Microcephaly Syndromes.</a> Semin Pediatr Neurol. 14 (3), 118-127 (2007).</li><li>Nicholas, A. K., 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=WDR62+is+associated+with+the+spindle+pole+and+is+mutated+in+human+microcephaly.">WDR62 is associated with the spindle pole and is mutated in human microcephaly.</a> Nat Genet. 42 (11), 1010-1014 (2010).</li><li>Pulvers, J. N., 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=Mutations+in+mouse+Aspm+(abnormal+spindle-like+microcephaly+associated)+cause+not+only+microcephaly+but+also+major+defects+in+the+germline.">Mutations in mouse Aspm (abnormal spindle-like microcephaly associated) cause not only microcephaly but also major defects in the germline.</a> Proc Natl Acad Sci U S A. 107 (38), 16595-16600 (2010).</li><li>Shao, Q., Herrlinger, S., 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=Zika+virus+infection+disrupts+neurovascular+development+and+results+in+postnatal+microcephaly+with+brain+damage.">Zika virus infection disrupts neurovascular development and results in postnatal microcephaly with brain damage.</a> Development. 143 (22), 4127-4136 (2016).</li><li>Li, C., 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=Zika+Virus+Disrupts+Neural+Progenitor+Development+and+Leads+to+Microcephaly+in+Mice.">Zika Virus Disrupts Neural Progenitor Development and Leads to Microcephaly in Mice.</a> Cell Stem Cell. 19 (5), 672 (2016).</li><li>Miki, T., Fukui, Y., Takeuchi, Y., Itoh, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+quantitative+study+of+the+effects+of+prenatal+X-irradiation+on+the+development+of+cerebral+cortex+in+rats.">A quantitative study of the effects of prenatal X-irradiation on the development of cerebral cortex in rats.</a> Neurosci Res. 23, 241-247 (1995).</li><li>Cragan, J. D., 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=Population-based+microcephaly+surveillance+in+the+United+States,+2009+to+2013:+An+analysis+of+potential+sources+of+variation.">Population-based microcephaly surveillance in the United States, 2009 to 2013: An analysis of potential sources of variation.</a> Birth Defects Res Part A Clin Mol Teratol. 106 (11), 972-982 (2016).</li><li>Cragan, J. D., 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=Baseline+Prevalence+of+Birth+Defects+Associated+with+Congenital+Zika+Virus.">Baseline Prevalence of Birth Defects Associated with Congenital Zika Virus.</a> MMWR Morb Mortal Wkly Rep. 66 (8), 219-220 (2017).</li><li>Mlakar, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zika+Virus+Associated+with+Microcephaly.">Zika Virus Associated with Microcephaly.</a> N Engl J Med. 374 (10), 951-958 (2016).</li><li>Jaenisch, T., Rosenberger, D., Brito, C., Brady, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Risk+of+microcephaly+after+Zika+virus+infection+in+Brazil,+2015+to+2016.">Risk of microcephaly after Zika virus infection in Brazil, 2015 to 2016.</a> Bull World Health Organ. 95 (3), 191-198 (2017).</li><li>Clancy, B., Darlington, R. B., Finlay, B. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translating+developmental+time+across+mammalian+species.">Translating developmental time across mammalian species.</a> Neuroscience. 105 (1), 7-17 (2001).</li><li>Driggers, R. W., 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=Zika+Virus+Infection+with+Prolonged+Maternal+Viremia+and+Fetal+Brain+Abnormalities.">Zika Virus Infection with Prolonged Maternal Viremia and Fetal Brain Abnormalities.</a> N Engl J Med. 374 (22), 2142-2151 (2016).</li><li>Semple, B. D., Blomgren, K., Gimlin, K., Ferriero, D. M., Noble-Haeusslein, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+development+in+rodents+and+humans:+Identifying+benchmarks+of+maturation+and+vulnerability+to+injury+across+species.">Brain development in rodents and humans: Identifying benchmarks of maturation and vulnerability to injury across species.</a> Prog Neurobiol. 106, 1-16 (2013).</li><li>Li, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zika+Virus+Infects+Neural+Progenitors+in+the+Adult+Mouse+Brain+and+Alters+Proliferation.">Zika Virus Infects Neural Progenitors in the Adult Mouse Brain and Alters Proliferation.</a> Cell Stem Cell. 19 (5), 593-598 (2016).</li><li>Vermillion, 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=Intrauterine+Zika+virus+infection+of+pregnant+immunocompetent+mice+models+transplacental+transmission+and+adverse+perinatal+outcomes.">Intrauterine Zika virus infection of pregnant immunocompetent mice models transplacental transmission and adverse perinatal outcomes.</a> Nat. Commun. 8, 14575 (2017).</li><li>Goodfellow, F., 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=Zika+Virus+Induced+Mortality+and+Microcephaly+in+Chicken+Embryos.">Zika Virus Induced Mortality and Microcephaly in Chicken Embryos.</a> Stem Cells Dev. 25 (22), 1-27 (2016).</li><li>Dick, G. W. A., Kitchen, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zika+Virus+(I).+Isolations+and+serological+specificity.">Zika Virus (I). Isolations and serological specificity.</a> Trans R Soc Trop Med Hyg. 46 (5), 509-520 (1952).</li><li>Shao, Q., Herrlinger, S., 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=The+African+Zika+virus+MR-766+is+more+virulent+and+causes+more+severe+brain+damage+than+current+Asian+lineage+and+Dengue+virus.">The African Zika virus MR-766 is more virulent and causes more severe brain damage than current Asian lineage and Dengue virus.</a> Development. , (2017).</li></ol>

Described here is a method for intracerebral inoculation of the ZIKV at embryonic, neonatal, and adult stages for the investigation of ZIKV-induced damages in brain development. While straightforward, there are a few considerations that investigators should take to ensure the quality of the study and the safety of those involved.
DENV is closely related to ZIKV in the flavivirus genus. DENV has not been causally linked with pediatric brain disorders in humans. DENV2 can successfully infect and...

Microcephaly is a condition resulting from defective brain development characterized by smaller than average head size in newborns. Children with microcephaly exhibit a range of symptoms which can include developmental delay, seizure, intellectual disability, hearing loss, vision problems, and problems with movement and balance, among others, depending on the severity of the disease and cause1,2,3. This condition is multifactorial in nature, with genetic, infectious agent, and environmental factors linked to causing microcephaly4,5,6,7,8,9. Prior to the 2015 - 2016 ZIKV outbreak, 8 children out of 10,000 births were diagnosed with microcephaly in the United States according to the CDC10. On February 1st of 2016 the World Health Organization declared the Zika virus a Public Health Emergency of International Concern due to the alarming increase in microcephaly diagnoses associated with ZIKV infection in mothers11,12. A recent study from the CDC on ZIKV cases in the United States suggests that maternal ZIKV infection results in a 20-fold increased risk for a child to develop microcephaly compared to uninfected individuals, and 4% of ZIKV infected mothers from the USA have resulted in children with microcephaly11. The rate of microcephaly-associated birth defects during pregnancy from ZIKV infection in Brazil have been reported to have affected up to 17% of babies in infected mothers, indicating that other factors in Latin America may be contributing to the increased risk13. While we know that the ZIKV can cause microcephaly and pathogenesis in the neural progenitor cell (NPC) population7,8,14, the complete pathogenesis of ZIKV in the developing brain remains elusive. It is important to develop animal models to further investigate the disease mechanisms underlying the brain abnormalities associated with ZIKV infection.
To directly study the effect that the ZIKV has on brain development, we first developed a mouse model using intracerebral inoculation of embryonic day 14.5 (E14.5) brain with ZIKV7. This stage was chosen as it is considered representative of the end of the first trimester in human gestation14. Pups can survive up to postnatal day 5 (P5) with this embryonic intracerebral injection method (~ 1 &#181;L of 1.7 x 106 tissue culture infective dose (TCID50/mL)). These postnatal pups exhibit a range of phenotypes similarly observed in infected human infants including enlarged ventricles, neuronal loss, axonal rarefaction, astrogliosis, and microglial activation12,15. A newborn mouse brain is relatively immature, akin to the developmental stage of the human brain at mid-gestation16, and mouse brain development includes a major postnatal component. To study later gestation stage infections, a method for postnatal infection is also described. Neonates infected with ZIKV at P1 are able to survive up to 13 days post-injection. Blood-born adult stage infection has been described in the mouse previously17 but requires the use of the interferon (IFN) regulatory factor (IRF) transcription factors IRF-3, -5, -7 triple knockout strain. This protocol describes a method of inoculating ZIKV intraventricularly to circumvent disabling the antiviral response of the murine model in the adult. While this circumvents the murine immune system, this route of injection does not directly mimic the typical route of infection. To address this discrepancy directly, the experimenter can perform an intrauterine infection of ZIKV instead of the intracranial route. Adopted from previous work18, we have briefly described this technique in this embryonic infection protocol.
The Zika virus strains implemented with this technique include the Mexican isolate MEX1-447,19 and the African isolate MR-766 isolated in 194720. Zika MEX1-44 was isolated in Chiapas, Mexico in January of 2016 from an infected Aedes aegypti mosquito. We obtained this virus with permission through the University of Texas Medical Branch at Galveston (UTMB). In addition, the Dengue virus serotype 2 (DENV2) was inoculated using this technique in a comparison study. DENV2, strain S16803 (sequence GenBank GU289914), was isolated from a patient sample from Thailand in 1974 and passaged in C6/36 cells. The virus was passaged twice in Vero cells by the World Reference Center for Emerging Viruses and Arboviruses (WRCEVA) before mouse injections. This demonstrates that this technique works equally well for diverse strains of ZIKV and other flaviviruses that may have an impact on brain development.

<table>
	<tbody>
		<tr>
			<td>Flexible Drive Shaft Drill Hanging Motor</td>
			<td>Leica</td>
			<td>39416001</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse Stereotax</td>
			<td>Kopf</td>
			<td>04557R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro4 Microsyringe Pump Controller</td>
			<td>WPI</td>
			<td>SYS-MICRO4</td>
			<td></td>
		</tr>
		<tr>
			<td>UMP3 UltraMicroPump</td>
			<td>WPI</td>
			<td>UMP3</td>
			<td></td>
		</tr>
		<tr>
			<td>Modulamp</td>
			<td>Schott</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Luer-lock tubing (19-gauge)</td>
			<td>Hamilton</td>
			<td>90619</td>
			<td></td>
		</tr>
		<tr>
			<td>Melting Point Capillary</td>
			<td>Kimble</td>
			<td>34500-99</td>
			<td>Glass needle</td>
		</tr>
		<tr>
			<td>Fluoro-Max: Red Fluorescent Microspheres</td>
			<td>Thermo Scientific</td>
			<td>R25</td>
			<td>No dilution; Use for practice injections</td>
		</tr>
		<tr>
			<td>10 &#181;L, Model 1701 LT SYR</td>
			<td>Hamilton</td>
			<td>80001</td>
			<td>for embryonic inoculation</td>
		</tr>
		<tr>
			<td>10 &#181;L, Model 1701 RN SYR, Small Removable NDL, 26s ga, 2 in, point style 2</td>
			<td>Hamilton</td>
			<td>80030</td>
			<td>for neonate/adult</td>
		</tr>
		<tr>
			<td>4-0 Ethilon Nylon Sutures</td>
			<td>Ethicon</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral Oil</td>
			<td>VWR</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>micropipette puller</td>
			<td>Sutter Instruments</td>
			<td>P-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette Grinder</td>
			<td>Narishige</td>
			<td>EG-44</td>
			<td></td>
		</tr>
		<tr>
			<td>Fastgreen FCF Dye</td>
			<td>Sigma</td>
			<td>F7252</td>
			<td>inject with 0.5% Dye</td>
		</tr>
		<tr>
			<td>Antibodies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Flavivirus group antigen antibody</td>
			<td>Millipore</td>
			<td>MAB10216</td>
			<td>ms IgG2a 1:400 (Figure 2, Figure 3)</td>
		</tr>
		<tr>
			<td>Pax6</td>
			<td>DBHB</td>
			<td>Pax6-s</td>
			<td>ms IgG1 1:20</td>
		</tr>
	</tbody>
</table>

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.

Representative images of our injection methods for the ZIKV inoculation of embryonic brain are shown in diagrams depicting intracerebral injections (Figure 1A) and intrauterine and intraplacental injections (Figure 1B), illustrating the way the pregnant dam and embryos should be viewed and oriented for surgery (embryonic inoculation protocol). Figure 2A exhibits ZIKV (MEX1-44) infection (immunostaine...

Watch this Scientific Journal Video about Establishing Mouse Models for Zika Virus-induced Neurological Disorders Using Intracerebral Injection Strategies: Embryonic, Neonatal, and Adult at JoVE.com

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

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 ...

The overall goal of the surgical methods presented here is to establish in vivo models of intracranial Zika virus infection. This method can help to address the key question in the field, such as pathology and the mechanism of Zika virus infection in the brain at both early and the later developmental stage as well as adult stage. The main advantage of this technique is that the direction infection of the brain can easily be achieved.The implications of this technique extend toward therapies or diagnoses of Zika virus-induced microcephaly or other neuro-developmental disorders. Although this method can provide insight into Zika virus pathogenesis, it can also be applied to study other flavivirus infection in the brain, such as dengue virus. Just prior to surgery, load the Zika virus injection mix into a prepared glass injection needle by first assembling the gas-tight 50-microliter injection syringe Luer-lock attachment and tubing.Saturate the assembled syringe and tubing with mineral oil. Once saturated, attach the needle and draw approximately six to seven microliters of virus into the needle. After anesthetizing a dam pregnant with E14.5 embryos, test the depth of anesthesia by pinching the toe or tail.When a surgical plane of anesthesia is achieved, insure that the dam is laying supine on a heating pad. Shave the surface of the abdomen and sterilize with iodine and alcohol three times. Drape the skin surrounding the surgical site with sterile cloths to avoid contamination of the incision and the instruments.Pinch and lift the skin of the abdomen with forceps, then use sterile scissors to make a 1 to 1 1/2-centimeter cut in the lower abdomen at the medial-sagittal line and expose the embryos. Next, cut a slit in the middle of a small sterile gauze and place over the surgical opening. Hydrate the opening and gauze with sterile saline.Then, pull the embryos through the slit to rest on the gauze, taking care to not remove more than four or five embryos at once. Hydrate the embryos with sterile saline prior to and throughout the inoculation to ensure they do not dry out. Illuminate the embryo to be injected with a lamp to visualize the head and skull sutures, then gently place a spatula under the head of the embryo to be injected and manipulate the embryo until the head is pushed up directly against the uterine wall.Hold the embryo in place with the non-dominant hand. Use the syringe to inject Zika virus or control medium into the lateral ventricles of the embryo brain. For the E11.5 stage intrauterine injection, use a one-milliliter syringe and a 27 gauge needle to inject Zika virus or a control medium into the intrauterine space.To improve retention of the pregnancy, avoid injecting the two embryos next to the ovaries and the two embryos next to the upper vagina. If performing an intrauterine injection, inject into the intrauterine space or into the placenta. Place the embryos back into the pregnant dam and fill the abdominal cavity with approximately 0.5 milliliters of sterile saline.Suture both the abdominal peritoneal muscle and external skin layer with 4-0 sterile sutures. Keep the dam on a heating pad and monitor while recovering from anesthesia. Thaw the virus on ice, then load the syringe by filling it with saline to reduce the dead volume in the syringe.Draw up to 0.75 microliters of air to separate the saline solution from the injection material. To set up a warmed, humidified recovery chamber, place a closable container with a sterile gauze on a heat box set at 37 degrees Celsius and soak with saline. After setting up the microinjector pump and controller, load the injection syringe with virus and sterilize the pump head mount of the surgical setup with 70%ethanol.Then, after cryo-anesthetizing the neonatal pups for five minutes, ensure sufficient anesthesia for injection by the lack of response to a tail or toe pinch. Place the pup on the head mount. Sterilize the surface of the head with iodine then ethanol, and then wipe dry with an alcohol wipe.Mark lambda with a black marker, then measure the distance from lambda to the edge of the eye and, using stereotaxic coordinates, calculate the 2/5 distance from lambda to the eye. Next, use a 26 gauge needle to carefully and superficially puncture the scalp and skull at the injection location and create an opening for the injection needle. Now lower the injection needle into the puncture site and, once the needle has just breached the skin, calculate a one-millimeter depth for injection into the lateral ventricle using the stereotaxic instrument.Once the needle is lowered to the injection depth, inject one microliter of virus at a rate of 10 nanoliters per second. Wait 30 seconds after injection, and then retract the needle in 0.5-millimeter increments by rotating the screw dorsally, waiting 30 seconds after each increment to reduce leakage of injected virus. After finishing the injection, transfer the pup to the pre-warmed, humidified chamber and monitor until recovered.Immobilize the head of an anesthetized adult mouse in the stereotaxic instrument and place a heating pad under the mouse. After a toe or tail pinch to insure a surgical plane of anesthesia, shave the scalp starting behind the eyes to the beginning of the ears. Sterilize the exposed skin with iodine and 70%ethanol.After making a 0.5-centimeter incision along the medial-sagittal line of the head in the sterilized location to expose bregma, clear the surface of the skull of meningeal tissues while simultaneously pushing the skin away from the injection sites. Align the surgical drill bit with bregma, and then identify the injection site by the stereotaxic coordinates. Using the control foot pedal, slowly drill into the skull until the hole is clear for the needle.Replace the drill bit with the microinjector pump and gas-tight 10-microliter syringe. Leave a small air bubble between the saline solution and the virus and draw up four to five microliters of virus. Lower the needle to 1.5 millimeters below the brain surface from the brain surface and inject one microliter of virus at a rate of 10 nanoliters per second, injecting the one microliter over 1 1/2 minutes.After removing the needle from the brain in increments as before, use forceps to loosen the skin and re-cover the exposed skull. Once the scalp has been sutured with 4-0 sutures, remove the mouse from the stereotax and monitor on a heating pad until recovered. Neuro-progenitor cells are immuno-labeled with Pax6, seen in red, and Zika virus via flavivirus antigens, seen in green, four days after injection at E18.5.The embryonic brain is infected with the MEX1-44 strain of Zika virus across all layers of the developing neocortex. The TCID50 assay using tissue from post-natal day three brain tissue inoculated at E14.5 shows that the Zika virus can efficiently replicate and grow in developing brains. This image shows MEX1-44 Zika virus infection at P13 cerebral cortex after P0 injection.Zika is detected using antibody against flavivirus group antigen seen in red. These images show the fluorescent beads that were used to practice injection location and lateral ventricle injection success in adults. While attempting this procedure, it's important to remember to keep track of the embryos injected, have appropriate controls, and keep the surgical wounds small.Following this procedure, other methods like intraperitoneal or arterial injection can be performed to answer the additional question like how Zika virus cross the placenta. After watching this video, you should have a good understanding of how to generate the in vivo mouse model of Zika virus infection in the brain.

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Neuroscience

Establishing Intracranial Brain Tumor Xenografts With Subsequent Analysis of Tumor Growth and Response to Therapy using Bioluminescence Imaging

Transplantation models using human brain tumor cells have served an essential function in neuro-oncology research for many years. In the past, the most commonly used procedure for human tumor xenograft establishment consisted of the collection of cells from culture flasks, followed by the subcutaneous injection of the collected cells in immunocompromised mice. Whereas this approach still sees frequent use in many laboratories, there has been a significant shift in emphasis over the past decade towards orthotopic xenograft establishment, which, in the instance of brain tumors, requires tumor cell injection into appropriate neuroanatomical structures. Because intracranial xenograft establishment eliminates the ability to monitor tumor growth through direct measurement, such as by use of calipers, the shift in emphasis towards orthotopic brain tumor xenograft models has necessitated the utilization of non-invasive imaging for assessing tumor burden in host animals. Of the currently available imaging methods, bioluminescence monitoring is generally considered to offer the best combination of sensitivity, expediency, and cost. Here, we will demonstrate procedures for orthotopic brain tumor establishment, and for monitoring tumor growth and response to treatment when testing experimental therapies.

Luciferase-modified human brain tumor xenografts can be established intracranially in athymic mice, with subsequent monitoring of tumor growth and response to therapy using bioluminescence imaging. In combination with survival analysis, bioluminescence monitoring is an essential research tool for pre-clinical testing of therapies being considered for treating brain tumors.

Transplantation models using human brain tumor cells have served an essential function in neuro-oncology research for many years. In the past, the most ...

Establishing Embryonic Mouse Neural Stem Cell Culture Using the Neurosphere Assay

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life span of the animal. They can potentially replace cells that are lost in the aging process or in the process of injury and disease. The existence of neural stem cells (NSCs) was first described by Reynolds and Weiss (1992) in the adult mammalian central nervous system (CNS) using a novel serum&#8208;free culture system, the neurosphere assay (NSA). Using this assay, it is also feasible to isolate and expand NSCs from different regions of the embryonic CNS. These in vitro expanded NSCs are multipotent and can give rise to the three major cell types of the CNS. While the NSA seems relatively simple to perform, attention to the procedures demonstrated here is required in order to achieve reliable and consistent results. This video practically demonstrates NSA to generate and expand NSCs from embryonic day 14-mouse brain tissue and provides technical details so one can achieve reproducible neurosphere cultures. The procedure includes harvesting E14 mouse embryos, brain microdissection to harvest the ganglionic eminences, dissociation of the harvested tissue in NSC medium to gain a single cell suspension, and finally plating cells in NSA culture. After 5-7 days in culture, the resulting primary neurospheres are passaged to further expand the number of the NSCs for future experiments.

This video protocol demonstrates the application of the neurosphere assay for the isolation and expansion of neural stem cells from the ganglionic eminences of embryonic day 14-mouse brain.

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life ...

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% ...

Cerebrovascular Casting of the Adult Mouse for 3D Imaging and Morphological Analysis

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). Animal models are necessary for studying the etiopathology and potential therapies of cerebrovascular diseases. Imaging the vasculature in large animals is relatively easy. However, developing vessel imaging methods of murine brain disease models is desirable due to the cost and availability of genetically-modified mouse lines. Imaging the murine cerebral vascular tree is a challenge. In humans and larger animals, the gold standard for assessing the angioarchitecture at the macrovascular (conductance) level is x-ray catheter contrast-based angiography, a method not suited for small rodents.
In this article, we present a method of cerebrovascular casting that produces a durable skeleton of the entire vascular bed, including arteries, veins, and capillaries that may be analyzed using many different modalities. Complete casting of the microvessels of the mouse cerebrovasculature can be difficult; however, these challenges are addressed in this step-by-step protocol. Through intracardial perfusion of the vascular casting material, all vessels of the body are casted. The brain can then be removed and clarified using the organic solvent methyl salicylate. Three dimensional imaging of the brain blood vessels can be visualized simply and inexpensively with any conventional brightfield microscope or dissecting microscope. The casted cerebrovasculature can also be imaged and quantified using micro-computed tomography (micro-CT)1. In addition, after being imaged, the casted brain can be embedded in paraffin for histological analysis.
The benefit of this vascular casting method as compared to other techniques is its broad adaptation to various analytic tools, including brightfield microscopic analysis, CT scanning due to the radiopaque characteristic of the material, as well as histological and immunohistochemical analysis. This efficient use of tissue can save animal usage and reduce costs. We have recently demonstrated application of this method to visualize the irregular blood vessels in a mouse model of adult BAVM at a microscopic level2, and provide additional images of the malformed vessels imaged by micro-CT scan. Although this method has drawbacks and may not be ideal for all types of analyses, it is a simple, practical technique that can be easily learned and widely applied to vascular casting of blood vessels throughout the body.

In this article, we present a simple, practical technique for cerebrovascular casting that is easy to perform and can be utilized to image the vascular tree of the adult mouse brain.

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). ...

Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear. Since there is no cell line that satisfactorily represents mammalian hair cells, research on hair cells relies on primary organ cultures. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice (Figure 1) 1-6. The utricle is a vestibular organ, and the hair cells of the utricle are similar in both structure and function to the hair cells in the auditory organ, the organ of Corti. The adult mouse utricle preparation represents a mature sensory epithelium for studies of the molecular signals that regulate the survival, homeostasis, and death of these cells.
Mammalian cochlear hair cells are terminally differentiated and are not regenerated when they are lost. In non-mammalian vertebrates, auditory or vestibular hair cell death is followed by robust regeneration which restores hearing and balance functions 7, 8. Hair cell regeneration is mediated by glia-like supporting cells, which contact the basolateral surfaces of hair cells in the sensory epithelium 9, 10. Supporting cells are also important mediators of hair cell survival and death 11. We have recently developed a technique for infection of supporting cells in cultured utricles using adenovirus. Using adenovirus type 5 (dE1/E3) to deliver a transgene containing GFP under the control of the CMV promoter, we find that adenovirus specifically and efficiently infects supporting cells. Supporting cell infection efficiency is approximately 25-50%, and hair cells are not infected (Figure 2). Importantly, we find that adenoviral infection of supporting cells does not result in toxicity to hair cells or supporting cells, as cell counts in Ad-GFP infected utricles are equivalent to those in non-infected utricles (Figure 3). Thus adenovirus-mediated gene expression in supporting cells of cultured utricles provides a powerful tool to study the roles of supporting cells as mediators of hair cell survival, death, and regeneration.

Mechanosensory hair cells are the receptor cells of the inner ear. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice. We present the dissection of the adult mouse utricle, and we demonstrate adenovirus-mediated infection of supporting cells in cultured utricles.

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear.  Since there ...

Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. Generally, the dyes are applied as solid crystals or by local pressure injection using glass pipettes. However, this can result in dilution of the dye and reduced labeling intensity, particularly when several hours are required for dye diffusion. Here we demonstrate a simple and low-cost technique for introducing fluorescent and ion-sensitive dyes into neurons using a polyethylene suction pipette filled with the dye solution. This method offers a reliable way for maintaining a high concentration of the dye in contact with axons throughout the loading procedure.

We describe a simple and low cost technique for introducing high concentration of fluorescent and calcium-sensitive dyes into neurons or any neuronal tract using a polyethylene suction pipette.

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

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 ...

Progenitor-derived Oligodendrocyte Culture System from Human Fetal Brain

Differentiation of human neural progenitors into neuronal and glial cell types offers a model to study and compare molecular regulation of neural cell lineage development. In vitro expansion of neural progenitors from fetal CNS tissue has been well characterized. Despite the identification and isolation of glial progenitors from adult human sub-cortical white matter and development of various culture conditions to direct differentiation of fetal neural progenitors into myelin producing oligodendrocytes, acquiring sufficient human oligodendrocytes for in vitro experimentation remains difficult. Differentiation of galactocerebroside+ (GalC) and O4+ oligodendrocyte precursor or progenitor cells (OPC) from neural precursor cells has been reported using second trimester fetal brain. However, these cells do not proliferate in the absence of support cells including astrocytes and neurons, and are lost quickly over time in culture. The need remains for a culture system to produce cells of the oligodendrocyte lineage suitable for in vitro experimentation.
Culture of primary human oligodendrocytes could, for example, be a useful model to study the pathogenesis of neurotropic infectious agents like the human polyomavirus, JCV, that in vivo infects those cells. These cultured cells could also provide models of other demyelinating diseases of the central nervous system (CNS). Primary, human fetal brain-derived, multipotential neural progenitor cells proliferate in vitro while maintaining the capacity to differentiate into neurons (progenitor-derived neurons, PDN) and astrocytes (progenitor-derived astrocytes, PDA) This study shows that neural progenitors can be induced to differentiate through many of the stages of oligodendrocytic lineage development (progenitor-derived oligodendrocytes, PDO). We culture neural progenitor cells in DMEM-F12 serum-free media supplemented with basic fibroblast growth factor (bFGF), platelet derived growth factor (PDGF-AA), Sonic hedgehog (Shh), neurotrophic factor 3 (NT-3), N-2 and triiodothyronine (T3). The cultured cells are passaged at 2.5e6 cells per 75cm flasks approximately every seven days. Using these conditions, the majority of the cells in culture maintain a morphology characterized by few processes and express markers of pre-oligodendrocyte cells, such as A2B5 and O-4. When we remove the four growth factors (GF) (bFGF, PDGF-AA, Shh, NT-3) and add conditioned media from PDN, the cells start to acquire more processes and express markers specific of oligodendrocyte differentiation, such as GalC and myelin basic protein (MBP). We performed phenotypic characterization using multicolor flow cytometry to identify unique markers of oligodendrocyte.

Primary, human fetal brain-derived, multipotential progenitor cells proliferate in vitro while maintaining the capacity to differentiate into neurons and astrocytes. This work shows that neural progenitors can be induced to differentiate through stages of the oligodendrocytic lineage by conditioning with select growth factors.

Differentiation of human neural progenitors into neuronal and glial cell types offers a model to study and compare molecular regulation of neural cell ...

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 ...