We gratefully acknowledge the support of the National Natural Science Foundation of China (Grant No. 31800903, awarded to W. Zheng) and the Wenzhou Science and Technology Project (no. Y2020426, awarded to Y. Y. Weng) for this work.

All animal procedures described in this study were conducted in accordance with the guidelines outlined in the NIH Guide for the Care and Use of Laboratory Animals and approved by the Institutional Animal Care and Use Committee at Wenzhou Medical University.
1. Animals
<ol>
	<li>Purchase male C57BL/6 mice aged 8-12 weeks and weighing 20-22 g.</li>
	<li>Obtain the transgenic Rosa-LSL (Lox-StoP-Lox)-tdTomato (Ai9) mouse line, and male A2ARflox/flox mice.</li>
	<li>Randomly assign the mice to two groups and house in cages, with a maximum of five mice per cage, under a standard 12 h light/12 h dark cycle for 1 week.</li>
	<li>Provide the mice with sufficient food and water and maintain them at a constant temperature at 25 &#176;C.</li>
</ol>
2. ChP-specific knockdown of A2ARs with AAV2/5-shRNA
<ol>
	<li>Weigh each mouse and write down the values.</li>
	<li>Anesthetize the mice intraperitoneally with 1% pentobarbital sodium at a dosage of 6-8 mL/kg equivalent to 60-80 mg/kg pentobarbital, and place the mouse on a heating pad to keep it warm. 
	NOTE: It is crucial to administer the correct dosage of pentobarbital sodium according to the mouse&#39;s weight to ensure successful surgery. The anesthesia should not be too deep, as it may result in mortality, but should not be too shallow, as the mouse may wake up during the procedure, potentially affecting the effectiveness of the virus injection. Check for the proper anesthesia dose by toe pinch and ensuring that mice do not respond.</li>
	<li>Use a specialized mouse shaver to trim the top hair of the anesthetized mice.</li>
	<li>Apply&#160;eye ointment on both eyes to prevent from drying, and fix the mice onto a stereotaxic apparatus to immobilize the brain. Cover the animal with a sterile drape.</li>
	<li>Thoroughly sterilize the skin of the head and neck of the mice with three rounds of iodophor disinfectant and 75% alcohol to reduce postoperative infection. Then apply 1% lidocaine topically to the scalp of mice and make a small incision to fully expose the skull under a microscope. 
	NOTE: Use sterile instruments throughout the procedure.</li>
	<li>Prepare two 10 &#181;L syringes: one with purchased AAV2/5-A2AR-shRNA (titer: 6.27 &#215; 109 vg/&#181;L) and the other with purchased AAV2/5-Scramble (titer: 6.21 &#215; 109 vg/&#181;L). 
	NOTE: When using the syringe, the front end needs to be connected to a glass capillary, and a range of 10 &#181;L can be obtained by controlling the length of the glass capillary.</li>
	<li>Use the microscope to locate the bregma and lambda and set the coordinate point (AP: -0.58; ML: &#177;1.10; DL: -2.20) on the 10 &#181;L syringe (Figure 1). The coordinate point is based on the mouse brain stereotaxic atlas15.</li>
	<li>Drill a small hole in the skull at the adjusted coordinate point. 
	NOTE: Under the microscope, drilling is performed using a sterile microdrill against the surface of the skull. After drilling, by removing the meninges covering the brain and exposing the underlying brain parenchyma, injury to the blood vessels is prevented. This step prevents the tip of the glass capillary from being broken off by the meninges. The diameter of the drilled hole should be sufficient to allow the needle tip to enter the brain tissue.</li>
	<li>Administer 2 &#181;L of AAV2/5-A2AR-shRNA virus into the brain ventricle via lateral injection at a steady rate of 100 nL/min. Keep the needle in place for 10 min before withdrawing. Note that the virus was administered with unilateral injection.</li>
	<li>Seal the injured skin using medical biofibrin glue promptly to prevent virus loss, which can be rinsed out with CSF after removing the needle if the glue is not applied quickly. Also, do not use cotton buds to wipe the CSF, as it may also cause virus loss.</li>
	<li>To facilitate recovery, place the mouse on a heating pad to maintain the body temperature at&#160;37.0 &#177; 0.5 &#176;C, and apply 1% lidocaine topically to the mouse&#39;s scalp. Allow the mice to recuperate for 2 weeks before inducing the EAE model, as this period is necessary for AAV2/5-A2AR-shRNA virus expression and subsequent A2AR knockdown.</li>
</ol>
3. ChP-specific knockdown of A2AR with a Cre/locus of X-overP1 (Cre/LoxP) system
NOTE: The following procedures can be achieved using the method described previously. Refer to steps 2.1-2.11 for detailed injection methods.
<ol>
	<li>Inject 2 &#181;L of CRE-TAT recombinase into each lateral ventricle of a Rosa-LSL (Lox-StoP-Lox)-tdTomato mouse as the experimental group and 2 &#181;L of sterile phosphate-buffered saline (PBS) as the control group. Using the same protocol as above (section 2), inject 2 &#181;L of CRE-TAT recombinase into each of the lateral ventricles of A2ARflox/flox mice along with 2 &#181;L of sterile PBS as a control group.</li>
	<li>Prepare frozen tissue sections and perform nuclear staining 2 weeks after the CRE-TAT recombinase injection. See steps 4.1-4.3 for more information.</li>
</ol>
4. Transcardial perfusion in mice
<ol>
	<li>Administer 60-80 mg/kg pentobarbital sodium to deeply anesthetize the mice. Confirm the plane of anesthesia by a toe pinch and perform transcardial perfusion using 40 mL of sterile PBS solution followed by 20 mL of 4% paraformaldehyde (PFA). 
	NOTE: Successful perfusion is indicated by a white color in the liver, while the presence of enlarged lungs or PBS outflow from the mouth during perfusion suggests a failure of the procedure.</li>
	<li>Quickly extract the mouse brain, ensuring minimal protein degradation.</li>
	<li>Submerge the mouse brain in 4% PFA/PBS overnight for post-fixation, followed by replacement with 30% sucrose PB solution for 72 h. 
	&#8203;NOTE: It is critical to avoid excessive dehydration of the brain tissue, so the brain should not be left in the sucrose PB solution for too long.</li>
</ol>
5. Frozen tissue sectioning and staining
<ol>
	<li>Embed the previously dehydrated mouse brain in optimal cutting temperature (OCT) glue, freeze the embedded brain, and use a sliding microtome to cut coronal sections with a thickness of 20 &#181;m.</li>
	<li>Place the brain sections on the glass slide. Once the brain sections are dry, store them in a -20 &#176;C refrigerator for subsequent experiments.</li>
	<li>Place each glass slide into a frame case and submerge the frame into a container filled with PBS to thoroughly rinse the slide. Gently rinse the brain slides with PBS solution three times, for 10 min each time. 
	NOTE: Caution should be exercised to prevent the brain sections from falling off the glass slide.</li>
	<li>Stain the brain slides with 4&#8242;,6-diamidino-2-phenylindole (DAPI) solution for 10 min.</li>
	<li>Rinse the brain slides with PBS solution for 5 min.</li>
	<li>Apply one drop of antifade mounting medium to the brain sections.</li>
	<li>Place a coverslip on the brain sections, seal the coverslip with nail polish, and analyze the sections using a conventional fluorescence microscope.</li>
</ol>
6. EAE induction
NOTE: Perform EAE induction after 2 weeks of the shRNA or CRE-TAT recombinase injection11.
<ol>
	<li>Create an aqueous solution by mixing 2.5 mg myelin oligodendrocyte glycoprotein (MOG35-55) with 2 mL of PBS. Prepare a complete Freunds adjuvant (CFA) oil solution by mixing M. tuberculosis (H37Ra) with incomplete Freunds adjuvant (IFA).</li>
	<li>Mix the aqueous and oil solutions in a 1:1 ratio. Use a tee pipe to whip the mixture into an oil-in-water state. 
	NOTE: The tee pipe is also called a three-way pipe. It is a plastic tube that can control the direction of flowing solution to fully mix the MOG35-55 and CFA.</li>
	<li>Use a high-speed homogenizer to make the MOG antigen emulsion for the EAE model under ice bath conditions.</li>
	<li>Anesthetize the mice intraperitoneally with 1% pentobarbital sodium at a dosage of 6-8 mL/kg equivalent to 60-80 mg/kg pentobarbital.</li>
	<li>Subcutaneously inject the MOG antigen emulsion into four different points (neck, back, left and right hips) at a volume of 10 mL/kg for a total of four injections each. 
	NOTE: It is important to carefully select the injection site, as different sites may have varying effects on the morbidity and mortality of mice. Additionally, repeated MOG injections can lead to immune tolerance, so the research team chose a single injection method to prevent this potential issue.</li>
	<li>Immediately inject 500 ng/mL of pertussis toxin (PT) intraperitoneally at a dosage of 5 mg/kg after MOG injection. 
	NOTE: PT increases the permeability of the blood-brain barrier (BBB) and facilitates the infiltration of T cells into the brain.</li>
	<li>After 48 h, intraperitoneally inject the PT solution at the same volume.</li>
</ol>
7. Neurological deficit score
<ol>
	<li>Evaluate and grade the mice daily using a rating scale16 ranging from 0 to 15 to assess the incidence and severity of EAE according to the following neurological deficits: 
	Tail: 0 indicates no signs, 1 represents a half-paralyzed tail, and 2 indicates a fully paralyzed tail. 
	Limbs: 0 indicates no signs, 1 represents a weak or altered gait, 2 indicates paresis, and 3 represents a fully paralyzed limb.</li>
	<li>Assign a quadriplegic animal having complete paralysis&#160;a score of 12, and assign mortality a score of 15.</li>
</ol>
8. Hematoxylin-eosin (H&#38;E) staining
<ol>
	<li>Take the dehydrated mouse brain and embed it in molten paraffin. Allow the paraffin block to cool and solidify for later use. 
	NOTE: The paraffin block should be completely dry and cool to avoid tissue damage.</li>
	<li>Cut the mouse brain paraffin block into 5 &#181;m thick slides. Place the paraffin mouse brain section on a glass slide and dry it in a 60 &#176;C oven for 3 h.</li>
	<li>Submerge the slides sequentially in xylene solution I for 10 min, xylene solution II for 10 min, 100% alcohol I for 3 min, 100% alcohol II for 3 min, 95% alcohol for 3 min, 90% alcohol for 3 min, 80% alcohol for 3 min, 70% alcohol for 3 min, and distilled water for 1 min.</li>
</ol>
9. Quantitative polymerase chain reaction (qPCR) analysis
<ol>
	<li>After perfusing the mice with PBS, remove the ChP from the ventricles and extract RNA with Trizol. Synthesize the cDNA using the first Strand cDNA Synthesis Kit.</li>
	<li>Carry out qPCR analysis using Ex Taq SYBR-green premix and a real-time PCR system. Use the following A2AR primers: forward - GCCATCCCATTCGCCATCA; reverse - GCAATAGCCAAGAGGCTGAAGA.</li>
</ol>

Selective Gene Silencing in the Choroid Plexus

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K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+functions+of+the+choroid+plexus-cerebrospinal+fluid+system.">Development and functions of the choroid plexus-cerebrospinal fluid system.</a> Nature Reviews: Neuroscience. 16 (8), 445-457 (2015).</li><li>Wolburg, H., Paulus, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Choroid+plexus:+biology+and+pathology.">Choroid plexus: biology and pathology.</a> Acta Neuropathologica. 119 (1), 75-88 (2010).</li><li>Solar, P., Zamani, A., Kubickova, L., Dubovy, P., Joukal, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Choroid+plexus+and+the+blood-cerebrospinal+fluid+barrier+in+disease.">Choroid plexus and the blood-cerebrospinal fluid barrier in disease.</a> Fluids Barriers CNS. 17 (1), 35 (2020).</li><li>Marques, 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=The+choroid+plexus+in+health+and+in+disease:+dialogues+into+and+out+of+the+brain.">The choroid plexus in health and in disease: dialogues into and out of the brain.</a> Neurobiology of Disease. 107, 32-40 (2017).</li><li>Myung, 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=The+choroid+plexus+is+an+important+circadian+clock+component.">The choroid plexus is an important circadian clock component.</a> Nature Communications. 9 (1), 1062 (2018).</li><li>Zhang, Y., 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+transgenic+FOXJ1-Cre+system+for+gene+inactivation+in+ciliated+epithelial+cells.">A transgenic FOXJ1-Cre system for gene inactivation in ciliated epithelial cells.</a> American Journal of Respiratory Cell and Molecular Biology. 36 (5), 515-519 (2007).</li><li>Johansson, P. 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B., 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=Cotransporter-mediated+water+transport+underlying+cerebrospinal+fluid+formation.">Cotransporter-mediated water transport underlying cerebrospinal fluid formation.</a> Nature Communications. 9 (1), 2167 (2018).</li><li>Zhu, L., 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=Klotho+controls+the+brain-immune+system+interface+in+the+choroid+plexus.">Klotho controls the brain-immune system interface in the choroid plexus.</a> Proceedings of the National Academy of Sciences. 115 (48), E11388-E11396 (2018).</li><li>Chen, X., 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=Different+serotypes+of+adeno-associated+virus+vector-+and+lentivirus-mediated+tropism+in+choroid+plexus+by+intracerebroventricular+delivery.">Different serotypes of adeno-associated virus vector- and lentivirus-mediated tropism in choroid plexus by intracerebroventricular delivery.</a> Human Gene Therapy. 31 (7-8), 440-447 (2020).</li><li>Konsman, J. 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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Magnetic+brain+stimulation+upregulates+adhesion+and+prevents+Eae:+MMP-2,+ICAM-1,+and+VCAM-1+in+the+choroid+plexus+as+a+target.">Magnetic brain stimulation upregulates adhesion and prevents Eae: MMP-2, ICAM-1, and VCAM-1 in the choroid plexus as a target.</a> International Journal of Neuroscience. 119 (9), 1399-1418 (2009).</li><li>Mills, J. H., Alabanza, L. M., Mahamed, D. A., Bynoe, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+adenosine+signaling+induces+CX3CL1+expression+in+the+brain+to+promote+experimental+autoimmune+encephalomyelitis.">Extracellular adenosine signaling induces CX3CL1 expression in the brain to promote experimental autoimmune encephalomyelitis.</a> Journal of Neuroinflammation. 9, 193 (2012).</li></ol>

The authors state that they have no competing financial interests or other disclosures to declare.

The research presented two distinct approaches for the targeted knockdown of ChP genes. The first approach involved the ICV injection of CRE-TAT, which contains Cre recombinase, into A2ARflox/flox mice. The second approach entailed ICV injection of AAV2/5 carrying shRNA of A2AR. By utilizing these two strategies, the work achieved the selective knockdown of A2AR within the ChP and was able to demonstrate the protective effects of inhibiting A2AR signaling in the ChP ...

The choroid plexus (ChP) was often thought to help maintain brain functional homeostasis by secreting cerebrospinal fluid (CSF) and brain-derived neurotrophic factor (BDNF)1,2. Increasing research over the last three decades has revealed that the ChP represents a distinct pathway for immune cell infiltration into the central nervous system (CNS).
The tight junctions (TJs) of the ChP, composed of a monolayer ChP epithelium, maintain immunological homeostasis by preventing macromolecules and immune cells from entering the brain3. However, under certain pathological conditions, the ChP tissue detects and responds to danger-associated molecular patterns (DAMPs) in the CSF and blood, leading to abnormal immune infiltration and brain dysfunction4,5. Despite its critical role, the ChP&#39;s small size and unique location in the brain make it difficult to study its function without affecting other brain regions. Therefore, manipulating gene expression specifically in the ChP is an ideal approach to understanding its function.
Initially, cyclization recombination enzyme (Cre) transgenic lines, which express Cre under the control of promoters specific to genes expressed in the ChP, were commonly used to delete target genes by breeding with floxed candidate genes6,7,8. For example, the transcription factor Forkhead box J1 (FoxJ1) is exclusively expressed in the ChP epithelium of the prenatal mouse brain7. Thus, the FoxJ1-Cre line was often used to delete genes located in the ChP6,9. However, the success of this strategy relies heavily on the specificity of the promoter. It was gradually discovered that the FoxJ1 expression pattern was not distinctive enough, as FoxJ1 was also present in ciliated epithelial cells in other parts of the brain and peripheral system7. To overcome this limitation, intra-cerebroventricular (ICV) injection of Cre recombinase was performed to deliver recombinase into the ventricles of floxed transgenic lines. This strategy showed high specificity, as evidenced by the presence of tdTomato fluorescence solely in the ChP tissue10,11. However, this method is still limited by the availability of floxed transgenic mouse lines. To address this issue, researchers have employed ICV injection of adeno-associated virus (AAV) to achieve ChP-specific knockdown or the overexpression of target genes12,13. A comprehensive evaluation of different AAV serotypes for ChP infection revealed that AAV2/5 and AAV2/8 exhibit strong infection abilities in the ChP, while not infecting other brain regions. However, AAV2/8 was found to infect the ependyma surrounding ventricles, whereas the AAV2/5 group showed no infection14. This method has the advantage of overcoming the limitations of acquiring floxed transgenic animals.
This article describes a step-by-step protocol for gene knockdown in the ChP using two methods: ICV of AAV2/5 carrying shRNA of the adenosine A2A receptor (A2AR) and Cre recombinase protein consisting of TAT sequence (CRE-TAT) recombinase to achieve ChP-specific knockdown of A2AR. The study findings suggest that knocking down A2AR in the ChP can alleviate experimental autoimmune encephalomyelitis (EAE). This detailed protocol provides useful guidance for ChP function studies and the specific knockdown of genes in the ChP.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>A2ARflox/flox mice</td>
			<td>State Key Laboratory of Ophthalmology, Optometry and Visual Science, Wenzhou Medical University</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AAV2/5-A2AR-ShRNA virus</td>
			<td>Shanghai Heyuan Biotechnology Co. LTD</td>
			<td>pt-4828</td>
			<td></td>
		</tr>
		<tr>
			<td>antifade mounting medium</td>
			<td>Beyotime Biotechnology</td>
			<td>0100-01</td>
			<td></td>
		</tr>
		<tr>
			<td>borosilicate glass capillary</td>
			<td>Beijing Meiyaxian Technology Co. Ltd</td>
			<td>B100-50-10</td>
			<td></td>
		</tr>
		<tr>
			<td>brain stereotaxic apparatus</td>
			<td>RWD, Shenzhen</td>
			<td>69100</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6 mice</td>
			<td>Beijing Vital Charles River Laboratory Animal Technology Company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CRE-TAT recombinase</td>
			<td>Millipore</td>
			<td>SCR508</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Absin</td>
			<td>B25A031</td>
			<td></td>
		</tr>
		<tr>
			<td>frozen slicing machine</td>
			<td>Leica</td>
			<td>CM1950</td>
			<td></td>
		</tr>
		<tr>
			<td>H37Ra</td>
			<td>Becton Dickinson and company</td>
			<td>231141</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton syringe</td>
			<td>Hamilton, American</td>
			<td>P/N:&#160;86259</td>
			<td></td>
		</tr>
		<tr>
			<td>Incomplete Freunds adjuvant</td>
			<td>Sigma</td>
			<td>F5506</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM900</td>
			<td></td>
		</tr>
		<tr>
			<td>MOG35-55</td>
			<td>Suzhou Qiangyao Biotechnology Co., LTD</td>
			<td>4010006243</td>
			<td></td>
		</tr>
		<tr>
			<td>OCT glue</td>
			<td>Epredia</td>
			<td>6502p</td>
			<td></td>
		</tr>
		<tr>
			<td>paraformaldehyde</td>
			<td>Chengdu Kelong Chemical Reagent Company</td>
			<td>30525-89-4</td>
			<td></td>
		</tr>
		<tr>
			<td>pentobarbital sodium</td>
			<td>Boyun Biotech</td>
			<td>PC13003</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette gun</td>
			<td>Eppendorf</td>
			<td>N45014F</td>
			<td></td>
		</tr>
		<tr>
			<td>PrimeScript 1st Strand cDNA Synthesis Kit</td>
			<td>Takara</td>
			<td>&#160;6110A</td>
			<td></td>
		</tr>
		<tr>
			<td>Real- Time PCR System</td>
			<td>BioRad</td>
			<td>CFX96</td>
			<td></td>
		</tr>
		<tr>
			<td>Rosa-LSL (Lox-StoP-Lox)-tdTomato mice</td>
			<td>Jackson Laboratory</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>sucrose</td>
			<td>Sangon Biotech</td>
			<td>A502792-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>super high speed homogenizer</td>
			<td>IKA</td>
			<td>3737025</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizol</td>
			<td>Invitrogen</td>
			<td>15596026</td>
			<td></td>
		</tr>
		<tr>
			<td>xylene solution</td>
			<td>Chengdu Kelong Chemical Reagent Company</td>
			<td>1330-20-7</td>
			<td></td>
		</tr>
	</tbody>
</table>

targeted knockdown of genes in the choroid plexus

The choroid plexus (ChP) serves as a critical gateway for immune cell infiltration into the central nervous system (CNS) under both physiological and pathological conditions. Recent research has shown that regulating ChP activity may offer protection against CNS disorders. However, studying the biological function of the ChP without affecting other brain regions is challenging due to its delicate structure. This study presents a novel method for gene knockdown in ChP tissue using adeno-associated viruses (AAVs) or cyclization recombination enzyme (Cre) recombinase protein consisting of TAT sequence (CRE-TAT). The results demonstrate that after injecting AAV or CRE-TAT into the lateral ventricle, the fluorescence was exclusively concentrated in the ChP. Using this approach, the study successfully knocked down the adenosine A2A receptor (A2AR) in the ChP using RNA interference (RNAi) or Cre/locus of X-overP1 (Cre/LoxP) systems, and showed that this knockdown could alleviate the pathology of experimental autoimmune encephalomyelitis (EAE). This technique may have important implications for future research on the ChP's role in CNS disorders.

ChP-specific A2AR knockdown by ICV injection of AAV2/5-shRNA or CRE-TAT 
The role of A2AR in the ChP as a powerful regulator of neural information in EAE pathogenesis remains unclear. Knocking down ChP-specific A2AR expression could shed light on the A2AR regulatory effects on the central immune system in EAE and other nervous system inflammations. This study used ICV injection of CRE-TAT to decrease A2AR expression in the ChP of A2AR...

Watch this Scientific Journal Video about Insights and Innovations in Gene Expression Manipulation Techniques for Choroid Plexus Research at JoVE.com

Author Spotlight: Insights and Innovations in Gene Expression Manipulation Techniques for Choroid Plexus Research 

Here, we describe a method to selectively alter gene expressions in the choroid plexus while avoiding any impact in other brain areas.

Targeted Knockdown of Genes in the Choroid Plexus

The choroid plexus (ChP) serves as a critical gateway for immune cell infiltration into the central nervous system (CNS) under both physiological and ...

targeted-knockdown-of-genes-in-the-choroid-plexus

Research

JoVE Journal

Neuroscience

799 Views.  Wenzhou Medical University. Here, we describe a method to selectively alter gene expressions in the choroid plexus while avoiding any impact in other brain areas.

Video: Author Spotlight: Insights and Innovations in Gene Expression Manipulation Techniques for Choroid Plexus Research 

Targeted Training of Ultrasonic Vocalizations in Aged and Parkinsonian Rats

Voice deficits are a common complication of both Parkinson disease (PD) and aging; they can significantly diminish quality of life by impacting communication abilities. 1, 2 Targeted training (speech/voice therapy) can improve specific voice deficits,3, 4 although the underlying mechanisms of behavioral interventions are not well understood. Systematic investigation of voice deficits and therapy should consider many factors that are difficult to control in humans, such as age, home environment, age post-onset of disease, severity of disease, and medications. The method presented here uses an animal model of vocalization that allows for systematic study of how underlying sensorimotor mechanisms change with targeted voice training. The ultrasonic recording and analysis procedures outlined in this protocol are applicable to any investigation of rodent ultrasonic vocalizations.
The ultrasonic vocalizations of rodents are emerging as a valuable model to investigate the neural substrates of behavior.5-8 Both rodent and human vocalizations carry semiotic value and are produced by modifying an egressive airflow with a laryngeal constriction.9, 10 Thus, rodent vocalizations may be a useful model to study voice deficits in a sensorimotor context. Further, rat models allow us to study the neurobiological underpinnings of recovery from deficits with targeted training.
To model PD we use Long-Evans rats (Charles River Laboratories International, Inc.) and induce parkinsonism by a unilateral infusion of 7 &mu;g of 6-hydroxydopamine (6-OHDA) into the medial forebrain bundle which causes moderate to severe degeneration of presynaptic striatal neurons (for details see Ciucci, 2010).11, 12 For our aging model we use the Fischer 344/Brown Norway F1 (National Institute on Aging).
Our primary method for eliciting vocalizations is to expose sexually-experienced male rats to sexually receptive female rats. When the male becomes interested in the female, the female is removed and the male continues to vocalize. By rewarding complex vocalizations with food or water, both the number of complex vocalizations and the rate of vocalizations can be increased (Figure 1).
An ultrasonic microphone mounted above the male's home cage records the vocalizations. Recording begins after the female rat is removed to isolate the male calls. Vocalizations can be viewed in real time for training or recorded and analyzed offline. By recording and acoustically analyzing vocalizations before and after vocal training, the effects of disease and restoration of normal function with training can be assessed. This model also allows us to relate the observed behavioral (vocal) improvements to changes in the brain and neuromuscular system.

Voice disorders are debilitating in aging and Parkinson disease. The ultrasonic vocalizations of rats, also affected by these conditions, can be used to study these voice disorders, their neural substrates, and the nature of functional recovery with behavioral intervention.

Voice deficits are a common complication of both Parkinson disease (PD) and aging; they can significantly diminish quality of life by impacting ...

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.
The Drosophila larva is an established model system to study thermal nociception, a sensory response to potentially harmful temperatures that is evolutionarily conserved across species1,2. The advantages of Drosophila for such studies are the relative simplicity of its nervous system and the sophistication of the genetic techniques that can be used to dissect the molecular basis of the underlying biology4-6 In Drosophila, as in all metazoans, the response to noxious thermal stimuli generally involves a "nocifensive" aversive withdrawal to the presented stimulus7. Such stimuli are detected through free nerve endings or nociceptors and the amplitude of the organismal response depends on the number of nociceptors receiving the noxious stimulus8. In Drosophila, it is the class IV dendritic arborization sensory neurons that detect noxious thermal and mechanical stimuli9 in addition to their recently discovered role as photoreceptors10. These neurons, which have been very well studied at the developmental level, arborize over the barrier epidermal sheet and make contacts with nearly all epidermal cells11,12. The single axon of each class IV neuron projects into the ventral nerve cord of the central nervous system11 where they may connect to second-order neurons that project to the brain.
Under baseline conditions, nociceptive sensory neurons will not fire until a relatively high threshold is reached. The assays described here allow the investigator to quantify baseline behavioral responses or, presumably, the sensitization that ensues following tissue damage. Each assay provokes distinct but related locomotory behavioral responses to noxious thermal stimuli and permits the researcher to visualize and quantify various aspects of thermal nociception in Drosophila larvae. The assays can be applied to larvae of desired genotypes or to larvae raised under different environmental conditions that might impact nociception. Since thermal nociception is conserved across species, the findings gleaned from genetic dissection in Drosophila will likely inform our understanding of thermal nociception in other species, including vertebrates.

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

A Visual Description of the Dissection of the Cerebral Surface Vasculature and Associated Meninges and the Choroid Plexus from Rat Brain

This video presentation was created to show a method of harvesting the two most important highly vascular structures, not residing within the brain proper, that support forebrain function. They are the cerebral surface (superficial) vasculature along with associated meninges (MAV) and the choroid plexus which are necessary for cerebral blood flow and cerebrospinal fluid (CSF) homeostasis. The tissue harvested is suitable for biochemical and physiological analysis, and the MAV has been shown to be sensitive to damage produced by amphetamine and hyperthermia 1,2. As well, the major and minor cerebral vasculatures harvested in MAV are of potentially high interest when investigating concussive types of head trauma. The MAV dissected in this presentation consists of the pial and some of the arachnoid membrane (less dura) of the meninges and the major and minor cerebral surface vasculature. The choroid plexus dissected is the structure that resides in the lateral ventricles as described by Oldfield and McKinley3,4,5,6. The methods used for harvesting these two tissues also facilitate the harvesting of regional cortical tissue devoid of meninges and larger cerebral surface vasculature, and is compatible with harvesting other brain tissues such as striatum, hypothalamus, hippocampus, etc. The dissection of the two tissues takes from 5 to 10 min total. The gene expression levels for the dissected MAV and choroid plexus, as shown and described in this presentation can be found at GSE23093 (MAV) and GSE29733 (choroid plexus) at the NCBI GEO repository. This data has been, and is being, used to help further understand the functioning of the MAV and choroid plexus and how neurotoxic events such as severe hyperthermia and AMPH adversely affect their function.

This video presentation shows a method of harvesting the two most important highly vascular structures that support forebrain function. They are the cerebral surface (superficial) vasculature along with associated meninges (MAV) and the choroid plexus which are necessary for cerebral blood flow and cerebrospinal fluid (CSF) homeostasis.

This video presentation was created to show a method of harvesting the two most important highly vascular structures, not residing within the brain ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls gastrointestinal motility. A procedure for the isolation and culture of a mixed population of neurons and glia from the myenteric plexus is described. The primary cultures can be maintained for over 7 days, with connections developing among the neurons and glia. The longitudinal muscle strip with the attached myenteric plexus is stripped from the underlying circular muscle of the mouse ileum or colon and subjected to enzymatic digestion. In sterile conditions, the isolated neuronal and glia population are preserved within the pellet following centrifugation and plated on coverslips. Within 24-48 hr, neurite outgrowth occurs and neurons can be identified by pan-neuronal markers. After two days in culture, isolated neurons fire action potentials as observed by patch clamp studies. Furthermore, enteric glia can also be identified by GFAP staining. A network of neurons and glia in close apposition forms within 5 - 7 days. Enteric neurons can be individually and directly studied using methods such as immunohistochemistry, electrophysiology, calcium imaging, and single-cell PCR. Furthermore, this procedure can be performed in genetically modified animals. This methodology is simple to perform and inexpensive. Overall, this protocol exposes the components of the enteric nervous system in an easily manipulated manner so that we may better discover the functionality of the ENS in normal and disease states.

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

We demonstrate a cell culture protocol for the direct study of neuronal and glial components of the enteric nervous system. A neuron/glia mixed culture on coverslips is prepared from the myenteric plexus of adult mouse providing the ability to examine individual neuron and glia function by electrophysiology, immunohistochemical, etc.

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls ...

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. Choroid plexus epithelial cells (CPECs) covering the surface of choroid plexus tissue harbor multiple unique cilia, but most of the functions of these cilia remain to be investigated. To uncover the function of CPEC cilia with particular reference to their motility, an ex vivo observation system was developed to monitor ciliary motility during embryonic, perinatal and postnatal periods. The choroid plexus was dissected out of the brain ventricle and observed under a video-enhanced contrast microscope equipped with differential interference contrast optics. Under this condition, a simple and quantitative method was developed to analyze the motile profiles of CPEC cilia for several hours ex vivo. Next, the morphological changes of cilia during development were observed by scanning electron microscopy to elucidate the relationship between the morphological maturity of cilia and motility. Interestingly, this method could delineate changes in the number and length of cilia, which peaked at postnatal day (P) 2, while the beating frequency reached a maximum at P10, followed by abrupt cessation at P14. These techniques will enable elucidation of the functions of cilia in various tissues. While related techniques have been published in a previous report1, the current study focuses on detailed techniques to observe the motility and morphology of CPEC cilia ex vivo.

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

In this study, a detailed light microscopic technique was optimized for real-time observation and analysis of the motion of CPEC cilia ex vivo together with an electron microscopic method for ultrastructural analysis.

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. ...

Localization of Odorant Receptor Genes in Locust Antennae by RNA In Situ Hybridization

Insects have evolved sophisticated olfactory reception systems to sense exogenous chemical signals. These chemical signals are transduced by Olfactory Receptor Neurons (ORNs) housed in hair-like structures, called chemosensilla, of the antennae. On the ORNs&#39; membranes, Odorant Receptors (ORs) are believed to be involved in odor coding. Thus, being able to identify genes localized to the ORNs is necessary to recognize OR genes, and provides a fundamental basis for further functional in situ studies. The RNA expression levels of specific ORs in insect antennae are very low, and preserving insect tissue for histology is challenging. Thus, it is difficult to localize an OR to a specific type of sensilla using RNA in situ hybridization. In this paper, a detailed and highly effective RNA in situ hybridization protocol particularly for lowly expressed OR genes of insects, is introduced. In addition, a specific OR gene was identified by conducting double-color fluorescent in situ hybridization experiments using a co-expressing receptor gene, Orco, as a marker.

Localization of Odorant Receptor Genes in Locust Antennae by RNA In Situ Hybridization

This paper describes a detailed and highly effective RNA in situ hybridization protocol particularly for low-level expressed Odorant Receptor (OR) genes, as well as other genes, in insect antennae using digoxigenin (DIG)-labeled or biotin-labeled probes.

Insects have evolved sophisticated olfactory reception systems to sense exogenous chemical signals. These chemical signals are transduced by Olfactory ...

Cochlear Surface Preparation in the Adult Mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

Targeted Neuronal Injury for the Non-Invasive Disconnection of Brain Circuitry

Surgical intervention can be quite effective for treating certain types of medically intractable neurological diseases. This approach is particularly useful for disorders in which identifiable neuronal circuitry plays a key role, such as epilepsy and movement disorders. Currently available surgical modalities, while effective, generally involve an invasive surgical procedure, which can result in surgical injury to non-target tissues. Consequently, it would be of value to expand the range of surgical approaches to include a technique that is both non-invasive and neurotoxic.
Here, a method is presented for producing focal, neuronal lesions in the brain in a non-invasive manner. This approach utilizes low-intensity focused ultrasound together with intravenous microbubbles to transiently and focally open the Blood Brain Barrier (BBB). The period of transient BBB opening is then exploited to focally deliver a systemically administered neurotoxin to a targeted brain area. The neurotoxin quinolinic acid (QA) is normally BBB-impermeable, and is well-tolerated when administered intraperitoneally or intravenously. However, when QA gains direct access to brain tissue, it is toxic to the neurons. This method has been used in rats and mice to target specific brain regions. Immediately after MRgFUS, successful opening of the BBB is confirmed using contrast enhanced T1-weighted imaging. After the procedure, T2 imaging shows injury restricted to the targeted area of the brain and the loss of neurons in the targeted area can be confirmed post-mortem utilizing histological techniques. Notably, animals injected with saline rather than QA do demonstrate opening of the BBB, but dot not exhibit injury or neuronal loss. This method, termed Precise Intracerebral Non-invasive Guided surgery (PING) could provide a non-invasive approach for treating neurological disorders associated with disturbances in neural circuitry.

The goal of the protocol is to provide a method for producing non-invasive neuronal lesions in the brain. The method utilizes Magnetic Resonance-guided Focused Ultrasound (MRgFUS) to open the Blood Brain Barrier in a transient and focal manner, in order to deliver a circulating neurotoxin to the brain parenchyma.

Surgical intervention can be quite effective for treating certain types of medically intractable neurological diseases. This approach is particularly ...